Molecular Motors

Molecular Motors Edited by M. Schliwa Also of Interest W. Ehrfeld, V. Hessel, H. Löwe Microreactors – New Technology...

Author: Manfred Schliwa

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Molecular Motors Edited by M. Schliwa

Also of Interest W. Ehrfeld, V. Hessel, H. Löwe

Microreactors – New Technology for Modern Chemistry (2000) ISBN 3-527-29590-9

S. P. Nunes and K.-V. Peinemann (Eds.)

Membrane Technology – the Chemical Industry (2001) SBN 3-527-28485-0

J. G. Sanchez Marcano and Th. T. Tsotsis

Catalytic Membranes and Membrane Reactors (2002) ISBN 3-527-30277-8

H. Schmidt-Traub (Ed.)

Chromatographic Separation – Fine Chemicals and Pharmaceutical Agents (planned 2003)

Molecular Motors Edited by Manfred Schliwa

Edited by Prof. Dr. Manfred Schliwa Ludwig-Maximilians-Universität Adolf-Butenandt-Institut Zellbiologie Schillerstrasse 42 80336 München Germany

This book was carefully produced. Nevertheless, editors, authors and publisher do not warrant the information contained therein to be free of errors. Readers are advised to keep in mind that statements, data, illustrations, procedural details or other items may inadvertently be inaccurate. Library of Congress Card No.: applied for A catalogue record for this book is available from the British Library. Bibliographic information published by Die Deutsche Bibliothek Die Deutsche Bibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data is available in the internet at http://dnb.ddb.de. c 2003 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim All rights reserved (including those of translation in other languages). No part of this book may be reproduced in any form – nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not specifically marked as such, are not to be considered unprotected by law. Printed in the Federal Republic of Germany. Printed on acid-free paper.

Composition Hagedorn Kommunikation, Viernheim Printing Druckhaus Darmstadt GmbH, Darmstadt Bookbinding Buchbinderei Schaumann GmbH, Darmstadt ISBN

3-527-30594 -7

Preface

Preface Editors of compilations such as this tend to stress in their prefaces that significant conceptual advances have been made recently, that novel technical developments have opened extraordinary opportunities for unprecedented discoveries and that the time seemed ripe to take stock and to point out developments which will advance the field in the near future. Well, all of this is true for this book too. It is also true that on such occasions we realize how much we have learned and yet how little we know. Since the publication nearly 40 years ago, of the landmark treatise on cell movement edited by Robert D. Allen and Noburô Kamiya and entitled Primitive Motile Systems in Cell Biology, the field has moved from the phenomenological to the mechanistic and from the largely structural to the primarily molecular. We have come to appreciate that at every level of complexity the cell operates through molecular machines. Some of these machines are single molecules that carry out one specific task, undergoing only small structural changes in the process. Others are macromolecular complexes composed of dozens, even hundreds of different components engaged in elaborate biochemical operations. Among the multitude of molecular machines of a cell, one group stands out owing to its ability to generate one of the hallmark characteristics of living systems: movement. The chapters of this book offer insights into the workings, interactions and functions of these remarkable molecules which are responsible for various forms of movement encountered in cells. The subdivision of the book into five sections developed naturally. First we learn about the basic designs of some of the most prominent cellular motors before considering their mechanochemistry; the role of motors in the context of elaborate cellular activities is considered next, followed by examples of defects which result when motors run ‘wild’; finally, biomotors are put into perspective with regard to nanobiotechnological applications and other types of molecular motors. The outcome is a pretty sizeable book, as can plainly be seen. Nevertheless, it is but an introduction to the subject, as other types of biological machines exist that could also, with some justification, be called motors but are not considered here for reasons of space. It is my hope, however, that salient features of cellular motors are covered even though gaps undoubtedly remain. I would like to express my sincere gratitude first and foremost to the authors who have managed to complete their chapters under pretty tight time constraints. I would also like to thank the staff at Wiley-VCH, in particular Dr. Andreas

V

VI

Preface

Sendtko, who have helped me in every respect and to Ursula Euteneuer for critical reading and helpful comments and discussions. Thanks to the efforts of everyone concerned less than one year has elapsed between conception of the book and the completion of the printed product. You might say all of us have motored along just fine. Manfred Schliwa September 2002

Contents

Contents Preface V List of Contributors XIX

Part 1 Basic Principles of Motor Design 1 1.1 1.2 1.2.1 1.2.2 1.2.3 1.3 1.3.1 1.3.2 1.3.3 1.3.4 1.3.5 1.3.6 1.4 1.4.1 1.4.2 1.5 1.6

The Myosin Superfamily: An Overview 3

2 2.1 2.2

Dynein Motors: Structure, Mechanochemistry and Regulation 45 Introduction 45

2.2.1 2.2.2

An Introduction to the Myosin Superfamily 3 Functional Properties of Myosins 7 Directionality and Processivity 7 Protein Motifs Found in Myosins 8 Myosin Regulation 10 Diverse Functions for Myosins 11 Non-muscle Contractile Structures 14 Cell Motility and Adhesion 15 Organelle/Cellular Component Transport 16 Maintenance of Actin-rich Extensions 21 Membrane Trafficking 24 Signal Transduction 26 Myosins in Disease 28 Griscelli Syndrome 28 Roles for Myosins in Hearing 29 New Myosins and Myosin Functions on the Horizon 31 Conclusions 32 References 33

Structural Organization of the Motor, Cargo-binding and Regulatory Components 46 Heavy Chains 48 Intermediate Chains 53

VII

VIII

Contents

2.2.3 2.2.4 2.2.5 2.2.6 2.2.7 2.2.7.1 2.2.7.2 2.2.7.3 2.2.7.4 2.2.8 2.3 2.4 2.5

Light Intermediate Chains 56 The LC8 Light Chain Class 57 The Tctex1/Tctex2 Light Chain Class 59 The LC7/roadblock Light Chain Class 61 Heavy Chain-associated Regulatory Light Chains 62 Light chain 1 62 Calmodulin-related light chains 63 Thioredoxins 64 p29 (cAMP-dependent phosphoprotein) 64 Light Chains Associated with Inner Arms I2/3 65 Mechanochemistry and Motility 65 Dynein Deficiencies and Disease 67 Conclusions 69 References 70

3 3.1 3.2 3.3 3.3.1 3.3.2 3.3.3 3.3.3.1 3.3.3.2 3.3.3.3 3.3.4 3.3.4.1 3.3.4.2 3.3.5 3.3.6 3.3.6.1 3.3.6.2 3.3.7 3.3.8 3.3.8.1 3.3.8.2 3.3.9 3.3.10 3.3.11 3.4 3.5 3.5.1 3.5.2 3.6 3.7

Kinesin Superfamily Proteins 79 Introduction 79

The Kinesin Superfamily Proteins 82 N-Kinesins 87 N-1 Kinesins 87 N-2 Kinesins 91 N-3 Kinesins 91 The Unc104/KIF1 family 91 The KIF13 family 92 The KIF16 family 92 N-4 Kinesins 92 The KIF3 family 93 The Osm3/KIF17 family 94 N-5 Kinesins 94 N-6 Kinesins 94 The CHO1/KIF23 family 95 The KIF20/Rab6 kinesin family 95 N-7 Kinesins 95 N-8 Kinesins 95 The Kid/KIF22 family 95 The KIF18 family 96 N-9 Kinesins 96 N-10 Kinesins 96 N-11 Kinesins 96 M-Kinesins 96 C-Kinesins 97 C-1 Kinesins 97 C-2 Kinesins 97 Orphans 98 Cargoes of KIFs; Specificity and Redundancy 98

Contents

3.8 3.9

Recognition and Binding to Cargoes 99 How to Determine the Direction of Transport 100 References 100

4 4.1 4.2 4.2.1 4.2.2 4.2.3 4.2.4 4.3 4.3.1 4.3.2 4.3.3 4.3.4 4.3.5 4.3.6 4.4 4.4.1 4.4.2 4.5

The Bacterial Flagellar Motor 111 Introduction 111 Structure 114 Propeller and Drive-shaft 117 Rotor 117 Stator 118 Rotor Stator Interactions 119 Function 120 Motor Driven by H and Na Ion Flux 121 Torque versus Speed 122 Independent Torque Generators 126

5 5.1 5.2 5.3 5.4 5.5 5.6 5.7 5.8

6 6.1 6.2 6.3

6.3.1 6.3.2 6.3.3 6.3.4 6.3.5

Proton Motive Force, Sodium-motive Force, Ion Flux 128 Reversibility 131 Steps? 131 Models 132 Conceptual Models 133 Kinetic Models 135 Summary 136 References 137 F1-Motor of ATP Synthase 141

Introduction 141 ATP Synthase 141 F1-Motor 142 Imaging of Rotation of F1-Motor 144 High-speed Imaging of F1 Rotation 145 New Crystal Structure for the F1-Motor 146 Catalysis and Rotation of F1-Motor 148 Perspectives 150 References 151 RNA and DNA Polymerases 153 Introduction 153

NTP Polymerization Mechanism 155 Basic Methods used to Study Polymerase Movement during Transcription 158 The Tethered Particle Motion Approach 158 The Surface Force Microscopy Technique 158 The Optical Tweezer Method 159 Method for Visualization of DNA Rotation during Transcription 161 Footprinting Approach 161

IX

X

Contents

6.3.6 6.4 6.5 6.6 6.7 6.8

7 7.1 7.2 7.3 7.3.1 7.3.2 7.3.3 7.4 7.5 7.6

Single Molecule Assay for DNA Polymerase 162 Mechanism of Force Generation for RNAP and DNAP 164 Molecular Model for RNAP Translocation 168 Possible Utilization of the Energy Released upon NTP Cleavage 171 Single-Molecule Studies and Molecular Mechanisms of Transcription Pausing and Arrest 172 Concluding Remarks 174 References 175 Helicases as Molecular Motors 179 Introduction 179 Basic Properties of Helicases 182 Mechanism of Helicase Activity 188 Unidirectional Translocation 188 Step Size of the Helicase 192 NA Strand Separation 192 HCV Helicase 194 Bacteriophage T7 gp4 Helicase 196 Conclusions 197 References 198

Part 2 Mechanochemistry 8 8.1 8.2 8.3 8.4 8.4.1 8.4.2 8.5 8.6 8.7

How Protein Motors Convert Chemical Energy into Mechanical Work 207 Introduction 207 A Brief Description of ATP Synthase Structure 208 The F1 Motor: A Power Stroke 209 The F0 Motor: A Brownian Ratchet 212 A Pure Brownian Ratchet 212 A Pure Power Stroke 214 Coupling and Coordination of Motors 216 Measures of Efficiency 218 Discussion 220

A1

Example Models to Illustrate the Difference between Ratchets and Power Strokes 221 Example 1: A power stroke without Brownian fluctuations 221 Example 2: A power stroke with Brownian fluctuations 222 Example 3: A Brownian ratchet that biases fluctuations 223 Example 4: A Brownian ratchet that rectifies fluctuations 224

A1.1 A1.2 A1.3 A1.4 A2

A Closer Look at Binding Free Energy 225 References 227

Contents

9 9.1 9.2 9.2.1 9.2.2 9.3 9.3.1 9.4 9.5

Molecular Motor Directionality 229 Introduction 229 Reversed Kinens 229 Chimeric Kinesin Motors 231 A Neck Mutant 233 Backwards Myosins 234 Chimeric Myosin Motors 235 Bidirectional Dyneins? 237 Perspectives 238 References 239

10 10.1 10.2 10.3 10.4 10.5 10.6 10.7 10.8 10.9 10.10 10.11 10.12 10.13 10.14

Kinesins: Processivity and Chemomechanical Coupling 243 Introduction 243 Kinesin Motility and Processivity 244 Biochemical Evidence for Kinesin Processivity 246

11

Quantitative Measurements of Myosin Movement In Vitro: The Reductionist Approach Carried to Single Molecules 271 Introduction 271 Quantitative In Vitro Assays for Myosin Movement Established the Motor Domain of Myosin 272

11.1 11.2 11.3 11.4 11.5 11.6 11.7

Step Size of Kinesin and its Path along the Microtubule 246 Kinesin Stoichiometry 247 Coordination between the Two Heads of Kinesin 248 Testing Processivity with One-headed Kinesin Mutants 249 ATP Hydrolysis Cycle of One-headed Kinesin 250 Structural Studies on Dimeric Kinesin 253 Two-headed Kinesin ATP Hydrolysis Cycle 254 Load Dependent Transitions 257 Ncd is a Non-processive Kinesin Family Member 259 A Processive Monomeric Kinesin, KIF1A 262 Unresolved Questions 264 References 266

Structural Studies Revealed Putative Pre-stroke and Post-stroke States of the Myosin Head 273 Single Molecule Analysis Revealed a Unitary Small Step in Motion as Myosin Interacts with Actin 275 Molecular Genetic Approaches Have Indicated Roles of Various Domains and Specific Residues of the Myosin Motor 277 Myosin V uses its Longer Lever Arm to Take a Larger Step along Actin 278 Conclusions and Perspectives 282 References 283

XI

XII

Contents

12

Structures of Kinesin Motor Domains: Implications for Conformational Switching Involved in Mechanochemical Coupling 287 Introduction 287 Structures of Kinesin Motor Domains 288 General Features of the Catalytic Core 290 The Nucleotide-Binding Active Site 291

12.1 12.2 12.2.1 12.2.2 12.2.2.1 N1, also called P-loop (G86xxxGKS/T, residue numbering according to rat kinesin) 291 12.2.2.2 N2 Switch 1 (N199xxSSR) 291 12.2.2.3 N3 Switch 2 (D232LAGSEKVGKT) 292 12.2.2.4 N4, (R14xRP) 292 12.2.3 Neck Linker, Neck and Hinge 292 12.3 Comparison with G-Proteins and Myosin 293 12.4 Mechanochemical Coupling from a Structural Point of View 294 12.5 Perspectives 300 References 301 13 13.1 13.2 13.3 13.4 13.5 13.6 13.7 13.8 13.9

13.10 13.11

Single Molecule Measurements and Molecular Motors 305 Introduction 305 Manipulation of Actin Filaments 306 Nanometry of Actin Filaments 308

Movement of Actin Filaments Caused by Single Myosin Molecules 309 Visualization of Single Molecules 310 Visualization of ATP Turnover and Mechano-chemical Coupling 312 Visualization of the Movement of Single Kinesin Motors 314 Visualization of the Processive Movement of Single Myosin Motors 317 Manipulation of Single Myosin Molecules with a Scanning Probe and Nanometry 319 Biased Brownian Movement 320 Concluding Remarks 321 References 322

Part 3 Functional Implications 14 14.1 14.2 14.3 14.4 14.5 14.6 14.6.1 14.6.2 14.6.3

Mitotic Spindle Motors 327

Microtubules, Motors and Mitosis 327 The Physical Nature of Mitotic Movements 329 MT Polymerization and Depolymerization as Mitotic Motors 331 Kinesins and Dyneins as Mitotic Motors 334 Functional Coordination of Mitotic Motors 338 Motor Action and Force-Generation during Mitosis 339 Mitotic Motors and Spindle Formation at Early Stages of Mitosis 339 Mitotic Motors and Force Generation in Prometaphase Metaphase 340 Mitotic Motors and Force Generation in Anaphase 343

Contents

14.7 14.8 14.9 14.10

Does a Spindle Matrix Facilitate the Function of Mitotic Motors? 345 Mitotic Motors and Intracellular Transport Systems 346 Mitotic Motors and the Spindle Assembly Checkpoint 349 Conclusions and Future Studies 350 References 351

15 15.1 15.2

The Roles of Molecular Motors in Generating Developmental Asymmetry 357 Introduction 357

16 16.1 16.2 16.3 16.3.1

Motors and Membrane Trafficking 377 Introduction 377

Localization of a L/R Determinant by Asymmetric Flow of Extraembryonic Fluid 357 15.2.1 Situs Inversus in Humans 357 15.2.2 Mice with Mutations in kif3a, kif3b, or lrd Lead to Nodal Flow Model 358 15.2.3 Inv Mutants Challenge the Nodal Flow Model 360 15.3 Asymmetric RNA Localization 361 15.3.1 A/P Patterning in Drosophila Oocytes and Embryos 361 15.3.1.1 Anterior localization of bcd mRNA in the oocyte 362 15.3.1.2 Posterior localization of osk mRNA within the oocyte by kinesin 364 15.3.1.3 Localization of mRNAs in blastoderm embryos 365 15.3.2 Yeast Mating Type Switching 366 15.4 Asymmetric Organelle Localization 368 15.4.1 Localization of the Fusome and Drosophila Oocyte Selection 368 15.4.2 Drosophila Oocyte Nuclear Migration 368 15.4.3 Lipid Droplet Migration in Drosophila Embryos 369 15.4.4 Nuclear Migration in Drosophila Photoreceptors 371 15.5 Future Directions 372 References 372

The Logic and Order of Membrane Trafficking 379 The Cytoskeleton and Motor Proteins in Membrane Trafficking 380 Role of the Cytoskeleton and Motor Proteins in Organelle Localization 380 16.3.2 Role of the Cytoskeleton in Membrane Trafficking Events 380 16.3.3 Role of Motor Proteins in Membrane Trafficking Events 382 16.4 Cooperation between Motors 383 16.4.1 Coordination of Movement along Microtubule and Actin Tracks 383 16.4.2 Coordination of Bidirectional Movement along Microtubule Tracks 385 16.4.3 Molecular Mechanisms for the Coordination of Motors on the Same Transport Cargo 387 16.5 Molecular Mechanisms of Motor–Cargo Linkage 388 16.5.1 Soluble Adaptor or Scaffolding Proteins as Motor Cargo Linkers 388 16.5.1.1 Other scaffolding complexes 392 16.5.2 Motor Cargo Linkage via Members of the Rab Family of Small G-proteins 393

XIII

XIV

Contents

Other Mechanisms for Linking Microtubule-based Motors to their Cargoes 395 16.5.3.1 Attachment to the membrane cytoskeleton 395 16.5.3.2 Attachment via integral membrane proteins 396 16.6 Regulation of Motor Activity 397 16.6.1 Motor Proteins must be Regulated at Several Steps of their Transport Cycle 397 16.6.2 Molecular Mechanisms for Regulating Motor Activity 399 16.7 Concluding Remarks 400 References 401 16.5.3

17 17.1 17.2 17.2.1

17.2.1.1 17.2.1.2 17.2.1.3 17.2.1.4 17.2.1.5 17.2.2 17.2.2.1 17.2.2.2 17.2.2.3 17.3 17.3.1 17.3.1.1 17.3.1.2 17.3.2 17.3.3 17.3.4 17.4 17.4.1 17.4.2 17.4.3 17.4.4

Regulation of Molecular Motors 411 Introduction 411

The Role of Phosphorylation in Regulating Molecular Motors 411 Phosphorylation can Control Motor Organelle or Motor Spindle Binding 412 Interaction of dynein with organelles can be regulated by phosphorylation 412 Interaction of dynein with dynactin can be regulated by phosphorylation 412 Interaction of kinesin with organelles can be regulated by phosphorylation 413 Interaction of kinesin family members with the spindle can be regulated by phosphorylation 414 The binding of myosin V to melanosomes is regulated by phosphorylation 414 The Activity of Motors may be Regulated by Phosphorylation 416 Phosphorylation of axonemal dynein inhibits its activity 416 Phosphorylation can activate or inhibit cytoplasmic dynein 416 Phosphorylation can activate or inhibit the activity of members of the kinesin family 417 The Role of G Proteins in Regulating Molecular Motors 419 G Proteins Mediate Motor Cargo Interactions 419 Rab27a recruits myosin Va to melanosomes 419 G proteins may recruit microtubule motors to organelles 421 G Proteins may Activate Motors 422 Motors may Bind Directly to G Proteins but the Function of these Interactions Remains Unclear 422 A Light Chain of Dynein may be Involved in Regulating G Protein GTPase Activity 423 Other Mechanisms of Regulation 424 Kinesin Folding 424 Lis 1 Interaction with Cytoplasmic Dynein 425 Motor Protein Regulation during the Cell Cycle 425 Motor Complexes and Coordination 426

Contents

17.5

Summary 427 References 427

18 18.1 18.2 18.2.1 18.2.1.1 18.2.2 18.3 18.3.1 18.3.2 18.4 18.4.1 18.4.2 18.4.3 18.4.4 18.4.5 18.4.6 18.5 18.5.1 18.5.2 18.6

Molecular Motors in Plant Cells 433 Introduction 433 Microtubule-based Motors 434 Kinesin-like Proteins 434 Phylogenetic analysis 443 Dyneins 447 Actin-based Motors 448 Myosins 448 Phylogenetic analysis 451 Cellular Roles of Motors 451 Cell Division 451 Cell Polarity and Morphogenesis 456 Cytoplasmic Streaming 457

Microtubule Dynamics and Organization 458 Intercellular Transport 459 Other functions 459 Regulation of Motors 460 Calcium/Calmodulin 460 Protein Phosphorylation 461 Concluding Remarks 461 References 462

Part 4 Motors in Disease 19 19.1 19.2 19.2.1 19.2.2 19.3 19.3.1 19.3.2 19.4 19.4.1 19.4.2 19.4.3 19.4.5 19.5 19.6

Myosin Myopathies 473

Introduction: Inherited Myosin Myopathy 473 Cardiac Myosin Heavy Chains 475 MyHC Structure and Function 475 Cardiac Muscle Regulation and Disease 477 Cardiac MyHC Myopathy 478 Functional Characterization of MyHC Motor Domain Mutations 478 Transgenic Models of Myosin-based FHC 480 MyHC Interacting Proteins and FHC 482 The Essential and Regulatory Light chains 482 Myosin Light Chain-based FHC 484 Myosin Binding Protein C-Based FHC 485 Titin-based Familial Hypertrophic Cardiomyopathy 487 Myosin-based Myopathies in Skeletal Muscle 488 Conclusions 489 References 490

XV

XVI

Contents

20 20.1 20.2 20.3 20.4

The Role of Dynein in Disease 497

21 21.1 21.2 21.3 21.4 21.5 21.5.1 21.5.1.1 21.5.1.2 21.5.1.3 21.5.1.4 21.5.2 21.5.2.1 21.5.2.2 21.5.2.3 21.5.3 21.5.3.1 21.5.3.2 21.5.3.3 21.5.4 21.5.4.1 21.5.4.2 21.5.5 21.5.5.1 21.5.5.2 21.5.5.3 21.6

Molecular Motors in Sensory Defects 511 Introduction 511

Dynein Functional and Structural Classes 497 Diseases Associated with Axonemal Defects 498 Role of a Cytoplasmic Dynein Light Chain in Retinitis Pigmentosa 500 Role of Cytoplasmic Dynein in the Smooth Brain Disease Lissencephaly 501 References 506

Development of the Visual and Auditory Sensory Systems 511 Visual Impairment 513 Hearing Impairment 514 Myosins Involved In Sensory Defects 515 Myosin VIIA 516 Structure, function, and expression of myosin VIIA 516 Shaker 1 Mice and Other Models 518 Usher Syndrome Type 1B 519 DFNB2 and DFNA11 520 Myosin VI 520 Structure, function, and expression of myosin VI 520 Snell’s waltzer mice and other models 522 DFNA22 524 Myosin XVA 525 Structure, function, and expression of myosin XVA 525 Shaker 2 526 DFNB3 527 MYH9 528 Structure, function, and expression of MYH9 528 DFNA17 528 Myosin IIIA 529 Structure, function, and expression of myosin IIIA 529 Myosin IIIA mutants 531 DFNB30 531 Concluding Remarks 532 References 533

Contents

Part 5 Beyond Biological Applications 22 22.1 22.2 22.2.1 22.2.2 22.2.3 22.2.4 22.2.5 22.3

Systematized Engineering of Biomotor-powered Hybrid Devices 541 Introduction 541 The Core Technologies 543 Nanoscale Directed Assembly 543 Molecular Energy Transduction 546 Control Mechanisms 549 Multimedia Device Construction 552 Engineering Issues 554 The Core Technologies as a Whole 556 References 557

23 23.1 23.2 23.3 23.4 23.5 23.6

Synthetic Molecular Motors 559 Introduction 559

Translational Synthetic Molecular Motors 559 Synthetic Rotary Molecular Motors 564 Chemically Driven Unidirectional Molecular Motor 567 Light-driven Unidirectional Molecular Motors 568 Conclusion and Prospects 575 References 575 Index 579

XVII

Also of Interest W. Ehrfeld, V. Hessel, H. Löwe

Microreactors – New Technology for Modern Chemistry (2000) ISBN 3-527-29590-9

S. P. Nunes and K.-V. Peinemann (Eds.)

Membrane Technology – the Chemical Industry (2001) SBN 3-527-28485-0

J. G. Sanchez Marcano and Th. T. Tsotsis

Catalytic Membranes and Membrane Reactors (2002) ISBN 3-527-30277-8

H. Schmidt-Traub (Ed.)

Chromatographic Separation – Fine Chemicals and Pharmaceutical Agents (planned 2003)

List of Contributors

List of Contributors Karen B. Avraham Department of Human Genetics and Molecular Medicine Sackler School of Medicine Tel Aviv University Tel Aviv 69978 Israel Richard Berry The Clarendon Laboratory University of Oxford Parks Road Oxford OX1 3PU UK Richard A. van Delden Department of Organic and Molecular Inorganic Chemistry Stratingh Institute University of Groningen Nijenborgh 4 9747 AG Groningen The Netherlands Sharyn A. Endow Duke University Medical Center Department of Microbiology P. O. Box 3020 Durham, NC 27710 USA

Ben L. Feringa Department of Organic and Molecular Inorganic Chemistry Stratingh Institute University of Groningen Nijenborgh 4 9747 AG Groningen The Netherlands Janice A. Fischer Section of Molecular Cell and Developmental Biology Institute for Cellular and Molecular Biology The University of Texas at Austin Austin, TX 78712 USA Leah T. Haimo Department of Biology Unversity of California Riverside, CA 92521 USA William O. Hancock Department of Bioengineering Pennsylvania State University 218 Hallowell Building University Park, PA 16802 USA

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Nobutaka Hirokawa Department of Cell Biology and Anatomy Graduate School of Medicine University of Tokyo Hongo 7-3-1, Bunkyo-ku Tokyo 113-0033 Japan

John P. Konhilas Department of Molecular, Cellular, and Developmental Biology University of Colorado at Boulder Campus Box 347 Boulder, CO 80309-0347 USA

Nataliya Korzheva The Public Health Research Institute Jonathon Howard International Center for Public Health Max Planck Institute of Molecular Cell Biology and Genetics Alex Goldfarb’s Laboratory 225 Warren Street Pfotenhauerstrasse 108 Newark, NJ 07103-3535 01307 Dresden USA Germany Yoshiharu Ishii Single Molecule Processes Project ICORP, JST 2-4-14 Senba-higashi, Mino Osaka 562-0035 Japan Michele C. Kieke Department of Genetics, Cell Biology and Development University of Minnesota 6-160 Jackson Hall 321 Church Street SE Minneapolis, MN55455 USA Stephen M. King Department of Biochemistry University of Connecticut Health Center 263 Farmington Avenue Farmington, CT 06030-3305 USA

Nagatoshi Koumura Department of Organic and Molecular Inorganic Chemistry Stratingh Institute University of Groningen Nijenborgh 4 9747 AG Groningen The Netherlands Leslie A. Leinwand Department of Molecular, Cellular, and Developmental Biology University of Colorado at Boulder Campus Box 347 Boulder, CO 80309-0347 USA Mikhail K. Levin Department of Biochemistry Robert Wood Johnson Medical School Piscataway, New Jersey NJ 08854 USA


E. Mandelkow Max-Planck-Unit for Structural Molecular Biology Notkestrasse 85 22607 Hamburg Germany A. Marx Max-Planck-Unit for Structural Molecular Biology Notkestrasse 85 22607 Hamburg Germany A. Mogilner Center for Genetics and Development Department of Mathematics University of California at Davis Davis, CA 95616 USA Carlo D. Montemagno Department of Bioengineering University of California Los Angeles Los Angeles, CA 90023 USA Arkady Mustaev The Public Health Research Institute International Center for Public Health Alex Goldfarb’s Laboratory 225 Warren Street Newark, NJ 07103-3535 USA Hiroyuki Noji Institute of Industrial Science University of Tokyo 4-6-1, Komaba Meguro-ku Tokyo 153-8505 Japan

George Oster Department of Molecular and Cell Biology 201 Wellman Hall University of California Berkley, CA 94720-3112 USA Smita S. Patel Department of Biochemistry Robert Wood Johnson Medical School Piscataway, New Jersey NJ 08854 USA A. S. N. Reddy Department of Biology and Program in Cell and Molecular Biology Colorado State University Fort Collins, Colorado 80523 USA J. M. Scholey Center for Genetics and Development Section of Molecular and Cellular Biology University of California at Davis Davis, CA 95616 USA Y.-H. Song Max-Planck-Unit for Structural Molecular Biology Notkestrasse 85 22607 Hamburg Germany Jacob J. Schmidt Department of Bioengineering University of California Los Angeles Los Angeles, CA 90023 USA

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James A. Spudich Department of Biochemistry and Department of Developmental Biology Stanford University School of Medicine Stanford, CA 94305 USA

Kristen J. Verhey University of Michigan Medical School Dept. of Cell and Dev. Biology 1335 Catharine St. Ann Arbor, MI 48109-0616 USA

Chin-Yin Tai Department of Cell Biology University of Massachusetts Medical School 377 Plantation St. Worcester, MA 01605 USA

Hongyun Wang University of California Santa Cruz 1156 High Street Santa Cruz, CA 95064 USA

Reiko Takemura Department of Cell Biology and Anatomy Graduate School of Medicine University of Tokyo Hongo 7-3-1, Bunkyo-ku Tokyo 113-0033 Japan Margaret A. Titus Department of Genetics, Cell Biology and Development University of Minnesota 6-160 Jackson Hall 321 Church Street SE Minneapolis, MN55455 USA Richard B. Vallee Department of Pathology Columbia University College of Physicians and Surgeons P & S 15-409 630 W. 168th St. New York, NY 10033 USA

Matthijs K. J. ter Wiel Department of Organic and Molecular Inorganic Chemistry Stratingh Institute University of Groningen Nijenborgh 4 9747 AG Groningen The Netherlands Toshio Yanagida Single Molecule Processes Project ICORP, JST 2-4-14 Senba-higashi, Mino Osaka 562-0035 Japan

Part 1 Basic Principles of Motor Design

1 The Myosin Superfamily: An Overview Michele C. Kieke and Margaret A. Titus

1.1

An Introduction to the Myosin Superfamily

Actin filaments and the myosin motors associated with them play important roles in many dynamic biological processes. The classic example of actin filaments and myosin at work is during skeletal muscle contraction. But the functions of actin and myosin extend to many other cellular events, such as motility, adhesion, endocytosis, cytoplasmic streaming, neuron growth, structural maintenance and polarization. Like molecular cars on an actin track, myosins transport organelles and other cellular components, such as mRNA. Myosins can also aid in the formation or maintenance of an organized actin-based structures (such as the stereocilia of hair cells), and play roles in intracellular signal transduction pathways (see Baker and Titus 1998, Mermall et al. 1998, Sokac and Bement 2000, Tuxworth and Titus 2000, for reviews summarizing myosin functions). Myosins are molecular motor proteins that use the energy from adenosine triphosphate (ATP) hydrolysis to generate force for directed movement along actin filaments (see Chapter 11 by Spudich and Chapter 13 by Ishii and Yanagida). Myosins are composed of one to two heavy chains, and one or more light chains. The heavy chain consists of several major domains and can include various other subdomains or protein motifs (Fig. 1.1). The relatively conserved N-terminal motor or ‘head’ domain has binding sites for both ATP and F-actin. A short region joining the head and neck (termed the ‘converter domain’) is believed to be responsible for producing the force required for movement. The neck domain contains one to six light chain binding regions termed IQ motifs, repeats of approximately 23 to 30 residues containing the sequence IQXXXRGXXXRK (Bähler and Rhoads, 2002). The divergent C-terminal globular ‘tail’ has been implicated in binding cargo and targeting the myosin to its proper location in the cell (Karcher et al., 2002). Some myosins also feature a coiled-coil domain that promotes heavy chain dimerization. The founding member of the myosin family, filament-forming class II muscle myosin, was discovered nearly a century ago, and its role in muscle contraction

4

1.1 An Introduction to the Myosin Superfamily

Myosin

Structural features* Motor

Neck

Tail

1-6 polybasic region

M1

+++

GPA

M2

SH3

C 1-2

M3 M4

SH3 x5

C

M5 M6

C x5

C

M7

SH3

x3

C

M8 x4

Zn+2

M9 x3

M10

x3

C

PH

C

C

x6

M11 M12

C x6

M13

+++

M14

++

M15 M16

SH3 x8

M17 M18

Key

Pro

ANK

chitin synthase domain

C

PDZ

C = coiled-coil region PH = pleckstrin homology domain SH3 = Src homology 3 domain Zn+2 = zinc-binding domain Pro = proline-rich region + + + = positively-charged region GPA = Gly, Pro, Ala-rich region = PEST site

ANK = ankyrin repeat

= rho-GAP domain = IQ motif = FERM domain = MyTH4 domain = N-terminal extension = protein kinase domain * not drawn to scale

1 The Myosin Superfamily: An Overview m Figure 1.1. Domain structure schematic for characterized myosin genes. Schematic illustrating

the variety of known structural motifs found in myosin genes. A general box diagram is given for each class, although individual members of the same class may vary depending on the organism and/or particular isoform.

has been studied extensively (Geeves and Holmes, 1999, Huxley, 2000). A combination of biochemical and molecular approaches has led to the identification of over 20 different myosin classes (Berg et al., 2001). Because of the extensive amount of knowledge acquired regarding the properties of myosin II it is referred to as ‘conventional’ myosin; all other types of myosin are referred to as ‘unconventional’. The first unconventional myosin, myosin I, was described in 1973 by Pollard and Korn (Pollard and Korn, 1973a, 1973b). They isolated a protein with enzymatic properties similar to myosin II (i. e. it exhibited actin-activated Mg2 -ATPase and ATP-sensitive binding to actin) from the common freshwater amoeba Acanthamoeba castellanii that had a lower molecular weight than muscle myosin II (125 versus 200 kD) and did not form filaments. In addition, it was determined that myosin I had one head rather than two. Careful analysis of this unusual molecule revealed that it was indeed a bona fide myosin (Korn, 1991). This work provided the first insights into the potential diversity and functions of the myosin superfamily. Myosin superfamily members are grouped into different classes based on phylogenetic analysis of motor domains (Fig. 1.2) (Berg et al., 2001, Cheney and Mooseker, 1992, Goodson and Spudich, 1993,). Each class is designated by a Roman numeral, largely in the order of their discovery (note that we will refer to myosins using Arabic numbers for simplicity). A total of 18 classes have been officially designated, but there are at least six novel myosins that have yet to be classified. Myosins have been found in a variety of organisms but no one class is universally expressed in all phyla. The yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe have a total of five myosin genes from three classes (M1, M2, M5). These myosins are shared by higher organisms, ranging from Caenorhabditis elegans (C. elegans) to mammals. The human genome includes about 40 myosin genes from 12 classes (Berg et al., 2001). M8, M11, and M13 are only found in plants (see Chapter 18 by Reddy; Reddy and Day, 2001), and M14 myosins are found in parasites such as Toxoplasma gondii and Plasmodium falciparum (Berg et al., 2001). A unique class of myosins (as yet undesignated) has been found in the ciliated protozoan Tetrahymena (Garcés and Gavin, 1998, Williams et al., 2000), suggesting that these organisms have a distinctive set of myosins. Cells typically express multiple myosins the expression of at least a dozen myosins in a single cell type has been described (Bement et al., 1994). This includes several different classes of myosin, as well as two or more isoforms of several classes. Myosins from the same class can have isoform-specific roles, such as M5 isoforms in mammalian cells and yeast (Reck-Peterson et al., 2000), or they can have functionally overlapping roles, such as the M1s in Dictyostelium discoideum and Saccharomyces (Geli and Riezman, 1996, Goodson et al., 1996, Jung et al., 1996, Novak et al., 1995).

5

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1.1 An Introduction to the Myosin Superfamily

Abbreviations: Ac: Acl: An: At: Ce: Dd: Dm:

Acanthamoeba castellani Acetabularia cliftonii Aspergillus nidulans Arabidopsis thaliana Caenorhabditis elegans Dictyostelium discoideum Drosophila melanogaster

Hs: Homo sapiens Lp: Limulus polyphemus Mm: Mus musculus Pf: Plasmodium falciparum Rn: Rattus norvegicus Sc: Saccharomyces cerevesiae Sp: Schizosaccharomyces pombe

Figure 1.2. Unrooted myosin superfamily phylogenetic tree. Phylogenetic tree from Berg et al., 2001, constructed using myosin motor domain sequences. Species names are listed in the Abbreviations table, and some gene names have been shortened to save space. Sequences

Tg: Toxoplasma gondii Tt: Tetrahymena thermophila CSM: Chitin synthase myosin HMWM: High molecular weight myosin MysPDZ: PDZ myosin

predicted in full or in part from genomic clones are indicated by an asterisk. Figure and legend text reprinted from Molecular Biology of the Cell (2001, 12: 780 794), with permission from the American Society for Cell Biology.

1 The Myosin Superfamily: An Overview

The existence of not only diverse myosin classes but also isoforms within those classes is a clear indication of the range of roles for this motor protein superfamily. Myosins have been implicated in many fundamental biological processes, and their functional significance is accented by the discovery that mutations in unconventional myosin genes in worms, flies, mice, and humans cause severe phenotypes such as deafness, blindness, and sterility. Despite the important and diverse roles for myosins, much remains to be learned about them at the molecular level. Only in the last decade have the intricacies of myosins been realized due to the characterization of properties such as structure, kinetics, localization, directionality, and putative functional roles. In the past five years, the rate of discovery in the myosin field has increased significantly we are learning much about what myosins do and how they work. This chapter will summarize what is currently known about the myosin superfamily members, including their cellular functions and their roles in various diseases.

1.2

Functional Properties of Myosins

The motor domains of myosins are relatively well-conserved, although kinetic properties can vary from class to class. Some of the myosins are short-duty motors that most likely function as part of an assembly of motors (e. g. M2). Others are processive and can move for long distances without releasing from the actin filament (e. g. M5). It should be noted, however, that not all members of the same class have similar properties. For example, mammalian M5 is highly processive while yeast M5 is not (Mehta et al., 1999, Reck-Peterson et al., 2001). Myosins can move toward one end of the actin filament or the other, a property referred to as ‘directionality’. In contrast to the conservation between motor regions, the tail regions are quite divergent between the different myosin classes. A variety of protein motifs are found in the myosin tail region and a few are also located in the motor domain. These kinetic and structural properties can offer insight into myosin functions. 1.2.1

Directionality and Processivity

The directionality and processivity of myosins contribute to their diverse cellular functions (Higuchi and Endow, 2002). Myosins move unidirectionally on actin filaments, either toward the plus-end or the minus-end of actin. The arrangement of actin filaments in the cell periphery is generally with the plus-ends (fast-growing) toward the plasma membrane. Myosins moving toward the plus-ends (i. e. M1, M5) would be expected to carry cargo and/or to localize at the cell periphery. On the other hand, myosins that move in the opposite direction, such as M6 and M9 (Inoue et al., 2002a, Wells et al., 1999), would be predicted to have complementary roles. Perhaps minus-end directed myosins provide a means to transport cargos

7

8

1.2 Functional Properties of Myosins

into the cell rather than toward the periphery, and thus provide an opposing activity for plus-end directed motors. In addition to the directionality of the motor, the time spent bound to actin filaments is an important consideration when thinking about myosin functions. ‘Duty ratio’ refers to the filament-bound state of myosin, or more specifically to the fraction of the ATPase cycle that myosin spends strongly bound to actin. Myosins that spend a significant proportion of the cycle tightly bound to actin have a ‘high-duty ratio’, while those that spend a small fraction of the cycle tightly bound are considered to have a ‘low-duty ratio’. Conventional muscle M2 is a low-duty motor (Geeves and Holmes, 1999), a property reflected in the fact that large assemblies of these motors work in coordination with each other during muscle contraction. A low-duty motor would not be suitable for persistent transport of cargo, as it would detach after a single step on actin. In contrast, motors with a high-duty ratio are good candidates to serve as long-range transport motors. Such single myosin molecules can translocate along an individual actin filament for long distances. It is also worth noting that high-duty ratio is a trait that would be advantageous for a motor suspected to provide force and maintain tension. High-duty ratio motors can be identified by detailed kinetic analyses or by in vitro motility assays of single myosin molecules. Myosins from at least three classes are processive M5 (Mehta et al., 1999), M6 (De La Cruz et al., 2001, Nishikawa et al., 2002 Rock et al., 2001), and M9b (Inoue et al., 2002a). M5 and M6 are two-headed myosins, while M9b is a single-headed motor. This structural difference suggests distinct mechanisms for processive movement. 1.2.2

Protein Motifs Found in Myosins

The myosins possess a variety of known protein motifs at either their N- or C-terminus (see Fig. 1.1) that offer insight into possible myosin function and localization. These motifs can be generally categorized as membrane localization or targeting, signaling, structural, or myosin-specific. A major effort is currently underway to identify the contribution of each of these regions to myosin function. Several classes of myosin have intriguing N-terminal domains. The motor domain of M3 has a protein kinase domain that is required for its function in the termination of the phototransduction signal cascade in the Drosophila eye (Bähler, 2000). M16 has a series of ankyrin repeats at its N-terminus (Patel et al., 2001), and M18 has an N-terminal PDZ domain (Furusawa et al., 2000, Yamashita et al., 2000). The role of these two different domains in mediating protein protein interactions suggests that the head may play a role in myosin localization apart from simply binding to actin. M15 has a unique extension at its N-terminus (Liang et al., 1999), the function of which remains to be determined but could play a role in its motor function and/or localization. The known membrane localizing motifs in the myosin tail are PH (pleckstrin homology), polybasic regions, and FERM (band 4.1/ezrin/radixin/moesin) do-


mains. PH domains bind to the charged head groups of phospholipids in the plasma membrane and are often involved in signal transduction pathways. Thus, myosins with PH domains (M10 and the novel myoM from Dictyostelium (Berg et al., 2000, Geissler et al., 2000, Oishi et al., 2000) might act as adapters or relay proteins in signal transduction cascades, functioning to localize signaling proteins to the plasma membrane. The M1s all have a polybasic domain that binds to anionic phospholipids with high affinity (Mermall et al., 1998, Titus, 1997), suggesting that this region directs the localization of this myosin class to a particular subset of intracellular membranes. It should be noted, however, that this domain also binds to actin and might be able to contribute to the association of M1s to actin filaments (Lee et al., 1999, Liu et al., 2000). FERM domains were first identified as the N-terminal region of the ERM (ezrin, radixin, moesin) family of actin-binding proteins that shares homology with the Nterminus of band 4.1 (Chishti et al., 1998). The band 4.1 protein family members participate in cell signaling events that regulate processes such as cell growth, cytoskeleton dynamics, and cell-substrate adhesion (Bretscher et al., 2000, Hoover and Bryant, 2000, Tsukita and Yonemura, 1999). The FERM domains of ERM proteins bind to phospholipids as well as the cytoplasmic region of transmembrane receptors such as the CD44 adhesion molecule, linking them to the actin cytoskeleton (Bretscher et al., 2000, Tsukita and Yonemura, 1999). Talin, another FERMdomain containing protein, binds both the cytoplasmic tail of integrins via its FERM domain and the actin cytoskeleton via its C-terminus (Horwitz et al., 1986), forming mechanical links between the two components (similar to the ERMs). The M7, M10, M12, and M15s each have one or two FERM domains in their C-terminus (Oliver et al., 1999). Several different classes of myosins contain motifs in their tail regions that are commonly found in signaling proteins. These include rho-GAP (rho-GTPase-activating protein) domains and SH3 (src homology 3) domains. M9s possess a rhoGAP domain (Müller et al., 1997, Post et al., 1998). The rho-GTPases have been reported to regulate actin cytoskeleton organization (Ridley, 2001), suggesting that M9 may act as both a motor and a signaling molecule. M4 and one subtype of M1 (the amoeboid type) each have a single SH3 domain (Oliver et al., 1999). The SH3 domain of the amoeboid M1 binds to proteins that recruit the Arp2/3 complex, implicating these motors in regulating or contributing to the directed polymerization of actin in yeast and Dictyostelium (Evangelista et al., 2000, Jung et al., 2001, Lechler et al., 2000, Xu et al., 1997). There are additional tail motifs that contribute either to the overall structure or function of given myosins. A number of unconventional myosins (M5, M6, M7, M8, M10, M11, M12, and M18) have stretches of predicted coiled-coil, indicating that these myosins function as dimers. The amoeboid M1s have a domain rich in the amino acids glycine, proline, and alanine (or serine or glutamate) that binds to actin with high affinity (Pollard et al., 1991). The M9s have one or more zinc-binding regions in the tail region (Müller et al., 1997, Post et al., 1998). Zinc-binding motifs are also found in several signaling proteins such as protein kinase C, raf kinase, and Vav (Chieregatti et al., 1998). Mammalian M5 and M10 both have PEST regions in

9

10

1.2 Functional Properties of Myosins

their tails (Berg et al., 2000, Espreafico et al., 1992), sequences enriched in the amino acids proline, glutamate, serine, and threonine. It has been suggested that the PEST regions are susceptible to cleavage by calpain, a calcium-dependent protease (reviewed in Rechsteiner and Rogers, 1996). If this cleavage occurs in vivo it would result in separation of the motor domain from the tail. One of the most ubiquitous and least characterized domains found in myosins is the MyTH4 (myosin tail homology) domain. MyTH4 domains are found not only in myosins (M4, M7, M10, M12, M15) but also in a kinesin from plants (Oliver et al., 1999) as well as in a protein that participates in signaling processes that occur during axon guidance (Huang et al., 2002). The precise role for the MyTH4 motif is not known, but preliminary evidence suggests that MyTH4 domains might bind to microtubules (Narasimhulu and Reddy, 1998, Oliver et al., 1999). In many cases the MyTH4 domain immediately precedes a FERM domain, suggesting that these two domains together form a functional unit. Evidence in support of a critical role for this domain comes from the finding that mutations in the MyTH4 domains of M7 and M15 cause disease (Liu et al., 1998b, Liang et al., 1999). Finally, both M5 and M6 have unique globular tail domains at their extreme Ctermini that direct the intracellular localization of each of these myosins (Buss et al., 2001, Reck-Peterson et al., 2000). In the case of M5a, there are several unique exons found only in M5a expressed in the skin (Separack et al., 1995). The skinspecific exon has been shown to play an essential role in targeting M5a to the melanosome (Wu et al., 1998). 1.2.3

Myosin Regulation

Motor function must be precisely regulated (see Chapter 17 by Haimo). The activity of myosins is regulated by either calcium or phosphorylation (Sokac and Bement, 2000). Most myosin light chains (LCs) are calmodulin or members of the calmodulin superfamily that bind to IQ motifs in the neck region. Myosins have from one to six IQ motifs per heavy chain (HC) and the LCs bind in the absence of calcium. Calcium binding to the associated calmodulin can result in a weakening of affinity and may result in LC loss. Binding also influences the actin-activated ATPase activity, actin binding, and can result in either reduced or increased translocation velocity of actin filaments in in vitro motility assays (Tauhata et al., 2001, Wolenski, 1995). The relationship between calcium binding to calmodulin LCs, actin binding, actin-activated Mg2 -ATPase activity (AAA) and in vitro motility is different for each myosin. In the case of M5, micromolar calcium concentration results in higher AAA and an increased affinity for actin, but decreased motility (Cheney et al., 1993, Tauhata et al., 2001). In contrast, higher calcium levels result in both decreased AAA and motility. The reader is referred to several recent reviews for a more detailed discussion of the effects of calcium-calmodulin regulation of various myosins (Barylko et al., 2000, Reilein et al., 2001, Sokac and Bement, 2000, Wolenski, 1995).


Regulation by HC phosphorylation occurs in the motor domain of amoeboid M1 and M6 at a residue called the ‘TEDS rule’ site (Bement and Mooseker, 1995) that resides 16 residues N-terminal from the conserved DALAK sequence located near the actin-binding site sequence. The TEDS site residue is usually either phosphorylatable (amino acids T or S) or acidic (E or D), thus the ‘TEDS’ designation. The importance of phosphorylation in regulating unconventional myosin activity is highlighted by studies of amoeboid M1s in fungi or amoebae. These motors require phosphorylation by a PAK family kinase that is regulated by small G-proteins for actin-activated ATPase activity (Brzeska and Korn, 1996). It should be noted that phosphorylation regulates the activity of only the lower eukaryotic forms of M1 because only these M1s have an S or T at the TEDS rule site. Mutation of the TEDS rule site to A in either yeast or Dictyostelium amoeboid M1 abolishes in vivo function (Novak and Titus, 1997, 1998 Wu et al., 1997a). Phosphorylation is not essential for the activity of all amoeboid M1s. Mutation of the TEDS rule site of Aspergillus MYOA to A results in a loss of virtually all AAA and motor activity, yet does not impair the in vivo function of this myosin (Liu et al., 2001). This suggests that some M1s may play a non-motor role in certain contexts. The higher eukaryotic M1s have an E or D at that site and their regulation is likely to occur via Ca2 -calmodulin rather than HC phosphorylation. Mammalian M6 is also phosphorylated by a PAK kinase at the TEDS rule site and this has been shown to moderately activate M6 motility in vitro as well the rate of Pi release (Buss et al., 1998, De La Cruz et al., 2001, Yoshimura et al., 2001). The exact relationship between changes in the rates of in vitro motility and alterations in the kinetic properties of phosphorylated M6 are not yet clear, but ongoing investigations should resolve this issue. Recent experiments have revealed that phosphorylation in the myosin tail region can regulate the activity of myosins not by affecting enzymatic activity, but by changing the affinity of the myosin for its cargo. Calcium/calmodulin dependent kinase II (CaMKII) phosphorylates a serine in the C-terminal organelle-binding domain of Xenopus M5 in a cell-cycle dependent manner (Karcher et al., 2001, Rogers et al., 1999). The onset of mitosis results in an increase in M5 tail phosphorylation and a concomitant decrease in the level of M5 associated with melanosomes in melanophores. The increased phosphorylation is predicted to cause cargo (melanosome) release from M5 and the treatment of mitotic Xenopus extracts with CaMKII inhibitors strongly inhibits M5-melanosome dissociation. HC phosphorylation in the myosin tail region may be one mechanism for regulating myosins involved in organelle transport.

1.3

Diverse Functions for Myosins

Myosins participate in diverse biological functions (Tab. 1.1). While a number of different roles for myosins have been defined over the past few years, it has become increasingly clear with the identification of additional novel myosins that

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12

1.3 Diverse Functions for Myosins Summary of putative myosin functions. A brief listing of the known functions of several different myosin classes. The question marks (?) indicate functions that have been inferred based on localization, initial phenotypic analyses and analysis of phenotypes resulting from ectopic expression of wild-type or mutant forms of a given myosin.

Table 1.1

Myosin

Putative role

Relevant organism or cell type/gene

Reference(s)

M1

Cell motility (leading edge)

Acanthamoeba, Dictyostelium, mammalian cells Intestinal epithelial cells/MYO1A Intestinal epithelial cells/MYO1A Dictyostelium, yeast Polarized epithelial cells/MYO1D Dictyostelium/myoB Stereocilia in mouse/MYO1C

1

Dictyostelium, mammal. cells, echinoderm Fission yeast/MYO2 and MYP2; budding yeast/MYO1 Drosophila and Dictyostelium Mammalian cells Acanthamoeba, Dictyostelium, mammalian cells

8

Microvilli structure Vesicle transport, endocytosis Endocytosis Endosome recycling Early endosome recycling Signal transduction M2

Cytokinesis

Morphogenesis Cortical bundles, stress fibers Cell motility (rear of cell)

2 3 4 5 6 7

9 10 11 12

M3

Cell integrity and structure Signal transduction Signal transduction (?)

Drosophila photoceptor cells/ninaC Drosophila eye/ninaC Limulus eye

13 14 15

M5

Secretory vesicle transport Vacuole transport/inheritance Golgi and peroxisome inheritance ASH1 mRNA transport Mitotic spindle orientation Particle transport SER and vesicle transport Pigment granule transport Particle transport, filopod extentsion Intracellular trafficking

Budding yeast/MYO2 Budding yeast/MYO2 Budding yeast/MYO2 Budding yeast/MYO2 Budding yeast/MYO2 Fisson yeast/MYO4 (MYO52) Neurons Frog, fish, mouse/M5a DRG neuron growth cones

16 17 18 19 20 21 22 23 24 25

Particle transport

Drosophila embryo/95F Drosophila oocyte/95F Drosophila developing sperm/95F

26 27 28

C. elegans developing sperm/spe-15 Mouse hair cells, stereocilia/Myo6, sv Epithelial cells Polarized epithelial cells

29 30 31 32

Dictyostelium/myoi Sensory hair cells/Myo7a, shaker-1

33 34 35

M6

Transport and membrane organization (?)

Clathrin-mediated endocytosis M7

Cell adhesion, filopod extension Lateral links, stereocilia organization Melanosome transport

Mammal. cells/M5b, M5c

Retinal pigment epithelia (RPE)

1 The Myosin Superfamily: An Overview Table 1.1

(continued).

Myosin

Putative role

Relevant organism or cell type/gene

Reference(s)

M9

Signaling to actin cytoskeleton (?)

Rat, human (GAP domain)

36

M10

Transport (?), filopod extension

Mammalian cells

37

M14

Gliding motility

Toxoplasma/TgMyoA and Plasmodium

38

The numbers correspond to the following references: 1. Fukui et al., 1989, Wessels et al., 1991, 20. Beach et al., 2000, Yin et al., 2000; 1996, Yonemura and Pollard, 1992; 21 Motegi et al., 2001, Win et al., 2000; 2. Cheney and Mooseker, 1992, Coluccio, 22 Bridgman, 1999, Evans et al., 1998, 1997, Tyska and Mooseker, 2002; Prekeris and Terrian, 1997, Tabb et al., 3. Fath and Burgess, 1993, Raposo et al., 1998; 1999, Durrbach et al., 2000; 23 Koyama and Takeuchi, 1981, Provance 4. Novak et al., 1995, Geli and Riezman, et al., 1996, Rodionov et al., 1998, 1996, Goodson et al., 1996, Jung et al., Rogers and Gelfand, 1998, Wei et al., 1996; 1997; 5. Huber et al., 2000; 24 Wang et al., 1996; 6. Neuhaus and Soldati, 2000; 25 Lapierre et al., 2001, Rodriguez and 7. García et al., 1998, Hasson et al., 1997, Cheney, 2002; Holt et al., 2002, Steyger et al., 1998; 26 Mermall and Miller, 1995; 8. see Fujiwara and Pollard, 1976, Ma27 Bohrmann, 1997; buchi and Okuno, 1977 and refer28 Hicks et al., 1999; ences in Robinson and Spudich, 2000; 29. Kelleher et al., 2000; 9. Bezanilla and Pollard, 2000, Motegi 30 Avraham et al., 1995, Deol and Green, et al., 2000; 1966, Self et al., 1999; 10. Mizuno et al., 2002, Young et al., 1993, 31 Biemsderfer et al., 2002, Hasson and Uyeda and Titus, 1997; Mooseker, 1994, Heintzelman et al., 11. see references in Burridge and 1994, Self et al., 1999; Chrzanowska-Wodnicka, 1996; 32 Buss et al., 2001, Inoue et al., 2002b, 12. Anderson et al., 1996, Eddy et al., 2000 Morris et al., 2002; Post et al., 1995, Verkhovsky et al., 33 Titus, 1999; 1995, Wessels et al., 1988; 34 Gibson et al., 1995, Hasson et al., 13. Hicks and Williams, 1992, Porter et al., 1997, Küssel-Anderman et al., 2000 1992; Self et al., 1998; 14. Montell and Rubin, 1988, Porter and 35 El-Amraoui et al., 2002 Liu et l., Montell, 1993, Porter et al., 1995, 1998a; Stephenson et al., 1983; 36 Müller et al., 1997, Reinhard et al., 15. Battelle et al., 1998; 1995, Wirth et al., 1996; 16. Johnston et al., 1991, Schott et al., 37 Berg and Cheney, 2002; 1999; 38 Dobrowolski and Sibley, 1996, Heint17. Hill et al., 1996; zelman and Schwartzman, 1997, 18. Hoepfner et al., 2001, Rossanese et al., Herm-Götz et al., 2002 Hettmann 2001; et al., 2000. 19. Bobola et al., 1996, Jansen et al., 1996, Long et al., 1997, Takizawa et al., 1997;

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1.3 Diverse Functions for Myosins

the full spectrum of functions for this family of motors has yet to be appreciated. Thus far, myosins have been implicated in a variety of processes that include cell motility and adhesion, intracellular transport, maintenance of actin-rich structures, membrane trafficking and signal transduction (see Baker and Titus, 1998, Mermall et al., 1998). 1.3.1

Non-muscle Contractile Structures

M2 forms filaments not only in muscle but in non-muscle cells as well. The nonmuscle M2 filaments are dynamic structures they can be assembled and disassembled in particular intracellular locations as needed (Tan et al., 1992). The critical role for M2 in cell motility will be discussed below; it is also involved in forming actin-rich contractile structures during processes such as cytokinesis, morphogenesis and wound healing (Bement, 2002, Robinson and Spudich, 2000, Young et al., 1993). M2 is found interspersed among actin filaments that form contractile bundles between adjacent epithelial cells and in the cortical actin network that supports cell structure. At the cell periphery, a contracting acto-M2 network is thought to prevent pseudopod formation, assist in the retraction of the rear of the cell and provide resting cortical tension (e. g. Eddy et al., 2000, Pasternak et al., 1989, Wessels et al., 1988). It is also found in actin filament stress fiber bundles that are thought to play a role in anchoring a cell to the substrate (Burridge and Chrzanowska-Wodnicka, 1996). Early studies showed the presence of M2 in the contractile ring at the equator of dividing cells and that the injection of function-blocking antibodies resulted in an inhibition of cytokinesis in echinoderm embryos (Fujiwara and Pollard, 1976, Mabuchi and Okuno, 1977). Subsequent molecular genetic studies in the Dictyostelium and yeast provided direct proof of a role for non-muscle M2 in cytokinesis (Guertin et al., 2002, Robinson and Spudich, 2000). While budding yeast and Dictyostelium only have a single M2, the fission yeast Schizosaccharomyces pombe has two M2s (Myo2p and Myp2p) that seem to play non-overlapping roles in cytokinesis (Bezanilla and Pollard, 2000, Motegi et al., 2000). Interestingly, in the case of Dictyostelium, M2 is only essential for cytokinesis when cells are in suspension where mutant cells become large and multinucleate. The cells are able to undergo a M2 independent cytokinesis when plated onto a substrate. Thus, Dictyostelium has more than one distinct mode of cytokinesis. One mode is dependent on M2 (‘cytokinesis A’), while another is M2-independent and instead requires adherence to a substrate (‘cytokinesis B’; see Gerisch and Weber, 2000, Uyeda et al., 2000). Finally, non-muscle M2 is required for morphogenesis in both Dictyostelium and Drosophila melanogaster (Uyeda and Titus, 1997, Young et al., 1993). For example, mutations in Drosophila M2 result in embryos with defects in dorsal closure, head involution and axon patterning (Young et al., 1993). Further analysis of the mutant embryos suggests M2 is involved in the maintenance of cell shape and cell sheet movement. Embryos mutant for myosin phosphatase, a negative regulator of non-muscle M2, also exhibit dorsal closure defects (Mizuno et al., 2002).


1.3.2

Cell Motility and Adhesion

Myosins play a central role in cell motility. Immunofluorescence experiments have shown that M2 is concentrated in the lamellae, posterior and cortical region of polarized, locomoting cells (Anderson et al., 1996, Post et al., 1995, Verkhovsky et al., 1995). Conversely, M1s are located at the leading edges of lamellipodial projections of migrating Dictyostelium and Acanthamoeba (Fukui et al., 1989, Yonemura and Pollard, 1992) and in filopodia, lamellipodia, and growth cones of mammalian cells (Lewis and Bridgman, 1996, Ruppert et al., 1995 Wagner et al., 1992). These findings suggest that M1 is involved in forward extension of the cell and that M2 functions in retraction at the rear of the cell body. Dictyostelium cells mutant for either M1 or M2 have defects in locomotion (Uyeda and Titus, 1997), further supporting the essential roles for myosins in cell motility. Both M1 and M2 are required for dictating where a pseudopod is formed. M2 filaments at the cortex inhibit protrusion and the loss of this myosin results in the extension of pseudopodia around the periphery of the cell and a 50 % decrease in velocity (Wessels et al., 1988). Dictyostelium lacking the M1s myoA or myoB are not deficient in extending pseudopodia, rather they make excess pseudopodia that are not extended in the typically ordered manner (i. e. one at a time) (Wessels et al., 1991, 1996). Similarly, chromophore-assisted laser inactivation (CALI) of M1 in neuronal growth cones causes expansion of lamellipodia (Wang et al., 1996). These observations suggest that M1s have a general role in regulating pseudopod formation, an actin-dependent process. The amoeboid M1s have recently been shown to be linked to the Arp2/3 actin-polymerization machinery and may aid in focusing the site of its action to direct the site of the growing pseudopod (Evangelista et al., 2000, Jung et al., 2001, Lechler et al., 2000, Lee et al., 2000). Characterization of a Dictyostelium M7 null mutant revealed a role for this class of myosin in cell adhesion (Tuxworth et al., 2001). Mutant cells displayed reduced contact between the cell surface and substrate during migration. Loss of adhesion to surfaces results in a defect in phagocytosis (Titus, 1999, Tuxworth et al., 2001) due to reduced binding to particles. Preliminary analysis suggests that the loss of adhesion is not due to a loss of cell surface receptors. Since the early stages of phagocytosis and cell motility are similar, it is not completely surprising that M7 is important for both processes. It has been proposed that this myosin participates in adhesion receptor complex assembly and disassembly at the plasma membrane (Tuxworth et al., 2001). Apicomplexan parasites such as Toxoplasma gondii and Plasmodium falciparium move by an unusual means termed gliding motility (Sibley et al., 1998). These cells tightly anchor themselves to their host cell and then propel themselves into the cell by translocating the sites of attachment towards the rear of the cell. No change in cell shape is involved in their movement, in contrast to amoeboid cells. Treatment of cells with actin depolymerizing drugs inhibits gliding motility (Dobrowolski and Sibley, 1996). These parasites uniquely express M14, a class of

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myosin that has a motor domain, virtually no neck, and a short tail (Heintzelman and Schwartzman, 1997, Hettmann et al., 2000) and one of these, TgMyoA, is localized to the plasma membrane (Herm-Götz et al., 2002, Hettmann et al., 2000). Recent biochemical analysis of TgMyoA (Herm-Götz et al., 2002) reveals that it is associated with a novel, relatively large LC (31 kD versus 20 kD) that is calmodulinlike and binds to the C-terminal 53 amino acids of the TgMyoA HC. The biochemical properties of this unusual myosin are remarkably similar to those of M2. In spite of a low degree of sequence similarity to other myosins in the converter and LC binding domains, TgMyoA propels actin filaments at a rate of Z 5 mm s 1 and has a step size of 5 nm. These data suggest that this myosin is capable of generating the forces required for propelling gliding motility. The development of an inducible expression system for Toxoplasma (Meissner et al., 2001) should facilitate analysis of the role of this fascinating myosin in gliding motility. Myosins contribute to various aspects of cell migration directed protrusion of the lamellipodium, adhesion to the substrate and retraction of the rear of the cell. The importance of an actomyosin-based mechanism for the propulsion of a wide array of cells is evidenced by the finding that an unusual myosin, M14, powers a unique form of motility. Given the complex series of events that occur during migration, it is likely that other myosins might also be found to participate in cell migration as new members of the myosin superfamily are studied. 1.3.3

Organelle/Cellular Component Transport

The proper localization of intracellular components at the necessary time(s) is important for cell function. Actin- and microtubule-dependent motors each play essential roles in ensuring that cellular components (cargos) reach the right place at the right time. Current evidence suggests that microtubule motors serve to translocate organelles for long distances while the actin-based motors move things locally (Brown, 2000, Rogers and Gelfand, 2000). Myosins appear to transport cellular cargos that range from vesicles to mRNA, and the best-studied of the transport myosins is M5. Evidence for cargo transport by M5s comes from the study of yeast as well as pigment cells from mouse, fish and frog (Brown, 2000, Rogers and Gelfand, 2000). The yeast Saccharomyces cerevisiae has two M5 genes (MYO2 and MYO4). The product of the MYO2 gene (Myo2p) is essential for viability. Yeast with the temperature sensitive (ts) mutation myo2-66 (a point mutation in the N-terminal actin-binding domain) exhibit an accumulation of secretory vesicles and arrest in a large, unbudded state at the restrictive temperature (Johnston et al., 1991, Lillie and Brown, 1994). It has been shown that Myo2p binds vesicles via its tail region and transports them along actin filaments from the mother cell to the bud (Catlett and Weisman, 1998, Reck-Peterson et al., 1999, Schott et al., 1999). Genetic data suggest that the Myo2p tail also binds to a kinesin homolog (Smy1p) and a vesicle-associated Rab protein (Sec4p) (Beningo et al., 2000, Lillie and Brown, 1992, Pruyne et al., 1998, Schott et al., 1999 Walch-Solimena et al.,


1997). The association between Myo2p and Smy1p is supported by co-localization and yeast two-hybrid experiments (Beningo et al., 2000, Lillie and Brown, 1994). These data contribute to the growing body of evidence that actin filament and microtubule systems cooperate. An M5 from fission yeast, Myo4p (also referred to as Myo52), has also been implicated in organelle transport. Myo4p/Myo52p is localized to small punctae that accumulate at sites of polarized growth at the ends of the cells and is also found at the septum. These particles appear to move rapidly within the cell. Deletion of myo4/myo52 results in rounder cells that exhibited an accumulation of internal vesicles (Motegi et al., 2000, Win et al., 2000). In addition to its role in transporting materials to sites of cell growth and division, Myo4p/Myo52p has also been found to play a role in vacuole fusion in response to osmotic stress (Mulvihill et al., 2001). Further work has revealed a function for M5 in vacuole inheritance in budding yeast (Hill et al., 1996). The myo2-66 ts mutant was found to exhibit aberrant vacuole partitioning and inheritance by the daughter cell prior to division. A screen for additional myo2 alleles resulted in the identification of the myo2-2 mutant yeast which was also defective for vacuole inheritance but did not display the cell growth defects observed with myo2-66 (Catlett and Weisman, 1998). The mutation was found to reside in a region of the Myo2p tail specifically required for vacuole binding (Catlett et al., 2000). It appears that Myo2p is required for two separate processes in yeast: the transport of secretory vesicles important for polarized cell growth and the segregation of vacuoles (an event not essential for polarized cell growth). This result has exciting implications for the identification of multiple cargos, and thus multiple cargo binding receptors, for the same molecular motor protein. Like vacuoles, peroxisomes and Golgi elements are also segregated into the yeast bud during cell growth. Movement of these organelles from mother to daughter requires the actin cytoskeleton, and it was demonstrated that Myo2p provides the means for their transport. In cells containing the myo2-66 ts allele, peroxisome movement stops abruptly at the non-permissive temperature (Hoepfner et al., 2001) and the inheritance of late Golgi elements is inhibited (Rossanese et al., 2001). In addition to its role as a transport motor, yeast Myo2p has been shown to participate in mitotic spindle orientation (Beach et al., 2000, Yin et al., 2000). Proper spindle orientation is important for accurate chromosome segregation. The tail region of Myo2p binds Kar9p (a microtubule-associated protein) in yeast two-hybrid and co-immunoprecipitation experiments, and Myo2p is required for Kar9p transport into the bud. It appears that Myo2p and Kar9p both play essential roles in spindle orientation. Current models propose that Kar9p binds to both microtubules and Myo2p and that Myo2p transports both into the growing bud (Beach et al., 2000, Yin et al., 2000). This analysis provides another link between the actin filament and microtubule cytoskeleton systems. The other M5 in budding yeast, Myo4p, has been shown to participate in the asymmetric localization of the transcriptional repressor, Ash1p (Bobola et al., 1996, Jansen et al., 1996). Ash1p represses HO endonuclease, the protein that in-

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duces mating type switching in yeast (Sil and Herskowitz, 1996). Thus, localization of Ash1p to the budding daughter cells suppresses mating type switching (Bobola et al., 1996, Jansen et al., 1996). ASH1 mRNA is asymmetrically segregated to the bud tip of the growing daughter cell (Long et al., 1997, Takizawa et al., 1997), and its movement into the bud tip can be monitored by tagging it with green fluorescent protein (GFP) (Beach et al., 1999, Bertrand et al., 1998). Myo4p and two other She proteins (She2p and She3p) co-localize with ASH1 mRNA (Long et al., 1997), and all four components co-immunoprecipitate from cell extracts (Böhl et al., 2000, Long et al., 2000, Münchow et al., 1999, Takizawa and Vale, 2000). Myo4p coimmunoprecipitates with the She3p adapter in the absence of RNA, while She2p (a second adapter) recruits the Myo4p She3p complex to ASH1 mRNA (Böhl et al., 2000, Long et al., 2000, Münchow et al., 1999, Takizawa and Vale, 2000). The association between She3p and Myo4p appears to be mediated by their coiled-coil regions (Böhl et al., 2000). M5 may also have a vesicle transport role in neurons, where it was found to move extruded axon smooth endoplasmic reticulum (SER) and brain vesicles in vitro (Evans et al., 1998, Tabb et al., 1998). Antibodies against the M5 head or tail were able to inhibit this movement (Evans et al., 1998, Tabb et al., 1998) and immunogold electron microscopy revealed the co-localization of M5 and a kinesin on the SER vesicles (Tabb et al., 1998), consistent with a model of actin filament and microtubule system cooperation. M5 has also been found to be associated with organelles in live neurons (Bridgman, 1999). Analysis of neurons from dilute mice lacking M5a reveals that organelles still undergo bidirectional movement along microtubules. However, loss of this motor causes an accumulation of organelles in regions of the cell enriched for dynamic microtubule ends, suggesting that M5a is responsible for dispersal of these organelles to actin-rich areas of the cell (Bridgman, 1999, Wu et al., 1998). The role of M5 in pigment granule transport has been well characterized in frog (Xenopus laevis), fish and mouse melanophores cell types that have traditionally served as model systems to study intracellular transport mechanisms (Tuma and Gelfand, 1999, Wu and Hammer, 2000). Pigment granules in Xenopus melanophores undergo rapid and tightly regulated aggregation and dispersal, creating changes in color in response to external stimuli (Tuma and Gelfand, 1999). Transport of pigment granules seems to involve both the actin filament and microtubule cytoskeletal systems (Rodionov et al., 1998, Rogers and Gelfand, 1998). Purified granules from frog melanosomes contain bound M5, kinesin II, and dynein, and have been shown to move along actin filaments (as well as microtubules) in vitro (Rogers et al., 1997, 1999) (Fig. 1.3a). Recent in vivo analysis of melanosome movement suggests that the motor action of kinesin and dynein are in a ‘tug-ofwar’ with M5 and that M5 activity is downregulated during the aggregation phase (Gross et al., 2002). Melanosome transport to the cell periphery by M5 has also been studied in mouse melanocytes (Wu and Hammer, 2000). Mouse melanosomes containing pigment are transported from the center of the cell to the periphery, where they are taken up by keratinocytes. The pigment is eventually deposited into the hair


nucleus

melanosome

-

+

melanosome

cytoplasmic dynein kinesin II

+

M5a

+

plus-end of microtubules

-

minus-end of microtubules actin filaments

microtubulefree zone

microtubules

a)

a)

F-actin M5a melanophilin

Rab27a

b)

b)

melanosome

Figure 1.3. Models for melanosome transport by M5a. (a) Schematic of melanosome transport in a Xenopus melanophore, highlighting the involvement of both microtubule-based motors (cytoplasmic dynein and kinesin II), and the actin-based motor M5a. In the dual filament model of transport proposed by Langford (discussed in Tuma and Gelfand, 1999), melanosomes are transported over a long distance from the center of the melanosome to the cell periphery on microtubules by dynein and kinesis. Once they reach the periphery, they are

transferred to the actin-filament cytoskeleton where local transport occurs via M5a. (b) A more detailed, molecular view of melanosome transport by M5a is diagrammed. A protein complex composed of Rab27a and melanophilin was shown to act as a receptor for M5a, targeting the motor to the melanosome (Fukuda et al., 2002, Hume et al., 2002, Provance et al., 2002, Wu et al., 2002a). Rab27a first binds the melanosome then recruits melanophilin, which subsequently recruits M5a into the complex.

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shaft, causing changes in coat color. Loss of M5a (dilute) in the mouse results in a lightened or ‘diluted’ coat color because the pigment granules are not transported to the dendritic tips of the melanocytes in both cultured cells (Koyama and Takeuchi, 1981, Provance et al., 1996) and in situ (Wei et al., 1997). M5 was shown to localize to the melanosomes (Lambert et al., 1998, Nascimento et al., 1997, Wu et al., 1997b). Expression of the globular tail domain in cultured melanocytes disrupts the binding between endogenous M5 and melanosomes, causing a clumping of these organelles in the central region of the cell that leads to the dilute phenotype (Wu et al., 2002b). Mutations at two other murine loci (ashen and leaden) cause the same phenotype as dilute a lightened coat color due to pigment granule transport defects (Silvers, 1979). The ashen gene was found to encode the Rab27a GTPase, and leaden encodes a Rab binding protein named melanophilin (Matesic et al., 2001, Wilson et al., 2000). The molecular relationship between the gene products of these loci has only recently been elucidated. The receptor for M5a (dilute) has been identified as a complex of two proteins, Rab27a (ashen) and melanophilin (leaden, also referred to as Slac2-a) (Fukuda et al., 2002, Hume et al., 2001, 2002, Provance et al., 2002, Wu et al., 2001, 2002a, 2002b). Rab27a was shown to first bind the melanosome, then recruit melanophilin, which subsequently recruits M5a into the complex (Fig. 1.3b). The C-terminal region of melanophilin binds to the C-terminal domain of M5a (Nagashima et al., 2002, Wu et al., 2002a). Binding of melanophilin to Rab27a occurs in a GTP-dependent manner, providing a means of controlling M5a association to the melanosome (Kuroda et al., 2002, Wu et al., 2002b). It is of interest to note that M7a is associated with melanosomes in retinal pigment epithelium (RPE) and that these melanosomes are not correctly distributed in the shaker-1 M7a mutant (Liu et al., 1998a). This observation, coupled with the recent identification of a rabphilin (El-Amraoui et al., 2002) as an M7a tail binding protein, suggests that M7a may also have a role in melanosome transport in the RPE. M6 has also been implicated in organelle transport, consistent with it being a high-duty ratio motor (Mermall et al., 1994). The fly M6, designated 95F, is present on moving particles of unknown identity in the syncytial blastoderm. These particles appear to move on linear tracks. An inhibitory polyclonal antibody against 95F was able to block the linear (non-random diffusion) motion of the particles. The effect was similar to what was observed following treatment of the embryos with either actin-depolymerizing agent cytochalasin D or ATP inhibitor dinitrophenol (DNP), suggesting that 95F M6 powers the movement of these particles. These particles appear to be associated with the invaginating plasma membrane and may be required for supplying membrane or other material to the growing invagination during the rapid cycles of cell division that occur in the early fly embryo (Mermall and Miller, 1995). Fluorescence time-lapse microscopy experiments with anti-95F antibodies revealed a role for fly M6 in the transport of cytoplasmic particles from the nurse cell to the oocyte (Bohrmann, 1997). Microinjection of mitochondria-specific dyes into the developing oocyte suggested that some of these particles were mitochondria. Partial loss of function mutations in the 95F gene revealed a role for M6


in sperm development (Hicks et al., 1999). The jaguar mutation disrupts the first exon of the 95F myosin, causing male sterility due to a defect in sperm individualization (the process of enclosing each spermatid with its own membrane from what was once a syncytium). Consistent with this phenotype, 95F is localized to the leading edge of the individualization complex (IC), an actin-rich structure that surrounds the nuclei as the process of individualization proceeds (Hicks et al., 1999). This region is enriched in vesicles and it is possible that 95F could participate directly in the transport of membranes to the IC. Alternatively, it may contribute to the organization or stabilization of a membrane-rich leading edge of the IC that is required for efficient delivery of plasma membrane to the sperm undergoing individualization. Interestingly, M6 is also critical for sperm development in C. elegans (Kelleher et al., 2000). Animals with a null mutation in the spe-15 gene are sterile and exhibit defects in the segregation of various cellular components during spermatogenesis. M6 is thus essential for transport and/or membrane reorganization during spermatogenesis in both flies and worms, and might play a conserved role despite the significant differences in sperm development between these two organisms. Myosins have also been implicated in nuclear movements. The unique myosin from the ciliated protozoan Tetrahymena thermophila, MYO1, has been found to be required for macronucleus elongation, a process essential for correct segregation of genetic material to daughter cells (Williams et al., 2000). This result raises the intriguing possibility that myosins play a role in the transport or motility of nuclei in some contexts. 1.3.4

Maintenance of Actin-rich Extensions

M1, M5, M6, M7, M10 and M15 are found in actin-rich cell extensions such as microvilli, filopodia and stereocilia. Their exact roles in the generation or maintenance of these structures remain unclear at this time. Several myosins have been proposed to provide materials, force and tension required to maintain these structures. One of the best characterized M1s is the brush border subtype MYO1A (better known as BBM1) that forms links between actin bundles in the microvillar core and overlying plasma membrane in brush-border intestinal microvilli (Coluccio, 1997, Mooseker and Cheney, 1995). It has been hypothesized to provide structural support to the microvilli, but recent analysis of GFP-tagged MYO1A in an epithelial cell line suggests that it is rapidly exchanged with a cytosolic pool (Tyska and Mooseker, 2002), raising questions about how this myosin might play a structural role. It has also been localized to Golgi-derived vesicles, suggesting that its role is largely to deliver membrane to the growing microvillus (Fath and Burgess, 1993) and that it might also contribute to the dynamic growth or changes in flexibility of this structure (Tyska and Mooseker, 2002). The Drosophila M3, ninaC (neither inactivation nor after potential C), was identified by the characterization of a vision mutant defective in photoreception. NINAC is expressed specifically in fly photoreceptor cells as two alternatively

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spliced proteins (Montell and Rubin, 1988). One form localizes to the microvillar extensions of photoreceptors (rhabdomeres), while the other form is found in cell bodies (Hicks and Williams, 1992, Porter et al., 1992). Cells null for NINAC undergo retinal degeneration (Porter et al., 1992). M3 was also recently identified in humans (Dosé and Burnside, 2000), but whether or not it functions in human vision is not yet apparent. M5 is found at the tips of microvilli and is also present in filopodia (Evans et al., 1997, Heintzelman et al., 1994). CALI of M5 in growth cones of dorsal root ganglion (DRG) neurons results in rapid filopod retraction (Wang et al., 1996), suggesting a role in delivery of materials required for filopod growth or for growth cone function. Fractionation studies and in vitro motility assays are consistent with this model of M5a action (Evans et al., 1998, Prekeris and Terrian, 1997). However, analysis of cultured dilute neurons did not reveal any defects in growth cone dynamics (Evans et al., 1997). The discrepancy between the CALI study and the analysis of cultured dilute neurons remains to be resolved, but it is possible that another M5 can compensate partially for the loss of M5a in the mutant mouse neurons. Dictyostelium M7 and mammalian M10 are both localized to the tips of filopodia (Berg et al., 2000, Tuxworth et al., 2001). Deletion of the Dictyostelium M7 results in a loss of filopodia extension (Tuxworth et al., 2001). It is not yet known if this is due to the loss of delivery of materials necessary for making filopodia (as appears to be the case for M5), or if this motor is required for stabilizing this type of actin-rich extension. Overexpression of M10 in COS-7 cells, in contrast, results in the production of longer filopodia in increased numbers (Berg and Cheney, 2002). The association of M10 with particles that move up and down along the filopodial shaft suggests that this motor might be required for the targeted delivery of materials to the growing filopod. The inner ear sensory hair cells in mice have actin-rich extensions (modified microvilli) called stereocilia at their apical surface (Fig. 1.4). These structures are organized into units or bundles that move together when stimulated. The bundles are deflected in response to sound vibrations and gravity, resulting in the opening of ion channels and eventually transduction of electrical signals to the brain (Holt and Corey, 2000). The Snell’s waltzer (sv) mouse mutant carries an intragenic deletion in the gene encoding M6 (Myo6) (Avraham et al., 1995). These animals are deaf, and also display hyperactivity and circling behavior (Deol and Green, 1966). M6 is localized to the base of the stereocilia and is enriched in the cuticular plate (a region of densely-packed actin filaments that the stereocilia are embedded into), as well as being diffusely distributed throughout the cytosol (Avraham et al., 1995, Hasson et al., 1997). Additionally, it is also found in the pericuticular necklace (a region at the periphery of the apical surface of the cell that is rich in vesicles) and is associated with punctae throughout the cytoplasm of the hair cells (Hasson et al., 1997). Molecular analysis of the inner ear hair cells in sv mice supports a role for M6 in maintaining stereocilia structure. Stereocilia bundles appear normal at birth, but by 3 days after birth the bundles become fused together and disorganized (Self


apical tip F-actin M1 = M1b = M6 = M7a

M15 stereocilia

(?)

= M15 = gated ion channel = tip link = cadherin

M7

= catenin = motor complex = vezatin

M6

M6

cuticular plate

perinuclear necklace (vesicle-rich)

F-actin

Figure 1.4. Schematic of myosins in the hair cell of the ear. Several myosins play critical roles in the mammalian ear mutations in M2, M3, M6, M7a and M15 have all been linked to human deafness. The apical tip of the hair cell features actin-rich extensions called stereocilia. M1 localizes near the tip of the stereocilia in close proximity to tip links, which are likely attached to gated ion channels that open upon deflection of the hair cell bundle following stimulation (Gillespie and Walker, 2001). M6 localizes to the base of the stereocilia, and is also enriched in the cuticular plate and perinuclear necklace (Avraham et al., 1995, Hasson et al., 1997). It is believed to provide the force

necessary to anchor the stereociliar membrane at the base of each extension. M7a localizes along the length of each stereocilium and is concentrated at site of lateral links, which are thought to be important for maintaining bundle integrity (Hasson et al., 1997). M7a may anchor cadherins at lateral link sites via an association with vezatin (Küssel-Anderman et al., 2000). Finally, M15 also localizes to stereocilia and is concentrated in the cuticular plate (Liang et al., 1999). Its role in the hair cell has not yet been elucidated. Note that the localization of neither M2 nor M3 in the sensory hair cells of the ear is as yet known.

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et al., 1999). One model for this fusion is that M6 provides the force required to anchor the membrane at the base of each individual stereocilium. The organization of actin filaments in the stereocilia and the minus-end directionality of M6 are such that this motor may help pull down the membrane around each stereocilium. In the absence of M6, discrete stereocilia form, but these structures are not preserved as the mice age resulting in stereocilia fusion. It should be noted that although M6 is expressed ubiquitously in mammals (Avraham et al., 1995, Hasson and Mooseker, 1994), the sv mice have no gross phenotypes other than deafness (Avraham et al., 1995, Deol and Green, 1966). However, their intestinal microvilli are shorter than those found in matched controls (Self et al., 1999), consistent with M6 playing an important role in the maintenance of these specialized actin-filled structures. Despite the aberrant microvillar length, no gross defects in digestion have been reported. It may be necessary to subject the mice to some type of stress in order to uncover the consequences of shorter microvilli. The hypothesized role for M6 in membrane reorganization during stereocilia bundle formation/maintenance, along with membrane reorganization during fly embryogenesis and sperm development, follows a recurring theme for M6 involvement in membrane remodeling and maintenance. As observed in stereocilia, M6 localizes at the base of microvilli (another type of actin-rich structure) in brush border epithelial cells (Biemsderfer et al., 2002, Hasson and Mooseker, 1994, Heintzelman et al., 1994) where it may provide force for anchoring the membrane around each microvillus. A role for M6 in endocytosis could also relate to various biological processes involving membrane restructuring. As a high-duty ratio motor (De La Cruz et al., 2001), M6 is a motor ideally adapted for maintaining membrane tension and providing force. M7a may also be important for aspects of stereocilia integrity. As with the Snell’s waltzer mouse, mice mutant for M7a (shaker-1) are deaf and display balance dysfunction (Gibson et al., 1995). Their hair cells extend stereocilia, but these are splayed apart (Self et al., 1998). Immunofluoresence studies have shown that M7a protein localizes along the length of each stereocilium and is concentrated at sites of lateral links that are believed to be important for maintaining the integrity of the stereocilia bundles (Hasson et al., 1995). Perhaps M7a is involved in organizing the stereocilia into bundles to provide rigidity during bundle deflections. M7a may be required to anchor cadherins at the lateral link sites via an association with vezatin, a ubiquitous, novel transmembrane protein (Küssel-Anderman et al., 2000). 1.3.5

Membrane Trafficking

The M1s have been suggested to play a role in vesicle transport and endocytosis based on localization studies. As mentioned above, MYO1A in intestinal epithelial cells is associated with Golgi-derived vesicles that potentially provide membrane to the base of microvilli (Fath and Burgess, 1993). MYO1A is localized to both endo-


somes and lysosomes and expression of a truncated myosin dominantly inhibits the delivery of endosomal contents to lysosomes (Cordonnier et al., 2001, Raposo et al., 1999). It has also been implicated in the trafficking of basolateral vesicles to the apical plasma membrane in polarized epithelial cells (Durrbach et al., 2000). The structurally similar MYO1D has been shown to play a role in trafficking of recycling endosomes (Huber et al., 2000). Interestingly, an amoeboid M1 from Dictyostelium (myoB) has also been implicated in recycling from early endosomes, suggesting that there is some basic conservation of M1 function (Neuhaus and Soldati, 2000). Kinetic analysis of MYO1A indicates that this mammalian M1 is a shortduty motor (Jontes et al., 1997); therefore, if it acts as an organelle motor, it should be present in a complex of motors. Such a complex has yet to be identified, and indeed a bona fide MYO1A binding protein remains to be found. The different trafficking roles that various M1 isoforms play suggest that there are specific targeting molecules for each one, but these have not yet been identified. Characterization of such proteins should provide interesting insights into how the targeting of individual myosins may contribute to exquisite specification of their function. Two M5 isoforms, M5b and M5c, have also been implicated in intracellular trafficking. M5b co-localizes with transferrin and Rab11a to intracellular perinuclear vesicles (Lapierre et al., 2001). Expression of the M5b tail alone disturbs the recycling of transferrin to the plasma membrane in both polarized and non-polarized cells. Two-hybrid analysis revealed that M5b interacts with Rab11a, but attempts to verify this interaction in vivo have not yet been successful (Lapierre et al., 2001). Similarly, M5c co-localizes with Rab8 on intracellular vesicles and expression of the tail domain alone perturbs transferrin trafficking (Rodriguez and Cheney, 2002). The available data suggest that M5c is associated with a compartment distinct from that of M5b, another indication that even within a class of myosins there can be precise functional specificity of individual isoforms. M6 has been suggested to participate in clathrin-mediated endocytosis in cultured polarized epithelial cells (Buss et al., 2001). Endocytosis is important for nutrient uptake in the cell, along with immune system defense against microorganisms and cell-surface receptor recycling. The actin cytoskeleton plays a role in endocytosis, perhaps by providing a structural framework for trafficking and/or supplying the track for myosin motors transporting vesicles. M6 tagged with GFP was shown to co-localize with clathrin-coated vesicles via its C-terminal tail in polarized Caco-2 cells (Buss et al., 2001). It was found in a protein complex with adaptor protein-2 (AP-2) by pull-down assays and co-immunoprecipitation. The AP-2 protein is a component of clathrin-coated vesicles associated with the plasma membrane. Overexpression of the M6 tail domain reduced the uptake of transferrin in transfected cells by over 50 % (Buss et al., 2001). In contrast, M6 is found in association with the Golgi complex in non-polarized cells as well as in the actin-rich ruffles of growth factor-stimulated cells (Buss et al., 1998). The difference in observed localization when compared to polarized cells has been attributed to the lack of an epithelial cell-specific insert in the globular tail region. These findings reiterate how specific residues in the myosin tail play a significant role in specifying subcellular localization and function.

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M6 has also been shown by two-hybrid screens and co-immunoprecipitation to associate with Disabled-2 (Dab2), a protein involved in endocytosis and signal transduction (Inoue et al., 2002b, Morris et al., 2002). The central region of Dab2 is important for AP-2 binding, while the C-terminal tail mediates binding to M6 (Morris et al., 2002). Dab2 transiently co-localizes with members of the low-density lipoprotein receptor (LDLR) family in clathrin-coated pits and is thought to regulate receptor protein trafficking (Morris and Cooper, 2001). It has been proposed that the M6 Dab2 interaction provides a link between the actin cytoskeleton and receptors undergoing endocytosis (Morris et al., 2002). It remains to be determined if M6 acts as a motor for endocytic vesicles or if it participates in the organization of an actin-based sub-membranous domain where initial sorting of endosomal contents occurs. 1.3.6

Signal Transduction

The role of myosins in signaling was first eludidated in the fly eye. Illumination of the fly eye with pulses of light results in a distinctive negative potential followed by a rapid return to baseline each time the eye is stimulated. The ninaC (M3) mutant has a larger negative potential upon stimulation followed by a slower return to baseline after cessation of the stimulus (Montell and Rubin, 1988, Stephenson et al., 1983). The N-terminal kinase activity, as well as calmodulin binding to sites in the C-terminal region, are required for normal phototransduction (Porter and Montell, 1993, Porter et al., 1995). The role of NINAC in normal phototransduction is separate from its role in maintaining the overall integrity of the rhabdomere microvilli (Porter et al., 1992). M3 is also found in Limulus polyphemus (horseshoe crab) eyes where it was identified as a phosphoprotein that undergoes clock-regulated phosphorylation (i. e. horseshoe crabs have circadian neural input that changes phototransduction in the eye, among other things) possibly by a cAMP-dependent protein kinase (Battelle et al., 1998). Interestingly, Limulus NINAC has also been shown to be a phosphoprotein that can be phosphorylated in vitro by protein kinase C (PKC) (Battelle et al., 1998). Mutation of PKC sites in the tail region results in defects in deactivation after the light stimulus has been shut off. Recent evidence suggests that NINAC is a component of the ‘signalplex’, a cluster of molecules required for phototransduction and that it binds to INAD (a PDZ protein) through its C-terminus (Wes et al., 1999). NINAC does not require INAD binding for localization to the rhabdomeres, but their interaction is essential for termination of the electrophysiological response to light (Wes et al., 1999). MYO1C (also known as myr 2, myosin 1b) plays an unusual role in signal transduction. This M1 localizes near the tips of the sensory hair cell stereocilia where a fine fiber known as the tip link extends from the side of one stereocilium to the top of the adjacent, shorter stereocilium (García et al., 1998, Hasson et al., 1997, Steyger et al., 1998; Fig. 1.4). Elegant biophysical studies have shown that the tip link is likely attached to a calcium channel that opens upon deflection of the hair bundle


following an auditory stimulation (Gillespie and Walker, 2001). The cell becomes depolarized, but if the stimulus persists adaptation occurs. Adaptation has been proposed to occur when a myosin molecule climbs up the actin filaments in the core of the stereocilium, acting to reset the tension on the tip link and close the channel. An exciting new chemical genetics approach has been used to test this model of MYO1C function (Holt et al., 2002). Introduction of a single mutation in the ATP binding pocket of MYO1C (Y61G) allows the mutant molecule to bind a modified ADP (NMB-ADP) that locks it into the rigor state. Wild-type MYO1C does not bind NMB-ADP and is not inhibited by it, whereas the Y61G-MYO1C functions normally in the presence of ATP but not with NMB-ADP (Gillespie et al., 1999). Application of NMB-ADP to mouse utricular hair cells expressing small amounts of Y61G-MYO1C has a dramatic effect it abolishes adaptation (Holt et al., 2002). These data provide strong evidence in support of MYO1C being the adaptation motor and suggest a novel role for myosins in the ion channel gating. Human and rat M9s are predicted to contain a GAP domain in their tail regions with approximately 30 % sequence identity to GAP proteins of the Rho small Gprotein subfamily (Reinhard et al., 1995, Wirth et al., 1996). Rho belongs to a family of signaling proteins in the Ras superfamily of monomeric GTPases that act as molecular ‘switches’ whose activity depends on their nucleotide conformation state (GTP-bound versus GDP-bound). Activation of Rho leads to the formation of actin filament bundles and focal adhesions, among other processes (Ridley, 2001). GAP proteins negatively regulate these processes by stimulating GTP hydrolysis of Rho, converting it from an active state (GTP-bound) to an inactive (GDPbound) state. M9s have been shown to stimulate the GTPase activity of Rho A, B and C (Chieregatti et al., 1998, Post et al., 1998, Reinhard et al., 1995), and as predicted the overexpression of M9 in cultured mammalian cells leads to a loss of stress fibers and focal adhesion contacts (Müller et al., 1997). The exact role for M9 in signal transduction is not known, but it is interesting to note that this motor has been shown to be processive and to move toward the minus-end of actin filaments (Inoue et al., 2002a). An unusual myosin from Dictyostelium may also play a role in intracellular signaling. MyoM (to date an undesignated myosin) has a functional GEF domain at its extreme C-terminus (Geissler et al., 2000, Oishi et al., 2000). While deletion of this myosin does not result in any overt phenotype, expression of the tail region alone resulted in a decrease in growth rates and a hypersensitivity to osmotic stress (Geissler et al., 2000, Oishi et al., 2000). Cells exposed to water extended broad, actin-filled protrusions with the myoM tail at their tip, suggesting that the GEF domain is capable of signaling to the actin cytoskeleton under conditions of stress (Geissler et al., 2000). It remains to be determined whether this phenotype reflects the true function of myoM, but the fact that the behavior of the cells is only affected under specific conditions suggests that the observed phenotype is not due to non-specific misregulation of GEF activity.

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1.4 Myosins in Disease

1.4

Myosins in Disease

The critical roles for unconventional myosins are highlighted by their connection to a variety of human disorders, primarily those related to sensory dysfunctions. A common theme of structural roles in the inner ear and retina emerge for three classes of myosins. Mutations in genes encoding M3, M6, M7 and M15 have been implicated in human hearing disorders and deafness in mice. Myosin functions in hearing and other cellular processes related to disease phenotypes are just beginning to be uncovered. Perhaps the best-known relationship between myosin mutation and disease is in familial hypertropic cardiomyopathy (FHC; see the Chapter 19 by Konhilas and Leinwand), an autosomal dominant condition characterized by shortness of breath, angina, and heart palpitations that can lead to heart failure, stroke and sudden death (Seidman and Seidman, 2001). FHC can be caused by mutations in b cardiac myosin HC. Myosin LC mutations have also been associated with human heart myopathies similar to FHC (Poetter et al., 1996). 1.4.1

Griscelli Syndrome

Griscelli syndrome is characterized by severe primary immunodeficiency along with pigment dilution of hair, eyebrows and eyelashes (Griscelli et al., 1978, Pastural et al., 1997). A role for M5 in this disorder was hypothesized to be due to the phenotypic similarities between Griscelli syndrome patients and the dilute mouse mutant (described above; also see Westbroek et al., 2001). For example, pigment accumulation has been observed in the center of Griscelli melanocytes (similar to the accumulation seen in dilute melanocytes; see Griscelli et al., 1978, Koyama and Takeuchi, 1981, Provance et al., 1996). M5a was subsequently shown to co-localize with melanosomes in cultured human melanocytes (Lambert et al., 1998). Genetic evidence supports a role for M5 in this disorder. The MYO5A gene maps to the disease locus 15q21 and at least two M5 mutations have been identified in Griscelli patients (type 1 Griscelli syndrome; Pastural et al., 1997, 2000). Some Griscelli patients were reported to have immunodeficiency disorders (Griscelli et al., 1978, Klein et al., 1994), but this phenotype was inconsistent with the mouse dilute phenotype. However, the RAB27A gene is quite close to MYO5A they are 1.6 cM apart and RAB27A has also been linked with this syndrome (type 2 Griscelli syndrome; Ménasché et al., 2000). Rab27a is localized with M5s on melanosomes (Hume et al., 2001, Wu et al., 2002a) and these two proteins co-immunoprecipitate from melanocyte extracts (Hume et al., 2001). Rab27a plays a role in granule exocytosis while M5a does not (Haddad et al., 2001). Thus, MYO5A mutations are associated with the pigmentation and neurological aspects of Griscelli disease, while RAB27A mutations are associated with the immunological disease form of Griscelli (Ménasché et al., 2000).


Mutations in the non-muscle M2 encoded by MYH9 have recently been implicated in three giant-platelet disorders May Hegglin anomaly (MHA), Sebastian syndrome (SBS) and Fechter syndrome (Consortium, 2000, Kelley et al., 2000). All three disorders are characterized by thrombocytopenia, large platelets and leukocyte inclusions (Consortium, 2000, Kelley et al., 2000). Based on similarities between the three disorders and mapping experiments, it was proposed that MHA, SBS and FTNS are allelic. One unique feature of FTNS is sensorineural deafness, consistent with a connection between Fechter syndrome and DFNA17 deafness disorder. 1.4.2

Roles for Myosins in Hearing

Approximately one in 1000 people experience severe or profound deafness at birth or during early childhood, and another one in 1000 children become deaf before they reach adulthood (Petit et al., 2001). Non-syndromic deafness patients are afflicted only with hearing loss, while syndromic cases exhibit hearing loss in combination with other defects. There are many genes involved in deafness disorders, including several encoding unconventional myosins (see Fig. 1.4; see also Chapter 21 by Avraham). M3, M6, M7 and M15 have fundamentally important functions in the auditory system as mutant alleles at these loci have been shown to co-segregate with human deafness. M7 plays a role in hearing and balance in humans, mice and zebrafish (Ernest et al., 2000, Gibson et al., 1995, Weil et al., 1995). M7a (MYO7A) has a highly restricted tissue distribution in mammals it is found almost exclusively in cells that have specialized actin-based structures such as the hair cells of the ear, retina, testis, lung and kidney (Hasson et al., 1995). Mutations in MYO7A result in two forms of human non-syndromic deafness, mutant loci DFNB2 (autosomal recessive; Weil et al., 1997) and DFNA11 (autosomal dominant; Liu et al., 1997) and a syndromic form of deafness, Usher’s syndrome type IB (Weil et al., 1995). Usher’s syndrome type IB is characterized by a gradual degeneration of the rods and cones in the retina, causing loss of vision in addition to hearing loss (see Eudy and Sumegi (1999) and Keats and Corey, (1999) for reviews). As mentioned above, the shaker-1 M7a mutant mice are deaf and display balance dysfunction (Gibson et al., 1995). Mutations in the Danio rerio (zebrafish) M7a gene, mariner, were found to exhibit circling behavior along with hair cell defects similar to those observed in shaker-1 mice (Ernest et al., 2000). Hair cells did extend stereocilia but these are splayed apart, suggesting that the links between adjacent stereocilia have been lost (Ernest et al., 2000). These data demonstrate the remarkable functional conservation of M7a through evolution. A recent electrophysiological study has uncovered a role for M7a in gating of transducer channels that cannot be attributed to the disorganization of the hair bundles (Kros et al., 2002), suggesting that this motor may play more than one role in stereocilia function. While analysis of the shaker-1 mutant mouse has provided insight into the role of M7a in hearing (Gibson et al., 1995, Kros et al., 2002, Self et al., 1998), it has not provided a clear explanation for the late onset retinitis pigmentosa exhibited by

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1.4 Myosins in Disease

Usher’s IB patients. A careful analysis of shaker-1 photoreceptors has revealed that opsin transport is decreased (Liu et al., 1999) and melanosome distribution is aberrant (Liu et al., 1998a). It has been suggested that the relatively short life-span of the mouse is not long enough for expression of the vision defect to become apparent (Liu et al., 1999). Mutations have been documented all along the length of the M7a HC in humans, as well as mice and zebrafish. In addition to confirming the importance of the motor domain, the identification of mutations in both the C-terminal MyTH4 and FERM domains indicate these domains are essential for function (Ernest et al., 2000, Liu et al., 1998b). Mutations in M15 also result in deafness both in humans and mice. MYO15 is mutant in human DFNB3, a non-syndromic, congenital deafness disorder (Wang et al., 1998). As is the case for M7a, mouse M15 has a restricted tissue distribution. In addition to the cochlea, it is expressed in the pituitary and other neuroendocrine tissue (Liang et al., 1999, Lloyd et al., 2001). The M15 homolog in mouse is also associated with a hearing loss phenotype; the shaker-2 mouse was found to have stereocilia that are approximately one-tenth the normal length (Probst et al., 1998). M15 is localized to the stereocilia of cochlear hair cells and is concentrated in the cuticular plate (Liang et al., 1999). Consistent with its localization, an abnormal organization of actin is also observed in the cochlear hair cells of shaker-2 mice, suggesting that this myosin plays a role in actin organization. M6 was first associated with hearing loss when a null mutation in murine Myo6 was discovered to cause the Snell’s waltzer phenotype (Avraham et al., 1995), as discussed above (Section 1.3.4). A missense mutation in the motor domain of M6a results in an autosomal dominant non-syndromic form of deafness, DFNA22, indicating that this myosin has a conserved role in hearing (Melchionda et al., 2001). The effect of this single mutation on the motor properties of M6a is not yet known. Less is known regarding the role of other myosins involved in hearing, as mouse models for several human deafness mutations are not yet available for study. The role of M3 in fly vision has already been discussed, but M3 also plays an important sensory role in humans. Mutation of M3a causes progressive hearing loss DFNB30 (Walsh et al., 2002). MYO3A is expressed in the sensory cells of the mouse ear, but more work remains to be done to determine how this motor protein contributes to the maintenance of hearing. Given the conservation of M3 from flies to Limulus to humans, it is tempting to speculate that it plays a role in regulating some aspect of signaling in hair cells. A mutation in MYH9, a non-muscle M2, has been reported to be responsible for an autosomal dominant form of deafness, DFNA17 (Lalwani et al., 2000). This missense mutation results in a single amino acid change (R705H) in a residue located in the critical SH1 helix, which is known to influence the ATPase activity of myosin. The underlying cause of deafness and the localization of this myosin to sensory cells are not currently known, but it will undoubtedly be of interest to determine how a myosin that plays a role in generating contractile forces contributes to hair cell function.


1.5

New Myosins and Myosin Functions on the Horizon

A considerable number of myosins have been identified but their functions have not yet been elucidated. The presence of interesting domains in their N- or C-termini and divergence in their motor domains suggests that they have the potential to play unique and interesting roles. As work in the field progresses, these roles will undoubtedly be uncovered. A comprehensive analysis of genes predicted by the human and fly genome projects (Berg et al., 2001) has revealed the existence of a novel human myosin predicted from contig AC023133. This myosin appears to have a unique N-terminal extension and a short tail region. Two novel myosins were identified in the fly genome, one designated 29CD that has an N-terminal extension and a short, proline-rich tail and another designated 95E that has a large insert in the motor domain loop 1 and a tail region comprised largely of basic residues. The diverse roles played by the cytoskeleton in fly development is likely to catalyze interest in studying the in vivo function of these novel myosins through a search for null mutants or use of double-stranded RNA interference (dsRNAi) to knock out their mRNAs. A novel human myosin, M18 or PDZ-myosin, was identified in a differential screen for genes expressed highly in bone marrow stromal cells that support hematopoietic cells (Furusawa et al., 2000). The tail region of this myosin is comprised largely of predicted coiled-coil sequences (suggesting that it is a dimer) and there is a PDZ domain at the N-terminus. The tissue distribution of this myosin is not yet known. It is localized to cytoskeletal elements in stromal cell lines as well as NIH3T3 cells, where it appears to be associated with both actin and microtubules as well as being distributed throughout the cell. The novel M16, also referred to as myr8, was identified in a study of neuronal myosins in the rat (Patel et al., 2001). Two forms of this myosin were identified. M16a has a shorter tail and is expressed predominantly in the nervous system. The longer M16b appears to be broadly expressed. Analysis of the tail sequence reveals that it is unique and does not contain any regions of predicted coiledcoil, suggesting M16b is a monomer. The N-terminus contains eight ankryin repeats that bind to protein phosphatases. Immunoprecipitation experiments revealed that M16b indeed associates with protein phosphatase 1 catalytic subunits 1a and 1g. This myosin has a punctate distribution in both neurons and astrocytes. M4, M12 and a novel unconventional myosin from scallop mantle (tissue comprised of both muscle and non-muscle cells) named ScunM have been identified either at the protein or gene level but no information regarding their tissue distribution (in the case of M12 and ScunM) or subcellular localization is available (Baker and Titus, 1998, Hasegawa and Araki, 2002, Horowitz and Hammer, 1990). These myosins appear to be unique to the organisms in which they were originally identified. The M4 gene was identified in Acanthamoeba and sequence analysis revealed the presence of a C-terminal SH3 domain as well as a single MyTH4 domain (Horowitz and Hammer, 1990). It was shown to co-precipitate

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1.6 Conclusions

with actin under rigor conditions and to be released from actin by ATP, indicating that it has the biochemical properties expected of a myosin. M12 was identified at the gene level through a comprehensive search of the C. elegans genome for myosin genes and is named HUM-4 (heavy chain of unconventional myosin) (Baker and Titus, 1997). HUM-4 has a 150-amino acid extension on the N-terminus. In addition, the P-loop and actin-binding sequences in the motor domain are more divergent than those found in other myosins. There are two MyTH4 domains in the tail but no clear FERM domains have been identified. No known mutations map near the hum-4 gene and efforts to inactivate gene function by dsRNAi have not resulted in any phenotype (J. P. Baker and M. A. Titus, unpublished data; WormBase www.wormbase.org). ScunM was identified in a search for unconventional myosin genes in scallop mantle tissue (Hasegawa and Araki, 2002). It is one of the smallest myosins, with the HC having a predicted molecular weight of 90 kD. Similar to the M14s, ScunM has a short tail consisting of only 46 amino acids and it lacks any LC binding domains. Phylogenetic analysis, however, reveals that this myosin is the founding member of a novel class and not an M14.

1.6

Conclusions

It is likely that efforts to sequence the genomes of various organisms will result in the discovery of additional novel myosins. The ability to express myosins in baculovirus and inactivate gene function by dsRNAi in organisms and cultured cells should permit rapid characterization of their biochemical properties and in vivo functions. In addition to identifying myosins based on conserved motor sequences, it is possible that large-scale efforts directed at crystallizing proteins will identify proteins that do not share the conserved myosin primary structure, but whose structure is highly similar to that of the myosin motor domain. Thus, a convergence of proteomic and genomic approaches is likely to expand our appreciation of the diversity of the myosin superfamily. A number of lessons can be learned from examining the myosin superfamily. The most striking one is the diversity of the family both in terms of the variety of different classes and the number of different isoforms within a class. This diversity is reflected in the wide range of functions observed for myosins of even a single class. There are classes of myosins that appear to be specific for plants and parasites, while only three myosin classes are conserved from fungi to humans. As evolution proceeded, it appears that higher eukaryotes acquired a more sophisticated complement of myosins. Some of which, such as M7, were preserved and others, such as M4 and M12, appear to have been lost. The importance of myosins is highlighted by their essential roles in fungi and their association with a number of human diseases, ranging from cardiomyopathies to deafness. An unexpected surprise that emerged from studies of myosins in mammals is the key role that they play in hearing, perhaps reflecting their ability to participate in an array of cellular functions.


Future studies of this large motor family are likely to reveal additional roles and operating principles as well as, undoubtedly, exotic new family members.

Acknowledgements

The authors would like to thank Shawn Galdeen for helpful comments on the manuscript and Dr. Richard Cheney for providing Figure 1.2. M. C. K. is supported by an NIH postdoctoral fellowship (NRSA) and work in the Titus laboratory is supported by the National Institutes of Health. M. A. T. is an Established Investigator of the American Heart Association. References Anderson, K. I., Y. L. Wang, and J. V. Small. 1996. Coordination of protrusion and translocation of the keratocyte involves rolling of the cell body. J. Cell Biol. 134: 1209 1218. Avraham, K. B., T. Hasson, K. P. Steel, D. M. Kingsley, L. B. Russell, M. S. Mooseker, N .G. Copeland, and N. A. Jenkins. 1995. The mouse Snell’s waltzer deafness gene encodes an unconventional myosin required for structural integrity of inner ear hair cells. Nature Genet. 11: 369 375. Bähler, M. 2000. Are class III and IX myosins motorized signalling molecules? Biochem. Biophys. Acta 1496: 52 59. Bähler, M. and A. Rhoads. 2002. Calmodulin signalling via the IQ motif. FEBS Lett. 513: 107 113. Baker, J. P. and M. A. Titus. 1997. A family of unconventional myosins from the nematode Caenorhabditis elegans. J. Mol. Biol. 272: 523 535. Baker, J. P. and M. A. Titus. 1998. Myosins: matching motors with functions. Curr. Op. Cell Biol. 10: 80 86. Barylko, B., D. D. Binns, and J. P. Albanesi. 2000. Regulation of the enzymatic and motor activities of myosin I. Biochem. Biophys. Acta 1496: 23 35. Battelle, B. A., A. W. Andrews, B. G. Calman, J. R. Sellers, R. M. Greenberg, and W. C. Smith. 1998. A myosin III from Limulus eyes is a clock-regulated phosphoprotein. J. Neurosci. 18: 4548 4559. Beach, D. L., E. D. Salmon, and K. Bloom. 1999. Localization and anchoring of mRNA in budding yeast. Curr. Biol. 9: 569 578.

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References myosin Va through its distinct coiled-coil regions. FEBS Lett. 517: 233 238. Narasimhulu, S. B. and A. S. N. Reddy. 1998. Characterization of microtubule binding domain in the Arabidopsis kinesin-like calmodulin binding protein. Plant Cell 10: 957 965. Nascimento, A. A. C., R. G. Amaral, J. C. S. Bizario, R. E. Larson, and E. M. Espreafico. 1997. Subcellular localization of myosin-V in the B16 melanoma cells, a wild-type cell line for the dilute gene. Mol. Biol. Cell 8: 1971 1988. Neuhaus, E. M. and T. Soldati. 2000. A myosin I is involved in membrane recycling from early endosomes. J. Cell Biol. 150: 1013 1026. Nishikawa, S., K. Homma, Y. Komori, M. Iwaki, T. Wazawa, A. H. Iwane, J. Saito, R. Ikebe, E. Katayama, T. Yanagida, and M. Ikebe. 2002. Class VI myosin moves processively along actin filaments backward with large steps. BBRC 290: 311 317. Novak, K. D. and M. A. Titus. 1997. Myosin I overexpression impairs cell migration. J. Cell Biol. 136: 633 648. Novak, K. D. and M. A. Titus. 1998. The myosin I SH3 domain and TEDS rule phosphorylation site are required for in vivo function. Mol. Biol. Cell 9: 75 88. Novak, K. D., M. D. Peterson, M. C. Reedy, and M. A. Titus. 1995. Dictyostelium myosin I double mutants exhibit conditional defects in pinocytosis. J. Cell Biol. 131: 1205 1221. Oishi, N., H. Adachi, and K. Sutoh. 2000. Novel Dictyostelium unconventional myosin, myoM, has a putative RhoGEF domain. FEBS Lett. 474: 16 22. Oliver, T. N., J. S. Berg, and R. E. Cheney. 1999. Tails of unconventional myosins. Cell. Mol. Life Sci. 56: 243 257. Pasternak, C., J. A. Spudich, and E. L. Elson. 1989. Capping of surface receptors and concomitant cortical tension are generated by conventional myosin. Nature 341: 549 551. Pastural, E., F. J. Barrat, R. Dufourcq-Lagelouse, S. Certain, O. Sanal, N. Jabado, R. Seger, C. Griscelli, A. Fisher, and G. de Saint Basile. 1997. Griscelli disease maps to chromosome 15q21 and is associated with mutations in the myosin-Va gene. Nature Genet. 16: 289 292. Pastural, E., F. Ersoy, N. Yalman, N. Wulffraat, E. Grillo, F. Ozkinay, I. Tezcan, G. Gediköglu,

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pleted Arabidopsis thaliana genome sequence. Genome Biol. 2: 1 17. Reilein, A. R., S. L. Rogers, M. C. Tuma, and V. I. Gelfand. 2001. Regulation of molecular motor proteins. Int. Rev. Cytol. 204: 179 238. Reinhard, J., A. A. Scheel, D. Diekmann, A. Hall, C. Ruppert, and M. Bähler. 1995. A novel type of myosin implicated in signalling by rho family GTPases. EMBO J. 14: 697 704. Ridley, A. J. 2001. Rho GTPases and cell migration. J. Cell Sci. 114: 2713 2722. Robinson, D. N. and J. A. Spudich. 2000. Towards a molecular understanding of cytokinesis. Curr. Op. Cell Biol. 10: 228 237. Rock, R. S., S. E. Rice, A. L. Wells, T. J. Purcell, J. A. Spudich, and H. L. Sweeney. 2001. Myosin VI is a processive motor with a large step size. PNAS 98: 13655 13659. Rodionov, V. I., A. J. Hope, T. M. Svitkina, and G. G. Borisy. 1998. Functional coordination of microtubule-based and actin-based motility in melanophores. Curr. Biol. 8: 165 168. Rodriguez, O. C. and R. E. Cheney. 2002. Human myosin-Vc is a novel class V myosin expressed in epithelial cells. J. Cell Sci. 115: 991 1004. Rogers, S. L. and V. I. Gelfand. 1998. Myosin cooperates with microtubule motors during organelle transport in melanophores. Curr. Biol. 8: 161 163. Rogers, S. L. and V. I. Gelfand. 2000. Membrane trafficking, organelle transport, and the cytoskeleton. Curr. Op. Cell Biol. 12: 57 62. Rogers, S. L., R. L. Karcher, J. T. Roland, A. A. Minin, W. Steffen, and V. I. Gelfand. 1999. Regulation of melanosome movement in the cell cycle by reversible association of myosin V. J. Cell Biol. 146: 1265 1275. Rogers, S. L., I. S. Tint, P. C. Fanapour, and V. I. Gelfand. 1997. Regulated bidirectional motility of melanophore pigment granules along microtubules in vitro. PNAS 94:3720 3725. Rossanese, O. W., C. A. Reinke, B. J. Bevis, A. T. Hammond, S. I. B., J. O’Connor, and B. S. Glick. 2001. A role for actin, Cdc1p, and Myo2p in the inheritance of late Golgi elements in Saccharomyces cerevisiae. J. Cell Biol. 153: 47 61. Ruppert, C., J. Godel, R. T. Müller, R. Kroschewski, J. Reinhard, and M. Bähler. 1995. Localization of the rat myosin I molecules myr 1 and myr 2 and in vivo targeting

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References of their tail domains. J. Cell Sci. 108: 3775 3786. Schott, D., J. Ho, D. Pruyne, and A. Bretscher. 1999. The COOH-terminal domain of Myo2p, a yeast myosin V, has a direct role in secretory vesicle targeting. J. Cell Biol. 147: 791 807. Seidman, J. G. and C. Seidman. 2001. The genetic basis for cardiomyopathy: from mutation identification to mechanistic paradigms. Cell 104:557 567. Self, T., M. Mahony, J. Fleming, J. Walsh, S. D. M. Brown, and K. P. Steel. 1998. Shaker1 mutations reveal roles for myosin VIIA in both development and function of cochlear hair cells. Development 125: 557 566. Self, T., T. Sobe, N. G. Copeland, N. A. Jenkins, K. B. Avraham, and K. P. Steel. 1999. Role of myosin VI in the differentiation of cochlear hair cells. Dev. Biol. 214: 331 341. Separack, P. K., J. A. Mercer, M. C. Strobel, N. G. Copeland, and N. A. Jenkins. 1995. Retroviral sequences located within an intron of the dilute gene alter dilute expression in a tissue-specific manner. EMBO J. 14: 2326 2332. Sibley, L. D., S. Hakansson, and V. B. Carruthers. 1998. Gliding motility: an efficient mechanism for cell penetration. Curr. Biol. 8: R12 R14. Sil, A. and I. Herskowitz. 1996. Identification of asymmetrically localized determinant, Ash1p, required for lineage-specific transcription of the yeast HO gene. Cell 84: 711 722. Silvers, W. K. 1979. Dilute and leaden, the plocus, ruby-eye, and ruby-eye-2. In: Coat Colors of Mice: A Model for Mammalian Gene Action and Interaction. Edited by W. K. Silvers. New York: Springer-Verlag, pp. 83 89; 104 107. Sokac, A. M. and W. M. Bement. 2000. Regulation and expression of unconventional myosins. Int. Rev. Cytol. 200:197 304. Stephenson, R. S., J. E. O’Tousa, N. J. Scavarda, L. L. Randall, and W. S. Pak. 1983. Drosophila mutants with reduced rhodopsin content. In: Biology of Photoreceptors. Edited by D. Cosens and D. Vince-Prue. Cambridge: Cambridge University Press, pp. 471 495. Steyger, P. S., P. G. Gillespie, and R. A. Baird. 1998. Myosin1b is located at tip link anchors in vestibular hair bundles. J. Neurosci. 18: 4603 4615. Tabb, J. S., B. J. Molyneaux, D. L. Cohen, S. A. Kuznetsov, and G. M. Langford. 1998. Trans-

port of ER vesicles on actin filaments in neurons by myosin V. J. Cell Sci. 111: 3221 3234. Takizawa, P. A., A. Sil, J. R. Swedlow, I. Herskowitz, and R. D. Vale. 1997. Actin-dependent localization of an RNA encoding a cell-fate determinant in yeast. Nature 389: 90 93. Takizawa, P. A. and R. D. Vale. 2000. The myosin motor, Myo4p, binds ASH1 mRNA via the adapter protein, She3p. PNAS 97: 5273 5278. Tan, J. L., S. Ravid, and J. A. Spudich. 1992. Control of nonmuscle myosin phosphorylation. Ann. Rev. Biochem. 61: 721 759. Tauhata, S. B. F., D. V. dos Santos, E. W. Taylor, M. S. Mooseker, and R. E. Larson. 2001. High affinity binding of brain myosin-Va to F-actin induced by calcium in the presence of ATP. J. Biol. Chem. 276: 39812 39818. The May-Hegglin/Fechtner Syndrome consortium 2000. Mutations in MYH9 result in May-Hegglin anomaly, and Fechtner and Sebastion syndromes. Nature Genet. 26: 103 105. Titus, M. A. 1997. Unconventional myosins: new frontiers in actin-based motors. Trends Cell Biol. 7: 119 123. Titus, M. A. 1999. A class VII unconventional myosin is required for phagocytosis. Curr. Biol. 9: 1297 1303. Tsukita, S. and S. Yonemura. 1999. Cortical actin organization: lessons from ERM (Ezrin/ Radixin/Moesin) Proteins. J. Biol. Chem. 274: 34507 34510. Tuma, M. C. and V. I. Gelfand. 1999. Molecular mechanisms of pigment transport in melanophores. Pig. Cell Res. 12: 283 294. Tuxworth, R. I. and M. A. Titus. 2000. Unconventional myosins: anchors in the membrane traffic relay. Traffic 1: 11 18. Tuxworth, R. I., I. Weber, D. Wessels, G. C. Addicks, D. R. Soll, G. Gerisch, and M. A. Titus. 2001. A role for myosin VII in dynamic cell adhesion. Curr. Biol. 11: 318 329. Tyska, M. J. and M. S. Mooseker. 2002. MYO1A (brush border myosin I) dynamics in the brush border of LL-PK1-CL4 cells. Biophys. J. 82: 1869 1883. Uyeda, T. Q. P. and M. A. Titus. 1997. The myosins of Dictyostelium. In: Dictyostelium: a Model System for Cell and Developmental Biology. Edited by Y. Maeda, K. Inouye, and I. Takeuchi. Tokyo, Japan: University Academy Press, pp. 43 64.

1 The Myosin Superfamily: An Overview Uyeda, T. Q. P., C. Kitayama, and S. Yumura. 2000. Myosin II-independent cytokinesis in Dictyostelium: its mechanism and implications. Cell Str. Func. 25: 1 10. Verkhovsky, A. B., T. M. Svitkina, and G. G. Borisy. 1995. Myosin II filament assemblies in the active lamella of fibroblasts: their morphogenesis and role in the formation of actin filament bundles. J. Cell Biol. 131: 989 1002. Wagner, M. C., B. Barylko, and J. P. Albanesi. 1992. Tissue distribution and subcellular localization of mammalian myosin I. J. Cell Biol. 119: 163 170. Walch-Solimena, C., R. N. Collins, and P. J. Novick. 1997. Sec2p mediates nucleotide exchange on Sec4p and is involved in polarized delivery of post-Golgi vesicles. J. Cell Biol. 137: 1495 1509. Walsh, T., V. Walsh, S. Vreugde, R. Hertzano, H. Shahin, H. Haika, M. K. Lee, M. Kanaan, M. C. King, and K. B. Avraham. 2002. From flies’ eyes to our ears: mutations in a human class III myosin cause progressive nonsyndromic hearing loss DFNB30. PNAS 99: 7518 7523. Wang, A., Y. Liang, R. A. Fridell, F. J. Probst, E. R. Wilcox, J. W. Touchman, C. C. Morton, R. J. Morell, K. Noben-Trauth, S. A. Camper, and T. B. Friedman. 1998. Association of unconventional myosin MYO15 mutations with human nonsyndromic deafness DFNB3. Science 280: 1447 1451. Wang, F. S., J. S. Wolenski, R. E. Cheney, M. S. Mooseker, and D. G. Jay. 1996. Function of myosin-V in filopodial extension of neuronal growth cones. Science 273:660 663. Wei, Q., X. Wu, and J. A. Hammer, III. 1997. The predominant defect in dilute melanocytes is in melanosome distribution and not cell shape, supporting a role for myosin V in melanosome transport. J. Mus. Res. Cell Mot. 18: 517 527. Weil, D., S. Blanchard, J. Kaplan, P. Guliford, F. Gibson, J. Walsh, P. Mburu, A. Varela, J. Levilliers, M. D. Weston, P. M. Kelley, W. J. Kimberling, M. Wagenaar, F. Levi-Acobas, D. Larget-Piet, A. Munnich, K. P. Steel, S. D. M. Brown, and C. Petit. 1995. Defective myosin VIIA gene responsible for Usher syndrome type 1B. Nature 374: 60 61. Weil, D., P. Küssel, S. Blanchard, G. Lévy, F. Levi-Acobas, M. Drira, H. Ayadi, and C. Petit. 1997. The autosomal recessive isolated deaf-

ness, DFNB2, and the Usher 1B syndrome are allelic defects of the myosin-VIIA gene. Nature Genet. 16: 191 193. Wells, A. L., A. W. Lin, L. Q. Chen, D. Safer, S. M. Cain, T. Hasson, B. O. Carragher, R. A. Milligan, and H. L. Sweeney. 1999. Myosin VI is an actin-based motor that moves backwards. Nature 401: 505 508. Wes, P. D., X. Z. S. Xu, H. S. Li, F. Chien, S. K. Doberstein, and C. Montell. 1999. Termination of phototransduction requires binding of the NINAC myosin III and the PDZ protein INAD. Nature Neurosci. 2: 447 453. Wessels, D., J. Murray, G. Jung, J. A. Hammer, III, and D. R. Soll. 1991. Myosin IB null mutants of Dictyostelium exhibit abnormalities in motility. Cell Mot. Cytoskel. 20: 301 315. Wessels, D., D. R. Soll, D. Knecht, W. F. Loomis, A. De Lozanne, and J. Spudich. 1988. Cell motility and chemotaxis in Dictyostelium amebae lacking myosin heavy chain. Dev. Biol. 128: 164 177. Wessels, D., M. A. Titus, and D. R. Soll. 1996. A Dictyostelium myosin I plays a crucial role in regulating the frequency of pseudopods formed on the substratum. Cell Mot. Cytoskel. 33: 64 79. Westbroek, W., J. Lambert, and J. M. Naeyaert. 2001. The dilute locus and Griscelli syndrome: gateways towards a better understanding of melanosome transport. Pig. Cell Res. 14: 320 327. Williams, S. A., R. E. Hosein, J. A. Carcés, and R. H. Gavin. 2000. MYO1, a novel unconventional myosin gene affects endocytosis and macronuclear elongation in Tetrahymena thermophila. J. Euk. Microbiol. 47: 561 568. Wilson, S. M., R. Yip, D. A. Swing, T. N. O’Sullivan, Y. Zhang, E. K. Novak, R. T. Swank, L. B. Russell, N. G. Copeland, and N. A. Jenkins. 2000. A mutation in Rab27a causes the vesicle transport defects observed in ashen mice. PNAS 97:7933 7938. Win, T. Z., Y. Gachet, D. P. Mulvihill, K. M. May, and J. S. Hyams. 2000. Two type V myosins with non-overlapping functions in the fission yeast Schizosaccharomyces pombe: Myo52 is concerned with growth polarity and cytokinesis, Myo51 is a component of the cytokinetic actin ring. J. Cell Sci. 114: 69 79. Wirth, J. A., K. A. Jensen, P. L. Post, W. M. Bement, and M. S. Mooseker. 1996. Human myosin-IXb, an unconventional myosin with

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References a chimerin-like rho/rac GTPase-activating protein domain in its tail. J. Cell Sci. 109: 653 661. Wolenski, J. S. 1995. Regulation of calmodulinbinding myosins. Trends Cell Biol. 5: 310 316. Wu, X. and J. A. Hammer, III. 2000. Making sense of melanosome dynamics in mouse melanocytes. Pig. Cell Res. 13: 241 247. Wu, X., B. Bowers, K. Rao, Q. Wei, and J. A. Hammer, III. 1998. Visualization of melanosome dynamics within wild-type and dilute melanocytes suggests a paradigm for myosin V function in vivo. J. Cell Biol. 143: 1899 1918. Wu, X., B. Bowers, Q. Wei, B. Kocher, and J. A. Hammer, III. 1997b. Myosin V associates with melanosomes in mouse melanocytes: evidence that myosin V is an organelle motor. J. Cell Sci. 110: 847 859. Wu, C., V. Lytvyn, D. Y. Thomas, and E. Leberer. 1997a. The phosphorylation site for Ste20plike protein kinases is essential for the function of myosin-I in yeast. J. Biol. Chem. 272: 30623 30626. Wu, X., K. Rao, M. B. Bowers, N. G. Copeland, N. A. Jenkins, and J. A. Hammer, III. 2001. Rab27a enables myosin Va-dependent melanosome capture by recruiting the myosin to the organelle. J. Cell Sci. 114: 1091 1100. Wu, X., K. Rao, H. Zhang, F. Wang, J. R. Sellers, L. E. Matesic, N. G. Copeland, N. A. Jen-

kins, and J. A. Hammer, III. 2002a. Identification of an organelle receptor for myosin-Va. Nature Cell Biol. 4: 271 278. Wu, X., F. Wang, K. Rao, J. R. Sellers, and J. A. Hammer, III. 2002b. Rab27a is an essential component of the melanosome receptor for myosin Va. Mol. Biol. Cell 13: 1735 1749. Xu, P., K. I. Mitchelhill, B. Kobe, B. E. Kemp, and H. G. Zot. 1997. The myosin I binding protein Acan125 binds the SH3 domain and belongs to the superfamily of leucine-rich repeat proteins. PNAS 94: 3685 3690. Yamashita, R. A., J. R. Sellers, and J. B. Anderson. 2000. Identification and analysis of the myosin superfamily in Drosophila: a database approach. J. Mus. Res. Cell Mot. 21: 491 505. Yin, H., D. Bruyne, T. C. Huffaker, and A. Bretscher. 2000. Myosin V orientates the mitotic spindle in yeast. Nature 406: 1013 1015. Yonemura, S. and T. D. Pollard. 1992. The localization of myosin I and myosin II in Acanthamoeba by fluorescence microscopy. J. Cell Sci. 102: 629 642. Yoshimura, M., K. Homma, J. Saito, A. Inoue, R. Ikebe, and M. Ikebe. 2001. Dual regulation of mammalian myosin VI motor function. J. Biol. Chem. 276: 39600 39607. Young, P. E., A. M. Richman, A. S. Ketchum, and D. P. Kiehart. 1993. Morphogenesis in Drosophila requires nonmuscle myosin heavy chain function. Genes Dev. 7: 29 41.

2 Dynein Motors: Structure, Mechanochemistry and Regulation Stephen M. King

2.1

Introduction

Dyneins are multicomponent complexes that function to translocate their attached cargo towards the minus end of microtubules. These enzymes are involved in a wide variety of fundamental cellular processes including, mitosis, vesicular transport, the assembly and motility of cilia and flagella, and the generation of left– right asymmetry in the developing embryo (see Karki and Holzbaur (1999) and King (2000a) for recent reviews). Cytoplasmic dynein is essential in mammals, and in dynein-null mutants embryonic development cannot progress past the blastocyst stage (Harada et al., 1998). However, dynein is apparently not essential for the viability of unicellular organisms such as the budding yeast Saccharomyces cerevisiae, where the presence of kinesin-related proteins with overlapping functions allows for growth at almost wild-type rates (Li et al., 1993). Surprisingly however, analysis of the Arabidopsis genome has revealed that both dynein and the associated dynactin complex, which promotes dynein-mediated vesicle motility, may be completely absent in higher plants (Lawrence et al., 2001). In general, dyneins are constructed around one to three heavy chains (HCs*; Z 520 kDa) that contain the ATPase and motor activities. The kinesin and myosin cytoskeletal motors are distantly related to the GTPase subunit of heterotrimeric G proteins (Kull et al., 1998). In contrast, dyneins are members of the ancient AAA family of ATPases (Neuwald et al., 1999) that includes a wide variety of proteins such as bacterial Clp proteases, mammalian N-ethyl maleimide-sensitive vesicle fusion protein, the d’ subunit of the Escherichia coli DNA polymerase III clamploader, the RuvB DNA motor that is responsible for branch migration during resolution of Holiday junctions and the microtubule-severing protein katanin. The mechanisms employed to generate dynein diversity and thus allow for attachment to distinct cellular cargoes and appropriate responses to complex regulatory inputs, also are distinct from those found in myosins and kinesins. For the latter enzymes, a single cell may contain a large number of related motors each derived from a different gene or set of genes and designed to fill a particular func-

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2.2 Structural Organization of the Motor, Cargo-binding and Regulatory Components

tional niche. However, there is only one major dynein HC isoform in the cytoplasm even though many different cargoes must be moved; cilia and flagella contain a wider isoform variety, as multiple distinct motor activities are apparently required to generate complex axonemal waveforms. In the case of dynein, attaching individual motors to appropriate cellular cargoes involves multiple accessory proteins that are differentially expressed in different tissue types and whose function is subject to regulation through a variety of mechanisms. In this chapter, I will discuss the composition and structure of dynein motors from both cytoplasm and cilia/flagella. I will also consider the mechanisms by which dynein motor and cargo-binding activities are regulated, and finally briefly review dynein mechanochemistry and force production.1)

2.2

Structural Organization of the Motor, Cargo-binding and Regulatory Components

All dyneins are built around one or more heavy chain motor units. There are two major classes of dyneins defined by the number of HCs present in the particle and by the various HC-associated polypeptides (Table 2.1). The first class includes cytoplasmic dynein, outer arm dynein and the I1 type of inner dynein arms. These complexes contain two or three HCs, at least two ICs that are members of the WD-repeat protein family, and a series of LCs from the LC8, Tctex1/Tctex2 and 1) Abbreviations used: AAA, ATPases associated

intermediate chain; NSF, N-ethyl maleimide with cellular activities; HC, heavy chain; IC, vesicle fusion protein. intermediate chain; LC, light chain; LIC, light Table 2.1 Composition of cytoplasmic and flagellar dyneins.

Component

Cytoplasmic dynein

Outer arm dynein

Inner arm dynein I1

Inner arm dynein I2/3

HC

2

2 3

2

1

IC

2

2 3

3

LIC

?

LCs Ca2 -binding LC1 Thioredoxin LC8 Tctex1/2 LC7/roadblock Actin p28 Centrin

2 Dynein Motors: Structure, Mechanochemistry and Regulation

Figure 2.1. Structure and composition of flagellar outer arm dynein. (a) Lane from a Coomassie blue-stained 5 15 % acrylamide gradient gel illustrating the polypeptide composition of the three-headed Chlamydomonas outer dynein arm. The a, b, and g HCs are not resolved using this particular gel system. Components of the trimeric docking complex required for assembly within the axoneme (DC1-3) also copurify with the dynein arm; the DC2 polypeptide (not indicated) co-migrates with IC2. This figure was prepared by Dr Miho Sakato (University of Connecticut Health Center). (b) Quick-freeze/ deep-etch micrograph of two outer arm dynein particles from Chlamydomonas flagella (Goodenough and Heuser, 1984). The basal region is formed from the N-terminal portion of the three HCs and a series of accessory IC and LC components. Stalks terminating in a small micro-

tubule-binding domain protrude from each large globular domain; together, these structures derive from the C-terminal portion of the HCs. This micrograph was kindly provided by Dr John Heuser (Washington University School of Medicine). (c) Model of the Chlamydomonas outer dynein arm indicating the location of the various polypeptide components (modified from DiBella and King (2001) c Academic Press). A series of LCs (LCs1, 3, 4 and 5) interact directly with the HCs and appear to be involved in the regulation of motor activity. The basal complex associated with the N-terminal HC domain comprises a series of WD-repeat ICs and members of the LC8, Tctex1/Tctex2 and LC7/roadblock families. All these basal components have close homologs in cytoplasmic dynein.

LC7/roadblock families. There are also dynein-specific components such as the LICs in cytoplasmic dynein, which have no known homologs in flagellar dyneins. Likewise the regulatory HC-associated LCs of the outer dynein arm have not yet been found in either cytoplasmic or inner arm I1 dyneins. The second major dynein class consists of a series of inner arm dyneins termed I2/I3 that contain

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monomeric HCs. Unlike other dyneins, these complexes contain actin and either centrin or p28 LCs. The oligomeric status of the HC(s) within the 1b class of cytoplasmic dynein required for retrograde intraflagellar transport is currently unknown (Pazour et al., 1999a, Porter et al., 1999). Furthermore, the accessory proteins within this dynein type also remain to be identified. 2.2.1

Heavy Chains

These proteins have a mass of Z 520 kDa (Z 44500 amino acids) and consist of several distinct domains. The C-terminal portions of these molecules comprise both the globular head (Z 12 nm in diameter) and an Z 9-nm stalk that terminates in a microtubule-binding domain. The N-terminal HC region forms an apparently flexible domain that mediates interactions with other HCs within the complex and is also involved in associations with other components that are required for assembly, cargo binding and regulation (Fig. 2.1). The original sequence analyses of outer arm and cytoplasmic dynein HCs (Gibbons et al., 1991, Koonce et al., 1992, Ogawa, 1991) identified four P-loops (or Walker A boxes) conforming to the general consensus G-X4 -G-K-T/S that is involved in nucleotide binding. However, only the first P-loop was completely conserved amongst both flagellar and cytoplasmic dynein HCs (Koonce et al., 1992). Recent analysis (Neuwald et al., 1999) revealed that the dynein HC is a member of the extensive AAA class of ATPases (see Ogura and Wilkinson (2001) for a review) and actually contains six AAA modules. The first four AAA domains are defined by the presence of P-loops 1 4. The P-loops in the last two AAA modules were not identified previously as they are highly degenerate and do not conform to the consensus. A map of the g HC from Chlamydomonas outer arm dynein indicating the location of the AAA units and other functional sites is shown in Fig. 2.2a. The high-resolution structures of several AAA proteins including NSF (Yu et al., 1998), DNA polymerase III clamploader d’ subunit (Guenther et al., 1997), and the bacterial Hs1VU ATP-dependent protease (Wang et al., 2001) have been solved. This has allowed the structure of individual AAA domains within the dynein complex to be modeled by homology (Mocz and Gibbons, 2001). In general, each dynein domain contains all the sequence motifs that define the main individual secondary structure elements required to form the core a/b fold and the extended ahelical domain (Fig. 2.2b). Domain-specific segments join these basic structural elements. Nucleotide is bound in the cleft formed between the two main folds of the AAA domain. The a/b portion contains the residues required for binding ATP including the glycine-rich Walker A box or P-loop that provides several phosphate ligands, and the Walker B box, which contains two acidic residues necessary for coordination of Mg2. The a-helical unit provides residues that detect the presence/absence of the terminal phosphate and also the sensor 2 region that undergoes alteration in conformation following nucleotide hydrolysis.


Figure 2.2. Structure and organization of the dynein heavy chain. (a) Diagram of the dynein HC indicating the six AAA domains, the microtubule-binding region and the section involved in both HC HC and HC IC interactions. The six AAA domains and the apparently unrelated C-terminal domain are proposed to each correspond to a globular sub-domain observed within the dynein head by (Samsó et al., 1998). The numbering on this figure is for the g HC from the Chlamydomonas outer dynein arm. Adapted from King (2000b). (b) An averaged negative stain image of the globular head domain from Dictyostelium cytoplasmic dynein HC is shown at left. This region has a diameter of 13.5 nm and contains seven distinct globular sub-domains surrounding an apparently empty central core. This micrograph was kindly provided by Dr Michael Koonce (Wadsworth Center). The model proposed by King (2000b) for the arrangement of AAA domains within the dynein head also is shown. The color code corresponds to the individual AAA units mapped in panel (a). (c) The Ca backbone for

the molecular model of the first AAA domain from the b HC of sea urchin sperm outer arm dynein (pdb accession 1HN5) is shown. The individual structural elements derived from the conserved AAA motifs are indicated and colored separately. Domain-specific segments outside of the AAA consensus regions are indicated in gray. This model was calculated by Mocz and Gibbons (Mocz and Gibbons, 2001) and is based on the known structures of three AAA proteins: NSF, the d’ subunit of the E. coli DNA polymerase III clamp loader and the HsIVU ATP-dependent protease. (d) Ribbon diagram of the sea urchin b HC AAA#1 molecular model indicating the predicted location of ATP bound between the core a/b structure and the terminal helical domain. (e) Expanded view of the ATP-binding region, showing three residues (K1858, T1859 and R2027) predicted to be involved in coordination of the triphosphate tail. In other AAA proteins acidic residues in the positions equivalent to D1904 and D1905 are required for coordination of the Mg2 ion.

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Several models for how the various AAA domains are arranged within the dynein head have been proposed (Fan and Amos, 2001, King, 2000b, Vale, 2000). All are based on the EM averaging analysis of Samsó et al. (1998), which revealed that the globular head consists of a seven-membered ring of discrete sub-domains. The models proposed by King and Vale are similar in that each AAA domain is assigned sequentially to a sub-domain within the ring. They differ in that Vale proposes that one sub-domain observed by EM derives from the MT-binding stalk structure located between AAA#4 and AAA#5, whereas King argues that this stalk adopts multiple conformations and is missing from the EM images due to averaging. In this latter model, the seventh sub-domain corresponds to the C-terminal Z 40-kDa segment of the HC that is not accounted for in the Vale model. Furthermore, the HC region that forms the microtubule-binding stalk is only sufficient to form a coiled coil of Z 10 nm (G. K. King and S. M. King, unpublished data) which is almost precisely the length of the structure observed by EM (Goodenough and Heuser, 1984); it does not contain sufficient mass to also form a globular sub-domain equivalent to that derived from an entire AAA module. The third model proposed by Fan and Amos (2001) suggests that there is actually a seventh dynein AAA domain located N-terminal of AAA#1, although this is not identified as such by the algorithm of Neuwald et al. (1999). In this model, the AAA modules are arranged such that the microtubule-binding stalks from the two heads can dimerize. This orientation places the first AAA domain far from the N-terminal stem domain and would require an extended structure traversing the AAA ring. ImmunoEM studies using AAA module-specific antibodies provide an obvious approach to resolving the domain arrangement within the dynein head. To date, the available evidence suggests that only the first, highly conserved AAA domain can both bind and hydrolyze ATP. For example, scission of the HC peptide backbone by vanadate-mediated photocleavage at the V1 site completely disrupts ATPase activity (Gibbons et al., 1987). In this reaction, monomeric metavanadate is associated with ADP and bound at the g-phosphate location within the active site where it acts as the chromophore for the cleavage reaction. Analysis of products generated during the analogous cleavage of myosin II revealed that scission first requires conversion of one P-loop residue (Ser in myosin) to the corresponding aldehyde (Cremo et al., 1991). Although the dynein HC contains multiple Ploops, the site of vanadate-mediated cleavage occurs within the first AAA domain (King and Witman, 1987, Lee-Eiford et al., 1986). It is also possible to cleave dynein HCs at one or more additional sites (termed V2) by UV irradiation in the presence of multimeric vanadate species and Mn2, which serves to suppress cleavage at the V1 site (King and Witman, 1987, Tang and Gibbons, 1987). The V2 site(s) also likely occur within AAA domains #3 or #4 (ambiguity here derives from uncertainties in the fragment masses determined by gel electrophoresis). Equilibrium partition studies to estimate the number of nucleotide binding sites per dynein HC suggested that all four sites with intact P-loops might be functional (Mocz and Gibbons, 1996). However, fluorescence anisotropy of dynein-bound methylanthraniloyl ATP indicated that only two nucleotide-binding sites had physiologically relevant association constants (Mocz et al., 1998). In vitro microtubule


translocation assays also support the idea that nucleotide (probably ADP) binding at additional sites regulates motor activity (Wilkerson and Witman, 1995, Yagi, 2000). Photoaffinity labeling studies of flagellar dynein HCs using photoactive ATP analogs also raised the possibility of two distinct regions involved in nucleotide binding (King et al., 1989, Mocz et al., 1988). ATP can adopt one of two conformations when bound to protein. In the syn form, the adenine base is stacked over the ribose, whereas in the anti orientation the base extends away from the sugar. In NSF, ATP is bound in the syn conformation (Yu et al., 1998). However, dyneins preferentially hydrolyze nucleotide analogs that adopt the anti conformation suggesting that this is the preferred orientation (King et al., 1989, Omoto and Nakamaye, 1989). As the adenine base binds in a generally hydrophobic pocket of AAA domains that makes few specific contacts with the nucleotide, there may have been little requirement for the base conformation of bound nucleotides to be conserved. One marked feature of AAA domain-containing proteins is that neighboring domains can exert a strong influence over enzymatic activity. For example, in one AAA unit of the NSF hexamer, a Lys side-chain from the adjacent AAA domain occupies the position near the terminal g-phosphate that is normally taken by the water molecule necessary for hydrolysis (Yu et al., 1998). Only when this residue is moved slightly to allow water entry, can nucleotide hydrolysis occur. Thus, nucleotide hydrolysis by the six AAA units within NSF can be coordinated. In the g HC of Chlamydomonas outer arm dynein, a basic residue occurs at the analogous position only in AAA#5 and thus could potentially regulate (or prevent) hydrolysis of nucleotide bound to AAA#4. The possibility that an LC regulates g HC ATPase directly via a similar mechanism is discussed further below. Evidence that dynein AAA domains other than AAA#1 affect enzymatic function comes from the cloning of a dynein HC (termed left right dynein) that is encoded at the murine inversus viscerum (iv) locus. Mice bearing this mutation show random organ placement (i. e. Z 50 % exhibit situs inversus and Z 50 % situs solitus) due to immotile nodal cilia and the consequent failure to impose a left right axis during embryonic development. The iv mutation is a single base pair alteration that results in a highly conserved Glu residue being changed to Lys in a region close to the sensor 2 motif of AAA domain #2 (Supp et al., 1997, 1999). The region of the HC responsible for ATP-sensitive microtubule binding is located between AAA domains #4 and #5. EM analysis indicates that it consists of an Z 9-nm stalk terminating in a small globular unit (Gee et al., 1997, Goodenough and Heuser, 1984, Koonce, 1997). This HC region contains two segments of Z 85 residues each that are predicted to form a coiled coil of Z 10 nm. Between these regions is an Z 120-residue domain that actually interacts with the microtubule. Surprisingly, although microtubule binding is a property common to all dyneins, the region responsible is not highly conserved amongst dyneins of different origins, suggesting that only a few conserved residues are actually required for the interaction. Mapping experiments have identified three small clusters of residues important for microtubule binding in (or near) a section with similarity to the microtubule-binding site of MAP1b (Koonce and Tikhonenko, 2000). Furthermore,

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mutation of several nearby residues altered nucleotide-dependent microtubule release suggesting a direct role for this small domain in determining dynein-microtubule affinity (Koonce and Tikhonenko, 2000). However, the question of how this affinity is modulated in response to ATP hydrolysis at a site i 20 nm distant from the hydrolytic domain remains unanswered. Two mutations within the b HC from Chlamydomonas outer arm dynein have been identified that suppress flagella paralysis caused by the lack of radial spokes or the central pair microtubule complex (Huang et al., 1982). Interestingly, these cause small deletions within the coiled coil segments forming the microtubule-binding stalk structure suggesting that this region exerts a regulatory function (Porter et al., 1994). The Z 160-kDa N-terminal region of the HC consists of Z 1780 residues and mediates both HC HC and HC IC interactions (Habura et al., 1999, Tynan et al., 2000b). These overlapping interaction sites are located approximately 400 800 residues from the N-terminus. Mutational analysis in Chlamydomonas has revealed that the large HC motor domain is not required for assembly of the dynein particle. The strain oda4 -s7 expresses a truncated form of the outer arm b HC that is only Z 160 kDa in size and is competent to assemble with the remaining components of this complex (Sakakibara et al., 1993). A circular dichroism study of this region (termed fragment B) from the b HC of sea urchin outer arm dynein revealed a relatively low a-helical content of Z 23 % which is consistent with a non-helical flexible structure (Mocz and Gibbons, 1990). Intriguingly, secondary structure predictions suggest a much greater helical content (i 60 %). There are several possible mechanisms by which nucleotide hydrolysis at AAA#1 might be converted to mechanical force at a site Z 20 nm distant. Conformational changes occurring in AAA#1 could be propagated through the other AAA domains ultimately causing movement of the microtubule-binding stalk (King, 2000b). Alternatively, the site of the power stroke might be the junction between the N-terminal stem and the globular head (Samsó et al., 1998). In this model, the entire AAA domain head and microtubule-binding stalk would move. An additional unresolved question concerns how microtubule-binding affinity is controlled at this distant interaction site and has led to the proposal that there is a rotational as well as linear component to the dynein power stroke. As described in more detail below, several dyneins do cause microtubule rotation during in vitro translocation indicating that they generate torque (Vale and Toyoshima, 1988). Both flagellar and cytoplasmic dynein HCs are phosphorylated in vivo; in both systems, modification occurred mainly on Ser residues (Dillman and Pfister, 1994, King and Witman, 1994). In optic nerve, the cytoplasmic dynein HC was more heavily phosphorylated in vesicles moving in an anterograde direction than in those obtained from the total cellular pool (Dillman and Pfister, 1994). This suggests that HC phosphorylation may affect dynein function by shutting down motor activity so that dynein-bearing vesicles can be moved in the anterograde direction. In the Chlamydomonas flagellum, I2/3 inner arms HCs are heavily modified (Piperno and Luck, 1981). However, within the outer arm only the a HC becomes phosphorylated (King and Witman, 1994, Piperno and Luck, 1981). Quantitative analysis suggests that there are a least six phosphorylated sites in this molecule.


These are located in two main segments of the HC; one immediately preceding the first AAA domain and the second located approximately 90 kDa from the C-terminus (King and Witman, 1994). Although far apart in the linear sequence, the model for HC organization proposed here places these two domains close to each other near the ATP hydrolytic module (see Fig. 2.2a,b). As label is incorporated at these sites when isolated axonemes are incubated with [g-32P] ATP, the kinase(s) and phosphatase(s) required for HC modification must be integral components of the flagellar axoneme and in close proximity to their target sites within the outer arm. Mutant studies in Chlamydomonas have identified a series of extragenic suppressors that rescue flagellar paralysis due to lack of radial spokes (Piperno et al., 1992). These proteins (termed the dynein regulatory complex) are located at the base of the I2/3 inner arm system near the radial spokes (Gardner et al., 1994). Defects in several of these proteins reduce the number of inner arms assembled into the axoneme and also appear to affect the level of phosphorylation present on inner arm HCs (Gardner et al., 1994, Luck and Piperno, 1989, Piperno et al., 1994). Thus, this protein complex is involved in transducing regulatory signals from the radial spoke/central pair complex to the I2/3 HCs. 2.2.2

Intermediate Chains

Dyneins built from two or more HCs also contain two ICs that are members of the WD-repeat protein family. These components are located at the base of the soluble dynein particle (King and Witman, 1990, Sale et al., 1985) and interact both with the HCs and also with three different classes of LCs. IC components from outer arm, inner arm I1 and cytoplasmic dyneins have been implicated in attaching the motor to specific cellular cargoes (King et al., 1991, Vaughan and Vallee, 1995, Yang and Sale, 1998), although they are not sufficient and additional adaptors, such as dynactin and the outer arm docking complex, are required to mediate attachment to specific cargoes (Holzbaur et al., 1991, Schroer and Sheetz, 1991, Takada and Kamiya, 1994). Some outer arm dyneins and inner arm I1 contain additional IC components that are not members of the WD-repeat family. These are discussed further towards the end of this section. Also, it is now clear that the Lis-1 protein interacts with dynein/dynactin to modify its function and may even be an integral component of specific subsets of cytoplasmic dynein (Faulkner et al., 2000, Liu et al., 1999, Smith et al., 2000). ICs from outer arm, inner arm I1 and cytoplasmic dyneins all contain seven copies of the WD-repeat motif within the C-terminal region of the molecule (Mitchell and Kang, 1991, Paschal et al., 1992, Wilkerson et al., 1995, Yang and Sale, 1998) (Fig. 2.3a). The consensus sequence for this motif was defined by Neer et al. (1994) and is illustrated in Fig. 2.3b. At the structural level, the WD repeats consist of multiple four b strand units that fold as antiparallel sheets (Fig. 2.3c). The WD repeat motif forms the outermost strand of one sheet and the three innermost strands of the neighboring sheet. In Gb, the seven copies of the motif yield a seg-

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Figure 2.3. Structure and organization of dynein intermediate chains. (a) Diagram of IC1 and IC2 (aka IC78 and IC69) from the Chlamydomonas outer dynein arm, IC140 from Chlamydomonas inner arm I1 and the 2c isoform of mammalian cytoplasmic dynein IC74. All these proteins are characterized by the presence of seven WD-repeats although there are genespecific N- and C-terminal extensions such as coiled coil domains. The segments of IC74-2c involved in binding the LC8 (Lo et al., 2001) and Tctex1 LCs (Mok et al., 2001), and the phosphorylation site (S84) (Vaughan et al., 2001) that controls interaction with dynactin are also indicated. Mutational analysis has demonstrated that the N-terminal region of IC140 is not essential for its function in vivo (Perrone et al., 1998). (From DiBella and King, 2001, c Academic Press). (b) Consensus sequence for

the WD-repeat (Neer et al., 1994; Smith et al., 1999). Multiple repeats come together to form a toroidal b-propeller structure. The color code for strands a d refers to the WD-repeat structure shown in (c). (c) Structure of one WD-repeat from the Gb subunit of heterotrimeric G protein (Sondek et al., 1996) (pdb accession 1GP2). The strands are color-coded to correspond to the consensus sequence shown in (b). This structure is stabilized by hydrogen-bonding between the four conserved residues indicated. (d and e) Top and side views of a ribbon diagram of the WD-repeat region of Gb (pdb accession 1GP2). One blade of the propeller structure is shown in magenta. The four b strands corresponding to one WD-repeat are indicated in orange and form the outermost strand of one blade and the three innermost strands of the adjacent blade.


mented toroidal structure (Fig. 2.3d, e) that forms multiple protein protein interfaces with both the Ga and Gg subunits (Sondek et al., 1996). This region of the dynein ICs is predicted to adopt a similar fold. The N-terminal sections of these ICs are gene-specific. For example, in IC74 of cytoplasmic dynein this domain binds directly to the p150glued component of dynactin (Vaughan and Vallee, 1995), whereas IC1 of Chlamydomonas outer arm dynein interacts with a-tubulin (King et al., 1991). Genetic analysis in Chlamydomonas indicates that both IC1 and IC2 (aka IC80/IC78 and IC69/IC70) are required for outer arm dynein assembly (Mitchell and Kang, 1991, Wilkerson et al., 1995). Similarly, IC140 is needed for assembly of inner arm I1 (Perrone et al., 1998). Furthermore, reversion analysis has identified IC2 alleles that allow for the restoration of outer dynein arm assembly but do not increase flagellar beat frequency to wild-type levels, suggesting that IC2 is also required for regulation of motor activity (Mitchell and Kang, 1993). Mammals contain two genes for IC74. Multiple isoforms are generated in various tissues and during development by alternative splicing and differential phosphorylation that yield different N-terminal segments (Paschal et al., 1992, Pfister et al., 1996a, 1996b). Currently, five alternatively spliced variants (designated 1a, 1b, 2a, 2b, and 2c) of the two mammalian IC74 genes are recognized. All five isoforms are found in neurons, although, only IC74-1a is neuron-specific (Pfister et al., 1996a). The IC74 isoforms found in cortical neurons and glia are differentially expressed and phosphorylated (Pfister et al., 1996b). In contrast, only the 2c isoform has been identified in liver (Vaughan et al., 2001). This suggests that isoform diversity may not directly correlate with functional diversity. Interaction of IC74-2c with p150glued is regulated by phosphorylation of Ser-84 (Vaughan et al., 2001). Mutant forms of this IC that mimic the dephosphorylated state bound to p150glued in vitro and disrupted dynein-mediated vesicular transport when expressed in cultured cells. Conversely, mutation of Ser-84 to Asp (resembles the phosphorylated state) yielded protein with reduced capacity for binding p150glued and did not affect dynein transport. Recently, an additional mechanism for suppressing dynein activity during apoptosis has been identified in mammalian cells. Following the apoptotic signal, dynein-based motility is shut down by proteolytic cleavage of IC74 (and p150glued of dynactin) by caspases (Lane et al., 2001). In addition to two WD-repeat ICs, sea urchin outer arm dynein also contains a 91.6-kDa modular protein (here termed suIC1 to distinguish it from the Chlamydomonas WD-repeat protein) consisting of an N-terminal thioredoxin followed by three copies of the nucleoside diphosphate kinase catalytic domain and a highly acidic C-terminal segment (Ogawa et al., 1996). SuIC1 is intimately associated with the b HC. Thioredoxin-related LCs have been identified in the Chlamydomonas outer arm and are discussed further below. However, to data the NDK motif has not been observed in dyneins from this organism, although flagella from Chlamydomonas do exhibit NDKase activity (Watanabe and Flavin, 1976). Additional ICs have also been identified in outer dynein arms from sperm of several other species including trout (Gatti et al., 1989), mussel and the ascidian Halocynthia (Ogawa

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et al., 1996). Immunological data suggests that some of these components also contain NDK modules, although their molecular characteristics remain to be fully elucidated. The presence of catalytic NDKs within dynein suggests a requirement for NTPs other than ATP. Inner arm I1 contains both the WD-repeat IC140 protein that is encoded at the IDA7 locus and is required for assembly, and also a phosphoprotein (IC138) (Habermacher and Sale, 1997). Reconstitution experiments rebinding dyneins from wild-type or radial spoke-deficient mutants onto dynein-depleted axonemes revealed that inner arm I1 is an essential regulator of microtubule sliding (Smith and Sale, 1992). Sliding rates were increased by addition of kinase inhibitors and decreased by phosphatase inhibitors and added kinase (Habermacher and Sale, 1997). IC138 is the only phosphoprotein present in the I1 complex suggesting that is involved in the control of I1 motor activity. Both cAMP-dependent protein kinase and casein kinase 1 have been found to modify IC138 (Yang and Sale, 2000). Furthermore, several mutants with defects in phototaxis contain electrophoretic variants of IC138 that differ based on their level of phosphorylation (King et al., 1994). 2.2.3

Light Intermediate Chains

The LICs are Z 53 57-kDa polypeptides that have so far been identified only in cytoplasmic dynein. In both mammals and birds, there are two distinct LIC proteins (termed LIC1 and LIC2 in mammals) derived from separate genes, each of which can be phosphorylated to yield an additional electrophoretic variant. These proteins contain a P-loop motif and around this site show limited homology to the nucleotide-binding region of ABC transporters, which are involved in translocation of various components across membranes (Gill et al., 1994, Hughes et al., 1995). However, there is not yet any evidence to indicate that LICs actually bind nucleotide. Outside of the ABC transporter homology region, LICs show no obvious similarity with any other proteins. LIC1, but not LIC2, has been found to bind directly to pericentrin, which is an integral component of the centrosome (Tynan et al., 2000b). This suggests that like both the ICs and LCs (see below), LICs also are involved in cargo attachment. Interestingly, point mutations within the LIC1 P-loop had no effect on this activity. Furthermore, binding of LIC1 and LIC2 to the dynein HC is mutually exclusive suggesting that there are two, and possibly three, distinct cytoplasmic dynein pools defined by their LIC content; that is with either LIC1, LIC2 or possibly neither LIC bound (Tynan et al., 2000b). Examination of mitotic (metaphase) extracts of Xenopus eggs revealed a dramatic decrease in the amounts of membrane-associated dynein and dynactin when compared to interphase extracts (Niclas et al., 1996). This decrease correlated with a reduction in the amount of minus-end directed vesicular trafficking observed. Although no changes in the phosphorylation of either HCs or ICs were identified, a large (i 12-fold) increase in phosphorylation of LICs was observed. This suggests


a potential role for LICs in regulating motor function through control of dyneinmembrane interactions (Niclas et al., 1996). 2.2.4

The LC8 Light Chain Class

LC8 was originally identified in the outer dynein arm from Chlamydomonas flagella (King and Patel-King, 1995b). Subsequently, this very highly conserved molecule (Z 90 % sequence identity between algae and humans) was also found in both brain cytoplasmic dynein (King et al., 1996a) and the I1 class of inner dynein arms (Harrison et al., 1998), and shown to be required for intraflagellar transport (Pazour et al., 1998). Intriguingly, Z 80 % of brain LC8 is not associated with dynein and this 89-residue polypeptide also interacts with a variety of other protein complexes such as neuronal nitric oxide synthase (Jaffrey and Snyder, 1996), myosin V (Espindola et al., 2000), flagellar radial spokes (Yang et al., 2001) and IkBa (Crepieux et al., 1997). LC8 has also been proposed to mediate the interaction of a wide variety of cellular and viral components with motor complexes. Suggested cargoes include the pro-apoptotic factor Bim (Puthalakath et al., 1999), rabies virus P protein (Raux et al., 2000) and Drosophila Swallow (Schnorrer et al., 2000). LC8 is essential in multicellular organisms. For example, in Drosophila LC8 partial loss-of-function mutations lead to female sterility, defects in wing and bristle development and disruption of axonal circuitry. More severe mutations lead to embryonic lethality following the induction of apoptosis (Dick et al., 1996a, Phillis et al., 1996). In Aspergillus, LC8 defects result in the failure of nuclear migration along the hyphae (Beckwith et al., 1998). In contrast, null mutations for LC8 in both Chlamydomonas and Saccharomyces cerevisiae are not lethal (Dick et al., 1996b, Pazour et al., 1998). Indeed, no growth defect has been observed in either organism, although Chlamydomonas mutants lacking LC8 are unable to assemble flagella due to the failure of intraflagellar transport and consequently are non-motile (Pazour et al., 1998). The Chlamydomonas outer dynein arm also contains a highly divergent LC8 homolog (termed LC6) that is as phylogenetically distant from LC8 as the versions of this molecule found in higher plants, nematodes and budding and fission yeast. Intriguingly, the LC6 protein is not essential for dynein assembly and null mutants exhibit only a very minor swimming defect (G. P. Pazour, A. Harrison, G. B. Witman and S. M. King, unpublished data; and see Pazour and Witman, 2000). In dynein and myosin V, zero-length crosslinking revealed that LC8 exists in situ as a dimer (Benashski et al., 1997). In vitro, the monomer dimer equilibrium is reversible with a Kd of 12 mM at pH 7 (Barbar et al., 2001). As the peptide-binding site is formed at the junction of the two monomers, conversion of dimer to monomer represents a potential mechanism for controlling LC8 peptide interactions. Although immunological analysis of brain cytoplasm indicated that a relatively large amount of LC8 was present (King et al., 1996a), the actual intracellular LC8 concentration is unknown and it is therefore unclear whether the monomer dimer transition occurs in vivo and plays any significant role in regulating LC8 activity.

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Figure 2.4. Structure of the LC8 dynein light chain. (a) Ribbon diagram of the LC8 dimer Xray structure solved by Liang et al. (1999) (pdb accession 1CMI). The dimer interface consists of two five-stranded b sheets. Intriguingly, the b3 strand from one monomer protrudes and interacts with the b sheet from the second monomer. The figure has been color-coded such that the strands forming each sheet are a single color. The two clefts formed at the dimer interface provide two identical peptide-binding surfaces. (b) Surface representation of LC8 indicating the peptide-binding cleft. Residues that form hydrogen bonds with the peptide are indicated in green, those that make hydrophobic contacts are in red. This figure is related to the ribbon diagram shown in (a) by a 90o rotation

about the x-axis. (c) Sequence alignment of the human LC8a, LC8b and LC8c isoforms defined by Wilson et al. (2001). The accession numbers are U32944, AI719240 and CAB41220, respectively. Positions at which LC8b and LC8c differ from the LC8a sequence are blocked in magenta and red, respectively. The secondary structure is indicated above the alignment and is color-coded for the leftmost monomer shown in panel (a). (d and e) Two views of the LC8 surface indicating the residues of LC8a that are changed in LC8b (magenta) and LC8c (red). Nearly all sequence changes affect residues exposed at the molecular surface and, except for V66 and H68, occur away from the peptidebinding cleft.


NMR and X-ray structures of the LC8 dimer in complex with a variety of binding peptides are now available (Fan et al., 2001, Liang et al., 1999, Tochio et al., 1998). Each monomer provides two a helices and five b strands which form an anti-parallel sheet (Fig. 2.4a). Importantly, at the dimer interface, the b3 strand from one monomer protrudes such that it hydrogen bonds to a four-stranded b sheet formed by strands b1, b2, b4 and b5 from the second monomer. Intermolecular hydrogen bond formation and the large hydrophobic surface area buried at the dimer interface (Z 1300 Å2) accounts for the high stability of the LC8 dimer (Fan et al., 2001, Liang et al., 1999). The symmetric structure of the LC8 dimer provides two identical peptide-binding channels formed at the dimer interface (Fig. 2.4b). These binding sites are relatively pliant and to date, the structural basis for the binding of two distinct consensus LC8 interaction sequences (K/R-X-T-Q-T and G-I-Q-V-D-R) has been elucidated (Fan et al., 2001; Liang et al., 1999). Both peptides form a b strand that hydrogen bonds to the LC8 b3 strands at the dimer interface. The interactions of the target peptide with LC8 mainly occur either through hydrogen bonds to backbone amides and carbonyls, or are hydrophobic in nature rather than being mediated via side chain side chain interactions. This suggests that elements outside of the peptide-binding surface must contribute to the specificity of LC8 target protein interactions. In vitro studies have revealed that the N-terminal region of IC74 (residues 1 289) from Drosophila is relatively unstructured, but that a significant increase in secondary structure content occurs following binding of LC8 (Makokha et al., 2002). Recently, it has been recognized that there are several distinct classes of LC8 (termed LC8a, LC8b and LC8c) in mammals that are expressed in different tissues (Wilson et al., 2001), and potentially may mediate interactions with distinct protein cargoes. The differences between these variants reside mainly on surface residues (Fig. 2.4c e). Interestingly, most substitutions (except for two in LC8c) also occur away from the known peptide binding channels further suggesting that other regions of the molecule may be involved in protein protein associations. 2.2.5

The Tctex1/Tctex2 Light Chain Class

The murine Tctex1 and Tctex2 proteins were originally identified as candidates for distorter proteins involved in the non-Mendelian transmission of variant forms of an Z 40-Mb region of mouse chromosome 17 known as the t haplotypes (Huw et al., 1995, Lader et al., 1989, O’Neill and Artzt, 1995). Due to recombination suppression by a series of large inversions, these variant haplotypes are inherited as a single unit. Nearly all the progeny of heterozygous males inherit the t haplotype form of the chromosome in a process known as transmission ratio distortion or meiotic drive (see Olds-Clarke (1997) and Silver (1993) for reviews). Furthermore, males bearing complementing t haplotypes are completely sterile; mice homozygous for non-complementing haplotypes are not viable due to the presence of recessive lethal mutations. This phenomenon arises due to defects in spermiogenesis such that sperm bearing the wild-type copy of chromosome 17 are defective

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whereas those bearing the t haplotype are apparently normal. Genetic studies suggested that this was caused by the action of a t mutant responder protein on a series of mutant distorters (Lyon, 1984). Identification of Tctex1 and Tctex2 as components of cytoplasmic and flagellar dynein provided the first indication that the underlying biological cause of this phenomenon might be disruption of dynein motor function (DiBella et al., 2001, Harrison et al., 1998, King et al., 1996b, Patel-King et al., 1997). Recently, the responder has been identified as a mutant version (Tcr) of a sperm motility kinase (Smok) with diminished catalytic activity and the strongest distorter has been found to encode an axonemal dynein HC (Fossella et al., 2000), providing further support for this hypothesis. Tctex1 and its close homolog rp3 (55 % sequence identity with Tctex1) have both been demonstrated to be integral components of cytoplasmic dynein that interact with the ICs at the base of the soluble particle (King et al., 1998, 1996b). Tctex1 is also present in the sperm flagellum (O’Neill and Artzt, 1995) and in Chlamydomonas is an integral component of the I1 class of inner dynein arm (Harrison et al., 1998). In cytoplasmic dynein, Tctex1 interacts with IC74 via a K/R-K/R-X-X-K/R consensus sequence (Mok et al., 2001). Tctex1 also has been found to bind several other cellular components including Doc-2 (Nagano et al., 1998), Fyn tyrosine kinase (Kai et al., 1997, Mou et al., 1998), rhodopsin (Tai et al., 1999) and the developmentally regulated GTPase NEDD-3 (A. Harrison and S. M. King, unpublished data). As Tctex1 is dimeric (DiBella et al., 2001) and therefore presumably contains two peptide-binding sites, this LC can mediate association of cytoplasmic dynein with specific cellular cargoes as has also been observed for LC8. Indeed, analysis of the rhodopsin Tctex1 interaction has provided evidence to support direct cargo attachment via this LC (Tai et al., 1999). Intriguingly, rhodopsin did not interact with rp3, and indeed overexpression of rp3 in Madin Darby canine kidney cells displaced the endogenous Tctex1 present in cytoplasmic dynein and disrupted apical Tctex1-mediated rhodopsin transport (Tai et al., 2001). This suggests that attachment of different LCs to a particular dynein particle represents one mechanism for defining cargo-binding specificity. Tctex2 was originally described as a sperm membrane protein (Huw et al., 1995). However, cloning of the Chlamydomonas homolog LC2 revealed that this protein is actually a component of the outer dynein arm (Patel-King et al., 1997) and defines one major branch within a protein family that includes Tctex1 and rp3. Subsequently, mammalian Tctex2 has been identified in cytoplasmic dynein from kidney and spleen but not in samples obtained from brain, liver or testis (DiBella et al., 2001). Examination of the EST database has revealed several additional Tctex2-related proteins in mammals that exhibit tissue-specific distributions. Identification of LC2 (Tctex2)-defective mutants in Chlamydomonas demonstrated that this LC class is essential for assembly of the outer dynein arm. Two insertional alleles at the oda12 locus have been identified (Pazour et al., 1999b). One allele is a complete null and fails to assemble any outer arms, while the second lacks the 3l end of the gene, is partially functional and allows for the assembly of a few arms at apparently random locations within the axoneme. These defects are readily rescued by transforming the oda12 mutant strain with additional copies of the wild-type gene.


Although Tctex1 clearly plays an important role in cytoplasmic dynein function, analysis of a Drosophila mutant revealed that this LC is not essential for viability. Flies homozygous for P-element insertions that disrupt the 5l-UTR of the only Tctex1 gene in this organism, reach adulthood but are male sterile as the sperm produced are non-motile (Caggese et al., 2001). This surprising result suggests either that Tctex1 function is not absolutely required for cytoplasmic dynein activity or that other cellular proteins can functionally replace Tctex1. In solution, Tctex1 is dimeric whereas Tctex2 is a monomer (DiBella et al., 2001). These observations agree with the apparent stoichiometry of these proteins within soluble dynein particles (King et al., 1996b, King and Witman, 1989). Mammalian cytoplasmic and Chlamydomonas inner arm I1 dyneins both contain two Tctex1-related LCs per particle, whereas there is only a single copy of the Tctex2 homolog LC2 within the outer dynein arm. However, it remains unclear whether Tctex1 and rp3 can form heterodimers and thus whether some cytoplasmic dynein particles contain one copy of each LC. Structural studies of the Tctex1 dimer in solution have revealed a secondary structure remarkably similar to that found for LC8, with two N-terminal a helices followed by four b strands that fold to form an antiparallel sheet (Mok et al., 2001, Wu et al., 2001). Thus, although the tertiary structure has not yet been solved, it seems likely that it will be closely related to that of LC8. 2.2.6

The LC7/roadblock Light Chain Class

Members of the third LC class associated with the intermediate chains at the base of the soluble dynein particle have been found in both cytoplasmic dynein and the flagellar outer arm (Bowman et al., 1999). Chlamydomonas LC7 is encoded at the ODA15 locus and is essential for the assembly of the outer dynein arm (Pazour and Witman, 2000). In Drosophila, mutations in the cytoplasmic dynein roadblock LC result in chromosome segregation defects, axonal degeneration and disruption of intracellular transport. Ultimately, these phenotypes lead to pupal lethality. Interestingly, one roadblock allele (termed roblz) produces a poisonous product that yields a phenotype (larval-lethality) more severe than that observed in the homozygous null strain (Bowman et al., 1999). There are six roadblock variants in Drosophila, including the bithoraxoid protein, which has been implicated in the development of thoracic and abdominal parasegments (Hogness et al., 1985; Lipshitz et al., 1987). However, it is not yet clear that all isoforms are dynein-associated. In mammals, there are two distinct forms of this LC (termed roadblock 1 and 2). To date, only the roadblock 2 LC has been found in cytoplasmic dynein from brain, although roadblock 1 is also competent to bind IC74 (Wilson et al., 2001). Intriguingly, expression of two human roadblock isoforms is dramatically altered in hepatocellular carcinoma tissues (Jiang et al., 2001). In both rats and monkeys, the expression of roadblock 1 in visual cortex is rapidly downregulated in response to visual stimulation, suggesting that roadblock protein is turned over rapidly and that

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roadblock expression in neurons is regulated by a mechanism tightly linked to sensory inputs (Ye et al., 2000). Recent sequence analysis has identified LC7/roadblock homologs not only in eukaryotes, but also in archaea and bacteria indicating that these proteins are of very ancient origin (Koonin and Aravind, 2000). Moreover, genetic analysis of the Myxococcus xanthus homolog (mglB) suggests a possible role for this superfamily in the regulation of NTPase function as it controls the activity of mglA which is a member of the Ras/Rab/Rho family of GTPases (Spormann and Kaiser, 1999). 2.2.7

Heavy Chain-associated Regulatory Light Chains

Thus far, potential regulatory LCs associated directly with the HCs have been identified only in the outer dynein arm. In Chlamydomonas these LCs fall into three distinct categories thioredoxin-like, calmodulin-like and a novel leucine-rich repeat protein (termed LC1) that is associated with the motor domain itself. Furthermore, a regulatory LC that controls dynein function in response to cAMP-dependent phosphorylation has been identified in Paramecium ciliary dynein. Together, these observations suggest that dyneins are subject to multiple regulatory inputs that control the activity of individual HCs.

Light chain 1 This Z22-kDa protein (Benashski et al., 1998) is a member of the SDS22 subclass of leucine-rich repeat proteins defined by Kajava (1998). Homologs have been identified in Chlamydomonas, sea urchins, Drosophila and mammals. In Chlamydomonas, two copies of LC1 are associated directly with the g HC and bind to the first two AAA domains. Furthermore, zero-length crosslinking revealed that this LC also interacts in situ with an additional axonemal protein of MrZ 45,000 that has not yet been identified. LC1 consists of an N-terminal helix, a central barrel formed from six bba motifs (the LRRs) that fold as a right-handed spiral, and a C-terminal helical domain that protrudes from the main protein axis (Fig. 2.5a). Structural comparisons revealed a strong similarity between LC1 and the U2A’ component of the human spliceosome with a backbone r. m. s. d. of 3.7 Å. This similarity includes the presence of a major hydrophobic patch on the large b sheet face of LC1 that in U2A’ binds a helical segment from U2BL. As the interaction between LC1 and the g HC is hydrophobic in nature, it is likely that this patch mediates the association (Fig. 2.5b,c). In this case, the protruding C-terminal domain will be oriented such that it would insert into the HC AAA domain. Located at the tip of this domain are two Arg residues that could make ionic contacts with g HC residues or possibly serve to control ATPase activity by displacing the water molecule adjacent to the g-phosphate that is required for hydrolysis. Small movements of these motifs could allow water entry and result in a round of ATPase activity. Alternatively, these Arg residues may act in a manner analogous to the ‘arginine fingers’ which have been observed in the GAPs associated with the Ras and Rho GTPases (Rittinger et al., 1997; 2.2.7.1


Figure 2.5 Structure of the motor domain-associated light chain. (a) Ribbon diagram of the mean LC1 structure (pdb accession 1DS9) determined by NMR spectroscopy (Wu et al., 2000). This 22-kDa protein consists of an Nterminal helix (a1), six bba motifs formed from the leucine-rich repeats that fold as a righthanded spiral and a C-terminal helical domain that protrudes from the main protein axis.

(From DiBella and King (2001) c Academic Press). (b and c) Two views of the molecular surface related by a 90o-rotation about the xaxis. Residues forming the hydrophobic patch found on the larger b sheet face that is predicted to bind the g HC (magenta strands in panel (a)) are indicated in green. The two Cterminal Arg residues that likely protrude into the motor domain are shown in red/orange.

Scheffzek et al., 1997). Upon GAP binding, the Arg residue contacts the g-phosphate and the O atom bridging the b- and g-phosphates of GTP and activates GTPase activity by stabilizing the transition state. In these systems, nucleotide hydrolysis is very slow in the absence of the GAP-derived Arg residue.

Calmodulin-related light chains The g HC from Chlamydomonas outer arm dynein is distinct in that it contains a second LC (LC4) associated with the N-terminal stem domain (Pfister et al., 1982, M. Sakato and S. M. King, unpublished data). This protein is a member of the calmodulin superfamily and contains four predicted helix-loop-helix motifs, two at either end of a central helical region (King and Patel-King, 1995a). However, only the N-terminal motifs conform precisely to the EF-hand consensus for Ca2 -binding loops. Direct Ca2 -binding assays revealed that LC4 indeed binds only one Ca2 with a KCa 3 q 10 5 M. In Chlamydomonas, the photoshock response involves an alteration in waveform and reversal of swimming direction that is signaled by a dramatic increase in in2.2.7.2

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traflagellar Ca2 to 10 4 M (Bessen et al., 1980). This response has been described as aberrant (Mitchell and Rosenbaum, 1985) or missing (Kamiya and Okamoto, 1985) in two different outer arm-deficient strains. Recent experiments using an in vitro microtubule binding assay suggest that Ca2 binding by LC4 may directly control ATP-dependent interactions of the g HC with microtubules (Sakato and King, 2001). Centrin, which is associated with the basal body-associated contractile fibers (Taillon et al., 1992), has been identified within a subset of the inner arms belonging to the I2/3 class (Piperno et al., 1992). This protein binds two Ca2 with high affinity (KCa 1.2 qx 10 6 M) and two with much lower affinity (KCa 1.6 qx 10 4 M) (Weber et al., 1994). Intriguingly, binding of this LC and p28 appears to be mutually exclusive; however, both appear to mediate association of actin with this inner arm class (Yanagisawa and Kamiya, 2001). An additional calmodulin-related protein with predicted EF-hands has been identified within the docking complex required for assembly of the Chlamydomonas outer dynein arm (Casey et al., 1998).

Thioredoxins Two distinct thioredoxin-related proteins LC5 and LC3 are associated with the Nterminal regions of the a and b HCs from the Chlamydomonas outer dynein arm (Patel-King et al., 1996). Furthermore, suIC1 from sea urchin sperm flagella outer arm dynein is a modular protein that contains a thioredoxin unit at the Nterminus (Ogawa et al., 1996). These proteins contain perfect copies of the redox-active site (W-C-G-P-C-K). Both Chlamydomonas LCs can be purified on the basis of a high affinity interaction with trivalent arsenic forming a covalent dithioarsine ring that may be reduced by mono- and dithiol reductants. Indeed, the outer dynein arm, but not the inner arms (which do not contain redox-active components), could be purified from a crude extract on the basis of this interaction (Patel-King et al., 1996). Recent analysis has revealed that ATPase activity of the g HC (but not the a or b HCs) is specifically enhanced by sulfhydryl oxidation (Harrison et al., 2002). Identification of these redox-active proteins in both sperm and Chlamydomonas flagella suggests that redox-based mechanisms are employed to manipulate dynein function and may be of general importance for flagellar motility. Intriguingly, a redox-activated cAMP-dependent tyrosine phosphorylation cascade has been proposed to control sperm capacitation (acquisition of swimming ability) in humans (Aitken et al., 1997). 2.2.7.3

p29 (cAMP-dependent phosphoprotein) Further evidence that LCs control dynein HC motor function has come from analysis of p29, which co-purifies with the outer arm from Paramecium cilia (Bonini and Nelson, 1990). The swimming velocity of this organism is increased when cells are treated with a membrane permeant cAMP analog. The p29 protein was 2.2.7.4


initially identified as the major axonemal polypeptide phosphorylated in a cAMPdependent manner (Hamasaki et al., 1991). Subsequent analysis determined that the rate of outer dynein arm-driven microtubule translocation was enhanced (by Z 2-fold) in samples treated with cAMP and protein kinase A, and in those where thiophosphorylated p29 had been rebound to the outer arm (Barkalow et al., 1994). 2.2.8

Light Chains Associated with Inner Arms I2/3

There are three accessory proteins associated with the various monomeric HCs that comprise this class of inner arms. Actin is found in all of these dyneins, although the role that it plays remains completely unknown (Kagami et al., 1990; Piperno et al., 1990). In Chlamydomonas, actin is encoded at IDA5 and mutations at this locus result in defects in several actin-requiring functions such as formation of the fertilization tubule required for mating (Kato-Minoura et al., 1997). These mutants also exhibit swimming defects as they lack several of the monomeric inner arms. However, two inner arm subclasses (termed b and g) remain competent to assemble within the axoneme as a result of incorporating a novel actin-like protein (65 % identical to actin) that is upregulated in the actin mutant and can functionally replace actin in these two subspecies (Kato-Minoura et al., 1998). Dyneins of this inner arm class differ in whether they contain centrin (see above) or p28 which is encoded at the IDA4 locus and is essential for inner arm assembly (LeDizet and Piperno, 1995; Piperno et al., 1990).

2.3

Mechanochemistry and Motility

The essential parameters of the dynein mechanochemical cycle have been determined for the intact outer dynein arm from Tetrahymena cilia using stopped flow and rapid chemical quench kinetic techniques, by Johnson and colleagues. This dynein contains three distinct HCs that likely have different motor properties (see below); unfortunately no information is available concerning the kinetics of cytoplasmic dynein mechanochemistry. The general pathway is shown in Fig. 2.6. In the absence of nucleotide, dynein is tightly bound to the microtubule in the rigor state. Binding of ATP and subsequent release from the microtubule are both rapid events occurring at 4.7 q 106 M 1 s 1 and 1000 s 1, respectively (Porter and Johnson, 1983). The rate of ATP hydrolysis yielding ADP and inorganic phosphate is 60 s 1 (Johnson, 1983). The large change in free energy required for dissociation of the dynein-microtubule complex was found to occur upon nucleotide binding not hydrolysis (Holzbaur and Johnson, 1986). Dissociation of the complex was also observed upon addition of the non-hydrolyzable nucleotide analog AMPPNP (Chilcote and Johnson, 1989). Identification of a pre-steady state phosphate burst derived from the rapid accumulation of dynein-ADP-Pi implied that

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2.3 Mechanochemistry and Motility

The dynein mechanochemical cycle. A scheme illustrating the mechanochemical cycle derived from the work of Johnson and colleagues on Tetrahymena outer arm dynein is shown. In the absence of ATP (rigor state), dynein (D) is tightly bound to the microtubule (M). ATP binding is fast and causes the rapid release of dynein from the microtubule. ATP is

Figure 2.6.

rapidly hydrolyzed, whereas release of both products, inorganic phosphate (P) and ADP, is relatively slow. Following phosphate release, dynein-ADP rebinds the microtubule and the rate-limiting step (ADP release) is coupled to conformational change. Redrawn and modified from Chilcote and Johnson (1989).

ATP hydrolysis is faster than product release which thus becomes the rate-limiting step in the cycle (Johnson, 1983; Shimizu et al., 1979; Takahashi and Tonomura, 1979). In the absence of microtubules, the rate of product release is Z 8 s 1. Kinetics of vanadate inhibition by formation of a dynein ADP vanadate complex demonstrated that ADP release occurs subsequent to phosphate release (Shimizu and Johnson, 1983). This was confirmed by Holzbaur and Johnson (1986), who examined the exchange of 18O from [18O]-phosphate to water. In order for the exchange reaction to occur, [18O]-phosphate must bind the dynein ADP complex. Thus, the phosphate generated by ATP hydrolysis must have already left the active site. The rates of ADP and phosphate release have been determined to be Z 20 s 1 and 60 70 s 1, respectively. As these values are similar, it is likely that both events are partially rate limiting. Importantly, the rate of ADP release is increased by the presence of microtubules and leads to enhanced microtubule activated ATPase activity (Omoto and Johnson, 1986). During the mechanochemical cycle, dyneins generate a force up to Z 4.7 pN and move along the microtubule with a step size of 8 nm (the length of one tubulin dimer) which is the same as kinesin (Hirakawa et al., 2000). At high motor densities, flagellar outer arm dyneins (containing two or three HCs) transport microtubules in an in vitro assay at rates 6 mm s 1 (Paschal et al., 1987a; Vale and Toyoshima, 1988) whereas rates for mammalian cytoplasmic dynein are Z 1.25 mm s 1 (Paschal et al., 1987b). The single HC present within the b/IC1 subunit of sea urchin outer arm dynein translocated microtubules at a rate of Z 10 mm s 1 (Sale and Fox, 1988). When beads coated with low densities of dynein (containing multiple HCs) were used, processive movement over long distances was observed (Wang et al., 1995), even though dynein has a low duty ratio and spends most of the mechanochemical cycle detached from the microtubule. This implies that action of the multiple heads within the dynein particle is coordinated. Single molecules of Tetrahymena outer arm dynein move processively only at low ATP concentrations below 20 mM, suggest-


ing the presence of two distinct motile activities within the complex (Hirakawa et al., 2000). However, nanometer-scale tracking revealed that unlike kinesin, dynein did not track along individual protofilaments but rather moved randomly across the available microtubule surface (Wang et al., 1995). When loosely bound to microtubules, dynein appears to undergo thermally driven one-dimensional diffusion along the microtubule (Vale et al., 1989). One important feature of dynein-based motility is that different HCs exhibit distinct motor properties. For example, the b/IC1 subunit from sea urchin outer arm dynein was found to translocate microtubules in an in vitro assay, but did not form a rigor bond with the microtubule when ATP was removed (Moss et al., 1992a). In contrast, the a HC subunit bound microtubules and co-sedimented with them in the presence and absence of ATP. Furthermore, incubation of both the a HC and intact outer arm dynein with microtubules resulted in ATP-dependent microtubule bundling suggesting that this HC forms both an ATP-sensitive rigor bond and also is involved in ATP-insensitive microtubule binding (Moss et al., 1992b). Recently, individual molecules of a single-headed inner arm motor (subspecies c from the Chlamydomonas flagellum) have been observed to act as processive motors even though the dynein has a low duty ratio and should be driven away from the microtubule by thermal diffusion (Sakakibara et al., 1999). Thus, at least in this dynein type additional mechanisms are present to ensure that the motor remains associated with the microtubule. Intriguingly, when placed under high load, this particular dynein slips backward on the microtubule suggesting that the action of this dynein type may be responsible for the high frequency (Z 300 Hz) vibration observed in quiescent axonemes (Kamimura and Kamiya, 1989). In vitro analysis has also revealed that outer arm dynein and several distinct inner arm species rotate the microtubule during translocation at rates of 1 3 Hz (Kagami et al., 1990; Vale and Toyoshima, 1988). This requires that torque be generated during the mechanochemical cycle and raises the possibility that rotation is due to biased tracking over the microtubule surface.

2.4

Dynein Deficiencies and Disease

Defects in the dyneins that power cilia and flagella lead to a wide variety of problems in multicellular organisms that reflect the many and diverse locations at which these organelles act. For example, flagella are required for male fertility, while cilia line the bronchial airways in the lungs, the ependymal lining of the brain and female reproductive tract. Furthermore, these organelles are needed to define the basic left right axis of the developing embryo (see Chapter 15 by Fischer). Genetic analysis in Chlamydomonas revealed that defects in dyneins and other ciliary structures such as the central pair microtubules and radial spokes lead to poorly motile or paralyzed flagella (see Mitchell (2000) for a review). Similarly, many instances have been reported in humans of immotile cilia or Kartagener’s syndrome in which cilia exhibit ultrastructural defects analogous to those observed

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2.4 Dynein Deficiencies and Disease

in Chlamydomonas mutants (Afzelius, 1979). Although many phenotypes such as poor sperm motility and severe bronchial problems are readily explained, an initially puzzling observation was that patients with immotile cilia syndrome often exhibit situs inversus where the major organs are positioned as the mirror image of their normal location. As discussed above, cloning of the mouse iv gene uncovered a single residue alteration within an axonemal dynein HC that leads to immotile cilia at the node of the developing embryo (Supp et al., 1997). Similarly, defects in a ciliary kinesin homolog KIF3B also result in inversion of organ placement in mice (Nonaka et al., 1998). Surprisingly, imaging of wild-type nodal ciliary motility did not reveal a standard ciliary waveform, but rather an apparently rigid ciliary shaft that rotated in a vortical manner (Nonaka et al., 1998). When small beads were added to the medium, this vortical ciliary motion swept them to one side of the embryo, where they accumulated. The current hypothesis proposes that if morphogen were added at the node, it also would be moved into a distinct gradient that could be sensed by cellular receptors and thus set up the required axial asymmetry needed for embryonic development (Nonaka et al., 1998). In iv mutant mice this motility fails and axis choice occurs essentially at random with the result that approximately half the progeny have inverted organ placement (see Mercola and Levin (2001) for a comprehensive review of the many issues involved). The multiple roles played by cytoplasmic dyneins, both alone and in combination with the adaptor complex dynactin, remain to be fully elucidated. However, to date there is clear evidence for the involvement of this microtubule motor in vesicular trafficking and maintenance of the Golgi apparatus. Furthermore, during mitosis dyneins localized at the cell cortex act to pull the spindle poles apart during anaphase B, while dyneins located at the kinetochore move cell cycle checkpoint proteins (Wojcik et al., 2001). Recently, this complex has also been found to be necessary for dissolution of the nuclear envelope where it appears to pull or tear the membrane and underlying lamina during prophase and prometaphase (Beaudouin et al., 2002; Salina et al., 2002). Intriguingly, several viruses, including Herpes and rabies, attach to dynein components such as LC8 and co-opt the motor machinery in order to move the invading particles toward the nucleus (Raux et al., 2000; Sodeik et al., 1997). Attempts to obtain homozygous mouse knock-outs lacking the cytoplasmic dynein HC failed to produce any viable offspring suggesting that this motor is absolutely necessary at an early stage in mammalian development (Harada et al., 1998). However, it has been possible to obtain cultured blastocysts lacking the cytoplasmic dynein HC. Analysis of these cells revealed a variety of transport defects including vesiculation of the Golgi apparatus and failure of endosomes and lyzosomes to accumulate in the perinuclear region (Harada et al., 1998). Similarly, analysis of total loss-of-function mutations in Drosophila revealed that the LC8 protein also is essential in multicellular organisms (Dick et al., 1996a). Partial loss-of-function mutants exhibit a pleiotropic array of defects in development and fertility (Dick et al., 1996a; Phillis et al., 1996). In Chlamydomonas, null mutants for LC8 are viable and indeed grow at wild-type rates (Pazour et al., 1998). However, these cells are unable to assemble flagella due to the failure of intraflagellar transport mediated by the 1b class of cytoplasmic dynein (Pazour et al., 1998, 1999a). Importantly, the LC8 pro-


tein is present in many enzyme complexes other than dynein (King et al., 1996a), such as myosin V (Espindola et al., 2000) and neuronal nitric oxide synthase (Jaffrey and Snyder, 1996). Thus care must be taken in ascribing all the phenotypes observed in these various mutants to defects in dynein-mediated events. Recently, an intriguing connection has been uncovered between the dynein motor and the devastating human disease lissencephaly (see Chapter 20 by Vallee and Tai). This condition is caused by reduction in the amount of LIS-1 protein and is characterized by aberrant brain development such that normal neuronal migration fails to occur and the brain surface remains smooth rather than becoming involuted (see Morris (2000) and Vallee et al. (2000) for further discussion). LIS-1 is a WD-repeat protein (similar to the dynein ICs and Gb), that localizes to the kinetochore, and interacts directly with the dynein complex. Intriguingly, when obtained from brain, LIS-1 co-purifies with platelet-activating factor (PAF) acetylhydrolase which deactivates PAF (Hattori et al., 1994), suggesting a requirement for dynein-based motility in the regulation of this lipid signaling molecule. Mice completely lacking the LIS-1 protein die during embryogenesis, whereas heterozygotes live but exhibit the characteristic severe neural phenotype (Hirotsune et al., 1998). Overexpression of LIS1 in mammalian cells causes a significant delay in mitosis, altered spindle orientation and affects dynein localization (Faulkner et al., 2000), whereas mutations in the Drosophila LIS-1 homolog lead to defects in oocyte maturation and neuronal development (Swan et al., 1999). Genetic analysis of the LIS-1 homolog NUDF in Aspergillus also revealed defects in nuclear migration consistent with decreased cytoplasmic dynein function (Xiang et al., 1995). Alterations of dynein cargo interactions can lead to pronounced disease in humans. For example, in photoreceptors rhodopsin must first be transported along microtubules to the base of the connecting cilium prior to being sent through the cilium and assembled into the membrane stacks. The C-terminal domain of rhodopsin is located on the outside of the rhodopsin-bearing vesicles and interacts directly with the Tctex1 LC (Tai et al., 1999). A number of retinitis pigmentosa mutations have been identified that occur within this C-terminal rhodopsin domain. Interestingly, these mutations do not affect the light harvesting ability of the mutant rhodopsin, but rather reduce the affinity of the protein for the Tctex1 LC (Tai et al., 1999). Current hypotheses suggest that the consequent decrease in rhodopsin transport contributes to the slow degradation of the membrane stacks and ultimately results in progressive blindness. Mapping studies have placed the human gene for Tctex1 near the retinal cone dystrophy-1 locus (Watanabe et al., 1996), suggesting that mutations in either the cargo itself or in the cargo-binding component of dynein can lead to retinal degeneration.

2.5

Conclusions

Although great progress has been made in understanding the composition of dynein and its mechanism of action, many important questions remain. For example, there is clearly much that remains to be learnt about the structure of dynein and

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References

how mechanical force is actually generated. Other major areas of current investigation focus on the mechanisms by which dyneins are attached to specific cellular cargoes and on how motor activity itself is regulated so as to result in useful work. With the powerful genetic, biochemical and microscopic techniques that are now being brought to bear on dynein, the future is bright and rapid progress in many of these areas can be anticipated.

Acknowledgements

I thank Drs John Heuser and Michael Koonce for the electron micrographs shown in Figs 2.1b and 2.2b, respectively, and Dr Miho Sakato for Fig. 2.1a. My laboratory is supported by grants GM51293 and GM63548 from the National Institutes of Health and by the Patrick and Catherine Weldon Donaghue Medical Research Foundation. References Afzelius, B. A. 1979. The immotile-cilia syndrome and other ciliary diseases. Int. Rev. Exp. Pathol. 19: 1 43. Aitken, R. J., D. Harkiss, W. Knox, M. Paterson, and D. S. Irvine. 1997. A novel signal transduction cascade in capacitating human spermatozoa characterized by a redox-regulated, cAMP-mediated induction of tyrosine phosphorylation. J. Cell Sci. 111: 645 656. Barbar, E., B. Kleinmann, D. Imhoff, M. Li, T. S. Hays, and M. Hare. 2001. Dimerization and folding of LC8, a highly conserved light chain of cytoplasmic dynein. Biochemistry 40: 1596 1605. Barkalow, K., T. Hamasaki, and P. Satir. 1994. Regulation of 22S dynein by a 29-kD light chain. J. Cell Biol. 126: 727 735. Beaudouin, J., D. Gerlich, N. Daigle, R. Eils, and J. Ellenberg. 2002. Nuclear envelope breakdown proceeds by microtubule-induced tearing of the lamina. Cell 108: 83 96. Beckwith, S. M., C. H. Roghi, B. Liu, and N. Ronald Morris. 1998. The ‘8-kD’ cytoplasmic dynein light chain is required for nuclear migration and for dynein heavy chain localization in Aspergillus nidulans. J. Cell Biol. 143: 1239 1247. Benashski, S. E., A. Harrison, R. S. Patel-King, and S. M. King. 1997. Dimerization of the highly conserved light chain shared by dynein and myosin V. J. Biol. Chem. 272: 0929 20935.

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References Pfister, K. K., R. B. Fay, and G. B. Witman. 1982. Purification and polypeptide composition of dynein ATPases from Chlamydomonas flagella. Cell Motil. 2: 525 547. Pfister, K. K., M. W. Salata, J. F. Dillman, E. Torre, and R. J. Lye. 1996a. Identification and developmental regulation of a neuron-specific subunit of cytoplasmic dynein. Mol. Biol. Cell. 7: 331 343. Pfister, K. K., M. W. Salata, J. F. Dillman, K. T. Vaughan, R. B. Vallee, E. Torre, and R. J. Lye. 1996b. Differential expression and phosphorylation of the 74-kDa intermediate chains of cytoplasmic dynein in cultured neurons and glia. J. Biol. Chem. 271: 1687 1694. Phillis, R., D. Statton, P. Caruccio, and R. K. Murphey. 1996. Mutations in the 8 kDa dynein light chain gene disrupt sensory axon projections in the Drosophila imaginal CNS. Development 122: 2955 2963. Piperno, G. and D. J. Luck. 1981. Inner arm dyneins from flagella of Chlamydomonas reinhardtii. Cell 27: 331 340. Piperno, G., K. Mead, M. LeDizet, and A. Moscatelli. 1994. Mutations in the ‘dynein regulatory complex’ alter the ATP-insensitive binding sites for inner arm dyneins in Chlamydomonas axonemes. J. Cell Biol. 125: 1109 1117. Piperno, G., K. Mead, and W. Shestak. 1992. The inner dynein arms I2 interact with a ‘dynein regulatory complex’ in Chlamydomonas flagella. J. Cell Biol. 118: 1455 1463. Piperno, G., Z. Ramanis, E. F. Smith, and W. S. Sale. 1990. Three distinct inner dynein arms in Chlamydomonas flagella: molecular composition and location in the axoneme. J. Cell Biol. 110: 379 389. Porter, M. E. and K. A. Johnson. 1983. Transient state kinetic analysis of the ATP-induced dissociation of the dynein-microtubule complex. J. Biol. Chem. 258: 6582 6587. Porter, M. E., R. Bower, J. A. Knott, P. Byrd, and W. Dentler. 1999. Cytoplasmic dynein heavy chain 1b is required for flagellar assembly in Chlamydomonas. Mol. Biol. Cell 10: 693 712. Porter, M. E., J. A. Knott, L. C. Gardner, D. R. Mitchell, and S. K. Dutcher. 1994. Mutations in the SUP-PF-1 locus of Chlamydomonas reinhardtii identify a regulatory domain in the b-dynein heavy chain. J. Cell Biol. 126: 1495 1507.

Puthalakath, H., D. C. Huang, L. A. O’Reilly, S. M. King, and A. Strasser. 1999. The proapoptotic activity of the Bcl-2 family member Bim is regulated by interaction with the dynein motor complex. Mol. Cell 3: 287 296. Raux, H., A. Flamand, and D. Blondel. 2000. Interaction of the rabies virus P protein with the LC8 dynein light chain. J. Virol. 74: 10212 10216. Rittinger, K., P. A. Walker, J. F. Eccleston, S. J. Smerdon, and S. J. Gamblin. 1997. Structure at 1.65 Å of RhoA and its GTPase-activating protein in complex with a transition-state analogue. Nature 389: 758 762. Sakakibara, H., H. Kojima, Y. Sakai, E. Katayama, and K. Oiwa. 1999. Inner-arm dynein c of Chlamydomonas flagella is a singleheaded processive motor. Nature 400: 586 590. Sakakibara, H., S. Takada, S. M. King, G. B. Witman, and R. Kamiya. 1993. A Chlamydomonas outer arm dynein mutant with a truncated b heavy chain. J. Cell Biol. 122: 653 661. Sakato, M., and S. M. King. 2001. Ca2 -specific modulation of the microtubule-binding activity of Chlamydomonas outer arm dynein. Mol. Biol. Cell 12: 313a. Sale, W. S. and L. A. Fox. 1988. Isolated b-heavy chain subunit of dynein translocates microtubules in vitro. J. Cell Biol. 107: 1793 1797. Sale, W. S., U. W. Goodenough, and J. E. Heuser. 1985. The substructure of isolated and in situ outer dynein arms of sea urchin sperm flagella. J. Cell Biol. 101: 1400 1412. Salina, D., K. Bodoor, D. M. Eckley, T. A. Schroer, J. B. Rattner, and B. Burke. 2002. Cytoplasmic dynein as a facilitator of nuclear envelope breakdown. Cell 108: 97 107. Samsó, M., M. Radermacher, J. Frank, and M. P. Koonce. 1998. Structural characterization of a dynein motor domain. J. Mol. Biol. 276: 927 937. Scheffzek, K., M. R. Ahmadian, W. Kabsch, L. Weismuller, A. Lautwein, F. Schmitz, and A. Wittinghofer. 1997. The Ras RasGAP complex: structural basis for GTPase activation and its loss in oncogenic Ras mutants. Science 277: 333 338. Schnorrer, F., K. Bohmann, and C. NussleinVolhard. 2000. The molecular motor dynein is involved in targeting swallow and bicoid RNA to the anterior pole of Drosophila oocytes. Nature Cell Biol. 2: 185 190.

2 Dynein Motors: Structure, Mechanochemistry and Regulation Schroer, T. A. and M. P. Sheetz. 1991. Two activators of microtubule-based vesicle transport. J. Cell Biol. 115: 1309 1318. Shimizu, T. and K. A. Johnson. 1983. Presteady state kinetic analysis of vanadate-induced inhibition of the dynein ATPase. J. Biol. Chem. 258: 13833 13840. Shimizu, T., I. Kimura, H. Murofushi, and H. Sakai. 1979. The initial burst of phosphate liberation by 30 S dynein ATPase from Tetrahymena cilia. FEBS Lett. 108: 215 218. Silver, L. M. 1993. The peculiar journey of a selfish chromosome. Trends Genet. 9: 250 254. Smith, E. F. and W. S. Sale. 1992. Regulation of dynein-driven microtubule sliding by the radial spokes in flagella. Science 257: 1557 1559. Smith, T. F., C. G. Gaitatzes, K. Saxena, and E. J. Neer. 1999. The WD-repeat: a common architecture for diverse functions. Trends Biol. Sci. 24: 181 185. Smith, D. S., M. Niethammer, R. Ayala, Y. Zhou, M. J. Gambello, A. Wynshaw-Boris, and L. H. Tsai. 2000. Regulation of cytoplasmic dynein behaviour and microtubule organization by mammalian Lis1. Nature Cell Biol. 2: 767 775. Sodeik, B., M. W. Ebersold, and A. Helenius. 1997. Microtubule-mediated transport of incoming Herpes simplex virus 1 capsids to the nucleus. J. Cell Biol. 136: 1007 1021. Sondek, J., A. Bohm, D. G. Lambright, H. E. Hamm, and P. B. Sigler. 1996. Crystal structure of a G-protein b/g dimer at 2.1 Å resolution. Nature 379: 369 374. Spormann, A. M. and D. Kaiser. 1999. Gliding mutants of Myxococcus xanthus with high reversal frequencies and small displacements. J. Bacteriol. 181: 2593 2601. Supp, D. M., M. Brueckner, M. R. Kuehn, D. P. Witte, L. A. Lowe, J. McGrath, J. Corrales, and S. S. Potter. 1999. Targeted deletion of the ATP binding domain of left right dynein confirms its role in specifying development of left right asymmetries. Development 126: 5495 5504. Supp, D. M., D. P. Witte, S. S. Potter, and M. Brueckner. 1997. Mutation of an axonemal dynein affects left right asymmetry in inversus viscerum mice. Nature 389: 963 966. Swan, A., T. Nguyen, and B. Suter. 1999. Drosophila Lissencephaly-1 functions with Bic-D and dynein in oocyte determination and nu-

clear positioning. Nature Cell Biol. 1: 444 449. Tai, A. W., J. Z. Chuang, C. Bode, U. Wolfrum, and C. H. Sung. 1999. Rhodopsin’s carboxyterminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 97: 877 887. Tai, A. W., J-Z. Chuang, and C-H. Sung. 2001. Cytoplasmic dynein subunit heterogeneity and its role in apical transport. J. Cell Biol. 153: 1499 1510. Taillon, B. E., S. A. Adler, J. P. Suhan, and J. W. Jarvik. 1992. Mutational analysis of centrin: an EF-hand protein associated with three distinct contractile fibers in the basal body apparatus of Chlamydomonas. J. Cell Biol. 119: 1613 1624. Takada, S. and R. Kamiya. 1994. Functional reconstitution of Chlamydomonas outer dynein arms from a- b and g subunits: requirement of a third factor. J. Cell Biol. 126: 737 745. Takahashi, M. and Y. Tonomura. 1979. Kinetic properties of dynein ATPase from Tetrahymena pyriformis. The initial phosphate burst of dynein ATPase and its interaction with ATP analogs. J. Biochem. 86: 413 423. Tang, W. Y. and I. R. Gibbons. 1987. Photosensitized cleavage of dynein heavy chains. Cleavage at the V2 site by irradiation at 365 nm in the presence of oligovanadate. J. Biol. Chem. 262: 17728 17734. Tochio, H., S. Ohki, Q. Zhang, M. Li, and M. Zhang. 1998. Solution structure of a protein inhibitor of neuronal nitric oxide synthase. Nature Struct. Biol. 5: 965 969. Tynan, S. H., M. A. Gee, and R. B. Vallee. 2000a. Distinct but overlapping sites within the cytoplasmic dynein heavy chain for dimerization and for intermediate chain and light intermediate chain binding. J. Biol. Chem. 275: 32769 32774. Tynan, S. H., A. Purohit, S. J. Doxsey, and R. B. Vallee. 2000b. Light intermediate chain 1 defines a functional subfraction of cytoplasmic dynein which binds pericentrin. J. Biol. Chem. 275: 32763 32768. Vale, R. D. 2000. AAA proteins. Lords of the ring. J. Cell Biol. 150: F13 F19. Vale, R. D., D. R. Soll, and I. R. Gibbons. 1989. One-dimensional diffusion of microtubules bound to flagellar dynein. Cell 59: 915 925. Vale, R. D. and Y. Y. Toyoshima. 1988. Rotation and translocation of microtubules in vitro

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References induced by dyneins from Tetrahymena cilia. Cell 52: 459 469. Vallee, R. B., N. E. Faulkner, and C. Y. Tai. 2000. The role of cytoplasmic dynein in the human brain developmental disease lissencephaly. Biochim. Biophys. Acta 1496: 89 98. Vaughan, K. T. and R. B. Vallee. 1995. Cytoplasmic dynein binds dynactin through a direct interaction between the intermediate chains and p150Glued. J. Cell Biol. 131: 1507 1516. Vaughan, P. S., J. D. Leszyk, and K. T. Vaughan. 2001. Cytoplasmic dynein intermediate chain phosphorylation regulates binding to dynactin. J. Biol. Chem. 276: 26171 26179. Wang, J., J. J. Song, M. C. Franklin, S. Kamtekar, Y. J. Im, I. S. Rho, C. S. Seong, C. H. Chung, and S. H. Eom. 2001. Crystal structures of the HS1VU peptidase-ATPase complex reveal and ATP-dependent proteolysis mechanism. Structure 9: 177 184. Wang, Z., S. Khan, and M. P. Sheetz. 1995. Single cytoplasmic dynein molecule movements: characterization and comparison with kinesin. Biophys. J. 69: 2011 2023. Watanabe, T., and M. Flavin. 1976. Nucleotidemetabolizing enzymes in Chlamydomonas flagella. J. Biol. Chem. 251: 182 192. Watanabe, T. K., T. Fujiwara, F. Shimizu, S. Okuno, M. Suzuki, E. Takahashi, Y. Nakamura, and Y. Hirai. 1996. Cloning, expression and mapping of TCTEL1, a putative human homolog of murine Tcte1, to 6q. Cytogenet. Cell Genet. 73: 153 156. Weber, C., V. D. Lee, W. J. Chazin, and B. Huang. 1994. High level expression in Escherichia coli and characterization of the EF-hand calcium-binding protein caltractin. J. Biol. Chem. 269:1 5795 15802. Wilkerson, C. G. and G. B. Witman. 1995. Dynein heavy chains have at least two functional ATP-binding sites. Mol. Biol. Cell. 6: 33a. Wilkerson, C. G., S. M. King, A. Koutoulis, G. J. Pazour, and G. B. Witman. 1995. The 78,000 M(r) intermediate chain of Chlamydomonas outer arm dynein is a WD-repeat protein required for arm assembly. J. Cell Biol. 129: 169 178. Wilson, M. J., M. W. Salata, S. J. Susalka, and K. K. Pfister. 2001. Light chains of mammalian cytoplasmic dynein: identification and characterization of a family of LC8 light chains. Cell Motil. Cytoskelton 49: 229 240.

Wojcik, E., Basto, R., Serr, M., Scaërou, F., Karess, R., and T. S. Hays. 2001. Kinetochore dynein: its dynamics and role in the transport of the Rough deal checkpoint protein. Nature Cell Biol. 3: 1001–1007. Wu, H., M. W. Maciejewski, S. E. Benashski, G. P. Mullen, and S. M. King. 2001. The 1H, 15 N and 13C resonance assignments for the Tctex1 dynein light chain from Chlamydomonas flagella. J. Biomol. NMR 20: 89 90. Wu, H., M. W. Maciejewski, A. Marintchev, S. E. Benashski, G. P. Mullen, and S. M. King. 2000. Solution structure of a dynein motor domain associated light chain. Nature Struct. Biol. 7: 575 579. Xiang, X., A. H. Osmani, S. A. Osmani, M. Xin, and N. R. Morris. 1995. NUDF, a nuclear migration gene in Aspergillus nidulans, is similar to the human LIS-1 gene required for neuronal migration. Mol. Biol. Cell 6: 297 310. Yagi, T. 2000. ADP-dependent microtubule translocation by flagellar inner-arm dyneins. Cell Struct. Funct. 25: 263 267. Yanagisawa, H. A. and R. Kamiya. 2001. Association between actin and light chains in Chlamydomonas flagellar inner-arm dyneins. Biochem. Biophys. Res. Commun. 26: 443 447. Yang, P. and W. S. Sale. 1998. The Mr 140,000 intermediate chain of Chlamydomonas flagellar inner arm dynein is a WD-repeat protein implicated in dynein arm anchoring. Mol. Biol. Cell 9: 3335 3349. Yang, P. and W. S. Sale. 2000. Casein kinase I is anchored on axonemal doublet microtubules and regulates flagellar dynein phosphorylation and activity. J. Biol. Chem. 275: 18905 18912. Yang, P., D. R. Diener, J. L. Rosenbaum, and W. S. Sale. 2001. Localization of calmodulin and dynein light chain LC8 in flagellar radial spokes. J. Cell Biol. 153: 1315 1326. Ye, F., S. Zangenehpour, and A. Chaudhuri. 2000. Light-induced down-regulation of the rat class 1 dynein-associated protein robl/ LC7-like gene in visual cortex. J. Biol. Chem. 275: 27172 27176. Yu, X., S. C. West, R. Jahn, and A. T. Brunger. 1998. Structure of the ATP-dependent oligomerization domain of N-ethylmaleimide sensitive factor complexed with ATP. Nature Struct Biol. 5:803 811.

3 Kinesin Superfamily Proteins Nobutaka Hirokawa and Reiko Takemura

3.1

Introduction

A cell is not simply a bag filled with cytoplasmic fluid surrounded by the plasma membrane in which membranous organelles such as the Golgi apparatus, endoplasmic reticulum and secretory vesicles float and through which newly synthesized proteins diffuse to reach their destination. Instead, cells transport and sort proteins and lipids after their synthesis to their proper destinations at appropriate velocities in membranous organelles and protein complexes using various kinds of motor proteins. The Neuron is composed of a cell body, dendrites and a long axon along the direction of impulse propagation. Because of the lack of protein synthesis machinery in the axon, most of the proteins necessary for the functioning of the axon and synaptic terminal must be transported down the axon after their synthesis into the cell body (Grafstein and Forman, 1980). Most proteins are conveyed in membranous organelles or protein complexes. In this sense intracellular transport in the axon is fundamental to Neuronal morphogenesis and functioning. Epithelial cells also develop a polarized structure, that is, apical and basolateral regions, to which certain proteins are specifically transported and sorted (Simons and Ikonen, 1997, Weims et al., 1997). Vectorial intracellular transport occurs not only in polarized cells such as Neurons and epithelial cells but rather in all types of cells (Hirokawa, 1996). Microtubules serve as rails for this transport and have a polarity with a fastgrowing or plus end and a slow-growing or minus end. They are well organized in cells. In nerve axons, microtubules are arranged longitudinally with the plus end oriented away from the cell body. In proximal dendrites, the polarity of microtubules is mixed, while in the distal end, the polarity is the same as in the axon. In epithelial cells, microtubules are organized with the plus end oriented toward the basement membrane. In most other cells such as fibroblasts, microtubules radiate from the cell center with the plus end oriented toward the periphery.

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Figure 3.1. Quick-freeze deep-etch electron micrographs showing short cross-bridges between membranous organelles and microtubules that are structural candidates for KIFs. Bar 50 nm. Reproduced from N. Hirokawa 1996, Trends Cell Biol. 6: 135 141, with permission.

Electron microscopy (EM) revealed short cross-bridge structures between the organelles and microtubules (Hirokawa, 1982), which are candidates for microtubule-associated motor proteins conveying the membranous organelles along microtubules (Fig. 3.1). Moreover, video-enhanced differential interference contrast microscopy revealed bidirectional transport of various types of membranous organelles (Allen et al., 1982, Schnapp et al., 1985). Based on these findings, kinesin superfamily proteins have been identified as motor proteins transporting cargoes along microtubule rails. ‘Conventional kinesin’, the first member of the kinesin superfamily that was identified, consists of two 120-kDa heavy chains (KHCs) and two 64-kDa light chains (KLCs), at least in animal species (Bloom et al., 1988, Brady, 1985, Cyr et al., 1991, Gauger and Goldstein, 1993, Schnapp et al., 1985, Stenoien and Brady, 1997, Vale et al., 1985, Wedaman et al., 1993). It has a rod-like structure composed of two globular heads (10 nm in diameter), a stalk, and a fan-like end, with a total length of 80 nm. The globular heads are composed of KHCs that bind to microtubules (Hirokawa et al., 1989; Scholey et al., 1989); the KLCs constitute the fan-like end (Hirokawa et al., 1989; Pfister et al., 1989). Complementary DNA (cDNA) encoding KHCs yields a protein of Z 975 amino acids of which

3 Kinesin Superfamily Proteins

Z 350 NH2 -terminal amino acids form the motor domain (which binds to microtubules and has ATPase activity), an a-helical coiled coil-rich stalk domain involved in dimer formation and a tail domain (Yang et al., 1989). A systematic molecular biological search of kinesin superfamily genes coding for proteins containing ATP-binding and microtubule-binding consensus sequences or screening using anti-pan kinesin antibodies led to the discovery of many kinesin superfamily proteins, KIFs (Aizawa et al., 1992, Cole et al., 1992, Hirokawa, 1993, Nakagawa et al., 1997, Tabish et al., 1995, Yang et al., 1997) (Figs 3.2 and 3.3). Combining molecular biological approaches with a BLAST search of proteins in public and private genome databases, a total of 45 KIFs were identified in mouse and human genomes (Miki et al., 2001; Figs 3.4, 3.5 and Tab. 3.1). In this chapter, we provide an overview of KIFs.

Figure 3.2. Structure of cDNAs from murine kinesin superfamily proteins (KIFs). Purple, motor domain; thin red line, ATP-binding consensus sequence; thick red line, microtubulebinding consensus sequence.

81

82

3.1 Introduction Table 3.1 All KIFs from mouse, human, D. melanogaster, C. elegans, S. cerevisiae, and selected other organisms.

H. sapiens

KIF1A

D. melanogaster

ATSV

KIF1B

CG8566

KIF2

CG1453

(-)

K11D9.1

(-)

RnKIF1D

Nakagawa et al., 1997, Nakajima et al., 2002, Rogers et al., 1997

XlXKIF2

Aizawa et al., 1992, FB, WB, Desai et al., 1999 Miki et al., 2001, FB

CAKin/KNSL6

Klp64D Klp68D

KIF3C

CG17461 KIF4

Klp3A

KRP85 KRP95

(-)

CgMCAK XlKCM1

Miki et al., 2001, Kim et al., 1997, Wordeman and Mitchison, 1995, Walczak et al., 1996

SpKRP85 SpKRP95

Aizawa et al., 1992, Yang et al., 1997, Rashid et al., 1995, Cole et al., 1993 Nakagawa et al., 1997, FB

Y43F4B.6

(-)

GgChrkin

KIF4B KIF5A KIF5B KIF5C

Reference

Nangaku et al., 1994, Zhao et al., 2001, FB

CG3219

KIF3A KIF3B

KIF4A

Others

Aizawa et al., 1992, Okada et al., 1995, Furlong et al., 1996, Otsuka et al., 1991

(-)

KIF2B KIF2C

S. cerevisiae

Unc104

KIF1C

KIF2A

C. elegans

Aizawa et al., 1992, Williams et al., 1995, WB, Wang and Adler, 1995 Miki et al., 2001, Ha et al., 2000

nKHC uKHC xKHC

KHC

Unc116

(-)

RnnKHC

Aizawa et al., 1992, Nakagawa et al., 1997, Kanai et al., 2000, Niclas et al., 1994, Yang et al., 1997, Saxton et al., 1991, Patel , et al., 1993

KIF6

(-)

(-)

(-)

Nakagawa et al., 1997

KIF7

(-)

(-)

(-)

Nakagawa et al., 1997

3 Kinesin Superfamily Proteins Table 3.1

(continued). H. sapiens

D. melanogaster

C. elegans

S. cerevisiae

Others

Reference

Eg5/KNSL1

Klp61F

BimC

Kip1 Cin8

AnBimC XlEg5

Nakagawa et al., 1997, Le Guellec et al., 1991, Enos and Morris, 1990, Roof et al., 1992, Hoyt et al., 1992

(-)

(-)

(-)

CrKlp1

Nakagawa et al., 1997, Piddini et al., 2001, Bernstein et al., 1994

Cmet Cana

(-)

Kip2

Nakagawa et al., 1997, Yen et al., 1991, Yucel et al., 2000, FB, Roof et al., 1992

KIF12

CG15844

(-)

(-)

Nakagawa et al., 1997, FB

KIF13A

Kin73

Klp4

(-)

Nakagawa et al., 1997, 2000, Li et al., 1997, Stewart et al., 1991

KIF8 KIF11

KIF9

KIF10

CENP-E

KIF13B

GAKIN

KIF14

HUMORFW

Klp38B

Klp6

(-)

Nakagawa et al., 1997, Molina et al., 1997, WB

KIF15

Hklp2

(-)

C06G3.2 C33H5.4

(-)

Nakagawa et al., 1997, Sueishi et al., 2000, WB

Klp98A

(-)

(-)

Nakagawa et al., 1997, Stewart et al., 1991

KIF16A

Nakagawa et al., 1997, Hanada et al., 2000

KIF16B

XlXklp4

Nakagawa et al., 1997, Vernos et al., 1993

KIF17

(-)

Osm3

(-)

Nakagawa et al., 1997, Setou et al., 2000, Shakir et al., 1993

KIF18A KIF18B

Klp67A

(-)

Kip3

Miki et al., 2001, FB, DeZwaan et al., 1997

KIF19A KIF19B

CG9913

(-)

CG12298

(-)

KIF20A

Rab6Kin

KIF20B

KlpMPP1

Miki et al., 2001, FB (-)

MmKlp174 Miki et al., 2001, Echard et al., 1998, FB Miki et al., 2001, Westendorf et al., 1994

83

84

3.1 Introduction Table 3.1

(continued). H. sapiens

KIF21A KIF21B

D. melanogaster

C. elegans

S. cerevisiae

CG5300

T01G1.1

(-)

Nod

(-)

(-)

XlXkid

Miki et al., 2001, Tokai et al., 1996, Antonio et al., 2000, Funabiki and Murray, 2000

Zen4A Zen4B

(-)

CgCHO1

Miki et al., 2001, Nislow et al., 1990, Adams et al., 1998, Raich et al., 1998, Kuriyama et al., 1994

CG17459

(-)

(-)

Miki et al., 2001, FB

(-)

(-)

(-)

Miki et al., 2001, Okamoto et al., 1998

(-)

Vab8

(-)

Miki et al., 2001, Wolf et al., 1998

Ncd

C41G7.2 M01E11.6 W02B12.7

Kar3

CgCHO2

Nakagawa et al., 1997, Ando et al., 1994, Endow et al., 1990, McDonald and Goldstein, 1990, WB, Wordeman and Mitchison, 1995, Kuriyama et al., 1995

(-)

Klp3

(-)

XlXCTK1

Nakagawa et al., 1997, Saito et al., 1997, Hanlon et al., 1997, Khan et al.,1997, Walczak et al., 1996

KIF22

Kid/KNSL4

KIF23

MKLP1/KNSL5 Pav

KIF24 KIF25

KNSL3

KIF26A KIF26B KIFC1

KIFC2

KIFC3

HSET/KNSL2

Others

Reference

Marszalek et al., 1999b, FB, WB

Nakagawa et al., 1997, Noda et al., 2001, Yang et al., 2001b

FB Flybase http://flybase.bio.indiana.edu:82/; WB, Wormbase, http://www.wormbase.org


Figure 3.3. Left: Panels showing main KIFs functioning in intracellular transport, as observed by low-angle rotary shadowing EM. Scale bar, 100 nm. Right: Schematic illustration of the

Figure 3.4. All mouse and human KIFs and phylogenetic analysis of mouse and human orthologs. Sequences were analyzed by the neighbor-joining method. Reproduced from H. Miki et al. 2001, PNAS 98: 7004 7011, with permission.

same KIFs based on EM studies or predicted from analysis of primary structures. Reproduced from N. Hirokawa 1998, Science 279: 519 526, with permission.

85

86

3.1 Introduction

Figure 3.5. Phylogenetic analysis of all KIFs expressed in mouse/human, D. melanogaster, C. elegans and S. cerevisiae. Amino acid sequences were aligned using maximum parisomy. Reproduced from H. Miki et al. 2001, PNAS 98: 7004 7011, with permission.


3.2

The Kinesin Superfamily Proteins

KIFs have been classified into three major types based on the position of the motor domain: NH2 -terminal motor domain type, middle motor domain type, and COOH-terminal motor domain type (referred to as N-kinesin, M-kinesin, and C-kinesin, respectively). All KIFs known or predicted to be transcribed in the human and mouse genomes are presented in a phylogenic tree along with the KIFs from S. cerevisiae, Drosophila melanogaster, and Caenorhabditis elegans (Fig. 3.5). The entire family is classified into 14 classes. N-kinesin consists of 11 classes, comprising 16 families. Most classes are composed of one family except N-3, N-4, N-6, and N-8. The N-3 class consists of Unc104/KIF1, KIF13, and KIF16 families. Members of the Unc104/KIF1 family are mostly monomeric, while members of the KIF13 family have different characteristics. The N-4 class is composed of the KIF3 and Osm3/KIF17 families. KIF3 is heterotrimeric, and Osm3/KIF17 forms homodimers, indicating that these two families are distinct within this class. The N-8 class consists of the KIF18 and Kid/KIF22 families. M-kinesin is composed of one class, the KIF2 family. C-kinesin is composed of C-1 and C-2 classes, each having one family. Most of the KIFs of other species including plant species can be categorized into these 14 classes.

3.3

N-Kinesins 3.3.1

N-1 Kinesins

The first reported kinesin, the kinesin heavy chain (KHC), is included in this group. There are three highly related family members in this group, KIF5A, KIF5B, and KIF5C, all of which form tetramers consisting of two heavy chains and two light chains (Aizawa et al., 1992, Hirokawa, 1998, Hirokawa et al., 1998, Kanai et al., 2000, Nakagawa et al., 1997, Navone et al., 1992, Nicklas et al., 1994, Yang et al., 1989). KIF5B is ubiquitously expressed in many tissues, while KIF5A and KIF5C are specifically expressed in the nervous system (Kanai et al., 2000, Miki et al., 2001, Niclas et al., 1994). The members of this family transport various organelles and macromolecular complexes within the cell (Hirokawa 1998, Miki et al., 2001, Terada and Hirokawa, 2000), including anterograde transported vesicles (Hirokawa et al., 1991) probably important for action potential propagation (Gho et al., 1992), mitochondria (Tanaka et al., 1998), lysosomes (Hollenbeck and Swanson 1990, Nakata and Hirokawa 1995), endocytic vesicles (Bananis et al., 2000), tubulin oligomers (Terada et al., 2000), intermediate filament proteins such as vimentin (Liao and Gunderson, 1998; Prahlad et al., 1998) and mRNA complexes (Brendza et al., 2000; Carson et al., 1997) (Fig. 3.6a,b). It may also be involved in cytoplasmic viral transport (Rietdorf et al., 2001).

87

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3.3 N-Kinesins

a)

b) Figure 3.6. Scheme of KIFs and their cargo Golgi network; TGN, trans-Golgi network; ECV, organelles in neurons (a) and in cells in general endosomal carrier vesicle. Black arrows indicate (b). In (b), neuron-specific KIFs and ubiquitous the direction of transport. KIFs are illustrated in the same cell. CGN, cis-


The exact molecular interaction involved in the binding of KIF5 to these cargoes has not been elucidated, but recently a direct interaction between KIF5 and some of the cargoes has been shown. KIF5 has been shown to bind directly to a group of scaffolding proteins of the JNK signaling pathway namely, the c-jun NH2 -terminal kinase (JNK)-interacting proteins (JIPs), JIP-1, JIP-2, and JIP-3 (also called JSAP1 and Sunday Driver, SYD) (Bowman et al., 2000, Byrd et al., 2001, Ito et al., 1999, Verhey and Rapoport 2001, Verhey et al., 2001). JIP-1 and JIP-2 interact with tetratricopeptide repeat (TPR) motifs of kinesin light chains through their C-termini. JIP-1 and JIP-2 are soluble proteins by themselves. Therefore, they do not directly connect kinesin to membrane organelles. Rather, they may serve as a link between other cargo membrane proteins and KIF5. One possible mechanism of interaction is that JIP-1 and JIP-2 mediate the interaction of KIF5 with vesicles containing ApoER2, the receptor for the Reelin ligand that controls Neuronal migration (Fig. 3.8). The role of JIP-3 (Sunday Driver, JSAP1) is less clear. JIP-3 interacts with TPR motifs of kinesin light chains through internal sequences, instead of the

Figure 3.7.

Schematic drawing of diverse functions of KIF3 in various cell types.

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C-terminus, and is proposed to be a membrane protein by itself (Bowman et al., 2000), but an unequivocal transmembrane domain has not yet been found. Amyloid precursor protein (APP), whose aberrant transport is thought to contribute to the development of Alzheimer’s disease, is another protein that was proposed to interact directly with KIF5 (Kamal et al., 2000). The interaction between APP and KIF5 is again through the TPR motifs of kinesin light chains (KLCs). It is proposed that APP serves as the membrane cargo receptor for KIF5 which links KIF5 to a particular subset of axon-transported vesicles. Thus far, reports of binding of KIF5 to other molecules seem to be focused on the TPR motifs of kinesin light chains. However, it was not clear whether KLC is the only site for the interaction between kinesin and cargoes. In some species such as fungi (Kirchner et al., 1999, Steinberg and Schliwa, 1995), kinesin light chains are absent, implying that KHCs alone are sufficient by themselves for binding to some cargoes (Seiler et al., 2000). An biochemical interaction between KHC and membranous organelles has been suggested (Skoufias et al., 1994). Very recently, GRIP1 (Glutamate Receptor Interacting Protein 1) has been shown to bind to the KHC tail domain and transport AMPA-type glutamate receptor (GluR2)-containing vesicles to dendrites (Setou et al., 2002) (Fig. 3.8). Furthermore, this study demonstrated

Figure 3.8.

Scheme of how KIFs recognize and bind cargoes.


clearly that GRIP1 steers kinesin to dendrites while JSAP1 (JIP-3, Sunday Driver) steers kinesin to axons through its interaction with KLC (Setou et al., 2002). KIF5B was also shown to interact with the actin-based myosin motor MyoVA, suggesting a coordination between these two motors (Huang et al., 1999). 3.3.2

N-2 Kinesins

This class includes KIF11, BimC, Eg5, and plays a role in mitosis (Blangy et al., 1995, Cole et al., 1994, Enos and Morris, 1990, Kapoor et al., 2000, Kapoor and Mitchison, 2001, Le Guellec et al., 1991, Miki et al., 2001, Sharp et al., 1999a). KIF11 (Eg5) was shown to form a homotetramer for bipolar spindle formation. The motors localize at the midzone of interpolar microtubules, where microtubules have an anti-parallel orientation. Because the tetramers have motor domains on both ends of the molecule, they are thought to crosslink anti-parallel microtubules and to allow them to slide against each other. 3.3.3

N-3 Kinesins

This class consists of three families, the Unc104/KIF1, KIF13, and KIF16 families (Miki et al., 2001). The Unc104/KIF1 family is unique, because most of the proteins in this family form monomeric motors.

The Unc104/KIF1 family There are four members in this family, namely KIF1A, KIF1Ba, KIF1Bb, and KIF1C (Hirokawa, 1998, Nakajima et al., 2002, Nangaku et al., 1994, Okada et al., 1995, Zhao et al., 2001). KIF1A and KIF1B are monomeric, a unique characteristic of these proteins compared to other KIFs. KIF1A was shown to be able to move processively as a monomer by a biased Brownian movement toward the microtubule plus end (Kikkawa et al., 2000, 2001, Okada and Hirokawa, 1999, 2000). C. elegans Unc104 is a homolog of mouse KIF1A (Hall and Hedgecock, 1991, Otsuka et al., 1991). KIF1Ba and KIF1Bb are splice variants which have similar N-terminal motor domains but are distinct in their cargo-binding domains. Interestingly, the cargo-binding domains of KIF1A and KIF1Bb have high homology, and both KIFs transport precursors of synaptic vesicles (Okada et al., 1995, Zhao et al., 2001) (Fig. 3.6a). The knock-out mice of KIF1A and KIF1B show an aberrant Neuronal function due to the defect in the transport of precursors of synaptic vesicles (Yonekawa et al., 1998, Zhao et al., 2001). This gene-targeting experiment demonstrated for the first time that defects in axonal transport due to a mutated motor protein can cause human peripheral neuropathy (Zhao et al., 2001). The KIF1B heterozygotes show progressive muscle weakness and motor disturbance similar to human neuropathies, and it was shown that patients with Charcot Marie Tooth disease type 2A carry a loss-of-function mutation in the motor 3.3.3.1

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domain of the KIF1B gene (Zhao et al., 2001). KIF1Ba transports mitochondria, but this role is shared with KIF5B (Tanaka et al., 1998) (Fig. 3.6a). KIF1C is dimeric, has a high homology to the mitochondrial motor KIF1Ba, and is believed to be involved in the transport of vesicles from the Golgi apparatus to the endoplasmic reticulum (Dorner et al., 1998). KIF1C was shown to be associated with 14-3-3 b, g, e and z proteins (Dorner et al., 1999). However, unexpectedly, cells from KIF1C (-/-) mice showed no significant difference from control cells in the transport of vesicles from the Golgi apparatus to the endoplasmic reticulum, suggesting that KIF1C is dispensable for retrograde transport (Nakajima et al., 2002). Other KIFs may share this function. Recently, KIF1C was reported to mediate mouse macrophage resistance to the anthrax lethal factor (Watters et al., 2001). KIF1C does not affect the cellular entry process of anthrax lethal toxin; therefore, events occurring later in the intoxication pathway are probably involved.

The KIF13 family KIF13A transports vesicles containing the mannose-6-phosphate receptor from the trans-Golgi network to the plasma membrane (Nakagawa et al., 2000) (Fig. 3.6b). The intracellular trafficking of the mannose-6-phosphate receptor has been well studied. One of the roles of the mannose-6-phosphate receptor is to function as a recognition signal for the transport of newly synthesized lysosomal enzymes from the trans-Golgi network; it is also involved in the uptake of external ligands on the cell surface. The interaction of KIF13A with the vesicle is mediated via direct interaction between the KIF13A tail and b-1-adaptin, a subunit of the AP1 adaptor complex (Nakagawa et al., 2000), and the direct interaction of AP-1 with the mannose-6-phosphate receptor has been demonstrated (Fig. 3.8). KIF13B (GAKIN) was reported to interact with the human lympohcyte homolog of the Drosophila disc large tumor suppressor protein (hDlg), and may participate in the translocation of hDlg from the cytoplasm to the ‘cap’ structure upon activation (Hanada et al., 2000). Among the MAGUK superfamily proteins, GAKIN binds to the guanylate kinase-like domain of PSD-95, but not to other MAGUKs. 3.3.3.2

The KIF16 family KIF14, KIF16A, and KIF16B are members of this family. The function of this group of KIFs has not been clarified yet (Miki et al., 2001). Because the tail domains as well as the expression patterns of KIF14, KIF16A, and KIF16B are different, these three KIFs may have separate functions. 3.3.3.3

3.3.4

N-4 Kinesins

This class is composed of the KIF3 family, which form heterotrimeric motors, and the Osm3/KIF17 family (Miki et al., 2001).


The KIF3 family KIF3A, KIF3B, and KIF3C constitute this family. Members of the KIF3 family form heterotrimeric complexes. Thus the KIF3A KIF3B heterodimer interacts at its tail domain with a non-motor protein, kinesin superfamily-associated protein 3 (KAP3) (Hirokawa 1998, Kondo et al., 1994, Pesavento et al., 1994, Rashid et al., 1995, Wedaman et al., 1996, Yamazaki et al., 1995, 1996) (Fig. 3.3). The KIF3A/B-KAP3 complex (also called kinesin II) transports vesicles and macromolecules (Hirokawa, 2000). In axons of mammalian Neurons, the complex transports vesicles which carry fodrin (Takeda et al., 2000), and choline acetyltransferase, a soluble protein in Drosophila (Ray et al., 1999). In melanophores, it is involved in the dispersion of pigment organelles called melanosomes (Gross et al., 2002, Reese and Haimo, 2000, Tuma et al., 1998) (Fig. 3.7). The KIF3 complex plays a vital role in the formation and maintenance of cilia and flagella in various organisms (Cole et al., 1998, Kozminski et al., 1995, Morris and Scholey, 1997, Piperno and Mead, 1997, Piperno et al., 1996, Walther et al., 1994). It transports macromolecular protein complexes from the bottom of the cilium to its distal tip along microtubules by a process called intraflagellar transport (Fig. 3.7). The components of these macromolecular complexes, called rafts, have been analyzed to some extent; rafts may contain as many as 15 polypeptides, but the details have not been completely clarified (Cole et al., 1998). An interesting aspect of the involvement of KIF3 in cilium formation is that it also plays an important role in the determination of the left right axis in early mouse development (Marszalek et al., 1999a, Nonaka et al., 1998, Takeda et al., 1999). During mouse development, there is a monocilium at the nodal cell that rotates to produce unidirectional leftward flow of extraembryonic fluid (Nonaka et al., 1998; Takeda et al., 1999). This nodal flow could generate concentration gradients of putative secreted morphogens in the extraembryonic fluid at the nodal region, which could switch on a gene cascade strictly expressed on the left side of the body and lead to left right asymmetry. KIF3 is required for the formation of this monocilium. In kif3A (–/–) or kif3B (–/–) mouse, nodal cilia cannot be generated, nodal flow is absent, and the left right axis formation is fundamentally impaired (Nonaka et al., 1998; Takeda et al., 1999). In retinal photoreceptor cells, the KIF3 complex is localized in the connecting cilium, which is located between the outer and inner segment (Whitehead et al., 1999). It was shown that the KIF3 complex participates in the transport of opsin from the inner to the outer segment at the connecting cilium, where microtubule plus-ends are pointed toward the outer segment (Marszalek et al., 2000). At the connecting cilium, opsin is localized at the plasma membrane, and the KIF3 motors probably drag the transmembrane protein along the underlying microtubules. The soluble protein, arrestin, also seems to be transported by the KIF3 complex. In migrating epithelial cells, KAP3 is reported to bind to the tumor suppressor gene, adenomatous polyposis coli (APC), and it is proposed that b-catenin is transported to the edges of migrating epithelial cells through this interaction (Jimbo et al., 2002). 3.3.4.1

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There is another form of the KIF3 complex, KIF3A/C KAP3. KIF3A/C KAP3 is expressed in the nervous system only (Muresan et al., 1998; Yang and Goldstein, 1998), while KIF3A/B KAP3 is ubiquitously expressed in many tissues. However, a gene-targeting experiment showed that the KIF3A/C-KAP3 complex is dispensable; therefore, the function of this KIF3 complex may be redundant (Yang et al., 2001a).

The Osm3/KIF17 family KIF17 is localized to the dendrites and transports vesicles containing N-methyl-Daspartate (NMDA)-type glutamate receptor 2B towards the microtubule plus-end to the postsynaptic site, where the receptor plays an important role in synaptic plasticity (Setou et al., 2000) (Fig. 3.2). In the dendrite, microtubules are of mixed polarity. The large protein complex containing mLin-10 (Mint1), mLin-2 (CASK), mLin-7 (MALS/Velis), and NR2B mediates the attachment of KIF17 to the vesicle. The tail domain of KIF17 interacts directly with the PDZ domain of mLin-10, which then sequentially interacts with mLin-2 (CASK), mLin-7, and the NR2B subunit in this order (Setou et al., 2000) (Figs 3.6a and 3.8). Osm-3 is closely related to KIF3, but forms a dimer (Shakir et al., 1993). In C. elegans, many Neurons have ciliated dendritic ends, and some of them are chemosensory receptors. Osm-3 transports ciliary components to these ciliated dendritic ends (Signor et al., 1999, Tabish et al., 1995). 3.3.4.2

3.3.5

N-5 Kinesins

This class includes KIF4A, KIF4B, KIF21A, and KIF21B, and plays roles in both vesicle transport and mitosis (Miki et al., 2001; Williams et al., 1995). KIF4A mRNA is abundantly expressed in juvenile Neurons, including differentiated immature Neurons, and the motor transports vesicles to growth cones (Sekine et al., 1994). These vesicles may contain L1, a cell adhesion molecule implicated in axonal elongation in immature Neurons (Peretti et al., 2000). KIF4 was shown to bind murine leukemia virus Gag proteins and may play a role in the transport of viruses in an infected cell (Kim et al., 1998). Chromokinesin, a chicken isolog of KIF4, is associated with chromosome arms and functions as a mitotic motor with DNA as its cargo (Wang and Adler, 1995). What KIF21A and KIF21B transport is not known, but they have different distribution patterns within a Neuron; KIF21A is localized throughout Neurons, but KIF21B is highly expressed in dendrites (Marszalek et al., 1999b). 3.3.6

N-6 Kinesins

CHO1/KIF23 and KIF20/Rab kinesin families constitute this class.


The CHO1/KIF23 family The KIF23 family, called by various names such as CHO1, MKLP, ZEN-4 or Pavarotti, depending on the species, appears to be involved in cytokinesis (Adams et al., 1998, Kuriyama et al., 1994, 2002, Nislow et al., 1990; Raich et al., 1998; Severson et al., 2000). Members of this family can bind to microtubules at their cargo-binding domain, and are, therefore, capable of bundling anti-parallel microtubules. During mitosis, they exist in the midbody region of the spindle in late anaphase and telophase, and probably establish the structure of the telophase spindle to provide a framework for the assembly of the contractile ring. They are also expressed in cultured Neurons (Ferhat et al., 1998, Sharp et al., 1997). 3.3.6.1

The KIF20/Rab6 kinesin family The Rab family of GTP-binding proteins regulates vesicular transport and membrane traffic. KIF20A (Rab6-KIF) was first reported as a motor protein that associates with Rab6 and participates in Golgi-derived vesicle transport (Echard et al., 1998). Moreover, it was revealed that KIF20 may also participate in cytokinesis (Fontijn et al., 2001, Hill et al., 2000). 3.3.6.2

3.3.7

N-7 Kinesins

KIF10 (CENP-E) plays an important role in mitosis (Cottingham and Hoyt, 1997, Roof et al., 1992, Yen et al., 1991, 1992). It associates with kinetochores and tethers the centromere to spindle microtubules. It is presumed to function in ‘congression’, in which chromosomes attached to the spindle microtubules oscillate and migrate to the spindle equator. Therefore, it plays an essential role in chromosome alignment at the spindle equator by tethering kinetochores to the dynamic microtubule plus end (McEwen et al., 2001, Schaar et al., 1997, Topper et al., 2001, Wood et al., 1997, Yao et al., 1997). CENP-meta has a similar function in Drosophila (Yucel et al., 2000). 3.3.8

N-8 Kinesins

This class includes the Kid/KIF22 and KIF18 families.

The Kid/KIF22 family KIF22 (Kid) is involved in mitosis (Tokai et al., 1996). It binds to the arms of chromosomes, not to the kinetochore, and plays a role in chromosome alignment at the equatorial plate during metaphase (Antonio et al., 2000, Funabiki and Murray, 2000, Levesque and Compton, 2001). It is proposed that Kid is responsible for the polar ejection force acting on chromosome arms and its degeneration is required for chromosome movement during anaphase. A highly homologous KIF, 3.3.8.1

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Nod, is required during female meiosis for chromosome alignment (Heald, 2000, Zhang et al., 1990). KIF19 is included in this family, but its function is not yet well characterized (Miki et al., 2001).

The KIF18 family This most recently discovered family of KIFs has not yet been characterized. 3.3.8.2

3.3.9

N-9 Kinesins

KIF12, a member of this family, is highly expressed in the kidney and may have a significant role in kidney cells. However, its function is not yet well characterized (Miki et al., 2001). 3.3.10

N-10 Kinesins

The KIF15 family forms this class that is predominantly expressed in the spleen and testis (Miki et al., 2001). Human KIF15 (Hklp2) may interact with the forkhead-associated domain of pKi-67, a widely used cell proliferation marker protein (Sueishi et al., 2000). 3.3.11

N-11 Kinesins

This class includes the KIF26 family that is related to Cos2 and Vab-8. Vab-8 has been implicated in axon outgrowth and cell migration (Wolf et al., 1998). Costal2 (Cos2) has been shown to be part of the hedgehog signaling cascade; therefore, it is involved in the determination of cell fate and the patterning of animal development (Ascano et al., 2002, Robbins et al., 1997, Sisson et al., 1997; Stegman et al., 2000; Wang et al., 2000). It forms a large multiprotein complex with Ser/Thr protein kinases, Fused (Fu) and the transcription factor Cubitus interruptus (Ci). Cos2 prevents Ci nuclear translocation by tethering it in the cytoplasm via binding to microtubules, whereas hedgehog stimulates Ci nuclear import.

3.4

M-Kinesins

M-kinesins have a motor domain at the center of the molecule. The KIF2 family belongs to this class; its members, KIF2A, 2B, and 2C, have functions in vesicle transport, mitosis, and regulation of microtubule dynamics (Kim et al., 1997, Miki et al., 2001, Noda et al., 1995; Figs 3.2 and 3.3). KIF2A plays a significant role in neurite outgrowth and transports vesicles containing a variant of the b-sub-


unit of the IGF-1 receptor, which is found on growth cones and is implicated in nerve cell development (Morfini et al., 1997; Fig. 3.6a). It may also participate in localization of lysosomes (Santama et al., 1998). The KIF2C homolog in Xenopus, XKCM1, has a microtubule-destabilizing activity. It influences microtubule dynamics within the cell and is thought to promote chromosome segregation during mitosis by depolymerizing microtubules at the kinetochore (Desai et al., 1999, Tournebize et al., 2000, Walczak et al., 1996). XKCM1 acts on a single protofilament, possibly via an interaction of the basic residues in XKCM1 with the acidic tail of tubulin (Niederstrasse et al., 2002). Mitotic centromere-associated kinesin (MCAK) identified in hamster is also a homolog of KIF2C (Wordeman and Mitchison, 1995). It is also a microtubule depolymerizing factor, consistent with its role in promoting chromosome segregation during mitosis (Maney et al., 2001).

3.5

C-Kinesins 3.5.1

C-1 Kinesins

Drosophila Ncd, S. cerevisiae Kar3, and KIFC1 are included in this family (Endow et al., 1990, Meluh and Rose, 1990). They are microtubule minus-end-directed motors and participate in meiosis, mitosis and karyogamy (Miki et al., 2001). They have a microtubule-binding site on the cargo-binding domain. During mitosis, Ncd may function by crosslinking and sliding anti-parallel spindle microtubules relative to one another, pulling them apart, and thereby opposing the effect of bipolar plus-end-directed microtubule motors (Sharp et al., 1999b). In meiosis, Ncd may play a role in establishing bipolar acentrosomal meiotic spindles (Cullen and Ohkura, 2001). Kar3 has an essential role in nuclear fusion during mating or karyogamy in S. cerevisiae. In addition, it has been implicated in several microtubule functions during the vegetative cell cycle including spindle assembly, mitotic chromosome segregation, microtubule depolymerization, kinetochore-motor activity and spindle positioning. These multiple functions may be performed by interaction of Kar3 with different associating proteins such as Cik1p or Vik1p (Barrett et al., 2000, Manning et al., 1999, Shanks et al., 2001, Troxell et al., 2001). 3.5.2

C-2 Kinesins

The KIFC2/C3 families constitute C-2 kinesins (Hanlon et al., 1997, Miki et al., 2001, Saito et al., 1997). KIFC2 is localized in the cell body and dendrites of Neurons (Saito et al., 1997). It participates in the transport in dendrites of multivesicular body-like membranous organelles, which are morphologically similar to multivesicular bodies but appear to lack markers of endocytic compartments (Fig. 3.6a). The classic multivesicular

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3.7 Cargoes of KIFs; Specificity and Redundancy

body exists in both axons and dendrites; therefore, the organelle transported by KIFC2 may constitute a novel class. It has not been rigorously demonstrated, but it is presumed that KIFC2 acts as a minus-end-directed motor. KIFC3 is a minus-end-directed microtubule motor protein that exists in kidney epithelial cells and other cell types. In epithelial cells, microtubule organization is different from that in other cell types; microtubule minus-ends are located at the apical surface of the cell. KIFC3 accumulates on the apical surface of epithelial cells and transports vesicles associated with annexin XIIIb, a previously characterized membrane protein for apically transported vesicles (Noda et al., 2001). A recent gene targeting study showed that KIFC3 plays a complementary role to cytoplasmic dynein in Golgi positioning and integration (Xu et al., 2002).

3.6

Orphans

These KIFs have no counterpart in Drosophila or C. elegans. KIF6 and KIF9 are located near the BimC family (Miki et al., 2001). KIF9 is reported to interact with the Ras-like GTPase Gem, which suggests that it may play a role in cell shape remodeling (Bernstein et al., 1994, Piddini et al., 2001). The distance between KIF9 and KIF6 is large. KIF7 has no evident homolog in Drosophila, C. elegans, or S. cerevisiae. Its function is as yet unknown, but it is predominantly expressed in the testis (Miki et al., 2001).

3.7

Cargoes of KIFs; Specificity and Redundancy

As mentioned above, KIFs convey various types of cargo. In some cases, a single KIF transports several distinct cargoes. For example, conventional kinesin (KIF5s) transports several cargoes such as mitochondria, lysosomes, vesicles containing Amyloid Precursor Protein (APP) and GAP43 (Kamal et al., 2000, 2001), ApoE2, the receptor for the Reelin ligand (Verhey et al., 2001), or the AMPA-type glutamate receptor GluR2 (Setou et al., 2002). Kinesin can also convey mRNA complexes (Brendza et al., 2000), tubulin oligomers (Terada et al., 2000) and vimentin intermediate filament protein complexes (Liao and Gunderson 1998, Prahlad et al., 1998). Another example is KIF3. KIF3 also conveys several different cargoes in distinct cell types. KIF3 transports vesicles associated with fodrin, important for neurite elongation in Neurons (Takeda et al., 2000), and it conveys protein complexes for the formation of cilia and flagella along microtubules in various ciliated cell types (Kozminski et al., 1995, Morris and Scholey, 1997). KIF3 is also a motor for the transport of pigment granules in pigmented cells in Xenopus (Gross et al., 2002, Reese and Haimo, 2000, Tuma et al., 1998) and is proposed to function in the trafficking of vesicles from the Golgi apparatus to the ER (LeBot et al., 1998). Thus,


KIF3 is another example of a motor that carries out different tasks depending on the type of cell (Fig. 3.7). How the same KIF can transport different cargoes is an interesting and important question that needs to be addressed. On the other hand, different KIFs sometimes redundantly convey similar cargoes. Mitochondria are transported by KIF1Ba (Nangaku et al., 1994) as well as by conventional kinesin (KIF5) (Kanai et al., 2000, Tanaka et al., 1998). KIF1A’s cargo is synaptic vesicle precursors (Okada et al., 1995), which are also conveyed by KIF1Bb (Zhao et al., 2001). Another example is lysosomes which have at least two microtubule plus-end motors, kinesin and KIF2Ab (Hollenbeck and Swanson, 1990, Nakata and Hirokawa, 1995, Santama et al., 1998). This redundancy could be one of the reasons for subtle phenotypes in knock-out mice in which some KIF genes have been knocked out.

3.8

Recognition and Binding to Cargoes

In many cases, a KIF has a specific cargo, while sometimes the same KIF, such as conventional kinesin and KIF3, conveys different cargoes. Thus, the fundamental question has been, how do KIFs recognize and bind to their cargoes? Studies of KIF17 and KIF13A clearly indicated for the first time how KIFs recognize and bind cargo molecules (Nakagawa et al., 2000, Setou et al., 2000). KIF17 transports NMDA-type glutamate receptor-containing vesicles through an interaction of its tail with mLin10-mLin2-mLin7-NR2B. KIF13A, on the other hand, recognizes and binds to mannose-6-phosphate receptor-containing vesicles via an interaction between the KIF13A tail-b1 adaptin-AP-1 adaptor complex and mannose-6-phosphate receptor (Fig. 3.8). In the case of conventional kinesin, it was demonstrated recently that it recognizes and binds cargoes such as APP- and GAP43-containing vesicles through KLC and APP interaction (Kamal et al., 2001), and ApoE2-containing vesicles through interaction between KLC, and JIP1 and JIP2 (Verhey et al., 2001). Kinesin also recognizes cargoes such as AMPA receptor subunit GluR2-containing vesicles through its interaction with KHC-GRIP1-GluR2 (Setou et al., 2002) (Fig. 3.8). Thus, kinesin uses KHC and KLC to discriminate between distinct cargoes. A similar strategy may be used by KIF3. Thus in terms of how KIFs recognize and bind cargoes, two methods can be distinguished. x

x

KIF tail scaffolding protein(s) or adaptor protein(s) membrane protein. This is applicable to KIF17-mLin10-mLin2-mLin7-NR2B, KIF13A-b1adaptin-AP-1adaptor complex-M6PR, KLC-JIP1/JIP2-ApoE2, the reelin receptor and KIF5 (KHC)GRIP1-GluR2 (Fig. 3.8). KIF tail membrane protein. Kinesin was reported to bind directly to membrane proteins through its interaction with KLC. Sunday Driver (also called JSAP1 and JIP3) (Bowman et al., 2000) and APP (Kamal et al., 2001) were proposed to be membrane proteins that directly bind to kinesin through KLC, but it needs to

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References

be demonstrated whether Sunday Driver (JSAP1 ad JIP3) is indeed a membrane protein. We are just beginning to understand how KIFs recognize and bind their specific cargoes. So far we only know that rather complex mechanisms are involved in these events. This may be related to how the association and dissociation between KIFs and cargoes are regulated. Another question is how KIFs recognize and bind to protein complexes such as tubulin and intermediate filament proteins, chromosomes and mRNAs. This is obviously a significant question that needs to be answered in the near future.

3.9

How to Determine the Direction of Transport

Another intriguing question is how a cell regulates the direction of transport, for example the transport to axons versus dendrites. Kinesin (KIF5) transports APPcontaining vesicles to the axon while the same motor conveys AMPA-receptor-containing vesicles to dendrites. In this case it was shown that GRIP1, which binds KHC and JSAP1, which binds KLC, steers kinesin to dendrites and axons, respectively (Setou et al., 2002). Thus, GRIP1 and JSAP1 are functioning as drivers for kinesin (KIF5). There are motors that transport cargoes mainly to dendrites. KIF17 conveys NMDA receptors mainly to dendrites (Setou et al., 2000) and KIFC2 transports a new type of multivesicular body-like organelle to dendrites (Saito et al., 1997). KIF21B was also proposed to be a motor for dendritic transport (Marszalek et al., 1999b). How differential transport is performed is clearly a fundamental question that needs to be solved in the near future. The mechanism is probably not simple and could involve several distinct events. Thus, interestingly, cells use many KIFs and very precisely control the direction and velocity of transport of various distinct cargoes that are fundamental in basic cellular functions. This field is obviously very important and rapidly advancing. Further studies need to be carried out to fully understand the mechanism of intracellular transport.

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4 The Bacterial Flagellar Motor Richard Berry

4.1

Introduction

Many species of bacteria actively navigate their environment, swimming to places where conditions are better (Armitage, 1999, Blair, 1995, Macnab, 1996,). They may be seeking out nutrients, light, oxygen, sites for invading a host or avoiding repellents such as toxic chemicals or damaging radiation. Many different swimming styles exist (Fig. 4.1), but most are based on the rotation of bacterial flagella. Exceptions include gliding bacteria such as myxococcus (Zusman et al., 1990) that move slowly over surfaces, and species of marine synechococcus that swim mysteriously without flagella at similar speeds to flagellated bacteria (Pitta et al., 1997, Samuel et al., 2001). In all cases flagella are passive, essentially rigid helices, propelled by a rotary motor at their base (Berry and Armitage, 1999, Blair, 1995, Macnab, 1996). The bacterial flagellar motor is powered by the flux of ions across the inner, or cytoplasmic, membrane of a bacterial cell envelope. Ion flux is driven by an electrochemical gradient, the proton motive force (pmf) or sodium-motive force (smf) in motors driven by H and Na respectively. (The electrochemical gradient consists of a voltage component and a concentration component, and is a key intermediate in the metabolism of both bacteria and higher organisms. The inside of a bacterial cell is typically at an electrical potential about 150 mV below the exterior and also typically contains a slightly lower concentration of H or Na ions.) Filaments rotate at speeds up to and beyond 1000 Hz (cycles per second) in swimming cells and each motor has a maximum power output on the order of 10 15 W, two or three orders of magnitude higher than other known ATP-driven molecular motors. If cells are attached to a surface by a single flagellar filament, or ‘tethered’, the motor turns the whole cell body at speeds around 10 Hz. The rotating heart of the motor is a set of rings in the cytoplasmic membrane, about 45 nm in diameter, and containing a total of a few hundred molecules of several different proteins. This rotor is surrounded by a ring of 8 to 16 independent torque generators, which are anchored to the cell wall and couple the flux of ions to rotation of the rotor.

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4.1 Introduction

Escherichia coli

Rhodobacter sphaeroides

Vibrio alginolyticus

Spirochaeta aurantia

Spirillum volutans Figure 4.1. Some swimming patterns of fla-

gellated bacteria. Different species of bacteria use rotating flagella to swim in different ways. In E. coli, several flagella form a bundle that propels the cell. R. sphaeroides swims with a single lateral flagellum, V. Alginolyticus with a

single polar flagellum contained in a membrane sheath. Spiral bacteria swim like corkscrews, using either periplasmic flagella contained within the outer membrane (Spirochaeta aurantia) or tufts of polar flagella (Spirillum volutans).

4 The Bacterial Flagellar Motor

Flagella of swimming bacteria were first recorded in the 1940s, using the South African sun as a light source for high-intensity dark-field microscopy (Pijper, 1948). Pijper’s films show bundles of flagella as a wave trailing behind swimming cells and suggest as the mechanism of propulsion, a propagating wave generated either by bending (as in eukaryotic flagella) or by rotation of a rigid helix. Early evidence for rotation is summarized by Berg and Anderson (1973), and conclusive proof was provided by Silverman and Simon (1974). Polystyrene beads attached to flagella moved from one side of the filament to the other as if rotating about an axis rather than staying on one side as would have happened if the filament were bending. Also, cells tethered by flagella to glass coverslips rotated about a fixed point when observed in a microscope. Cells rotated even with hook or filament mutations that did not allow cells to swim, ruling out the possibility that they were swimming around an inert point of attachment to the slide. Swimming is not enough to allow bacteria to navigate; they also need to sense which way to go and how to get there. The basic strategy appears to be common to many different species. Flagellar motors are able to switch spontaneously, for example either reversing direction or switching between ‘run’ and ‘stop’ states. Switches lead to a change in swimming direction and occur stochastically at intervals of several seconds. Navigation is accomplished by modulation of the intervals between changes of direction according to temporal changes in attractant or repellent levels. For example, Escherichia coli is propelled by a co-ordinated bundle of several counterclockwise-rotating flagella, with motors located at random on the cell surface (Macnab, 1996). When one or more of the motors switches to rotation in the opposite direction it is expelled from the bundle and the new pattern of forces on the filament causes it transiently to assume a kinked shape, changing the swimming direction in the process. These events, called tumbles, have been studied for decades, although the details of how motor switches lead to changes in swimming direction have only recently been revealed (Turner et al., 2000). The cell compares conditions sensed within the last second to those over the preceding several seconds, and suppresses tumbles when conditions are improving (Segall et al., 1986). This strategy is tailored to the physical constraints imposed by the small size of the cell, which loses track of its direction over several seconds due to Brownian motion and is too small to tell at which of its ends the concentration of a chemical attractant is higher (Berg and Purcell, 1977). In other species the details are slightly different. Rhodobacter sphaeroides has a single flagellum which coils up when the motor stops, re-orienting the cell in the process, while other species simply swim in the opposite direction upon motor reversal. The basic strategy of increasing the likelihood of a change of direction when conditions deteriorate and/or reducing it when conditions improve, however, appears to be the same. Much work has been done on the protein signaling pathway in E. coli that communicates changes in sensory receptor complexes to the flagellar motor. All of the components are identified, and the details of their interactions are perhaps better understood than any other protein network. Much is still to be learned, both in E. coli and in other species with related but more complex pathways, but this will not

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4.2 Structure

be covered here. More details may be found in several reviews on the topic (Armitage, 1999, Blair, 1995, Stock and Surette, 1996). This chapter is concerned with the flagellar motor itself; its structure, measurements of its rotation, and models of how it works. Although experiments on tethered bacteria in the 1970s were the first observations of single molecular motors, considerably more is now known about the detailed mechanisms of the ATP-driven motors actin/myosin, kinesin/microtubules and F1-ATPase. Because the flagellar motor is a large complex, present in the cell in low numbers, and assembled spanning the entire cell envelope, it has not been possible to re-constitute the motor in vitro. Because of the size of the motor and its location in the membrane, X-crystallography has to date yielded the structure of only the C-terminal domain of one rotor protein, FliG. In addition, with the pmf rather than ATP hydrolysis as the driving force for rotation, experiments to measure in solution rate constants of chemical steps in the motor cycle are not possible. Similar difficulties exist for nature’s other ion-driven motor, the membrane-spanning FO motor of ATP-synthase, where rotation has been inferred but never observed directly. Despite these constraints, much has been learned about this fascinating biological electrical rotary motor using a range of electron microscopy, genetic and biophysical techniques. This chapter summarizes what we know.

4.2

Structure

The bacterial flagellum is one of the most complex structures found in bacteria, with 40 50 genes involved in its expression, assembly and control. The details of flagellar genetics and assembly are reviewed by Macnab (1996). It includes cytoplasmic membrane proteins and proteins that anchor it to the peptidoglycan cell wall, as well as structural proteins that assemble into rings, a drive shaft and a bushing that carries the shaft through the outer membrane. There is also an export apparatus with homology to type III secretion systems (Aizawa, 2001) that is required to form the extracellular parts of the flagellum, but this is not believed to have any role in the rotary mechanism of the motor. Not counting the hook and filament, which are part of the propeller rather than the motor itself, nor the LP-ring and the rod, which are the drive-shaft, the flagellar motor contains on the order of 250 protein molecules of six different types (Thomas et al., 1999). It is about 50 nm in diameter and spans the entire cell envelope from the cytoplasm, through the cytoplasmic membrane, periplasm, cell wall and outer membrane (Fig. 4.2). The molecular weight of the C-ring alone is about 5 MDa (Francis et al., 1999). The flagellar motor has been something of a test object for single-particle techniques in electron microscopy, and its structure is known to a resolution approaching 20 Å. Most of the recent electron microscopy has been carried out on the enteric bacterium Salmonella typhimurium, but indications are that the structure of the motors of other species are essentially the same.


Figures 4.3a and b show side-on and end-on views of the rotor obtained by transmission cryo-electron microscopy (cryo-EM) (Francis et al., 1994, Thomas et al., 1999), and Fig. 4.3c shows a scanning electron microscope image of rotors visualized by fast-freeze and thin-film metal replica techniques (Khan et al., 1998). Figure 4.3d shows the stator, a ring of particles left in the membrane after freeze-fracture (Khan et al., 1988). The structure of the entire motor inferred from these images is shown in Fig. 4.2, with the locations indicated of various flagellar structural genes that have been identified by immuno-gold labelling and mutational studies. The rotor, drive shaft (rod) and propeller (hook, filament) are shown in white, the stator is shaded.

Hook

Filament

L-ring P-ring Outer membrane Cell wall

Cytoplasmic membrane N

Rod

C

MS-ring C-ring

Protein

Size

Number of copies

MotB

34 kD

8x2

MotA

32 kD

8x4

FliF

61 kD

26

FliG FliM FliN

37 kD 38 kD 15 kD

26/34 34 68/136

45 nm

A schematic diagram of the Hdriven flagellar motor showing the various components and the proteins associated with the torque-generating apparatus. The area shown in white rotates, while the stator, shaded gray, is attached to the cell wall. Interactions between certain charged residues in the cyto-

Figure 4.2.

plasmic domain of MotA and the C-terminal domain of FliG are crucial for torque generation. The aspartic acid residue asp32 in the membrane-spanning domain of MotB is crucial for ion flux. The structure and mechanism of Na motors are thought to be very similar to those of H motors.

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4.2 Structure A)

C)

B)

D)

Figure 4.3. (a) Side-on projection of a 3-D model of the rotor obtained by alignment and averaging of cryo-electron micrographs of single rotors. (b) End-on view of a single rotor, an image of the type used in (a). (c) Scanning EM of thin-film metal replicas of several rotors. (d) Freeze-fracture EM of a ring of stator particles in the cytoplasmic membrane. The dark scale bar is 45 nm and applies to Figs 4.2a, b and d.

The white scale bar in Fig. 2c is 50 nm. (Figure 4.3a reprinted from Francis et al. (1994) with permission from Elsevier Science Publishers; Fig. 4.3b reprinted with permission from Thomas et al. (1999). c 1999 National Academy of Sciences, USA; Fig. 4.3c reprinted from Khan et al. (1998). c Elsevier Science, USA; Fig. 4.3d reprinted from Khan et al. (1988) with permission from Elsevier Science Publishers).


4.2.1

Propeller and Drive-shaft

The filament is the propeller, joined to the motor by a universal joint called the hook and a drive shaft called the rod. The L- and P-rings are a bushing that carries the drive shaft through the cell wall and outer membrane. It is not certain whether the L- and P-rings rotate with the rotor or are fixed to the cell envelope. The filament is typically 5 10 mm long, 20 nm wide, and composed of over 10,000 copies of a single protein, flagellin, with a molecular weight between 25 60 kDa. Some species may have more than one flagellin, but the proteins are usually closely related. Filaments grow from the distal end, with monomers transported through a central channel running the entire length of the filament before being incorporated by a capping complex (Yonekura et al., 2001). The filament can undergo transformation between different helical forms, induced torsionally or by changes in pH (Kamiya and Asakura, 1982, Turner et al., 2000). This has been explained in terms of a model in which the filament is composed of 11 helical protofilaments, each of which can exist in either long or short forms (Calladine, 1982). The model predicts 12 helical conformations, with any number from 0 to 11 of short protofilaments, all adjacent, and the rest of the protofilaments in the long form. The model has been confirmed by light microscopy, cryo-EM and X-ray crystallography (Samatey et al., 2001). Namba and Vonderviszt (1997) have published a detailed review of the structure of the filament and other flagellar components. 4.2.2

Rotor

Torque is believed to be generated by interactions between the stator particles in the cytoplasmic membrane and the protein FliG, located at the junction of the MS- and C-rings. Figure 4.3a shows a side-on projection of a 3-D model of the rotor, derived by re-orienting and averaging 123 separate cryo-EM images of individual rotors (Francis et al., 1994). All details of the periodicity around the rotation axis are lost in this method, as there was insufficient resolution in the individual images for azimuthal alignment. One notable feature is the extremely low density of the link between the MS- and C-rings. Indeed the C-ring was not seen in early images of the rotor, presumably because this link was broken during the relatively harsh treatment used to isolate rotors. Figure 4.3b shows an end on view of the C-ring in a single rotor. Three such images were seen in the original experiments, two of which had 34-fold and the third 33-fold symmetry (Thomas et al., 1999). Subsequent work with over-expressed MS- and C-ring proteins shows azimuthal C-ring symmetry ranging from 31 to 38, with a peak around 34 (S. Khan, unpublished data). The MS-ring is composed of a single protein, FliF, and is believed to have 26- rather than 34fold symmetry (Jones et al., 1990, Sosinsky et al., 1992). Remarkably, a deletion-fusion mutant missing 10 % of the C-terminus of FliF and 28 % of the N-terminus of FliG, with the remaining parts of the two proteins fused together into a single poly-

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4.2 Structure

peptide, still produces functional motors. The mutant rotors have a smaller C-ring than the wild-type, consistent with 31 repeating units of normal size, and rotate more slowly. This has led to a model for rotation in which the eight ‘surplus’ units in the C-ring (five in the mutant) constitute defects in the interface between the MS- and C-rings, and these defects are passed around the rotor by the stator particles to generate rotation (Thomas et al., 1999). This model makes the curious prediction that the MS and C-rings rotate at different speeds, in the ratio 8 : 34. The prediction has not been tested, as motor rotation has only ever been measured via rotation of the hook and filament relative to the cell body. The exact orientation and number of copies of the different proteins in the rotor is not known. Quantitative immunoblot analysis combined with known molecular weights and rotor symmetries indicates a ‘best-guess’ model in which the MS-ring is a hub formed by 26 copies of FliF, 26 or 34 copies of FliG connect to this hub at the N-terminal and generate torque by interaction with the stator at the C-terminal. The rest of the C-ring is attached to FliG, consists of and 34 and 68 or 136 copies of FliM and FliN respectively, and controls motor switching as well as possibly interacting with FliG in torque generation (Matthews et al., 1998, Thomas et al., 1999). 4.2.3

Stator

Figure 4.3d shows a scanning EM image of a ring of studs left in the cytoplasmic membrane after freeze fracture (Khan et al., 1988). The number of studs per ring is variable, but averages around 11. This variability may be the result of freeze fracture pulling some studs into the other face of the membrane, or it may reflect real variability in the number of Mot complexes around the flagellum. Similar rings containing 14 to 16 studs were first seen at the poles of Aquaspirillum serpens, an organism with polar tufts of flagella (Coulton and Murray, 1978), and have also been found in an alkalophilic bacillus (Khan et al., 1992), presumably the stator of a Na motor. The central hole in the rings has the same diameter as the MSring and the particle in the middle of the hole matches the diameter of the flagellar rod. Mutants lacking either of the integral membrane proteins MotA or MotB produce apparently normal flagella which do not rotate, and in which no rings of studs can be found. Thus it appears that the ring of studs is the stator of the flagellar motor, and contains at least the proteins MotA and MotB. The model shown in Fig. 4.2 is deduced from the relative sizes of rings in the rotor and in the membrane and should be treated with a little caution as there is no direct evidence of the position of the C-ring relative to stator. Sequence analysis indicates that MotB has one membrane-spanning alpha-helical region near the N-terminus and a peptidoglycan-binding region at the C-terminus. MotA has four membrane-spanning regions and a large cytoplasmic domain. Tryptophan scanning mutagenesis of the membrane helices indicates which residues tolerate replacement by the bulky hydrophobic tryptophan side chain, and are thus likely to face the membrane. The pattern of these residues has led to a model in which the helix of MotB inserts obliquely into a bundle with the MotA


helices, together forming a stator unit which is anchored to the cell wall by the peptidoglycan binding site in MotB (Sharp et al., 1995). Although the Mot proteins show no homology to any previously identified proton translocating proteins, various lines of evidence indicate that the MotA/B stator particles are the proton channels of the motor. Cells that over-express wild-type Mot proteins show reduced growth rates that are attributable to leakage of protons across the cytoplasmic membrane. Paralysed mot mutants do not show the reduction in growth rate and give considerably reduced proton fluxes in artificially energized membrane vesicles made from over-expressing cells (Blair and Berg, 1990, Sharp et al., 1995, Zhou et al., 1998b). Inferred rates of proton flux are low when compared to expected rates based on levels of expression and the energetics of flagellar rotation. This is not surprising as the over-expressed Mot proteins are not incorporated into motors, whereas normal levels of proton flux are expected to depend upon coupling to rotation in complete motors. The best studied Na-driven flagellar motor is the polar motor of the marine bacterium Vibrio alginolyticus (Yorimitsu and Homma, 2001). This species swims with a single polar motor in media of low viscosity, such as sea water, and also expresses multiple lateral motors driven by H for swimming in viscous environments such as on the surfaces of fish. The proteins PomA and PomB in the Na-driven motor appear to be equivalent to MotA and MotB, with similarities in sequence, topology and function. Genetic fusion and gel purification and filtration studies indicate that Pom proteins are associated into complexes containing four molecules of PomA and two of PomB, and that each of these complexes may represent two functional torque-generating units (Yorimitsu and Homma, 2001). Indications are that MotA and MotB in H motors are arranged in the same way (Braun and Blair, 2001), and the results of experiments in which the number of units in H motors is altered while measuring rotation speeds are consistent with complexes containing two functional torque generators. In addition, Na motors have two other genes, motX and motY, (discovered before the pom genes) without which the motors do not rotate (McCarter, 1994a,b). These have no sequence homology to other motor genes. Like MotB, MotY is predicted to have a single membrane-spanning region and a peptidoglycan binding region. Over-expression of MotX in E. coli inhibits growth, but only in media of high Na concentration (where Na conduction would collapse the pmf). Amiloride, a specific blocker of the Na motor, blocks the growth inhibition, indicating that MotX can function as a Na channel. MotX and MotY are assumed to be part of the torque generator, but little further is known about their location or role. 4.2.4

Rotor Stator Interactions

The interactions between rotor and stator that generate torque have not been measured directly, but mutational analysis has shed some light on which protein domains are involved. A screen of all conserved acidic and basic residues in the proteins that are involved in torque generation (MotA, MotB, FliG, FliM, FliN) re-

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4.3 Function

vealed only one that is essential for rotation aspartate 32 in the membrane-spanning region of MotB (Zhou et al., 1998b). This residue is thought to be part of the proton conducting pathway of the motor. However, three charged residues in FliG and two in the cytoplasmic domain of MotA were found that affected rotation when replaced by other residues. Mutations in these residues are highly synergistic, and in particular changes in one protein are able to restore defects in rotation caused by changes in the other protein. The charge of the residues involved is crucial. For example reversing either the charge at position 281 in FliG or at position 98 in MotA impairs rotation considerably more than reversing both charges together. This indicates that the torque-generating mechanism relies on electrostatic interactions between the rotor and stator at these residues (Zhou et al., 1998a). Some support for this hypothesis is provided by the crystal structure of the C-terminal domain of FliG from a thermophilic bacterium (Lloyd et al., 1999), which is consistent with a model in which the key charged residues are on a face that interacts with the stator. FliG fusions using different parts of the protein from different bacteria show that the C-terminal domain of FliG from the thermophile was able to work in place of the E. coli domain. Similarly, the C-terminal domains of FliG from E. coli and the Na motor of Vibrio cholerae have been shown to be functionally interchangeable (Gosink and Hase, 2000).

4.3

Function

A motor is a machine that generates mechanical work from some other form of energy. The natural unit of energy for molecular motors is the thermal energy, kT, where k is Boltzmann’s constant and T is the absolute temperature. At room temperature, kT is about 4 q 10 21 J, and any possible motion that a molecular motor can make (that is, each ‘degree of freedom’) will be shaken by Brownian motion such that it contains half this amount of energy. The chemical energy released by hydrolysis of a single ATP molecule is about 20 kT, while a proton crossing the cytoplasmic membrane of bacteria releases electrochemical energy equal to the proton charge times the pmf. With a typical pmf of 150 mV, this is about 6 kT. Thus it is clear that molecular motors are subject to Brownian motion in as much as the energy changes involved in their mechanisms are in the same order of magnitude as the thermal energy. The work done by a force F moving distance d is simply the product Fd, where the movement is in the same direction as the force. Molecular motors have dimensions of several nanometres (1 nm 10 9 m) and exert forces of several piconewtons (1 pN 10 12 N), and in these units kT 4 pN nm. Torque is defined as the product of a force and the perpendicular distance to an axis of rotation, and therefore also has units of pN nm. This is not the same as energy however, as here the force and distance in question are perpendicular to each other, rather than parallel. In terms of energy, torque is best thought of as the work done per radian, where one radian (equal to about 57 h) is the angle for which the parallel distance moved by a force acting tangentially


to a circle is the same as the perpendicular, radial distance to the center. Since the angle in radians is the ratio of tangential to radial distance, angles are dimensionless and thus torque has the same dimensions (pN nm) as energy. In the bacterial flagellar motor, work is done at a rate equal to the product of torque and angular speed, and electrochemical energy is consumed at a rate equal to the product of pmf and proton flux. There are two complimentary approaches to understanding the mechanism of a motor. One is to look at what it is made of and how the parts join up, as in the previous section on motor structure. The other approach is to watch how it moves. New techniques have allowed the observation of single mechanical steps in the cycle that couples the hydrolysis of a single ATP ‘fuel’ molecule to motion in single kinesin, myosin and F1-ATPase motors (Noji et al., 1997). Similar experiments on the flagellar motor have not yet been successful, due mainly to the small and variable size of the fuel quantum, the fast motor cycle (10 100 q faster than known ATP motors) and the difficulty of re-constituting the complex structure in an in-vitro assay that allows the level of control necessary to measure single steps. What has been measured is how torque and speed depend upon pmf, proton flux and each other. These measurements allow models of the mechanism to be tested and are beginning to narrow down the wide range of possible mechanisms that have been proposed. 4.3.1

Motor Driven by H and Na Ion Flux

Aside from its rotary motion, the most striking difference between the flagellar motor and linear molecular motors of eukaryotes is that the flagellar motor is powered by transmembrane ion fluxes rather than ATP hydrolysis. The ion-motive kT Ci ln force consists of two components, and may be written as pmf C , q Co where C is the transmembrane voltage (inside minus outside), Ci and Co are the activities of the ions inside and outside the cell respectively, kT is the thermal energy and q the charge of the ion. The electrical component is simply the voltage, C; the chemical component is due to different ion concentrations on either side of the membrane and is represented by the second term on the right-hand side of the equation. Early evidence for a pmf driven motor in bacteria was based upon addition of agents that prevent the generation of ATP or of components of the pmf (Glagolev and Skulachev, 1978, Manson et al., 1977). Cells swam without ATP and without one or other component of the pmf, but not without both components of the pmf. These experiments did not rule out the possibility that the agents acted directly or indirectly on the motor by a route independent of ATP or the pmf. Conclusive evidence came when it was shown that flagella in cell envelopes containing no cytoplasm (‘ghosts’), isolated from E. coli, could rotate if an artificial diffusion potential was generated across the cytoplasmic membrane by the addition of K and valinomycin (Ravid and Eisenbach, 1984). The ion dependence of Na motors

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4.3 Function

was first demonstrated in alkalophilic bacillus (Hirota and Imae, 1983) and in V. alginolyticus (Chernyak et al., 1983), by demonstrating insensitivity to proton uncouplers but sensitivity to sodium concentration, sodium uncouplers and sodium channel blockers. The chemical and electrical components of the pmf or smf are equivalent in terms of torque generation in tethered cells (Manson et al, 1980, Hirota and Imae, 1983). Aside from the identity of the driving ion, Na motors seem to work with essentially the same mechanism as H motors. Lithium supports motor rotation in Na motors (Liu et al., 1990), and deuterium in H motors (Chen and Berg, 2000b), in both cases with normal torque at low speeds but considerably reduced torque at higher speeds. The most compelling evidence that the motors have a common mechanism is the existence of functional chimeras containing components of both types of motor (Yorimitsu and Homma, 2001). PomA can be replaced by MotA from the H motor of R. sphaeroides, and in addition the membrane-spanning part of PomB can be replaced by the corresponding part of MotB, and the resulting chimeric motors work in V. alginolyticus. The ion selectivity is for Na, indicating that Na selectivity is conferred by the rotor, MotX and MotY, or the periplasmic part of PomB. Severely impaired motility was also observed in Vibrio cholerae with MotA and MotB from E. coli replacing the Na motor components PomA and PomB, either in the presence or absence of MotX and MotY (Gosink and Hase, 2000). Selectivity of this hybrid motor is probably for H. This would indicate that neither MotX, MotY nor the rotor, confers ion selectivity, at least in this case. All attempts to date to rescue mot mutations in H-motor species with Na-motor stator components have failed. The identification of MotA/B and PomA/B complexes as the flagellar motor ion channels had led to the assumption that these proteins determine ion selectivity. If so, the results from chimeric motor studies restrict the selectivity determinant to the periplasmic domain of PomB, which had not previously been thought to be part of the channel. Clearly the question of ion selectivity is still to be resolved, in particular the role of MotX and MotY in Na motors. If the acidic residue asp32 in MotB is indeed a crucial part of the proton channel, the chimera data indicate that the corresponding residue in R. sphaeroides at least, is able to use Na in place of the proton. 4.3.2

Torque versus Speed

To understand mechanism of the flagellar motor we need to understand the mechanochemical cycle what the different states of the motor are and how they couple ion flux and rotation. In the absence of direct measurements on single steps in the cycle, the relationship between torque and speed is one of the best probes of the flagellar mechanism. Data can be compared to the predictions of models with different coupling mechanisms and used to eliminate models that do not fit. The earliest torque speed measurements recorded broad peaks in the frequency spectrum of fluctuations in light intensity, taken from images of a population of swimming Streptococcus cells (Lowe et al., 1987). A peak at 100 Hz repre-


sented the vibration of cell bodies at the frequency of the rotating flagellar bundle, while a second, low frequency peak was due to counter-rotation of the cell body. The motor rotation rate is the sum of these two frequencies. By measuring rotation rates in media of different viscosity, an approximately linear decrease in torque between 50 and 100 Hz was found, extrapolating to zero torque at around 110 Hz. Parallel measurements of bundle rotation rates using this technique and linear swimming speeds indicated that the two were directly proportional, allowing swimming speed to be used as an indicator of rotation rate. The rotation of single flagella in swimming V. alginolyticus cells has been measured using laser dark-field microscopy (Fig. 4.4a). Extremely high speeds have been recorded, up to 1700 Hz in media with 300 mM Na (Magariyama et al., 1994). As with E. coli, swimming speed varies linearly with motor rotation rate at the population level (Magariyama et al., 1995). However, swimming speed alone is not a reliable indicator of the rotation rate of individual flagellar motors as the constant of proportionality depends upon the variable length of flagella. Experiments which control the speed of the motor by altering the viscosity of the medium can only access a limited range of speeds. Using the technique of electrorotation, an external torque can be applied to the motor via the body of a tethered cell, allowing a far wider range of speeds to be examined in a single motor. In particular, the motor can be studied under conditions where it is made to rotate backwards or forwards at speeds higher than the zero-torque speed, by the electrorotation torque. These regimes are not accessible in cells rotating under their own power. The phenomenon of electrorotation has been known for a long time, but has only recently been applied to studies of the bacterial flagellar motor (Berg and Turner, 1993, Berry and Berg, 1998, Washizu et al., 1993). Essentially, a rapidly rotating electric field polarizes the cell body and the surrounding medium. Due to the high frequency of rotation (several MHz ), the polarization lags behind the electric field, and thus a torque is exerted upon the cell. This torque is proportional to the square of the electric field amplitude. Figure 4.4c illustrates schematically the electrorotation apparatus of Berg and Turner (1993). The gap between the electrodes was 70 mm, and up to 75 V could be applied to each opposing pair of electrodes, 90h out of phase, at frequencies in the MHz range. This allowed enough torque to be applied with electrorotation to spin tethered cells at speeds up to 1000 Hz in either direction. Even at these relatively high speeds, Reynolds number for a tethered cell is of the order of 10 3, which means that inertial forces are negligible. Therefore the rotation rate of the flagellar motor is always directly proportional to the total torque, with the constant of proportionality being the rotational viscous drag coefficient of whatever is rotating. The torque generated by the flagellar motor was measured by measuring rotation speeds with the same applied torque, before and after breaking the motor. Motors were either broken mechanically or de-energized by ionophores or ultraviolet irradiation that killed the cells. After breaking, motors did not rotate on their own and rotated at speeds proportional to the applied torque when electrorotated. The motor torque is thus proportional to the difference in speeds recorded at the same applied torque, before and after breaking. Absolute values of torque can be calculated by estimating the viscous drag coeffi-

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4.3 Function

A)

B)

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Torque (pN nm)

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o

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11.2 C

-200

0

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Rotation rate (Hz) Figure 4.4. The torque speed relationship of

the flagellar motor. Fast rotation of single motors has been measured in several different ways. (a) Under laser dark-field illumination (large arrow), single helical flagella appear as a series of bright bands (ovals) that travel away from the cell body as the helix rotates. A typical time-course of light intensity recorded through a slit (dark bar) placed in front of the cell image is shown. The broad peak marks the passage of the cell body and the rapid oscillations mark the passage of the rotating filament. (b) Polystyrene beads of diameter 0.3 1.0 mm can be attached to truncated flagella and their rotation recorded by monitoring the deflection of a focussed laser

beam. (c) A large torque can be applied to tethered cells using the technique of electrorotation. Oscillating voltages are applied to four micro-electrodes, 90h out of phase, to generate a rotating electric field at the cell. This exerts a torque (light arrow) that may be many times larger than the motor torque (dark arrow) and may be in either direction. (d) The flagellar motor generates nearly constant torque up to a certain speed which depends upon temperature. At higher speeds less torque is generated and at very high speeds the motor resists rotation. (Figure 4.4d adapted from Figure 6 in Berry and Armitage (1999) with permission from Elsevier Science Publishers).


cient of the tethered cell (Meister and Berg, 1987), or alternatively, as in Fig. 4.4d, by reference to stall torque measured with a calibrated optical trap (Berry and Berg, 1997). Slightly lower absolute values of torque are predicted based on the rotation rates of polystyrene beads attached to truncated flagella (Ryu et al., 2000). Figure 4.4d summarizes torque speed curves obtained from E. coli at different temperatures. The most unusual feature, compared to torque speed curves for other molecular motors, is the wide range of speeds over which the torque is nearly constant. The torque plateau has been confirmed in experiments where high speeds are achieved without electrorotation by attaching small polystyrene beads to truncated flagella (Chen and Berg, 2000a, Fig. 4.4b). By rapid exchange of media of different viscosity, the speed of rotation of single flagellar motors was varied up to Z 300 Hz. Another notable feature of the torque speed relationship is the continuity of torque either side of zero speed. This has been confirmed independently by using optical tweezers to stall tethered cells and then to measure the torque they exert either when pushed slowly backwards or allowed to rotate slowly forwards (Berry and Berg, 1997). The result is in contrast to similar measurements on kinesin, in which it was found that single motors could be stalled by a force of 5 pN, but would not slip backwards even when 13 pN was applied (Coppin et al., 1997). The continuity of torque either side of stall in the flagellar motor is evidence against a ratchet-like mechanism in which there is an irreversible step in the motor cycle. The torque plateau indicates that internal processes in the torque-generating cycle of the motor, (for example the motion of protons through the motor or possibly torque-generating conformational changes) are not rate limiting at these speeds. Rather, the rotation rate is limited mechanically by the load on the motor. At higher speeds torque is much reduced, indicating that internal processes are now rate limiting. At the zero-torque speed these internal processes dissipate all the energy available to the motor and the output power goes to zero. As an analogy, imagine a person riding a bicycle with one gear and no freewheeling. The ‘internal processes’ are the motion of the rider’s legs on the pedals and the output torque is what the rider applies to the back wheel via the chain. At low speeds and high loads, for example riding uphill, the rider can push the pedals with all his force and will produce constant torque. But if he is going very fast downhill, there comes a point where he can only just move his legs around fast enough to keep up with the pedals and is not pushing at all. Here, all his efforts are being dissipated in the motion of his legs. This interpretation is confirmed by the temperature dependence of the torque speed curves and also by experiments in which substitutions of non-physiological ions reduced motor torque in swimming cells (fast rotation) but not in tethered cells (slow rotation) (Chen and Berg, 2000, Liu et al., 1990). The rate-limiting steps at high speed are thermally activated and dependent upon the driving ion, as expected for internal electrochemical processes; but at low speeds the rate-limiting steps are independent of temperature and ion type, as expected for mechanical relaxation of a strained torque-generating state.

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4.3 Function

4.3.3

Independent Torque Generators

Good evidence exists that the particles in the stator ring of Fig. 4.3d are independent torque generators containing the proteins MotA and MotB. ‘Resurrection’ experiments have been carried out in E. coli in which wild-type MotA or MotB is expressed from an inducible plasmid in tethered cells of a non-motile mutant in the same mot gene (Blair and Berg, 1988, Block and Berg, 1984). These cells initially do not rotate, but start to rotate and then speed up in equal speed increments over the course of several minutes as the wild-type protein is expressed and incorporated into the motor (Fig. 4.5a). The maximum number of speed increments seen is eight, and after eight increments motor speeds remain steady at levels slightly faster than the speed of wild-type cells harvested during exponential growth. Comparing this to up to 16 particles typically seen in stator rings suggests that each increment may correspond to the addition of more than one particle and also that exponentially growing bacteria may not possess a full complement of force-generating units. In experiments where flagellar motors resurrect after being disrupted either mechanically or electrically (Berry et al., 1995, Fung and Berg, 1995), equal speed increments have been observed that are less than one-eighth of the maximum speed seen for the same cell. The question of the maximum number of torque-generating units per motor remains unresolved, but perhaps the most likely explanation of the data is that there are up to 16 units which come and go in pairs (in undamaged motors). This is consistent with the structural model in which a Mot complex contains two torque-generating units. No resurrection experiment has ever seen more than eight speed increments, which would be expected if the smaller speed increments seen with damaged motors corresponded to incorporation of complexes containing one damaged and one intact unit. Torque is generated by independent force-generating units in Na- as well as in H -driven motors. Induced expression of PomA in a pomA background restored swimming speeds from zero to wild-type levels over the course of about 10 min (Asai et al., 1997). Muramoto and co-workers (Muramoto et al., 1994) observed stepwise decreases in speed after ultraviolet irradiation of tethered cells of bacillus in the presence of a photoreactive analog of amiloride, a known inhibitor of Na channels and Na flagellar motors. These results are interpreted as the successive inactivation of independent units upon irreversible binding of the amiloride analog and suggest that there are between five and nine units per motor. Resurrection experiments with polystyrene beads attached to truncated flagella confirm that torque-generating units act independently at high as well as at low speeds (Ryu et al., 2000). Figures 4.5b and c show resurrection of motors attached to beads of diameter 1.0 and 0.3 mm respectively. The 1.0-mm data are essentially the same as those from tethered cells. At each step a new Mot complex adds a torque corresponding to 10 Hz, regardless of the other complexes and regardless of the increase in motor speed from 10 Hz for the first step to 60 Hz for the last step. The data with 0.3-mm beads are different. The first complex generates 104 Hz worth of torque, whereas after the last step the motor has six complexes


A) Speed (Hz)

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Figure 4.5. Independent torque generators. Resurrection of mot mutants in E. coli. (a c) Incorporation of torque-generating complexes after induction of wild-type mot genes in (a) tethered cells, (b) cells labelled with 1.0-mm beads and (c) cells labelled with 0.3-mm beads. (d) Torque speed relationships for different numbers of complexes from 1 to 5. The solid lines are to guide the eye. Dashed lines show the load lines for 1.0- and 0.3-mm beads and

connect data points derived from raw data as shown in (b) and (c) respectively. The torque speed curves all appear to have the same shape and differ only in the absolute values of torque. This indicates that complexes generate torque independently and with a high duty ratio. (Figure 4.5a Blair and Berg (1988); Figs 4.5b d adapted from Figures 1 and 3 in Ryu et al. (2000).

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4.3 Function

and is rotating at 230 Hz, so each complex is generating about 40 Hz worth of torque. The observed reduction in the torque generated per unit can be explained by considering the torque speed curve shown in Fig. 4.4e (which shows that the motor at room temperature generates about half as much torque at 230 Hz as it does at 104 Hz), so long as the torque speed dependence of each unit is the same as that of the whole motor. By estimating the drag coefficients of beads of different sizes, torque speed curves can be constructed for different numbers of Mot complexes from 1 to 5 (Fig. 4.5d). Comparison with a simple model indicates: that the torque speed relationship is property of individual torque-generating complexes, that there is no ‘freewheeling’ (complexes have a high duty ratio), that each complex acts independently, and that its rate-limiting transition at low load cannot be speeded up by torque imparted by other units via the rotor. The only interaction between the torque generators appears to be that they share the viscous load and thus increase the speed of the motor compared to a single generator driving the same size polystyrene bead. 4.3.4

Proton Motive Force, Sodium-motive Force, Ion Flux

So far there has only been one successful measurement of the proton flux through the flagellar motor. The change in the rate of proton uptake by a population of cells, monitored via changes in the pH of a weakly buffered medium in which the cells were swimming, was measured when flagellar rotation was stopped by adding anti-filament antibody to crosslink the filaments of different motors (Meister et al., 1987). It was deduced that the rotation-dependent proton flux is proportional to rotation rate, and corresponds to the transit of approximately 1200 protons per revolution of the motor. Patch-clamp techniques, which in principle offer the most attractive prospect of controlling pmf and measuring proton fluxes through flagellar motors, are impractical for a number of reasons. The small size of bacteria and the presence of the outer membrane makes patch-clamping the cytoplasmic membrane very difficult. It is possible to create giant spheroplasts by lysis of the cell wall, but the outer membrane often remains attached (Buechner et al., 1990, Martinac et al., 1990) and spheroplasts are non-motile, presumably because the cell wall plays a crucial function in anchoring the stator of the flagellar motor. Even if patch-clamping were possible, 1200 protons per revolution of a single motor rotating at 100 Hz amounts to a current of about Z 105 protons per second, or Z 0.01 pA, two orders of magnitude smaller than the single-channel currents typically recorded in patch-clamp experiments. Thus the possibility of directly measuring proton fluxes through single motors seems remote. Direct control of the membrane voltage at the flagellar motor has been demonstrated only once. E. coli were grown with cephalexin to produce cells tens of microns long with a single cytoplasmic compartment (Fung and Berg, 1995). Long cells were sucked into a custom-pulled micropipette with a narrow constriction about 10 mm from the tip, where they became stuck. The part of the cell membrane inside the pipette was made permeable by a proton ionophore in the pipette and a


transmembrane voltage was applied to motors on the external part of the cell by voltage-clamping the inside of the pipette. The arrangement and equivalent electrical circuit are shown in Figs 4.6a and b. Each cell lasted a few minutes, until diffusion of the ionophore collapsed the membrane voltage in the external part of the cell. Dead cells of normal size were attached to flagella on the external part of the cell as markers of rotation, essentially equivalent to tethered cells. The rotation rate of these markers was proportional to the applied voltage, as shown in Fig. 4.6c. Earlier work (in which the pmf in streptococcus was controlled using the K ionophore valinomycin and a K diffusion potential) showed a linear dependence of torque on pmf, extending all the way to zero pmf in tethered cells, with no threshold for rotation (Khan et al., 1985). A complication revealed by the voltage-clamp experiments is that torque-generating units inactivate upon de-energization and re-activate independently over a few minutes following re-energization. This is consistent with earlier work (Armitage and Evans, 1985) and explains an apparent threshold pmf for rotation that had been seen in earlier work upon re-energization but not upon de-energization of flagellar motors. It is not known whether inactivated units remain attached to the flagellar motor, or whether they diffuse away and have to be replaced from a circulating pool in the membrane. A fixed stoichiometry (1200 protons per revolution), the linear relationship between torque and pmf in tethered cells and the torque plateau shown in Fig. 4.4d, are all consistent with the interpretation that the motor is close to equilibrium at these speeds. In each revolution, the electrochemical energy input is equal to 1200 protons times the pmf, and the work done is equal to 2p radians times the torque. If the torque is proportional to pmf and does not depend directly on speed, then the input and output energies are proportional and thus the efficiency is constant. With a pmf of 150 mV and a plateau torque of 4000 pN nm, input and output energies per revolution come to 29,000 and 25,000 pN nm respectively, indicating that the efficiency is close to 1, and thus that the motor works close to equilibrium in the torque plateau. Measurements of the effect of pmf on motor torque at high speeds would be very informative, but to date have not been made. Various authors report threshold membrane potentials for swimming and saturation of swimming speed at potentials larger than 100 mV. However, this does not necessarily reflect a non-linearity between motor torque and pmf in the low-load, high-speed conditions of swimming cells. Rather, the threshold and saturation effects may reside in the hydrodynamics of swimming propelled by rotating flagella or may be due to uncertainty in the membrane voltage obtained using diffusion potential methods. Other authors have measured the dependence of motor speed on external pH and Na concentration (Chen and Berg, 2000b, Hirota and Imae, 1983). Both Na and H motors fail at extremes of pH, but to varying degrees in different experiments and it is not possible to distinguish specific effects of H concentration from general effects of extreme pH. The swimming speed of bacteria powered by Na-motors increases with Na concentration up to about 100 mM, but again these results are hard to interpret in detail because the internal ion concentration and membrane voltage are

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4.3 Function

A)

Rout

B)

Vm

Rin Vctrl

Raccess

Rleak

C) 4

2

Speed (Hz)

130

1

0 -50

-100

-150

Protonmotive Force (mV) Figure 4.6. Torque versus protonmotive force.

Torque and protonmotive force are proportional at high loads. (a) Long E. coli cells are sucked into a pipette with a constriction near the mouth. The inside of the pipette is held at Vctrl by a voltage clamp, and the section of the cell inside the pipette is made permeable to protons using an ionophore. A dead cell of normal size is attached to a flagellum in the part of the cell outside the pipette as a marker of rotation. (b) The equivalent circuit, from which the

membrane voltage in the outside part of the cell, Vm, can be deduced (after measuring the current which changes as the ionophore diffuses to the outside part of the cell and in the process reduces Rout). (c) Torque is proportional to the speed of rotation of the marker, which is shown as a function of the pmf (Vm) for markers on two different cells. (Data and figures adapted from Figures 1b and d and Figure 4 in Fung and Berg, 1995).


likely to vary with external ion concentration as a consequence of cellular homeostasis mechanisms (Hirota and Imae, 1983, Shioi et al., 1980). 4.3.5

Reversibility

Another remarkable feature of the flagellar motors of many species is their reversibility. Switches are fast, occurring within 1 ms in single filaments (Kudo et al., 1990) and the motor generates similar torque in either directional mode (Berry et al., 1995). Switches occur at random time intervals and individual switches of different motors on the same cell are not at all correlated, indicating that the stochastic process controlling motor switching occurs at the level of the individual motor (Ishihara et al., 1983, Macnab and Han, 1983). Switching is controlled by binding of the active form of the chemotaxis protein CheY to FliM in the Cring. In the absence of CheY, at room temperature, motors in E. coli rotate exclusively counterclockwise (CCW). Active CheY increases the probability of clockwise (CW) over CCW rotation in E. coli, presumably by reducing the free energy of the CW state relative to the CCW state. However, CW rotation can be observed in mutants lacking CheY if the temperature is reduced close to 0 hC, indicating an entropic contribution to the free energy difference between different rotational states (Turner et al., 1996). Current indications are that the basic torque-generating cycle of the motor is reversible with respect to the direction of proton flux. Berg et al. (1982) found a mutant streptococcus which did not switch in response to changes in cytoplasmic pH, but rotated in the opposite direction upon reversal of a pmf generated with valinomycin and a K diffusion potential. Using the voltage clamp technique, Fung and Berg (1995) found that reversing the direction of the pmf in E. coli usually caused motors to stop rotating, inactivating upon removal of the normal pmf. In five cases out of 17 however, motors reversed direction and rotated a few times before stopping. (It is not clear why streptococcus motors did not also de-activate upon reversal of the pmf). Thus the bacterial flagellar motor appears to be reversible at two levels. First, it possesses two modes which couple proton influx in normally energized cells to rotation in either direction. Second, each mode can probably be driven in either direction depending on the direction of the pmf. However, the effects of reversing the pmf upon the basic mechanism of the flagellar motor are confused, in E. coli at least, by the disassembly of motors in the absence of a normal pmf. 4.3.6

Steps?

A great deal has been learned about the mechanism of ATP-driven molecular motors by observing single mechanical steps corresponding to the hydrolysis of a single ATP molecule. Kinesin takes steps of 8-nm per ATP hydrolysed, equal to the ab-tubulin repeat length along the microtubule. F1-ATPase takes 120 h-steps,

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4.4 Models

corresponding to the three-fold symmetry of the motor. Based on the 34-fold symmetry of the C-ring, one might expect to see steps of around 11h in the rotation of a flagellar motor powered by a single torque generator. However, if 1200 protons cross the membrane per revolution in a wild-type motor with approximately 10 torque generators, one would expect steps of 3h for each proton in a single-generator motor. Technical difficulties have so far defied all attempts to measure directly the steps in the rotation of the flagellar motor and the possibility remains that the motor mechanism is such that there are no physical steps to be seen. It is however, possible to infer the stochastic behavior of the motor from fluctuations in rotation speed. Variance analysis of the time intervals taken by tethered cells to cover a fixed number of revolutions reveals fluctuations that are intrinsic to the flagellar motor mechanism (Berg et al., 1982). Under the assumption that rate-limiting events in the motor cycle occur at random times (a Poisson process), the data indicate that each revolution includes around 400 such events. Contributions to the variance from processes other than motor stepping would make this number an underestimate, while any regularity in the intervals between events would mean that 400 is an overestimate. A more sophisticated experiment, using electrorotation to control cell rotation rates (Samuel and Berg, 1995), excluded the possibility that the variance was due to free rotational Brownian motion superimposed upon a smooth motor rotation. As in the earlier work, these results were consistent with about 400 randomly occurring events per revolution of the motor. When the experiment was repeated in a resurrection strain in which motors contained between one and four torque generators, the predicted number of events was proportional to the number of generators, with about 50 per generator per revolution (Samuel and Berg, 1996). Whether these ‘events’ correspond to the transit of single protons or to some other chemical or mechanical step in the working cycle of the motor, remains to be seen.

4.4

Models

The level of detail in our current understanding of the structure and function of the bacterial flagellar motor, based on experimental results, leaves quite a lot of room for alternative models of the motor mechanism. Several discussions of competing models may be found (Berg and Turner, 1993, Berry, 2000), here we give an overview of the different modelling approaches that have been taken and what they can tell us.


4.4.1

Conceptual Models

Figure 4.7a shows three different ways in which the flagellar motor could generate torque. In the ‘turbine’ mechanism, protons interact simultaneously with rows of components on both the rotor and the stator. These rows are tilted with respect to one another and protons flow into the cell by travelling at their point of intersection. Proton influx makes the rotor and stator slide relative to each other so that the intersection of the tilted rows follows the proton as it passes through the motor. The tilted rows on the rotor could be lines of charged residues, so for example alternate negatively and positively charged lines would attract and repel a transient ion, coupling its transit to rotation (Berry, 1993). Alternatively, the lines on the rotor and stator could be residues that each constitute half a binding site for the ion. Binding sites are formed only at the intersection of rotor and stator lines and thus a transient ion would carry the intersection with it across the membrane, again leading to rotation (Lauger, 1988). The structure of the C-terminal domain of FliG shows alternating positive and negative charges on the face that interacts with MotA, which could form alternating lines of charges. In the ‘turnstile’ mechanism, ions are channelled onto the rotor from outside the cell (Meister et al., 1989). The rotor then moves, either under the effect of some force on the charged ions or simply due to thermal fluctuations. Each ion is allowed to pass into the cytoplasm, completing the motor cycle, only after the rotor has moved a certain distance in the right direction. With the ion gone, the rotor is locked to the stator again and the motor has made one step. This mechanism is favored for the proton-translocating FO rotary motor of ATP-synthase, with a conserved aspartate residue in the c-subunit identified as the proton-binding site on the rotor. However, it is worth noting here that extensive mutagenesis of Cring proteins has failed to identify any possible proton binding sites on the rotor of the flagellar motor (Zhou et al., 1998a). The defining element of a ‘crossbridge’ model is that the stator undergoes a conformational change while bound to the rotor, driven by ion translocation, which generates elastic strain in the crossbridge and causes the rotor to rotate. The rotor must then release the stator so that the conformational change can be reversed without reversing the rotation. Conformational changes in the cytoplasmic domain of MotA (containing the key residues that interact with the rotor) have been linked to protonation of the aspartate residue at position 32 in MotB (asp32) (Kojima and Blair, 2001). Conformational changes are inferred from changes in the pattern of proteolysis fragments of MotA caused by mutations in asp32 of MotB and the link to protonation is based on the observation that the most effective mutations were those that most closely mimic a neutral, protonated aspartate residue at position 32. In addition, changes were blocked by dominant non-rotating mutations of proline 173 in MotA, which also block proton flux though the Mot complex. This has led to a crossbridge model in which transient protons bind to and neutralize asp32, leading to conformational changes of MotA that exert torque on the rotor. The same conformational changes should

133

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4.4 Models

A)

Turbine

Turnstile

Crossbridge

B)

A

B

Rotor/stator interaction energy

f Angle Figure 4.7. Some models of the motor mechanism. (a) Three different types of model for the mechanism of the flagellar motor. In a ‘turbine’ model, ions flowing in the stator interact with tilted lines of charges on the rotor. In a ‘turnstile’ model, ions are deposited onto the rotor by one type of stator channel and can only complete a transit if the rotor rotates, carrying them to a second type of stator channel. In a crossbridge model, conformational changes of the stator driven by ion transit exert torque

upon the rotor. (b) Models can be understood in terms of motor states with different rotor stator interaction energy profiles. For example, a transition from state A to state B as shown by the arrow requires the input of chemical energy, which will be released as work as the motor in state B relaxes towards its new angle of minimum energy. (Figure 4.7a adapted from Figure 7 in Berry and Armitage (1999) with permission from Elsevier Science Publishers).


also couple the rotor angle to proton access to asp 32, although there are no structural details to indicate how this might be achieved. In practice, the boundaries between these categories become blurred when a detailed treatment is attempted. Nonetheless, they form a useful guide to describing and understanding specific mechanisms and help to illustrate the similarities and key differences between competing models. In any of the above schemes, the most likely mechanism for motor switching is a co-ordinated transformation of the Cring which reverses the handedness of the motor’s helical symmetry. 4.4.2

Kinetic Models

Regardless of the conceptual, structural or physical details of a model, an explicit mathematical description is needed in order to yield predictions that can be compared with experimental data. The classical approach, which can be traced to early models of muscle (Huxley and Simmons, 1971), is to define distinct chemical states for the motor, each with a characteristic functional dependence of free energy upon the rotor angle. Thermally-activated transitions between these states are allowed at appropriate angles and are driven by proton translocation. Transition rates depend indirectly on external load, which alters the probability of different rotor angles at which, in turn, the transition rates are different. An illustrative example is shown in Fig. 4.7b. Energy profiles for two states are shown, with energy minima separated by an angle f. Minima could be the rotor angles at which opposite charges on the rotor and stator align in a turbine model or alternatively the angle of minimum elastic strain in a crossbridge model. A transition from the energy minimum of state A to state B requires electrochemical energy and leads to torque generation as the motor in state B relaxes to its new angle of minimum energy. This type of mechanism is known as a ‘powerstroke’, in that the electrochemical energy directly drives a transition to a torque-generating state. Any model is fully specified by the energy profiles of the motor states, the transition rates between states as functions of rotor angle and the rotational diffusion coefficient of the rotor. In this way, models that are structurally different or that generate force via different physical interactions can be identical to each other in terms of the predicted behavior of the motor, and it is a matter of semantics whether to consider them as different physical manifestations of the same motor mechanism or as different mechanisms with the same functional characteristics. Once the model is specified, Monte-Carlo simulations of the equation of motion of the rotor coupled with the allowed kinetic transitions can be used to obtain predicted trajectories of the motor, while state occupancies, averaged torques and speeds can be found by numerical solution of the associated reaction diffusion equations (Elston and Oster, 1997). For a simpler treatment, some authors assume tight coupling between proton flux and rotation and combine chemical transitions and the subsequent mechanical relaxation into a single step, with a rate constant that depends explicitly on the mechanical load (Meister et al., 1989). Fitting such a model to the experimental torque speed relationship indicates that the flagellar

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4.5 Summary

motor operates with a ‘powerstroke’ mechanism. That is, there is a step in the cycle that converts most of the electrochemical energy of the proton directly into rotation (Berry and Berg, 1998). This is in contrast to ‘thermal ratchet’ models in which the role of the free energy supplied by the influx of protons is to ‘save’ thermal fluctuations in a certain direction rather than to create directly a torque-generating state. As more experimental data is collected on the performance of the flagellar motor, models will need to be extended and adapted to interpret these data and to narrow down the range of possible mechanisms.

4.5

Summary

A great deal has been learned about the structure and mechanism of the bacterial flagellar motor over the last 25 years. The structure of the rotor and stator are known to about 20 Å, and the first X-ray crystal structure of a part of the motor has been solved. Mutagenesis has revealed the topology of motor proteins and some details of how they interact with each other. The torque speed relationship has been measured both in the wild-type and in motors containing different numbers of torque-generating complexes. Complexes act independently and each generates about 500 pN nm of torque at speeds anywhere between 100 Hz backwards and about 160 Hz forwards at room temperature. At higher speeds and lower temperatures, motors run slower. The maximum output power of the motor is about 2 q 106 pN nm s 1, at least two orders of magnitude higher than any other known molecular motor. At low speeds, torque is proportional to pmf and the efficiency is close to 1, indicating that the motor is close to equilibrium. Motors can switch direction or can be driven backwards by reversing the direction of the pmf if the switch is disabled. However, there is still much to be learned. More crystal structures of motor parts are needed and these need to be fitted into ever-improved EM images of the whole motor. The comparison of H- and Na-driven motors, and in particular a better understanding of hybrid motors, should reveal more about the role of ions in the torque-generating mechanism. Measurement of the torque speed relationship at different values of pmf or smf would be highly informative, as would any new information on the flux of ions through the motor. Hopefully the application of new techniques such as optical tweezers to the study of motors that run on a single torque-generating complex will reveal single steps in motor rotation corresponding to individual events in the motor cycle. As the structural and functional data accumulate, it should eventually be possible to understand in detail the physical and biochemical principles that underlie the working of this remarkable biological, rotary, electric motor.


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4 The Bacterial Flagellar Motor Matthews, M. A. A., H. L.Tang, and D. F. Blair. 1998. Domain analysis of the FliM protein of Escherichia coli. J. Bacteriol. 180: 5580 5590. McCarter, L. L. 1994a. MotX, the channel component of the sodium-type flagellar motor. J. Bacteriol. 176: 5988 5998. McCarter, L. L. 1994b. MotY, a component of the sodium-type flagellar motor. J. Bacteriol. 176: 4219 4225. Meister, M. and H. C. Berg. 1987. The stall torque of the bacterial flagellar motor. Biophys. J. 52: 413 419. Meister, M., S. R. Caplan, and H. C. Berg. 1989. Dynamics of a tightly coupled mechanism for flagellar rotation. Biophys. J. 55: 905 914. Meister, M., Lowe, G., and Berg, H. C. 1987. The proton flux through the bacterial flagellar motor. Cell. 49: 643-650. Muramoto, K., S. Sugiyama, E. J. Cragoe Jr., and Y. Imae. 1994. Successive inactivation of the force-generating units of sodium-driven bacterial flagellar motors by a photoreactive amiloride analog. J. Biol. Chem. 269: 3374 3380 Namba, K. and F. Vonderviszt. 1997. Molecular architecture of bacterial flagellum. Q. Rev. Biophys. 30: 1 65. Noji, H., R. Yasuda, M. Yoshida, and K. Kinosita. 1997. Direct observation of the rotation of F1-ATPase. Nature 386: 299 302. Pijper, A. 1948. Bacterial flagella and motility. Nature 161: 200 201. Pitta, T. P., E. E.Sherwood, A. M. Kobel, and H. C. Berg. 1997. Calcium is required for swimming by the nonflagellated cyanobacterium Synechococcus strain WH8113. J. Bacteriol. 179: 2524 2528. Ravid, S. and M. Eisenbach. 1984. Minimal requirements for rotation of bacterial flagella. J. Bacteriol. 158: 1208 1210. Ryu, W. S., R. M. Berry, and H. C. Berg. 2000 Torque generating units of the flagellar motor of Escherichia coli have a high duty ratio. Nature 403: 444 447. Samatey, F. A., K. Imada, S. Nagashima, F. Vonderviszt, T. Kumasaka, M. Yamamoto, and K. Namba. 2001. Structure of the bacterial flagellar protofilament and implications for a switch for supercoiling. Nature 410: 331 337. Samuel, A. D. T. and H. C. Berg. 1995. Fluctuation analysis of rotational speeds of the bacterial flagellar motor. Proc. Natl. Acad. Sci. USA 92: 3502 3506.

Samuel, A. D. T. and H. C. Berg. 1996. Torquegenerating units of the bacterial flagellar motor step independently. Biophys. J. 71: 918 923. Samuel, A. D, J. D. Petersen, and T. S. Reese. 2001. Envelope structure of Synechococcus sp. WH8113, a nonflagellated swimming cyanobacterium. BMC Microbiol. 1: 4. Segall, J. E., S. M. Block, and H. C. Berg. 1986. Temporal comparisons in bacterial chemotaxis. Proc. Natl. Acad. Sci. USA. 83: 8987 8991. Sharp, L. L., J. Zhou, and D. F. Blair. 1995. Tryptophan-scanning mutagenesis of MotB, an integral membrane protein essential for flagellar rotation in Escherichia coli. Biochemistry 34: 9166 9171. Shioi, J-I., S. Matsuura, and Y. Imae. 1980. Quantitative measurements of proton motive force and motility in Bacillus subtilis. J. Bacteriol. 144: 891 897. Silverman, M. and M. Simon. 1974. Flagellar rotation and the mechanism of bacterial motility. Nature 249: 73 74. Sosinsky, G. E., N. R. Francis, D. J. DeRosier, J. S. Wall, M. N. Simon, and J. Hainfeld. 1992. Mass determination and estimation of subunit stoichiometry of the bacterial hook basalbody flagellar complex of Salmonella typhimurium by scanning transmission electron microscopy. Proc. Natl. Acad. Sci. USA 89: 4801 4805. Stock, J. B. and M. G. Surette. 1996. Chemotaxis. In: Escherichia coli and Salmonella: Cellular and Molecular Biology. Edited by F. C. Neidhardt, R. Curtiss, I, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger. Washington: ASM, pp. 1103 1129. Thomas, D. R., Morgan, D. G., and D. J. DeRosier. 1999. Rotational symmetry of the C ring and a mechanism for the flagellar rotary motor. Proc. Natl. Acad. Sci. USA 96: 10134 10139. Turner, L., S. R. Caplan, and H. C. Berg. 1996. Temperature-induced switching of the bacterial flagellar motor. Biophys. J. 71: 2227 2233. Turner, L., W. S. Ryu and H. C. Berg. 2000. Real-time imaging of fluorescent flagellar filaments. J. Bacteriol. 182: 2793 2801. Washizu, M., Y. Kurahashi, H. Iochi, O. Kurosawa, S-I. Aizawa, S. Kudo, Y. Magariyama,

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References and H. Hotani. 1993. Dielectrophoretic mea- stator in the bacterial flagellar motor. Proc. Natl. Acad. Sci. USA 95: 6436 6441. surement of bacterial motor characteristics. Zhou, J., L. L. Sharp, H. L. Tang, S. A. Lloyd, S. IEEE Trans. Ind. Applicat. 29: 286 294. Billings, T. F. Braun, and D. F. Blair. 1998b. Yonekura, K., S. Maki-Yonekura, and K. Function of protonatable residues in the flaNamba. 2001. Structure analysis of the flagellar cap-filament complex by electron cryo- gellar motor of Escherichia coli: a critical role for Asp 32 of MotB. J. Bacteriol. 180: microscopy and single-particle image analy2729 2735. sis. J. Struct. Biol. 133: 246 253. Yorimitsu, T. and M. Homma. 2001. Na-driv- Zusman, D. R., M. J. McBride, W. R. McCleary, en flagellar motor of Vibrio. Biochim. Biophys. and K. A. O’Connor. 1990. Control of directed Acta 1505: 82 93. motility in Myxococcus xanthus. In: Biology of Zhou, J., S. A. Lloyd, and D. F. Blair. 1998a. the chemotactic Response. Edited by J. P. ArElectrostatic interactions between rotor and mitage and J. M. Lackie. Cambridge, UK: CUP, pp. 199 219.

5 F1-Motor of ATP Synthase Hiroyuki Noji

5.1

Introduction

The F1-motor is the soluble portion of ATP synthase and acts as a rotary motor that generates rotary torque using the energy derived from ATP hydrolysis. The rotation of the isolated F1-motor was directly observed in the light microscope and its mechanical properties have been studied in detail; the motor rotates in discrete 120 hsteps, each of which is driven by the hydrolysis of one ATP molecule and generates a constant torque of 40 pN nm irrespective of a load. Recently, two important contributions were made: (1) high-speed imaging of the rotation and (2) novel crystal structures of the F1-motor. The results revealed that a single 120 h-step is composed of 90 h- and 30 h-substeps, which are driven by ATP binding and, presumably, the release of ADP or phosphate, respectively. In this brief review, a model for catalysis and rotation of the F1-motor is proposed based on these new insights.

5.2

ATP Synthase

ATP synthase is a ubiquitous enzyme located in the inner membranes of mitochondria, thylakoid membranes of chloroplasts or the plasma membranes of bacteria. As implicated by the binding-change mechanism proposed by P. D. Boyer, ATP synthase employs the mechanical rotation of its subunits to convert the electrochemical potential of protons across membranes (DmH), which is generated by respiration or photo-reaction, into chemical energy for ATP synthesis (Boyer, 1993). This enzyme comprises two motors sharing a common rotor shaft (Fig. 5.1) (Noji and Yoshida, 2001). The subcomplex protruding from the membrane, the F1-motor, can generate rotary torque using the energy of ATP hydrolysis. Its subunit composition is a3b3g1d1e1and the Mr is Z 380,000. The F0 -motor, a membrane-embedded subcomplex, generates torque that is coupled to proton movement down DmH. The bacterial F0 has a Mr of Z 150,000 and the subunit structure a1b2c10 14. The

142

5.3 F1-Motor

Figure 5.1. Schematic diagram of the ATP synthase. Left, side view of ATP synthase. The ATP synthase is composed of the F1- and the F0 motor sharing a common rotary shaft (light gray). A stator connects the two motors (dark gray) so that they do not slip. Right, the two

motors can be reversibly separated. The F1motor generates a rotary torque using the energy of ATP hydrolysis. The F0 -motor utilizes a proton flow for torque generation. The directions of the motors are opposite to each other.

g subunit is the major part of the F1 rotor shaft and attaches to the rotor part of the F0 -motor. The stator stalk composed of b and d subunits also connects F1 to F0 and keeps the stator from spinning with the rotor. Under physiological conditions where the driving force of the F0 -motor surpasses that of the F1-motor, the F0 motor rotates the common shaft in its intrinsic direction so as to reverse the rotational direction of the F1-motor, thereby catalyzing ATP synthesis. When the driving force of the F1-motor dominates, the F1-motor reverses the F0 -motor and protons are pumped to the opposite side of the membrane.

5.3

F1-Motor

F1 is dissociated from F0 under low ionic conditions and becomes the soluble enzyme that only hydrolyses ATP; hence the name F1-ATPase. The catalytic sites are mainly located in the b subunit, but the minimum complex that shows active and stable ATPase activity is the a3b3g subcomplex. The crystal structure of the a3b3g subcomplex of bovine mitochondrial F1 shows that three a and three b subunits are arranged alternately in a hexamer ring thereby forming a large central cavity in which half of the coiled-coil structure of the g subunit is inserted (Abrahams et al., 1994). According to the recently reported structure of the F1 F0c complex of yeast ATP synthase, the other half of the coiled-coil portion of the g subunit ex-

5 F1-Motor of ATP Synthase

a)

b) Ribbon representation of the a subunit (green), the b subunit (gray) and the g subunit (red) in the 1994 crystal structure of the F1-motor. (a) Side-view. (b) View from membrane side.

Figure 5.2.

143

144

5.4 Imaging of Rotation of F1-Motor

tends into the rotor part of the F0 -motor (Stock et al., 1999). The structure reported in 1994 reveals that the three catalytic sites on the b subunits differ in their nucleotide-binding states. The first site is occupied by Mg-AMP-PNP (a non-hydrolyzable analog of ATP), the second by Mg-ADP, and the third is empty (no bound nucleotide). The sites are termed bATP, bADP and bempty, respectively. These structural features are consistent with the prediction of the binding-change mechanism: three catalytic sites should be in the three different nucleotide states and cooperative interconversion of the states causes the rotation of the g subunit.

5.4

Imaging of Rotation of F1-Motor

The rotary motion of the g subunit of the F1-motor was directly observed with a singlemolecule imaging technique (Noji et al., 1997). To visualize the rotation, F1 molecules from a thermophilic bacterium (Bacillus strain PS3) were attached to a glass surface. As a marker for rotation 1 to 4 mm long fluorescently labeled actin filaments were attached to the g subunit of F1-motors (Fig. 5.3, left). Depending on the ATP concentration, the rotation of actin filaments at 0.2 10 revolutions s 1 was observed in the light microscope. The direction of the rotation was always counterclockwise when viewed from the Fo side, consistent with the data from the crystal structure in which one b undergoes transition from bATP to bADP, then to bEmtpy. In this experimental system, the F1-motor rotates an actin filament against hydrodynamic friction. Therefore the rotary torque was calculated from the hydrodynamic friction which is proportional to the cube of the filament length. These experiments showed that the F1motor generates a constant torque of about 40 pN nm irrespective of the load, the ATP concentration or the rotary angle (Yasuda et al., 1998). At low ATP concentrations (I 600 nM), actin filaments showed a stepwise rotation. This is because the limiting reaction step is ATP binding. Thus the F1-motor waits for ATP at the fixed position, rotates 120h upon arrival of ATP, and then waits for the next ATP. A 120 h-step is consistent with the three-fold arrangement of catalytic b subunits in the a3b3 hexamer. A statistical analysis of the observed steps confirmed that the hydrolysis of one ATP molecule is sufficient to make one 120 h-step (Yasuda et al., 1998). Furthermore, it was also shown that the viscous load on the rotation did not affect the apparent rate constants of ATP binding. This is interesting because ATP binding is a torque-generating step, as discussed below. In one experiment, the apparent rate constant of the ATP binding reaction could be affected by a load. This discrepancy is discussed in the last paragraph. The F1-motors from Escherichia coli and chloroplasts were also shown to be rotary motors (Hisabori et al., 1999, Noji et al., 1999). There was no obvious difference between the observed rotations of these F1-motors. The mechanical properties of the different F1-motors seem to be conserved among species.


Figure 5.3. Experimental systems for singlemolecule observation of the rotation of the F1motor. Left, a long actin filament (Z 5 mm) modified with fluorescent dyes is attached to the g subunit of the F1-motor which is immobilized on a cover slip. Rotary movement of the filament can be observed with a fluorescence

microscope. Only part of the filament is depicted. Right, a small marker, a gold bead of 40 nm in diameter, is attached to the g subunit. Its viscous friction is 10 3 to 10 4 times smaller than that of an actin filament. Rotary movement of beads was observed by laser dark-field optics.

5.5

High-speed Imaging of F1 Rotation

Our group has observed the rotation of F1-motors at the light microscopic level using either actin filaments or plastic beads as markers. However, the viscous friction imposed on the large marker prevented fast rotation of the F1-motor and obscured the stepping behavior. Recently, we have developed a new experimental system to improve the visualization of rapid rotation. By using a smaller marker, a colloidal gold bead of 40 nm diameter, the viscous friction is reduced by a factor of 10 3 to 10 4 compared to actin filaments (Fig. 5.3, right) (Yasuda et al., 2001). Thus, the marker does not essentially impede the rotation of the motor any more. The motion of a gold bead was imaged by laser dark-field microscopy using a high-speed camera at Z 8000 frames per second. By this high-speed imaging, it was revealed that the F1-motor rotates at 130 Hz at saturating ATP concentrations. This was expected from the rate of ATP hydrolysis measured in solution. Surprisingly, the 120 h-stepping rotation could be further resolved into 90 h- and 30 h-substeps (Fig. 5.4). Kinetic analysis of the observed rotation showed that ATP binding drives a 90 h-substep, and a 30 h-substep is generated by an additional reaction following a mechanically silent reaction as shown in the following reaction scheme. ATP binding 90 hsubstep reaction 1 reaction 2 30h-substep In the reaction scheme, ‘reaction 1l is mechanically silent, and ‘reaction 2l generates the 30 h-substep. The two reactions take 1 ms. Previous biochemical studies have shown that the step that hydrolyzes ATP does not release energy. The ATP in

145

146

5.6 New Crystal Structure for the F1-Motor

Time-course of the rotation of a gold bead at 20 mM ATP. The curves in the panel are continuous. The gray horizontal lines are placed 30¡ below the black lines. The circles in the inserts indicate a projection of Z 0 h- and Z 90 h-dwell points on a circular trajectory.

Figure 5.4.

the catalytic site is in equilibration with ADP Pi in roughly a 2 : 1 ratio, thus energy levels of both states are comparable to each other. The hydrolysis step is, therefore, a likely candidate for ‘reaction 1l, and product(s) release (ADP, phosphate, or both) is ‘reaction 2l which generates a 30 h-substep. Indeed, the crystal structure suggests that the F1-motor changes its conformation drastically upon ADP release; the conformation of the b subunit with nucleotide was different from that without nucleotide (Fig. 5.5a,b).

5.6

New Crystal Structure for the F1-Motor

A new crystal structure for the F1-motor was presented by Walker and Leslie’s group in 2001; it suggested the existence of a possible intermediate state between the 90 h-substep and the 30 h-substep (Menz et al., 2001). They had reported several crystal structures for the F1-motor prepared under different conditions before this new structure. However, all these structures are essentially identical to that reported in 1994 except for minor differences. In contrast, the new structure reported in 2001 has two remarkable features. First, the conformation of the b subunit is different. In the structures presented previously, one b subunit without bound nucleotide exhibited an ‘open’ conformation, in which the crevice between the Cterminal domain and the nucleotide-binding domain was open (Fig. 5.5a), whereas two b subunits with a bound nucleotide were in a ‘closed’ conformation in which the C-terminal domain was lifted up to the nucleotide-binding domain (Fig. 5.5c). In the new structure, two b subunits bind ADP AlF4 at their catalytic sites repre-


a)

Figure 5.5. Three different conformations of the b subunit. The central g subunit (red), the b subunit (gray and green) and the a subunit are viewed from the side. The two a-helices in the C-terminal domain of the b subunit are colored green to emphasize the conformational differences. In the 1994 crystal structure one b subunit without bound nucleotide is in the ‘open’ conformation (a). In the 2001 structure, the b subunit that corresponds to the ‘open b subunit’ in the 1994 structure, binds ADP and SO2– 4 and assumes the ‘halfclosed’ conformation (b). The two b subunits that bind an analog of ATP (AMPPNP) or ADP are in the ‘closed’ conformation (c).

b)

c)

147

148

5.7 Catalysis and Rotation of F1-Motor

senting a ‘closed’ form similar to the 1994 structure. The third b subunit binds ADP and SO2– 4 and, instead of showing the ‘open’ conformation as in the 1994 structure, lifts the C-terminal domain to take on a ‘half-closed’ conformation (Fig. 5.5b). This structure might represent a catalytic intermediate state after hydrolysis but before release of ADP and phosphate, because the aArg373 intrudes between the binding sites of ADP and phosphate to prevent binding of ATP but to allow binding of ADP and phosphate. Second, when compared to the 1994 structure, the g subunit appears twisted. Although the angle of the a helix around the C-terminus shows only 1h deviation, the middle region and the protruding portion rotate 20h in the clockwise direction, which is opposite to the rotational direction of the F1-motor. These regions have contact with the C-terminal domain of the ‘half-closed’ b subunit. Therefore, open movement of this domain seems to induce a 20 h-rotation of the middle and the protruding portions of the g subunit when the ‘half-closed’ b subunit transforms to the open conformation upon release of products. Thus, the difference between the 1994 and 2001 structures suggests the existence of an intermediate state between the 90 h- and 30 h-steps, which is consistent with our observations.

5.7

Catalysis and Rotation of F1-Motor

As seen in the crystal structures, the three b subunits exhibit different conformations, i. e. they differ in their catalytic state. The interconversion between the catalytic states of the three b subunits is thought to generate the rotation of the g subunit. I propose a scheme that combines both the 1994 and 2001 structures (Fig. 5.6). The arrow in Fig. 5.6 indicates the position of the protruding parts of the g subunit. In the 1994 structure, two of three b subunits bind an analog of ATP and ADP respectively, and both appear in the ‘closed’ conformation. However in state 1, I have depicted the b subunit (green) which originally bound ADP according to the 1994 structure, as the one that binds ADP and phosphate instead. There are two reasons for this: (1) the interface between the b subunit and the neighboring a subunit is tightly closed and the bound ADP is encapsulated inside the protein, thus the b subunit with ADP has the highest affinity for nucleotide. (2) Biochemical studies have shown that the hydrolysis reaction occurs only at the highest affinity site and is in equilibrium with ATP synthesis. When the empty b subunit (blue) which is in the ‘open’ conformation, binds ATP, it does so without a large conformational change (state 2, first docking). Then, it transforms from an ‘open’ to a ‘closed’ conformation (state 3, induced fit) to reach the final binding state (state 4, second docking). The induced fit accompanies the gradual transformation of the C-terminal domain, keeping direct contact with the g subunit as seen in the crystal structures (see Fig. 5.5). Therefore the b subunit pushes the g subunit, causing it to rotate during the transformation. Triggered by the rotation, the neighboring b subunit (green) changes the conformation from ‘closed’ to ‘half-closed’, in which the ATP is irreversibly hydrolyzed into ADP and phosphate. In the state referred to as second docking, the middle and pro-


Figure 5.6. Model for catalysis and rotation of the F1-motor. Three b subunits surround the central g subunit (yellow arrow). ‘TP’, ‘DPpP’, ‘DP’, and ‘P’ denote ATP, ADP and phosphate, in the equilibrium with ATP synthesis, ADP and phosphate. The transformation from state 1 to state 7 corresponds to a single 120 h-step comprising 90 h- and 30 h-substeps. For details, see text.

truding portions of the g subunit rotate 90h relative to the original angle, whereas the C-terminal region rotates by 120 h. Subsequently, the b subunit occupied by ATP (orange) is transformed into the next highest affinity state for ATP hydrolysis, closing its interface with the neighboring a subunit (state 5). This takes 1 ms. This state corresponds to the one in the 2001 structure. In the next 1 ms ADP and phosphate are released from the b subunit in the ‘half-closed’ conformation (green; state 6). This b subunit then changes to the relaxed state, or ‘open’ conformation. This process is accompanied by a 30 h-rotation of the middle and protruding parts of the g subunit (state 7).

149

150

5.8 Perspectives

In my model, the F1-motor directly converts the binding energy released upon ‘induced fit’ into mechanical work against viscous friction. This means that the F1-motor dissipates binding energy into the medium as heat, without storing the energy in its conformation. However, another model seems to be possible in which the F1-motor rotates the g subunit after completion of the induced fit. In this case, the F1-motor should store the binding energy in its conformation and then release the energy as the driving force for the rotation. The stored energy could be converted not only into the generation of torque, but also into the energy for the release of the bound ATP in reverse. This is because the F1-motor is a reversible motor that is induced to release ATP by the torque of the F0 -motor during ATP synthesis. The possibility that the F1-motor releases bound ATP before rotation would increase when the F1-motor coupled to a large marker, such as an actin filament. Therefore, the latter model disagrees with the observation that an actin filament attached to the g subunit does not affect the apparent rate constant of ATP binding (Yasuda et al., 1998). In contrast, in my model, the F1-motor does not store binding energy, instead it is dissipated as heat. The viscous load on the actin filament only makes the induced fit slow. Thus, our model can explain that the apparent rate constant of ATP binding is less sensitive to the viscous load. Oster’s group has also proposed essentially a similar concept to the ‘induced fit’ described here and has called it ‘binding transition’ (Oster and Wang, 2000). They employ the term ‘binding transition’ to explain the high efficiency of energy conversion by the F1-motor and the constant torque irrespective of the rotary angle (see Chapter 8). It should be noted that our model is based on the so-called ‘tri-site’ model in which the occupation of all three catalytic sites with nucleotide is necessary for rapid catalysis and rotation, because the F1-motor in the 2001 structure has nucleotide bound to all three catalytic sites. P. D. Boyer rejects the ‘tri-site’ model and instead proposes that the occupation of two catalytic sites is sufficient to support rapid catalysis and rotation (‘bi-site’ model) (Boyer, 2002). On the other hand, the ATP-induced change of tryptophan fluorescence at the catalytic sites supports the tri-site model (Ren and Allison, 2000, Weber and Senior, 2001). Experiments using a simultaneous observation of the binding of fluorescently labeled ATP and the rotation of the F1-motor should settle this question.

5.8

Perspectives

I think the most prominent feature of the F1-motor is not just rotary movement, but its reversibility. The ATP hydrolysis reaction of the F1-motor is tightly coupled to the rotation of the g subunit; we can therefore control the catalytic reactions of the F1-motor by rotating the g subunit with an external force. In the near future, the molecular mechanism of this motor will be studied in more detail with single-molecule manipulation techniques (e. g. optical tweezers, magnetic tweezers, AFM). It will also be a challenge to observe directly the rotation of the proton-


driven F0 -motor. For this we need a truly novel experimental system in which a voltage can be applied to a membrane containing the F0 -motor while visualizing its rotation with the help of a suitable marker.

Acknowledgements

I thank Drs T. Nishizaka, E. Muneyuki and U. Schliwa for critical reading of the manuscript, and M. Yoshida, K. Kinosita, Jr., R. Yasuda, K. Adachi and H. Itoh for useful discussions.

References Abrahams, J. P., A. G. W. Leslie, R. Lutter, and J. E. Walker. 1994. Structure at 2.8-Å resolution of F1-ATPase from bovine heart mitochondria. Nature 37: 621 628. Boyer, P. D. 1993. The binding change mechanism for ATP synthase some probabilities and possibilities. Biochim. Biophys. Acta 1140: 215 250. Boyer, P. D. 2002. Catalytic site occupancy during ATP synthase catalysis. FEBS Lett. 512: 29 32. Hisabori, T., A. Kondoh, and M. Yoshida. 1999. The g subunit in chloroplast F1-ATPase can rotate in a unidirectional and counter-clockwise manner. FEBS Lett. 463: 35 38. Menz, R. I., J. E. Walker, and A. G. W. Leslie. 2001. Structure of bovine mitochondrial F1ATPase with nucleotide bound to all three catalytic sites. Cell 106: 331 341. Noji, H., R. Yasuda, M. Yoshida, and K. Kinosita, Jr. 1997. Direct observation of the rotation of F1-ATPase. Nature 386: 299 302. Noji, H., K. Hasler, W. Junge, K. Kinosita, Jr, M. Yoshida, and S. Engelbrecht. 1999. Rotation of Escherichia coli F1-ATPase. Biochem. Biophys. Res. Commun. 260: 579 599.

Noji, H. and M. Yoshida. 2001. The rotary machine in the cell, ATP synthase. J. Biol. Chem. 276: 1665 1668. Oster, G. and H. Wang. 2000. Reverse engineering a protein: the mechanochemistry of ATP synthase. Biochim.Biophys. Acta, 1458: 482 510. Ren, H. and W. S. Allison. 2000. Substitution of bGlu201 in the a3b3g subcomplex of the F1ATPase from the thermophilic Bacillus PS3 increases the affinity of catalytic sites for nucleotides. J. Biol. Chem., 275: 10057 10063. Stock, D., A. G. W. Leslie, and J. E. Walker. 1999. Molecular architecture of the rotary motor in ATP synthase. Science 286: 1700 1705. Yasuda, R., H. Noji, K. Kinosita, Jr, and M. Yoshida. 1998. F1-ATPase is a highly efficient molecular motor that rotates with discrete 120 degree steps. Cell 93: 1117 1124. Yasuda, R., H. Noji, M. Yoshida, K. Kinosita, Jr, and H. Itoh. 2001. Resolution of distinct rotational substeps by submillisecond kinetic analysis of F1-ATPase. Nature 410: 898 904. Weber, J. and A. E. Senior. 2001. Bi-site catalysis in F1-ATPase: does it exist? J. Biol. Chem. 276: 35422 35428.

151

6 RNA and DNA Polymerases Nataliya Korzheva and Arkady Mustaev

6.1

Introduction

Transcription and replication in the living cell are among the most fundamental processes that constitute the essence of life. Replication of DNA is accomplished by DNA polymerases (DNAP) that synthesize exact copies of genomic DNA during cell division, while the expression of genetic information encoded in DNA is provided by cellular RNA polymerase (RNAP). Both kinds of enzymes perform the same type of reaction polymerization of nucleoside triphosphates (NTPs), suggesting similarities in their catalytic mechanisms. From a mechanistic point of view polymerases can be viewed as molecular machines that perform directional mechanical work against dissipative forces during polymerization. Such forces are hydrodynamic drag and impediments imposed by specific sequences encoded in DNA as well as by numerous proteins that hold on to DNA such as histones, regulatory factors and so forth. From this perspective both DNA and RNA polymerases can be compared with the other extensively studied molecular motors reviewed in the other chapters of this book such as myosins, kinesins, dyneins and bacterial flagellar motors. Some important mechanical and thermodynamic parameters of polymerases and other molecular motors are shown in Tab. 6.1. The interest in mechanical considerations of polymerases has been kindled by recent observations that the force generated by these enzymes greatly exceeds that of other known molecular motors. This consideration seems to be extremely useful since it means that general kinetic and thermodynamic concepts which were developed for more conventional molecular motors, can be applied to NTP polymerizing enzymes to better understand the mechanisms of DNA and RNA synthesis. Comparison of the properties of RNA and DNA polymerases reveals principal differences to kinesin-like motor proteins (Julicher and Bruinsma, 1998). These differences reflect the different purposes of these two kinds of enzymes: while motor proteins perform fast active transport in cells, the purpose of NTP polymer-

154

6.1 Introduction Table 6.1.

Mechanical and thermodynamic parameters of polymerases and other molecular

motors. Motor

Force

Step size, nm

Velocity, nm/s

Efficiency, %

RNAP

14–25 pN

0.34

3.4–17

9–15

DNAP

34 pN

0.34

34–340

23

Kinesin

6 pN

8

800–3000

40–60

Myosin

3–5 pN

5–15

3000

12–42

F1-ATPase

40 pN/nm

120 h

4 rps

100

izing enzymes is the accurate synthesis of RNA and DNA using DNA as a template. There is an important difference in symmetry. In the case of kinesins and myosins the direction of movement is dictated by the polarity of cytoskeletal filaments. Polymerases are mechanically and structurally more complex in that they perform a variety of different types of movements along the template. RNA polymerase uses double-stranded DNA as a template, which is non-polar structure and in principle permits motion in both directions. In each particular case the direction of movement is determined by the polarity of a promoter particular sequences in DNA where RNAP starts transcription. In the case of DNAP the direction of DNA synthesis is determined by the polarity of replication. Another important difference is the size of individual stepping events. For kinesin the characteristic stepping distance (8 nm) is equal to the periodicity of tubulin monomers along the microtubule (Svoboda and Block, 1994). The corresponding step size for RNA and DNA polymerase is the distance between two base pairs in the DNA double helix, which is 0.34 nm. An important characteristic of motor enzymes is their velocity. Considering the polymerization rate for bacterial RNAP of 40 50 nt s 1 (Uptain and Chamberlin, 1997), the calculated linear velocity will be 14 17 nm s 1. This value is at least 50 times less than that for kinesin. For eukaryotic RNAPs this rate is even lower. In contrast, for phage RNA polymerases and some DNA polymerases the polymerization rate exceeds 500 1000 nt s 1 (170 340 nm s 1), which is comparable to the velocity of kinesin. A significant difference has been observed in the forces generated by polymerases and other motors (Tab. 6.1). The forces of 14 to 25 pN for bacterial RNAP (Wang et al., 1998, Yin et al., 1995) and 34 pN for DNAP (Wuite et al., 2000) are much larger than those for kinesin 6 pN (Meyhofer and Howard, 1995) or myosin 3 to 5 pN (Finer et al., 1994). On the other hand, the efficiency of energy conversion per single step movement is 1.5 to 3 times lower than for kinesin and myosin. Another difference is in the manner in which the chemical energy is provided and consumed. RNA and DNA polymerase motion is driven by the polymerization

6 RNA and DNA Polymerases

reaction and the energy is provided by all four NTPs. Each polymerization step is directly coupled to the forward motion of the enzyme. Exceptions are some exotic cases where upon transcription at a certain sequence, RNAP repeatedly slips back without losing RNA from the active site, thus synthesizing a homopolymeric RNA without advancing forward (Ba et al., 2000, Pal and Luse, 2002, Uptain et al., 1997). Stalled polymerase does not consume or save chemical energy. Kinesin, on the other hand, can consume ATP and store the energy since ATP hydrolysis is not directly coupled to movement. Polymerases, whose activity can be modulated by numerous protein factors, are more chemically complex than other motors (Gelles and Landick, 1998). RNAP functions as the primary target for the regulation of gene expression and responds to the intrinsic signals encoded in DNA sequences such as termination and pausing sites. Our current understanding of the mechanisms of polymerization is mostly based on crystallographic and biochemical kinetic studies coupled with micro-mechanical properties of directly observed movement of single molecules along the template during transcription and DNA synthesis. In this review we will consider the following aspects of DNAP and RNAP functioning as molecular motors: the mechanisms of force generation, experimental approaches to studying polymerase movement and possible molecular mechanisms of polymerase translocation.

6.2

NTP Polymerization Mechanism

RNAP performs NTP polymerization on double helical DNA using one particular strand as a template, while DNAP uses single-stranded DNA. RNAP is an obligatory processive enzyme; each RNA molecule, whose length can be up to 106 nucleotides (nt), is synthesized in its entirety by a single molecule of RNAP moving processively along the template. Once an RNA chain is released by one RNAP molecule, no other polymerase is able to catch and elongate it as the nucleic acid scaffold of the transcription complex is thermodynamically unstable and must be supported in functional configuration by the enzyme. In contrast, DNAPs are much less processive and are able to catch the 3l end of the synthesizing DNA chain released by other DNAPs. Processivity of DNAP can be greatly increased by protein factors. Figure 6.1 shows the schematic structure of a transcription elongation complex (TEC) and DNAP complex with the template and primer. The transcription cycle starts after RNAP occasionally bumps into DNA and slides along the duplex until a promoter is encountered. The recognition and engagement with a promoter requires protein initiation factors which make sequence-specific DNA contacts (Gross et al., 1998). Transition from the initiation to the elongation phase of RNA synthesis is accompanied by relinquishing the sequence-specific contacts and establishing a new type of contact where RNA polymerase does not have a particular affinity to any DNA sequence. This transition

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6.2 NTP Polymerization Mechanism

Figure 6.1. Nucleotide polymerization cycles and schemes of transcription elongation complex (A) and DNA polymerizing complex (B). Schemes I to III represent different functional states of the nucleotide addition cycle. State IV(A) shows the backtracked elongation complex. State IV(B) shows the complex where the 3l end of DNA is caught by the exonuclease site. The nucleic acid scaffold is surrounded by RNAP (A) and consists of downstream (at the right) and upstream (at the left) DNA duplexes,

RNA/DNA hybrid, 5l single stranded RNA, and transcription bubble. For DNAP (B) only downstream ssDNA and upstream DNA duplex constitute the scaffold. The catalytic center is represented by i and i1 sites. The arrows (A, state I) show the formation or disruption of base pairing and indicate the transition to the pre-translocated state II. The arrows (A, state II) indicate the transition in the opposite direction (to state I).


is followed by dramatic stabilization of the complex that makes it highly processive. Nucleic acid protein interactions in TEC are of topological nature and allow free sliding of the enzyme along the template. During elongation RNAP functions in association with the DNA template and RNA product. Within the complex, synthesized RNA forms a short hybrid 8 9 base pairs (bp) long (Korzheva et al., 1998, Nudler et al., 1997) with the transcribed DNA strand, while the other DNA strand is separated from the template. The resulting DNA melting region of 12 13 nt is called the ‘transcription bubble’ (Zaychikov et al., 1995). The configuration of nucleic acids within the bubble region is maintained constant due to interaction with the protein. In the catalytically competent state the 3l RNA terminus occupies the i site of the active center, while the i1 site is vacant for incoming NTP. DNAP slides along a single DNA strand during replication, synthesizing its exact complementary copy which is always base-paired to the template (Fig. 6.1B). The catalytic cycle of a single RNA chain elongation step as shown in Fig. 6.1A I, involves NTP binding (III), phosphodiester bond formation as a result of nucleophilic displacement of the pyrophosphate residue from the NTP molecule by RNA 3l hydroxyl (II), and translocation of a new 3l RNA nucleotide residue from the i1 to the i site to vacate the NTP binding center (I). It is important to stress here that it is not clear yet whether pyrophosphate release precedes or is followed by translocation. The same nucleotide addition cycle is also observed for DNAP (Fig. 6.1B, I III). The majority of the available data suggests that both RNAP and DNAP move along the DNA monotonously, whereby upon each single nucleotide addition step the enzyme advances by 0.34 nm the size of a single nucleotide residue. In the normal elongation complex the 3l RNA end oscillates between the i and i1 sites of the active center (Fig. 6.1, I and II) which is evidenced by the ability of the stalled TEC to resume elongation or undergo processive pyrophosphorolysis upon addition of NTPs or PPi, respectively. Besides the sliding accompanying nucleotide addition steps or pyrophosphorolysis, RNAP is involved in other kinds of movement not connected with catalysis. Thus, the enzyme can occasionally translocate to at least one position downstream of the i1 site to a position i2 (‘hypertranslocated’ state), leaving the 3l RNA end in the i–1 position (Gelles and Landick, 1998, V. Epshtein et al., 2002). Movement in the opposite direction upstream of the i site, known as backtracking (Fig. 6.1A IV), occurs spontaneously at certain sequences and will be considered in Section 6.7. A similar kind of movement, which results in disruption of base pairing of the 3l end of DNA with the template, occurs in the case of DNAP as well, when the 3l end of the DNA product becomes occasionally captured at the exonuclease center of the enzyme, which resides at a distance of 2 nm from the polymerization center (Fig. 6.1B IV; Joyce and Steitz, 1994).

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6.3 Basic Methods used to Study Polymerase Movement during Transcription

6.3

Basic Methods used to Study Polymerase Movement during Transcription

Important properties of transcription and replication complexes that have provided insight into the mechanism of translocation were revealed using two basic approaches: (1) single molecule studies of the influence of mechanical force on polymerase translocation, and (2) classical biochemical approaches that have provided information about the positioning and microscale movement of the enzymes along the template during the nucleotide addition step. A remarkable breakthrough of the past decade was the development of experimental techniques that allowed manipulation of single protein molecules and detection of their movement. Examples of the application of such techniques to transcription studies are the direct observation of linear movement of a single molecule of RNAP relative to DNA during transcription (Davenport et al., 2000, Guthold et al., 1999, Schafer et al., 1991, Wang, 1999) and during a onedimensional diffusional search for a promoter (Guthold et al., 1999, Harada et al., 1999, Kabata et al., 1993), as well as during DNA rotation (Harada et al., 2001). Below we will present a brief description of these methods and discuss their validity and resolution. 6.3.1

The Tethered Particle Motion Approach

The tethered particle motion approach is based on the visualization by light microscopy of Brownian motion of a plastic bead or colloidal gold particle attached to the upstream end of the DNA template (Schafer et al., 1991). The extent of Brownian motion is used to determine the length of the DNA between the bead and RNAP during transcript elongation (Yin et al., 1994). This method is arguably the least perturbational; it neither exerts force on DNA nor significantly alters its conformation (Gelles and Landick, 1998). Since bead diffusion is more rapid than the mean velocity of DNA translocation in a polymerase catalytic cycle, tethering of the bead should not significantly affect the translocation reaction in the case when the bead is attached further from the DNA molecule than the DNA persistence length. (The persistence length of Z 150 bp is the characteristic distance over which the orientation of the double helical axis is changed by random bending.) 6.3.2

The Surface Force Microscopy Technique

The surface force microscopy (SFM) method offers the possibility of high-resolution imaging of single macromolecules and their complexes attached to flat surfaces. Unlike conventional high-resolution electron microscopy, which can operate only in vacuo, SFM can be performed in any environment air or liquid (Drake


et al., 1989) at natural physiological conditions of cells and biomolecules. This method also allows the forces of intramolecular and intermolecular interactions in the pN range to be measured. Transcription complexes of RNAP have been imaged using SFM both in air (Rees et al., 1993) and fluid (Guthold et al., 1994). Continuing development of this technique led to the monitoring of the movement of RNAP during RNA synthesis (Kasas et al., 1997). This was achieved by taking advantage of TECs immobilized on mica surfaces. While RNAP is always bound tightly to this surface, the binding of DNA to mica can be regulated by the concentration of Zn2 in the medium. Repetitive changing of the Zn2 concentration using a flow cell ‘turned on’ RNA synthesis for a while and then ‘froze’ it by increasing the binding force of the DNA to the surface, thus making it suitable for visualization. Alternation of these steps allowed time-lapse imaging of the transcription complex. This approach enables sequential imaging of individual complexes moving at rates of 0.5 2 bases s 1. The resolution ability of this method is restricted by the acquisition time, which is 30 60 s, but is believed to be improved. 6.3.3

The Optical Tweezer Method

Figure 6.2A shows the design of the experiment which allowed to measure the force exerted by a single RNAP molecule during transcription (Wang et al., 1998, Yin et al., 1995). The defined transcription elongation complex was assembled in solution by mixing RNAP, an incomplete set of substrates and DNA. This complex was irreversibly absorbed through RNAP on the glass surface of the microscope flow cell. Polystyrene beads 0.5 mm in diameter were attached to DNA termini in the direction of RNAP movement during transcription. The complex was identified visually by movement of the bead using light microscopy. The microscope was equipped with an optical trapping interferometer (‘optical tweezers’) which could exert a calibrated force on a bead while simultaneously measuring its displacement. Using ‘tweezers’, the bead was moved to the optimal position, slightly above the glass surface in the trap center. During RNA synthesis immobilized RNAP threads DNA through itself causing movement of the bead away from the trap center, which develops tension in the DNA between the bead and the polymerase. The optical trap acts as a linear spring attached to the reference frame, therefore during transcription the bead adopts the position where the force of the trap is balanced by the force exerted by RNAP (Fig. 6.2B). Thus the force generated by RNAP can be calculated directly by measuring of the displacement of the bead.

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Techniques for studying polymerase movement during transcription. (A) Experimental design used by Wang et al. (1998) and Yin et al. (1995) to observe single RNAP molecules pulling a DNA template. RNA polymerase was fixed to the surface after ternary complex formation. The beads were attached to the transcriptionally downstream end of DNA and then optically trapped. The bead displacement (x) was observed as time progress from t1 to t3. (A,C) Reprinted with permission from Science (Wang, M. D. et al., Science 282: 902–907, Figure 1, A and Figure 2, C). Copyright 1998 American Association for the Advancement of Science. (B) The mechanical equivalent of the experimental geometry shown in (A) (Yin et al., 1995). The optical trap acts as a spring of stiffness atrap attached to a stationary reference frame and exerts a force Ftrap on the bead. The bead adopts a position where Ftrap is balanced by the force Ftc exerted by the polymerase, acting through DNA. Polymerase, attached to another reference frame, and DNA act as a spring of stiffness atc. By calibrating the optical trap stiffness and determining the displacement of the bead, it is possible to measure Ftrap and thus Ftc. (B) Reprinted with permission from Science (Yin, H. et al., Science 270: 1653–1657, Figure 6.2.

Figure 1, C). Copyright 1995 American Association for the Advancement of Science. (C) The transcript length at different loads applied to the ternary complex (Wang et al., 1998). The bottom trace shows the movement of DNA through RNAP extracted from the bead position data. This movement is interrupted by transcriptional pauses (marked by arrows). After algorithmic deletion of pauses, the records show the elongation rate to be remarkably constant up to external loads nearly sufficient to stall the complex (at 23 pN). (D) Observation of DNA rotation by the RNA polymerase (Harada et al., 2001). RNA polymerase is attached to a glass surface. The magnetic bead at the downstream end of the DNA was pulled upwards by the disk-shaped neodymium magnet. Daughter fluorescent beads served as markers of rotation. (D) Reprinted with permission from Nature (Harada, Y. et al., Nature 409: 113–115, Figure 1, A). Copyright 2001 Macmillan Magazines Ltd. (E) The chemical footprinting approach aimed to detect the position of bound protein on the DNA. DNA labeled at the end undergoes chemical degradation under single hit conditions. After separation of the products the ‘gap’ indicates the exact position of the protein.


6.3.4

Method for Visualization of DNA Rotation during Transcription

During transcription RNAP threads the right-handed double helix of DNA, which causes clockwise DNA rotation. An approach has been suggested, which can visualize relative rotation between RNAP and DNA directly and in real-time using an optical microscopy technique based on the tethered particle method (Harada et al., 2001). The design of the experiment is shown in Fig. 6.2D. The RNAP in the complex with template DNA and RNA product was stalled by substrate deprivation at a particular position on the template and then attached to a glass surface. A magnetic bead of diameter 850 nm coated with streptavidin was attached to the downstream end of the DNA where terminal nucleotide residues were biotinylated. Then, all four NTPs were added to allow further transcription. The rotation was observed directly by labeling the bead with smaller fluorescent beads. The bead was pulled upward at Z 0.1 pN with a magnet in order to confine the rotation in a horizontal plane and to restrain the DNA from supercoiling, which would interfere with torque transmission to the end bead. At the conditions used at least Z180 consecutive revolutions have been observed, suggesting that thousands of base pairs can be transcribed without extensive rotational slippage. A magnetic field of a different polarity could be applied to the bead, thus facilitating or impeding the rotation caused by RNAP. The dependence of the rotation rate on the polarity and intensity of the magnetic field can provide important information about the transcription mechanism. The real-time observation of rotation opens up the possibility of resolving individual transcriptional steps. Transcription of one base will produce a linear translocation of 0.34 nm, which is extremely difficult to resolve. However, the accompanying rotation is as much as 35h and is easily detectable. RNAP reaches a maximal rotation rate of Z 0.2 revolutions s 1. To rotate both DNA and a bead of diameter 850 nm in bulk water at this speed, a torque of about 5 pN nm is required. This value is much less than that of the conventional rotary motor F1-ATPase (40 pN nm) (Yasuda et al., 1998; see also Chapter 5 by Noji). Thus far, single-molecule transcription methods are able to measure DNA translocation with a precision of a few tens of nanometers roughly 100 bp. It is believed that precision in future experiments will be substantially increased by improved instrumentation and modifications to experimental designs. Despite the fact that the experiments use preparations of surface-immobilized RNAP, kinetic properties of transcription (e. g. elongation and rotation rates, transcription pause lifetimes) are identical or nearly identical to those inferred from conventional solution studies (Harada et al., 2001, Schafer et al., 1991). Thus, results from two classes of studies can be combined to achieve a coherent picture of the biochemical and mechanical aspects of polymerase function.

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6.3.5

Footprinting Approach

This approach is based on the use of agents which cleave the DNA chemically or enzymatically. That part of the DNA which is protected by the bound protein is inaccessible for cleavage. In the chemical version of the method cleavage conditions are used where less than one ‘hit’ occurs per DNA chain (Fig. 6.2E). Analyzing the degradation products by gel electrophoresis in denaturing conditions allows the determination of the exact position of the protein relative to the labeled DNA terminus. Another version of this approach is based on the ability of exonuclease III to digest one DNA strand in the DNA duplex starting from one end. Cleavage stops when the exonuclease encounters the protein in the complex. The position where the exonuclease has made contact with the protein and been stopped, can be easily mapped by analysis of the digestion products using reference degradation products as a sequence marker. 6.3.6

Single Molecule Assay for DNA Polymerase

Single molecule studies allow observation of the movement of the T7 DNAP along DNA and the effect of template stress on both polymerizing and exonuclease activity of the enzyme. In this study DNA consisting of single- and double-stranded segments was used (Wuite et al., 2000). To measure polymerase activity the optical-trap described above was applied (Fig. 6.3A). Because single-stranded (ss) and double-stranded (ds) DNAs differ in length at any given tension (Fig. 6.3B), interconversion between these two forms changes the tension of the molecule if the end-to-end distance of the template is held constant. Alternatively, the end-to-end distance of the molecule changes if the tension is held constant. The end-to-end distance of ssDNA (Fig. 6.3B) is shorter than that of dsDNA for tensions below 6.5 pN (‘crossover point’) because ssDNA, despite having about twice the contour length of dsDNA, is more retractile due to its greater flexibility. Above 6.5 pN, however, contour length predominates over entropy and ssDNA is longer than dsDNA. The force extension curve of a molecule that is partly ssDNA and partly dsDNA can be fit to a linear combination of ssDNA and dsDNA stretching curves. Under constant end-to-end distance, DNA synthesis causes an increase in template stress which in turn reduces the rate of polymerization (Fig. 6.3C, top curve). Treatment of the time velocity curve (Fig. 6.3C, bottom curve) reveals ‘bursts’ of activity consistent with the loading of the enzyme molecule onto the 3l end of the growing chain, replicating processively and falling off. Polymerization stops when the template tension exceeds 34 pN.


Figure 6.3. Optical trap set-up for observation of DNA synthesis. (A) DNA fragment containing both single and double strand segments is attached between two beads, one held on the tip of a glass pipette, the other is in an optical trap. The end-to-end length of the DNA is obtained by video imaging of the bead positions, and the force (F) was measured using the change in light momentum, which exits the dual-beam trap. (B) Force-extension data for

dsDNA and ssDNA. (C) Replication of a ssDNA template under constant tension. Upper curve, conversion to dsDNA plotted as a fraction of ssDNA left in the template versus time. Lower curve, polymerization rate obtained by differentiating the upper curve. (A, B, C) Reprinted with permission from Nature (Wuite, G. et al., Nature 404: 103–106, Figure 1A, B and Figure 2A). Copyright 2000 Macmillan Magazines Ltd.

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6.4 Mechanism of Force Generation for RNAP and DNAP

6.4

Mechanism of Force Generation for RNAP and DNAP

What causes directional movement of RNAP and DNAP along DNA during polymerization reaction? And how tremendous force which, if related to the mass of the motor, exceeds that for contemporary jets and rockets by factor 107–108 (Marden and Allen, 2002) is generated? The force exerted by RNA polymerase is 109 times greater than the weight of the enzyme itself and 104 times greater than the weight of bead used in “optical tweezers” experiment (see above). The size of the bead that could be “lifted” by single RNAP molecule is big enough to be seen by naked eye. This is a remarkable example of how a large force links “micro” and “macro” worlds. Depending on the transcribed sequence the translocation, followed by each nucleotide addition step could be energy-consuming process that has to be balanced. The energy required for translocation can be supplied by polymerization reaction itself. Conventional wisdom suggests that NTPs could be the source of required energy. Two different sources of energy could be potentially involved. One is the binding energy of NTP to the active center, which is in range of 8 to 9 kcal/mol (Guajardo and Sousa, 1997). Another kind of energy in the range of 7.2 kcal/mol is liberating upon the chemical reaction of phosphodiester bond formation and release of pyrophosphate (Yager and von Hippel, 1987). Two principal mechanisms, which consider either first or second source of energy as a principal contributor to translocation, have been suggested. One hypothesis is that RNAP operates by a “Brownian Ratchet” mechanism, proposed previously for other motors (Cordova et al., 1992). According to this mechanism the enzyme rapidly oscillates between neighboring template positions (pre- and posttranslocated states), whereby 3lRNA end occupies either i1 or i site of the active center respectively (Figure 1, I and II). At a certain moment of time NTP enters the active center (when the i1 site is vacant) and locks the polymerase in posttranslocated (catalytically competent) state, so that the enzyme cannot slide back to pretranslocated position (III). In this case the NTP acts as a “pawl” of the ratchet preventing backward movement of the enzyme and reentering of 3lRNA end to i1 site. After incorporation into RNA of monophosphate residue of NTP the process repeats thus causing step-by-step directional movement of the enzyme along DNA. Another mechanism for translocation that could be conceived for RNAP is “Power Stroke”, in which the enzyme is rigidly coupled to the template and cannot slide back and forth spontaneously. In this case the enzyme is pushed forward by conformational change within it, caused by some external influence. For instance, in RNA polymerase this conformational change can be caused by and energetically coupled with the reaction of phosphodiester bond formation by analogy with those motors where ATP hydrolysis induces structural rearrangements. The question about how polymerizing enzymes actually move, by using either of these mechanisms or by some intermediate mechanism requires adequate research tools and the knowledge about different aspects of structure and functioning of elongation and replication complexes.


Some properties of RNAP elongation and DNAP polymerizing complexes allow to draw immediate conclusion about the mobility of enzymes. The mentioned above efficient pyrophosphorolytic cleavage of RNA and DNA in the complexes deprived of the substrates, which represents the reaction truly reverse to NTPs polymerization suggests that 3lRNA terminus more or less freely oscillates between i and i1 sites. It could be argued, that pyrophosphate itself can induce reverse translocation of 3lRNA terminus to i1 site, otherwise fixed in i site. However, the experiments of Fe2 mediated affinity cleavage of RNA in TEC indicate that translocation equilibrium exists even in the absence of pyrophosphate (Epshtein et al., 2002). This fact is indicative for the random motion of polymerases within the complexes and shows that translocation step is not strictly energetically coupled to polymerization. As it is seen from Figure 6.1 (the translocation from stage II to I) one-step advancement of RNAP is accompanied by melting of 1 bp of downstream DNA duplex and simultaneous restoring of basepairing at the upstream edge of transcription bubble. At the same time this translocation leads to shortening of RNA/DNA hybrid by one bp. Since free energy (DG) for the formation of a single base pair ranges from 0.9 to 3.4 kcal/mol (Stryer, 1995), the translocation could be either favored or impeded depending on value of DG which in the extreme cases constitutes –1.6 and 5.9 kcal/mol, respectively. The corresponding constants for translocation equilibrium (2x101 – 5x10 –5) calculated from the Arrenius equation Keq e DG/RT show that in TEC RNAP predominantly exists in pretranslocated state, and that the relative fraction of RNAP, which can bind NTP at the active site is only fractions of percent. It should be noted that translocation is not likely to be favored by enhanced binding of 3lRNA terminus at i site as soon as the affinity of nucleoside-5l-monophosphates (NMPs) (which can be viewed as a model of 3lterminal RNA residue) to this site is low, as can be judged by their Km values in abortive initiation reaction (Szafranski et al., 1985). A few important observations relevant to the mechanism of RNAP translocation were made using optical tweezers studies. First, it was found that during RNA synthesis RNAP is able to move forward against the significant applied force. It was found also, that the rate of polymerization was almost independent on the applied force up to the forces closed to stall (Figure 6.2C). Extremely important set of experiments with the above described technique allowed to relate the free energy of the chemical reaction of phosphodiester bond formation to the force generated by RNAP (Yin et al., 1995). The free energy liberated during chemical reaction of single nucleotide addition to RNA is connected with concentrations of NTP and PPi by the equation: DG DGo RTln[NTP]/[PP i]. The stall forces were measured for TEC in the presence of NTP and the different concentrations of PPi (1 mM – 1 mM), which ranged calculated DG values from 7.2 to 3.1kcal/mol. Surprisingly, the measured stall forces were nearly identical under these conditions. On the other hand the force generated by molecular motor (F) is a linear function of DG (Fisher and Kolomensky, 1999): F DG/a, where a is a distance between the neighboring base pairs in DNA helix (0.34nm). Therefore the conclusion from this experiment

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was that the energy available from the polymerization reaction itself is not used in force-generating step. At this point it is appropriate to consider the results of the elegant study performed on RNAP from phage T7 (Guajardo et al., 1998), which revealed the particular step in nucleotide addition responsible for translocation. First, defined elongation complexes were obtained using incomplete set of substrates. Then the system was supplemented with NTP analog, which was able to bind normally to the active center and form a base pair with the template, but was unable to incorporate into RNA. According to footprinting data (Figure 6.4), the addition of NTP analog resulted in translocation of the enzyme in the direction of RNA synthesis by a distance equal to the size of one nucleotide residue. The simplest explanation of these results is provided by mentioned above “Brownian ratchet” mechanism. As it was pointed above (see also Huang and Sousa, 2000), in TEC translocational equilibrium is strongly shifted to pretranslocated state where 3lRNA end occupies i1 site of the catalytic center (Figure 6.1, II) that is energetically more favorable. On the other hand NTP substrate can successfully compete with 3lRNA end for the binding to the i1 site of the catalytic center due to high affinity. Indeed, the average binding energy calculated from the corresponding Km values is in the range from 8 to 9 kcal/mol which is quite significant to displace 3lRNA end to i site and thus provide the translocation. According to this model NTP binding to the active center is the force-generating step in transcription. From this point of view it is clear why the measured stall force was not influenced by PPi, simply because at used PPi concentrations NTP binding is not affected. There could be one obvious way to test this hypothesis in single molecule manipulation assay whereby instead of NTP its analogs with impaired affinity to the active center but unaffected DGhydr would be used. Lowered free binding energy for those analogs should diminish corresponding generated force. In another version of this experiment mutant RNAP with decreased Km for NTP may be used as well. The kinetic models of RNAP movement consistent with the results of single molecule studies have been developed (Wang, 1998B, Julicher and Bruinsma, 1998) So, available data presented above strongly support the notion that NTP binding to the active center is the force-generating step in transcription. Subsequent catalytic reaction of phosphodiester bond formation and pyrophosphate release from thermodynamic point of view has a very important consequence: it eliminates the “pawl” of the ratchet. It is evidenced by the fact that the major fraction of the binding energy of original NTP (which acts as a “pawl” of the ratchet) is associated with its pyrophosphate moiety (Guajardo and Sousa, 1997). This mechanism also explains why the force-generating step (translocation of the enzyme caused by NTP binding) is not load-dependent over most of the force range. The mechanical work, which is done by the applied force against the moving polymerase at the values close to stall (25 pN) can be calculated as: A Fs a and is equal to about 1 kcal/mol, which is very close to the energy of the weakest base pair. The disruption of even one base pair at the 3l end of the hybrid facilitates the backtracking process, the movement of the enzyme in the reverse to the synthesis direction (see chapter 6.7). The applied stall force stops polymerization as soon as it


Figure 6.4. Translocation of RNAP induced by NTP. (A) Scheme of the experiment. Structure of transcriptional bubble associated with the enzyme is presented. The position of the enzyme relative to DNA in the absence (top) and presence of NTP (bottom) is indicated by vertical dashed lines. The direction of exonuclease digestion of the template DNA strand in the complexes and position of radioactive label are shown. (B) Electrophoretic analysis of the radioactive exonuclease degradation products

in different elongation complexes in the presence or absence of NTP. Horizontal numbers indicate the length of RNA in the complex. The vertical numbers indicate the size of DNA degradation products. Continuous diagonal lines link the position of upstream edge of RNAP footprint with corresponding degradation products. (B) Reprinted from JMB, 303: 347–358, J. Huang and R. Sousa, Figure 1A, Copyright 2000, by permission of the publisher Academic Press an imprint of Elsevier Science.

induces and acts in the direction of backtracking. The binding of incoming NTP in the i1 site in the case of backtracking is impossible as soon as this site and the way to the active center are occupied by RNA chain. The stronger base pairing in the hybrid at the active center, which can be observed on poly-dG or poly-dC templates, will certainly increase the stall force, that can be checked experimentally. For DNAP passive sliding mechanism was suggested on the basis of kinetic evidence. The model proposed the existence of two conformational states of the enzyme. In “open” conformation the enzyme is able to slide along the template,

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while in the “closed” conformation, induced by NTP binding the mobility is restricted (Johnson, 1993). This model has been recently supported by direct crystallographic data, which demonstrated that NTP binding causes a conformational changes, which could restrict the lateral mobility of DNAP on the template (Doublie, et al., 1998a,b). Single molecule studies revealed that DNAP could work against significant template tension equal to 34 pN (Wuite et al, 2000). Enhanced sensitivity of polymerization rate to the applied force suggested that the rate-limiting step was affected. The replication rate of T7 DNA polymerase is about 100 bases/sec at the lowest tension of the template, it is becoming 200 bases/sec at about 6 pN, further increase in tension causes the rate to decrease until polymerization stalled. Considering crystallographic data (Doublie, 1998b) the authors explain this phenomenon by the effect of template tension on closing the “fingers” which sets up the enzyme for catalysis or by disruption the interactions of template residues at the active center. Noteworthy, when template tension is increased above 40 pN, a fast exonucleolysis is initiated, which is becoming force independent above 42 pN and is about 100 times faster than observed at zero tension. It is known that the exo rate is limited by the escape of 3l end from the polymerization site and it is thought that several base pairs must be melted to allow binding of DNA 3l end to the exonuclease site. Forces greater than 40 pN are also known to promote fraying in dsDNA. Presumably, the template tension increases the escape rate from or decreases the binding rate to the polymerization site due to facilitated melting of the growing 3l end. Based on the experiments shown in Figure 6.3 (Wuite et al, 2000), we should specify here that as soon as completely stretched ssDNA is twice longer than the same segment in double helical structure, the external force can pull out the template residue from the active site to a maximal distance a, which is about 0.34 nm. The stall force (34 pN) calculated as A Fsa is equal to 1.4 kcal/mol and therefore can disrupt the weak base pairs in dsDNA.

6.5

Molecular Model for RNAP Translocation

Understanding the details of polymerases as molecular machines will definitely come from a combination of biochemical, biophysical and structural studies. The recent success in determination of the crystal structure of polymerases and their functional complexes with DNA led to the development of some working models to explain the translocational mechanism at the molecular level. A structural model of the RNAP elongation complex is shown in Fig. 6.5. According to the model, the downstream DNA duplex is accommodated in a deep ‘trough’ formed by the b’ subunit and topped by a domain of the b subunit known as the ‘roof’ (Korzheva et al., 2000). The RNA/DNA hybrid is enclosed in the channel formed by the largest subunits of the enzyme between the catalytic Mg2 ion at its downstream end and the ‘rudder’ loop of the b’ subunit at its upstream end. Cata-


Views of the TEC model. The model consists of three components: (1) the Taq core RNAP crystal structure shown as color-coded molecular surfaces (b, cyan; b’, pink; a, white; v, dark gray, catalytic Mg2, magenta sphere), (2) the DNA template (template strand, red; non-template strand, yellow) and (3) the RNA transcript (gold) and incoming nucleotide substrate (green). The direction of transcription is Figure 6.5.

indicated. (A) View perpendicular to the main active-site channel, which runs roughly horizontal. Parts of the protein structure are colored (b flap in green, b’ rudder in rose). (B) View down the secondary channel, showing the path for diffusion of the incoming nucleotide substrate (green) into the active site or 3l RNA exit path upon backtracking. The catalytic Mg2 ion is just visible to the left of nucleotide substrate.

lytic Mg2 also marks the beginning of the ‘secondary channel’ (Fig. 6.5B), which has been proposed to supply the substrates to the active center and to accommodate the 3l-terminal segment of ssRNA released upon RNAP backtracking. The secondary channel branches off the main channel by the ‘F-bridge’, which is part of the active center (see also Fig. 6.6). After separation from the DNA template, the 5l segment of ssRNA exits through the channel underneath the ‘flap’, a long, flexible loop of the b subunit. The downstream portion of the non-template DNA strand within the bubble, resides in the channel between two structural domains of the b subunit, while the upstream portion is accessible to the solvent. The available data do not allow the precise positioning of the upstream DNA duplex, but according to footprinting studies this DNA may be flexible in the TEC. Crystallographic imaging of yeast polymerase (Cramer et al., 2001) revealed essentially the same structure of the active site as in the TEC model described above. The only prominent difference was the conformation of the F-bridge. In the yeast structure it is straight, while in Thermus aquaticus (Taq) it is bent. As straight and bent conformations of the F-bridge are structurally compatible with pre- and post-translocated states of RNAP, respectively, Kornberg and co-workers (Gnatt et al., 2001) suggested that a change from a straight to a bent conformation is involved in translocation. This suggestion has been further supported by the demonstration that these two conformations of the F-bridge do really exist in functional TECs (Epshtein et al., 2002). Figure 6.6 shows the presumed mechanism for RNAP translocation during a single nucleotide addition cycle. In it a principal

169

170

6.5 Molecular Model for RNAP Translocation

Figure 6.6. Transcription cycle and the proposed translocation mechanism. RNA (pink) base-paired to DNA template (yellow, with bases 1, 2 and 3 numbered) translocates relative to the active center (symbolized by Mg2 flanked by the i and i1 sites). The b’ subunit Fbridge (green), which is supported by the helices of the G-loop (white), separates the main channel for the DNA duplex (yellow: purple) from the secondary channel for the nucleotides, backtracking or abortive RNA release. The evolutionarily conserved crucial amino acids of the b’ subunit are indicated: Lys789 (red), Met932 (blue), Thr790 and Ala791 (cyan) (the numbering is that for E. coli). (A) The transcription

complex before phosphodiester bond formation. The F-bridge is relaxed. (B) The pretranslocation stage. The pink arrow represents the trajectory for RNA during backtracking or abortive release. (C) Translocation. The F-bridge bends causing Lys789 to enter the i1 site. The bending is induced by Met932 swapping contacts with Lys789. Side chains 790 and 791 clash with the DNA base in the i1 site inducing its translocation together with the rest of the RNA/DNA scaffold of the TEC. (D) Relaxation. The F-bridge returns to the straight helix conformation vacating the i1 site for the next nucleotide (pink arrow) or for reverse translocation.

structural element that is responsible for translocation is represented by the Fbridge which is in close contact with G and G’ helixes. In the straight conformation of the F-helix (Fig. 6.6A) the active site is vacant for NTP and incorporates the incoming substrate into RNA (Fig. 6.6B). During subsequent structural rearrangements intrusion of the Met932 residue of the b’ subunit in between the G- and F-helixes results in the bending of the F-helix and the flipping-out of Lys789, Ser791, and Ala792 residues towards the active center. This rearrangement is coupled with translocation of RNAP (Fig. 6.6C) because the flipped residues of the helix collide with the edge of the RNA/DNA hybrid. It should be noted, though, that this model does not assume a strict coupling between the switch of the Fbridge and translocation of RNA. Indeed, for the nucleotide substrate to enter the active site the F-bridge must be in a straight conformation without any reverse


translocation of RNA (Fig. 6.6D). This sequence of transitions has been termed ‘swing-gate mechanism’. Upon the switch the Lys789 residue swings in and out of the active center, thus either making it accessible for, or blocking out, the substrate. It should be emphasized that relaxation of the F-bridge not only allows nucleotide entry into the i1 site but also permits the establishment of the base pair (Fig. 6.6D). In fact, flipping of the DNA base into the active center should block both back translocation of RNA and bending of the F-helix, thus keeping the active center in a catalytically competent state. It is not clear yet whether this transition of the F-bridge is triggered by the catalytic reaction; it is equally unclear what its actual role in NTP polymerization is. The crucial role of those transitions for the functioning of the enzyme is supported by the inhibition of RNAP activity by aamanitin and streptolydigin which are believed to freeze the F-helix in straight or bent conformations, respectively (Bushnell et al., 2002, V. Epshtein, et al., 2002).

6.6

Possible Utilization of the Energy Released upon NTP Cleavage

What is the energy liberated during the catalytic reaction used for? Obviously the utilization of energy-rich compounds such as NTPs is necessary for efficient RNA synthesis. The favorable DG of the polymerization reaction makes it practically irreversible. The fact that this energy is not used for force generation solely means that it does not contribute to a limiting step of the enzyme’s translocation. However, a mechanism could be envisioned in which this energy is used for translocation in some specific cases. The basis for the possible existence of such a mechanism comes from the fact that the limiting step in elongation is pyrophosphate release (Erie et al., 1992), which means that after phosphodiester bond formation, pyrophosphate is retained by the active center. This should cause significant electrostatic repulsion between the bound pyrophosphate and the negatively charged phosphate at the newly formed 3l RNA end. This repulsion can act as a ‘power stroke’ and drive translocation of RNAP. This process would be energetically justified as the large fraction of the energy that is liberated during NTP decay originates from electrostatic repulsion of negatively charged phosphates of the triphosphate chain (Stryer, 1995). For translocation to occur, the vector of the repulsion force must be collinear with the direction of translocation, which imposes strict constraints on the possible arrangement of the 3l RNA end and the leaving pyrophosphate. Indeed, the SN2 mechanism of pyrophosphate displacement for the catalytic reaction (Armstrong et al., 1979) dictates the position of this moiety consistent with this mechanism of translocation (Fig. 6.7). As seen in the co-crystal structure of T7 DNAP with template, primer and substrate (Doublie et al., 1998b), the pyrophosphate residue of NTP at the catalytic center is secured in the pocket and therefore could withstand the electrostatic repulsion required for successful translocation. Under normal conditions such translocation could not contribute to the directional work performed by the enzyme, because of the tendency to relocate back to the pre-translocated state before the next substrate can

171

172

6.7 Single-Molecule Studies and Molecular Mechanisms of Transcription Pausing and Arrest

Putative mechanism of RNAP translocation caused by electrostatic repulsion between the negatively charged phosphate of the RNA 3l terminus at the i site and pyrophosphate secured at the i1 site of the catalytic center. The positions of NTP substrate in Figure 6.7.

the active center and Mg2 ions were modeled into the RNAP II crystal structure (Gnatt et al., 2001) by analogy with DNAP (Doublie et al., 1998). Conserved Asp residues of the b’ subunit holding Mg ions are indicated according to the E. coli numeration.

bind at the i1 site of the active center and rectify the movement. Indeed, the estimated relaxation time is about 10 6 s (Wang et al., 1998). However, this process may be an efficient means for driving over the roadblocks represented by DNA-bound proteins. When the polymerase collides with a roadblock the ‘power stroke’ exerted according to this mechanism, can push off an obstacle, causing its dissociation from DNA. The ‘Brownian ratchet’ mechanism simply might not work in such cases where the translocation is blocked by the obstacle and the active center is simply not vacant for the NTP. An obvious way to verify this hypothesis would be to check the efficiency of polymerase passage through the roadblocks at different PPi concentrations. In this case a gradual increase of PPi concentration would cause a progressive reduction of the generated force and the efficiency of passage.

6.7

Single-Molecule Studies and Molecular Mechanisms of Transcription Pausing and Arrest

A remarkable property of RNAP motion during transcription is its highly variable rate (Fig. 6.2C), which can range from 0 to 50 nucleotides per second. Stops or pause sites can interrupt continuous movement in many positions. These sites are very important for regulation at the elongation stage and may precede other elongation events such as backtracking, arrest and termination (Artsimovich and Landick, 1998, Landick, 1997, 1999). Conventional biochemical experiments indicate that during elongation, the 3l end of the RNA can be displaced from the enzyme’s active center via disrupted RNA/DNA base pairing (Fig. 6.1, IV) inducing different stages of backtracking


from a pre-translocated TEC to a complex that has backtracked many base pairs along the template in the upstream direction (Komissarova and Kashlev, 1997a,b, Landick, 1999, Nudler et al., 1997, Palangat et al., 1998). Because arrest occurs at or near pause sites, it is possible that these processes are related. A polymerase transcribing faster is more likely to resist pausing and arrest. The simplest mechanistic explanation for backtracking lies in the energetics of base-pairing interactions within the region of the transcription bubble and the RNA/DNA hybrid. Nucleic acid protein interactions do not seem to contribute to this process since they are not sequence specific and allow free sliding. The elemental act of transcription is accompanied by the breaking of base pairs at the downstream edge of the transcription bubble and at the upstream edge of the RNA/DNA hybrid, as well as the establishment of base pairs at the upstream edge of the bubble and the downstream edge of the hybrid. As the base-pairing energy ranges from -0.9 to -3.4 kcal mol 1 for different bases, at heterogeneous templates the forward translocation can be either favored or impeded depending on the particular sequence transcribing. The impedance of translocation would result in the tendency of RNAP to lose 3l RNA from the active center and to backtrack to a register where the base-paring interactions are most favorable (Fig. 6.1, IV). This process could be reversible (if the backtracking proceeds a moderate distance) or irreversible (if RNAP slides back too far). Reversible backtracking should cause a more or less continuous pause in RNA synthesis while irreversible backtracking leads to arrest. Single-molecule studies (Wang et al., 1998, Yin et al., 1995) support this view. As mentioned previously, large applied forces stopped polymerization. After the stall, followed by force reduction, only a small fraction of RNAP molecules was able to resume RNA synthesis, often with significant lag ranging from 0 to 30 s. Notably, backtracking can occur not only in substrate-deprived complexes but also in the course of normal transcription. It is clear that the applied force facilitates this process as it acts in the direction of backtracking. While passing ‘normal’ sequences RNAP can resist the force, but abruptly stops when encountering such backtracking sites. RNAP can probably generate more significant forces on the template lacking backtracking sites. Indeed, for some complexes the values of stall forces were much greater than average. They likely were detected on the DNA segments occasionally missing unfavorable sites. It should be pointed out that the maximal possible force generated by polymerases (140 160 pN), when calculated from the equation f DG/a (Fisher and Kolomensky, 1999), is much higher than the value actually observed. The most likely reason for that is the propensity of RNAP to backtrack, which seems to be predetermined by the details of construction of this molecular machine. The 3l RNA end is poorly supported in the active center and can easily thread through the ‘secondary channel’ (Fig. 6.5B) during backtracking. This channel represents a ‘pore’ about 3 nm in length and is believed to supply the substrates to the active center. The possibility of backtracking is reduced at high NTP concentrations since in this case the complementary substrate taps that pore upon binding at the active center, thus keeping the RNA in place. Even at high concentrations of NTPs the

173

174

6.8 Concluding Remarks

active center occasionally becomes temporarily vacant (since substrate selection is required). This leaves the possibility for backtracking, the latter being greatly accelerated by an applied external force. Detailed experiments on the influence of the applied force on transcription pausing and arrest revealed that a paused state is a kinetic intermediate between forward elongation and an arrested state. A moderate external force (I9 pN) did not affect the fraction of molecules which leave the main elongation pathway to pause. Neither did this force affect the efficiency of pause and arrest. However, in the high-force regime (9 to 15 pN), the incidence of arrest during a pause nearly tripled for most of the pause positions as compared to the low-force regime (Davenport et al., 2000). Higher external forces accelerate the rate at which molecules that are already paused can convert into the arrested state, perhaps by inducing backtracking of the complex, or by decreasing the rate at which these paused molecules can escape the pause to re-enter elongation (Wang et al., 1998). These studies also showed that RNAP molecules possess different intrinsic transcription rates and different propensities to pause and stop (Davenport et al., 2000). The switching between alternative states of elongation may be the basis for yet another level of transcriptional control and regulation in both bacteria and eukaryotes.

6.8

Concluding Remarks

What is the mechanism of energy conversion that polymerases use to generate such a great force? Let us consider the amount of RNAP equal to one mole (500 kg), which is comparable to the mass of a real motor. Such amount of the enzyme would produce a force of 1013 N (Z16 pN x 6 x 1023), which is hard to comprehend. It would burn 350 kcal of energy per second (Z7 kcal x 50 n/sec), which is equal to consumption of about 50 gallons of gas per hour. At the same time this “motor” would move at the speed undetectable by naked eye – just 1 meter a year! (as soon as the velocity is still equal to that for a single RNAP molecule). The power of this motor calculated as W = fappv, (where fapp is a load close to stall and v is a linear velocity) would be just about 40 kW (1013 N x 17 x 10 –9 m/sec) which is equal to 54 horse powers. So, it can be concluded that the mystery of high force generation is high-energy consumption combined with extremely slow motion. In this way polymerase could be compared to a motor with a very high transmission ratio used, for instance, in road pavers. Another distinguishing feature of polymerases as motors is their high regulation, which particularly for RNAP, is not limited just by the availability of ‘fuel’ in the medium. The rate of RNAP elongation is controlled by the transcribed DNA sequence as well as by protein factors which upon binding to the enzyme, modulate the rate of polymerization thus affecting the catalytic activity. The significance of the unusually high force generation for function of the polymerase remains unsolved. For RNA polymerase the most probable use of that force


is to drive over the roadblocks. DNA polymerases, on the other hand, are assisted during replication by a ‘front squad’ of accessory proteins such as helicases that unwind the double helix and deal with the roadblocks, thus enabling the enzyme to do its job, which is accurate DNA synthesis. Probably in this case ‘strong muscles’ are of no use, but affect the energetics of the polymerization reaction. However, a recent study (Delagoutte and von Hippel, 2001) revealed that helicase movement along DNA, and DNA unwinding, are greatly accelerated by the trailing DNA polymerase, which replicates the leading DNA strand. This finding invites the exciting idea that the force exerted by DNAP during polymerization pushes the helicase forward, thus increasing the rate of replication. In principle, the force generated by DNAP upon DNA synthesis could itself promote efficient DNA unwinding as in the case of RNAP. However, in the replication process polymerization and helicase functions are separated.

Acknowledgements

This work was supported by NIH grants GM49242 and GM30717 to Alex Goldfarb.

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Cramer, P., Bushnell, D. A., Kornberg, R.,D. 2001. Structural basis of transcription: RNA polymerase II at 2.8 angstrom resolution. Science 292: 1863 7186. Davenport, R.,J., Wuite, G. J., Landick, R., and Bustamante, C. 2000. Single-molecule study of transcriptional pausing and arrest by E. coli RNA polymerase. Science 287: 2497 2500. Delagoutte, E. and von Hippel, P. H. 2001. Molecular mechanisms of the functional coupling of the helicase (gp41) and polymerase (gp43) of bacteriophage T4 within the DNA replication fork. Biochemistry 40: 4459 4477. Doublie, S. and Ellenberger ,T. 1998a.The mechanism of action of T7 DNA polymerase. Curr. Opin. Struct. Biol. 8: 704 712. Doublie, S., Tabor, S., Long, A. M., Richardson, C. C., and Ellenberger, T. 1998b. Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 A resolution. Nature 391: 251 258. Drake, B., Prater, C. B., Weisenhorn, A. L., Gould, S. A., Albrecht, T.,R., Quate, C. F., Cannell, D. S., Hansma, H.,G., and Hansma, P. K. 1989. Imaging crystals, polymers, and

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6 RNA and DNA Polymerases Landick, R. 1997. RNA polymerase slides home: pause and termination site recognition. Cell 88: 741 744. Landick, R. 1999. Shifting RNA polymerase into overdrive. Science 284: 598 599. Marden, J. H. and Allen, L. R. 2002. Molecules, muscles, and machines: universal performance. Proc. Natl Acad. Sci. USA 2002. 99: 4161 4166. Meyhofer, E. and Howard, J. 1995. The force generated by a single kinesin molecule against an elastic load. Proc. Natl Acad. Sci. USA 92: 574 578. Nudler, E., Mustaev, A., Lukhtanov, E., and Goldfarb, A. 1997. The RNA DNA hybrid maintains the register of transcription by preventing backtracking of RNA polymerase. Cell 89: 33 41. Pal, M. and Luse, D. S. 2002. Strong natural pausing by RNA polymerase II within 10 bases of transcription start may result in repeated slippage and reextension of the nascent RNA. Mol. Cell Biol. 22: 30 40. Palangat, M., Meier, T. I., Keene, R. G., and Landick, R. 1998.Transcriptional pausing at 62 of the HIV-1 nascent RNA modulates formation of the TAR RNA structure. Mol. Cell 1: 1033 1042. Rees, W. A., Keller, R. W., Vesenka, J. P, Yang, G., and Bustamante, C. 1993. Evidence of DNA bending in transcription complexes imaged by scanning force microscopy. Science 260: 1646 1649. Schafer, D. A., Gelles, J., Sheetz, M. P., and Landick, R. 1991. Transcription by single molecules of RNA polymerase observed by light microscopy. Nature 352: 444 448. Stryer, L. 1995. Biochemistry (4th edn). New York: Freeman. Svoboda, K. and Block, S. M. 1994. Force and velocity measured for single kinesin molecules. Cell 77: 773 784. Szafranski, P. II, Smagowicz, W. J., Wierzchowski, K. L. 1985. Substrate selection by RNA polymerase from E. coli. The role of ribose and 5l-triphosphate fragments, and nucleotides interaction. Acta Biochim. Pol. 32: 329 349.

Uptain, S. M. and Chamberlin, M. J. 1997. Escherichia coli RNA polymerase terminates transcription efficiently at rho-independent terminators on single-stranded DNA templates. Proc. Natl Acad. Sci. USA 94: 13548 13553. Uptain, S. M., Kane, C. M., and Chamberlin, M. J. 1997. Basic mechanisms of transcript elongation and its regulation. Annu. Rev. Biochem. 66: 117 172. Wang, M. D.1999. Manipulation of single molecules in biology. Curr. Opin. Biothechnol. 10: 81 86. Wang, M. D., Schnitzer, M. J., Yin, H., Landick, R., Gelles, J., and Block, S. M. 1998. Force and velocity measured for single molecules of RNA polymerase. Science 282: 902 907. Wuite, G. J., Smith, S. B., Young, M., Keller, D., and Bustamante, C. 2000. Single-molecule studies of the effect of template tension on T7 DNA polymerase activity. Nature 404: 103 106. Yin, H., Landick, R., and Gelles, J. 1994. Tethered particle motion method for studying transcript elongation by a single RNA polymerase molecule. Biophys. J. 67: 2468 2478. Yin, H., Wang, M. D., Svoboda, K., Landick, R., Block, S. M., and Gelles, J. 1995. Transcription against an applied force. Science 270: 1653 1656. Yager, T. D. and von Hippel, P. H. 1987. In: Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology vol 2, Edited by F. C. Neidhardt et al. Washington DC: American Society for Microbiology, pp. 1241 1275. Yasuda, R., Noji, H., Kinosita, K. Jr, and Yoshida, M. 1998. F1-ATPase is a highly efficient molecular motor that rotates with discrete 120 degree steps. Cell 93: 1117 1124. Zaychikov, E., Denissova, L., and Heumann, H. 1995. Translocation of the Escherichia coli transcription complex observed in the registers 11 to 20: ‘jumping’ of RNA polymerase and asymmetric expansion and contraction of the ‘transcription bubble’. Proc. Natl Acad. Sci. USA 92: 1739 1743.

177

7 Helicases as Molecular Motors Mikhail K. Levin and Smita S. Patel

7.1

Introduction

Helicases are molecular motor proteins that use the energy of NTP1) hydrolysis to unidirectionally translocate along NA and separate (unwind) the complementary strands of the NA duplex. Helicases can also destabilize the secondary structure of RNA, remove NA associated proteins, and thread NA through various pores. By performing these functions, helicases serve as integral parts of the cellular machinery responsible for DNA replication, repair, recombination, transcription, ribosome biogenesis, translation, RNA splicing, RNA editing, RNA transport, RNA degradation, bacterial conjugation, and viral packaging/unpackaging (de la Cruz et al., 1999, Lüking et al., 1998, Matson, 1991). Helicase structure and function has been widely discussed in the literature (Bird et al., 1998, Carpousis et al., 1999, Eisen and Lucchesi, 1998, Frei and Gasser, 2000, Geourjon et al., 2001, Hall and Matson, 1999, Hoffmann-Berling, 1982, Labib and Diffley, 2001, Lei and Tye, 2001, Lohman, 1992, 1993, Lohman and Bjornson, 1996, Marians, 2000, Marintcheva and Weller, 2001, Soultanas and Wigley, 2000, 2001). In this chapter, we briefly review helicase structure and function and focus on recent models of unidirectional translocation and NA unwinding. We discuss the helicases from HCV and from T7 bacteriophage in greater detail. Since the discovery of the DNA unwinding activity in Escherichia coli in 1976 (Abdel-Monem and Hoffmann-Berling, 1976, Abdel-Monem et al., 1976), it has become apparent that helicases perform a broad range of functions in all living organisms. A recent study has revealed 134 ORFs coding for helicase-like proteins in the genome of the yeast, Saccharomyces cerevisiae (Shiratori et al., 1999). Extrapolating these results to other organisms, it can be assumed that approximately

1) Abbreviations: ADPNP, adenosine 5l-(b,g-imido)triphosphate; BM, Brownian motor; bp, base pair;

ds, double-stranded; EDTA, ethylenediaminetetraacetic acid; HCV, hepatitis C virus; NA, nucleic acid; NTP, nucleotide triphosphate; ORF, open reading frame; SDS, sodium dodecylsulfate; SF, superfamily; ss, single-stranded; TMP-PCP, deoxythymidine (b, g, methylene)triphosphate.

180

7.1 Introduction Table 7.1.

Helicase-related human genetic disorders.

Helicase

Family

Disease/Symptoms

References

XPB (ERCC2), XPD

SF2, RAD25

Xeroderma pigmentosum (blindness and deafness, light sensitivity, developmental disabilities), trichothiodystrophy (brittle hair), Cockayne syndrome (light sensitivity, short statue, premature aging)

(Sung et al., 1993)

ATRX (X-linked nuclear protein)

SF2

X-linked a-thalassemia/mental retardation syndrome (severe psychomotor retardation, characteristic facial features, genital abnormalities), Juberg Marsidi syndrome (mental retardation, growth failure, characteristic facies, sensorineural deafness, microgenitalism)

(Stayton et al., 1994)

BLM (RECQL3)

SF2, RecQ

Bloom syndrome (dwarfism, immunodeficiency, UV light sensitivity)

(Ellis et al., 1995)

Spinal muscular atrophy with respiratory distress

(Sebastiani et al., 1995)

CATF1 RAD54

SF2

Various cancers

(Kanaar et al., 1996)

WRN (RECQL2)

SF2, RecQ

Werner syndrome (scleroderma, cataracts, progeria, hypogonadism, diabetes)

(Gray et al., 1997)

RTS (RECQL4)

SF2, RecQ

Rothmund Thomson syndrome (skin atrophy, malformations of the teeth, nails, and bone)

(Kitao et al., 1999)

DNMT3B

SF2, ATRX-like Immunodeficiency centromeric instability facial anomalies syndrome

(Xu et al., 1999)

HAGE

SF2, DEAD

Tumor-specific expression

(Martelange et al., 2000)

Twinkle

T7 gp4-like

Autosomal dominant progressive external ophthalmoplegia (adPEO), severe retarded depression, avoidant personality, multiple mtDNA deletions

(Spelbrink et al., 2001)

BRIP1 (BACH1)

SF2, DEAH

Early-onset breast cancer, ovarian cancer

(Cantor et al., 2001)

SMARCAL1

SF2

Schimke immunoosseous dysplasia (short stature, proteinuria with progressive renal failure, cerebral ischemia)

(Coleman et al., 2000)

7 Helicases as Molecular Motors

2 % of ORFs in a eukaryotic genome code for helicases. Mutations in genes coding for helicases result in several human diseases including cancer, premature aging and others (Tab. 7.1) (Furuichi, 2001, Mohaghegh and Hickson, 2001, Nakura et al., 2000). This is not surprising, considering that helicases perform diverse functions and are involved in almost all DNA and RNA metabolic processes. Similar to higher organisms, many viruses code for proteins with helicase conserved motifs. These helicases play key roles in viral life cycles. Most are believed to be involved in viral genome replication, although some are required during other stages of the viral life cycle, such as genome packaging/unpackaging (Kadaré and Haenni, 1997, Lüking et al., 1998). Viral helicases are therefore attractive targets for antiviral therapy (Borowski et al., 2002, Crute et al., 2002, De Francesco et al., 2000, Hong et al., 2000, Littlejohn et al., 1998, Yao and Weber, 1998). As shown in Tab. 7.2, helicase-like proteins are found in the genomes of about half of animal virus families. Some viruses code for multiple helicases that belong to different superfamilies. Table 7.2 shows that helicases from three superfamilies are distributed among different viral classes and families. For example, SF3 helicases can be Table 7.2.

Animal viruses and associated helicases*.

Viral genome

dsDNA

Viral Family

Adenoviridae Herpesviridae Iridoviridae Papovaviridae Poxviridae

DNA and RNA RT

Hepadnaviridae Retroviridae

ssDNA

Circoviridae Parvoviridae

dsRNA

Reoviridae Birnaviridae

ssRNA ()

Arenaviridae Astroviridae Bunyaviridae Caliciviridae Coronaviridae Flaviviridae Orthomyxoviridae Picornaviridae Togaviridae

ssRNA (-)

Filoviridae Paramyxoviridae Rhabdoviridae

* Fields et al., 1996, Gorbalenya and Koonin, 1993.

Helicase superfamily SF1 SF2 SF3

X

other

X X X X

X X

X X X X X X

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182

7.2 Basic Properties of Helicases

found in () ssRNA, ssDNA, and dsDNA viruses. This highlights the fact that the evolution of viruses is a complex multifactor process that includes multiple origins and acquisitions of genes from host organisms (Fields et al., 1996).

7.2

Basic Properties of Helicases

Helicase function requires multiple enzymatic activities. Helicases hydrolyze ATP in the presence of Mg2, although some show a preference for other nucleotides such as GTP or dTTP. The NTPase activity in the absence of NA is low; however, it is stimulated many fold in the presence of NA. The NA-stimulated NTPase activity is coupled to unidirectional translocation of the helicase along the NA. Several in vitro assays provide evidence for unidirectional translocation. The dependence of steady state and pre-steady state kinetics of NTP hydrolysis on the length of a NA substrate can be used to measure the rate of the translocation (Dillingham et al., 2000, Raney and Benkovic, 1995, Young et al., 1994a, 1994b). Helicases have been shown to displace streptavidin bound to biotin linked to an end of a ssDNA. A 5l 3l helicase preferentially displaces the streptavidin bound to the 3l-end of the DNA whereas a 3l 5l helicase displaces the streptavidin bound to the 5l-end of the DNA indicating translocation directionality (Morris and Raney, 1999, Morris et al., 2002). However, the most common way to visualize translocation is by monitoring the unwinding of dsNA. Most helicases require a stretch of ssNA of specific polarity adjacent to the duplex region to initiate NA strand separation (Fig. 7.1). NA unwinding assays are typically performed by mixing a helicase with a partially duplex substrate and initiating the reaction by adding NTP. The reaction is terminated with an agent that denatures and dissociates the helicase from the NA, but leaves the remaining duplex substrate intact (for example, a mixture of EDTA and SDS). The completely unwound NA strands are resolved from the duplex substrate by native polyacrylamide gel electrophoresis. It is assumed that when the reaction is terminated, the incompletely unwound duplexes renature and appear as dsNA substrate on the gel. The strand separation process can also be measured in real time by using fluorescently labeled DNA and monitoring fluorescence or loss of resonance energy transfer as the strands are separated (Bjornson et al., 1994, Cheng et al., 2001, Eggleston et al., 1996, Houston and Kodadek, 1994, Raney et al., 1994). The unwinding substrate specificity is often used to determine the directionality of translocation. Accordingly, helicases that require a 3l ssNA tail to initiate NA un-

Figure 7.1.

Schematic of an unwinding reaction catalyzed by a 3 to 5l helicase. The helicase (gray circle) initiates unwinding by binding to the 3l-tail, hydrolyzing NTP, translocating along one strand of ss/dsNA substrate, and separating the strands of the substrate.


winding are assumed to translocate from 3l to 5l, and vice versa. This method has its shortcomings since some helicases need both 5l and 3l non-complementary tails to initiate NA unwinding, for example hexameric ring helicases like T7 phage gp4 helicase (Ahnert and Patel, 1997), T4 phage gp41 helicase (Richardson and Nossal, 1989), and bacterial DnaB helicase (LeBowitz and McMacken, 1986). Yet some helicases can initiate unwinding from a blunt end. Helicases are classified according to their biological function, NA substrate specificity, directionality of translocation and oligomeric state. Most helicases show a specificity for DNA or RNA and are classified accordingly. Others like the HCV helicase can unwind both dsDNA and dsRNA, and also hybrid duplexes (Gwack et al., 1997). Helicases can be grouped by their directionality of translocation into 3l to 5l and 5l to 3l. Oligomeric state divides the helicases into two established groups: ring (hexameric) helicases and all others (Patel and Picha, 2000). Studies of amino acid sequence homology revealed conserved helicase sequence motifs and laid the ground for a more detailed classification of helicases (Gorbalenya and Koonin, 1993). Two of these conserved motifs, Walker A and Walker B or motifs I and II are present in many NTP binding proteins (Walker et al., 1982). According to sequence homology, helicases are classified into families and superfamilies. For example, three families defined within the SF2 (DEAD, DEAH, and DexH) are named after variations in their Walker B conserved motif. Sequence homology classification does not correlate with biochemical properties of helicases. For example, helicases with different directionality and substrate specificity often belong to the same SF (Gorbalenya and Koonin, 1993). This suggests that small changes in helicase primary structure can be responsible for directionality and substrate specificity. It is likely that helicases with related primary structures share a general mechanism of translocation and unwinding. Therefore classification by sequence homology is useful for understanding the mechanism of helicase activity. Structural analysis provides an important tool for studying helicases. The number of available crystal structures of helicases has dramatically increased in recent years (Tab. 7.3). The helicases characterized structurally share similarities in their three-dimensional folds (recA-like folds). A helicase typically has several domains. Each helicase molecule has a single NTP binding site and a distinct NA binding site. Since the NTPase activity modulates the NA binding affinity, and vice versa, these two sites are allosterically linked. Several crystal structures of HCV helicase are available, and all show a very similar protein conformation (Cho et al., 1998, Kim et al., 1998, Yao et al., 1997, 1999). Figure 7.2 shows the structure of HCV NS3 helicase domain (NS3h). The three subdomains form a claw-like structure, and two of these subdomains resemble the 1A and 2A domains of PcrA (Fig. 7.2, pink and light green respectively). Conserved motifs I, II and III form the ATP binding site, while motifs V and VI are positioned so that they can be involved in transmitting a conformational change from the ATP binding site to the DNA binding site. The protein DNA contacts are distributed throughout the length of the protein-bound dU8 substrate. All the basic amino acid residues of the NS3h DNA binding site are positioned within hydrogen- or ionic- bonding distance

183

184


Table 7.3.

Atomic structures of helicases in the Protein Data Bank*.

PDB ID

Helicase

Organism

Res [Å]

Reference

Comments

1PJR

PcrA

Bacillus stearothermophilus

2.5

(Subramanya et al., 1996)

1HEI

NS3h

HCV

2.1

(Yao et al., 1997)

Ca2

1UAA

Rep

Escherichia coli

3.0

(Korolev et al., 1997)

DNA

1A1V

NS3h

HCV

2.2

(Kim et al., 1998)

DNA, SO24

8OHM

NS3h

HCV

2.3

(Cho et al., 1998)

1B79

DnaB

Escherichia coli

2.3

(Fass et al., 1999)

1C4O

UvrB

Thermus thermophilus

1.5

(Machius et al., 1999)

1CR0, 1CR1, 1CR2, 1CR4

Gene 4

Bacteriophage T7

2.3

(Sawaya et al., 1999)

dTTP, dATP, dTDP

1CU1

NS3

HCV

2.5

(Yao et al., 1999)

PO34

1D8B

SGS1

Saccharomyces cerevisiae

n/a

(Liu et al., 1999)

NMR

1D9X, 1D9Z

UvrB

Bacillus caldotenax

2.6

(Theis et al., 1999)

ATP, Mg2

1JWE

DnaB

Escherichia coli

n/a

(Weigelt et al., 1999)

NMR

1QHG, 1QHH PcrA


2.5

(Soultanas et al., 1999)

ADPNP, Mg2

1QVA

eIF4A


2.5

(Johnson and McKay, 1999)

2PJR, 3PJR

PcrA


2.9

(Velankar et al., 1999)

ss/dsDNA, ADPNP, SO24

1E0J, 1E0K

Gene 4

Bacteriophage T7

3.0

(Singleton et al., 2000)

ADPNP, Mg2

1F08

E1

Bovine papillomavirus

1.9

(Enemark et al., 2000)

1FUK, 1FUU

e/F4A


1.75

(Caruthers et al., 2000)

1E9R, 1E9S

TrwB

Escherichia coli

2.4

(Gomis-Ruth et al., 2001)

1G8Y

RepA

Escherichia coli

2.4

(Niedenzu et al., 2001)

1GM5

RecG

Thermotoga maritima

3.24

(Singleton et al., 2001)

DNA, ADP, Mg2

1HV8

MjDEAD Methanococcus jannaschii

3.0

(Story et al., 2001)

SO24

1IN4, 1IN5, 1IN6, 1IN7, 1IN8, 1J7K

RuvB

Thermotoga maritima

1.6– 2.0

(Putnam et al., 2001)

ADP

1JEQ, 1JEY

Ku

Homo sapiens

2.7

(Walker et al., 2001)

DNA

SO24

* All structures were determined by X-ray crystallography except indicated otherwise in Comments. Important crystallization co-factors are shown in Comments.


Figure 7.2. Conserved motifs and the crystal structure of HCV helicase. (A) Helicase subdomains (pink, green and blue) and conserved motifs (I, II, III, V, VI) are shown. (B) Stereo view of HCV helicase complexed with oligodeoxynucleotide (dark blue), a sulfate ion (dark blue), and subdomains (same color as in A). Also shown are the residues of the conserved motifs (same color as in A) and the residues making contacts with the ssDNA (orange). The structure was visualized in Swiss-PdbViewer (www.expasy.ch/spdbv/) from the Protein Data Bank entry 1A1V (Kim et al., 1998), and rendered in POV-Ray (www.povray.org).

from the DNA phosphoryl groups and make about half of all the contacts of the phosphoryl groups. About 60 % of protein DNA contacts occur through the phosphoryl groups and 20 % through ribose groups while only about 20 % of interactions occur through the DNA bases. This observation is consistent with the fact that NS3 binds ssDNA in a sequence independent manner. The structural features responsible for discriminating between DNA and RNA substrates are not apparent from any of the helicase crystal structures. The Rep helicase and PcrA helicase crystal structures have been solved in the presence of ssDNA and ADPNP. These structures reveal the presence of four sub-domains. The 1A and 2A sub-domains are homologous to each other and contain the RecA-fold. Residues from the conserved helicase motifs line the interface of these sub-domains and bind NTP. The ssNA binds in a groove that is formed by 1A and 2A sub-domains. Domain movements between 1A and 2A and rigid body rotation of subdomain 2B have also been observed in different structures and these have been implicated as intermediates of the helicase reaction (Velankar et al., 1999). The ring helicases assemble from six identical (most ring helicases) or non-identical (MCM helicase) subunits. The three-dimensional structures of ring helicases from various organisms have been characterized extensively by electron microscopy, revealing that the six subunits are arranged in a toroidal shape with the cen-

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186



Figure 7.4. Conformational changes involved in the stepping mechanisms. The six steps represent the minimal number of conformational changes needed to support unidirectional translocation. The mechanism does not represent the kinetics of translocation and some steps can be concerted.

tral channel large enough to accommodate a strand of DNA or RNA (Fig. 7.2; Patel and Picha, 2000). The crystal structures of ring helicases including RepA, RuvB, and a fragment of T7 gp4 protein reveal that the helicase domain fold is similar to the RecA protein (Fig. 7.3). The T7 helicase NTP binding site as well as most conserved motifs appear at the subunit interface. In addition, a critical arginine (R522) from a neighboring subunit is within hydrogen-bonding distance of the gamma phosphate of NTP bound at the interface and implicated in transducing conformational changes between subunits of the hexamer (Singleton et al., 2000). Consistently, the biochemical data indicate that the subunits coordinate their activities (Hingorani et al., 1997). Very little detailed structural information is available about the NA binding site. Electron microscopy studies of ring helicases and biochemical studies indicate that the amino acids in the central channel are involved in binding the NA (Egelman et al., 1995, Morris and Raney, 1999, VanLoock et al., 2002). A future challenge is to determine the structure of a ring helicase bound to appropriate NA substrates, which will greatly help in extracting mechanistic information related to the pattern of NTP hydrolysis by the six subunits and how NTPase facilitates movement of the ring along the NA.

m Figure 7.3.

Crystal structure of bacteriophage T7 gp4 helicase domain. A) shows the six helicase subunits in a ring with a C2 symmetry and four ADPNP (dark blue) bound at the subunit interface. Symmetrical subunits are shown in the same color. (B) Helicase conserved motifs of T7 gp4 protein. Gray area represents the helicase domain shown in the crystal structure. (C) Stereo view from inside the ring of subunits A and B complexed to ADPNP (dark blue) and Mg2+ (green, space filling). The helicase conserved motifs are shown in the same color as in (B). The subunits are colored the same as in (A). Amino acid residue R522 is shown. The structures were visualized in Swiss-PdbViewer (ww.expasy.ch/spdbv/) from the Protein Data Bank entry 1E0J (79), and rendered in POV-Ray (www.povray.org).

187

188

7.3 Mechanism of Helicase Activity

7.3

Mechanism of Helicase Activity

In order to unwind a long stretch of duplex NA, a helicase has to unidirectionally translocate while locally separating the strands of NA. Thus, unwinding is viewed as a combination of translocation and strand separation processes. Both of these processes are fueled by the energy released from NTP hydrolysis. While the detailed mechanisms of translocation and strand separation are not known, generally, the unwinding process occurs as follows: The helicase bound near the junction of ss and duplex parts of the NA substrate cycles through different NTP ligation states. E p E · NTP p E · NDP · Pi p E · NDP p E NTP binding, hydrolysis and product release act as a switch that induces conformational changes on the helicase NA binding site, forcing it to change its affinity for the NA or to perform a power stroke. These conformational changes drive the unidirectional translocation and unwinding in a stepwise fashion (Fig. 7.1). Several mechanisms of NTPase-coupled translocation and unwinding have been proposed for different helicases (Kim et al., 1998, Lohman and Bjornson, 1996, Lohman et al., 1998, Patel and Picha, 2000, Singleton et al., 2000, Soultanas and Wigley, 2001, von Hippel and Delagoutte, 2001). Some of the differences in the proposed mechanisms reflect the diverse biochemical properties of different helicases, such as oligomeric state, the interactions with ssNA versus duplex NA, and the effect of NTP ligation state on the NA binding properties. Considering that helicases show a significant degree of structural homology, it would be desirable to arrive at a general model that, with minor variations, can explain the properties of all helicases. The general model should address the mechanism of unidirectional movement as well as NA strand separation. It should be adaptable to different classes of helicases, explain translocation directionality and helicase function. 7.3.1

Unidirectional Translocation

Most commonly, translocation is viewed as a stepping process similar to the one shown in Fig. 7.4. Stepping requires at least two NA binding sites that independently bind and release NA and change the distance between each other. In the case of a monomeric helicase, the two hands attached to the black circles represent two parts of its NA binding site that move relative to each other. In the case of a dimeric helicase, the hands represent NA binding sites on two subunits of a helicase complex. The conformational changes in the NA binding sites, shown in Fig. 7.4, occur in response to NTP binding, hydrolysis and product release at the NTP binding site of the helicase. In an oligomeric helicase, the NTPase activity at each subunit controls its NA binding site; thus, coordinated NTPase activity leads to coordinated binding and release of NA. Note that in a monomeric helicase, both NA binding sites are under the control of a single NTP binding site.


Free energy, DG, plotted as a function of ssNA length. Solid line shows the free energy E0 of helicase bound to ssNA. Dotted line shows the free energy of the helicase-ssNA-ATP complex, EATP. The black circles represent the position of the helicase on ssNA. (i) Free helicase. (ii) Tight binding of the helicase to ssDNA is associated with a forward movement. (iii and iv) helicase-ATP complex is weakly bound to ssDNA and can diffuse along it in either direction. (iil) if ATP is hydrolyzed while the helicase is in a favorable position (iv), helicase rebinding is associated with a step forward. The helicase is in the same conformation as in (ii), but one step ahead. Figure 7.5.

In the stepping mechanism, a helicase undergoes a round of conformational changes (from i to vi) after which it appears in the same conformational state (i’), one step away from its original position. The cycle starts with the first NA binding site tightly bound to NA (closed hand, i) and the second binding site weakly bound to NA (open hand). In a power stroke motion, the second binding site moves away from the first one (ii), followed by tight binding of the second site (closing, iii), opening of the first site (iv), a conformational change (power stroke) bringing sites back together (v), closing of the first site (vi), and, finally, opening of the second site (i’). Thus one cycle of a stepping mechanism is completed in six conformational changes controlled by ATPase cycle. Stepping is an easily understood and accepted model of translocation since it is frequently observed in the macroscopic world; for example, most terrestrial animals translocate by stepping. At the same time, stepping is a complex process involving many coordinated movements. As applied to a monomeric helicase, all six conformational changes of the stepping mechanism must occur under the influence of the single NTP binding site. In addition, stepping mechanism ignores the stochastic nature of the microscopic world. Macroscopically-observed enzymatic reactions are composed of a large number of microscopic stochastic events. Biological systems have evolved to utilize this randomness to achieve a non-random directed result. It is likely, that helicases as well as other molecular motors are no exception. It has been shown that Brownian motion can be integrated into a model of a molecular motor leading to a simpler, more flexible, and more robust design (Astumian, 2000, 2001, Huxley, 1998, Oster, 2002).

189

190


A Brownian motor (BM) uses the thermal fluctuations from its surroundings to achieve unidirectional movement. The notion of a BM does not contradict the second law of thermodynamics, because the energy of NTP hydrolysis puts a bias on the system. There are many ways by which a BM can bias random thermal movement, and we discuss one of these in relation to the mechanism of helicases. Consider the interactions between the helicase and the ssNA. Normally, one assumes that the free energy of complex formation is constant along the length of ssNA. Suppose that the NA binding site of the helicase interacts with the ssNA in a way that creates a periodic dependence of binding energy along the length of NA (saw tooth profile). If the saw tooth profile is asymmetric, that is, one side of the tooth is steeper than the other (Fig. 7.5, solid line), most binding events of the free helicase (i) to ssNA will be accompanied by a movement of the helicase from left to right. The binding associated with the movement brings the system to its local minimum (ii). The depth of the energy minimum is such that the helicase cannot escape it due to the thermal fluctuations. Suppose now that when NTP binds to the helicase, it causes a conformational change in its NA binding site. As a result, the ssNA binding free energy becomes weaker and constant along the ssNA length (Fig. 7.5, dotted line). This allows the helicase to slide along ssNA randomly in either direction because of Brownian motion (iii and iv). The random movement of the helicase continues only for a short time. After NTP gets hydrolyzed and the NTPase products are released, the asymmetric saw tooth binding energy profile is restored and the helicase rebinds to the lattice tightly. The resulting state of the helicase after rebinding depends upon its position at the end of its weakly bound state. If Brownian motion moved the helicase backward (position iii), the helicase will end up in the same position that it

Figure 7.6. Thermodynamics of NA strand separation. Relative free energies of species involved in the NA unwinding reaction. Strand separation of free dsNA is thermodynamically unfavorable (solid line) because of inter-strand interactions. Strand separation, when coupled to helicase binding to the ssNA product, is thermodynamically favorable, but may be slow because of the relatively high free energy of the transition state, DG³ (dashed line). Helicase may accelerate strand separation by binding and stabilizing the transition state (dotted line).


started, that is no net movement (ii), after the restoration of the saw tooth energy profile. However, there is a significant probability that the helicase diffuses forward (position iv). In this case, the helicase will end up one step forward from its original position upon tight binding to ssNA. Thus, repeated binding and hydrolysis of NTP results in a net unidirectional translocation of the helicase along the ssNA. Computer simulation of a BM mechanism created by F.-J. Elmer et al. is available.1) The efficiency of a motor is defined as the fraction of productive steps per round of NTP hydrolysis. Several factors can affect the efficiency of a Brownian motor. The greater the asymmetry of the saw tooth energy profile the greater the efficiency of the motor, since it will increase the fraction of helicase molecules that reach position iv and make a step forward. The efficiency is also highly dependent on the rate of NTP hydrolysis which determines the life time of the weakly bound state (Fig. 7.5, positions iii and iv). A helicase can move forward even against a moderate external force as long as a significant fraction of the helicase can diffuse forward to position iv. The stepping and BM translocation models are distinguished by the fact that stepping requires at least two NA binding sites that bind and release NA independently and move relative to each other. Although BM benefits from two or more coordinated NA binding sites, a BM with just one NA binding site can translocate unidirectionally. Stepping and BM translocation models are also distinguished by the origin at which the force required for unidirectional translocation, is generated. In the stepping model, the force is generated at the junction of the NA binding sites. This force moves the NA binding sites relative to each other, producing unidirectional movement. Two kinds of forces are at work in a Brownian motor. One is generated at the NA binding site during tight binding of the helicase to NA. The other is Brownian motion that can move the helicase in any direction along the NA while the helicase binding is in a flat energy profile. The BM model for monomeric helicases predicts limited processivity of translocation and unwinding in vitro because the helicase holds on to the ssNA only via one binding site, and therefore it is more likely to dissociate during the sliding phase. Experimental observations support this prediction (Ali and Lohman, 1997, Dillingham et al., 2000). A dimeric or hexameric helicase with two or six ssNA binding sites can implement a BM mechanism with a greater efficiency. This is especially true if the helicase subunits coordinate their ATPase cycles and take turn binding and hydrolyzing ATP. In fact, a coordinated dimer should appear as if it is stepping, because its subunits are taking turns moving forward. The idea of BM or molecular ratchet model is well established in the field of ion pumps and filament motors, while only a few researchers consider a BM mechanism for helicases (Doering et al., 1995). The BM translocation model is consistent with most known facts about helicases. Therefore the BM model is preferred over the stepping model as a simpler, flexible, and a more evolvable (more likely to appear in an evolutionary process) model. 1) http://monet.physik.unibas.ch/Zelmer/bm/

191

192


7.3.2

Step Size of the Helicase

An important characteristic of a molecular motor is its step size, determination of which can help establish the mechanism of the motor. Several methods have been used to estimate the step size of helicases. The step size of a helicase has been estimated by measuring the single turnover kinetics of NA unwinding and fitting the resulting kinetics to a stepping equation (Ali and Lohman, 1997). Estimation of the number of steps (or the step size) from appearance of the final product of the reaction relies on the assumption that all the unwinding species have identical stepping properties. Ideally, helicase molecules that have the same primary structure should behave in an identical manner. However, if multiple populations exist that have different stepping rates then the unwinding kinetics will appear to indicate a fewer kinetic steps than it actually does, thus exaggerating the step size. The lower limit of the step size can be determined by measuring the coupling ratio, which is the number of bases the helicase can travel per NTP hydrolyzed. For an accurate estimation of step-size there is a need to develop methods, such as single molecule methods, which will allow observation of the helicase molecule and a direct measurement of its step size. 7.3.3

NA Strand Separation

Separating the strands of a duplex NA involves breaking the hydrogen bonds that hold the bps together. Although this is a thermodynamically unfavorable process with an average DG of Z 10 kJ mol 1 base 1 at physiological temperature, the NA bp open and close spontaneously. Since opening of bp allows exchange of imino protons, the opening rates can be measured by proton NMR. The rates of spontaneous bp opening range from much more than 1000 s 1 (at 15 hC) for terminal bps, to about 30 s 1 for bps in the middle of the duplex region (Gueron and Leroy, 1995). Even though individual bps open at a high rate, the equilibrium for this reaction is shifted towards bp formation (Fig. 7.6, solid line). In order to make the unwinding process thermodynamically favorable under physiological conditions, the helicase must stabilize the open bps by binding to ssNA (Fig. 7.6, dashed line). The helicase does this while moving forward in a stepwise fashion, as described in the previous section. It has been proposed that strand separation can occur either actively or passively (Lohman, 1992). According to an active mechanism, the helicase increases the rate of bp opening in addition to stabilizing the ssNA product by binding. The rate of bp opening can be increased by lowering the energy of the transition state (Fig. 7.6, dotted line), which can occur by helicase binding to an unwinding intermediate. If the helicase unwinds NA by the ‘passive’ mechanism, by definition, it does not change the rate of bp opening. One way the helicase can stabilize the unwinding transition state is by binding to a distorted duplex region. In fact many researchers view interactions of the helicase


Figure 7.7.

The Kd of HCV helicase with various DNA substrates determined by fluorimetric

titration.

with dsNA as an indication of an active unwinding mechanism (Velankar et al., 1999, Wong and Lohman, 1992). Another way to actively disrupt bps is by unidirectional translocation. It has been shown that several helicases can dislodge streptavidin from biotin that is covalently linked to the end of ssDNA (Morris and Raney, 1999, Morris et al., 2002), and also dissociate other proteins bound to RNA (Jankowsky et al., 2001, Linder et al., 2001). Considering that the biotin– streptavidin complex is an unnatural substrate for helicases, these results indicate that helicases can disrupt the bonds that lie in their path without specifically interacting with the bonded moieties. It is difficult to distinguish between active and passive helicase mechanisms simply from the kinetics of NA unwinding, because the rate of spontaneous bp opening exceeds the rate of translocation and unwinding by any helicase or replicating system. Instead of a block on the path of a helicase, base pairing should be viewed as a fluctuating force directed against the movement of the helicase. Binding to the duplex NA and destabilizing it may become useful for a helicase that unwinds with a step size of many bps. Although it is unclear how this mechanism would improve the efficiency of a helicase relative to one that unwinds as a wedge which translocates between two strands of NA while pushing them apart without any specific interactions. It is impossible to say if the latter constitutes active or passive unwinding because it contains features of both. We propose that any protein capable of unidirectional translocation along one strand of dsNA while excluding the other strand can unwind dsNA by the wedge mechanism. Therefore, translocation

193

194

7.4 HCV Helicase

is the primary activity of a helicase and helicases should be defined as proteins that can unidirectionally translocate along a strand of nucleic acid.

7.4

HCV Helicase

The genome of HCV (Flaviviridae family) is translated into a single polypeptide cleaved into 10 proteins. NS3 protein combines protease and helicase activities on its N- and C-terminal domains, respectively. Another interesting feature of this helicase is that it can bind and unwind both DNA and RNA substrates (Gwack et al., 1997, Tai et al., 1996). The helicase activity of NS3 is essential for the virus (Kolykhalov et al., 2000) and it is postulated to play a role in viral replication. The crystal structure discussed earlier shows that NS3 can bind to ssDNA eight nucleotides long. Our results show that although the minimal binding site of NS3 on ssDNA is eight bases, NS3 can tightly bind ss/dsDNA substrates with a much shorter 3lssDNA tail, but not a 5l-ssDNA tail (Fig. 7.7). Additional data suggests that NS3 binding to ss/dsDNA substrate with a short 3l-ssDNA tail is accompanied by the separation of several bps, which was observed in the absence of ATP hydrolysis. We propose that tight binding of NS3 helicase to ssDNA is associated with a movement from 3l to 5l along the 3l-DNA strand, which provides directionality. In other words, the free energy binding profile of NS3 and ssDNA is tilted towards the 5lend, as shown in Fig. 7.5, positions i and ii. This property arises from the interaction of the DNA binding site of NS3 with ssDNA. Interestingly, the movement is not powered by ATP hydrolysis. In fact, it occurs in the absence of ATP. Instead, the movement is powered by the binding energy of NS3 to ssDNA. As discussed earlier, movement along ssDNA can lead to duplex strand separation even without specific interactions with the duplex (wedge mechanism). After NS3 is tightly bound to ssDNA, the system becomes trapped in its lowest energy state with any further movement and subsequent cycles of strand separation becoming impossible. Naturally, we looked at the effect of ATP on NS3 affinity to DNA. Experiments with a non-hydrolyzable ATP analog showed that NS3 ATP complex has a reduced affinity for the DNA substrate (Fig. 7.7), whereas the presence of ADP does not affect the affinity of NS3 for DNA. Thus binding of ATP switches NS3 to a different conformation that has a weaker affinity for DNA. The weaker binding state allows the structural elements of the DNA binding site to move randomly and NS3 protein may diffuse along the DNA strand (Fig. 7.5, positions iii and iv). Following ATP hydrolysis and product release, NS3 tightly re-binds the DNA substrate, and with a certain probability, moves forward. Thus, the cycling of NS3 between tightly bound (ii) and weakly bound (iii and iv) states, under the control of the ATPase cycle, leads to unidirectional translocation. Since the helicase movement to position (ii) occurs along the decreasing energy slope, it can be considered a power stroke. Brownian motion however, plays a key role in this mechanism giving the helicase a chance to move over the barrier to the


Figure 7.8.

Model of T7 gp4 helicase bound to ss/dsDNA substrate.

next energy well. Therefore HCV helicase can function as a combination of power stroke and Brownian ratchet described by Astumian (1997), Mogilner et al. (2001), Wang and Oster (2002). The proposed mechanism of translocation and unwinding is similar to those described previously for NS3 and other helicases in the way the ATPase cycle modulates the interaction of the DNA with the helicase (Bjornson et al., 1996, Kim et al., 1998, Velankar et al., 1999). According to the mechanism proposed here for NS3, the energy of ATP hydrolysis is not used directly for translocation or DNA strand separation. Instead, the energy of ATP binding is used to recover the enzyme from the energetic well of a tightly bound state while ATP hydrolysis, a reaction with a DG 0, triggers helicase rebinding linked to forward motion. Therefore, the energy of ATPase is used for unwinding and translocation indirectly. The proposed mechanism may be applicable to other helicases, although they may have phases of their ATPase cycle and DNA binding affinity coordinated in a different manner; for example, ATP binding may switch the helicase into a tight ssDNA binding state rather than the weak binding state as observed for the NS3 helicase.

7.5

Bacteriophage T7 gp4 Helicase

As discussed earlier, T7 gp4 helicase is a ring helicase. It is encoded by the bacteriophage and used for phage DNA replication and recombination. Although gp4 functions as a part of the replication complex, the purified helicase is capable of unwinding dsDNA efficiently using the energy of dTTP hydrolysis (Matson and Richardson, 1983, Patel et al., 1992). Biochemical studies indicate that T7 gp4 can translocate along ssDNA in a unidirectional manner (Tabor and Richardson,

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1981). Recently we have measured the rate constant of translocation as 130 bases s 1 (at 18 hC) and the dTTPase activity coupled to translocation is 50 s 1. This indicates that the helicase translocates on an average three bases per dTTP hydrolyzed (Kim et al., 2000). The mechanism of unidirectional movement is not known, but it appears to require the cooperative action of the six subunits. There is a negative cooperativity in dTTP binding, such that only three to four nucleotides bind tightly (Hingorani and Patel, 1996). Mutant poisoning (Patel et al., 1994) and pre-steady state kinetic experiments (Hingorani et al., 1997) have indicated that the subunits of the ring act in a cooperative manner. Sequential NTP hydrolysis has been observed by pre-steady state kinetic experiments in the absence of ssDNA (Hingorani et al., 1997). Similarly, in the presence of ssDNA, the NTP hydrolysis at one subunit is coupled to product release at the other subunit (unpublished results), which suggests a sequential mechanism of NTP hydrolysis. The pattern of sequential hydrolysis is not clear. That is, the number of subunits participating in hydrolysis as well as the number of groups hydrolyzing sequentially needs to be (one at a time, or several at a time) determined. Several possible mechanisms of NTP hydrolysis by the ring helicases have been discussed (Patel and Picha, 2000). The role of NTPase activity in ring helicases is similar to that of other helicases, in that NTP acts as a molecular switch. T7 gp4 binds ssDNA in the presence of dTMP-PCP, a non-hydrolyzable analog of dTTP with a Kd around 10 nM and dTTP hydrolysis and Pi release leads to weakening of the interactions with the ssDNA (Hingorani and Patel, 1993). Thus DNA binding and release events are catalyzed by dTTP binding and Pi release steps, but it is not clear how these events lead to unidirectional translocation. The crystal structure of the gp4 fragment reveals that three adjacent subunits are in different conformational states (Fig. 7.3). Thus, NTPase-induced conformational changes in the subunits and cooperative action of these subunits must play a vital role in translocation and unwinding. The ring structure of T7 gp4 hexamer and the mode of ssDNA binding within the central channel result in a topologically linked helicase DNA complex (Egelman et al., 1995, Yu et al., 1996). This structure of encircling the ssDNA track confers high processivity to T7 gp4 helicase and other ring helicases. High processivity of translocation distinguishes the ring helicases from helicases such as E. coli UvrD and PcrA helicases which do not assemble into rings. Exceptions include RecBCD that may not encircle the DNA, but shows a high processivity of DNA unwinding (Bianco et al., 2001). Processivity (P) is defined as the probability of making a step forward on the NA lattice divided by the probability of dissociation from that position on the lattice. The reported average processivity of UvrD is about 10 bp (P 0.9) during dsDNA unwinding (Ali and Lohman, 1997), and PcrA shows a similarly low processivity of translocation on ssDNA (Dillingham et al., 2000). The average processivity of ssDNA translocation by T7 gp4 hexamer on the other hand is 0.99996 (Kim et al., 2002), a number close to 1, which indicates that the probability of T7 helicase dissociating from ssDNA during translocation is very low. The average number of steps taken along ssDNA before dissociation


(N) can be determined from the relationship, P (N - 1)/N. We calculate N 33.3 q 103 steps or T7 gp4 helicase travels 66.7 kb (with a step size of two bases) on an average along ssDNA before dissociating. T7 gp4 helicase by itself is highly processive and association with T7 DNA polymerase may make it even more efficient. Another factor that may increase the processivity of translocation is the coordination of activity among the hexamer subunits, as discussed above. Gp4 ring helicase interact with only one strand of the DNA, which binds within the central channel of the ring, and excludes the complementary strand from the central channel (Ahnert and Patel, 1997; Fig. 7.3). This mode of DNA binding minimizes immediate re-annealing of the DNA strands after they are separated. In contrast to helicases such as Rep and PcrA, T7 gp4 shows little affinity for duplex DNA. Thus, it appears that unidirectional movement of gp4 helicase along ssDNA plays a crucial role in DNA unwinding. Preliminary measurements indicate that T7 gp4 unwinds dsDNA with a rate of about 40 bp s 1. Thus, T7 gp4 helicase translocates along ssDNA at a rate about three times faster than the measured rate of DNA unwinding. A simple way to unwind DNA is by the wedge mechanism discussed in 7.3.3. Nevertheless additional, structural and biochemical studies are necessary to elucidate the mechanisms of translocation and unwinding.

7.6

Conclusions

Genome sequencing and comparison of primary structures has unraveled a large number of putative helicases. Future studies will focus on determining the biochemical activities of these putative helicases and defining their cellular roles in various NA metabolism processes. Recently many helicases have been associated with human diseases; however, the link between a defective helicase and its manifestation in a disease such as cancer or premature aging, remains a subject of intense study. Viral helicases are attractive targets for drug therapy; therefore the basic understanding of the structure and mechanism of viral helicases will aid the process of drug discovery. Since the discovery of a helicase in 1976, a large amount of information about helicases has been accumulated. Nevertheless we lack a general mechanism that explains NA unwinding and other functions performed by helicases. Despite all the differences in their properties, it is likely that all helicases operate by a similar mechanism. We believe that unidirectional translocation is the basic activity of all helicases responsible for dsNA strand separation and other functions such as protein dissociation, NA transport and packaging. Of all the translocation mechanisms proposed, a BM model is the simplest and the most versatile, since it can be applied to helicases with different structures and functions. To shed new light on the mechanism of unidirectional translocation and NA unwinding, new methodologies will be applied to helicases such as analysis of single molecules that will allow direct observation of the helicase reaction. Detailed information as well as general mechanisms of helicase activity will emerge from these efforts.

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Acknowledgements

We would like to thank C. M. Drain, A. V. Persikov and N. M. Stano for critical discussions of the manuscript. This work was supported by NIH grant GM55310.

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References Singleton, M. R., S. Scaife, and D. B. Wigley. 2001. Structural analysis of DNA replication fork reversal by RecG. Cell 107: 79 89. Soultanas, P. and D. B. Wigley. 2000. DNA helicases: ‘inching forward’. Curr. Opin. Struct. Biol. 10: 124 128. Soultanas, P. and D. B. Wigley. 2001. Unwinding the ‘Gordian knot’ of helicase action. Trends Biochem. Sci. 26: 47 54. Soultanas, P., M. S. Dillingham, S. S. Velankar, and D. B. Wigley. 1999. DNA binding mediates conformational changes and metal ion coordination in the active site of PcrA helicase. J. Mol. Biol. 290: 137 148. Spelbrink, J. N., F. Y. Li, V. Tiranti, K. Nikali, Q. P. Yuan, M. Tariq, S. Wanrooij, N. Garrido, G. Comi, L. Morandi, L. Santoro, A. Toscano, G. M. Fabrizi, H. Somer, R. Croxen, D. Beeson, J. Poulton, A. Suomalainen, H. T. Jacobs, M. Zeviani, and C. Larsson. 2001. Human mitochondrial DNA deletions associated with mutations in the gene encoding Twinkle, a phage T7 gene 4-like protein localized in mitochondria. Nature Genet. 28: 223 231. Stayton, C. L., B. Dabovic, M. Gulisano, J. Gecz, V. Broccoli, S. Giovanazzi, M. Bossolasco, L. Monaco, S. Rastan, and E. Boncinelli. 1994. Cloning and characterization of a new human Xq13 gene, encoding a putative helicase. Hum. Mol. Genet. 3: 1957 1964. Story, R. M., H. Li, and J. N. Abelson. 2001. Crystal structure of a DEAD box protein from the hyperthermophile Methanococcus jannaschii. Proc. Natl Acad. Sci. USA 98: 1465 1470. Subramanya, H. S., L. E. Bird, J. A. Brannigan, and D. B. Wigley. 1996. Crystal structure of a DExx box DNA helicase. Nature 384: 379 383. Sung, P., V. Bailly, C. Weber, L. H. Thompson, L. Prakash, and S. Prakash. 1993. Human xeroderma pigmentosum group D gene encodes a DNA helicase. Nature 365: 852 855. Tabor, S. and C. C. Richardson. 1981. Template recognition sequence for RNA primer synthesis by gene 4 protein of bacteriophage T7. Proc. Natl Acad. Sci. USA 78: 205 209. Tai, C. L., W. K. Chi, D. S. Chen, and L. H. Hwang. 1996. The helicase activity associated with hepatitis C virus nonstructural protein 3 (NS3). J. Virol. 70: 8477 8484. Theis, K., P. J. Chen, M. Skorvaga, B. Van Houten, and C. Kisker. 1999. Crystal struc-

ture of UvrB, a DNA helicase adapted for nucleotide excision repair. EMBO J. 18: 6899 6907. VanLoock, M. S., A. Alexandrov, X. Yu, N. R. Cozzarelli, and E. H. Egelman. 2002. SV40 Large T antigen hexamer structure. Domain organization and DNA-induced conformational changes. Curr. Biol. 12: 472 476. Velankar, S. S., P. Soultanas, M. S. Dillingham, H. S. Subramanya, and D. B. Wigley. 1999. Crystal structures of complexes of PcrA DNA helicase with DNA substrate indicate an inchworm mechanism. Cell 97: 75 84. von Hippel, P. H. and E. Delagoutte. 2001. A general model for nucleic acid helicases and their ‘coupling’ within macromolecular machines. Cell 104: 177 190. Walker, J. E., M. Saraste, M. J. Runswick, and N. J. Gay. 1982. Distantly related sequences in the alpha- and beta-subunits of ATP synthase, myosin, kinases and other ATP-requiring enzymes and a common nucleotide binding fold. EMBO J. 1: 945 951. Walker, J. R., R. A. Corpina, and J. Goldberg. 2001. Structure of the Ku heterodimer bound to DNA and its implications for doublestrand break repair. Nature 412: 607 614. Wang, H. and Oster, G. (2002). Ratchets, power strokes, and molecular motors. Applied Physics A. (in press). Weigelt, J., S. E. Brown, C. S. Miles, N. E. Dixon, and G. Otting. 1999. NMR structure of the N-terminal domain of E. coli DnaB helicase: implications for structure rearrangements in the helicase hexamer. Structure Fold. Des. 7: 681 690. Wong, I. and T. M. Lohman. 1992. Allosteric effects of nucleotide cofactors on Escherichia coli Rep helicase-DNA binding. Science 256: 350 355. Xu, G. L., T. H. Bestor, D. Bourc’his, C. L. Hsieh, N. Tommerup, M. Bugge, M. Hulten, X. Qu, J. J. Russo, and E. Viegas-Pequignot. 1999. Chromosome instability and immunodeficiency syndrome caused by mutations in a DNA methyltransferase gene. Nature 402: 187 191. Yao, N., P. Reichert, S. S. Taremi, W. W. Prosise, and P. C. Weber. 1999. Molecular views of viral polyprotein processing revealed by the crystal structure of the hepatitis C virus bifunctional protease-helicase. Structure Fold. Des. 7: 1353 1363.

7 Helicases as Molecular Motors Yao, N. and P. C. Weber. 1998. Helicase, a tar- Young, M. C., D. E. Schultz, D. Ring, and P. H. get for novel inhibitors of hepatitis C virus. von Hippel. 1994a. Kinetic parameters of the Antivir. Ther. 3: 93 97. translocation of bacteriophage T4 gene 41 Yao, N. H., T. Hesson, M. Cable, Z. Hong, A. D. protein helicase on single-stranded DNA. J. Kwong, H. V. Le, and P. C. Weber. 1997. Mol. Biol. 235: 1447 1458. Structure of the hepatitis C virus RNA heli- Yu, X., M. M. Hingorani, S .S. Patel, and E. H. case domain. Nature Struct. Biol. 4: 463 467. Egelman. 1996. DNA is bound within the Young, M. C., S. B. Kuhl, and P. H. von Hippel. central hole to one or two of the six subunits 1994b. Kinetic theory of ATP-driven transloof the T7 DNA helicase. Nature Struct. Biol. 3: cases on one-dimensional polymer lattices. J. 740 743. Mol. Biol. 235: 1436 1446.

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Part 2 Mechanochemistry

8 How Protein Motors Convert Chemical Energy into Mechanical Work George Oster and Hongyun Wang ‘Biologists observe things that cannot be explained. Theorists explain things that cannot be observed’. Aharon Katchalsky 8.1

Introduction

Imagine living in a world where a Richter 9 earthquake raged continuously. In such an environment, engines would be unnecessary. You would not need to even pedal your bicycle: you would simply attach a ratchet to the wheel preventing it from going backwards and shake yourself forwards! At the scale of proteins, Brownian motion is even more furious, and proteins evolved to take advantage of this enormous supply of energy. Feynman showed that the familiar mechanical ratchet could not work in an isothermal environment, lest it violate the Second Law of Thermodynamics (Feynman et al., 1963), and so motor proteins must employ a different strategy to convert random thermal fluctuations into a directed force: they use chemical energy via intermolecular forces to capture ‘favorable’ configurations. The way in which proteins do this is dictated by three factors: their size, the strength and range of intermolecular bonds at physiological conditions and the magnitude of the Brownian fluctuations that constitute their thermal environment. These determine the energy, length and time scales on which protein motors can operate. Roughly speaking, motor proteins trap thermal fluctuations in two ways: by biasing diffusion of small, angstrom-sized, steps (‘small ratchets’), and by rectifying nanometer-sized or larger, diffusional displacements (‘big ratchets’). For reasons that will become clear, biasing a sequence of small Brownian fluctuations is generally called a power stroke, while rectifying a large thermal displacement is called a Brownian ratchet. Said another way, Brownian ratchets move down free energy landscapes in steps much larger than kBT, while power strokes move in steps comparable to or smaller than kBT.1) The distinction is imprecise, but useful, since it 1) The quantity kBT measures the thermal energy

of Brownian motion; its value is

1 kBT Z 4.1 pN nm 4.1 q (25 hC).

10-21

J at 298 K

208

8.2 A Brief Description of ATP Synthase Structure

delineates two extremes in the general mechanism by which proteins use intermolecular attractions to convert chemical energy into mechanical work.1) There are only a few motors that can be regarded as being pure power stroke motors or pure ratchets; most protein motors employ a combination of the two strategies. However, these ‘thoroughbreds’ are good illustrations of the principles. In fact, evolution has designed one protein that joins both ratchet and power stroke motors into one remarkable device: F0F1 ATPase, or ATP synthase, the machine that manufactures the fuel that powers many other protein machines. We will use this protein as our running example. Since this volume is aimed primarily at biologists, our discussion will be mostly qualitative and heuristic. However, it is important to realize that the explanatory cartoons we use are supported by extensive calculations. Omitting them is akin to leaving out the ‘Materials and Methods’ section in an experimental paper: assertions without authority are simply opinions. The supporting quantitative arguments are, perforce, contained in the citations.

8.2

A Brief Description of ATP Synthase Structure

In order to discuss the workings of the F0 and F1 motors we give a brief account of their structure, which is summarized in Fig. 8.1. A more complete account is given in Chapter 5 by Noji in this volume. ATP synthase comprises two rotary motors acting in opposition, each operating on an entirely different physical mechanism. The F0 motor is contained in the membrane-bound portion and employs as its energy source a transmembrane electromotive force. The F1 motor is contained in the soluble portion and is driven by the hydrolysis of ATP. The F1 motor is constructed of a coiled-coil shaft (the g-subunit) surrounded by a hexamer composed of alternating a and b subunits. Nucleotide binding sites nestle in the cleft between subunits; however, only three of the sites are catalytically active, the other three bind, but do not hydrolyze, ATP. The catalytic sites are contained mostly in the b-subunit, with a few but crucial residues contributed by the a subunit. The catalytic sites hydrolyze sequentially and drive the rotation of the g-shaft. Movies showing the motion of the hexamer and the rotation of the g-shaft can be downloaded from a web site given in (Oster and Wang, 2000a; Wang and Oster, 1998). The F0 motor is composed of two portions. Between 10 and 14 c-subunits (depending on the species and/or conditions) are assembled into a cylinder attached to the g-shaft and e-subunit. It interfaces with a second transmembrane assembly consisting of the a and b subunits. By convention, the g-cn-e assemblage is called the ‘rotor’, and the remainder of the protein (the a3b3 hexamer and the d-b2 -a complex) comprises the ‘stator’, although each rotates in the opposite direction. 1) We have included a short Appendix with sev-

eral simple examples that illustrate the differ-

ence between a ‘power stroke’ and a ‘Brownian ratchet’.

8 How Protein Motors Convert Chemical Energy into Mechanical Work δ

δ

β α

F1

α

α3β3 b2

γ

γ

ε

ε c10-14

Fo

β

a

Membrane

a

H+ The structure of ATP synthase. The panel on the left shows a composite based on the pdb coordinates of the known subunits (Pedersen et al., 2000); the right-hand panel is the corresponding structure represented sche-

Figure 8.1.

cytoplasm

b2

c10-14

Membrane

periplasm

matically showing the relative rotation of the F0 and F1 motors and the direction of ion flow through the a-cn subunit interface. Subscripts denote the subunit stoichiometries.

8.3

The F1 Motor: A Power Stroke

The study of protein ATPases has led to a few generalizations that help us understand the mechanism by which these molecular motors convert the energy residing in the nucleotide g-phosphate bond into a directed mechanical force. x

x

x

x

At physiological conditions, the free energy of hydrolysis of one ATP is Z 20 24 kBT; of this, about 8 9 kBT is enthalpic, the balance being entropic. Almost all nucleotide binding sites are nestled in the cleft between protein subunits. The nucleotide is grasped by loops emanating from a parallel b-sheet. In many motors, the force-generating step is associated with the binding of nucleotide to the catalytic site. We propose that this is true of all ATPase motors. In particular, for the F1 motor this is the only way to reconcile all of the biochemical and mechanical measurements with its high mechanical efficiency. After the force has been generated by ATP binding, an ATP is tightly bound in the catalytic site. The role of hydrolysis is to break the tightly bound ATP into two products and weaken the binding so the products can be released and the force-generating cycle can be repeated.

In the F1 motor, the ATP binding site lies asymmetrically in the cleft between the b and a subunits, the majority of the catalytic sites residing in the b subunit. The power stroke is accomplished by a hinge bending motion that swings the top of the b subunit down towards the bottom portion. The bending motion of the b subunit can be measured by the motion of helices B and C that emanate from the b

209

210

8.3 The F1 Motor: A Power Stroke

Figure 8.2. The F1 power strokes are accomplished by the hinge bending motions of b subunits, which are driven by ATP binding to the catalytic sites. (a) During the hinge bending motion, the top part of b rotates about 30h toward the bottom part. This rotation closes the

angle between helices B and C. (b) The hinge bending motion of each b subunit pushes on the off-axis section of the g shaft. The rotation of the g shaft is driven by the coordinated hinge bending motions of all three b subunits (Oster and Wang, 2000a,b, Wang and Oster, 1998).

sheet of the catalytic site, as shown in the ribbon diagram in Fig. 8.2a. The bending of the b subunit by about 30h rotates the central g shaft by pushing on its off-axis section, much like turning a crankshaft (Fig. 8.2b). The energy for the power stroke derives from the nucleotide hydrolysis cycle, which consists of four major steps: Binding

CS + ATP PI Release

! 3

! 1

Hydrolysis

!

CS ATP

2

CS ADP PI (8:1)

ADP Release

CS ADP + PI

! 4

CS + ADP

The nucleotide binding step 1 is the force-generating step and should more properly be expressed as a sequence of binding steps: CS + ATP ! CS ATP ! ! CS ATP

(8:2)

|{z} Binding Transition

Here the progression from weak to tight binding is symbolized by the increasing size of the bonding symbol indicating the progressive annealing of 15 20 hydrogen bonds (and some hydrophobic interactions at the sugar end). The mechanism driving the hinge bending motion of the b is illustrated schematically in Fig. 8.3. Immediately after it diffuses into an open catalytic site, the ATP is bound only weakly. The catalytic site wraps around the ATP by forming more bonds which

8 How Protein Motors Convert Chemical Energy into Mechanical Work

Brownian motion

Figure 8.3. The ATP binding transition from weak to tight proceeds as the catalytic site grasps the ATP in a grip of hydrogen bonds (the ‘binding zipper’). As binding progresses, the catalytic site closes up and pulls the top part of the b subunit toward the bottom part. In this way, the binding free energy is converted to a power stroke with nearly a constant force. During the power stroke some of the binding energy is stored in the elastic deformation of the b-sheet, which acts like a spring. This energy is released during the unbending motion to aid product release and return the subunit to its open state.

Nucleotide

β-sheet Spring

tightens the catalytic site and pulls down the top part of b toward the bottom part. The bonds between the ATP and the catalytic site are formed more or less sequentially, the formation of each bond corresponding to a small drop in binding free energy, that drives a small fraction of the hinge bending motion (Bockmann, 2002, Oster and Wang, 2000a; Sun et al., 2002). When all the bonds have been formed, the ATP is tightly bound in the catalytic site. The overall process from weak to tight binding is called the ‘binding transition’. During the binding transition, the ATP binding free energy is utilized efficiently to generate a bending motion with a nearly constant torque about the hinge region near the b-sheet. The binding transition has two important features (Oster and Wang, 2000a,b, Sun et al., 2002): x

The binding free energy decreases (the binding becomes stronger) nearly monotonically and smoothly during the binding transition. This drives the hinge bending motion of the b subunit and consequently the g shaft rotation in the hydrolysis cycle. In the reverse synthesis cycle, the unbending of the b subunit is driven by the g shaft rotation in the opposite direction, powered by the F0 motor. When the top of the b subunit is forced up and away from the bottom portion, the binding free energy increases (the binding becomes weaker). This progressively releases the nucleotide from the catalytic site (Antes et al., in press).

211

212

8.4 The F0 Motor: A Brownian Ratchet x

By the end of the binding transition, approximately 6 10 kBT of elastic energy is stored in the b-sheet whose loops grasp the nucleotide. Note that the catalytic site should be flexible but not elastic, lest it dissipate too much of the binding free energy by elastic recoil.

To summarize, the power stroke is driven by progressive capturing of relatively small Brownian motions that anneal the nucleotide into the catalytic site. Product release is accomplished by using the free energy of hydrolysis to weaken the product binding so that thermal fluctuations can knock them out of the catalytic site. Thus Brownian motion drives both the power and exhaust strokes.

8.4

The F0 Motor: A Brownian Ratchet

The F0 motor is driven by the ion-motive force, Dmc across a bacterial or mitochondrial inner membrane. The ion-motive force consists of an ion concentration gradient and an electrical potential difference. In most organisms, the ion is a proton. However, much information has been gleaned from anaerobic bacteria whose F0 is driven by a sodium ion motive force. In either case, the transmembrane chemical potential difference in millivolts is given by: Dmc+

kB T log10 c + p ± log10 c + c + Dc = 2.3 e |{z}

[mV]

(8:3)

DpH

where e is the electronic charge, [c]p the ion (sodium or proton) concentration of the periplasm expressed in molars, [c]c the ion concentration of the cytoplasm and Dc is the electrical potential difference across the membrane. Equation 8.3 is a thermodynamic relationship that implies that a concentration difference is equivalent to an electrical potential at equilibrium. Since motors operate out of equilibrium, this turns out not to be true in general. However, Eq. 8.3 points out the need for a mechanism for transforming both an entropic and electrical potential into mechanical work. The basic principle that accomplishes this can be illustrated by the ‘toy’ model shown in. Fig. 8.4. 8.4.1

A Pure Brownian Ratchet

First, consider a rod with a linear array of negative charges (e. g. a DNA strand, etc.) that can freely diffuse through a hydrophilic pore embedded in a membrane as shown in Fig. 8.4a. If a potential, Dc, is imposed across the membrane, then the rod will be propelled to the left and can perform work against a load force, FL. This can be considered as nearly a pure power stroke. The motion of the rod can be viewed as being driven by a driving potential, f, tilted to the left in Fig. 8.4a. The slope of the driving potential gives the motor force driving the rod


Principle of the F0 motor. (a) A pure power stroke. A rod of negative electrical charges passes through a polar transmembrane pore. An electrical potential, Dc imposed across the membrane drives the line of charges to the left. (b) A pure Brownian ratchet. The charged rod passes through a hydrophobic membrane pore. A high concentration of counterion charges (Na or H) on the right can bind to the negative sites and neutralize them so that they can pass through the pore. A second positive ion on the left (not shown) neutralizes the membrane potential. The low concentration of counterions on the left ensures that the bound charges dissociate but do not quickly rebind, so that the bare site cannot re-enter the pore. The attractive bonds between the rod charge and the aqueous solvent molecules rectify the rod’s diffusion. We call this a Brownian ratchet. (c) The power stroke and ratchet can be combined using an L-shaped ‘stator’. An aqueous input channel (colored white) connects via a polar channel (to the right) to the output reservoir. The body of the stator is hydrophobic so that ions cannot leak across the membrane. With this design, the motor has both power stroke and ratchet components. Figure 8.4.

Δψ

kBT FL

φ

(a)

kBT FL

φ (b)

Δψ Dy

FL

kBT (c)

to the left: FM - df/dx. The rod will stall when the motor force equals the load force: FM FL. Because the rod is very small, its motion is stochastic (indicated in Fig. 8.4 by the Gaussian force labeled kBT). In fact, the net drift of the rod driven by the potential is deeply buried in its random motion: at any time the rod is only a tiny bit more likely to move to the left than to the right, and the mean square of the instantaneous velocity is several orders of magnitude larger than the net drift velocity. So the net movement down the potential gradient can only be detected

213

214

8.4 The F0 Motor: A Brownian Ratchet

by looking at correlations over many steps. This is illustrated by example 3 in the Appendix. 8.4.2

A Pure Power Stroke

Next, consider the situation in Fig. 8.4b where the rod passes through a hydrophobic pore. Moreover, there is a difference in the concentration of a positive counterion (e. g. sodium or protons) between the two sides of the membrane. The counterion can bind to and neutralize the negative charges (binding sites) on the rod. A difference in the concentration of a second ion (e. g. potassium), which cannot bind to the sites on the rod, neutralizes the membrane potential. A bare binding site is negatively charged and cannot move into the hydrophobic pore, for that would entail shedding its hydration shell at a considerable energetic cost.1) Because of the high concentration on the right side, the binding sites will be largely neutralized and so the rod can diffuse freely to the left through the pore. However, once a neutralized site has emerged from the pore to the left, the bound positive charge will quickly dissociate and leave the binding site unoccupied. Because of the low concentration on the left side, the binding site is likely to remain unoccupied, which prevents it from moving back into the pore. Thus diffusion to the right is rectified by the ion concentration difference, and the rod moves stochastically to the left, driven by a pure Brownian ratchet. The driving potential, f, in this case looks like a staircase. When the concentration on the left is much lower than that on the right, each step of the staircase potential is much larger than kBT, so that reverse steps are unlikely. This is illustrated by example 4 in the Appendix. The load force, FL, opposing the motion has the effect of ‘tilting’ the driving potential so that the rod must diffuse uphill, against the load force. The two driving forces can be combined as shown in Fig. 8.4c. Here the membrane is horizontal and so the motor must be augmented by a fixed ‘stator’ assembly. The stator body is hydrophobic with two exceptions. First, there is a large aqueous channel that permits the ions from the high concentration reservoir to access the binding sites. Second, there is a polar channel connecting the aqueous channel to the low concentration reservoir. Since the aqueous channel is isopotential with the high concentration reservoir, this arrangement converts the transmembrane potential drop from vertical to horizontal. In this way the membrane potential and the ion concentration difference act in tandem to move the rod to the left, the former driving a power stroke and the latter driving a Brownian ratchet. The actual F0 motor works somewhat differently from the idealized version in Fig. 8.4c. Fig. 8.5a shows two more modifications that are required to make the arrangement resemble the sodium driven F0 motor of the bacterium P. modestum (Dimroth et al., 1999, Oster et al., 2000). The linear array of charged binding sites is first wrapped around a cylinder. This cylinder consists of 10 14 c-subunits, 1) The energy cost of moving a negative charge

into the pore is approximately DG z 45 kBT (Israelachvili and Ninham, 1977).


215

Periplasm

Rotor Dielectric Barrier

Periplasm

Δψ

Membrane Cytoplasm

Input Channel

Rotor Charges

Stator

Δμ -2.3

Stator charge Cytoplasm

Figure 8.5. Operation of the F0 motor (Dimroth et al., 1999). (a) Simplified geometry of the sodium driven F0 motor showing the path of ions through the stator. This is the same arrangement as in Fig. 8.4c, but with the charge array wrapped around a cylinder that is free to rotate in the plane of the membrane with respect to the stator. In addition, a ‘blocking charge’ has been added to the stator to prevent the leakage of charge from the high concentration reservoir (periplasm) to the low concentration reservoir (cytoplasm). (b) Free energy diagram of one rotor site as it passes through the rotor stator interface. Step 1 p 2, the rotor diffuses to the left, bringing the empty (negatively charged) site into the attractive field of the positive stator charge. 2 p 3, once the site

R227

is captured, the membrane potential biases the thermal escape of the site to the left (by tilting the potential and lowering the left edge). 3 p 4, the site quickly picks up an ion from the input channel neutralizing the rotor. 4 p 5, an occupied site being nearly electrically neutral can pass through the dielectric barrier. If the occupied site diffuses to the right, the ion quickly dissociates back into the input channel as it approaches the stator charge. 5 p 6, upon exiting the stator the site quickly loses its ion. The empty (charged) site binds solvent and cannot pass backwards into the low dielectric of the stator. The cycle decreases the free energy of the system by an amount equal to the electromotive force.

depending on the organisms and/or conditions. Second, a ‘blocking charge’ (R232) is present on the stator that prevents leakage of ions between the two reservoirs that would dissipate the ion gradient unproductively. The presence of this blocking charge alters the picture of the rotation of the rotor with respect to the stator substantially. Fig. 8.5b shows the potential experienced by one binding site of the F0 motor. The presence of the blocking charge creates an electrostatic potential well that attracts the rotor binding site as soon as it diffuses into the polar channel. Once trapped in the well, the rotor depends on Brownian fluctuations to escape. However, the membrane potential biases escape by lowering the left side of the electrostatic well so that the rotor charge is much more likely to escape to the left than to the right. Once it escapes into the aqueous input channel, it is quickly neutralized by the positive ions so that it can move freely across the hydrophobic barrier. Note that the membrane potential only biases the Brownian fluctuations to the left, but does not actually drive a power stroke as in Fig. 8.4a. In summary, the F0 motor qualifies as a Brownian ratchet since it rectifies large thermal fluctuations (or long diffusions). The energy for rectification derives from the ion concentration gradient via the short range interactions between ions and

kBT e

ΔpNa +

216

8.5 Coupling and Coordination of Motors

the binding sites (binding and unbinding). It also uses the membrane potential to bias thermal fluctuations that release the rotor site from the attraction of the stator blocking charge: it takes less energy to hop out to the left than to the right. This might be thought of as a partial power stroke, so we see that the classification into power stroke and ratchet is largely a question of definition. An interesting class of ratchet motors is those that use the principle of trapping Brownian fluctuations during their assembly to perform a ‘1-shot’ motor task. Examples include the acrosome of Limulus and Thyone, the spasmoneme of Vorticella (Mahadevan and Matsudaira, 2000), and the polymerizaton of actin that propels lamellipodial protrusion and certain intracellular parasites, such as Listeria and Shigella (Mogilner and Oster, 1996a,b). An important corollary of the ratchet principle, and one that dramatically distinguishes molecular motors from other machines, is that using thermal fluctuations to go uphill in free energy amounts cools off the immediate environment. If the ratchet shown in Fig. 8.4b is coupled to work against a conservative load force, then the process is endothermic: heat is absorbed from surrounding fluid to increase the potential energy of the external agent that exerts the load force. By contrast, if the motor shown in Fig. 8.4a is coupled to work against a viscous load, then the process is exothermic: energy from the electrical potential goes to produce the drift velocity of the rod, which in turn is converted to heat by viscous friction. These microscopic thermal energy transactions lead to some surprising properties of molecular motors. For example, it is possible for the motor to perform more work on a viscous load than the free energy it derives from a reaction cycle! These counterintuitive properties are discussed in more detail in the references (Oster and Wang, 2000b, Wang and Oster, 2002a,b).

8.5

Coupling and Coordination of Motors

Most ATPase motors have more than one catalytic site. During the motor operation, each catalytic site hydrolyzes ATP and contributes to force generation. These catalytic sites generally do not operate independently, but act in concert with other catalytic sites. Two heads of a kinesin dimer are coordinated with each other to generate unidirectional motion and to ensure processivity. ATP synthase has three catalytic sites. AAA motors are hexamers of ATPases, sometimes stacks of two hexamers. The chaperonin Groel is a stacked pair of heptameric rings, each with seven catalytic sites and the portal protein may even be a dodecameric ring of 12 ATPases. Generally, the catalytic sites must act in concert, either firing sequentially, as in F1, or as in Groel, simultaneously in each ring, but alternating between the rings. This coordination is necessary for the proper operation of the motors, but how is it accomplished? In all cases, motor ATPase catalytic sites are too far apart to communicate in any other way than via elastic strain through the intervening protein structure. Although the details of strain coordination differ, a clue can be found in the correla-


tion between the ADP release at one catalytic site and the ATP binding at another. In F1, the strain-induced release of ADP arises from two possible sources. First, the g shaft is asymmetric, so that at every rotational position it strains each catalytic site differently. Second, the intrinsic asymmetry of the protein structure allows the catalytic site to radiate strain differently to the two neighbor sites. The catalytic sites are located in the cleft between adjacent a and b subunits, with the majority of the binding residues in the b subunit (Menz et al., 2001, Stock et al., 2000). This asymmetry allows ATP binding at one catalytic site to propagate a conformational change to the b portion of next catalytic site in the direction of motor rotation, but propagates a different conformational change to the a portion of the previous catalytic site. Thus, one catalytic site can affect the reactions on two neighboring catalytic sites differently. The conformational change directed to the b portion of the next catalytic site can lower the free energy barrier for ADP dissociation, readying it for the next hydrolysis cycle. Indeed this may be the primary structural feature determining the direction of rotation. Because of the unequal symmetry between the F0 and F1 motors, elastic coupling plays an additional role. The F1 motor has three catalytic sites and rotates in three major 120h steps, each with a brief pause at 90 h. On the other hand, the F0 motor has a rotational symmetry that varies between 10 and 14, depending on the source. Therefore, there is no unique ‘stoichiometry’ between the two rotational motions. This symmetry mismatch is not a problem for the motors since the g-shaft that couples them is torsionally flexible. In synthesis, this allows the stochastic F0 motor to deliver a smooth torque to F1, which minimizes dissipation as the nucleotide is unzipped from the catalytic site (Junge et al., 2001; Oster and Wang, 2000a). Thus elastic coupling between subunits of a protein motor provides the means for both coordinating the catalytic cycles and buffering the independent Brownian motions of the subunits. In walking motors the determination of directionality depends on asymmetric structural features of the heads that alternate depending on whether the head is leading or trailing. However, the situation is more complicated since each head has two binding partners: nucleotide and track. One proposal is that strain is generated by binding of the forward head to the track, triggers release of product from that head (Uemura et al., 2002). Also strain is relayed to the rear head via the connecting structure to lower the energy barrier for a particular step in the reaction cycle (for example, hydrolysis, or product release). This particular reaction step, in turn, triggers the release of rear head from the track (Hancock and Howard, 1999). Thus, binding to and release from the track are correlated with the relative positions of the heads. The alternating phases of the two hydrolysis cycles generate unidirectional motion (Peskin and Oster, 1995).

217

218

8.6 Measures of Efficiency

8.6

Measures of Efficiency

We have discussed the molecular principles by which protein motors convert chemical bond energy into mechanical work. However, while the general principles may be the same for all motors, the detailed mechanisms are quite diverse. Therefore, it is frequently useful to determine the efficiency of a motor to provide clues as to its mechanism. The most common mechanical measurement performed on protein motors is to vary the load (the force resisting the motion) and measure the speed. In general, two kinds of load experiments are carried out on protein motors in order to determine their load velocity behavior. In one type, a load is applied to resist the motor’s progress using a laser trap or the elastic stylus of an atomic force microscope. In this case the motor is working against a conservative force (i. e. derivable from a potential energy function) that depends only on the displacement of the motor, f –6f/6j). A second, and generally more experimentally convenient method, is to vary the viscous resistance of the fluid environment of the motor. This is a dissipative force that depends on the motor velocity. The information gleaned from the two kinds of measurements yield different information (Oster and Wang, 2000a, Wang and Oster, 2002a,b). The thermodynamic efficiency, hTD, is generally defined as the ratio of the work done by the motor to the energy input: hTD =

f hdi ± DG

(8:4)

where DG is the free energy drop in one reaction cycle (e. g. from hydrolyzing one ATP, or passing one proton through the motor), and fphdi is the reversible work done by the motor against the conservative load force, f, in one reaction cycle. Here hdi is the average distance covered per reaction cycle, sometimes called the ‘step size’. fphdi is the energy output from the motor because it goes to increasing the potential energy of the external agent that exerts the conservative force. Thus, the thermodynamic efficiency is the ratio of energy output to energy input and it measures the energy conversion efficiency when the motor is operating reversibly, i. e. ‘infinitely slowly’. For a motor working against a constant force (e. g. a laser trap force clamp), Eq. 8.4 can be generalized to the steady state (Oster and Wang, 2000a, Wang and Oster, 2002b) by defining an efficiency: ha

f h vi ± DG hr i

(8:5)

Here hri is the average reaction rate (e. g. hydrolysis cycles per second or average proton flux) and hvi is the average velocity. Strictly speaking, Eq. 8.5 is not a thermodynamic quantity since the steady state need not be the equilibrium state. Nevertheless, it is a well-defined quantity that is less than unity.


In general, the average step size, hdi depends on the load force, f. When hdi is independent of f, we say the motor is tightly coupled. In that case, each reaction cycle is, on average, coupled to a fixed displacement (one motor step) regardless of the load force. A tightly coupled motor has two properties: x x

When the motor is stalled the chemical reaction is also stopped. Increasing the load beyond the stall force drives the motor in the opposite direction and also reverses the chemical reaction.

For a tightly coupled motor, one can show that the stall force is given by fstall -DG/ hdi. At stall, the thermodynamic efficiency is 100 %. When the motor is operating close to stall, the thermodynamic efficiency approaches 100 %, regardless of the motor mechanism. Conversely, a high thermodynamic efficiency suggests only that the motor motion and the chemical reaction are tightly coupled (Wang and Oster, 2002a). Equation 8.5 applies only to the situation where the motor is working against a constant load. For macroscopic motors, inertia is important and the effect of Brownian fluctuations is negligible. Therefore, they tend to move at roughly a constant velocity (at least on short time scales), and so the friction force on the motor is approximately constant. In this case, the friction force can be treated effectively as a conservative force and the efficiency given by Eq. 8.5 is well defined. Thus, for a macroscopic motor, we generally do not have to worry whether it is working against a conservative force or a friction force. That is, for a macroscopic motor moving with roughly a constant velocity, both a viscous drag and a conservative load simply oppose the motor motion and they have approximately the same effect on the motor. For a protein motor, the situation is very different. Because protein motors are very small, the effect of inertia is negligible and the motor is driven mostly by the random Brownian force. The instantaneous velocity changes direction very rapidly and its absolute value is several orders of magnitude larger than the average velocity. Consequently, the drag force on the motor is stochastic and cannot be treated as a conservative force. The effect of a conservative load force on the motor is different from that of a viscous drag force that simply opposes the motor motion in any direction and whose magnitude increases with the velocity.1) Therefore, for protein motors, it is necessary to distinguish the case where the motor is loaded with a conservative force and the case where the motor is loaded with a viscous drag. When a protein motor works against a viscous load, the thermodynamic efficiency defined above does not apply. A commonly employed measure of efficiency in this case is the Stokes efficiency, defined by replacing f in Eq. 8.5 with the average viscous drag force, fD zhvi. Here z kBT/D is the drag coefficient and D is the diffusion coefficient, which can be computed or measured independently (Wang and Oster, 2002a): 1) A conservative load force tends to drive the

motor backwards (opposite to the positive motor direction) and its magnitude is independent of the velocity. A viscous drag simply damps the motor velocity, while the Brownian force and/or the chemical reaction excite the

velocity. If the chemical reaction is halted, the viscous drag will relax the motor to thermal equilibrium with the surrounding fluid, while the conservative force will drive the motor in the opposite direction.

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8.7 Discussion

hStokes =

z hvihdi z hvi2 ; or hStokes = ± DG hr i ± DG

(8:6)

Although zhvi2 has the dimension of energy per unit time, zhvi2 is not the rate of the work done by the motor motion on the fluid medium1). Indeed, it is not clear what energy per unit time zhvi2 measures. However, since zhvi2 increases with hvi, the quantity zhvi2 does measure some aspect of the motor’s mechanical performance. So the Stokes efficiency is the ratio of this ‘mechanical performance’ index to the energy supply. Of course, for Eq. 8.6 to be a valid measure of efficiency, it has to satisfy hStokes J 100 %. This is true, but the proof is not trivial. One can show that Eq. 8.6 measures how close the motor comes to delivering a constant force (Oster and Wang, 2000a, Wang and Oster, 2002a,b). Given the free energy supply – DG, the maximum average velocity is achieved if – DG is utilized to generate a roughly constant driving force (independent of motor position). When the driving force is constant, the Stokes efficiency is 100 %. When the Stokes efficiency is close to 100 %, the driving force is close to a constant force (Wang and Oster, 2002b). A high Stokes efficiency for the F1 motor was one of the factors that implicated the progressive binding of ATP as the force-generating process, for only in this fashion could a constant torque be generated (Oster and Wang, 2000a). To summarize, if the thermodynamic efficiency, hTD, is close to 1, this means that the motor motion is tightly coupled to the chemical reaction. However, when the Stokes efficiency, hStokes, is close to 1, this means that the motor is delivering a nearly constant force. Since the Stokes and thermodynamic efficiencies measure different properties of the motor, it is worthwhile to measure both efficiencies experimentally.

8.7

Discussion

The basic principle underlying force generation in molecular motors seems at first glance pretty simple: proteins use short range attractive intermolecular forces to bias small thermal fluctuations or to rectify larger diffusions. The former we call ‘power strokes’, the latter ‘Brownian ratchets’. But the details are all important, and they are devilishly diverse. At some level of abstraction the operation of a molecular motor can be viewed as the motion of a point moving down a multidimensional free energy surface (Bustamante et al., 2001, Oster and Wang, 2000a, Wang et al., 1998). Indeed, many models start from this viewpoint and try and deduce what the laws of physics can say about the general properties of the surface. While useful from the viewpoint of theory, models at this level of abstraction are likely to be unsatisfying to biologists who seek a more mechanistic understanding, 1) zhvi should not be confused with zhv i. By the equipartition theorem of statistical meachanics, 2

2

zhv2i equals zkBT m 1 at equilibrium, where m is the motor mass. zhv2i is several orders of magnitude larger than zhvi2.

A1 Example Models to Illustrate the Difference between Ratchets and Power Strokes

akin to how an engineer would understand the design and operation of an automobile motor. This involves dealing with the details of protein structure. The study of molecular motors, more than most other areas in biology, draws on a diversity of fields. Biochemistry elucidates the kinetics of the energy supplying reactions and thermodynamics establishes constraints on the energetic transactions. Mutation studies isolate the key functional amino acids and mechanical measurements provide criteria that circumscribe possible mechanisms. Finally, microscopy and X-ray crystallography provides the sine qua non structures, for it is nearly impossible to deduce how a machine works without knowing what it looks like. However, all of these studies combined cannot produce a mechanistic theory of how motor proteins work; this requires the unifying power of mathematical modeling. For until the operation of a motor can be formulated as equations and solved, our knowledge is, at best, qualitative and uncertain, hardly better than a plausible cartoon that may, or may not, obey the laws of chemistry and physics and in any event cannot be compared quantitatively with experiments.

Appendix


Here we discuss four model examples to illustrate the role of Brownian fluctuations and the classification of power strokes and Brownian ratchets. For simplicity, we consider the one-dimensional motion of an object.

A1.1

Example 1: A power stroke without Brownian fluctuations

Suppose the object is deterministically moved forward a fixed unit distance, Dx, per unit time, Dt (e. g. 0.01 nm in 1 ms). The trajectory of the object is shown in Fig. A1a. The object moves forward uniformly with a velocity of (Dx)/(Dt). This model example is a pure power stroke without Brownian fluctuations. It is mostly relevant for macroscopic motors. Because of the large inertia, macroscopic motors tend to move at a roughly constant velocity and the effect of Brownian fluctuations is negligible. For protein motors, because of the small size, at room temperature the motion is dominated by Brownian fluctuations. One may argue that this is a hypothetical example of a power stroke protein motor at zero temperature.

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A1.2 Example 2: A power stroke with Brownian fluctuations

a

400

b 400

300

x

300

200

x

100

200

100

0

0

0

100

200

t

300

400

0

100

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t

c

400

d

400

300 300

x

200

x 200

100 100 0 0 0

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200

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0

t (a) Trajectory of the object for model 1. The object moves forward uniformly with an average velocity of hvi Dx/Dt Dx/1. (b) Trajectory of the object for model 2. The object is driven by a constant force and is subject to Brownian fluctuations. The average velocity of the object is hvi Dx/Dt. (c) Trajectory of the object for model 3. The object is moved by Brownian fluctuations, and the net drift comes from biasing fluctuations. The average velocity of the object is hvi 0.92pDx/ Dt. The free energy consumption per unit length

Figure A1.

1000

2000

3000

4000

t is the same as that in Example 2. Examples 2 and 3 have similar statistical behaviors, and so it is experimentally difficult to distinguish between them. (d) Trajectory of the object for model 4. The object is moved by Brownian fluctuations, and the net drift comes from rectifying large fluctuations. The free energy consumption per unit length is the same as that in Examples 2 and 3 but the average velocity of the object is hvi 0.1 Dx/Dt, significantly lower than that of Examples 2 and 3.

Since protein motors can only function in a certain temperature range and in a certain fluid medium, this model is not really relevant for protein motors, and so we must modify it to incorporate Brownian fluctuations.

A1.2

Example 2: A power stroke with Brownian fluctuations

Suppose, in addition to the deterministic unit forward displacement per unit time, in each unit time the object makes n 10 independent random moves. Each random move is one unit displacement either forward or backward with equal prob-


ability, p (e. g. flipping a fair coin: p 0.5). This random motion is added to simulate Brownian fluctuations. The trajectory of the object is shown in Fig. A1b. The average velocity of the object is hvi Dx/Dt, but the motion is stochastic. If we view the displacement in each unit time as the ‘instantaneous velocity’ for that unit time, then the instantaneous velocity is much larger than the average velocity. The inset shows the details of the trajectory; it is evident that the deterministic drift is buried in the random motions and can be detected only by looking at long time correlations. The number of random moves per unit time is proportional to the diffusion coefficient, D, of the object: 2DDt = n(Dx)2. This allows us to calculate the diffusion coefficient from which the drag coefficient, z, is computed from the Einstein relation: z kBT/D. In this example, the constant force driving the deterministic forward motion is f zhvi and the free energy consumption per unit length is zhviDx. For n 10, the free energy consumption per unit length is 0.2 kBT. A model such as this is appropriate for describing a charged object, such as a DNA strand, driven through a fluid medium by an electrical potential gradient.

A1.3

Example 3: A Brownian ratchet that biases fluctuations

In the two examples above, the net drift is caused directly by a driving force (e. g. an electric potential gradient), and the Brownian fluctuations do not affect the net drift. Next, we consider situations where the net drift is actually caused by biasing or rectifying Brownian fluctuations. In the absence of other influences, the forward and backward fluctuations have the same probability. However, if internal barriers are established to block, partially or completely, the backward fluctuations, then the object is more likely to fluctuate forward. Suppose in each unit time, Dt, the object makes 10 independent random moves (fluctuations). Each random move is 1 unit displacement, Dx, either forward or backward with equal probability (p 0.5). Suppose that each time the object passes a multiple of 5 q Dx, a barrier is established at that location. At a barrier, the object can fluctuate forward in two ways: a forward Brownian fluctuation and a backward Brownian fluctuation that is reflected by the barrier. The object can move back past the barrier only if a backward Brownian movement ‘breaks’ the barrier. The probability of breaking a barrier depends on the strength of the barrier. Let pf be the probability of forward fluctuation at the barrier and pb be that of backward fluctuation. DG/kBT log(pf/pb) gives the free energy (in units of kBT) required to break the barrier. If we use pf 0.73 and pb 0.27, the corresponding free energy drop at the barrier is 1 kBT. Since the barriers are separated by 5 q Dx, the free energy consumption per unit length is 1/5 0.2 kBT, the same as that used in Example 2. The trajectory of the object is shown in Fig. A1c. The average velocity of the object is hvi 0.92 q Dx/Dt, similar to Example 2. Because of the small free energy drop (1 kBT) associated with each barrier, it only partially blocks backward fluctuations.

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A1.4 Example 4: A Brownian ratchet that rectifies fluctuations

The inset shows the details of the trajectory. This model is an example of a Brownian ratchet that biases small fluctuations. The motion of the object is stochastic and indistinguishable from that of Example 2 where the object is driven by a constant force and subject to Brownian fluctuations. In this example, the object is directly moved by Brownian fluctuations. The net drift results from biasing fluctuations. For this reason, this model can be classified as a Brownian ratchet. However, it has the same phenomenological behavior as the power stroke motor in Example 2, and so it is very difficult and unnecessary to experimentally distinguish it from a power stroke motor.

A1.4

Example 4: A Brownian ratchet that rectifies fluctuations

Finally, consider the model Example 3, but now suppose that in each unit time, Dt, the object makes 10 independent random moves (fluctuations). Each random move is one unit displacement, Dx, either forward or backward with equal probability (p 0.5). Now suppose that each time the object passes a multiple of 100 q Dx, a barrier of 20 kBT is established at that location. The barrier is very high: the probability of a forward fluctuation is pf 1 – 3.8 q 10 11 z 1 and the probability of a backward fluctuation that surmounts the barrier is pb 3.8 q 10 11 z 0. Since the barriers are 100 q Dx apart, the free energy consumption per unit length is 20/100 0.2 kBT, the same as that in Examples 2 and 3. The trajectory of the object is shown in Fig. A1d. The average velocity of the object is hvi 0.1Dx/Dt, significantly lower than that of Examples 2 and 3. Because of the large free energy drop associated with each barrier, it almost completely blocks backward fluctuations. Thus, this model is a Brownian ratchet that rectifies large fluctuations. The stochastic motion of the object is different from that of Examples 2 and 3. The object advances slowly in large ratchet steps: once it passes a multiple of 100Dx, it almost never goes back. One can add a ‘load’ to the above examples by decreasing the probability of a forward fluctuation and increasing that of a backward fluctuation at every location. One can also combine the power stroke and ratchet by adding both a deterministic motion and larger barriers, or alternatively by adding both small barriers and larger barriers. But this would complicate the model and obscure the simplicity of the distinctions we are trying to illustrate. However, we should point out that there is a way to use the time series data itself to estimate how much of the motor driving force can be ascribed to ratchet and power stroke contributions. This involves constructing an Effective Driving Potential from the time series data of the motor; this is discussed in Wang and Oster (2002a).

A2 A Closer Look at Binding Free Energy


The simple description given in the text of ATP binding to the catalytic site of F1 or of ions binding to the rotor of F0, conceals a great deal of complexity because it neglects the role of solvent effects. Charges in aqueous solution are always hydrated, surrounded by a shifting cohort of hydrogen bonded waters. Before ATP can bind to the catalytic site to initiate the hydrolysis cycle (or sodium binding to the rotor charge in the F0 motor cycle), both must shed their water coats. The shedding of the water coats is progressive as the hydrogen bonds form between ATP and the catalytic site. Before each hydrogen bond can form, the hydration

TΔS

6 7 9 8 5

3 2

4 1

−ΔG 2

-H

Enthalpic and entropic changes during desolvation and binding can be followed by plotting - DH, vs. TDS. In these coordinates, the free energy change, DG, is plotted as linear contour lines decreasing up and to the right. The formation of one hydrogen bond between the enzyme and nucleotide can be represented schematically as a reversible path showing a single desolvation and binding process. In state 1, the enzyme and nucleotide sites are hydrated (solid circles). Removing a water molecule from one site entails an enthalpic increase, DH12. This is followed by an entropic increase, TDS23, as the water escapes into solution. Finally, the removed water hydrogen bonds with other waters resulting in an enthalpic decrease, DH34. Similar changes accompany the release of a water molecule from the other site during the Figure A2.

transition from state 4 to state 7. Now the two empty sites must be brought close together, entailing an entropic decrease, TDS78, and an enthalpic decrease, DH89 as the sites bind. Thus the overall free energy change, DG19, has enthalpic and entropic components (black bars) that depend on many factors, especially whether the water binds more strongly to the sites than to other waters. Note that the sequence 1 p 9 is only meant to show the enthalpic and entropic changes. It does not represent the actual sequence of what occurs during the desolvation and binding process. In particular, 2 p 3 (the removed water diffusing into solution) and 3 p 4 (the removed water bonding with other waters) occur simultaneously and cannot be separated.

225

226


water molecules must be shed from the donor and receptor just before they form the bond (otherwise they will be re-hydrated quickly). Consider the overall process of the formation of one hydrogen bond between ATP and the catalytic site. This entails a number of energetic and entropic transactions. We can plot the process schematically as the path shown in Fig. A2. (This path does not represent the actual non-equilibrium process, but rather a ‘reversible work’ path to illustrate the separate entropic and enthalpic transactions). We see that, even in the simplest case, a single association event entails four enthalpic and four entropic changes when water molecules break their hydrogen bonds to charged sites, escape into solution, re-bond with other waters, and finally two sites associate. Clearly, solvent effects can tip the free energy balance, but it is seldom easy to compute how. The binding transition that generates the power stroke as embodied in Eq. 8.2 and Fig. 8.3 is also an oversimplified description. The hydrogen bonds are not arrayed in a linear sequence (as suggested by the term ‘binding zipper’), nor are they either ‘on’ (zipped) or ‘off’ (unzipped). The bonding surfaces are complex, hydrogen bonds have a finite range and angular dependence, and thermal motions create a stochastic pattern of graded bonding interactions. Nevertheless, molecular dynamics studies demonstrate that the free energy changes gradually and nearly linearly as the nucleotide unbinds or binds to the catalytic site.


References Antes, I., et al. 2002. The unbinding of ATP from the catalytic site of F1-atpase. Biophys. J. (In Press). Bockmann, R. 2002. Nanoseconds molecular dynamics simulation of primary mechanical energy transfer steps in F1-ATP synthase. Nat. Struct. Biol. 9: 198 202. Bustamante, C., et al. 2001. The physics of molecular motors. Acc. Chem. Res. 34: 412 420. Dimroth, P., et al. 1999. Energy transduction in the sodium F-ATPase of Propionigenium modestum. Proc. Natl. Acad. Sci. USA 96: 4924 4929. Hancock, W. and J. Howard. 1999. Kinesin’s processivity results from mechanical and chemical coordination between the ATP hydrolysis cycles of the two motor domains. Proc. Natl Acad. Sci. USA 96: 13147 13152. Israelachvili, J. and B. Ninham. 1977. Intermolecular forces the long and short of it. J. Colloid Interface Sci .58: 14 25. Junge, W., et al. 2001. Inter-subunit rotation and elastic power transmission in F0F1-ATPase. FEBS Lett. 251: 1 9. Mahadevan, L. and P. Matsudaira. 2000. Motility powered by supramolecular springs and ratchets. Science 288: 95 99. Menz, R., et al. 2001. Structure of bovine mitochondrial F1-ATPase with nucleotide bound to all three catalytic sites: implications for the mechanism of rotary catalysis. Cell 106: 331 341. Mogilner, A. and G. Oster. 1996a. Cell motility driven by actin polymerization. Biophys. J. 71: 3030 3045. Mogilner, A. and G. Oster. 1996b. The physics of lamellipodial protrusion. Euro. Biophs. J. 25: 47 53.

Oster, G. and H. Wang. 2000a. Reverse engineering a protein: The mechanochemistry of ATP synthase. Biochim. Biophys. Acta 1458: 482 510. Oster, G. and H. Wang. 2000b. Why is the efficiency of the F1 ATPase so high? J. Bioenerg. Biomembr. 332: 459 469. Oster, G., et al. 2000. How F0 -ATPase generates rotary torque. Proc. Roy. Soc. 355: 523 528. Pedersen, P., et al. 2000. ATP Synthases in the year 2000: evolving views about the structures of these remarkable enzyme complexes. J. Bioenerget. Biomembr. 32: 325–332. Peskin, C. S. and G. Oster. 1995. Coordinated hydrolysis explains the mechanical behavior of kinesin. Biophys. J. 68: 202s 210s. Stock, D., et al. 2000. The rotary mechanism of ATP synthase. Curr. Opin. Struct. Biol. 10: 672 679. Sun, S., et al. 2002. Elastic energy storage in F1-ATPase. Biophys. J. (In press). Uemura, S., et al. 2002. Kinesin microtubule binding depends on both nucleotide state and loading direction. Proc. Natl Acad. Sci. USA 99: 5977 5981. Wang, H. and G. Oster. 1998. Energy transduction in the F1 motor of ATP synthase. Nature 396: 279 282. Wang, H. and G. Oster. 2002a. Ratchets, power strokes, and molecular motors. Appl. Phys. A 75: 315–323. Wang, H. and G. Oster. 2002b. The Stokes efficiency for molecular motors and its applications. Europhys. Lett. 57: 134 140. Wang, H., et al. 1998. Force generation in RNA polymerase. Biophys. J. 74: 1186 1202.

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9 Molecular Motor Directionality Sharyn A. Endow

9.1

Introduction

Molecular motors (Howard, 2001) are specialized enzymes that use the energy from ATP hydrolysis to move along a filament. Remarkably, motor proteins move unidirectionally along their filament. The cytoskeletal motors, the focus of this chapter, fall into three groups the kinesins, dyneins, and myosins that move towards either the fast polymerizing/depolymerizing plus ends or more stable minus ends of microtubules or actin filaments. Other molecular motors, such as helicases and RNA polymerases, also show polarity of movement along their substrate and may employ mechanisms for directionality similar to those emerging from work on the cytoskeletal motors. Because the plus ends of microtubules or actin filaments are located distal to the nucleation sites, at the periphery of the cell while the minus ends are adjacent to the centers of nucleation, usually more internal to the cell, directional movement allows the cytoskeletal motors to perform specific cellular functions. For example, conventional kinesin, a plus-end microtubule motor, is thought to be the motor involved in fast axonal transport of vesicles and organelles from the neuronal cell body to the presynaptic region of the axon (Amaratunga et al., 1995, Hirokawa et al., 1991, Hurd and Saxton, 1996), while cytoplasmic dynein, a minus-end motor, transports membranous organelles in the opposite direction, from the distal regions of the axon to the cell body (Martin et al., 1999, Schnapp and Reese, 1989, Wang et al., 1995).

9.2

Reversed Kinesins

The discovery of the first kinesin (Brady et al., 1982, Vale et al., 1985a), known as conventional kinesin, and the demonstration of its plus-end motility (Vale et al., 1985b) led to the idea that kinesin would be responsible for plus-end movement along microtubules in the cell and the previously discovered dyneins would per-

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9.2 Reversed Kinesins

form minus-end microtubule movement (Vale et al., 1985b). This idea was largely dismantled with the discovery that Ncd, a Drosophila motor which is clearly related to kinesin by sequence homology, moves to microtubule minus ends (McDonald et al., 1990, Walker et al., 1990). The minus end directionality of Ncd is consistent with the presence of microtubule minus ends at the spindle poles (Euteneuer and McIntosh, 1981, Telzer and Haimo, 1981) and the role of the motor in focusing the ends into poles in oocyte meiotic spindles (Endow and Komma, 1997, 1998, Matthies et al., 1996) and attaching centrosomes to poles in embryo mitotic spindles (Endow and Komma, 1996). The kinesin motors are now known to comprise a large family of related proteins that perform different roles in vesicle and organelle transport (Hirokawa, 1998), and spindle assembly and chromosome motility during meiosis and mitosis (Endow, 1999b, Hunter and Wordeman, 2000, Inoué and Salmon, 1995). Kinesin proteins, including orthologs of Ncd, have now been found in a broad range of evolutionarily diverse organisms that include yeasts, mammals and plants (Kim and Endow, 2000). These motors are thus likely to exist in all eukaryotes. The minus-end kinesins differ from the plus-end kinesins in domain organization: the motor or catalytic domains with their highly conserved nucleotide-binding motif and adjacent microtubule-binding site are located at the C terminus of the coiled-coil dimerization domain in the minus-end kinesins, while they are present at the N terminus of the coiled coil in the plus-end kinesins (Fig. 9.1). The functional implications of this ‘reversed’ domain organization are still uncertain, but, together with structural differences in how the motor is joined to the coiled coil, the altered interactions between the catalytic and coiled coil

Figure 9.1. Kinesin and Ncd structures. Structures of dimeric conventional kinesin (PDB 3KIN) (Kozielski et al., 1997) and dimeric Ncd (PDB 2NCD) (Sablin et al., 1998) are shown. The motor or catalytic domains with their highly conserved ATP- and microtubule-binding regions are N-terminal to the coiled-coil stalk of kinesin but C-terminal to that of Ncd. The neck of conventional kinesin (black) consists of an Z 12-residue neck linker shown space-filled (rat

kinesin V327-T338) that forms extensive contacts with the catalytic domain, together with the proximal, less stable region of the coiled coil (rat kinesin A339-K354) (Kozielski et al., 1997). The neck of Ncd (black) comprises the distal, less stable end of the coiled coil (R335N348) (Endow, 1999c, Sablin et al., 1998), which makes extensive contacts with the catalytic domains.

9 Molecular Motor Directionality

domains may underlie differences in motor directionality. We do know that the correlation between minus-end directionality and having a C-terminal motor domain is invariant so far all the C-terminal motor kinesins whose directionality has been determined have been found to be minus-end motors (Endow, 1999a). It therefore seems likely that the C-terminal motor kinesins will all turn out to be minus-end motors. 9.2.1

Chimeric Kinesin Motors

The finding, more than 10 years ago, of a kinesin motor with the opposite polarity of movement as the rest of the motor proteins in its family opened the field to studies of the molecular basis of directionality and the relationship between motor directionality and function. Despite this long lead time, progress in identifying the region of the kinesin motor proteins required for directionality has been made only relatively recently. This information has come through the construction and analysis of chimeras between plus-end conventional kinesin and minus-end Ncd. The first functional chimeras were reported in 1997, 7 years after the finding that Ncd was a minus-end motor. The long gap in time was largely due to the inability of workers to produce functional hybrid motors because it was not apparent where the junctions between the motors should be made. It was only after reports of the first crystal structures for conventional kinesin and Ncd motor domains that conserved structural elements could be identified and the correct junctions between the two motors made. In one case, however, the correct junctions were made by chance (Henningsen and Schliwa, 1997). The hybrid motors that were genetically engineered from kinesin and Ncd replaced the conserved motor domain of conventional kinesin (Case et al., 1997, Henningsen and Schliwa, 1997) or Ncd (Endow and Waligora, 1998) with the corresponding region from the motor of opposite directionality. Unexpectedly, the direction of movement of the chimeric motors was, in each case, the same as the nonmotor region of the construct (Fig. 9.2). These findings gave rise to the idea that the non-motor region of conventional kinesin or Ncd directs the motor to the microtubule plus or minus end. Based on its unusual properties, it was thought likely that the region involved in motor directionality was the neck, just adjacent to the conserved catalytic domain or motor core. The role of the Ncd neck in motor directionality was demonstrated by mutating two residues at the neck/motor junction in an Ncd kinesin chimera consisting of the Ncd non-motor region joined to a kinesin motor domain, which moved to the microtubule minus end. The mutated chimera was a slow plus-end motor that reversed the directionality of the original chimera (Endow and Waligora, 1998) (Fig. 9.2). The slow plus-end motility of the mutated chimera was attributed to inactivation of the Ncd neck together with conformational changes within the motor core that displaced the motor towards the plus end. Slow plus-end movement was subsequently reported for neck-mutated native conventional kinesin or Ncd (Case et al., 2000, Sablin et al., 1998; Fig. 9.2). The slow plus-end motility of the mutants was again attributed to conformational

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9.2 Reversed Kinesins

x x

Kinesin

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xx

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Motor core

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xx

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Figure 9.2. Kinesin and Ncd motors used in directionality studies. Motor directionality was analyzed by constructing chimeras between plus-end conventional kinesin and minus-end Ncd (Case et al., 1997, Endow and Waligora, 1998, Henningsen and Schliwa, 1997) and mutating the hybrid motors (Endow and Waligora, 1998), or mutating native kinesin or Ncd (Case et al., 2000, Sablin et al., 1998). The catalytic domain or motor core of conventional kinesin

x x Mutations

is white; that of Ncd is black. The catalytic domains of conventional kinesin and Ncd both show slow plus-end motility, as revealed by the mutated Ncd-kinesin chimera (bottom) or the mutated native motors. This slow plus-end movement of conventional kinesin may be amplified by the neck and/or neck linker. The neck of Ncd is required for minus-end directionality of the Ncd kinesin chimera and native Ncd motor (bottom).

changes of the conserved motor core that results in a plus-end displacement of the motor along the microtubule. The nature of the proposed conformational changes is not known, but they may be associated with ADP release or ATP binding by the motor. The studies of chimeric motors (Endow and Waligora, 1998), mutated chimeric motors (Endow and Waligora, 1998) and a neck-mutated native motor (Sablin et al., 1998), have led to the conclusion that residues of the Ncd neck are needed for minus-end directionality. The Ncd neck must somehow overcome the slow plusend movement intrinsic to the motor core to direct movement towards the microtubule minus end. The findings for conventional kinesin are less clear, because the stalk/neck could simply amplify the slow plus-end movement of the catalytic domain rather than conferring plus-end directionality on the motor. The neck of conventional kinesin differs from the neck of Ncd in that it consists of an Z 12-residue region, the neck linker, together with the proximal end of the coiled-coil stalk, instead of the distal end of the coiled-coil stalk as in Ncd (Fig. 9.1). The Ncd neck also shows extensive interactions with the catalytic domains of the dimeric motor, which is not observed for the helical region of the neck of conventional kinesin (Fig. 9.1). Instead, the neck linker of conventional kinesin shows extensive interactions with residues of the motor core in the dimer crystal structure (Kozielski et al., 1997). Thus, the mechanism of directionality determination could differ for Ncd and conventional kinesin due to differences in their neck structures and interactions of their necks with the catalytic domains.


9.2.2

A Neck Mutant

Further information regarding the role of the Ncd neck in motor directionality has come from an analysis of a neck mutant that converts minus-end Ncd into a motor that lacks directionality and moves in either direction on microtubules (Endow and Higuchi, 2000). Gliding assays of the neck mutant showed movement towards either the microtubule plus or minus end. Movement in either direction was also observed in gliding assays when a motor core residue that touches the neck residue in crystal structures was mutated. Single-motor laser trap assays revealed a directional conformational or angle change of wild-type Ncd that occurs upon binding of the motor to a microtubule. This displacement was biased towards the minus end for wild-type Ncd, but occurred in either the plus or minus direction for the Ncd neck mutant. The interpretation of these results is that a movement, probably of the stalk/neck, occurs upon binding of the Ncd motor to a microtubule. The directional bias of this movement is dependent on neck motor core interactions and causes wild-type Ncd to move towards the microtubule minus-end (Endow and Higuchi, 2000). These findings imply that the stalk/neck acts like a lever arm to amplify movements of the catalytic core. There is now considerable evidence that a region of the kinesins, either the neck linker of conventional kinesin or the stalk/neck of the kinesin proteins, acts mechanically to amplify small displacements or angle changes of the catalytic domain (Case et al., 2000, Endow and Higuchi, 2000, Endow and Waligora, 1998, Sablin et al., 1998). Unlike the myosins, however, direct evidence that the coiled-coil stalk/neck of the kinesins acts like a lever arm is still lacking. This idea, in fact, has been largely dismissed for conventional kinesin because the neck linker at the end of the helical stalk represents a flexible domain between the stalk and catalytic core that was disordered in the first crystal structure of a conventional kinesin motor domain (Kull et al., 1996). It is therefore difficult to envision how the stalk could amplify force produced by the catalytic domain. Instead, the neck linker itself is thought to amplify the movements of the conventional kinesin catalytic domain (Case et al., 2000, Rice et al., 1999). Another possibility is that the stalk could amplify movements of the catalytic domain and transduce force when the neck linker is immobilized by interactions with the catalytic domain, as observed in subsequent crystal structures of conventional kinesin (Kozielski et al., 1997, Sack et al., 1997) (Fig. 9.1). The neck of Ncd and other C-terminal kinesin motors, in contrast, lacks a neck linker and consists of the distal end of the a-helical coiledcoil stalk joined directly to the conserved motor domain (Fig. 9.1). The absence of a neck linker in Ncd allows the catalytic domain to interact extensively with the stalk/ neck, whereas the catalytic domain interacts with the neck linker but not the stalk/ neck in conventional kinesin. This means that the stalk/neck could function somewhat differently in the plus- and minus-end kinesin motors, but could potentially act to propel the motor either towards the plus or minus end of the microtubule by amplifying the directional components of movement.

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9.3 Backwards Myosins

9.3

Backwards Myosins

Compared to the kinesins, minus-end myosins are relative newcomers to the motors field. A myosin that moves ‘backwards’ relative to the known myosins was discovered only in 1999, 9 years after Ncd was found to be a reversed kinesin motor. Myosin VI, the first minus-end myosin to be discovered, was expressed and tested for directionality in gliding assays after workers noted that the protein predicted from the gene sequence contained a 53-residue insertion that is not present in other myosins. They then demonstrated that myosin VI moves to the pointed or minus ends of actin filaments, instead of the barbed or plus ends (Wells et al., 1999). The insertion that distinguishes myosin VI from other myosins is present next to the so-called ‘converter’, an element of myosin adjacent to the catalytic domain that appears to be analogous to the neck linker of conventional kinesin (Fig. 9.3). Myosin VI differs from other myosins not only by the presence of the 53-residue insertion next to the converter, but also by smaller insertions in the motor domain and in the number of IQ motifs, or light chain-binding sites. However, the motor domain of myosin VI is present at the N terminus of the coiled-coil rod as in the plus-end myosins, rather than the C terminus as in Ncd and other minus-end kinesin motors. Myosin VI thus differs in overall structure from the minus-end kinesins with their reversed domain organization. This means that a C-terminal motor domain is not required for minus-end directionality of motor movement. Further, the mechanism of directionality reversal is likely to differ for the minus-end myosins and kinesins, based on the structural differences between the motors. A second backwards myosin, myosin IXb, has been reported recently (Inoue et al., 2002). Remarkably, myosin IXb is a single-headed processive motor (Inoue et al., 2002, Post et al., 2002). The directionality of myosin IXb has yet to be con-

Myosin structure. A structure of myosin II, or conventional myosin, is shown (PDB 1B7T) (Houdusse et al., 1999). The catalytic domain with its highly conserved ATP- and actin-binding sites is at the N-terminus of the ahelical rod (dark gray). The converter (black), a

Figure 9.3.

rigid a/b domain that includes the first three turns of the rod, joins the catalytic domain to the rod. Light chains are shown associated with the rod. The structure shown is a monomer; structures of dimeric myosin motors have not yet been reported.


firmed by another laboratory, however, and the mechanism of its processivity still remains to be determined. The evolutionary relationship between myosins VI and IX also deserves further attention the two subfamilies arise from the same branch of at least one myosin family tree (Hodge and Cope, 2000), raising the question of whether the motors arose independently in evolution. 9.3.1

Chimeric Myosin Motors

Following the discovery that myosin VI is a minus-end motor, workers turned to investigation of the molecular basis of directionality of the myosins, keeping in mind the finding that the neck of the kinesins is implicated in directionality of motor movement. The myosins differ from the kinesins in the structure of the region between the conserved catalytic domain and the a-helical coiled coil, which has been referred to as the neck of the kinesin motors (Endow, 1999c, Kozielski et al., 1997, Sablin et al., 1998). In the myosins, the SH1/SH2 helix at the C terminus of the conserved catalytic domain is followed by the converter and the IQ motifs, or light-chain binding sites, which vary in number in different myosins and separate the converter from the coiled-coil rod. The IQ motifs can be thought of as comprising the helical region of the neck. The strategy adopted by myosin workers for mapping the region of the motor required for directionality was the same as that used by kinesin workers, and involved constructing chimeras between minus-end myosin VI and a plus-end myosin. The first such chimeras were reported recently and were made between myosin VI and myosin V, a plus-end motor (Homma et al., 2001). The constructs substituted regions of myosin VI or myosin V with the corresponding regions from the motor of opposite directionality. Two hybrids replaced the myosin VI motor with or without the SH1/SH2 helix with those of myosin V, and a third hybrid replaced the myosin VI motor, SH1/SH2 helix and converter with those of myosin V (Fig. 9.4). The three chimeras, all of which contained the 53-residue insertion unique to myosin VI, were all plus-end motors. A fourth chimera, that was complementary to the previous three, replaced the myosin V motor, SH1/SH2 helix and converter with those of myosin VI. This chimera, which lacked the myosin VI 53-residue insertion, was a minus-end motor (Fig. 9.4). The conclusion reached in this report was that the motor core determines the direction of myosin movement. This conclusion is not fully warranted, however. The hybrid motors that were reported show that the 53-residue insertion is not sufficient or necessary for minus-end directionality and that the plus-end motility of myosin V resides in the motor core. But the only minus-end chimera reported still contained myosin VI neck residues, including the converter, although it was missing the 53-residue insertion (Homma et al., 2001) (Fig. 9.4). It is thus possible that the myosin VI neck residues that were retained in the hybrid are needed for minus-end directionality, rather than elements within the motor core. The velocity of the minus-end chimera was 5 6-fold reduced compared to native myosin VI, suggesting that the insertion could serve to amplify the minus-end movement of

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9.3 Backwards Myosins

Myosin V

Plus-end chimeras

Motor core SH1/SH2 Converter Insertion IQ motif

Myosin VI

Minus-end chimera

Figure 9.4. Myosin hybrids used in directionality studies. Myosin motor directionality has been analyzed by constructing chimeras between myosin V, a plus-end motor, and myosin VI, a minus-end motor (Homma et al., 2001). The myosin V catalytic domain or motor core, converter and IQ motifs or light chain binding sites, are white; those of myosin VI are black. Three chimeras, all of which contained the myosin V motor core, were plus-end motors, indicating that plus-end motility is associated

with the myosin V motor core. The three hybrid motors all contained the 53-residue insertion unique to myosin VI, indicating that the insertion is not sufficient for minus-end directionality. One chimera, containing the myosin VI motor core, SH1/SH2 helix and converter, was a minus-end motor. This hybrid did not contain the myosin VI 53-residue insertion, indicating that the insertion is not required for minus-end directionality.

the motor. Together with the fact that the insertion does not occur in the known plus-end myosins, the slow velocity of the minus-end chimera lends support to the idea that the insertion facilitates, but is not sufficient for, minus-end motility. Work on the kinesins has shown that plus-end motility appears to be a ‘default’ mode of the motor, presumably due to conformational changes in the catalytic domain that displace the motor towards the plus end (Case et al., 2000, Endow and Waligora, 1998, Sablin et al., 1998). Because this may also be true of the myosins, further constructs are needed to determine whether minus-end directionality of myosin VI maps to the conserved catalytic domain. Three-dimensional reconstructions of myosin VI-decorated actin filaments made from cryoelectron microscopy images have been interpreted to show that the myosin VI lever arm rotates in the opposite direction to that of myosin II, explaining the minus-end directionality of myosin VI (Wells et al., 1999). The converter is believed to undergo structural changes that result in rotation of the light chain-binding domain (Houdusse et al., 1999). Because it lies at the base of the coiled-coil rod,


a model in which the 53-residue insertion adjacent to the converter alters the rotation of the lever arm seems plausible. However, the structure of the insertion is not known and nor are the structural elements with which it interacts, and this information could provide valuable clues regarding the role of the insertion. An alternate possibility is that determinants of motor directionality exist in the myosin motor core, as concluded by other workers (Homma et al., 2001). This would mean that structural elements of the catalytic domain or their movements during the hydrolysis cycle must differ between the plus- and minus-end myosins. Structural differences between the motor domains of the plus-end myosins and myosin VI do exist and may underlie the differences in directionality of the motors, but further work is needed to determine their effects on myosin motility. The finding of a second putative minus-end myosin (Inoue et al., 2002) may provide information that could help elucidate the molecular basis of myosin directionality.

9.4

Bidirectional Dyneins?

Although dynein was the first microtubule motor to be identified (Gibbons and Rowe, 1965), studies on the dyneins have lagged behind those of other cytoskeletal motors. This is primarily due to the extremely large size of the dyneins and the difficulty of purifying or expressing sufficient quantities of the proteins for biochemical or structural studies. Axonemal dyneins and the later discovered cytoplasmic dynein are minus-end microtubule motors (Paschal and Vallee, 1987, Sale and Satir, 1977) and the expectation, until recently, has been that the dyneins will all be minus-end motors. However, the finding of kinesin and myosin motors that move to the minus end of their filament, in contrast to other members of their families, has raised the possibility that plus-end dyneins could exist. To date, there is no conclusive evidence for unidirectional plus-end motility of a dynein, but there are several reports that provide suggestive evidence for bidirectional movement of specific dyneins. The first of these reports concerned a cytoplasmic dynein from the freshwater amoeba, Reticulomyxa (Euteneuer et al., 1988). Active transport attributed to the dynein was observed along polarity-oriented microtubule bundles from lysed amoebae, and was bidirectional. The motility could be switched to unidirectional minus-end transport by phosphorylation, raising the possibility that phosphorylation of the motor regulates its directionality. Further work on the Reticulomyxa motor has yet to be reported, but would presumably involve in vitro motility assays using the purified motor and polarity-labeled microtubules. More recently, a second report of dynein bidirectionality has emerged. Microtubules interacting with de-membranated flagellar axonemes have been reported to show oscillatory backand-forth movement in laser trap assays that has been attributed to outer arm dyneins (Shingyoji et al., 1998). The motility, interpreted to be due to alternating forward and backwards processive stepping by axonemal dyneins along the microtubule, differed from one-dimensional diffusional movement in that it was ATP-de-

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9. Perspectives

pendent and appeared to consist of multiple successive steps in one direction, followed by multiple successive steps in the opposite direction. The movement was attributed to one or a few outer dynein arms and could also be observed in axonemes treated with high salt, which removes outer arms but exposes inner dynein arms. Thus, the oscillating movement could also be a property of inner arm dyneins. These observations are intriguing, but are complicated by several factors. First, there are now many kinesin microtubule motors that have been discovered in various organisms. These kinesins, most of which are plus-end motors, are associated both with cytoplasmic and axonemal microtubules and may be responsible for some or all of the plus-end motility observed in lysed cell preparations or de-membranated axonemes. The large number of kinesin motors in most organisms means that it will be necessary to use purified dynein motors to demonstrate properties of their motility. Second, the dyneins are structurally more complex than the kinesins or even the myosins. The heavy chains are large, 300 400 kDa, and contain multiple ATP-binding motifs (Gibbons et al., 1991, Ogawa 1991), only one of which appears to be involved in force generation by the motor. A model of dynein heavy chain structure, based on sequence homology to proteins whose structures have been determined, consists of an unusual ring of six modules (Mocz and Gibbons, 2001). Axonemal outer arm dyneins consist of two different heavy chains (Sale et al., 1985, Tang et al., 1982), with a third distinctive heavy chain in Tetrahymena and Chlamydomonas (Goodenough and Heuser 1984, Johnson and Wall, 1983), and it is not known whether nucleotide hydrolysis and movement of the heavy chains along the microtubule are coordinately regulated with one another. These structural characteristics suggest that motility of the dyneins could differ substantially from that of the kinesins or myosins. For example, the backwards component of the oscillatory movement, towards the microtubule plus end, observed by Shingyogi et al. (1998) may not be coupled to ATP hydrolysis and thus may not be analogous to steps along the microtubule by conventional kinesin. On the other hand, the finding of a kinesin mutant that moves in either direction along the microtubule (Endow and Higuchi, 2000) provides a precedent for a bidirectional motor and suggests that such variant motors might occur naturally as a consequence of weakened neck interactions with the catalytic domain.

9. 5

Perspectives

It is now apparent from functional studies that the neck of conventional kinesin is not only required for directionality, possibly to amplify movements of the catalytic domain (Case et al., 2000), but it is also essential for processivity (Hancock and Howard, 1998) and coordinated regulation of ATP hydrolysis (Hackney, 1994, Jiang et al., 1997). The neck of Ncd has been implicated in conferring minus-end directionality on the motor (Endow and Waligora, 1998, Endow and Higuchi, 2000, Sablin et al., 1998). The region of myosin analogous to the neck linker of conven-


tional kinesin, the converter, lies between the coiled-coil rod and the catalytic domain. Based on X-ray crystallography studies, the myosin converter contains a flexible joint that acts as a hinge for movement of the lever arm (Dominguez et al., 1998, Houdusse et al., 1999, Rayment et al., 1993). Movements of the kinesin stalk/neck and those of the myosin rod are likely to bias motor directionality, based on findings for Ncd (Endow and Higuchi, 2000) and are probably tightly coupled to nucleotide release or binding by the motor. Disabling the neck uncouples the catalytic domain from the stalk or rod, affecting basic aspects of motor function, such as processivity and regulation of ATP hydrolysis. Further studies should not only illuminate the mechanical basis of motor directionality, but provide insights into the motor mechanism.

References Amaratunga, A., Leeman, S. E., Kosik, K. S. and Fine, R. E. 1995. Inhibition of kinesin synthesis in vivo inhibits the rapid transport of representative proteins for three transport vesicle classes into the axon. J. Neurochem. 64: 2374 2376. Brady, S. T., Lasek, R. J. and Allen, R. D. 1982. Fast axonal transport in extruded axoplasm from squid giant axon. Science 218: 1129 1131. Case, R. B., Pierce, D. W., Hom-Booher, N., Hart, C. L. and Vale, R. D. 1997. The directional preference of kinesin motors is specified by an element outside of the motor catalytic domain. Cell 90: 959 966. Case, R. B., Rice, S., Hart, C. L., Ly, B. and Vale, R. D. 2000. Role of the kinesin neck linker and catalytic core in microtubule-based motility. Curr. Biol. 10: 157 160. Dominguez, R., Freyzon, Y., Trybus, K. M. and Cohen, C. 1998. Crystal structure of a vertebrate smooth muscle myosin motor domain and its complex with the essential light chain: visualization of the pre-power stroke state. Cell 94: 559 571. Endow, S. A. 1999a. C-terminal motor kinesin proteins. In: Guidebook to the Cytoskeletal and Motor Proteins. Edited by Kreis, T. and Vale, R. Oxford, UK: Oxford University Press. Endow, S. A. 1999b. Microtubule motors in spindle and chromosome motility. Eur. J. Biochem. 262: 12 18. Endow, S. A. 1999c. Determinants of molecular motor directionality. Nature Cell Biol. 1: 163 167.

Endow, S. A. and Higuchi, H. 2000. A mutant of the motor protein kinesin that moves in both directions on microtubules. Nature 406: 913 916. Endow, S. A. and Komma, D. J. 1996. Centrosome and spindle function of the Drosophila Ncd microtubule motor visualized in live embryos using Ncd GFP fusion proteins. J. Cell Sci. 109: 2429 2442. Endow, S. A. and Komma, D. J. 1997. Spindle dynamics during meiosis in Drosophila oocytes. J. Cell Biol. 137: 1321 1336. Endow, S. A. and Komma, D. J. 1998. Assembly and dynamics of an anastral:astral spindle: the meiosis II spindle of Drosophila oocytes. J. Cell Sci. 111: 2487 2495. Endow, S. A. and Waligora, K. W. 1998. Determinants of kinesin motor polarity. Science 281: 1200 1202. Euteneuer, U. and McIntosh, J. R. 1981. Structural polarity of kinetochore microtubules in PtK1 cells. J. Cell Biol. 89: 338 345. Euteneuer, U., Koonce, M. P., Pfister, K. K. and Schliwa, M. 1988. An ATPase with properties expected for the organelle motor of the giant amoeba, Reticulomyxa. Nature 332: 176 178. Gibbons, I. R. and Rowe, A. 1965. Dynein: a protein with adenosine triphosphatase activity from cilia. Science 149: 424 426. Gibbons, I. R., Gibbons, B. H., Mocz, G. and Asai, D. J. 1991. Multiple nucleotide-binding sites in the sequence of dynein b heavy chain. Nature 352:640-643. Goodenough, U. and Heuser, J. 1984. Structural comparison of purified dynein proteins

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References with in situ dynein arms. J. Mol. Biol. 180: 1083 1118. Hackney, D. D. 1994. Evidence for alternating head catalysis by kinesin during microtubulestimulated ATP hydrolysis. Proc. Natl Acad. Sci. USA 91: 6865 6869. Hancock, W. O. and Howard, J. 1998. Processivity of the motor protein kinesin requires two heads. J. Cell Biol. 140: 1395 1405. Henningsen, U. and Schliwa, M. 1997. Reversal in the direction of movement of a molecular motor. Nature 389: 93 96. Hirokawa, N. 1998. Kinesin and dynein superfamily proteins and the mechanism of organelle transport. Science 279: 519 526. Hirokawa, N., Sato-Yoshitake, R., Kobayashi, N., Pfister, K. K., Bloom, G. S. and Brady, S. T. 1991. Kinesin associates with anterogradely transported membranous organelles in vivo. J. Cell Biol. 114: 295 302. Hodge, T. and Cope, M. J. 2000. A myosin family tree. J. Cell Sci. 113: 3353 3354. Homma, K., Yoshimura, M., Saito, J., Ikebe, R. and Ikebe, M. 2001. The core of the motor domain determines the direction of myosin movement. Nature 412: 831 834. Houdusse, A., Kalabokis, V. N., Himmel, D., Szent-Györgyi, A. G. and Cohen, C. 1999. Atomic structure of scallop myosin subfragment S1 complexed with MgADP: a novel conformation of the myosin head. Cell 14: 459 470. Howard, J. 2001. Mechanics of Motor Proteins and the Cytoskeleton. Sunderland, MA: Sinauer Associates, Inc. Hunter, A. W. and Wordeman, L. 2000. How motor proteins influence microtubule polymerization dynamics. J. Cell Sci. 113: 4379 4389. Hurd, D. D. and Saxton, W. M. 1996. Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila. Genetics 144: 1075 1085. Inoué, S. and Salmon, E. D. 1995. Force generation by microtubule assembly/disassembly in mitosis and related movements. Molec. Biol. Cell 6: 1619 1640. Inoue, A., Saito, J., Ikebe, R. and Ikebe, M. 2002. Myosin IXb is a single-headed minusend-directed processive motor. Nature Cell Biol. 4: 302 306. Jiang, W., Stock, M. F., Li, X. and Hackney, D. D. 1997. Influence of the kinesin neck

domain on dimerization and ATPase kinetics. J. Biol. Chem. 272: 7626 7632. Johnson, K. A. and Wall, J. S. 1983. Structure and molecular weight of the dynein ATPase. J. Cell Biol. 96: 669 678. Kim, A. J. and Endow, S. A. 2000. A kinesin family tree. J. Cell Sci. 113: 3681 3682. Kozielski, F., Sack, S., Marx, A., Thormählen, M., Schönbrunn, E., Biou, V., Thompson, A., Mandelkow, E.-M. and Mandelkow, E. 1997. The crystal structure of dimeric kinesin and implications for microtubule-dependent motility. Cell 91: 985 994. Kull, F. J., Sablin, E. P., Lau, R., Fletterick, R. J. and Vale, R. D. 1996. Crystal structure of the kinesin motor domain reveals a structural similarity to myosin. Nature 380: 550 555. Martin, M., Iyadurai, S. J., Gassman, A., Gindhart, J. G. J., Hays, T. S. and Saxton, W. M. 1999. Cytoplasmic dynein, the dynactin complex, and kinesin are interdependent and essential for fast axonal transport. Mol. Biol. Cell 10: 3717 3728. Matthies, H. J. G., McDonald, H. B., Goldstein, L. S. B. and Theurkauf, W. E. 1996. Anastral meiotic spindle morphogenesis: role of the Non-Claret Disjunctional kinesin-like protein. J. Cell Biol. 134: 455 464. McDonald, H. B., Stewart, R. J. and Goldstein, L. S. B. 1990. The kinesin-like ncd protein of Drosophila is a minus end-directed microtubule motor. Cell 63: 1159 1165. Mocz, G. and Gibbons, I. R. 2001. Model for the motor component of dynein heavy chain based on homology to the AAA family of oligomeric ATPases. Structure 9: 93 103. Ogawa, K. 1991. Four ATP-binding sites in the midregion of the b heavy chain of dynein. Nature 352: 643 645. Paschal, B. M. and Vallee, R. B. 1987. Retrograde transport by the microtubule-associated protein MAP 1C. Nature 330: 181 183. Post, P. L., Tyska, M. J., O’Connell, C. B., Johung, K., Hayward, A. and Mooseker, M. S. 2002. Myosin-IXb is a single-headed and processive motor. J. Biol. Chem. 277: 11679 11683. Rayment, I., Rypniewski, W. R., Schmidt-Base, K., Smith, R., Tomchick, D. R., Benning, M. M., Winkelmann, D. A., Wesenberg, G. and Holden, H. M. 1993. Three-dimensional structure of myosin subfragment-1: a molecular motor. Science 261: 50 58.

9 Molecular Motor Directionality Rice, S., Lin, A. W., Safer, D., Hart, C. L., Naber, N., Carragher, B. O., Cain, S. M., Pechatnikova, E., Wilson-Kubalek, E. M., Whittaker, M., Pate, E., Cooke, R., Taylor, E. W., Milligan, R. A. and Vale, R. D. 1999. A structural change in the kinesin motor protein that drives motility. Nature 402: 778 784. Sablin, E. P., Case, R. B., Dai, S. C., Hart, C. L., Ruby, A., Vale, R. D. and Fletterick, R. J. 1998. Direction determination in the minus-enddirected kinesin motor ncd. Nature 395: 813 816. Sack, S., Müller, J., Marx, A., Thormählen, N., Mandelkow, E.-M., Brady, S. T. and Mandelkow, E. 1997. X-ray structure of motor and neck domains from rat brain kinesin. Biochemistry 36: 16, 155 16,165. Sale, W. S. and Satir, P. 1977. Direction of active sliding of microtubules in Tetrahymena cilia. Proc. Natl Acad. Sci. USA 74: 2045 2049. Sale, W. S., Goodenough, U. W. and Heuser, J. E. 1985. The substructure of isolated and in situ outer dynein arms of sea urchin sperm flagella. J. Cell Biol. 101: 1400 1412. Schnapp, B. J. and Reese, T. S. 1989. Dynein is the motor for retrograde axonal transport of organelles. Proc. Natl Acad. Sci. USA 86: 1548 1552. Shingyoji, C., Higuchi, H., Yoshimura, M., Katayama, E. and Yanagida, T. 1998. Dynein arms are oscillating force generators. Nature 393: 711 714.

Tang, W. J., Bell, C. W., Sale, W. S. and Gibbons, I. R. 1982. Structure of the dynein-1 outer arm in sea urchin sperm flagella. I. Analysis by separation of subunits. J. Biol. Chem. 257: 508 515. Telzer, B. R. and Haimo, L. T. 1981. Decoration of spindle microtubules with dynein: evidence for uniform polarity. J. Cell Biol. 89: 373 378. Vale, R. D., Reese, T. S. and Sheetz, M. P. 1985a. Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility. Cell 42: 39 50. Vale, R. D., Schnapp, B. J., Mitchison, T., Steuer, E., Reese, T. S. and Sheetz, M. P. 1985b. Different axoplasmic proteins generate movement in opposite directions along microtubules in vitro. Cell 43: 623 632. Walker, R. A., Salmon, E. D. and Endow, S. A. 1990. The Drosophila claret segregation protein is a minus-end directed motor molecule. Nature 347: 780 782. Wang, C., Asai, D. J. and Robinson, K. R. 1995. Retrograde but not anterograde bead movement in intact axons requires dynein. J. Neurobiol. 27: 216 226. Wells, A. L., Lin, A. W., Chen, L-Q., Safer, D., Cain, S. M., Hasson, T., Carragher, B. O., Milligan, R. A. and Sweeney, H. L. 1999. Myosin VI is an actin-based motor that moves backwards. Nature 400: 505 508.

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10 Kinesins: Processivity and Chemomechanical Coupling William O. Hancock and Jonathon Howard

10.1

Introduction

Movement is fundamental to cellular function, and to understand the molecular basis of cellular behavior and to design strategies to treat disease, it is important to understand the motor proteins that underlie cellular motion. Kinesins are molecular motors that use the energy derived from ATP hydrolysis to transport organelles and vesicles along intracellular microtubules. Conventional kinesin, an axonal transport motor, was the first kinesin to be identified and has been the most intensively studied. Subsequently, a number of other related motors have been identified by sequence homology to the kinesin motor domain; this kinesin family of motor proteins (often termed kinesin-like proteins, kinesin family proteins or unconventional kinesins) numbers 4 in the human genome (Kim and Endow, 2000, Miki et al., 2001) and includes motors that transport vesicles, organelles, protein complexes and chromosomes. A primary goal of research on kinesins and other motor proteins is to understand the transduction of chemical energy into mechanical work at the level of single protein molecules. These investigations require a range of techniques including mechanical measurements on individual motor molecules, enzyme kinetic studies, structural biology, and theoretical modeling of motor mechanisms. In the last decade and a half a considerable research effort by laboratories around the world has brought forth an outline of the mechanism of kinesin mechanochemistry and provided insights into the cellular role of various kinesin family members. This chapter will describe the experiments that underlie our current understanding of kinesin chemomechanical coupling and processivity and point to the unresolved questions that drive current and future research in this area. Conventional kinesin’s processive behavior, the motor’s ability to walk many steps along a microtubule without dissociating, is well established. This property is important for long distance transport as in the movement of vesicles along axons in nerve cells. Many of the kinetic steps by which kinesin transduces ATP hydrolysis into mechanical motion are understood, although there are still a num-

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10.2 Kinesin Motility and Processivity

ber of questions regarding the precise biochemical transitions that lead to force production and uncertainties regarding the critical steps that underlie processivity. The body of knowledge on conventional kinesin has set the paradigm to which the mechanism of other kinesin family members are compared. Studies on these non-conventional kinesins show that, while the fundamental steps in the chemomechanical cycle appear to be conserved, there are significant differences in chemomechanical coupling and processivity that adapt these motors to their diverse cellular functions. This chapter begins by describing some of the key experiments on the motility, processivity and biochemistry of conventional kinesin. These findings put constraints on models of chemomechanical coupling, eliminating many possible mechanisms. Next, experiments investigating the nature of coordination between the two heads of kinesin are described, which lead to a model of the ATP hydrolysis cycle for an individual kinesin head. Structural studies, which provide key clues for defining a kinetic model for two-headed kinesin, are then described followed by a discussion of the current model for the two-headed kinesin chemomechanical cycle that accounts for coupling ATP hydrolysis to movement and accounts for the processive behavior of dimeric kinesin. Finally, the conventional kinesin mechanism is compared to that of two kinesin family members: Ncd, a non-processive dimeric motor and KIF1A, a processive monomeric motor.

10.2

Kinesin Motility and Processivity

The first investigations into kinesin motility utilized the in vitro gliding assay (Fig. 10.1), in which motors are adsorbed to a glass surface and the movement of microtubules is observed by video-enhanced differential interference or fluorescence microscopy (Allen et al., 1981, 1985). An important insight into the mechanism of kinesin motility was the observation that kinesin is a processive motor. Processivity is defined as a motor’s ability to move multiple steps along its track without dissociating. The evidence for kinesin processivity came from dilution experiments in which a reduction in the motor surface density resulted in a proportional decrease in the rate that microtubules landed on the surface and started to move (Howard et al., 1989). At the lowest motor concentrations, microtubules were observed swiveling around nodal points on the surface and detaching from the surface when their trailing end moved over these points. This proportionality between activity and motor density coupled with the swiveling behavior demonstrated that individual kinesin motors could move multiple steps along microtubules without detaching. This processive movement of conventional kinesin is very different from skeletal muscle myosin motility assays where multiple motors are required to move an actin filament (Uyeda et al., 1990), and is consistent with kinesin’s role as a long distance transport motor. A second motility assay uses micron-scale glass or polystyrene beads, to which motors are attached, to visualize the movement of individual or groups of motors

10 Kinesins: Processivity and Chemomechanical Coupling In vitro motility assays used to study kinesin motility. (A) The microtubule gliding assay in which motors are adsorbed to a glass surface via the motor tail or other domain, and microtubules are observed gliding over the surface of motors. Motor surface density can be varied as can solution conditions to test the nucleotide dependence of movement. (B) The bead assay in which microtubules are immobilized on a surface and motors are adsorbed to glass or polystyrene beads.

Figure 10.1.

along immobilized microtubules or axonemes (Block et al., 1990). By decreasing the motor-to-bead ratio such that each bead, on average, has one or fewer motors attached, the bead assay confirmed that individual kinesin molecules can move processively along microtubules, and gave a more precise estimate of the distance moved per encounter of a motor with a microtubule. In the bead assay, the run length was found to be approximately 1 mm (Block et al., 1990), providing bounds for the off-rate of kinesin from microtubules. Processivity was confirmed by bead tracking experiments (Coppin et al., 1997, Kojima et al., 1997, Svoboda et al., 1993), which show that kinesin takes up to one hundred 8-nm steps along the microtubule before dissociating (Fig. 10.2A). This stepping behavior will be discussed further below.

(A)

(B) ATP

k bi

k cat ADP

N k off

Demonstration of mechanical and chemical processivity. (A) Kinesin stepping along a microtubule. A single kinesin motor was adsorbed to a glass bead. The bead was held in an optical trap at a low ATP concentration and its position was detected using a quadrant photodiode detector. The motor takes many 8-nm steps along the microtubule without falling off: the number of steps defines the

Figure 10.2.

extent of mechanical processivity. Data courtesy of Nick Carter and Rob Cross. (B) Evidence for chemical processivity. The motor binds to the microtubule with an on-rate equal to kbi and hydrolyzes N ATP at a turnover rate kcat before detaching with a rate koff. The number of ATP hydrolyzed per encounter, N, defines the extent of chemical processivity.

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10.4 Step Size of Kinesin and its Path along the Microtubule

10.3

Biochemical Evidence for Kinesin Processivity

Processive motility by conventional kinesin was supported by biochemical experiments demonstrating that dimeric kinesin hydrolyzes many ATPs per encounter with a microtubule (Hackney, 1995a). The initial evidence for this chemical processivity came from ATPase measurements in which the maximum microtubule-stimulated ATPase rate, kcat, was quite high (76 s 1 for the two-headed molecule) while the concentration of microtubules necessary for half-maximal activation, KM, was low (240 nM (Hackney, 1995a) to 30 nM (Hackney, 1994b), depending on buffer conditions and motor concentration), indicating that dimeric kinesin motors have a high affinity for their microtubule tracks. These data implied that if motors detached from microtubules once per hydrolysis cycle, then the bimolecular on-rate for motors binding to microtubules (kbi kcat/KM Z 3 q 108 – 3 q 109 M 1 s 1,) would need to be considerably faster than the calculated diffusion limited on-rate (2 – 3 q 107 M 1 s 1) (Hackney, 1995b). This discrepancy could be resolved if, instead of detaching once per ATP hydrolysis, the motor hydrolyzed many ATP per encounter with the microtubule. To calculate the number of ATP hydrolyzed per encounter, the bimolecular on-rate for motor binding to microtubules (kbi) was determined by a different method that takes advantage of the fact that kinesin binds ADP tightly in the absence of microtubules (off rates Z 0.01 s 1) but releases ADP rapidly when it encounters a microtubule (Hackney, 1988). To determine kbi, motors loaded with radioactive ADP were combined with various concentrations of microtubules, and the release of bound ADP was monitored over time. Because motor binding to microtubules is the rate-limiting step, this measurement provides the true bimolecular rate constant, kbi for motor binding. Dividing kcat/KM from the ATPase measurements by kbi gives the number of ATP hydrolyzed per encounter, N. For dimeric conventional kinesin, the number was found to be 120 ATP per encounter (Hackney, 1995a; Fig. 10.2B). These biochemical data also provide an estimate for the rate motors detach from their microtubule tracks. Dividing the maximal ATPase rate of 76 s 1 per dimer by the number of ATP hydrolyzed per encounter gives an estimated detachment rate of 0.6 s 1, which agrees reasonably with the duration of runs seen in the bead motility assay (Block et al., 1990).

10.4

Step Size of Kinesin and its Path along the Microtubule

An important constraint to defining the mechanism of kinesin motility was to define the path that kinesin takes as it steps along microtubules. The first experiment to define the path of kinesin took advantage of the fact that in microtubules with different numbers of protofilaments, the protofilament axis varies in relation to the microtubule axis. In some microtubules the protofilaments have a left-handed supertwist, others right-handed, while in some microtubules the protofilaments are parallel to the microtubule axis. By defining conditions that favor specific pro-

10 Kinesins: Processivity and Chemomechanical Coupling

tofilament numbers and tagging microtubules to allow microtubule rotation to be visualized in the microtubule gliding assay, Ray et al. (1993) showed that kinesin follows the protofilament axis with great fidelity. This fidelity was examined another way by attaching motors to submicron-scale beads and tracking the bead position with nanometer precision along immobilized microtubules (Berliner et al., 1995). As the beads moved down microtubules the position of the bead perpendicular to the microtubule axis hardly changed, indicating that the motors moved parallel to the protofilament axis and only rarely switched protofilaments. Because the tubulin dimers that make up a protofilament have a repeating 8-nm periodicity, or 4 nm per a or b monomer, the steps that kinesin takes along a microtubule were predicted to be a multiple of 4 nm, although other stepping mechanisms that utilize more than one protofilament could lead to different values. Using an optical tweezer and nanometer-scale positional detection of a motorcoated bead, Svoboda and coworkers observed steps of 8 nm as single kinesin motors moved a bead over an immobilized microtubule (Svoboda et al., 1993). This measurement defined the unitary mechanical event in kinesin motility and showed that the step size for conventional kinesin was independent of the ATP concentration and the load.

10.5

Kinesin Stoichiometry

A necessary first step towards understanding chemomechanical coupling is to define the coupling ratio: how many ATPs are hydrolyzed per 8-nm step that kinesin takes along a microtubule? Possible mechanisms range from those that have high fuel efficiency where each ATP hydrolysis event leads to a burst of steps to those with low fuel efficiency where numerous ATP are hydrolyzed without a concurrent mechanical step. A mechanism in which ATP hydrolysis is tightly coupled to movement predicts that one ATP is hydrolyzed per 8-nm step. Early kinesin ATPase experiments (Hackney, 1988) indicated that the motor hydrolyzes ATP much slower than expected based on the motility speed. However, it was recognized that fulllength kinesin takes on a folded conformation in solution, allowing the tail to interact and presumably inhibit the ATPase rate of the heads (Hackney et al., 1992). This was confirmed by measuring the ATP hydrolysis rate of truncated kinesins (Hackney, 1994b) or full-length motors bound to glass beads (Coy et al., 1999a, 1999b). In these cases, the ATPase rate of nearly 100 ATP s 1 for dimeric kinesin correlates with the motility rate (800 nm s 1) and measured step size (8 nm) (Coy et al., 1999b). Hence, under conditions where the mechanical load is small, kinesin is a tightly coupled motor and hydrolyzes one ATP per 8-nm step. The stoichiometry of one ATP per step was also inferred by measuring the frequency of steps at various ATP concentrations, analyzing the distribution of dwell times between steps and comparing the data to various models of coupling (Hua et al., 1997; Schnitzer and Block, 1997). At low ATP concentrations individual steps are observed instead of bursts of steps and the rate of stepping varies linearly

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10.6 Coordination between the Two Heads of Kinesin

with ATP concentration, ruling out models in which ATP binding leads to more than one step. Similarly, because the dwell times between steps are exponentially distributed at low ATP concentrations, mechanisms that involve multiple ATP binding per step are also discounted. In summary, both observations of stepping frequencies and correlation of ATPase rates with movement speeds together demonstrate that under low load (I 1 pN), conventional kinesin hydrolyzes one ATP per step. This puts bounds on chemomechanical coupling models described below. However, it is still an open question whether this 1 : 1 stoichiometry holds when the motor is moving against high external loads. Additionally, this stoichiometry does not necessarily apply to other motors in the kinesin family whose chemomechanical coupling mechanisms are tuned for different cellular roles.

10.6

Coordination between the Two Heads of Kinesin

While mechanical and biochemical experiments can clearly demonstrate kinesin processivity, uncovering the mechanistic basis in terms of structural and biochemical states and transitions between these states is more challenging. The eventual goal in understanding chemomechanical coupling in motor proteins is to formulate a kinetic model detailing the steps of the ATP hydrolysis cycle including rate constants governing transitions between these states and connections to structural transitions. However, uncovering these states and transitions is made difficult by the presence of kinesin’s two heads and uncertainties regarding the nature of their interactions and coordination. By loading kinesin’s two heads with labeled ADP, combining these motors with microtubules, and monitoring the rate of ADP dissociation under various conditions, Hackney (1994a) showed that the two heads of kinesin are not kinetically independent. These experiments again relied on the slow ADP release rate of microtubule-free kinesin (Z 0.01 s 1) (Hackney, 1988), which is accelerated over three orders of magnitude by microtubule binding. The key result was that when motors and microtubules were combined in the absence of any free nucleotide, only half of the bound ADP was released at a fast rate and the other half released slowly, suggesting that one head can bind to the microtubule and release its nucleotide while the other is prevented from binding. If ATP is added, the remaining bound ADP is rapidly released, showing that ATP binding or hydrolysis catalyzes a transition in the microtubule-stimulated ATP hydrolysis cycle. This singly bound state, presumed to be an intermediate on the kinetic pathway for microtubule-stimulated ATP hydrolysis, demonstrates that kinesin’s two heads do not operate independently. The finding that in the presence of ATP both heads rapidly released their bound nucleotide left unresolved the question of whether ATP binding or ATP hydrolysis is necessary for nucleotide release by the second head. This question was answered by measuring the rate and extent of bound-ADP release in the presence of saturating concentrations of the non-hydrolyzable ATP analog, AMP-PNP (Ma and Taylor,


1997a). In AMP-PNP, the microtubule-stimulated release of the second bound ADP was 30 s 1, significantly faster than the rate measured in either no nucleotide or saturating ADP. However, the ADP release rate is 3 12-fold slower in AMPPNP than it is in ATP (Crevel et al., 1999, Hackney, 2002, Ma and Taylor, 1997a). Additionally, ATP hydrolysis by a bound monomer head, an analog of the bound head in a dimer, has been shown to be quite rapid (i 200 s 1, (Ma and Taylor, 1997b), 100 s 1 (Jiang and Hackney, 1997)) compared to the ADP release rate in the presence of AMP-PNP. Thus, it is likely that ATP hydrolysis by the bound head precedes binding and ADP release by the tethered head. These data are incorporated into the two-headed hydrolysis model developed below.

10.7

Testing Processivity with One-headed Kinesin Mutants

Although experiments on two-headed kinesin provide the framework for understanding processivity, the interdependency of the two heads invariably masks important features of the hydrolysis cycle. For this reason, a considerable push was made to investigate the motility and biochemical characteristics of various oneheaded kinesin constructs. Initially, simple deletions were made to the coiled-coil and dimerization domain, and these truncation mutants tested for microtubule gliding (Stewart et al., 1993). These results suggested that, if attached to suitable artificial tails, one-headed kinesins were motile, but the motility speeds were very slow, the ATPase rates were very fast, and it appeared that the activity of the monomers depended upon exactly where in the sequence the truncation was made. A second approach that successfully demonstrated one-headed kinesin motility was to fuse the kinesin head to a biotin-binding protein and attach these motors to streptavidin-coated beads (Berliner et al., 1995). These beads, coated with many one-headed kinesin motors, moved along microtubules but fluctuated in their lateral position significantly more than beads coated with two-headed kinesin. Importantly, when the motor surface density was decreased, no motility was observed, consistent with a lack of processivity. However, the lack of any activity at low motor numbers limited any conclusions regarding the mechanism of processivity. In an effort to retain as much of the wild-type kinesin structure as possible, Hancock and Howard created a single-headed kinesin construct that retained the full coiled-coil domain, rod and tail, but which contained only one head domain (Hancock and Howard, 1998). At high surface densities in the microtubule gliding assay, these one-headed kinesin heterodimers exhibited robust gliding activity with speeds roughly 1/8th that of wild-type kinesin. To test whether one-headed kinesin is processive, the surface density of motors was systematically decreased and the activity, defined as the rate that microtubules land on and begin to move over the surface, was measured. When the motor surface density of wildtype kinesin was varied in this way, the fall in the microtubule landing rate was

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10.8 ATP Hydrolysis Cycle of One-headed Kinesin

proportional to the motor density and at the lowest motor densities observed (calculated as I 10 motors mm 2), microtubules were observed pivoting over individual points on the surface, indicative of individual kinesin motors processively moving microtubules. In contrast, one-headed kinesin exhibited a steep dependence of landing rate on motor density and at the lowest motor densities no motility was observed, demonstrating that this one-headed kinesin heterodimer is not a processive motor. By fitting the landing rate curves to models of cooperative motor movement, it was estimated that a minimum of four to six one-headed kinesins are necessary to move a microtubule. These data suggest that the processivity of the kinesin dimer is due to the coordinated motion of its two heads. An important insight into kinesin processivity came from the behavior of oneheaded kinesin heterodimers under conditions where movement was not seen. At low motor densities, microtubules still bound to motors but did not move. This behavior was predicted by the hand-over-hand model if the two heads are coordinated such that the first head does not release until the second head binds, then deleting the second head should result in a long-lived bound state by the remaining motor domain. The kinetics of one-headed kinesin detachment was analyzed in greater detail by adsorbing kinesin heterodimers to glass beads at densities of one motor per bead and observing the rate of binding and unbinding from microtubules (Hancock and Howard, 1999). As a comparison, wild-type kinesin, which takes approximately 100 steps per second, requires that each head must detach at a rate of at least 50 s 1 during the walking cycle. In contrast, the detachment rate of one-headed kinesin heterodimer in saturating ATP concentrations was 3 s 1. Hence the second head accelerates detachment of the first head by at least an order of magnitude. This experiment demonstrates a second way in which the motion of the heads are coordinated (the first being the half-site release experiment). The slow detachment of the one-head-bound state is presumably an adaptation to increase the processivity of the kinesin dimer.

10.8

ATP Hydrolysis Cycle of One-headed Kinesin

Combining ATPase and microtubule binding data from one-headed kinesin heterodimers together with kinetic data from dimeric and truncated monomeric kinesins, it is possible to construct a model for the one-headed kinesin hydrolysis cycle. This chemomechanical model for a single kinesin head is a necessary first step towards building a model for the two-headed kinesin hydrolysis cycle, and it provides a framework in which to understand processivity. Here we will step through the kinetic model for one-headed kinesin (Fig. 10.3A) along with the evidence supporting each transition. Kinesin heads in solution bind ADP tightly (Hackney, 1988) and microtubule binding causes rapid release of this bound nucleotide (at 50 300 s 1; Crevel et al., 1999, Gilbert et al., 1998, Hackney, 2002, Ma and Taylor, 1997a) and strong attachment of the motor to the microtubule. The next step, ATP binding, is also


One-headed kinesin and myosin and detaches in the ATP state. For clarity, other hydrolysis cycles. Kinesin binds to microtubules nucleotide states and reverse transitions are in the ADP state and likely detaches in the ADP- not shown. Pi state, while myosin binds in the ADP-Pi state

Figure 10.3.

rapid at physiological nucleotide concentrations: the bimolecular on-rate has been measured to be 0.8 3 mM 1 s 1 (Gilbert et al., 1998, Hackney, 2002, Ma and Taylor, 1997b). This ATP state also binds strongly to microtubules as evidenced by the high affinity between kinesin and microtubules in the presence of the non-hydrolyzable ATP analog AMP-PNP (Lasek and Brady, 1985). The structural change underlying one-headed kinesin movement most likely occurs upon or immediately

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10.8 ATP Hydrolysis Cycle of One-headed Kinesin

following ATP binding, based on mechanical data from two-headed kinesin discussed below. Following ATP binding, hydrolysis occurs rapidly (i 200 s 1 (Ma and Taylor, 1997b), 100 s 1 (Jiang and Hackney, 1997)) leaving the motor bound to the microtubule in the ADP-Pi state. All of the steps up to this point are generally agreed upon and consistent with kinetic studies using a range of one-headed kinesin constructs. The detachment step for one-headed kinesin is not as broadly agreed upon, however, though the bulk of the evidence points to detachment in the ADP-Pi state and subsequent attachment in the ADP state following phosphate release. Full-length one-headed kinesin heterodimer hydrolyzes ATP at a slow rate (Z 3 ATP s 1), which is similar to the detachment rate in the presence of ATP (2.9 s 1; Hancock and Howard, 1999). Thus, one-headed kinesin detaches once per hydrolysis cycle and both the detachment rate and the ATPase rate are slower than in the native two-headed motor. A simple explanation for these results is that one-headed kinesin detaches in the ADP-Pi state. Pi would then unbind from the detached head, completing the hydrolysis cycle. A similar model has also been proposed in which Pi release occurs when the motor is bound to the microtubule, but is immediately followed by microtubule detachment (Cross et al., 2000). An alternate one-headed kinesin cycle, motivated by kinesin’s relatively low microtubule affinity in the ADP state (Crevel et al., 1996, Romberg and Vale, 1993), holds that the motor detaches in the ADP state (Hackney, 2002, Rice et al., 1999, Vale and Milligan, 2000). However, because motors attach in the ADP state as well, the entire hydrolysis cycle would occur while kinesin is bound to the microtubule and would therefore produce no net movement. To obtain a net displacement during a hydrolysis cycle, a motor must undergo a conformational change (a power stroke) while bound to the microtubule, followed by a recovery stroke while detached. Hence, to harness movement from this cycle, kinesin ADP would need to have two distinct biochemical and structural states a pre-power stroke state that releases ADP rapidly upon binding to the microtubule and a post-power stroke state that releases ADP slowly and allows the head time to detach in the ADP state. There is no structural or biochemical evidence for these two states. One apparent inconsistency in the literature is the one-headed kinesin ATPase rate. Full-length one-headed kinesin constructs display slow ATPase kinetics (3 s 1; Hancock and Howard, 1999), while truncated monomeric heads display hydrolysis rates as fast or faster than each head in dimeric kinesin (64 to 89 s 1; Huang and Hackney, 1994, Jiang and Hackney, 1997, Moyer et al., 1996). However, the monomers display aberrant or no motility and in some cases they hydrolyze many ATP per microtubule binding event (Jiang and Hackney, 1997). The disparate models can be reconciled by positing that Pi release is slow for a microtubulebound head, and acts as a checkpoint such that detachment from the microtubule precedes Pi release as argued above. In truncated monomer kinesins, this checkpoint is lost and Pi release followed by ADP release and subsequent ATP binding results in multiple ATP molecules being hydrolyzed without the motor moving or detaching from the microtubule.


To understand the one-headed hydrolysis cycle of conventional kinesin, it is helpful to compare it to the myosin hydrolysis cycle (Fig. 10.3B). Myosin binds to actin tightly in the ADP and no nucleotide states and binds weakly in the ATP and ADPPi states (Lymn and Taylor, 1971). In the standard cycle, the motor hydrolyzes ATP in the detached state, binds to actin in the ADP-Pi state, and undergoes a weak-tostrong transition coincident with Pi release. Following ADP release, ATP binding leads to rapid detachment and recovery to the pre-power stroke conformation is associated with hydrolysis (reviewed by Howard (2001)). The one-headed kinesin hydrolysis cycle presented above is similar to the myosin cycle, except that the states are shifted 90h such that attachment occurs in the ADP state and detachment occurs in the ADP-Pi state. With one possible exception (Matthies et al., 2001), all kinesin family members tested to date, share the same overall hydrolysis cycle with the ATP and nucleotide-free states binding strongly to microtubules and ADP and ADP-Pi states binding more weakly.

10.9

Structural Studies on Dimeric Kinesin

In addition to biochemical and mechanical experiments, structural studies on dimeric kinesin in solution and attached to microtubules provide the third set of constraints necessary to define a chemomechanical model for kinesin. The crystal structure of dimeric kinesin with ADP bound to both heads shows that the two heads have approximate two-fold symmetry in which the heads are related by a 120 h-rotation instead of a 180 h-rotation expected for exact two-fold symmetry (Kozielski et al., 1997). The heads are held together by a coiled-coil dimerization domain and the so-called neck linker that connects the core motor domain to the dimerization domain. In the crystal structure where no microtubules are present and ADP is bound to each head, the coiled-coil is closed and in each head the linker is held tightly against the core motor domain, such that a considerable rearrangement would be necessary for both heads to simultaneously bind to a microtubule. Cryoelectron microscopic (cryoEM) reconstructions of dimeric kinesin motors attached to microtubules show that the relationship between the bound and free heads varies depending on what nucleotides are bound to each head (Fig. 10.4). In the absence of nucleotide, one head of the dimeric motor is bound to the microtubule and the tethered head is pointing to the plus-end of the microtubule and leaning to the left of the bound head (Arnal and Wade, 1998, Hirose et al., 1996). In AMP-PNP the tethered head is positioned on the right side of the bound head, suggesting that there is a conformational change following ATP binding to the bound head. This structural change is supported by experiments on monomeric kinesin in which the carboxyl terminus is labeled with a gold particle (Rice et al., 1999): in AMP-PNP the gold particle is located in the position where the tethered head is found in cryoEM reconstructions. From ADP release experiments, it is expected that in AMP-PNP the tethered head can bind to the microtubule and release its bound ADP. Thus one might ex-

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10.10 Two-headed Kinesin ATP Hydrolysis Cycle

Kinesin conformational changes during the ATP hydrolysis cycle. Images are digital reconstructions from electron micrographs of frozen samples. Both heads can be seen, one attached to the microtubule (A) and one free (F). The position of the tethered head changes with nucleotide state, particularly from the nu-

Figure 10.4.

cleotide-free to AMP-PNP state, due to conformational changes in the bound head or in the interface between the two heads (arrowed). Image by courtesy of Dick Wade, reproduced from Arnal and Wade (1998) with permission from Cell Press.

pect that, instead of finding the second head tethered in AMP-PNP reconstructions, it would instead be bound to the next binding site along the microtubule. In fact, there is less mass attributed to the second head in cryoEM reconstructions in AMP-PNP, suggesting either that the second head is mobile or that it is bound to the microtubule (Hoenger et al., 2000). This is one piece of evidence in support of a kinesin state in which both heads may be bound to the microtubule. Another piece of evidence in support of a two-heads-bound conformation comes from optical tweezer measurements where motors were attached to glass beads and the rupture force and stiffness compared in no nucleotide versus AMP-PNP. In the presence of the ATP analog, the stiffness and the force required to pull the motor off of the microtubule were twice that measured in no nucleotide (Kawaguchi and Ishiwata, 2001), suggesting that both heads bind in AMP-PNP while only one can bind in the absence of nucleotide. Hence, there is both structural and mechanical evidence that when one head binds ATP, the kinesin dimer can achieve a two-head-bound state.

10.10

Two-headed Kinesin ATP Hydrolysis Cycle

Combining mechanical, biochemical and structural data for two-headed kinesin together with insights from the one-headed hydrolysis cycle, a consistent picture for the two-headed kinesin hydrolysis cycle emerges. A chemomechanical model for


conventional kinesin, originally developed by Hancock, Shief and Howard (Hancock and Howard, 1999, Schief and Howard, 2001), is presented here (Fig. 10.5) and includes some features that are broadly agreed upon and other features that are still being debated. This model provides a framework for understanding which steps are load dependent, which transitions are rate limiting, which transitions are critical for processivity, and which features of the cycle are modified in kinesin-related proteins that display novel motility characteristics. In solution, kinesin has a high affinity for ADP and binding to a microtubule causes the release of ADP from one of the two heads. This transition (state 1) causes one head to bind tightly to the microtubule in a conformation that prevents the second head from binding. This step is supported by biochemical data discussed above (Hackney, 1995a) and by cryoEM structural studies that show that in the absence of nucleotide one head is bound to the microtubule and one head remains free (Arnal and Wade, 1998, Hirose et al., 1996). In this one-headbound state, it is presumed that the conformation of the bound head is such

0

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5

DP

φ

D

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6

T

ADP ATP ADP·Pi

φ

D D

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0 nm 8 Model of the two-headed kinesin hydrolysis cycle, combining biochemical, structural and motility data. The neck-linker is shown as a rod that alternates between a docked (1) and a free conformation (e. g. 2), and a coiled-

Figure 10.5.

φ

0 nm 8 coil that alternates between a closed (1) and partially open (e. g. 2) conformation. For details see text. Figure reprinted from Schief and Howard (2001) with permission from Elsevier Science.

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10.10 Two-headed Kinesin ATP Hydrolysis Cycle

that the free head is prevented from interacting with the microtubule. Accordingly, the neck linkers for both heads are shown in their structured state, tightly associated with the core head domain. The second step, ATP binding, occurs rapidly at physiological nucleotide concentrations (kon estimated at 0.3 3 mM 1 s 1; Gilbert et al., 1998, Hackney, 2002, Ma and Taylor, 1997b). Two bodies of evidence suggest that ATP binding induces a conformational change in the bound head that accounts for at least part of the 8-nm displacement along the microtubule during each step. As discussed above, when kinesin containing bound ADP is combined with microtubules in the presence of AMP-PNP, the release of the second ADP molecule is much faster than in the absence of any nucleotide. Although the rate is not as fast as in the presence of saturating ATP concentrations, this result nonetheless indicates that ATP binding leads to a conformational change that permits the second head to bind the microtubule and release its nucleotide. The second piece of evidence suggesting that ATP binding induces a conformational change in the bound head comes from mechanical experiments using microneedles or optical tweezers to impose a mechanical load on a motor. At low ATP concentrations, where ATP binding to the head is the rate-limiting step in the hydrolysis cycle, the velocity of movement slows with imposed force (Meyhofer and Howard, 1995, Svoboda et al., 1993). Thus, a transition coincident or associated with ATP binding is load dependent as discussed further below. Following ATP binding, the nucleotide is hydrolyzed to ADP Pi and the second head rapidly attaches and releases its bound ADP locking the motor into a twohead-bound conformation (state 5). This transition is thought to involve a significant conformational change to allow the two heads to span the 8 nm between successive tubulin subunits. A restructuring of the neck linker in the forward head is a plausible explanation, but it is also possible that the linker in the bound head is stretched, or that the coiled-coil dimerization domain also partially unfolds to permit both heads to bind. In any case, the rear head is thought to be under considerable strain in this two-heads-bound state, which accelerates its detachment. To complete the hydrolysis cycle, the rear head is detached and swung towards the plus-end of the microtubule to ready it for the next hydrolysis cycle (state 1l). Presently there is insufficient evidence to strictly define this portion of the pathway, although available evidence points to a preferred sequence. The simplest interpretation is that the strain in the two-heads-bound state (state 5) causes the rear head to detach from the microtubule and a restructuring of the neck linker and coil following Pi release swings the rear head to a forward tethered position (state 1l). It is formally possible, however, that ATP binding is rapid enough to allow ATP to bind to the forward head before the rear head is detached, followed rapidly by rear-head detachment and Pi release (Rice et al., 1999). This pathway may be preferred at high external loads, where the external force pulling backward slows the detachment of the rear head, allowing time for the forward head to bind ATP. Another possible transition out from state 5 is for Pi to be released before rearhead detachment. Because the ADP affinity of kinesin bound to microtubules is very low (koff is 50 300 s 1; Gilbert et al., 1998, Hackney, 2002, Ma and Taylor,


1997a), ADP would be rapidly released, followed by ATP binding and hydrolysis without an associated step. These futile hydrolysis cycles may occur at high external loads, where the rear head detaches slowly enough to permit extra hydrolysis cycles before detachment. The 1 : 1 stoichiometry at low load confirms that they never or only rarely occur at low loads, and this is why the pathway shown in Fig. 10.5 in which rear-head detachment precedes Pi release, is preferred. Processive kinesin motors take approximately 100 steps along a microtubule without detaching (Block et al., 1990, Hackney, 1995a), meaning that there is a 1 % probability of detachment per hydrolysis cycle. What limits kinesin processivity? Because motors bind tightly in both the ATP and nucleotide-free states (detachment rates 0.001 s 1; Hancock and Howard, 1999) it is unlikely that detachment occurs from those states or any state in which both heads are bound. Detachment most likely occurs from state 3, in which the bound head is in the ADP-Pi state and hence any mechanical load that increases the duration the motor spends in state 3 should decrease the extent of processivity. It should be noted that other models that include different intermediate states in the hydrolysis cycle predict different paths for detachment (Hackney, 2002, Yajima et al., 2002), and hence this feature of the hydrolysis cycle is not entirely resolved.

10.11

Load Dependent Transitions

A primary question in understanding kinesin mechanochemistry is: what are the structural changes that underlie the 8-nm step that occurs for each hydrolysis cycle? Is there one transition that covers the entire distance? Is the 8-nm instead made up of a number of substeps each associated with a biochemical transition? An associated question is: what transition is rate limiting at high loads? The way to explore these questions has been to determine which transitions in the hydrolysis cycle vary with external load. The first evidence for a load-dependent step came from mechanical experiments using flexible microneedles or optical tweezers to impose external loads on individual kinesin motors. The speed of movement was found to decrease linearly with imposed load (Fig. 10.6) both at high ATP concentrations and, importantly, at low ATP concentrations as well (Meyhofer and Howard, 1995, Svoboda et al., 1993). Because ATP binding is the rate-limiting step at low ATP concentrations, it follows that small forces that slow movement must be slowing ATP binding or a step closely associated with ATP binding. If ATP binding is irreversible, this slowing could be achieved by slowing the on-rate for ATP binding. Alternatively if ATP binding is reversible, then the load could accelerate the ATP off-rate or could slow a step subsequent to ATP binding, allowing more time for ATP to dissociate and achieving the same effect of a slowed biochemical transition. To investigate the effects of external load and ATP binding on kinesin movement in more detail, Visscher et al. (1999) developed an optical trap with feedback control whereby the sustained movements of individual motors could be observed

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Kinesin velocity as a function of external load. A microtubule was attached to a flexible microneedle with known stiffness and the deflection of the microneedle by an individual kinesin was measured. (A) Raw data showing a number of motor-microtubule interactions in which the motor deflects the needle up to a maximal force and then eventually detaches from the microtubule. (B) Velocity as a

Figure 10.6.

50

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function of force at saturating ATP concentrations (top curve) and limiting ATP concentrations (bottom curve). (C) Expanded view of data at limiting ATP concentrations showing that the velocity decreases with load even at limiting ATP concentrations. Data reprinted from Meyhofer and Howard (1995). Copyright 1995, National Academy of Sciences, USA.

under constant external load. It was observed that at high loads that slowed the motor stepping rate, the ATP concentration necessary to achieve half-maximal velocity increased. Again, the data were interpreted as an external load either decreasing the on-rate for ATP binding or otherwise altering the rate constant of a step closely associated with ATP binding. The data were described by a model of ATP binding to the microtubule-bound head, the subsequent binding of the tethered head to the microtubule and the eventual detachment of the first head (Schnitzer et al., 2000). To account for the ATP dependence, a composite state was invoked in which the tethered head undergoes a 4-nm displacement from a rear to forward position and the two positions exist in a rapid equilibrium such that the external load affects the probability that the tethered head will be in the forward position according to Boltzman-statistics. The result of this composite state is that by mass action there is a greater probability of ATP unbinding at high loads, account-


ing for the apparent drop in ATP affinity, while at low loads binding of the tethered head to the next binding site is favored. Thus, the presence of a major structural transition coincident with or immediately following ATP binding that would confer load-dependence is supported by biochemical (section 10.6), electron microscopic (section 10.9) and mechanical (this section) studies. However, the force velocity curve at saturating ATP concentrations, which is consistently found to be linear (Coppin et al., 1997, Kojima et al., 1997, Meyhofer and Howard, 1995, Svoboda et al., 1993) or concave down (Visscher et al., 1999), implies there is at least one more force-dependent conformation (Keller and Bustamante, 2000). One candidate for a second load-dependent transition is a structural rearrangement following unbinding of the rear head (state 6 in Fig. 10.5). Because the dimeric motor is thought to be highly strained when both heads are bound, release of the rear head should pull the dimerization domain forward and an external load pulling towards the minus end of the microtubule should slow the detachment, while an external load in the direction of movement should accelerate it. In support of this idea, it was observed that for a motor moving against an elastic load, a force pulling the motor away from the microtubule in a perpendicular direction leads to an increase in the velocity at high loads (Gittes et al., 1996). This result suggests a perpendicular force can accelerate detachment of the rear head, which may be rate limiting at high loads. The magnitude and kinetics of other load-dependent transitions in the kinesin chemomechanical cycle are an area of ongoing investigation.

10.12

Ncd is a Non-processive Kinesin Family Member

In contrast to kinesin’s processive behavior, the kinesin-related protein Ncd lacks processivity in either motility or biochemical experiments. These differences are consistent with the different cellular functions of these two motors: conventional kinesin is a long-distance transport motor while Ncd is involved in meiosis and mitosis where presumably many motors cooperate in spindle formation and maintenance (Endow et al., 1994). While the atomic structures of the kinesin and Ncd head domain are strikingly similar (Kull et al., 1996, Sablin et al., 1996), the attachment to their dimerization domains is quite different (Kozielski et al., 1997, Sablin et al., 1998), a feature that directly contributes to the opposite direction of movement compared to conventional kinesin, and which may explain the lack of processivity of Ncd as well. By comparing structural and chemomechanical models of Ncd and conventional kinesin it should be possible to identify the critical features that underlie processivity of conventional kinesin. The first experiments that demonstrated a lack of Ncd processivity measured the dependence of the landing rate on the surface motor density similar to the approach taken for one-headed kinesin described above (deCastro et al., 1999). Due to the lack of a comparable control motor to confirm that the dimeric molecule binds to the surface in a functional way, the landing rate in ATP was compared

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to that in AMP-PNP, which promotes strong binding to the microtubule. Consistent with individual Ncd motors being sufficient to bind microtubules to the surface, in AMP-PNP the landing rate was found to be proportional to the surface motor density. This control experiment confirmed that motors were bound to the surface in an active conformation and it defined the motor dilution necessary for single motor interactions with microtubules. When the experiments were repeated in ATP, much higher motor densities were required to obtain equivalent landing rates, and the landing rate fell off steeply with motor density, indicative of many motors being required to move a microtubule. A more direct assay of the stepping behavior of Ncd utilized an optical tweezer system to bring a microtubule into contact with individual surface-adsorbed Ncd motors (deCastro et al., 2000). By attaching silica beads to both ends of a microtubule and immobilizing motors on a third bead attached to the surface of the flow chamber, the microtubule could be positioned to interact with individual motor molecules and the displacements measured directly. Instead of the stereotyped staircase of 8-nm steps seen for conventional kinesin, Ncd produced unitary steps of approximately 9 nm. To observe these steps, the duration of binding events was prolonged by decreasing the ATP concentration and multiple events were averaged to increase the signal to noise ratio. The initial binding, which was monitored by a decrease in the amplitude of fluctuations in the bead position, involved little or no displacement of the motor along the microtubule axis. However, at the end of the binding event there was a measurable displacement of roughly 9 nm along the microtubule axis followed by rapid detachment. The duration of the entire binding event varied with ATP concentration, consistent with the nucleotide-free motor holding to the microtubule and awaiting ATP binding. In contrast, the delay between the displacement and detachment, although obscured somewhat by the signal averaging, did not appear to vary with ATP concentration. These mechanical transients are consistent with ATP binding causing a conformational change in the bound Ncd head, which is followed rapidly by hydrolysis and detachment from the microtubule in the ADP-Pi state. These mechanical events for dimeric Ncd are what might be expected from a non-processive one-headed conventional kinesin. Following binding and ADP release the motor is tightly bound to the filament, and ATP binding and hydrolysis lead to detachment (Hancock and Howard, 1999). Kinetic models of the kinesin hydrolysis cycle predict displacements upon ATP binding of 1 4 nm (Schief and Howard, 2001, Schnitzer et al., 2000), smaller than those observed from Ncd. It is possible that mechanical amplification by the Ncd dimerization domain could underlie this large observed step size. Biochemical experiments on dimeric Ncd also point to a non-processive mechanism and highlight differences from processive kinesin. The first biochemical evidence against Ncd being processive comes from experiments where Ncd motors are first incubated with microtubules in the absence of nucleotide to promote a strongly bound Ncd microtubule complex. When this complex is rapidly mixed with ATP, there is an initial burst of hydrolysis that has an amplitude equal to the concentration of motors present, and which is followed by a slower steady


state rate (Foster and Gilbert, 2000). This stoichiometric burst is consistent with microtubule-bound motors binding and hydrolyzing one molecule of ATP and then detaching from the microtubule, with later hydrolysis events requiring the motor to reattach to the microtubule. The second piece of evidence is that the detachment of Ncd from microtubules following rapid mixing with ATP has a fast rate of 13 s 1 (Foster and Gilbert, 2000), considerably faster than the overall hydrolysis rate of 2 s 1 (Foster et al., 1998). Finally, equilibrium binding experiments showed that in ADP plus added Pi the microtubule affinity of Ncd is lower than in ADP alone (Foster et al., 1998), suggesting that the motor detaches in the ADP-Pi state and then rapidly releases its Pi. This detachment step agrees with much of the conventional kinesin data (Hancock and Howard, 1999, Rosenfeld et al., 1996) although in both cases direct demonstration of detachment in the ADP-Pi state has proven difficult. In summary, the key feature of the Ncd hydrolysis cycle that prevents processivity is that following ATP hydrolysis, motor detachment from the microtubule (most likely in the ADP-Pi state) occurs at a faster rate than attachment of the second head. To prevent diffusion of the motor away from the microtubule, most models of alternating hand-over-hand processivity require that during some portion of the walking cycle both heads must bind simultaneously to the microtubule. In conventional kinesin a structural transition in the neck region, melting from a b-sheet conformation to a random coil or opening of a coiled-coil, is thought to provide the flexibility needed to allow both heads to bind simultaneously to a microtubule (Kozielski et al., 1997). Available structural evidence for Ncd suggests that the two heads are more tightly associated with one another than are the two heads of conventional kinesin, and thus may not be able to simultaneously bind to a microtubule. First, in the crystal structure of two headed Ncd (Sablin et al., 1998), the heads possess two-fold symmetry and are held together tightly against the coiled-coil dimerization domain by multiple associations between residues in the core motor domain and those in the coiled-coil. Second, the neck-linker that joins the catalytic core to the neck-coil is considerably shorter for Ncd (Endow and Waligora, 1998, Sablin et al., 1998), which would necessitate a considerable uncoiling of the neck coil to allow the tethered head to bind to the next microtubule binding site. Third, in cryoEM reconstructions of dimeric Ncd bound to microtubules (Arnal et al., 1996, Sosa et al., 1997), which show the tethered head pointing towards the minus end of the microtubule, the tethered head is considerably more well-defined than for conventional kinesin, indicating that it is relatively immobile. Hence, all available structural evidence suggests that the two Ncd heads remain tightly associated, which would prevent both heads from simultaneously binding to a microtubule and preclude processive movement.

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10.13 A Processive Monomeric Kinesin, KIF1A

10.13

A Processive Monomeric Kinesin, KIF1A

Because the prevailing model for conventional kinesin processivity involves coordination between the two heads, it came as quite a surprise when a monomeric kinesin, KIF1A, was shown to be processive (Okada and Hirokawa, 1999, 2000). KIF1A, which is thought to be involved in vesicle transport in neurons, has been shown by hydrodynamic analysis to be monomeric in solution (Okada et al., 1995). Motility of individual KIF1A molecules was observed by fusing the KIF1A head and conventional kinesin neck linker to green fluorescent protein and observing individual motors moving along immobilized microtubules (Okada and Hirokawa, 1999). The motors bound microtubules for long durations and diffused back and forth with a net movement towards the microtubule plus-end such that over time the motors accumulated at one end of the microtubules. The key to KIF1A processivity is that the motor domain has a second microtubule binding site consisting of a loop of positively charged residues. This ‘Kloop’, named because of its numerous lysine residues, is thought to interact with the negatively charged carboxyl tail of tubulin, termed the ‘E-hook’ because it contains numerous glutamate residues. A body of evidence supports the notion that the K-loop provides a tether that holds the motor on the microtubule during the detachment phase of the hydrolysis cycle. First, when the K-loop of KIF1A or the E-hook of tubulin was deleted, the processive behavior was abolished. Second, mutants in which some of the lysine residues were deleted showed a graded decrease in microtubule affinity with deletion of lysines. Third, if the K-loop was inserted into the homologous sequence of a conventional kinesin monomer, this normally non-processive head displayed processive movement with positional fluctuations in both directions but a clear net displacement over time. Fourth, the cryoEM reconstruction of KIF1A bound to a microtubule, aided by alignment with the crystal structure of the conventional kinesin head, shows an interaction between the Kloop and E-hook (Kikkawa et al., 2000). The motility of KIF1A is very different from the precise stepping of conventional kinesin. KIF1A makes rapid runs with amplitudes of hundreds of nanometers in both directions and periodically pauses for hundreds of milliseconds or more. This movement can be quantitatively described as one-dimensional diffusion superimposed on a steady directed component. The proposed mechanism involves diffusional events where the motor is loosely associated to the microtubule via its Kloop, together with a conventional kinesin power stroke to bias the diffusion towards the microtubule plus-end (Okada and Hirokawa, 2000). Consistent with this, when movement was analyzed in the presence of ADP only, the diffusive component continued, but the net movement towards the plus-end was abolished. Biochemical experiments (Okada and Hirokawa, 2000) provide insight into the walking mechanism of KIF1A. The microtubule-stimulated ATPase rate of KIF1A is 110 s 1 and when compared to the 140 nm s 1 net velocity towards the microtubule plus-end, indicates that the motor most likely takes many futile steps that result in ATP hydrolysis but no net displacement. To determine the ex-


tent of chemical processivity (the number of ATPs hydrolyzed per encounter with the microtubule), the approach developed for conventional kinesin was employed. By comparing the predicted motor-microtubule association rate (kbi) from ATPase measurements to the actual encounter rate measured by release of labeled ADP following microtubule binding, it was determined that KIF1A hydrolyzes nearly 700 ATP per encounter with a microtubule. Thus KIF1A is five-fold more chemically processive than conventional kinesin. Finally, the time constant for motor dissociation from the microtubule (6.3 s) calculated from the kinetic data agreed well with the detachment time constant measured from the motility assays. Linking the biochemistry with the observed diffusive motility indicates that ATP hydrolysis and movement are only weakly coupled in KIF1A. In addition to KIF1A motility, processive movement has also been reported for a monomeric kinesin construct derived from conventional kinesin (Inoue et al., 2001). When monomeric kinesin heads were fluorescently labeled and observed in a low ionic strength motility buffer, Inoue et al. observed microtubule binding and short-distance movement along microtubules (mean displacement 40 nm), suggesting these truncated heads work by a similar processive mechanism as KIF1A. When a fusion protein was constructed with a protein that normally binds weakly to microtubules, the run lengths were increased, consistent with adding a second nonspecific binding site to the motor. The high ATPase rates and diffusive character of the movement suggests a similar loose coupling between ATP hydrolysis and movement as seen in KIF1A. However, these results directly conflict with observations made on almost identical one-headed kinesin constructs from a number of laboratories where no processive motility was observed (Berliner et al., 1995, Hancock and Howard, 1998, Okada and Hirokawa, 1999, Pierce et al., 1999, Vale et al., 1996, Young et al., 1998). At present, the differences in experimental technique and data analysis that underlie these conflicting results are yet to be reconciled. KIF1A processivity clearly demonstrates that there are other mechanisms besides the hand-over-hand model to achieve processivity. It also sheds light on the question of whether conventional kinesin motility uses a concerted conformational change in the head domains or a diffusional mechanism in order for the free head to find its next binding site. An outstanding question is whether a one-headed processive motor can move against an external force. During the diffusive phases of the motility cycle, the motor would be expected to slip backwards against a load. Experiments using individual KIF1A motors bound to beads and held in an optical trap should shed light on this question. One possibility is that during vesicle transport in vivo multiple motors are working together such that the probability of diffusing and/or slipping backwards under load is greatly diminished. An important caveat to the KIF1A work is that other laboratories have failed to detect processive movement by homologous monomeric kinesins. Using a similar single molecule fluorescence assay, Pierce et al. (1999) failed to detect processive movement in UNC104, a KIF1A homolog from C. elegans, and Rogers et al. (2001) failed to detect processive movement in KIF1D. One possibility is that the conventional kinesin neck linker which is fused to KIF1A, contributes to processivity

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possibly by acting as a second positively charged tether. In support of this idea, it was shown that duplicating the neck sequence or adding extra positively charged residues to the neck coil of conventional kinesin increased the processive run length while adding negative charges reduced processivity (Romberg et al., 1998, Thorn et al., 2000). Also, cleaving the negatively charged E-hook from tubulin was found to decrease processivity in both wild-type and mutant conventional kinesin (Thorn et al., 2000, Wang and Sheetz, 2000), consistent with the idea that a second electrostatic interaction in addition to the normal kinesin-microtubule interface may help to keep motors associated with microtubules during processive movement. Hence, to understand whether single motor processivity of KIF1A can actually occur in cells it will be necessary to demonstrate processive movement of a KIF1A construct that does not contain the conventional kinesin neck linker domain. Recent experiments by Tomishige et al. (2002) suggest that KIF1A is able to dimerize and that in vivo motor function is best accounted for by this dimeric species. First, artificial dimerization of Unc104, the C. elegans ortholog of KIF1A, resulted in single-molecule motility velocities similar to those found in high-density assays and observed in vivo. This contrasts with the slow directed velocity of the biased diffusion observed by Okada et al. Second, increasing the concentration of KIF1A/Unc104 motors containing the wild-type dimerization domain in this single-molecule assay resulted in similar fast motility, indicating that dimerization of native motors is possible at sufficiently high motor concentrations. Third, using an optical tweezer assay, individual dimeric KIF1A/Unc104 motors were found to walk in 8 nm steps along the microtubule and generate similar forces to conventional kinesin. And fourth, in a multimotor liposome motility assay, KIF1A/Unc104 motors containing the putative dimerization domain transported the liposomes long distances at high speeds, while motors lacking the dimerization domain required higher motor concentrations for motility and never achieved high velocities. Hence, although Tomishige et al. did observe diffusional motility of KIF1A/Unc104 when fused to a positively charged neck linker, confirming the work of Okada et al., KIF1A/Unc104 motility in vivo is best explained by reversible dimerization, which activates the motors and may act as a mechanism for regulation. Interestingly, some myosins may use a similar electrostatic mechanism to stay attached to actin. A number of studies have suggested that, in addition to the conventional actin binding site on the myosin heavy chain, in many muscle types there is a positively charged Z 40 amino acid N-terminal extension of the myosin essential light chain (ELC), located at the base of the myosin head, that interacts with actin as well. Crosslinking studies (Sutoh, 1982) provided the first evidence for this interaction, and cryoEM reconstructions of myosin heads bound to actin filaments provide additional support: a region of the ELC was seen stretching roughly 8 nm across the myosin head and binding to the actin filament (Milligan et al., 1990). Furthermore, this interaction was demonstrated to have functional consequences. In myosins in which the N-terminal ELC extension was naturally absent or was mutated or deleted, the muscle shortening velocity (Sweeney, 1995) and the ATPase rate in solution were accelerated (Timson et al., 1998), while the apparent actin affinity was decreased (Timson et al., 1998). These data


suggest that in some myosin isoforms this second actin binding site increases the duration that myosin is bound to actin during each hydrolysis cycle, perhaps by helping to orient the myosin head to optimize actin binding. Hence, it appears that this performance-enhancing feature, a non-specific filament binding site that complements the primary filament binding site, may be a general feature of cytoskeletal motor proteins.

10.14

Unresolved Questions

To gain further insights into kinesin chemomechanical coupling, future investigations will require the convergence of single molecule mechanical studies, biochemical investigations and quantitative modeling. There are a number of mechanical questions to be resolved. First, how do external forces alter the biochemical rates in the hydrolysis cycle? Mechanical forces can alter both forward and reverse rate constants and the magnitude of the effect will depend on the precise direction of the applied forces. The influence of forces with components orthogonal to the direction of motion should give insight into the three-dimensional conformational changes underlying movement. A second mechanical question concerns the existence of substeps. The available biochemical and mechanical data predict there are at least two substeps in the kinesin walking mechanism. By examining the position of kinesin-coated beads stepping along a microtubules with microsecond time resolution, Nishiyama et al. (2001) recently found evidence for two 4-nm substeps in each 8-nm kinesin step. Further resolution of these substeps will likely require mutant motors that either have structural alterations that magnify subtle movements or have altered biochemical properties that slow the transitions sufficiently to observe the substeps. A third mechanical question concerns the overall conformational change that occurs in two-headed kinesin during its walking cycle. Because the two heads in conventional kinesin are identical peptides, it might be thought that they undergo identical conformational changes during their ATP hydrolysis cycles (a symmetric hand-over-hand mechanism). This differs from walking mechanisms analogous to left and right feet marching along a microtubule: a symmetric mechanism would mean that the two heads would rotate around one another, introducing a one-half twist per step (Howard, 1996). Such twisting up of the motors is not consistent with high-density assays, which show that tens to hundreds of motors can move microtubules for many microns: if each head rotated 180h per 8 nm, an impossible number of twists would be accumulated in each motor. Even in single-molecule assays where a motor takes up to 100 steps, symmetric models would imply an unfeasible amount of torsion (50 twists), which is expected to cause the microtubules in gliding assays to swivel in one direction during translocation across the surface. Hua et al. (2002), using surface-immobilized kinesin and low ATP levels to slow the stepping rate, carefully looked for evidence of microtubule rotation during

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References

translational movement and found no evidence of such twisting. Taken together, these observations imply that either the rotational strain is relieved during each step (asymmetric hand-over-hand model) or that the motor steps using some other mechanism that introduces no rotational strain in the two-headed motor. Motors with different directionality have already provided insights into structural aspects of the kinesin power stroke and continued characterization of kinesin family motors should provide further insight into the fundamentals of chemomechanical coupling in kinesins. A corollary of this is understanding population effects in non-processive motors because these motors work as aggregates in cells it will be important to understand motor properties that emerge when motors work together in aggregate. These questions will require the convergence of single-molecule measurements, enzyme-kinetic studies, structural investigations and theoretical modeling. The answers gleaned from these investigations will help us to understand not only kinesin mechanochemistry, but will help define the physical and molecular principles underlying all cellular movement.

Acknowledgements

W. O. H. acknowledges support from the Whitaker Foundation. J. H. acknowledges support from the NIH (AR-405930), the Max Planck Society and the Human Frontier Science Program. The authors thank Bill Schief for helpful comments on the manuscript.

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References Huang, T. G. and D. D. Hackney. 1994. Drosophila kinesin minimal motor domain expressed in Escherichia coli. Purification and kinetic characterization. J. Biol. Chem. 269: 16493 16501. Inoue, Y., A. H. Iwane, T. Miyai, E. Muto, and T. Yanagida. 2001. Motility of single oneheaded kinesin molecules along microtubules. Biophys. J. 81: 2838 2850. Jiang, W. and D. D. Hackney. 1997. Monomeric kinesin head domains hydrolyze multiple ATP molecules before release from a microtubule. J. Biol. Chem. 272: 5616 5621. Kawaguchi, K. and S. Ishiwata. 2001. Nucleotide-dependent single- to double-headed binding of kinesin. Science 291: 667 669. Keller, D. and C. Bustamante. 2000. The mechanochemistry of molecular motors. Biophys. J. 78: 541 556. Kikkawa, M., Y. Okada, and N. Hirokawa. 2000. 15 Å resolution model of the monomeric kinesin motor, KIF1A. Cell 100: 241 252. Kim, A. J. and S. A. Endow. 2000. A kinesin family tree. J. Cell Sci. 113: 3681 3682. Kojima, H., E. Muto, H. Higuchi, and T. Yanagida. 1997. Mechanics of single kinesin molecules measured by optical trapping nanometry. Biophys. J. 73: 2012 2022. Kozielski, F., S. Sack, A. Marx, M. Thormahlen, E. Schonbrunn, V. Biou, A. Thompson, E. M. Mandelkow, and E. Mandelkow. 1997. The crystal structure of dimeric kinesin and implications for microtubule-dependent motility. Cell 91: 985 994. Kull, F. J., E. P. Sablin, R. Lau, R. J. Fletterick, and R. D. Vale. 1996. Crystal structure of the kinesin motor domain reveals a structural similarity to myosin. Nature 380: 550 555. Lasek, R. J. and S. T. Brady. 1985. Attachment of transported vesicles to microtubules in axoplasm is facilitated by AMP-PNP. Nature 316: 645 647. Lymn, R. W., and E. W. Taylor. 1971. Mechanism of adenosine triphosphate hydrolysis by actomyosin. Biochemistry 10: 4617 4624. Ma, Y. Z. and E. W. Taylor. 1997a. Interacting head mechanism of microtubule-kinesin ATPase. J. Biol. Chem. 272: 724 730. Ma, Y. Z. and E. W. Taylor. 1997b. Kinetic mechanism of a monomeric kinesin construct. J. Biol. Chem. 272: 717 723. Matthies, H. J., R. J. Baskin, and R. S. Hawley. 2001. Orphan kinesin Nod lacks motile properties but does possess a microtubule-

stimulated ATPase activity. Mol. Biol. Cell 12: 4000 4012. Meyhofer, E. and J. Howard. 1995. The force generated by a single kinesin molecule against an elastic load. Proc. Natl Acad. Sci. USA 92: 574 578. Miki, H., M. Setou, K. Kaneshiro, and N. Hirokawa. 2001. All kinesin superfamily protein, KIF, genes in mouse and human. Proc. Natl Acad. Sci. USA 98: 7004 7011. Milligan, R. A., M. Whittaker, and D. Safer. 1990. Molecular structure of F-actin and location of surface binding sites. Nature 348: 217 221. Moyer, M. L., S. P. Gilbert, and K. A. Johnson. 1996. Purification and characterization of two monomeric kinesin constructs. Biochemistry 35:6321 6329. Nishiyama, M., E. Muto, Y. Inoue, T. Yanagida, and H. Higuchi. 2001. Substeps within the 8nm step of the ATPase cycle of single kinesin molecules. Nature Cell Biol. 3: 425 428. Okada, Y. and N. Hirokawa. 1999. A processive single-headed motor: kinesin superfamily protein KIF1A. Science 283: 1152 1157. Okada, Y., and N. Hirokawa. 2000. Mechanism of the single-headed processivity: diffusional anchoring between the K-loop of kinesin and the C terminus of tubulin. Proc. Natl Acad. Sci. USA 97: 640 645. Okada, Y., H. Yamazaki, Y. Sekine-Aizawa, and N. Hirokawa. 1995. The neuron-specific kinesin superfamily protein KIF1A is a unique monomeric motor for anterograde axonal transport of synaptic vesicle precursors. Cell 81: 769 780. Pierce, D. W., N. Hom-Booher, A. J. Otsuka, and R. D. Vale. 1999. Single-molecule behavior of monomeric and heteromeric kinesins. Biochemistry 38: 5412 5421. Ray, S., E. Meyhofer, R. A. Milligan, and J. Howard. 1993. Kinesin follows the microtubule’s protofilament axis. J. Cell Biol. 121: 1083 1093. Rice, S., A. W. Lin, D. Safer, C. L. Hart, N. Naber, B. O. Carragher, S. M. Cain, E. Pechatnikova, E. M. Wilson-Kubalek, M. Whittaker, E. Pate, R. Cooke, E. W. Taylor, R. A. Milligan, and R. D. Vale. 1999. A structural change in the kinesin motor protein that drives motility. Nature 402: 778 784. Rogers, K. R., S. Weiss, I. Crevel, P. J. Brophy, M. Geeves, and R. Cross. 2001. KIF1D is a fast non-processive kinesin that demonstrates

10 Kinesins: Processivity and Chemomechanical Coupling novel K-loop-dependent mechanochemistry. EMBO J. 20: 5101 5113. Romberg, L., D. W. Pierce, and R. D. Vale. 1998. Role of the kinesin neck region in processive microtubule-based motility. J. Cell Biol. 140: 1407 1416. Romberg, L. and R. D. Vale. 1993. Chemomechanical cycle of kinesin differs from that of myosin. Nature 361: 168 170. Rosenfeld, S. S., B. Rener, J. J. Correia, M. S. Mayo, and H. C. Cheung. 1996. Equilibrium studies of kinesin-nucleotide intermediates. J. Biol. Chem. 271: 9473 9482. Sablin, E. P., R. B. Case, S. C. Dai, C. L. Hart, A. Ruby, R. D. Vale, and R. J. Fletterick. 1998. Direction determination in the minus-enddirected kinesin motor ncd. Nature 395: 813 816. Sablin, E. P., F. J. Kull, R. Cooke, R. D. Vale, and R. J. Fletterick. 1996. Crystal structure of the motor domain of the kinesin-related motor ncd. Nature 380: 555 559. Schief, W. R. and J. Howard. 2001. Conformational changes during kinesin motility. Curr. Opin. Cell Biol. 13: 19 28. Schnitzer, M. J. and S. M. Block. 1997. Kinesin hydrolyses one ATP per 8-nm step. Nature 388: 386 390. Schnitzer, M. J., K. Visscher, and S. M. Block. 2000. Force production by single kinesin motors. Nature Cell Biol. 2: 718 723. Sosa, H., D. P. Dias, A. Hoenger, M. Whittaker, E. Wilson-Kubalek, E. Sablin, R. J. Fletterick, R. D. Vale, and R. A. Milligan. 1997. A model for the microtubule-Ncd motor protein complex obtained by cryo-electron microscopy and image analysis. Cell 90: 217 224. Stewart, R. J., J. P. Thaler, and L. S. Goldstein. 1993. Direction of microtubule movement is an intrinsic property of the motor domains of kinesin heavy chain and Drosophila ncd protein. Proc. Natl Acad. Sci. USA 90: 5209 5213. Sutoh, K. 1982. Identification of myosin-binding sites on the actin sequence. Biochemistry 21: 3654 3661. Svoboda, K., C. F. Schmidt, B. J. Schnapp, and S. M. Block. 1993. Direct observation of ki-

nesin stepping by optical trapping interferometry. Nature 365: 721 727. Sweeney, H. L. 1995. Function of the N terminus of the myosin essential light chain of vertebrate striated muscle. Biophys. J. 68: 112S 118S. Thorn, K. S., J. A. Ubersax, and R. D. Vale. 2000. Engineering the processive run length of the kinesin motor. J. Cell Biol. 151: 1093 1100. Timson, D. J., H. R. Trayer, and I. P. Trayer. 1998. The N-terminus of A1-type myosin essential light chains binds actin and modulates myosin motor function. Eur. J. Biochem. 255: 654 662. Tomishige, M., D. R. Klopfenstein and R. D. Vale. 2002. Conversion of Unc104/KIF1A kinesin into a processive motor after dimerization. Science 297: 2263–2267. Uyeda, T. Q., S. J. Kron, and J. A. Spudich. 1990. Myosin step size. Estimation from slow sliding movement of actin over low densities of heavy meromyosin. J. Mol. Biol. 214: 699 710. Vale, R. D. and R. A. Milligan. 2000. The way things move: looking under the hood of molecular motor proteins. Science 288: 88 95. Vale, R. D., T. Funatsu, D. W. Pierce, L. Romberg, Y. Harada, and T. Yanagida. 1996. Direct observation of single kinesin molecules moving along microtubules. Nature 380: 451 453. Visscher, K., M. J. Schnitzer, and S. M. Block. 1999. Single kinesin molecules studied with a molecular force clamp. Nature 400: 184 189. Wang, Z., and M. P. Sheetz. 2000. The C-terminus of tubulin increases cytoplasmic dynein and kinesin processivity. Biophys. J. 78: 1955 1964. Yajima, J., M. C. Alonso, R. A. Cross, and Y. Y. Toyoshima. 2002. Direct long-term observation of Kinesin processivity at low load. Curr. Biol. 12: 301 306. Young, E. C., H. K. Mahtani, and J. Gelles. 1998. One-headed kinesin derivatives move by a nonprocessive, low-duty ratio mechanism unlike that of two-headed kinesin. Biochemistry 37: 3467 3479.

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11 Quantitative Measurements of Myosin Movement In Vitro: The Reductionist Approach Carried to Single Molecules James A. Spudich

11.1

Introduction

The challenge of the postgenomic era is to understand the roles in vivo of every gene product and its molecular mechanism of action. The complexity of understanding a gene product in these ways is exemplified by the enormous interdisciplinary effort mobilized to elucidate the roles of the actin-based molecular motor myosin and how it transduces chemical energy into mechanical movement. No enzyme has been studied by more diverse approaches than myosin. Biophysics, biochemistry, physiology, classical genetics, and molecular genetics have all made their contributions. Hugh E. Huxley combined X-ray diffraction and electron microscopy studies on muscle and suggested the swinging cross-bridge model of muscle contraction in 1969 (Huxley, 1969). In the following decade, many important findings shaped our understanding of myosin function. Tension transient measurements using intact muscle (Huxley and Simmons, 1971), structural studies including those based on 3D-reconstructions from electron micrographs (Moore et al., 1970), actin-activated myosin ATPase kinetics (Lymn and Taylor, 1971), the development of nucleotide analogs to probe the ATPase cycle (Goody et al., 1972, Goody and Hofmann, 1980, Yount et al., 1971), and other approaches have all been important methods used to probe the myosin molecule. By the 1980s, time-resolved X-ray diffraction on living muscle fibers using synchrotron radiation, revealed changes in cross-bridge order that are kinetically consistent with tension development (Huxley et al., 1982). ATPase assays provided the kinetics of all of the rate and equilibrium constants, and thus the free energy changes associated with steps of the actin-activated myosin ATPase cycle. These are well understood features of the cross-bridge mechanism.

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11.2 Quantitative In Vitro Assays for Myosin Movement Established the Motor Domain of Myosin

11.2

Quantitative In Vitro Assays for Myosin Movement Established the Motor Domain of Myosin

Until the early 1980s, the field was limited by the lack of a quantitative in vitro assay to measure the essential function of myosin its ATP-driven movement along actin. First and foremost one must have a quantitative in vitro assay for the function of interest. Our early attempts in the 1970s to establish a quantitative in vitro motility assay were frustrating due to the difficulty in obtaining convincing, reproducible, quantitative movement of purified actin and myosin. One approach was to try to observe movement of polystyrene beads coated with purified myosin along surface-attached purified actin filaments that had been oriented by one means or another. The lack of convincing movement could have been due to improperly organized actin filaments, myosin-coated beads that were not functional, or to the possibility that movement required other components of the muscle sarcomere. Interestingly, this approach eventually worked (Spudich et al., 1985), but it was necessary to first backtrack and create a more complex assay than that currently used. To test whether actin filament organization was the problem, we relaxed our insistence that the system be totally pure, and we used the well-defined actin cables in Nitella (Kersey et al., 1976) as tracks to test movement of beads coated with myosin purified from rabbit muscle. This experiment worked on the first try and led to a quantitative measurement of movement of myosin along actin filaments in vitro (Sheetz and Spudich, 1983, Sheetz et al., 1984). The myosin-coated beads were clearly functional. Knowing this, it became easier to find ways to replace the complex Nitella substratum with purified, oriented actin filaments. We established more rigorous conditions to obtain good orientation, and for the first time observed quantitative movement of myosin-coated beads on actin filaments with a totally defined, purified system (Spudich et al., 1985). Strikingly, only pure actin and myosin and ATP were required to obtain movement at velocities essentially the same as that observed in live muscle under zero load. About the same time, Yanagida and coworkers demonstrated that single actin filaments labeled with phalloidin-tetramethylrhodamine can be visualized under a fluorescence microscope (Yanagida et al., 1984). Using this observation, we reversed the orientations of actin and myosin in our assay. We showed that movement of fluorescent actin filaments along myosin molecules attached to a surface could be easily quantitated (Kron and Spudich, 1986). Quantitative in vitro motility assays, developed from 1983 to 1986, have not only been invaluable tools but have led to the discovery and characterization of the kinesin family of molecular motors (Vale et al., 1985). The discoveries of a large family of kinesin motors and studies that led to a compelling model of how kinesin works (Rice et al., 1999), are pivotal in understanding the roles of molecular motors in vivo and how molecular motors work (Goldstein and Philp, 1999, Schliwa and Woehlke, 2001, Vale and Milligan, 2000) (see Chapter 10). For the actin-based myosin motor, quantitative in vitro motility assays provided the evidence needed to rule out some models of contraction, such as those involv-

11 Quantitative Measurements of Myosin Movement

ing conformational changes in the tail region of the molecule (Harrington, 1979). The cross-bridge itself (Subfragment 1 of myosin, or S1; Fig. 11.1) was shown directly and unequivocally to be the motor domain of myosin (Toyoshima et al., 1987). The tail of myosin-II has the critical function of forming bipolar thick filaments that anchor the S1 motor domains to a macromolecular assembly used in muscle contraction and in contractile processes such as cytokinesis in non-muscle cells (Robinson and Spudich, 2000). To understand the motor activity, attention focused on S1.

11.3

Structural Studies Revealed Putative Pre-stroke and Post-stroke States of the Myosin Head

One of the most important breakthroughs of the last decade in the field was the determination of the high resolution crystal structures of G-actin (Kabsch et al., 1990) and of S1 (Dominguez et al., 1998, Houdusse et al., 2000, Rayment et al., 1993a, Smith and Rayment, 1996). S1 from the conventional myosin II consists of a catalytic domain and a light chain binding domain (Fig. 11.1). The catalytic domain contains a nucleotide binding site of the P-loop variety that is closely associated with switch-I and switch-II helices. The nucleotide site is Z 4 nm away from the actin binding site, and these two sites communicate with one another via the switch I and switch II helices, which move in response to the state of the nucleotide, especially the presence or absence of Pi in the active site. Extending from the edge of the catalytic domain furthest from the actin binding site is an alpha helix that is surrounded by two calmodulin-like light chains. This light chain binding region has been suggested to act like a lever arm, which amplifies smaller conformational changes near the nucleotide binding site by swinging through a large angle from a pre-stroke state to a post-stroke state (Fig. 11.1). Strikingly, X-ray crystallography revealed at least two distinct states of the lever arm, one a putative prestroke state and another a putative post-stroke state (Dominguez et al., 1998, Houdusse et al., 2000, Rayment et al., 1993b, Smith and Rayment, 1996; Fig. 11.1). For an excellent review on the structural mechanism of muscle contraction, see Geeves and Holmes (1999). The asymmetric shape of S1 allowed the docking of the X-ray crystal structure of S1into low resolution 3D images generated by electron microscopy of the actin S1 complex (Rayment et al., 1993a,b, Schroeder et al., 1993). For some forms of myosin II, 3D reconstructions from electron micrographs in the presence and absence of ADP, revealed different conformations of the lever arm, suggesting that there is a small additional stroke that derives from the release of ADP from the active site (Jontes et al., 1995, Whittaker et al., 1995). While the static electron microscope and X-ray crystal structures are essential towards understanding how S1 works, important contributions have come from dynamic studies of nucleotide-dependent S1 conformational changes. Probes placed at the most reactive cysteine near the active site showed very little change in orien-

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11.3 Structural Studies Revealed Putative Pre-stroke and Post-stroke States of the Myosin Head

Pre-stroke state A

Pre-stroke state B

ADP.Pi

ADP.Pi

ATP

Pi

Post-stroke state

Figure 11.1. Pre- and post-stroke states of S1. stroke state. Pre-stroke state B was predicted

Structures modeled from the pre-stroke and post-stroke S1 crystal structures (Dominguez et al., 1998, Houdusse et al., 2000, Rayment et al., 1993a,b, Smith, 1996) are shown, with the catalytic domain of S1 kept in a fixed orientation. The lever arm moves through an angle of Z 70h going from pre-stroke state A to the post-

from FRET measurements (Shih et al., 2000). The red dye shows the position of an acceptor dye for the FRET measurements. The green and yellow dyes show two independent positions for placing a donor dye on the regulatory light chain. Figure modified from Shih et al. (2000).

tation during contractile events (Cooke et al., 1982). These results led to the suggestion that the banana-shaped S1 as observed by electron microscopy may actually swing not at the actin binding site, but rather in the middle of the S1 (Cooke, 1986). The subsequent crystal structures drove home this modified swinging cross-bridge model the major change in orientation was suggested to occur between the catalytic domain and the light chain-bound lever arm (Rayment et al., 1993a,b). In order to visualize the suggested change using a dynamic approach, molecular genetic manipulation of the S1, removing existing reactive cysteines and placing new ones at sites that should reveal large changes in lever arm orientation, was necessary. Using this approach, an Z 70-degree rotation of the lever arm was apparent with at least three distinct states of the lever arm position (Shih et al., 2000).


11.4

Single Molecule Analysis Revealed a Unitary Small Step in Motion as Myosin Interacts with Actin

The swinging cross-bridge model is only consistent with a small, Z 10-nm step in motion when myosin interacts with actin. In 1985, Yanagida et al. reported a much larger step (Z 100 nm) in motion for each ATP hydrolyzed, using single muscle sarcomeres from which the Z-lines, structures that anchor the thin filaments in the sarcomere, had been removed by protease treatment (Yanagida et al., 1985). Debate ensued for more than a decade about the size of the step taken when myosin interacts with actin, with some experiments suggesting a step size of Z 10 nm (Toyoshima et al., 1990, Uyeda et al., 1990, 1991) and others Z 100 nm or more (Harada et al., 1990, Saito et al., 1994, Yanagida and Harada, 1988). The large step sizes from the experiments of Yanagida and his colleagues led them to suggest that the actin filament may be the true motor rather than the myosin molecule. Such a concept was extended by Schutt and coworkers to a specific model of a ribbon to helix transition in the actin filament structure, with the myosin S1 acting simply as an anchor to link the thick and thin filaments together during the actin structural change (Schutt et al., 1995). Thus, the actin, rather than the myosin, was proposed to be the motor. Clearly, if the step size were considerably larger than Z 10 nm, then the conventional swinging cross-bridge model could not be correct. At times, nearly identical experiments using the in vitro motility assay system seemed to provide very different answers for the step size. Toyoshima et al. (1990), for example, measured Z 10 nm movement per ATP hydrolyzed per myosin head by measuring total movement in the in vitro motility assay and simultaneously measuring the total ATP hydrolyzed in the same time interval. Harada et al. (1990), on the other hand, carried out seemingly the same experiment and measured Z 100 nm of movement per ATP hydrolyzed per myosin head. One major issue in measurements using an ensemble of myosin molecules on the surface is the estimate of the number of myosin molecules that are bound in a functional manner, oriented properly, and within reach of the actin filament as it moves along the surface. The calculation of the step size per ATP hydrolyzed per molecule depends critically on this estimate. Given these complexities and other uncertainties that derived from these indirect measurements of the step size using an ensemble of myosin molecules, direct measurement of the step that occurs upon a single interaction between myosin and actin was necessary. A refinement of the in vitro motility assay permitted the direct measurement of myosin steps, one molecule at a time (Finer et al., 1994). This system used laser trapping to reduce the experiment to the single molecule level to establish that there is a unitary step in motion for conventional myosin-II of Z 5 15 nm, each step most probably linked to the turnover of a single ATP (Finer et al., 1994, although see Yanagida and Iwane (2000) and Ishii and Yanagida (Chapter 13) for an alternative view). An optical trap consists of a laser that is focused through a lens, usually a high numerical aperture microscope objective. A somewhat simplified view of the basis of the trapping considers that light has

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11.4 Single Molecule Analysis Revealed a Unitary Small Step in Motion as Myosin Interacts with Actin

momentum. As the photons are diffracted through a particle, like a small plastic bead, they are deflected away from the beam, which results in an equal and opposite momentum imparted on the bead directed into the laser beam. The bead is imaged on quadrant photodiode detectors, which reliably detect single displacements of the bead on the order of nanometers with millisecond time resolution. Our first design of a dual beam optical trap used acousto-optic deflectors to perform feedback on the position of the trap, but used mirror translations and separated beam paths for steering the traps (Finer et al., 1994). We are currently designing a system that uses a single set of two commercially available, orthogonal acousto-optic deflectors to create multiple traps and to rapidly adjust their position in the sample plane (Molloy, 1998, Visscher et al., 1996). This design allows control of positioning, position clamping, modulation of trap stiffness, and force clamping by changing the input signal to the acousto-optic deflectors. For a detailed review of this dual beam optical trap and of optical trapping in general, see Rice et al. (2002). Ishii and Yanagida (Chapter 13) describe this optical trapping approach and other approaches for single molecule analysis, and I refer you to their chapter for useful illustrations regarding the various methods. Such single molecule analyses, to some extent pioneered by the need to measure parameters of molecular motors (Finer et al., 1994, Svoboda et al., 1993), is becoming a field of its own. One might consider single molecule analysis the most modern form of biochemistry, where one no longer needs to infer what an enzyme is doing by examining an ensemble of enzyme molecules. An ensemble approach necessarily provides data about the average behavior of the molecules being examined and eliminates the ability to observe much of the interesting dynamics going on at the single molecule level. Thus, single molecule analysis is reductionism reduced to its most fundamental level. Yanagida and his coworkers have contributed in major ways to advance the field of single molecule analysis applied to molecular motors, including the use of total internal reflection fluorescence microscopy (TIRFM) to simultaneously measure nucleotide binding to and release from myosin and the stroke that occurs upon binding of myosin to actin (Funatsu et al., 1995, Ishijima et al., 1998). Together with Ron Vale, Yanagida and colleagues also measured the movement of single fluorescently-labeled kinesin molecules using TIRFM (Vale et al., 1996), which is the best way to determine the processivity of a molecular motor. The cumulative results from many laboratories over the last decade have led us back firmly to the view that myosin is indeed the motor and that major changes in the architecture of the actin filament do not occur. A most convincing piece of evidence for this in my view, is the observation that myosins can move along actin filaments that are firmly glued in place on an avidin-coated surface, where each actin monomer in the filament is biotinylated (Mehta et al., 1999, Rief et al., 2000, Sakamoto et al., 2000). Major changes in actin filament structure are not possible in this configuration. Furthermore, there is now widespread agreement that there is a small unitary step in motion (Spudich, 2001, Isshi and Yanagida, Chapter 13), although Yanagida and his colleagues express the view that the motor may take a number of such small steps for each ATP hydrolyzed, and


that the motor somehow remembers that it hydrolyzed an ATP molecule sometime in its past history (Isshi and Yanagida, Chapter 13). Isshi and Yanagida (Chapter 13) describe the myosin as the motor but they conclude that their results ‘indicate that the movement of molecular motors is driven by thermal motion rather than structural changes occurring in the motor molecules’. That thermal motion is important in the stepping process is an idea that stems back to the 1957 A. F. Huxley model (Huxley, 1957) and was revisited in a review by Vale and Oosawa (1990). Recent work on other myosin family members, myosin V and myosin VI, has clearly demonstrated a diffusive element to the stepping process (Moore et al., 2001, Nishikawa et al., 2002, Rock et al., 2001, Spudich and Rock, 2002, Veigel et al., 2002), but this is only one facet of the overall step. For myosin V, for example, the major part of the 36-nm step size is best described as resulting from a conformational change in the rather long lever arm (see below). Nucleotide-dependent conformational changes in the myosin molecule almost certainly provide both a major portion of the step in motion, depending on the myosin type, as well as the directionality of the movement by biasing the motor in that particular direction.

11.5

Molecular Genetic Approaches Have Indicated Roles of Various Domains and Specific Residues of the Myosin Motor

Another important development in the study of myosin has been the use of molecular genetics to dissect the roles of various domains and specific residues of S1 (for review, see Ruppel and Spudich, 1996). For example, the coupling of phosphate release with actin binding, and the coupling of the conformational change of the lever arm swing with phosphate release were emphasized studying a myosin II with a serine to leucine mutation near the active site (Murphy et al., 2001). The mutation causes premature release of phosphate, before the S1 binds tightly to actin, which results in an uncoupling of the ATP hydrolysis cycle and the mechanical motion. These data provide support for the view that the system is engineered to assure that the conformational change of the lever arm position from a prestroke state to a post-stroke state occurs only after the myosin has bound strongly to the actin filament. The data assembled from these multifaceted approaches applied to this research area best fits the following model (Fig. 11.2). The myosin II cross-bridge binds to ATP, and then releases its attached actin filament. The cross-bridge then hydrolyzes the ATP, and primes itself in preparation for a productive working stroke. Actin rebinding triggers phosphate release, which in turn prompts the myosin cross-bridge to return to its starting conformation, in a motion termed the ‘power stroke’. This power stroke involves a relatively fixed catalytic domain bound to actin and swinging of the light chain binding region through a considerable angle, providing a working stroke of 5 15 nm. The net result is that the attached actin filament is translocated in the direction of its minus (pointed) end. While Fig. 11.2A is simplified to show only forward directions around the cycle, reversi-

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11.6 Myosin V uses its Longer Lever Arm to Take a Larger Step along Actin

Figure 11.2. The actin-activated myosin-II AT-

Pase cycle. (A) The states of the myosin S1 domain that are strongly attached to actin are shown in red. For myosin-II the strongly attached state time is only Z 5 % of the total cycle time. That is, the myosin spends most of its time off the actin or only weakly associated (yellow states). The S1 structures shown were modeled based on various crystal structure determinations (Dominguez et al., 1998, Hou-

dusse et al., 2000, Rayment et al., 1993a.b, Smith, 1996). The S1s and F-actin are drawn to scale. (B) All of the myosin and actin myosin nucleotide states are shown. All steps are reversible. A, actin; M, myosin, AM, the actin myosin complex. A primary path corresponding to the structures shown in Fig. 11.2A is AM to AM.ATP to M.ATP to M.ADP.Pi to AM.ADP.Pi to AM.ADP to AM.

bility of the steps is established and important (Fig. 11.2B). For example, rebinding of myosin ATP and myosin ADP to actin is rapid, and the complexes actin myosin ATP and actin myosin ADP are important intermediates. While this model is widely accepted, interpretation of data has been at times difficult and controversial. This is partly due to the relatively small step size of the muscle type myosin II and the fact that the molecule spends a very short time in a strongly bound state to actin.

11.6

Myosin V uses its Longer Lever Arm to Take a Larger Step along Actin

The clearest evidence for nucleotide-dependent coupling of conformational changes in myosin leading to stepping of myosin along an actin filament has resulted from initial genetic experiments on mutant mice with reduced hair coloring. The authors of that work determined that the defective gene leading to the ‘dilute phenotype’ coded for a new member of the myosin family, myosin V (Mercer et al., 1991). This myosin has a typical catalytic domain for a myosin molecule but the lever arm is three times longer than that of myosin II (Cheney et al., 1993). The tail of the myosin V is very different from that of myosin II, presumably because


its role is to bind to vesicular cargo rather than to form bipolar thick filaments. The unique feature of the myosin V S1, the long lever arm, should, according to the swinging cross-bridge model, allow for a much longer step size than that observed for myosin II. In a relatively short time, studies of myosin V have revealed its probable mechanism of action, and further clarified how the myosin family of molecular motors works. The advantages of studying the myosin V motor are that it is built to take an exceedingly large step along actin and it remains tightly bound to actin for a large part of its ATPase cycle. These characteristics allow the myosin V to move processively along an actin filament. These properties are essential when one considers the task that this molecule performs in vivo. In melanocytes, melanin-containing vesicles are presumably carried along actin filaments anchored in the cell cortex. Not all faces of the actin filaments are available to the myosin and yet it walks processively along the actin in order to move the melanin-containing vesicles into the dendritic spines of the melanocytes, from where they are taken up by keratinocytes. It is not surprising then that the step size of myosin V is Z 36 nm (Mehta et al., 1999, Moore et al., 2001, Rief et al., 2000, Spudich and Rock, 2002, Veigel et al., 2002) (Fig. 11.3), the long-pitch helical pseudorepeat of the actin filament. Furthermore, myosin V moves processively along actin filaments in vitro even when those filaments are tightly adhering to a surface along one face of the filament (Mehta et al., 1999, Rief et al., 2000, Sakamoto et al., 2000). In this configuration it is impossible for myosin-V to follow the long pitch actin helix as it moves along the actin; it clearly must step from one helical crossover point of the filament to the other, as viewed looking straight down onto the surface-attached actin filament. The conformation of the myosin V while strongly bound to actin has been revealed by electron microscopy (Walker et al., 2000). The authors point out that in low concentrations of ATP, when the myosin

Experimental scheme of the force feedback enhanced laser trap. A feedback loop keeps the distance between the polystyrene bead center (gray curve) and the laser trap center (lower black curve) constant as the

Figure 11.3.

myosin V molecule steps along the actin filament. Thus, the myosin V is kept under constant load as it moves ( Rief et al., 2000, Visscher et al., 1999). Figure is from Rief et al. (2000).

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11.6 Myosin V uses its Longer Lever Arm to Take a Larger Step along Actin

heads both remain bound to the actin filament, the molecule often appears to be straining forward, in the form of a ‘telemark skier’. A combination of biochemical studies and single molecule analyses on myosin V provide compelling evidence that the dwell time between successive 36-nm steps is limited by ATP binding at sub-saturating ATP concentrations and by ADP release in the presence of saturating ATP (De La Cruz et al., 1999, 2000, Rief et al., 2000). Affinity constants and on and off rate constants for ATP and ADP calculated from traditional biochemical approaches are remarkably consistent with those revealed by single molecule analysis (De La Cruz et al., 1999, Trybus et al., 1999, Wang et al., 2000). By examining the myosin V molecules one at a time, one can distinguish mechanistically between slowing the rate of stepping due to sub-saturating ATP concentrations versus due to competition of binding of ADP and ATP for the active site (Rief et al., 2000). An unusually low myosin V ATPase activity reported by Ando and coworkers (Sakamoto et al., 2000) led them to suggest that myosin V may move 400 nm for each ATP hydrolyzed. Myosin V ATPase has subsequently been shown to be much higher, 13 s 1 (De La Cruz et al., 1999). This value is consistent with one Z 36-nm step per ATP hydrolysis and the observed velocity of Z 0.5 mm s 1. An elegant recent study on myosin V using single molecule analysis and polarization of fluorescence of lever arm labeled molecules has allowed Goldman and his colleagues to essentially visualize the myosin V stepping directly (Forkey et al., 2002). Thus, the combination of biochemical, structural and single molecule analyses of the last few years provides strong evidence for a mechanism of movement of this myosin along an actin filament that involves armover-arm 36-nm steps that are limited by the release of ADP from the rearward head (Fig. 11.4). Studies on myosin V have also revealed another element in the unitary step that involves a Brownian ratchet type of highly diffusive motion (Moore et al., 2001, Veigel et al., 2002). This is in keeping with suggestions of a significant Brownian ratchet component to molecular motor movement (Huxley, 1957, Rice et al., 1999, Vale and Oosawa, 1990, Isshi and Yanagida, Chapter 13). Thus, the working stroke provided by rotation of the long lever arm is likely to be about 20 to 25 nm. The rest of the 36-nm stride distance results from a diffusive Brownian motion that allows the head to find and bind to the appropriate actin site along the helical actin filament. Thus, part of the step involves a conventional lever arm stroke resulting from a conformational change in the myosin head, while another part derives from Brownian motion. The distribution between these two mechanistic forms of motion is overwhelmingly in the Brownian motion direction for myosin VI (Nishikawa et al., 2002, Rock et al., 2001), where a very short lever arm may result in a relatively small part of the overall step, probably to bias the directionality of the movement of this processive motor. Yanagida, Ikebe and colleagues (Tanaka et al., 2002) showed that a chimeric molecule consisting of a myosin V catalytic domain, only one IQ motif of the myosin V neck region (corresponding to a very short lever arm), and part of a smooth muscle myosin tail can take relatively large steps. Analysis of their data indicates that the steps are closer to Z 30 nm, not 36 nm, many backward steps are seen, and a histogram of the number of


A model for the nucleotide-dependent processive stepping of myosin V along an actin filament. As illustrated on the lower right, myosin V dwells with both heads attached to the actin filament, the leading head with ADP and the trailing head in rigor. ATP binding to the trailing head promotes its dissociation from actin, and forward movement of the released head then discharges intramolecular strain. The previous leading head then becomes the trailing head (lower left). Note that the trailing head moves 72 nm to reach its new site of attachment, but this results in only a 36-nm step of the body of the myosin V or of any cargo attached to the cargo attachment domain. The new, detached leading head quickly hydrolyzes

Figure 11.4.

ATP and then binds actin. Force generation follows either actin binding or phosphate release, which itself occurs either concomitant with or immediately following actin binding. These steps are fast relative to ADP release. At this point, one finds the molecule in its kinetically dominant state: both heads bound to actin and ADP, the leading head in a pre-stroke configuration and the trailing head in a poststroke configuration (upper right). The leading head is stressed against the direction of motion and the trailing head is stressed along this direction, an asymmetry that should bias the following ADP release to occur at the trailing head and not the leading head in this state.

events with a particular step size is much broader than that of the native myosin V. This diffusive character of the stepping and the significant amount of backward stepping is very reminiscent of the movement of myosin VI along actin (Nishikawa et al., 2002, Rock et al., 2001). This unusual chimeric construct seems to have provided the myosin V with characteristics that are not those of the native myosin V molecule. The latter shows a fairly sharp histogram of stepping of 36 nm plus or minus about one actin subunit on a protofilament of the F-actin, and it shows very few backward steps. It is as if the Brownian motion aspect of the native myosin V

281

282

11.7 Conclusions and Perspectives

movement has been accentuated in this myosin V smooth muscle chimeric molecule, and the higher level of precision of the normal movement of the native myosin V molecule has been lost. A single IQ motif myosin V, generated by simply removing five of the six IQ motifs from the native gene, results in a molecule that can only produce small steps in motion, as would be predicted by the lever arm hypothesis. Moreover, two-headed myosin V molecules constructed with only four light chains per head are still processive, but their step size is reduced to 24 nm. (T. Purcell, C. Morris, J. A. Spudich, and H. L. Sweeney, Proc. Natl. Acad. Sci. USA, in press).

11.7

Conclusions and Perspectives

In summary, many essential elements of the mechanism of myosin action are now generally agreed on. Myosin, not actin, is the motor. The motor domain is the myosin head domain. Myosin takes small steps along the actin filament. Large changes in actin filament structure do not occur during the translocation. Large conformational changes occur in the myosin head domain in a nucleotide-dependent manner. The conformational change in myosin II predicted by structural studies and dynamic biophysical measurements can account for much, if not all, of the step in motion observed by single molecule analysis. Some myosin family members show considerable diffusive searching during the stepping. One remaining question in the field relates to the extent of the contribution of motion derived from Brownian motion. Ishii and Yanagida (Chapter 13) take the extreme view that none of the motion derives from the conformational change. Another remaining issue is whether the system is tightly coupled, where each step involves one ATP hydrolysis event, or whether it is conceivable that there might be multiple steps from one ATP hydrolysis event (Ishii and Yanagida, Chapter 13). As described above, evidence for tight coupling is strongest in the case of myosin V, where the data is extremely difficult to explain in any other way (Rief et al., 2000). What needs to be done? While a crystal structure of F-actin has not been obtained, the Holmes et al. model of F-actin is well founded (Holmes et al., 1990, Lorenz et al., 1993, 1995) and likely to be correct for the most part. A crystal structure of the F-actin S1 complex, however, is essential. The high-resolution structure of the actin-bound state of S1 will reveal important aspects of the actin myosin interface and clarify how the communication between the actin binding site, the nucleotide site and the lever arm occurs. Details of strain-dependent changes in nucleotide affinities, so critical in models of myosin function, need to be understood. Extensive mutational analysis will help answer these and other remaining questions. In particular, modern tools, and especially single molecule analyses, will provide the essential dynamic measurements for further understanding how the myosin enzyme transforms the energy from ATP hydrolysis into mechanical movement.


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sin complex and its implications for muscle contraction. Science 261: 58 65. Rayment, I., W. R. Rypniewski, K. SchmidtBase, R. Smith, D. R. Tomchick, M. M. Benning, D. A. Winkelmann, G. Wesenberg, and H. M. Holden. 1993b. Three-dimensional structure of myosin subfragment-1: a molecular motor. Science 261: 50 58. Rice, S., A. W. Lin, D. Safer, C. L. Hart, N. Naber, B. O. Carragher, S. M. Cain, E. Pechatnikova, E. M. Wilson-Kubalek, M. Whittaker, E. Pate, R. Cooke, E. W. Taylor, R. A. Milligan, and R. D. Vale. 1999. A structural change in the kinesin motor protein that drives motility. Nature 402: 778 784. Rice, S., T. Purcell, and J. A. Spudich. 2002. Building and using optical traps to study properties of molecular motors. Methods Enzymol. (in press). Rief, M., R. S. Rock, A. D. Mehta, M. S. Mooseker, R. E. Cheney, and J. A. Spudich. 2000. Myosin-V stepping kinetics: a molecular model for processivity. Proc. Natl Acad. Sci. USA 97: 9482 9486. Robinson, D. N. and J. A. Spudich. 2000. Towards a molecular understanding of cytokinesis. Trends Cell Biol. 10: 228 237. Rock, R. S., S. E. Rice, A. L. Wells, T. J. Purcell, J. A. Spudich, and H. L. Sweeney. 2001. Myosin VI is a processive motor with a large step size. Proc. Natl Acad. Sci. USA 98: 13655 13659. Ruppel, K. M. and J. A. Spudich. 1996. Structure-function analysis of the motor domain of myosin. Ann. Rev. Cell Dev. Biol. 12: 543 573. Saito, K., T. Aoki, and T. Yanagida. 1994. Movement of single myosin filaments and myosin step size on an actin filament suspended in solution by a laser trap. Biophys. J. 66: 769 777. Sakamoto, T., I. Amitani, E. Yokota, and T. Ando. 2000. Direct observation of processive movement by individual myosin V molecules. Biochem. Biophys. Res. Commun. 272: 586 590. Schliwa, M. and G. Woehlke. 2001. Molecular motors. Switching on kinesin. Nature 411: 424 425. Schroeder, R. R., D. J. Manstein, W. Jahn, H. Holden, I. Rayment, K. C. Holmes, and J. A. Spudich. 1993. Three-dimensional atomic model of F-actin decorated with Dictyostelium myosin S1. Nature 364: 171 174.

11 Quantitative Measurements of Myosin Movement Schutt, C. E., M. D. Rozycki, J. K. Chik, and U. Lindberg. 1995. Structural studies on the ribbon-to-helix transition in profilin: actin crystals. Biophys. J. 68: 12S 17S; discussion 17S 18S. Sheetz, M. P. and J. A. Spudich. 1983. Movement of myosin-coated fluorescent beads on actin cables in vitro. Nature 303: 31 35. Sheetz, M. P., R. Chasan, and J. A. Spudich. 1984. ATP-dependent movement of myosin in vitro: characterization of a quantitative assay. J. Cell Biol. 99: 1867 1871. Shih, W. M., Z. Gryczynski, J. R. Lakowicz, and J. A. Spudich. 2000. A FRET-based sensor reveals large ATP hydrolysis-induced conformational changes and three distinct states of the molecular motor myosin. Cell 102: 683 694. Smith, C. A., and Rayment, I. 1996. X-ray structure of the magnesium (II) ADP.vanadate complex of the Dictyostelium discoideum myosin motor domain to 1.9 A resolution. Biochemistry 35: 5404 5417. Spudich, J. A. 2001. The myosin swinging cross-bridge model. Nature Rev. Mol. Cell Biol. 2: 387 392. Spudich, J. A. and R. S. Rock. 2002. A crossbridge too far. Nature Cell Biol. 4: E8 E10. Spudich, J. A., S. J. Kron, and M. P. Sheetz. 1985. Movement of myosin-coated beads on oriented filaments reconstituted from purified actin. Nature 315: 584 586. Svoboda, K., C. F. Schmidt, B. J. Schnapp, and S. M. Block. 1993. Direct observation of kinesin stepping by optical trapping interferometry. Nature 365: 721 727. Tanaka, H., K. Homma, A. H. Iwane, E. Katayama, R. Ikebe, J. Saito, T. Yanagida, and M. Ikebe. 2002. The motor domain determines the large step of myosin-V. Nature 415: 192 195. Toyoshima, Y. Y., S. J. Kron, E. M. Mc Nally, K. Niebling, C. Toyoshima, and J. A. Spudich. 1987. Myosin subfragment-1 is sufficient to move actin filaments in vitro. Nature 328: 536 539. Toyoshima, Y. Y., S. J. Kron, and J. A. Spudich. 1990. The myosin step size: measurement of the unit displacement per ATP hydrolyzed in an in vitro assay. Proc. Natl Acad. Sci. USA 87: 7130 7134. Trybus, K. M., E. Krementsova, and Y. Freyzon. 1999. Kinetic characterization of a mono-

meric unconventional myosin V construct. J. Biol. Chem. 274: 27448 27456. Uyeda, T. P. Q., S. J. Kron, and J. A. Spudich. 1990. Myosin step size estimation from slow sliding movement of actin over low densities of heavy meromyosin. J. Mol. Biol. 214: 699 710. Uyeda, T. Q., H. M. Warrick, S. J. Kron, and J. A. Spudich. 1991. Quantized velocities at low myosin densities in an in vitro motility assay. Nature 352: 307 311. Vale, R. D. and R. A. Milligan. 2000. The way things move: looking under the hood of molecular motor proteins. Science 288: 88 95. Vale, R. D. and F. Oosawa. 1990. Protein motors and Maxwell’s demons: does mechanochemical transduction involve a thermal ratchet? Adv. Biophys. 26: 97 134. Vale, R. D., T. Funatsu, D. W. Pierce, L. Romberg, Y. Harada, and T. Yanagida. 1996. Direct observation of single kinesin molecules moving along microtubules. Nature 380: 451 453. Vale, R. D., T. S. Reese, and M. P. Sheetz. 1985. Identification of a novel force-generating protein, kinesin, involved in microtubulebased motility. Cell 42: 39 50. Veigel, C., F. Wang, M. L. Bartoo, J. R. Sellers, and J. E. Molloy. 2002. The gated gait of the processive molecular motor, myosin V. Nature Cell Biol. 4: 59 65. Visscher, K., S. P. Gross, and S. M. Block. 1996. Construction of multiple-beam optical traps with nanometer-level position sensing. IEEE J. Sel. Top. Quant. Electr. 2: 1066 1076. Visscher, K., M. J. Schnitzer, and S. M. Block. 1999. Single kinesin molecules studied with a molecular force clamp. Nature 400: 184 189. Walker, M. L., S. A. Burgess, J. R. Sellers, F. Wang, J. A. Hammer, J. Trinick, and P. J. Knight. 2000. Two-headed binding of a processive myosin to F-actin. Nature 405: 804 807. Wang, F., L. Chen, O. Arcucci, E. V. Harvey, B. Bowers, Y. Xu, J. A. Hammer, and J. R. Sellers. 2000. Effect of ADP and ionic strength on the kinetic and motile properties of recombinant mouse myosin V. J. Biol. Chem. 275: 4329 4335. Whittaker, M., E. M. W. Kubalek, J. E. Smith, L. Faust, R. A. Milligan, and H. L. Sweeney.

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References 1995. A 35-A movement of smooth muscle induced by a myosin crossbridge during myosin on ADP release. Nature 387: one ATP hydrolysis cycle. Nature 316: 748 751. 366 369. Yanagida, T. and Y. Harada. 1988. Minimum Yanagida, T., M. Nakase, K. Nishiyama, and F. structural unit required for energy transducOosawa. 1984. Direct observation of motion tion in muscle. Adv. Exp. Med. Biol. 226: of single F-actin filaments in the presence of 277 287. myosin. Nature 307: 58 60. Yanagida, T. and A. H. Iwane. 2000. A large Yount, R. G., D. Ojala, and D. Babcock. 1971. step for myosin. Proc. Natl Acad. Sci. USA 97: Interaction of P–N–P and P–C–P analogs of 9357 9359. adenosine triphosphate with heavy meroYanagida, T., T. Arata, and F. Oosawa. 1985. myosin, myosin, and actomyosin. BiochemisSliding distance of actin filament try 10: 2490 2496.

12 Structures of Kinesin Motor Domains: Implications for Conformational Switching Involved in Mechanochemical Coupling Y.-H. Song, A. Marx and E. Mandelkow

12.1

Introduction

The family of kinesin-like proteins has currently more than 250 known members (kinesin home page: www.proweb.org/kinesin) and a large number of these kinesins has been detected using molecular biological methods (Kim and Endow, 2000). For example, at least 45 different transcripts of KIFs are expressed in the human body, 38 of these alone are found in human brain tissue (Miki et al., 2001). The founding member of the family, ‘conventional’ kinesin, comprises three domains: the N-terminal motor domain, the central stalk and the C-terminal light chain-binding domain (Goldstein and Philp, 1999, Woehlke and Schliwa, 2000). The motor domain is around 325 residues long and can be located at various places within the molecule (N-terminal, internal, or C-terminal; for an overview see Chapter 3 by Hirokawa and Takemura). These proteins are responsible for various intracellular transport processes but all of them share a common feature, the kinesin motor domain. Different kinesin-related proteins can transport vesicles in opposite directions along microtubules: Conventional kinesin is a highly processive microtubule-dependent motor that moves vesicles along microtubules in the plus direction towards the cell periphery. There are also kinesin related proteins like Ncd or Kar3 which can move their cargo in the opposite direction towards the cell center (Endow, 1999). The molecular basis of directional movement is discussed in the chapter by Endow. This chapter is concerned with the comparison of the known structures of kinesin motor domains and related folds in myosin and Gproteins. An attempt is made to gain more insight into what may be the conformational alteration along the pathway of the ATPase cycle which is related to the generation of force along microtubules (see also Chapter 10 by Hancock and Howard). Illustrations of structures and animations can be found at www.mpasmb-hamburg.mpg.de/ktdock.

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12.2 Structures of Kinesin Motor Domains

12.2

Structures of Kinesin Motor Domains

The structures of the motor domain have been analyzed crystallographically on several kinesin head constructs from different kinesins (Table 12.1). The processive monomeric kinesin Kif1A so far is the only one for which the structure has been solved in two different nucleotide states, the AMPPCP- and the ADPbound form (pdb: 1I6I and 1I5S; Kikkawa et al., 2001). The head domain of kinesin can be subdivided into a catalytical core domain of Z 325 residues responsible for the ATPase activity and the binding to microtubules, a ‘linker’ region (residues Z 325 340) which connects to the ‘neck’ (residues Z 340 370) and the beginning of the coiled-coil stalk (Fig. 12.1a and 12.2a; numbers refer to rat brain kinesin un-

12 Structures of Kinesin Motor Domains Compilation of kinesin and kinesin-related motor constructs used in X-ray crystallography structure determinations.

Table 12.1.

Construct

Sourceb

HsK349

H. sapiens KHC

RnK354

PDB code

Reference

2 349

1BG2

Kull et al. (1996)

R. norvegicus KHC

2 354

2KIN

Sack e al. (1997)

RnK379 (dimer)

R. norvegicus KHC

2 379

3KIN

Kozielski et al. (1997)

NcK355

N. crassa KHC

1 355

2GOJ

Song et al. (2001)

Eg5

H. sapiens KSP (HsEg5)

1 368

Kif1A (ADP)

Residues

M. musculus KIF1A

1II6

Turner et al. (2001)

1 355

a

1I5S

Kikkawa et al. (2001)

a

1I6I

Kikkawa et al. (2001)

Kif1A (AMPPCP)

M. musculus KIF1A

1 355

Ncd

D. melanogaster Ncd

355 700

Ncd (dimer)

D. melanogaster Ncd

281 700

2NCD

Sablin et al. (1998)

Ncd (dimer)

D. melanogaster Ncd

295 700

1CZ7

Kozielski et al. (1999)

Kar3 wt

S. cerevisiae Kar3

383 729

3KAR

Gulick et al. (1998)

Kar3 wt

S. cerevisiae Kar3

372 729

1F9T

Yun et al. (2001)

Kar3-N650K

S. cerevisiae Kar3

383 729

1F9U

Yun et al. (2001)

Kar3-R598A

S. cerevisiae Kar3

383 729

1F9V

Yun et al. (2001)

Kar3-E631A

S. cerevisiae Kar3

383 729

1F9W

Yun et al. (2001)

Sablin et al. (1996)

a

Chimeric construct with additional six residues from the neck linker of mouse conventional kinesin and a His tag.

b

Source names according to the Kinesin Home Page (www.proweb.org/kinesin).

m Figure 12.1. Structure of kinesin. (A) X-ray

structure of a monomeric construct of rat kinesin containing 354 residues of the motor domain, including the neck linker and the initial part of the neck helix (pdb code: 2KIN, Sack et al., 1997). The motor domain contains an eightstranded central b-sheet b1–b8) with a topological order of the b-strands 2-1-8-3-7-6-4-5, surrounded by six a-helices, three on either side. This fold represents a ‘Walker’ fold of a nucleotide-binding protein. (Walker et al., 1982). The nucleotide (ADP) is located at the upper end (ball-and-stick model). The neck linker b9 b10 is closely attached to the body (‘docked’). (B) Ribbon diagram of a hypothetical hybrid structure of kinesin and Ncd, showing the different relationships between the core motor domain and the neck. The core motor domain (b0–a6, gray) is taken from the rat kinesin and agrees closely with that of Ncd. The C-terminal ‘neck linker’ b9 b10 and ‘neck helix’ a7 of rat kinesin are shown in red and the N-terminal ‘neck linker’ and ‘neck helix’ of Ncd in green. Kinesin is seen from the opposite side

of the microtubule-binding surface. The second head in the Ncd dimer structure would be rotated by 180h around the Ncd neck helix. The second head in the rat brain kinesin dimer structure would be rotated by Z 120h around the kinesin neck helix. (C) Superposition of X-ray crystal structure of rat brain kinesin (3KIN; Kozielski et al., 1997) and NMR solution structure of neck-hinge peptide K357-D386 in several variations (Seeberger et al., 2000). Both structures agree in the common region of the neck helix K357 to W370, but the X-ray structure disappears beyond that point due to disorder, and the NMR structure reveals several short structural elements (extended chain, partial helical turns) linked by flexible joints (several structure solutions shown in orange). Due to this flexibility, the motor domains could easily rotate or tilt, relative to the stalk, which would be important in adopting the proper orientation for binding to a microtubule. Further structures and models can be viewed at www.mpasmb-hamburg.mpg.de/ktdock.

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290


less otherwise stated). The ‘neck linker’ of 10 15 residues is highly conserved among all plus-ended motors. In the minus-directed motor Ncd, a ‘neck helix’ with only a short ‘neck linker’ emanates at the N-terminus (Sablin et al., 1998; Fig. 12.1b). A surprising feature of the structure of kinesin motor domains is its similarity to myosin, an actin-dependent motor, and Ras, a G-protein, which share a similar fold in their core (Kull et al., 1996, 1998, Sablin et al., 1996). 12.2.1

General Features of the Catalytic Core

The folds of all known kinesin structures within the catalytic core are similar, as expected from the sequence homology (around 40 % identity between kinesins). The central eight-stranded b-sheet is surrounded by six a-helices (a1 a6), three on either side. Figures 12.1 and 12.2 represent the overall fold of the motor domain as ribbon diagrams. Analysis of the conformational variations among the kinesin structures shows that they are concentrated in certain areas of the kinesin molecule (Kull and Endow, 2002, Sack et al., 1999). We have now extended these analyses into the kinesin structures which have been published since then. The highest deviations between kinesins are found in those regions which have been proposed to change conformation during the ATPase cycle, namely switch 1, switch 2, and the microtubule-binding surface. Switch 1 and 2 are in close proximity to the active site (Fig. 12.1a). There are also large structural differences in the regions of the neck linker and the neck. These regions are largely undefined in most kinesin structures, which indicates a highly mobile structure in this region. A well-ordered neck linker (docked onto the body of the motor domain) and a-helical neck is found in rat kinesin (monomeric and dimeric, with bound ADP; Kozielski et al., 1997, Sack et al., 1997). The neck linker is more disordered, semi-detached but visible in NcKin (Song et al., 2001), Eg5 (Turner et al., 2001) and with reservation in Kif1A complexed with AMPPCP (Kikkawa et al., 2001), and it is invisible due to disorder in the structure of human kinesin (Kull et al., 1996). Eg5 is an exceptional case, as the neck linker is almost perpendicular to that expected for the docked conformation. In Ncd, a C-terminal motor, the non-helical linker is very short and enters immediately into the neck helix, in contrast to the extended linker of plus-end directed motors (Sablin et al., 1998). Finally, in the case of Kif1A, the neck linker has been truncated and partially replaced by residues from the sequence of mouse conventional kinesin (from within b9 to the start of b10 according to the structure of rat kinesin), plus a His-tag. Thus, the structure of the linker must be considered with caution.

12 Structures of Kinesin Motor Domains

12.2.2

The Nucleotide-Binding Active Site

The active site is formed by four structural elements, termed N1 N4 (Sablin et al., 1996). They include the highly conserved residues which are essential for the proper binding of ATP and its hydrolysis. The overall topology of this region is similar to that of myosin and the GTPases (Table 12.2). Table 12.2. Nucleotide-binding motifs in kinesin, myosin and G-proteins (p21ras). Pdb codes and

sequence numbering: rat kinesin: 2KIN, (Sack et al., 1997); chicken smooth muscle myosin: 1BR1, (Dominguez et al., 1998); p21ras: 1QRA, (Scheidig et al., 1995). Protein

Nucleotide

N1 P-loop

N2 Switch 1

N3 Switch 2

N4 base

Kinesin (rat)

ATP

G86QTxxGKS/T

N199xxSSR

D232xxGxE

R14xRP

Myosin (chicken smooth muscle)

ATP

G177ESxxGKS/T

N242xxSSR

D465xxGxE

N125P

P21ras

GTP

G10xxxxGKS

T35

D57xxG

N116KxD

N1, also called P-loop (G86xxxGKS/T, residue numbering according to rat kinesin) This is a common phosphate-binding motif for proteins which bind a mononucleotide (Schulz, 1992). The b-phosphate of adenine or guanosine di- or tri-phosphate is wrapped by this motif, where the conserved lysine (K92) with its positive charge supplies the proper electrical environment for the negatively-charged phosphate. The serine or threonine (T93) coordinates the cofactor, a Mg2 ion. This motif is essential for nucleotide binding, and the structure is highly conserved among ATP or GTP-binding proteins (rms I 0.3 Å), so that the structures of such proteins can be superimposed on the basis of this motif. 12.2.2.1

N2 Switch 1 (N199xxSSR) The term switch 1 is used in analogy with GTP-binding proteins. This compares to a3a in kinesin, however, the ‘switch 1 region’ should include a3, L9, and a3a. This is one of the regions where the structural variations of kinesins are most pronounced (Sack et al., 1999, Song et al., 2001). As shown by Figs. 12.3 and 12.5, the lengths of a3 and a3a differ significantly in different kinesin structures. It is remarkable that a lengthening of a3 is accompanied by a shortening of a3a and vice versa, e. g. the helix a3 in NcKin is 23 residues long compared to 16 residues in RnKin resulting in a 35-Å-long helix. The position of L9 critically determines the size of the nucleotide-binding pocket (Song et al., 2001). In the case of Kar3 and Ncd, this loop is bent inwards to such an extent that the size of the active site is significantly reduced. 12.2.2.2

291

292


N3 Switch 2 (D232LAGSEKVGKT) The switch 2 region contains the conserved LAGSE-motif belonging to the loop 11, located closely to both the nucleotide and the microtubule-binding region defined by helix a4, loop 12, helix a5, and loop 13 (sometimes called ‘switch 2 cluster’). This is also the region which undergoes conformational changes during the ATPase cycle. In all the structures besides the structure of fast fungal kinesin (NcKin) and the R598A mutant of Kar3 (Yun et al., 2001) loop 11 is invisible. Comparing the structures of the helix a4 in Kif1A and NcKin and others, it is likely that helix a4 alters its length during the ATPase cycle. 12.2.2.3

N4, (R14xRP) The motif N4 is responsible for the coordination of the base moiety. The proline forms a stacking interaction with the adenosine ring, and the aliphatic side chain of arginine makes a hydrophobic interaction. 12.2.2.4

12.2.3

Neck Linker, Neck and Hinge

This region is located at the end of a6, residue Z 325 for RnKin (Z 329 for NcKin) and was largely disordered in most of the other known kinesin structures. However, RnKin exhibits an extended neck linker (b9, b10, residues Z 325 339) and an a-helical neck (339 370) which leads into the coiled-coil stalk domain and is responsible for the dimerization of kinesin. This region undergoes large structural transitions during the movement of kinesin and is probably intrinsically mobile (Case et al., 2000, Rice et al., 1999). Data from EPR and FRET measurements suggest that in the ATP state the neck linker is attached and fixed to the body of the motor domain (‘docked’), whereas it is detached and mobile in the ADP state (Rice et al., 1999, Rosenfeld et al., 2001). This has led to a model of structural changes transmitted from the nucleotide-binding site to the neck linker and hence to the rest of the stalk (Vale and Milligan, 2000). In this model, nucleotide dependent upward and downward movement of the switch 2 helix a4 (‘relay helix’) causes the binding or detachment of the conserved Ile (I327 in RnKin) onto the motor core. This leads to ‘docking’ or ‘undocking’ of the neck linker to the core. In the ATP bound state (hypothetically represented by the rat kinesin structure – but see below) loop 12 of the switch 2 cluster allows the root of the neck linker to bind and, thus, docking of the whole neck linker is initiated. In the ADP/nucleotide-free state (likened to the structure of human kinesin) a shift of helix a4 and loop12 prevents binding of the neck linker. The model is partly supported by the structures of Kif1A which show more order in the presence of AMPPCP (an ATP analog) than with ADP (Kikkawa et al., 2001). On the other hand, the model is at variance with the structure of RnKin where the neck linker and neck are docked and highly ordered in the ADP state (Sack et al., 1997), suggesting that alternative models of conformational relays should also be considered. Pro-


gress in this area is hampered by the lack of structures in different states of nucleotide binding and switching. Interestingly, the neck linker contributes to the stability of the neck helix by formation of two helix capping motifs (Tripet and Hodges, 2002). Thus, it seems that structural effects induced by nucleotide processing can now be traced from the active site via switch 2, switch 2 cluster (a4, L12, a5), and neck linker down to the neck helices. That the neck coiled-coil is more than a passive element that binds the motor to the rest of the molecule has been demonstrated by a kinesin mutant with additional positive charges in the neck region that displays ultra-processive movement (Thorn et al., 2000). Similarily, cleavage of the negatively charged C-terminus of tubulin alters the electrostatic interaction with kinesin and results in dramatically decreased run lengths (Wang and Sheetz, 2000). The coiled-coil stalk of kinesin is interrupted by several non-a-helical regions which could operate as hinges, allowing the stalk to kink and swivel (Seeberger et al., 2000, Thormahlen et al., 1998). The first hinge is predicted to comprise residues Z 370 410 (Grummt et al., 1998, Tripet et al., 1997), and indeed the neck helix terminates at W370 in the crystal structure. The NMR structure of the peptide K357 D386, containing the second half of the neck helix and the first half of the hinge, corroborates this picture (Fig. 12.1c). It shows that the neck helix is indeed a-helical in both structures, indicating that the helix propensity is high enough to stabilize the neck even in the absence of the core motor domain. Beyond the neck helix (Z residue 370), disorder and flexibility sets in, as might be expected of a true hinge (Seeberger et al., 2000).

12.3

Comparison with G-Proteins and Myosin

As Table 12.2 shows the nucleotide of kinesin is embedded in four structural elements that share remarkable structural homologies with GTPases and myosin (Sack et al., 1999, Vale, 1996). It is therefore tempting to speculate that they may also share the same reaction mechanism for the hydrolysis of the nucleotide triphospates. It is known that the detailed sub-steps, such as the overall rate-limiting step or the conformation with highest affinity to the interaction partners (tubulin, actin, or GAP), are different. For example, there is a principal difference in the nucleotide-dependent affinity of myosin or kinesin to its filaments, i. e. myosin complexed with ATP only has a low affinity to F-actin while kinesin in complex with ATP binds strongly to microtubules. Fortunately in the case of myosins (D. discoideum myosin II and smooth muscle myosin II) and G-proteins (e. g. P21ras, EF-Tu) the structures are solved in complexes with different nucleotides and nucleotide analogs (Geeves and Holmes, 1999, Sprang, 1997). These studies have revealed that the movements of switch 1 and 2 motifs mediate the events of structural modifications along the path of nucleotide hydrolysis. These movements are also accompanied by the reversible formation of critical salt bridges (best documented in the case of myosin). Both G-proteins and myosin have a ‘closed’ conformation

293

294

12.4 Mechanochemical Coupling from a Structural Point of View

of the nucleotide-binding pocket when they are in complex with nucleotide triphosphate analogs. An exception is myosin complexed with MgADP-BeF3 which can occur in both open and closed conformations (Fisher et al., 1995, Geeves and Holmes, 1999). In the ‘closed’ conformation the amide nitrogen of the conserved glycine in switch 2 (corresponding to G235 in rat brain kinesin) forms a hydrogen bond with an oxygen of the g-phosphate. In the ‘open’ conformation of the active site with bound ADP or GDP, the peptide bond of the conserved glycine flips away from the pocket. In the case of G-proteins, e. g. P21ras, an enormous movement of 15 Å of the switch 1 region follows this switching mechanism of the gphosphate sensing region during nucleotide hydrolysis. It has been shown that the disruption of Mg2 coordination is the key step for dissociation of the nucleotide GDP in P21ras. This takes place via the GEF-mediated displacement of the switch 1 region of about 15 Å (Boriack-Sjodin et al., 1998) Such a movement can be observed in NcKin in comparison with switch 1 of other kinesins (Fig. 12.6).

12.4

Mechanochemical Coupling from a Structural Point of View

Kinesin is a molecular motor which transduces chemical energy into movement by hydrolysis of ATP. There are three fundamental states for the enzyme: ATP-bound, ADP-bound and nucleotide-free active sites. The first two states exhibit only small structural differences between the presence and the absence of g-phosphate at the active site, but this small difference triggers a cascade of structural alterations. These alterations propagate from the P-loop to the switch 1 and switch 2 regions until they end in a distant region of the enzyme which then induces the dramatic movement of domains comparable to that seen in EF-Tu or in the swing of the lever arm in myosin (Hilgenfeld, 1995, Houdusse et al., 2000). During the path of these conformational changes, the enzyme modulates its affinity to the protein partners, e. g. microtubules in the case of kinesin, F-actin for myosin, and the cofactors GEF or GAP for the G-proteins (Hilgenfeld, 1995, Holmes and Geeves, 2000, Scheffzek et al., 1998). The case of kinesin is even more complex because it is a highly processive motor and can take many steps along a microtubule before it dissociates, implying that the interaction between the two motor subunits matters as well. This issue is discussed elsewhere (see J. Howard). To understand the relay of chemical into mechanical energy, a key question is how the conformational changes take place during the reaction cycle. In the case of myosin it has been possible to study the structures of the motor domain ( myosin subfragment S1) complexed with different nucleotides, including ADP, and different ATP-analogs (Bauer et al., 2000, Furch et al., 1999, Gulick et al., 2000). The comparison between these structures has revealed that the structure of the active site comprises one of the extreme conformational states, either the OPEN or the CLOSED form. These terms refer to the width of the nucleotide binding pocket, as measured by the distance between the first glycine in the P-loop (corresponding to G86 in RnKin) and the conserved glycine in the switch 2 region


(corresponding to G235 in RnKin). It is thought that the closed form corresponds to the tight binding of ATP. The conformational changes in myosin are limited to parts of switch 2 and the switch 2 helix and regions of the two reactive thiols, i. e. the switch 2 helix bends during nucleotide hydrolysis, leading to a change in relative orientations of structure elements around the end of the switch 2 helix. The other changes of the rest of the motor domain take place by rigid-body rotations of secondary and tertiary structure elements (Holmes and Geeves, 2000). In kinesin, translation and amplification of the small local changes at the nucleotide sensing switch regions into large scale effects seems to rely on a similar mechanism involving the switch 2 helix (a4). However, at the end it is not the rotation of a rigid lever as in myosin that takes effect, but the modulation of surface properties which control zippering of the flexible neck linker onto the core domain (Kull and Endow, 2002, Vale and Milligan, 2000). In order to understand the mechanism of kinesin ATPase, it would be helpful to know how different types of kinesin react in different stages of the ATPase cycle. Unfortunately, for kinesin all structures so far determined are complexed with ADP except the monomeric processive motor protein, Kif1A, which is complexed with the non-hydrolyzable ATP-analog AMPPCP (Kikkawa et al., 2001). However, functionally important structural elements may also be identified by analyzing variances between kinesin constructs from different sources. Correlation of structural variances with differences in activity (e. g. directionality, processivity, velocity) is a promising tool to gain further insight into the basic mechanisms. Similarily, comparing native and mutant structures of the Kar3 motor domain (Yun et al., 2001) has been very informative. It has shown that key structural elements like the switch 2 helix a4 and the switch 1 region a3-loop9-a3a are highly variable, and pointed to a path of interactions linking the nucleotide to the microtubule binding site by a series of salt bridges. Interruption of this link leads to uncoupling of ATP hydrolysis from microtubule binding. Kinetic and biochemical measurements on the salt bridge mutant R210A of a dimeric motor construct from Drosophila kinesin (corresponding to the R598A mutant of Kar3), further demonstrated that this link is essential for proper coordination of the two heads, ensuring high processivity with tight coupling of ATP hydrolysis and motor stepping (Farrell et al., 2002). In summary, comparison of the kinesin structures from different sources has revealed several interesting features that might give hints about the structural modifications which occur during nucleotide hydrolysis: (1) Each kinesin structure that has been determined so far shows a unique structure for the switching region. (2) The length of the switch 1 helix, a3 is different and the helix a3a shows different secondary structures: a-helix or b-strand. (3) The ‘relay’ helix, a4, is variable in length and can be tilted around 20h between different structures. (4) Loop 11 in the switch 2 region is mostly disordered except for NcKin, but the length of the disordered loop differs (Fig. 12.2). (5) There are different sets of salt bridges present in different structures. (6) The ‘neck linker’ region is mostly disordered but it is ‘docked’ close to the core of the molecule in RnKin-ADP (Sack et al., 1997), semidocked in NcKin-ADP (Song et al., 2001) and possibly docked in Kif1A-AMPPCP (Kikkawa et al., 2001).

295

296


Figure 12.2. Structural comparisons of kinesin kinesin overlap remarkably well with the corre-

and myosin. Ribbon diagrams of the NcKin355 (pdb: 1GOJ; Song et al., 2001) and Dictyostelium discoideum myosin motor domain (pdb: 1MMA; Gulick et al., 1997) showing the overall fold (both in the presence of ADP). Similar structural elements are shown in color. Kinesin and myosin were superimposed by aligning the residues of the P-loop (NcKin 88–94, myosin 178–184). Seven strands of the eight-stranded b-sheet (blue) and six surrounding a-helices of

sponding structural elements of myosin. The upper panel shows a ‘front’ view of kinesin where the microtubule-binding surface is in the background. The lower panel shows the ‘rear’ view into the microtubule-binding elements (green), as seen from the microtubule surface. Note that in the structure of NcKin the neck linker is semi-detached from the body of the motor domain (‘semi-docked’), and the neck does not fold as an a-helix (compare Fig. 12.1a).

The various kinesin structures suggest that the switching mechanism has additional sub-steps, compared to myosin, where the structures have been determined in different nucleotide states and where the switching region undergoes only some tilting and rotating movement. The salt bridges observed in kinesins and myosins suggest that there are conserved charged residues in the g-phosphate-sensing switch region. Different sets of salt bridges can be observed in different structures between these conserved charged residues (Song et al., 2001). The switches are intrinsically mobile and therefore it is possible that any of the salt bridges observed so far may form at a certain stage during the path of the ATPase. Nonetheless, we may be able to observe only a restricted set of salt bridges in a given structure.

12 Structures of Kinesin Motor Domains α3

Switch 1

L9

α3a

β6

L10

β7

Kif1a(AMPPCP) Kif1a(ADP) NcK355 RnK354 HsK349 Eg5(mol 2) Eg5(mol 1) Kar3 Ncd

188SYNDIQDLMDSGNKARTVAATN----MNETSSRSHAVFNIIFTQKRHDAETNITTEKVSKISLVD 188SYNDIQDLMDSGNKARTVAATN----MNETSSRSHAVFNIIFTQKRHDAETNITTEKVSKISLVD 179SVQEVYEVMRRGGNARAVAATN----MNQESSRSHSIFVITITQKNVETG----SAKSGQLFLVD 176SPEEVMDVIDEGKANRH----VAVTNMNEHSSRSHSIFLINIKQENVETEK----KLSGKLYLVD 175SPDEVMDTIDEGKSN----RHVAVTNMNEHSSRSHSIFLINVKQENTQTEQ----KLSGKLYLVD 209NKDEVYQILEKGAAKRTTAATLM----NAYSSRSHSVFSVTIHMKETTIDGEE-LVKIGKLNLVD 209NKDEVYQILEKGAAKRTTAAT----LMNAYSSRSHSVFSVTIHMKETTIDGEE-LVKIGKLNLVD 269SEEMVEIILKKANKL----RSTASTASNEHSSRSHSIFIIHLSGSNAKTG----AHSYGTLNLVD 524DPNHLRHLMHTAKMN----RATASTAGNERSSRSHAVTKLELIGRHAEKQE----ISVGSINLVD

Kif1a(AMPPCP) Kif1a(ADP) NcK355 RnK354 HsK349 Eg5(mol 2) Eg5(mol 1) Kar3 Ncd

α5 L11 α4 L12 LAGSERADSTGAKGTRLKEGANINKSLTTLGKVISALAEMDSGPNKNKKKKKTDFIPYRDSVLTWLLRENL LAGSERADSTGAKGTRLKEGANINKSLTTLGKVISALAEMDSGPNKNKKKKKTDFIPYRDSVLTWLLRENL LAGSEKVGKTGASGQTLEEAKKINKSLSALGMVINALTDGKS-----------SHVPYRDSKLTRILQES LAGSEKVSKTGAEGAVLDEAKNINKSLSALGNVISALAEGTK-----------THVPYRDSKMTRILQDSLD LAGSEKVSKTGAEGAVLDEAKNINKSLSALGNVISALAEGS------------TYVPYRDSKMTRILQD LAGSENIGRSGAVDKRAREAGNINQSLLTLGRVITALVERTP------------HVPYRESKLTRILQDSL LAGSENIGRSGAVDKRAREAGNINQSLLTLGRVITALVERTP------------HVPYRESKLTRILQDSL LAGSERINVSQVVGDRLRETQNINKSLSCLGDVIHALGQPDS---------TKRHIPFRNSKLTYLLQYSL LAGSE-SPKT---STRMTETKNINRSLSELTNVILALLQ------------KQDHIPYRNSKLTHLLMPS

Switch 2

Sequence alignment of switch 1 and switch 2 of kinesins according to their structures. The sequences shown start from the switch 1 region (a3-L9-a3a) and end by a5. This includes the switch 2 cluster. Regions of ahelix are in grey boxes, b-sheet bold and un-

Figure 12.3.

derlined, and disordered (invisible) white on black background. Note that the lengths of a3, a3a, and a4 are variable, as well as the disordered part of L11, whereas the surrounding structure elements are well conserved.

From this observation one can deduce that the motor proteins may be in a dynamic conformational state. The role of the track (microtubule or F-actin) is probably to pave the way out of this non-directed dynamic conformational path to proceed in an ordered direction resulting in an ATP-driven motor activity (Song et al., 2001, Vale and Milligan, 2000). Here we venture one step further, positioning different kinesin structures in an (incomplete) cycle of potential structural modifications along the pathway of the ATPase cycle, assuming that variances between different crystal structures are indicative of the structural changes that are induced by ATP hydrolysis (cf. Table 12.3 for the hypothetic limiting states with ATP and ADP). Table 12.3. Regions of kinesin related to switching and their structural modifications. The table shows structural features of Kif1A complexed with ADP (pdb: 1I5S), with AMPPCP (pdb:1I6I) and other known kinesin structures complexed with ADP.

Structural elements

‘ADP-state’

‘ATP-state’

a3

Long

Short

L9

Out

In

a3a

Short

Long

L11

More ordered

Disordered

a4

Long

Short

b9-b10

Undocked

Docked

297

298


Kif1A-AMPPCP (pdb code: 1I6I) and NcKin (pdb code: 1GOJ) are positioned at the corner posts of the conformational events (Fig. 12.4 and 12.5). The structure of Kif1A-AMPPCP does not show the interactions which were recognized to be essential for the hydrolysis of nucleotide triphosphate in G-proteins and myosin (Pai et al., 1990, Smith and Rayment, 1996). After the binding of NTP, hydrolysis of the bond of the bridge oxygen between b- and g-phosphate occurs, resulting in the peptide-flip of the conserved glycine. Such peptide flips have been observed

Figure 12.4. Hypothetical states of kinesin

structures during the ATPase cycle. Kinesin structures are ordered in a circle as if each of them would represent one of the putative intermediates of kinesin structures during the path of the ATPase. Note that all the structures are in the same orientation (viewed from the microtubule surface) and only the switch 1 and 2 regions are colored in red. The switch 1 and 2 regions are the structural elements which undergo dramatic changes during the path of the kinesin ATPase (see text and Fig. 12.5). The sequence is (1) Kif1A-AMPPCP, 1I6I (2) RnKin, 2KIN, (3) Kif1A-ADP, 1I5S, (4) Kar3, 3KAR, (5) Eg5-molecule 1, 1II6, (6) Eg5-molecule 2, 1II6, (7) NcKin, 1GOJ.

Figure 12.5. Switching mechanism. Pairwise

structural alignments of kinesin structures with the structure of Kif1A-AMPPCP (pdb:1I6I). Only the regions of switch 1 and 2 are shown. The description below is always to compare to the AMPPCP state of Kif1A. The structure of Kif1AAMPPCP is in red and the others are in green. The sequence of the structures is the same as in Figure 4. Note that in the reference structure Kif1A-AMPPCP complex, the region of a3a adopts a b-hairpin structure. (a) RnKin: a3a has two turns and a4 is tilted. (b) Kif1A in complex with ADP: the switch 1 helix a3 is elongated by half a turn, but the helix is not continuous, see text. Helix a4 (the relay helix) is lengthened and tilted by about 20 h. (c) Kar3: a4 has the same conformation and length as the Kif1A structure complexed with ADP. (d) Eg5 (molecule 1): a3 is lengthened and continuous and the a3a-region is helical. (e) Eg5 (molecule 2): a4 is even shorter than in Kif1A-AMPPCP, but the orientation is like that of the Kif1A-ADP complex. (f) NcKin: a3 is elongated by three turns and a4 is extended but not continuously by one more turn.


in both G-proteins and myosin (pdb:1VOM versus 1MMD). Since the structure of Kif1A in complex with AMPPCP does not feature those essential interactions for the transition state the structure might be representing a collision complex with ATP (Cooke, 1986, Kull and Endow, 2002). The structure of NcKin is assigned to the other end of the cycle because this structure shows a wide open active site with large displacement of the switch 1 region. The Mg2 cofactor plays an important role in nucleotide exchange and hydrolysis of small G-proteins (Mittal et al., 1996). Crystallographic studies of the Ha Ras Sos complex have revealed that the nucleotide exchange factor GEF dislocates the switch 1 region, thereby disrupting the Mg2 coordination (Boriack-Sjodin et al., 1998). This results in the destabilization of GDP binding and accelerates GDP dissociation. Figure 12.6 shows a similar displacement of the switch 1 region if one compares Kar3 with NcKin. To a lesser extent this is also the case when the structure of NcKin is superimposed on that of HsKin and RnKin. Given that ADP/ ATP exchange is rate limiting in the overall reaction, this may explain why NcKin has a higher ATPase rate constant than other kinesins.

Figure 12.6. Displacement of loop 9 of switch 1 region: Superposition of kinesin structures in the region of loop 9. The orientation of the view is rotated 90h around the vertical axis and 30h around the horizontal axis compared with the view in Fig. 12.1a. (a) NcKin versus RnKin and HsKin (b) NcKin versus Kar3 and Ncd (c) Superposition of NcKin and Kar3 showing the

displacement of loop 9 in the two structures. The distance between residues A197 (NcKin) and A587 (Kar3) is about 15 Å. (d) Superposition of Ras-GTP and Ras-Sos structures. The displacement of switch 1 is also about 15 Å as measured by the movement of residue Y32 (Boriak-Sjodin et al., 1998).

299

300

12.5 Perspectives

The other kinesin structures are positioned between these two reference points according to the appearance of switch 1 and switch 2, i. e. considering the length of switch 1 and the length and angle of switch 2-helix (Figs. 12.4 and 12.5). The order of the structural transitions is arbitrary. However, it is worth noting the following: (1) Switch 1 helix, a3, can alter its length. The region of switch 1 can undergo structural transition from a-helix (a3a) to small anti-parallel b-strands. (2) The switch 2 helix, a4, can simultaneously suffer a tilting and a rotating movement. (3) Loop 11 in the switch 2 region can undergo a structural transition between an ‘ordered’ and a ‘disordered’ conformation.

12.5

Perspectives

In this survey we have summarized the structures of the currently known kinesin motor domains, with particular emphasis on the elements that make up the nucleotide binding site and may be involved in transforming chemical energy into mechanical work. We have left out many other structural features, in particular the interactions between kinesin and microtubules which have been deduced from cryo-electron microscopy of kinesin-microtubule complexes (for a review, see Mandelkow and Hoenger, 1999), and the regulatory features that reside in the kinesin tail and associated light chains (for a review, see Woehlke and Schliwa, 2000). But even within the restricted aspect of switching within the motor domain we are currently limited to educated guesses. The reason is that different kinesins, although similar in core features, can be quite different in certain aspects, and it is not clear whether these differences are meaningful in terms of reaction pathways. We have tried to put some of these differences into an ordered scheme, but this should only be regarded as a basis for discussion. There is only one kinesin structure solved in two nucleotide binding states (Kif1A; Kikkawa et al., 2001), with ADP or with AMP-PCP, but it is not clear to what extent this mimics ATP, and moreover the kinesin has a hybrid sequence. One kinesin structure has been solved with several functionally important mutations (Kar3; Yun et al., 2001). However, since all structures are with bound ADP there is a question of how they are related to the ATPase cycle. Spectroscopic evidence (Rice et al., 1999, Rosenfeld et al., 1996) indicates that the neck linker is docked onto the body of the motor domain in the ATP state but becomes unzippered in the ADP state, and this has been incorporated into a model of switching (Vale and Milligan, 2000). Consistent with this, most kinesin structures (with bound ADP) have an undocked disordered neck linker and neck; however, the one structure with a clearly docked neck linker is also in an ADP-state (Sack et al., 1997), making the docking/undocking paradigm vulnerable. Furthermore, the question of head head interactions, essential for the half-site reactivity of kinesin (Gilbert et al., 1998, Hackney, 1996), cannot be addressed for lack of solved structures, but it is interesting to note that the known dimer structures (rat brain kinesin and Ncd; Kozielski et al., 1997, Sablin et al., 1998) show the two heads in positions where they cannot simultaneously interact


with the microtubule surface and suggest that major conformational rearrangements have to take place upon microtubule binding. In this situation, there is clearly a need for more kinesin structures in different nucleotide states. This has been achieved in the case of several other proteins which also depend on nucleotide-dependent switching and share common structural elements, such as myosin or G-proteins (Kull and Endow, 2002, Vale and Milligan, 2000). These examples offer the perspective that the mechanochemical cycle of kinesin can be solved eventually, provided that we find ways to overcome the resistance of kinesin to crystallization.

Acknowledgements

We thank Eva-Maria Mandelkow, Jens Müller, Susan Gilbert and Jon Kull for many stimulating discussions and critical reading of the manuscript.

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Miki, H., Setou, M., Kaneshiro, K. and Hirokawa, N. 2001. All kinesin superfamily protein, KIF, genes in mouse and human. Proc. Natl Acad. Sci. USA 98: 7004 7011. Mittal, R., Ahmadian, M. R., Goody, R. S. and Wittinghofer, A. 1996. Formation of a transition-state analog of the Ras GTPase reaction by Ras-GDP, tetrafluoroaluminate, and GTPase-activating proteins. Science 273: 115 117. Pai, E. F., Krengel, U., Petsko, G. A., Goody, R. S., Kabsch, W. and Wittinghofer, A. 1990. Refined crystal structure of the triphosphate conformation of H-ras p21 at 1.35 Å resolution: implications for the mechanism of GTP hydrolysis. EMBO J. 9: 2351 2359. Rice, S., Lin, A. W., Safer, D., Hart, C. L., Naber, N., Carragher, B. O., Cain, S. M., Pechatnikova, E., Wilson-Kubalek, E. M., Whittaker, M., Pate, E., Cooke, R., Taylor, E. W., Milligan, R. A. and Vale, R. D. 1999. A structural change in the kinesin motor protein that drives motility. Nature 402: 778 784. Rosenfeld, S. S., Jefferson, G. M., King, P. H. 2001. ATP reorients the neck linker of kinesin in two sequential steps. J. Biol. Chem. 276: 40167 40174. Rosenfeld, S. S., Correia, J. J., Xing, J., Rener, B. and Cheung, H. C. 1996. Structural studies of kinesin-nucleotide intermediates. J. Biol. Chem. 271: 30212 30221. Sablin, E. P., Case, R. B., Dai, S. C., Hart, C. L., Ruby, A., Vale, R. D. and Fletterick, R. J. 1998. Direction determination in the minus-enddirected kinesin motor ncd. Nature 395: 813 816. Sablin, E. P., Kull, F. J., Cooke, R., Vale, R. D. and Fletterick, R. J. 1996. Crystal structure of the motor domain of the kinesin-related motor ncd. Nature 380: 555 559. Sack, S., Kull, F. J. and Mandelkow, E. 1999. Motor proteins of the kinesin family. Structures, variations, and nucleotide binding sites. Eur. J. Biochem. 262: 1 11. Sack, S., Muller, J., Marx, A., Thormahlen, M., Mandelkow, E. M., Brady, S. T. and Mandelkow, E. 1997. X-ray structure of motor and neck domains from rat brain kinesin. Biochemistry 36: 16155 16165. Scheffzek, K., Ahmadian, M. R. and Wittinghofer, A. 1998. GTPase-activating proteins: helping hands to complement an active site. Trends Biochem. Sci. 23: 257 262.

12 Structures of Kinesin Motor Domains Scheidig, A. J., Franken, S. M., Corrie, J. E., Reid, G. P., Wittinghofer, A., Pai, E. F. and Goody, R. S. 1995. X-ray crystal structure analysis of the catalytic domain of the oncogene product p21H-ras complexed with caged GTP and mant dGppNHp. J. Mol. Biol. 253: 132 150. Schulz, G. 1992. Binding of nucleotides by proteins. Curr. Opin. Struct. Biol. 2: 61 67. Seeberger, C., Mandelkow, E. and Meyer, B. 2000. Conformational preferences of a synthetic 30mer peptide from the interface between the neck and stalk regions of kinesin. Biochemistry 39: 12558 12567. Smith, C. A. and Rayment, I. 1996. X-ray structure of the magnesium II .ADP.vanadate complex of the Dictyostelium discoideum myosin motor domain to 1.9 Å resolution. Biochemistry 35: 5404 5417. Song, Y.-H., Marx, A., Muller, J., Woehlke, G., Schliwa, M., Krebs, A., Hoenger, A. and Mandelkow, E. 2001. Structure of a fast kinesin: implications for ATPase mechanism and interactions with microtubules. EMBO J. 20: 6213 6225. Sprang, S. R. 1997. G protein mechanisms: insights from structural analysis. Annu. Rev. Biochem. 66: 639 678. Thormahlen, M., Marx, A., Sack, S. and Mandelkow, E. 1998. The coiled-coil helix in the neck of kinesin. J. Struct. Biol. 122: 30 41. Thorn, K. S., Ubersax, J. A. and Vale, R. D. 2000. Engineering the processive run length of the kinesin motor. J. Cell Biol. 151: 1093 1100. Tripet, B. and Hodges, R. S. 2002. Helix capping interactions stabilize the N-terminus of the kinesin neck coiled-coil. J. Struct. Biol. 137: 220 235.

Tripet, B., Vale, R. D. and Hodges, R. S. 1997. Demonstration of coiled-coil interactions within the kinesin neck region using synthetic peptides. Implications for motor activity. J. Biol. Chem. 272: 8946 8956. Turner, J., Anderson, R., Guo, J., Beraud, C., Fletterick, R. and Sakowicz, R. 2001. Crystal structure of the mitotic spindle kinesin Eg5 reveals a novel conformation of the neck-linker. J. Biol. Chem. 276: 25496 25502. Vale, R. D. 1996. Switches, latches, and amplifiers: common themes of G proteins and molecular motors. J. Cell Biol. 135: 291 302. Vale, R. D. and Milligan, R. A. 2000. The way things move: looking under the hood of molecular motor proteins. Science 288: 88 95. Walker, J. M., Saraste, M., Runswick, M. J. and Gay, N. J. 1982. Distantly related sequences in the alpha- and beta-subunits of ATP synthase, myosin, kinases and other ATP-requiring enzymes and a common nucleotide binding fold. EMBO J. 1: 945 951. Wang, Z. and Sheetz, M. P. 2000. The C-terminus of tubulin increases cytoplasmic dynein and kinesin processivity. Biophys. J. 78: 1955 1964. Woehlke, G. and Schliwa, M. 2000. Walking on two heads: The many talents of kinesin. Nature Reviews Molecular Cell Biology 1: 50 58. Yun, M., Zhang, X., Park, C. G., Park, H. W. and Endow, S. A. 2001. A structural pathway for activation of the kinesin motor ATPase. EMBO J. 20: 2611 2618. Internet addresses on kinesin structure: www.proweb.org/kinesin; www.mpasmb-hamburg.mpg.de/ktdock

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13 Single Molecule Measurements and Molecular Motors Yoshiharu Ishii and Toshio Yanagida

Single molecule detection was developed to measure the unique operation of molecular motors. The function of molecular motors has been ascribed to the movement of a single motor molecule along a partner protein track. Manipulating single protein tracks and single motors and imaging single molecules has allowed the behavior of such unitary machines to be measured. The mechanical responses to the energy input from the hydrolysis of ATP molecules have been determined and the underlying mechanism explored. The results indicate that the movement of molecular motors is driven by thermal motion rather than structural changes occurring in the motor molecules. Thermal Brownian motion must be biased in one direction. In this chapter, we summarize the methods and results of single molecule measurements of molecular motors.

13.1

Introduction

Almost 30 years after the proposal of the sliding filament theory of muscle, the movement of actin filaments over myosin immobilized on coverslips was first visualized under a microscope (Harada et al., 1990, Kron and Spudich, 1986). This was accomplished by visualizing actin filaments labeled with phalloidin tetramethylrhodamine by fluorescence microscopy (Yanagida et al., 1984). Phalloidin, a mushroom toxin, binds stoichiometrically to actin molecules and serves to stabilize the actin molecules in the filament form at low concentrations, at which point the motility assay and the single molecule measurements are performed. The in vitro assay has proven to be a model system in which both biochemical and mechanical properties of the interaction of myosin and actin can be studied using purified proteins. This system contrasts with conventional measurement systems which use single muscle fibers and proteins in solution. The mechanical properties can be measured in a muscle fiber but the system is not defined biochemically. The biochemical properties can be measured in solution, while the mechanical properties cannot be studied.

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13.2 Manipulation of Actin Filaments

The sliding movement of actin and myosin can be basically ascribed to the operation of a unit machine that contains a single myosin molecule and an actin filament. In the in vitro assay, the smooth movement of an actin filament involves interactions with many myosin molecules. Thus, it is not possible to determine which myosin molecules interact with the actin filament and when and where ATP molecules are hydrolyzed. By manipulating single actin filaments and single myosin molecules, it has been possible to study the interaction between myosin and actin in a controlled manner. Examining the single unit machine offers many advantages over ensemble measurements. Myosin and actin function dynamically. Myosin undergoes a series of chemical reactions during ATP hydrolysis. It changes the interaction with actin cyclically in association with these chemical reactions. In addition, all biomolecules are inevitably exposed to thermal perturbations. These processes are stochastic. In ensemble measurements involving large number of molecules, only average values can be determined and the dynamic changes, which are important in relation to function, are hidden and obscured in the averaging process. Single molecule measurements have overcome these difficulties and have allowed the dynamic behavior of protein molecules to be measured (Ishii and Yanagida, 2000).

13.2

Manipulation of Actin Filaments

Actin filaments can be captured and manipulated by a laser trap (Finer et al., 1994) or a glass microneedle (Ishijima et al., 1996; Fig. 13.1). A laser trap is a technique used to manipulate protein molecules using the laser in a non-invasive manner. When a small bead is irradiated by a focused laser, the incident laser beam is deflected on the surface and radiation pressure is exerted. The total radiation pressure exerted on the entire sphere is always directed towards the focal point of the laser, so the beads are attracted to the focal point and follow the focal position of the laser when it is moved (laser trap). The beads also behave as if they are connected via a spring; i. e. the total trap force exerted on the beads is proportional to the distance from the focal point when their position deviates from the focal point (nanometry). Utilizing these properties of the laser trap, an actin filament can be manipulated via two beads attached at either end by a double trap, and the movement generated by myosin can be measured (Finer et al., 1994; Fig. 13.1a). For the mechanical measurements, the actin filament is suspended taut and brought into contact with myosin molecules sparsely fixed on the glass surface. The actin filaments with the beads at their ends move against the trap force in the presence of ATP. Thus, the displacement of the actin filaments generated by myosin can be measured by monitoring the changes in the position of the beads attached to them. The force generated can be estimated if the stiffness of the laser trap is known. The movement generated by single myosin molecules could be detected if the number of myosin molecules on the glass surface was reduced to one. The stiffness of the laser trap is dependent on the power of the laser. For a bead

13 Single Molecule Measurements and Molecular Motors

1 mm in diameter, the trap force is several tens of pN and the stiffness is Z 0.1 pN nm 1 when the power of the laser is several hundred mW. This stiffness is sufficient to measure the movement caused by a single myosin molecule. The displacement generated with a single ATP molecule is approximately 10 nm. Thus, the force will be less than 8 pN because the free energy released from the hydrolysis of a single ATP molecule is Z 80 pN nm 1. Microneedles are another method used to manipulate actin filaments. Thin glass microneedles can be prepared and attached to the ends of the actin filaments (Ishijima et al., 1996; Fig. 13.1b). The premise of the measurement using microneedles is the same as that for the laser trap. When actin filaments move as a result of the interaction with myosin, the microneedle bends and at the same time a restoring force is applied to the microneedle resisting the movement. The strength of the force is proportional to the displacement, with the stiffness of the microneedle as a proportional constant. When the force caused by myosin is balanced with the restoring force applied by the microneedle, the movement ceases. The displacement of the actin filaments can be measured and the force can be estimated when the stiffness of the microneedle is known. The stiffness of a microneedle is dependent on its shape. Long, thin glass microneedles are more flexible. A microneedle 50 mm in length and 0.3 mm in diameter, which is the typical length used for the measurements, gives a stiffness of 1 pN nm 1.

Manipulation of actin filaments. (a) An actin filament is manipulated via two beads attached at each end. (b) A microneedle is attached at one end of an actin filament to manipulate it. For nanometry, the displacement

Figure 13.1.

of the actin filament produced by single myosin molecules can be measured by recording the displacement of the beads or the tip of a microneedle with nanometer accuracy.

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13.3 Nanometry of Actin Filaments

13.3

Nanometry of Actin Filaments

Given that the displacement of the molecular motors is in the order of nanometers, the change in the position of the beads or the tip of the microneedle must be measured with nanometer accuracy. The image of the bead or the tip of the microneedle can be enlarged approximately 1000 times and projected onto a pair of photodiodes. The change in the position of the bead or tip of the microneedle results in a change in the difference of the intensities of each photodiode. Therefore, the difference in signals between multiple photodiodes is measured to estimate the displacement of the bead or microneedle. Thus the displacement can be determined with nanometer accuracy, which is much less than the diffraction limit. The diffraction limit is the minimum distance that two spots can be apart to be resolved as separate in the light microscope. As demonstrated in these measurements, it is possible to trace a moving spot with an accuracy greater than the diffraction limit. In these measurements, the displacement of the actin filaments is determined by measuring the displacement of a marker such as a bead or microneedle. If the connection between the myosin molecules and the beads or the microneedle is not tight and some compliant linkages exist between them, damping of the actual displacement of myosin and thermal fluctuations can occur. In addition, the position of the bead or microneedle is subject to thermal fluctuation. The mean square displacement I x2 i of a bead or microneedle can be related to the stiffness of the laser trap or microneedle, K, by the equation K I x2 i/2 kBT/2. According to this equation, the root mean square distance is 6.43 nm for a stiffness of 0.1 pN nm 1 and 2.03 nm for a stiffness of 1 pN nm 1. To measure the displacement with nanometer accuracy, it is important to make the system as stiff as possible. The beads or microneedle may respond slowly to the fast movements of the protein molecules. This response determines the time resolution. The response time of the system, t, is related to the friction coefficient b and the stiffness K as t b/ K. The friction coefficient depends on the shape and size of the probe. For a long and flexible glass microneedle with a diameter of 1 mm and a length of 1 cm, (stiffness 0.01 pN nm 1), for example, the temporal resolution is only 1 s. To increase the temporal resolution, a short thin glass microneedle is necessary. For a microneedle with a length of 50 mm and a diameter of 0.3 mm (which gives a stiffness of 1 pN nm 1), the temporal resolution can be increased to almost 1 ms. In contrast, the viscous drag on a bead is small because a bead is smaller than a microneedle. For a bead 1 mm in diameter, the temporal resolution is as good as 0.1 ms. In addition to this advantage of time resolution, the laser trap experiments are much easier to carry out than microneedle experiments. The stiffness of the laser trap can be easily changed by changing the laser power. In the microneedle measurements, microneedles must be individually handcrafted and the stiffness of each individual microneedle must be determined.


13.4

Movement of Actin Filaments Caused by Single Myosin Molecules

The actin filament repeats steps and returns to its original position, while myosin repeats the binding to and dissociation from the actin filament (Fig. 13.2). The profile of the displacement trace is dependent on the ATP concentration. At high concentrations of ATP (in the mM range) the actin filaments showed rapid transient movement. As the ATP concentration decreased to the order of mM, the duration of the displacement became longer, while the size of the displacement remained the same. At very low ATP concentrations (in the nM range), individual events caused by a single myosin and ATP molecule can be identified. The duration time of the displacement increased from Z 0.2 s at 1 mM ATP to Z 20 s at 10 nM ATP. The ATP concentration-dependent trace of the displacement has been interpreted as the mechanical step in relation to the chemical reaction of ATP hydrolysis. The ATP binding step is dependent on the ATP concentration. The time for ATP molecules to bind is longer at low ATP concentrations and much shorter at high ATP concentrations. This is thought to be the mechanism by which single molecules ‘learn’ the ATP concentration of the system. The binding of ATP to myosin triggers the dissociation of myosin from the actin filament resulting in the bead returning to its original position. Myosin actually moves the actin filament when the products of ATP hydrolysis, ADP and Pi, are released. The relationship between the mechanical events and the chemical reaction has been confirmed directly by simultaneously measuring the movement of the motor and the turnover of ATP molecules, as outlined below. One important parameter that the mechanical measurements of myosin have provided is the size of the myosin step (step size) caused during the hydrolysis

Time course of the displacement of actin filaments produced by single myosin molecules. Upper trace: the white line represents the raw data and the black line the same

Figure 13.2.

data after filtration with a low-path filter (20 Hz bandwidth). The stiffness was calculated from the variance of bead fluctuations (bottom panel).

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13.5 Visualization of Single Molecules

of a single ATP molecule (Ishii et al., 2002, Spudich, 1994). The position of the bead fluctuated thermally around a mean position, which was dependent on the stiffness of the trap as mentioned above (Molloy et al., 1995). Once myosin was strongly bound to the actin filament, the stiffness of the system increased and the thermal motion of the beads greatly decreased. By measuring the variance of the displacement trace, it is therefore possible to determine when myosin interacts with actin. However, the size of the step could not be determined for individual displacements, because the starting position of the displacements cannot be measured. The displacement relative to the mean position of the bead has a wide distribution, reflecting the thermal fluctuation of the bead. Using this technique only mean displacements can be obtained. The size of the displacement caused by a single myosin molecule upon ATP hydrolysis varies among research groups, the reported values ranging from 5 to 20 nm (Finer et al., 1994, Guilford et al., 1997, Molloy et al., 1995, Tanaka et al., 1998). Many experimental factors can affect the results of the step size. In the measurements, myosin molecules are immobilized on the glass surface. These immobilization procedures may affect the activity of myosin. Direct attachment of the myosin head to the glass surface may alter its activity as suggested by the results of in vitro motility assays (Iwane et al., 1997). To avoid the direct artificial attachment of the head portion of the myosin to the glass surface, the head is placed in a myosin myosin rod co-filament, which can be prepared so as to contain only one or two heads in a single filament 5 8 mm in length. The use of such filaments has an additional advantage. It has been reported that the step size is dependent on the angle between the myosin and the actin filament (Tanaka et al. 1998). The step size is maximal when the two filaments are parallel, but approaches zero when the two filaments are positioned at right angles. In the myosin myosin rod filament, the orientation of the myosin head relative to the actin filament is known. In the situation where the myosin head molecules are placed in a random orientation, the displacement of actin filaments would range between the maximum value at an angle of 0h and the minimum value at an angle of 90 h(Yanagida et al., 2000). The average value would be less than the maximum value.

13.5

Visualization of Single Molecules

Biomolecules including protein molecules and small chemical compounds such as ATP are much smaller than the diffraction limit. It is impossible to visualize them using conventional light microscopy. In order to visualize such small molecules they are labeled with fluorescent probes and then the fluorescence is monitored. In 1995 a single fluorophore attached to a biomolecule in aqueous solution was first visualized after the background noise was greatly reduced by eliminating the noise from different parts of the microscope (Funatsu et al., 1995). Many microscopes have also been developed to decrease the background noise. The basic concept behind these microscopes is that only local regions are illuminated to ex-


cite the molecules of interest. To visualize the behavior or motion of biomolecules at work, real-time imaging in aqueous solution is also a prerequisite. Total internal reflection fluorescence microscopy (TIRFM) is a technique which illuminates only the surface area between the solution and the coverslip, allowing both local illuminating and real-time imaging (Funatsu et al., 1995; Fig. 13.3). When incident laser light is above the critical angle, the light is totally reflected at the interface with the aqueous solution, and an evanescent field is created 100 to 200 nm from the surface of the aqueous solution. Thus, only the fluorescent molecules positioned near the glass surface are excited and emit fluorescence. Large numbers of fluorescent molecules outside the region are not excited and therefore the background noise is greatly reduced. A single fluorophore is imaged as a spot, but the size of the spot appears to be enlarged to several hundreds of nanometers due to diffraction, even though the size of the molecule is in the order of a few nanometers. Because of this, spatial resolution is limited. While the fluorophores are continuously illuminated, single fluorophores are repeatedly excited and emit fluorescence within nanoseconds. A single spot therefore represents the integration of a large number of photons emitted from a single fluorophore. Single fluorophores can suddenly cease to emit fluorescence after a certain amount of exposure to irradiation, because they become irreversibly damaged as a result of the photochemical reaction. The average time for a photobleaching process to occur depends on the power of the laser. When a laser with higher power is used, photobleaching occurs in a short time, although the fluorescence is initially brighter. The single stepwise drop of the fluorescence is a good test to confirm that the fluorescent spot was from a single fluorophore. Fluorescence intensity can also be used to determine whether the fluorescence spots arise from single molecules. Histograms of the fluorescence intensity from single spots show a large peak corresponding to multiples of the minimum intensity of a single molecule.

Total internal reflection fluorescence microscopy. An evanescent field is created within 100 Z 200 nm from the interface between the glass surface and the solution when the laser light strikes at an angle greater than the critical angle. Only molecules in the vicinity of the glass surface are excited and emit fluorescence.

Figure 13.3.

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13.6 Visualization of ATP Turnover and Mechano-chemical Coupling

13.6

Visualization of ATP Turnover and Mechano-chemical Coupling

Fluorescent spots remain in the same position when single biomolecules are immobilized on a glass surface and their motion can be traced only when they move slowly (not greater than tens of mm s 1). Rapidly moving fluorophores cannot be traced and only contribute to the background noise. The binding of fluorescent ATP to the protein molecules can be visualized in single molecule imaging, because the mobility of bound fluorescent ATP is different from that free in solution (Funatsu et al., 1995; Fig. 13.4). ATP can be chemically modified with a fluorescent probe. The fluorescent ATP molecules can be visualized as spots when bound to immobilized myosin. However, fluorescent molecules cannot be observed when they are free in solution due to the rapid Brownian movement. The time span between binding and dissociation has a characteristic distribution, reflecting stochastic behavior. The histogram of the time course can be fitted to an exponential curve and the decay time (dissociation rate) can be compared with ATPase rates measured in solution at saturating concentrations of ATP. This assay allows the measurement of ATP binding at the single molecule level. However, the range of concentrations of fluorescent molecules is limited, because all fluorescent ATP molecules in solution contribute to the background noise even when evanescent illumination is used. Typically fluorescent ATP is used at concentrations between 10 and 20 nM. Of course, the binding affinity must be high enough to observe binding at such low concentrations.

Imaging of the turnover of individual fluorescent ATP molecules produced by the interaction of single myosin molecules in a myosin filament with a single actin filament. (Top) a picture for an actin filament trapped by a laser via two beads attached at both ends and a myosin myosin-rod co-filament. (Second) fluorescence image of myosin and actin filament. The lower panels show sequential images of an association and dissociation event of ATP (ADP) with the same myosin head.

Figure 13.4.


Given that both mechanical and chemical processes can be measured at the single molecule level, the coupling between them can be directly determined by simultaneous measurements (Ishijima et al., 1998). The direct determination of the coupling is possible only when single molecule detection techniques are used, as individual chemical and mechanical events are not averaged. The data of the simultaneous measurements of the biochemical and mechanical events at low time resolution (Z 30 ms) have shown that the chemical reaction and the mechanical events appear to be coupled (Fig. 13.5). When ATP binds, myosin dissociates from actin and myosin generates the step movement upon ADP dissociation. The coupling between the binding of ATP and the dissociation of myosin has been confirmed at higher time resolution (I 10 ms). However, a time difference between the ADP release and the generation of movement became apparent. This difference had a particular distribution at higher time resolution. In half the cases, the generation of the displacement occurred at the same time as the release of ADP, while in the other half the mechanical displacement was delayed by I 0.5 s after the release of ADP.

Direct determination of the coupling between chemical and mechanical events. The ATP turnover monitored by the fluorescence of Cy3-ATP (bottom panel) and the movement of myosin (top panel) were simultaneously measured at the single molecule level. The dissociation of myosin from actin fi-

Figure 13.5.

laments (decrease in the displacement) is coupled with the binding of ATP (increase in fluorescence) and the generation of movement (increase in the displacement) is coupled with the dissociation of the products of the ATP hydrolysis reaction (decrease in fluorescence).

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13.7 Visualization of the Movement of Single Kinesin Motors

It should be noted that it is not possible to experimentally distinguish photobleaching from the dissociation of fluorescent ADP in a single event. In both cases, the fluorescent spots disappear suddenly. The laser power can be decreased to decrease the rate of photobleaching and the probability that the photobleaching occurs before the dissociation takes place becomes lower. In these experiments the turnover of fluorescent ATP by myosin and the decay time for the photobleaching of Cy3-ATP either directly attached to the glass or bound to myosin in the presence of vanadate, can be compared with the turnover time. Thus, the probability that photobleaching occurs prior to nucleotide release can be reduced to a few percent. The free energy released at the step where the products of ATP hydrolysis are dissociated, must be stored until the generation of the displacement. The storage of the energy, or a prolonged memory effect, can be explained with the existence of two or more different conformational states, between which the protein undertakes slow transitions. Evidence for these multiple conformational states and spontaneous slow transitions between them has come from the studies of single molecule spectroscopy for many proteins including myosin (Wazawa et al., 2000). Protein molecules have several local minimal states with high-energy barriers between them.

13.7

Visualization of the Movement of Single Kinesin Motors

The visualization of single molecules also allows the observation of the movement of single fluorescently-labeled motor molecules along a protein track. For visualization, the motor molecules must move for long distances without dissociating. Kinesin, a microtubule-based motor, is one such processive motor. The movement of single molecules of fluorescently-labeled kinesin along microtubules has been visualized (Vale et al., 1996). A single kinesin molecule moves for Z 500 nm in 1 Z 2 s. The velocity is similar to that of microtubules moving over kinesin molecules attached to a coverslip. The processive movement of kinesin has been explained on the basis of its double-headed structure; one of the two heads moves one step while the other head stays attached to the microtubule. The two heads operate cooperatively and the steps alternate. If this is the case, then single-headed kinesin should not be able to move for long distances. However, single molecule imaging showed that some of the single-headed kinesin molecules did in fact move along the microtubules for long distances, indicating that two heads are not essential for processive movement (Fig. 13.6). The data analysis of the time course of the displacement has shown that the movement of single-headed kinesin resulted from diffusion and was biased in one direction (Inoue et al., 2001, Okada and Hirokawa, 1999). For these single-headed kinesin molecules it has been suggested that additional interactions with microtubules occur so that the motor protein molecules are kept on the track of microtubules during movement. KIF1A, a single-headed motor of the kinesin superfamily, has a lysine-rich positively-charged loop (the K loop)


Movement of single kinesin molecules along microtubules. The trace of the fluorescently-labeled kinesin was recorded and the displacement along microtubules is plotted as a function of time. K351 is a kinesin construct which lacks the C-terminal coiled-coil re-

Figure 13.6.

gion (single headed kinesin) and biotin-dependent transcarboxylase (BDTC) is fused to K351 (K351BDTC). K411 is a double headed kinesin which shows processive movement over a long distance in one direction.

that is thought to interact with negative charges on the C-terminus of microtubules. Deletion mutants of single-headed kinesin also showed diffusion-driven movement and only traveled small distances of 50 nm in less than 1 s. The travel distance and interaction time were enhanced to 500 nm and 2 s, respectively, when they were fused with a peptide called biotin-dependent transcarboxylase (BDTC) which interacts with microtubules. BDTC does not affect the movement of twoheaded kinesin. In the images of the movement of single kinesin molecules, we have frequently observed more than two kinesin molecules moving simultaneously along a single microtubule. The images have shown that there is a tendency for several kinesin molecules to move together suggesting that there is some communication between kinesin molecules via the microtubules (Muto, 2001). Microtubules may be ‘activated’ by the binding of a single kinesin molecule to assume a high affinity state, resulting in cooperative binding of kinesins.

315

316

13.7 Visualization of the Movement of Single Kinesin Motors

However, although the movement of single molecules has been visualized, it has not been possible to observe the unitary movement of kinesin due to limitations in the accuracy of single molecule imaging techniques. The experimental accuracy of imaging is restricted to several hundred nm due to the diffraction limits and the video rate of 1/30 of a second. These limitations have been overcome using a laser trap system where it has been possible to observe the stepwise movement of kinesin. The use of fluorescent beads instead of single fluorophores enhanced the contrast of the images. The laser trap ‘cooled’ the thermal motion of the beads and the use of a pair of photodiodes allowed the measurement of the displacement to be undertaken with nanometer accuracy (Svoboda et al., 1993). Kinesin attached to a bead was brought into contact with microtubules in the presence of ATP. The number of kinesin molecules attached to a bead was estimated statistically. The distribution of kinesin molecules on beads was consistent with Poisson statistics as indicated by the probability of a moving bead at various molar ratios of kinesin to beads. The average number of kinesin molecules on a bead used for the experiments and the probability that more than two kinesin molecules attached to a bead can be estimated. Kinesin moves along microtubules in regular steps of 8 nm (Fig. 13.7). The steps are primarily in the forward direction and only occasionally in the backward direction. As kinesin molecules moved against an increased load produced by the laser, the average time of a step as well as the number of the backward steps increased. Eventually, at a stall force of 7 8 pN the number of steps in the forward and backward directions became equal. When this occurred the molecule dissociated from the microtubules and

x laser

bead kinesin microtubules

Stepwise movement of kinesin attached to beads. The displacement of kinesin can be monitored by the laser trap method. The displacement trace shows 8-nm steps. Backward movement was observed at high load.

Figure 13.7.


returned to its original position. Both forward and backward movements occurred at the same chemical state; the single cycle of the ATP hydrolysis was coupled to either forward or backward steps and the direction could be determined stochastically. Thus, the thermal movements, which have been clearly observed for singleheaded kinesin, are the driving force even for the stepwise movement of kinesin. Substeps of the 8-nm step of kinesin have been uncovered using laser trap measurements with small beads (Nishiyama et al., 2001). Decreasing the size of the beads improved the time resolution, although the images of the bead on the detector became fainter. Dark field illumination was used to brighten the images. The 8nm step could be resolved into fast and slow substeps, each corresponding to a displacement of Z 4nm.

13.8

Visualization of the Processive Movement of Single Myosin Motors

In the case of muscle myosin, it was impossible to detect the sliding movement with the imaging techniques described above, because muscle myosin readily dissociates from actin filaments at every cycle of the ATPase reaction. Recently, different types of myosin behavior have been discovered. Myosin V, VI and IX were found to move processively and it has been possible to visualize the movement of single myosin molecules when they are fused to green fluorescent protein (GFP; Fig. 13.8). Myosin VI, for example, moves 240 nm along a single actin filament in 0.44 s (Nishikawa et al., 2002). Interestingly, the directionality of myosin VI is opposite to the direction of other myosins (Wells et al., 1999). One of the ends

Processive movement of myosin V and myosin VI. The movement of myosins V and VI was visualized by means of green fluorescent protein (GFP). Sequential images indicate the movement of these myosin molecules. Myosin V and myosin VI move in opposite directions. The pointed end of the actin filament was fluorescently labeled.

Figure 13.8.

317

318

13.8 Visualization of the Processive Movement of Single Myosin Motors

of the actin filament (pointed end) was fluorescently marked and the direction of the movement of myosin could be confirmed for individual myosin molecules. Given that myosin V and VI are processive motors, displacement records were obtained by trapping these myosins like kinesin, as well as by double trapping the actin filaments. A consecutive step movement was observed which was consistent with other processive motors. The advantage of processive myosins is that it is possible to determine the size of individual steps without disturbance from thermal fluctuations, with the exception of the first step. Measuring the step size of myosin V and VI allowed the testing of a model developed on the basis of structural studies of myosin heads. In this so-called lever arm model, a long a-helix (the neck domain) connecting the motor domain and the tail domain is thought to rotate relative to the motor domain, thus acting as a lever (Rayment et al., 1993). This model predicts that the size of the displacement would be proportional to the length of the neck domain, or lever arm. Myosin V has a long neck domain, which contains six repeats of the calmodulin or light chain binding site as compared to the two binding sites in muscle myosin. The step size of myosin V was shown to be 36 nm, while that of muscle myosin is 5 Z 20 nm, a finding in agreement with the lever-arm model (Mehta et al., 1999). However, a deletion mutant, in which the neck domain is shortened to only one calmodulin-binding site, performed the same large step as the original myosin V motor (Tanaka et al., 2001; Fig. 13.9). A similar result has been obtained using myosin VI. Myosin VI in its natural form has only one calmodulin-binding site and also produces a large step (Nishikawa et al., 2002, Rock et al., 2001). These observations of large steps in myosins with a short lever arm cannot be explained by the lever-arm model. A step size of 36 nm matches the helical pitch of the two-stranded actin filament,

The step movement of myosin V and its deletion mutant. The displacement was measured by a double trapping method. (Top) wild type myosin V. (Bottom) the mutant was obtained by deleting a large part of the neck domain.

Figure 13.9.


indicating that the architecture of actin filaments plays an important role in the movement of myosin. Thus, the relationship between structure and function can be studied by combining single molecule detection with cell biology and protein engineering. Even if the structural change, i. e. swinging of the lever arm, occurs as predicted based on the X-ray crystallographic studies, these studies indicate that the structural change does not drive the mechanical movement. The structural change of the neck domain may play another role, for example in regulating the kinetic steps as a strain sensor.

13.9

Manipulation of Single Myosin Molecules with a Scanning Probe and Nanometry

In order to measure the details of the displacement of motors, we have developed a new technique in which single muscle myosin molecules instead of actin filaments are captured and manipulated (Kitamura et al., 1999; Fig. 13.10). A single myosin head fragment, S1, was attached to a scanning probe and the displacement of the probe along immobilized actin filaments was measured. The probe was brought into contact with actin bundles. The use of actin bundles instead of filaments makes the manipulation for the interaction easier and the system stiffer. The stiffness of this system was 1 pN nm 1 compared to a stiffness of 0.2 pN nm 1 for the laser trap system, given the root mean square displacement of 2.0 nm for the scanning probe system as compared to 4.5 nm for the laser trap system. The response time was I 0.4 ms. This improvement was critical for detecting the details of the movement of single myosin molecules. To capture single myosin S1 molecules, they are marked by attaching a fluorescent dye to them. The fluorescently-labeled S1 molecules were sparsely placed on a glass slide and scanned using a probe. As a scanning probe, a whisker made from ZnO and 10 nm in diameter with sharp edges, was attached to the tip of a thin glass needle. A single myosin S1 molecule was caught by the tip of the scanning

Figure 13.10. Measurement of 5.5-nm steps in the displacement generated by myosin S1. A single myosin S1 molecule was attached to the tip of a cantilever and brought into contact with actin filaments. Given that this system is stiffer than the laser trap system, steps in the rising phase of the displacement record are identified. Backward steps have been occasionally observed, as indicated by an arrow.

319

320

13.10 Biased Brownian Movement

probe. Confirmation that only a single S1 molecule had been captured, was achieved by monitoring the fluorescence intensity and the photobleaching behavior. The possibility that non-fluorescent myosin molecules and molecules which were labeled but had already been photobleached, were attached in addition to a single fluorescent myosin molecule cannot be ruled out. However, the probability that additional non-fluorescent molecules were attached was estimated to be marginal based on the labeling ratio of myosin after all experimental precautions had been taken. Myosin S1 was attached tightly to a probe via the biotin avidin system. Given that this molecular glue was attached to the light chain of S1, which is a considerable distance from both the catalytic domain and the binding domain for actin on the myosin head, the attachment to the probe is thought not to interfere with motor functions. The displacement record of muscle myosin measured with this system was indistinguishable from the trace of actin filaments obtained from double trap measurements, if the data were plotted on the same time scale (Fig. 13.10). The rising phase of the displacement could be expanded, because the time resolution was considerably better for the scanning probe method. Within a single displacement during the hydrolysis of a single ATP molecule, there was stepwise movement. In a single displacement, there were one to five regular steps, giving rise to a total displacement of Z 5 to 30 nm. The average number of steps in a single displacement was three, corresponding to a total displacement of 13 nm. The size of a single step was 5.5 nm, coinciding with the interval between adjacent actin monomers on one strand of an actin filament.

13.10

Biased Brownian Movement

In addition to the variation in the total number of steps in a single displacement, evidence has been obtained to suggest that the rising phase of the displacement is stochastic. The interval time between steps also varied and was independent of the ATP concentration, whereas the time between the displacements was ATP concentration-dependent. The dwell time between steps was dependent on the temperature so that the 5.5-nm steps could be distinguished more clearly at lower temperatures. Approximately 10 % of the total number of steps was in the backward direction. Backward movements increased with an increase in the load. In the experimental system used, the load applied could be increased by increasing the stiffness of the microneedle. At high loads, the number of steps in a single displacement was smaller and the interval time between the steps was larger, while the size of the steps remained the same. Thus, the reciprocal of the interval time between the steps decreased with an increase in the load. This relationship is basically similar to the force velocity relationship in muscle, indicating that the unit machine consisting of a single myosin molecule and a single actin filament is sufficient to explain the mechanical properties of muscle.


The available data indicate that the sliding movement of myosin is driven by thermal movement. However, thermal Brownian motion is random. The direction of the movement must be biased and the energy released from ATP hydrolysis is used to bias it. Otherwise, the second law of thermodynamics would be violated. In both myosin and kinesin, the directionality of the movement can be explained by a difference in the activation energy between forward and backward movement. The probability of movement is proportional to exp(– e/kBT) when e is the activation energy, kB is Boltzmann’s constant, and T is the absolute temperature. The ratio of the probability for forward and backward movement r nforward/nbackward can be written by the difference in the activation energy between forward and backward movement, as r exp(– De/kBT), where De eforward – ebackward. Analysis of the data shows that the difference in the activation energy was 2 Z 3 kBT for myosin and 5 kBT for kinesin at zero load. A model simulation further shows that the directionality ratio can reach 9 : 1 when the potential difference between neighboring actin monomers is only 10 % of the energy barrier between neighboring actin monomers. The question arises as to how this direction-dependent potential profile is created. One interesting idea is that the binding of a myosin molecule may activate several neighboring actin monomers in a direction-dependent manner, so that the myosin molecule is attracted more to one direction than to the other. In the case of the kinesin microtubule system, such cooperativity in the binding was demonstrated based on the imaging of the binding of single kinesin molecules along microtubules, as described above. The binding of kinesin molecules was induced to a greater extent in front of the moving kinesin molecule. In this model, a statistically biased potential is assumed to cause the biased motion. Another explanation for biased Brownian movement is based on dynamic fluctuations of the potential. Myosin and actin dynamically change their structure and the interactions that occur between them. It has been shown that the movement of Brownian particles under the asymmetrical potential is biased by fluctuating potential unless the fluctuation is random. There is an optimum frequency at which the movement is biased. In the model where a structural change results in movement, different structures of myosin are assumed to correspond to the different states of the ATPase cycle and the structural change is associated with the transition between these states. In contrast, in the biased Brownian model, the movement is thought to occur in one state (ADP.Pi state) in the cycle of ATP hydrolysis. Thus, it is possible to simulate the movement of myosin.

13.11

Concluding Remarks

New findings in the field of biosciences have been frequently initiated by the development of new techniques. Single molecule detection has allowed us to monitor the dynamic behavior of biomolecules, which is hidden in the average values of en-

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References

semble measurements where it is not possible to measure the sequence of the mechanical and biochemical events. Single molecule detection techniques have completely changed the style of research in this field. Combining this technique with the techniques of protein engineering and cell biology, the possibility for new types of measurements has increased. However, many unanswered questions still remain and additional new questions have been raised. There is still room for further improvement of established techniques and the development of new ones. For example, fluorescent dyes with greater stablity would be very useful. In addition, experimental systems that have increased stiffness would increase the experimental resolution for nanometry. Molecular motors are typical molecular machines. Although protein molecules of nanometer size are exposed to thermal agitation, they function very efficiently. There is increasing evidence to suggest that these molecules do not operate against thermal perturbation, but rather harness thermal energy to perform their functions. We have started measuring the behavior of molecular machines in which the biomolecules assemble and interact. Single molecule detection will be the technique of choice to unveil the underlying mechanisms.

References Finer J. T., R. M. Simmons, and J. A. Spudich. 1994. Single myosin molecule mechanics: Piconewton forces and nanometre steps. Nature 368: 113 119. Funatsu T., Y. Harada, M.Tokunaga, K. Saito, and T. Yanagida. 1995. Imaging of Single Fluorescent Molecules and individual ATP turnovers by single myosin molecules in aqueous solution. Nature 374: 555 559. Guilford, W. H., D. E. Dupuis, G. Kennedy, J. B. Patlack, and D. M. Warshaw. 1997. Smooth muscle and skeletal muscle myosin produce similar unitary forces and displacement. Biophys. J. 72: 1006 21. Harada Y., K. Sakurada, T. Aoki, D. D. Thomas, and T. Yanagida. 1990. Mechanochemical coupling in actomyosin energy transduction studied by in vitro movement assay J. Mol. Biol. 216: 49 68. Inoue Y., A. H. Iwane, T. Miyai, E. Muto, and T. Yanagida. 2001. Motility of single one-headed kinesin molecules along microtubules. Biophys. J. 81: 2838 2850. Ishii Y. and T. Yanagida. 2000. Single molecule detection in life science Single Mol. 1: 5 16. Ishii, Y., S. Esaki, and T. Yanagida. 2002. Experimental studies of the myosin actin motor. Appl. Phys. A 75: 325 330.

Ishijima, A., H. Kojima, H. Higuchi, Y. Harada, T. Funatsu, and T. Yanagida. 1996. Multiple and single-molecule analysis of the actomyosin motor by nanometer-piconewton manipulation with a microneedle: Unitary steps and forces. Biophys. J. 70: 383 400. Ishijima A., H. Kojima, T. Funatsu, M. Tokunaga, H. Higuchi, and T. Yanagida. 1998. Simultaneous measurement of chemical and mechanical reaction. Cell 92: 161 171. Iwane,A. H., K. Kitamura, M. Tokunaga, and T. Yanagida. 1997. Myosin subfragment-1 is fully equipped with factors essential for motor function. Biochem. Biophys. Res. Commun. 230: 76 80. Kitamura K., M. Tokunaga, A. H. Iwane, and T. Yanagida. 1999. A single myosin head moves along an actin filament with regular steps of 5.3 nanometres. Nature 397: 129 134. Kron, S. J. and J. A. Spudich. 1986. Fluorescent actin filaments move on mysin fixed to a glass surface. Proc. Natl. Acad. Sci. USA 83: 6272 6276. Mehta, M., R. S. Rock, A. D. M. Rief, J. A. Spudich, R. S. Mooseker, and R. E. Cheney. 1999. Myosin-V is a processive actin-based motor. Nature 400: 590 593. Molloy J. E., J. E. Burns, J. Kendrick-Jones, R. T. Tregear, and D. S. C. White. 1995. Movement

13 Single Molecule Measurements and Molecular Motors and force produced by a single myosin head. Nature 378: 209 212. Muto, E. 2001. Is microtubule an active participant in the mechanism of motility? Biophys. J. 80: 513a. Nishikawa S., K. Homma, Y. Komori, M. Iwaki, T. Wazawa, A. H. Iwane, J. Saito, R. Ikebe, E. Katayama, T. Yanagida, and M. Ikebe. 2002. Class VI myosin moves processively along actin filaments backward with large steps. Biochem. Biophys. Res. Commun. 290: 311 317. Nishiyama, M., E. Muto, Y. Inoue, T. Yanagida, and H. Higuchi. 2001. Substeps within the 8nm step of the ATPase cycle of single kenisin molecules. Nature Cell Biol. 3: 425 428. Okada Y. and N. Hirokawa. 1999. A processive single-headed motor: Kinesin superfamily protein KIF1A. Science 283: 1152 1157. Rayment I., W. R. Rypniewski, K. SchmidsBase, R. Smith, D. R. Tomchick, M. M. Benning, Winkelmann, D., A. G. Wesenberg, and H. M. Holden. 1993. Three-dimensional structure of myosin subfragment-1: a molecular motor. Science 261: 50 58. Rock R. S., S. E. Rice, A. L. Wells, T. J. Purcell, J. A. Spudich, and H. L. Sweeney. 2001. Myosin VI is a processive motor with a large step size. Proc. Natl. Acad. Sci. USA 98: 3655 13659. Spudich J. A. 1994. How molecular motors work. Nature 372: 515 518. Svoboda, K., C. F.Schmidt, B. J. Schnapp, and S. M. Block. 1993. Direct observation of ki-

nesin stepping by optical trapping interferometry. Nature 365 721 727. Tanaka H., A. Ishijima, M. Honda, K. Saito, and T. Yanagida. 1998. Orientation dependence of displacements by a single oneheaded myosin relative to the actin filament. Biophys. J. 75: 1886 94. Tanaka H., K. Honma, A. H. Iwane, E. Katayama, R. Ikebe, J. Saito, T. Yanagida, and M. Ikebe. 2001. The motor somain determines the large step of myosin-V. Nature 415: 192 195. Vale R. D., T. Funatsu, D. W. Pierce, L. Robmerg, Y. Harada, and T. Yanagida. 1996. Direct observation of single kinesin molecules moving along microtubules Nature 380: 451 453. Wazawa T., Y. Ishii, T. Funatsu, and T. Yanagida. 2000. Spectral sluctuation of a single fluorophore confugated to a protein molecule. Biophys. J. 78: 1561 1569. Wells A. L., A. W. Lin, L.-Q. Chen, D. Sater, S. M. Caln, T. Hasson, B. O. Carragher, R. A Milligan, and H. L. Sweeney. 1999. Myosin VI is an actin-based motor that moves backwards. Nature 401: 505 508. Yanagida T., M. Nakase, K. Nishiyama, and F. Oosawa. 1984. Direct observation of motion of single F-actin filaments in the presence of myosin. Nature 307: 58 60. Yanagida, T., K. Kitamura, H. Tanaka, A. H. Iwane, A. Ishijima, and S. Esaki. 2000. Single molecule analysis of the actomyosin motor. Curr. Opin. Cell Biol. 12: 20 25.

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Part 3 Functional Implications

14 Mitotic Spindle Motors J. M. Scholey and A. Mogilner

14.1

Microtubules, Motors and Mitosis

Mitosis, the process by which identical copies of the replicated genome are distributed to the daughter products of each nuclear division, depends upon the action of the mitotic spindle, a protein machine that uses microtubules (MTs) and MT-based motor proteins to assemble itself and to segregate sister chromatids (Karsenti and Vernos, 2001, Mitchison and Salmon, 2001, Wittman et al., 2001). Spindle morphogenesis begins during prophase and pro-metaphase when MTs, motors, chromosomes and centrosomes interact and self-organize into a bipolar structure (Fig. 14.1) which by metaphase consists of pairs of sister chromatids aligned on the spindle equator facing opposite spindle poles. During the subsequent anaphase, sister chromatids are moved to opposite poles while the poles themselves move further apart and finally, during telophase, the nuclear envelope re-assembles around the segregated sisters. The movement of chromosomes and the positioning of spindle poles throughout mitosis depend upon mitotic motors, proteins that use nucleotide hydrolysis to generate force and directed motion. Mitotic motors include polymerizing and depolymerizing MTs that exert pushing and pulling forces, respectively as well as some members of the dynein and kinesin families, which generate force in the spindle by stepping along the MT polymer lattice. Here we discuss general principles of force generation by dynamic MTs and motor proteins and their deployment in the spindle. We do not present a comprehensive review of the recent literature on mitotic motors which is covered in other reviews (Banks and Heald, 2001, Brunet and Vernos, 2001, Heald, 2000, Hidebrandt and Hoyt, 2000, Sharp et al., 2000b). MTs are the major cytoskeletal filaments of the spindle and the structural organization of spindle MTs has been elucidated using careful electron microscopic analysis which reveals that spindle MTs comprise two overlapping radial arrays emanating from spindle poles with their plus ends distal, forming the astral, kinetochore and interpolar MT bundles (McIntosh and McDonald, 1989; Fig. 14.1). Spindle MTs use GTP hydrolysis to facilitate two types of dynamic behavior, dy-

328

14.1 Microtubules, Motors and Mitosis

Figure 14.1. Events of mitosis. (A) Prophase.

Dynein motors pull on astral MT generating outward force. Ncd motors cross-link interpolar MTs and develop inward force. The sum of the forces drives the centrosomes apart. (B) Prometaphase. Both Ncd, and bipolar kinesin motors cross-link the interpolar MTs. The chromosome is captured by the MT polymer and transported poleward (possibly, by dynein motors). (C) Metaphase. The chromosomes are aligned at the ‘equator’. This alignment is the result of dynamically coupled forces generated at the sister kinetochores (1 and 2) and of the

polar ejection forces developed at the MT plus end/chromokinesin complexes at the chromosome arms (3). (D) Anaphase. Segregated chromosomes are pulled poleward by a force generated at the kinetochores and coupled to MT’s plus ends disassembly. At the same time, interpolar MTs switch from MT flux to MT sliding due to an inhibition of MT depolymerization at the poles, the motor Ncd is turned off, and thus the poles separate further. (E) Telophase. Chromosomes are separated, and nuclear envelopes form.

14 Mitotic Spindle Motors

namic instability in which MTs grow and shrink by polymerizing and depolymerizing at their plus ends, and poleward flux in which MT plus-ends facing the spindle equator polymerize while their minus ends located at the poles depolymerize. It is clear that the dynamic properties of MTs are critical for spindle morphogenesis and also for generating forces for mitotic movements (Inoue and Salmon, 1995). Spindle formation and function also depends upon the action of multiple MTbased motor proteins, enzymes that couple ATP hydrolysis to the generation of force and motion relative to MT tracks (Sharp et al., 2000a). It is clear that these motors act by a variety of mechanisms to coordinate chromosome movements, acting for example by a ‘sliding filament mechanism’ and sliding adjacent MTs in relation to one another, driving the intracellular transport of chromosomes or vesicles along MTs, modulating the dynamic properties of spindle MTs, or regulating progression through mitosis by acting as components of the spindle assembly checkpoint. Given this rich repertoire of MT and motor functions in the spindle, a current challenge is to understand how the activities of the individual components are coordinated to produce a precision machine capable of segregating chromatids with the fidelity observed in cells. In this review we discuss current ideas about the physical nature of mitotic movements, mechanisms of force generation by MT dynamics and motor proteins in the spindle, and how these force-generating elements are deployed in the spindle to produce this impressive protein machine.

14.2

The Physical Nature of Mitotic Movements

Mitotic movements occur at the microscopic scale under conditions, where viscosity (Purcell, 1977) and thermal fluctuations (Berg, 1983, Mogilner et al., 2002) play dominant roles. Given the stochastic character of physical processes occurring at this scale, the reliability and precision with which the mitotic spindle operates is quite remarkable. The spindle machinery operates in an aqueous environment. A water molecule is about 0.1 nm in radius, while globular proteins are two orders of magnitude larger. This size difference suggests that the fluid can be treated as a continuum. When an object of size l is moving through the fluid with velocity u, the fluid acceleration around the object is characterized by the term r(du/dt), where r is density, and t is time, and the viscous drag on the object has the magnitude h(d2u/dx2), where h is the dynamic viscosity, and x is the spatial coordinate. In this situation, l and t l/u are the characteristic spatial and temporal scales, respectively, and the orders of magnitude of the inertial and viscous forces on the fluid are (ru2/l) and h(u/l2), respectively. The ratio of these forces is the dimensionless Reynolds number: Re ul/n, where n h/r is the fluid specific viscosity (in water, n Z 106 mm2 s 1). The characteristic size and rate of movement in mitosis are l Z 1 mm and u Z 1 mm s 1, so mitosis is characterized by very low Reynolds numbers: Re Z 10 6 (see Tab. 14.1). In this limit, the viscous force is dominant, while the inertial force is negligible. In a sense, spindle movements are governed by Aristo-

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14.2 The Physical Nature of Mitotic Movements Table 14.1

Physical parameters of the spindle environment.

Symbol

Definition

Value and unit

Re

Reynolds number

Z 10

hc

Effective viscosity of the cytoplasm

100-300 cP (0.1–0.3 pNps/mm2)

m

Characteristic size of a molecular motor protein

Z 10–100 nm

rch

Radius of the chromosome

Z 0.2 mm

lch

Length of the chromosome

Z 6 mm

zch

Viscous drag coefficient of the chromosome

Z 10 pNps/mm

Dch

Effective diffusion coefficient of the chromosome

Z 10

d

Size of tubulin dimer

Z 8 nm

dm

MT diameter

Z 25 nm

lm

Characteristic average length of MTs

Z 10 mm

Ns

Characteristic number of spindle MTs

Z 100

Nf

Characteristic number of MTs in kinetochore and interpolar fiber

10–20

t

MT turnover time in mitosis

20–60 s

Vmt

Characteristic rates of MT growth

10–50 mm/min

DG

Strain energy stored in the MT lattice from the GTP hydrolysis

Z 26 pNpnm/dimer

Fm

Characteristic force that can be generated by a single MT or motor

Z 1–10 pN

Fpr

Characteristic outward (dynein) and inward (Ncd) forces in prophase

Z 10–100 pN

T

Characteristic duration of various events in mitosis

Minutes

Nch

Number of chromosomes

Z 10

Fej

Polar ejection force per MT polymer

Z 1 pN

Fk

Poleward kinetochore force

0.1 pN (min), 100s pN (max)

Vpol

Characteristic rate of poleward movement in anaphase

1–4 mm/min

Vmot

Characteristic rates of motor transport

10–50 mm/min

lmt

MT persistence length

1–5 mm

lch

Chromosome persistence length

10–100 mm

C

Tubulin concentration

10–20 mM

kBT

Thermal energy

Z 4 pNpnm

6

4

mm2/s


telian mechanics (F t u), rather than by Newtonian physics (F t du/dt): the velocity (not the acceleration) of motion is proportional to the applied force. At low Reynolds numbers, the viscous force Fv resisting the object’s motion is proportional to the velocity according to Stokes’s formula: Fv – zu. Here the viscous drag coefficient is z (geometric factor)p hl in the case of a spherical object with radius l, z 6pphl, whereas for a cylinder of length l and radius r moving sidewise, z 4pphl/(ln(l/r) 0.5). For small proteins, the cytoplasm appears aqueous, and its effective microscopic viscosity is close to that of water, h Z 1 cP 10 3 pN mm 2 and thus for a motor protein of characteristic size l Z 10 nm, zpr Z 6pphl Z 10 4 pN mm 1. For objects the size of a Drosophila chromosome (radius of arm, rch Z 0.2 mm and length lch Z 6 mm (Marshall et al., 2002)), the effective cytoplasmic viscosity arises mainly from cytoskeletal deformation rather than aqueous shearing, and the corresponding macroscopic viscosity is two orders of magnitude greater, than that of water, hc Z 200 cP 0.2 pN mm 2 (Alexander and Rieder, 1991, Marshall et al., 2002). Consequently, if the chromosome is pulled sidewise, its effective drag coefficient is zch Z 10 pN mm 1. A protein moving through the fluid is acted on by frequent and uncorrelated momentum impulses arising from the thermal motions of the fluid. This leads to a ‘random walk’, when the protein makes extremely frequent and short steps (of length Z 0.01 nm and duration Z 10 13 s (Mogilner et al., 2002)) in a random direction. The resulting Brownian movement is equivalent to the diffusion of the protein. The corresponding diffusion coefficient is given by the Einstein Relation: D kBT/z. Here kBT is the so-called thermal energy that serves as a gauge of energy in the microscopic world. At the temperature of a spindle, kBT z 4 pN nm, the diffusion coefficient of a globular protein is Dp Z 10 mm2 s 1, and that of a chromosome is Dch Z 10 4 mm2 s 1. In the intracellular world, diffusion is very effective over small time intervals and short distances, while forced drift is more effective at greater times and longer distances. The reason for this effect pis that the distance traveled by diffusion grows as a square root of time, d Z Dt, unlike the drift increasing proportionally with time, d Z ut. Thus for a globular protein it would take Z 10 s to move randomly over 10 mm faster than if it were moved the same distance by a unidirectional molecular motor. However, for larger objects, this is not so. A chromosome would need a few days to move the same distance (t Z (10 mm)2/10 4 mm2 s 1)! Clearly, the chromosomes cannot move effectively by diffusion in the course of mitosis (whose characteristic time is minutes), and instead the biased walk of mitotic motors underlies the movement of chromosomes and spindle poles.

14.3

MT Polymerization and Depolymerization as Mitotic Motors

Spindle MTs are built from ab-tubulin heterodimers each containing two molecules of GTP, one of which is concealed in the a-subunit while the other is bound to the b-subunit and exposed to water. Within the MT polymer lattice,

331

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14.3 MT Polymerization and Depolymerization as Mitotic Motors

these subunits are organized into a helical B-lattice of usually 13 protofilaments (pf) that form a hollow tube of diameter 25 nm with a discontinuity, or seam, between two of the 13 pfs (Nogales et al., 1999). Along a pf, all subunits point in the same direction (ab-ab-ab) which gives the MT a structural polarity and by convention the b-tubulin end is called the plus end while the a-tubulin end is called the minus end. This structural polarity is crucial for MT function because the two ends have different polymerization kinetics with the plus ends polymerizing and depolymerizing faster than the minus ends, and because it constrains the directionality of kinesin and dynein motors which move unidirectionally along the polymer lattice. The polymerization and depolymerization of spindle MTs coupled to GTP hydrolysis underlie dynamic instability and flux (Desai and Mitchison, 1997, Mitchison, 1989, Mitchison and Kirshner, 1984a,b). Flux is the term used to describe the movement of tubulin subunits from MT plus-ends facing the spindle equator to the MT minus-ends facing the poles and is thought to depend upon MT polymerization at the plus ends and MT depolymerization at the minus ends being coupled to poleward translocation of the polymer lattice. Dynamic instability describes the behavior of the ends of individual MTs which alternate between phases of polymerization (growth) and depolymerization (shrinkage) with the transitions from growth to shrinkage being termed catastrophes, and the converse transitions being termed rescues. Thus, dynamic instability is characterized by four parameters: rates of growth and shrinking and frequencies of catastrophes and rescues. While individual MTs are engaged in this stochastic behavior, a population of MTs can be described by its length distribution and average number as determined using these parameters together with the effective nucleation rate (Dogterom and Leibler, 1993). In the spindle, MTs are nucleated on g-tubulin ring complexes located in the centrosome of amphiastral spindles (Schiebel, 2000). The a-ends of tubulin heterodimers bind to the g-tubulin ring complexes and consequently the plus ends grow radially outward and display dynamic instability. The end of a growing MT can act as a motor that generates a pushing force. For example, experiments with MTs growing inside liposomes showed that polymerization of MTs can generate enough force to deform the membrane (Fygenson, 1995, Hotani and Miyamoto, 1990). More recently, Dogterom and Yurke (1997) showed that MTs polymerizing against the wall of a chamber could generate a pushing force of several pN. Thus in the spindle, a centrosome-nucleated MT could, in principle, polymerize at its distal plus end and exert a force that pushes a chromosome away from the pole, during pro-metaphase congression, for example. Theoretical modeling has demonstrated that such pushing forces can be explained by a thermal ratchet mechanism (Mogilner and Oster, 1999, van Doorn et al., 2000). According to the elastic polymerization ratchet model, a MT growing against an obstacle is involved in Brownian motion and consequently it undulates and bends very frequently. When the MT is bent, a gap appears between its tip and the obstacle. If a large enough gap persists for a sufficiently long time interval, a tubulin dimer can intercalate into the gap and assemble onto the tip of the growing poly-


mer. This increases the MT’s length so when the longer polymer’s tip contacts the obstacle again, the MT remains bent and the corresponding elastic force pushes the obstacle forward. The energy for this pushing force is supplied by the binding free energy of GTP dimers that associate at the growing tip of the MT and is used to rectify the Brownian motion of the tip. Strictly speaking, the force is generated by thermal fluctuations of the MT, and the binding free energy is used to rectify its thermal bending. If a MT assembles against no resistance, then its elongation rate is simply Vmt d(konC – koff), where d 8 nm/13 Z 0.6 nm is the MT length increment associated with the addition of a dimer to the tip of one of 13 pfs, C is GTP tubulin concentration, and kon and koff are the subunit association/dissociation rates, respectively. Detailed statistical physical analysis (Mogilner and Oster, 1999) shows that the thermal bending undulations of the fiber are much faster than the process of dimer assembly. When a MT polymerizes against a load force FL which resists MT growth, the effective association rate konC is modified by a probability p(FL, d) of there being a gap of width d or greater between the MT tip and the obstacle: Vmt d(konC p(FL, d) – koff). This equation with the specified function p(FL, d) gives the force-velocity relation for a polymerizing MT pushing against a load. At low loads the MT can polymerize rapidly, but as the load force increases it will slow down the rate of growth, until polymerization stalls. The stall force corresponds to the maximal pushing force achieved when growth is stopped. Near the stall, the probability function is given by the Boltzmann factor: p(FL, d) Z exp[FL d/kBT]. Thus the elongation rate decreases exponentially with increasing load, and the stall force can be found from the balance of the effective association and dissociation rates: Fs Z (kBT/d)pln[konC/koff ]. The order of magnitude of the pushing force is common for all thermal ratchet mechanisms and is equal to the thermal energy divided by the step of polymerization, Fs Z kBT/d Z 6.5 pN. The logarithmic factor is of the order of unity in a wide range of the system’s parameters, so one MT fiber can generate the pushing force from a few to tens of pN. While polymerizing MTs can generate a pushing force, depolymerizing MTs can develop pulling forces. For example, using in vitro assays, Coue et al. (1991) observed that the depolymerizing ends of MTs could pull particles at rates of almost 1 mm s 1 against estimated viscous forces of Z 10 pN, and subsequently Lombillo et al. (1995a) found that plastic beads coated with plus end-directed MT motors remain attached to the plus ends of depolymerizing MTs and are carried towards the MT minus ends as the polymer shortens. These results suggest that, in the spindle, it is possible that plus end-directed MT motors on the kinetochore could attach to the plus end of a depolymerizing centrosome-bound MT during anaphase in a way that allows the depolymerizing MT to pull the chromosome to the pole. The nature of the pulling force associated with MT depolymerization remains elusive. The earliest theory by Hill (1985) suggested that the tip of the depolymerizing MT is associated with a docking protein having the form of a sliding collar that allows tubulin dimers to dissociate freely from the MT tip. The interior of this ‘collar’ has high affinity for the MT lattice and consequently when subunits dissociate from the tip of the MT, the binding free energy gradient drives the dock-

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14.4 Kinesins and Dyneins as Mitotic Motors

ing protein toward the MT minus end producing a ‘pulling’ force. Another possibility is the ‘conformation wave’ model proposed by Mitchison (1988), in which the elastic force from the pfs curving outward at the disassembling plus end can drive the sliding collar toward the minus end. Finally, Peskin and Oster (1995) developed a quantitative model in which a bead coated with high affinity tubulin-binding proteins undergoes diffusion along the MT polymer lattice. The binding energy gradient prevents the bead from detaching from the plus end of the MT and as it rolls it weakens the bonds between neighboring tubulin dimers and facilitates depolymerization. All such models allow estimates of the order of magnitude of the pulling force developed by MT depolymerization. The origin of this force is very likely a combination of the thermal ratchet mechanism (Brownian motion of a docking protein ratcheted by disassembly at the end of the MT) and the protein elasticity associated with a conformational change in tubulin. The former component can be estimated as kBT/d Z 6.5 pN modified by the logarithmic factor of order 1, and the latter can be estimated from the above by the strain energy stored in the MT lattice from the GTP hydrolysis, DG Z 26 pN nm/dimer (Inoue and Salmon, 1995), divided by d: 26 pN nm/(8 nm/13) Z 45 pN. So, both pushing polymerization and pulling depolymerization forces can range from a few pN to a few tens of pN per MT fiber. These forces are comparable in magnitude to those generated by kinesin and dynein motors, and they are likely to play significant roles in driving mitotic movements.

14.4

Kinesins and Dyneins as Mitotic Motors

Natural selection has created motor proteins that have specialized motor domains capable of converting chemical energy into the generation of force and movement (Howard, 2001, Mogilner et al., 2002). Many of these molecular motors walk vectorially along MT tracks using nucleotide hydrolysis as a fuel and thereby generate forces for mitotic movements (Brunet and Vernos, 2001, Hildebrandt and Hoyt, 2000, Sharp et al., 2000b, c). The generation of force and motion by molecular motors is thought to depend on a mechanical cycle consisting of the power stroke in which the bound motor domain changes its conformation and generates force, alternating with the recovery stroke when the motor domain detaches from the MT and undergoes a diffusive search for the next binding site on the track. This produces a biased random walk whose directionality is determined by the polarity of the MTs and stereospecificity of the motor’s binding to the MT. Tight coupling of these events to a cycle of hydrolysis make the resulting mechanochemical cycle irreversible and unidirectional, so that motors move either towards the plus or the minus ends of MT tracks, corresponding to movement towards or away from the spindle poles. A striking property of molecular motors that distinguishes them from macroscopic motors is the overwhelming importance of thermal fluctuations. For this reason,


all motor proteins must be regarded as ‘Brownian machines’ in which force generation and movement depend on Brownian motion as well as the elastic (and other physical) forces associated with the power stroke. The forces developed by individual molecular motors near stall can be estimated using thermodynamic arguments similar to those used above to estimate the magnitude of forces generated by MT polymerization and depolymerization. The ratchet part of the force is of the order of kBT/d Z 1 pN, where d Z 8 nm is now the size of the motor’s ‘step’. The active power stroke force is limited from the above by the energy of ATP hydrolysis, DGh Z 80 pN nm, divided by the step size: DGh/d Z 10 pN. Thus, molecular motors can develop forces in the range of a few pN. The rate of free movement of the molecular motors is limited by the rates of the associated ATP hydrolysis cycle and the time of the recovery stroke. Normally, ten to hundreds of cycles/steps take place every second, so the motors advance at Z 0.1 1 mm s 1. The motor’s displacement along the MT track is not steady, because stochastic processes of chemical reactions and searches for binding sites govern the steps of the mechanochemical cycle. One quantity that can be monitored as the motor advances is the variance of its displacement, Var[x(t)] about the mean, ‹x(t)› Vt. Normally, the variance grows linearly with time: Var[x(t)] 2Deff t, where Deff is the effective diffusion coefficient of the motor. The greater the number of reaction processes associated with each mechanochemical cycle, the less random the motor’s ‘walk’ becomes. In addition to the force velocity relations and statistical properties, motor proteins are characterized by the duty ratio, which is the fraction of time when the motor domains are bound to MTs and developing force (or, roughly speaking, ratio of the time of the power stroke to the time of the cycle). Most of these relations and parameters have not been determined for mitotic motors. Several members of the kinesin and dynein families function as mitotic motors (Brunet and Vernos, 2001, Dujardin and Vallee, 2002, Goldstein, 2001, Hildebrandt and Hoyt, 2000, Hirokawa et al., 1998, Holzbaur and Vallee, 1994, Karki and Holzbaur, 1999, Sharp et al., 2000a, Vale and Fletterick, 1997). The founding member of the kinesin family, conventional kinesin (Howard, 1996), is an intracellular transport motor capable of traveling long distances along a microtubule without dissociating (Howard et al., 1989) and which may drive transport along spindle fibers. It has two heads motor domains at the N-terminus of the heavy chain and moves in a fascinating head-over-head fashion toward the plus end: while one head is attached, another is ‘searching’ for the next binding site in the plus-end direction. Kinesin-related proteins share the same highly conserved motor domain but outside the motor domain they differ. Motors that have N-terminal motor domains (KIN-N) move towards the plus end of MTs, those with C-terminal motor domains (KIN-C) move towards the minus end, whereas motors from the KIN-I subfamily which contain internally located motor domains seem to destabilize MT ends (Desai et al., 1999, Schroer, 2001). Dyneins are structurally unrelated to kinesins, but they also use ATP hydrolysis to move (in a minus-end direction) and generate force. The dynein motor domain consists of six AAA (ATPases) domains only one of which hydrolyzes ATP (King, 2000), and a short but structurally

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14.4 Kinesins and Dyneins as Mitotic Motors

complex stalk that contains MT-binding sites (Gee et al., 1997). Dynein can produce force by a conformational change in the head applying tension to the rigid stalk and pulling on the microtubule. Alternatively, the stalk might rotate about a fulcrum located within the head, like a windshield wiper (Gee et al., 1997). Mitotic motors act by a variety of mechanisms (Brunet and Vernos, 2001, Heald, 2000, Sharp et al., 2000a). For example, it has long been proposed that force is generated in the spindle by motor-driven MT sliding (McIntosh et al., 1969, Scholey et al., 2001, Sharp et al., 2000a), and accordingly, members of the plus-end-directed bipolar (BimC) family of kinesins oligomerize to form bipolar homotetramers with motor domains positioned at opposite ends of a central rod, that are thought to be capable of cross-linking MTs throughout the spindle and sliding apart anti-parallel MTs within interpolar MT bundles (Sharp et al., 1999). On the other hand, members of the minus-end-directed C-terminal kinesin family form homodimers, containing C-terminal motor domains, linked by a coiled-coil rod to a tail that contains nucleotide-insensitive MT binding sites. Thus, C-terminal kinesins are also thought to be capable of cross-linking and sliding adjacent MTs (Karabay and Walker, 1999), possibly acting on interpolar MTs to draw the poles together (Sharp et al., 2000a). Similarly dynein appears to be capable of sliding MTs relative to adjacent MTs or relative to cortical actin filaments, exerting pulling forces on spindle poles (Dujardin and Vallee, 2002). There also exist mitotic motors that transport chromosomes as cargo along the surface lattice of spindle MTs. For example, the plus-end-directed motor, CENP-E and the minus-end-directed motor, dynein, both localize to kinetochores, and are likely to participate in chromosome congression and segregation (Sharp et al., 2000b, Savoian et al., 2000, Yucel et al., 2000). Several presumptively plus-end-directed motors, collectively referred to as chromokinesins, bind to chromosome arms as cargo, where they are thought to provide forces that push chromosomes towards the metaphase plate (Brunet and Vernos, 2001) and some intracellular transport proteins appear to move vesicles and protein complexes along spindle MTs (Section 14.8). Finally it is clear that mitotic motors can regulate spindle MT assembly dynamics. For example, the KIN-I motor, XKCM1/MCAK, localizes to kinetochores where it is thought to induce disassembly of kinetochore MTs (Desai et al., 1999) whereas the orphan motor, CENP-E is thought to be able to use its plusend-directed motor activity to anchor kinetochores to the shortening plus ends of MTs during anaphase (Lombillo et al., 1995b). Thus XKCM1/MCAK motors could induce the shortening of kinetochore-to-pole MTs during anaphase, and this could work in concert with the plus-end anchoring activity of CENP-E to transduce MT shortening into poleward forces on chromosomes. The diversity of mitotic motors can be appreciated by considering a single organism, Drosophila melanogaster which has 36 MT-based motors and of these, 11 are strong candidates for being mitotic motors (Table 14.2). Cytoplasmic dynein, which is localized to cortical structures and kinetochores in syncytial blastoderms, has been implicated in spindle pole positioning, poleward chromosome movements and the transport of checkpoint proteins (Savoian et al., 2000, Sharp et al.,

14 Mitotic Spindle Motors Table 14.2

Mitotic motors in Drosophila embryos.

Motor Protein

Cytogenetic position

Structure

Function

1. Cytoplasmic dynein

64C

Pole pole separation; poleward chromosome motion; checkpoint

2. Bipolar kinesin, KLP61f

61F

Cross-linking spindle MTs; pole pole separation

3. C-terminal kinesin, Ncd

99C

Cross-linking spindle MTs; pulling poles together; pole organization

4. Pav KLP (MKLP1)

64B

?

Mid-zone organization; cytokinesis

5. KLP 3A

3A

?

Pole pole separation; nuclear positioning; mid-zone organization

6. CENP-meta 7. CENP-ana

32E

8. KIN-I: KLP59C (XKCM1) 9. KIN-I: KLP10A

59C 10A

10. Chromokinesin: KLP54D 11. Chromokinesin: KLP38B

54D

Checkpoint; coupling to depolymerizing MTs; congression x

x

38B

?

x

x

Depolymerizing kinetochore’s fibers at the kinetochores Depolymerizing astral MTs at spindle poles Antipolar chromosome movement Pole pole separation

2000a, b, Wojcik et al., 2001). The bipolar kinesin, KLP61F and the C-terminal kinesin, Ncd are thought to cross-link MTs within interpolar MT bundles in embryonic spindles, where they generate antagonistic outward and inward forces on spindle poles (Sharp et al., 1999, 2000a). Two other kinesins, KLP3A and PavKLP localize to the spindle inter-zone (Adams et al., 1998, Williams et al., 1995), where they may also associate with interpolar MT bundles and contribute to the generation of forces that position spindle poles. Like dynein, the kinesins, KLP59D, KLP10A (both KIN-I motors), CENP-meta and CENP-ana, are candidates for being kinetochore motors (Desai et al., 1999, Yucel et al., 2000). There are three chromokinesins that are candidates for driving the transport of chromosome arms towards the spindle equator (Brunet and Vernos, 2001) namely KLP38B, KLP54D and Nod. Of these, Nod appears to function specifically in the female oocyte meiotic spindle, and there is no evidence for a mitotic role.

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14.5

Functional Coordination of Mitotic Motors

An important issue facing mitosis researchers is why the mitotic spindle uses so many motors. Our current view is that cells use multiple mitotic motors in parallel to generate a delicate balance of complementary and antagonistic forces (Hoyt and Gieser, 1996, Sharp et al., 2000a). These ideas emerged initially from elegant genetic studies carried out in yeast, but they have recently been extended and refined by exploiting the Drosophila embryo, where it is possible to visualize and quantify specific mitotic movements at high temporal and spatial resolution in the presence and absence of specific motor inhibitors (Hildebrandt and Hoyt, 2000, Sharp et al., 2000a). The results argue that specific mitotic movements are not driven by individual mitotic motors acting alone, but instead depend on shifts in the balance of forces generated by multiple motors. For example, in Drosophila embryos, when spindle pole positioning is measured as a function of time, pole separation proceeds in a complex fashion, with stops, starts and rate changes that are thought to reflect changes in the net force acting on the poles, and these net forces in turn reflect the action of multiple MT-motors that serve to position the poles (Sharp et al., 2000a). When these forces balance one another, spindle pole spacing is maintained under isometric tension in a quasi-stable steady state structure. During mitosis, the spindle appears to pass through a series of these steady-state structures, at which points multiple complementary and antagonistic motors precisely balance one another. Transitions from one steady state to the next are thought to reflect the up- or downregulation of subsets of motors, which alters the net force acting on the spindle poles, allowing a specific mitotic movement that is visible as a change in the spacing of the spindle poles. We refer to this model as the multiple motor-dependent transient steady state model for spindle pole positioning (Sharp et al., 2000a). Transient motor-generated steady-state structures may also serve as a mechanism for chromosome positioning during chromosome capture, congression and segregation. Progression through this pathway could plausibly be signaled by the density and polarity patterns of MTs surrounding chromosome arms and kinetochores. The response of a chromosome to its position would, in turn, be determined by the relative strength of the poleward versus plateward forces generated by motors positioned on these structures together with MT polymerization depolymerization dependent forces. Thus, chromosomes would always tend to move toward a specific steady state or balance position, which could be altered at specific stages of the cell cycle by alterations in the activity of specific motors as well as by subtle changes in spindle and chromosome structure. For example, it is easy to imagine how a balance of kinetochore motor-, chromokinesin- and MT polymerization depolymerization-generated forces could position chromosomes on the equator during the metaphase steady state, but testing this hypothesis and discerning the details require more work (Kapoor and Compton, 2002; Section 14.6.2).


14.6

Motor Action and Force-Generation during Mitosis

Spindle formation and chromosome segregation (Fig. 14.1) involves a complex interplay between dynamic MTs and motor proteins associated at spindle poles, kinetochores and chromosome arms, and is likely to be dependent on forces generated by both MT polymerization depolymerization and by molecular motor action (Banks and Heald 2001, Brunet and Vernos, 2001, Heald, 2000, Hunter and Wordeman, 2000, Inoue and Salmon, 1995, Scholey et al., 2001, Sharp et al., 2000a). In what follows, we discuss the motor and MT-related mechanisms in mitosis. 14.6.1

Mitotic Motors and Spindle Formation at Early Stages of Mitosis

During interphase, MTs are nucleated from g-tubulin ring complexes associated with the centrosome, forming a single radial array (Schiebel, 2000). A subset of these MTs have their minus ends anchored on the centrosome, while others appear to be released by katanin-dependent severing, but maintain a physical association with centrosomes via MT cross linking motors (Heald 2000, Merdes et al., 1996). The formation of a bipolar spindle is associated with an increase in the dynamic instability properties of MTs (Salmon et al., 1984), the migration of the two centrosomes around the nuclear envelope (Sharp et al., 2000a) and the organization of centrosome-associated MTs into a bipolar array (Karsenti and Vernos, 2001). Motor proteins play important roles in many aspects of this self-organization process, although the details appear to vary between systems (Heald et al., 1997, Karsenti and Vernos, 2001, Sharp et al., 2000a). For example, in Drosophila embryos during interphase prophase, centrosomes separate around the surface of the nuclear envelope at an initial fast rate that slows down as the poles separate until the centrosomes come to lie a few microns apart on opposite sides of the nucleus where they are maintained for 2 3 min in the ‘prophase steady state’ (Sharp et al., 2000a). Function inhibition experiments suggest that these events depend upon antagonistic outward and inward forces generated by cortical dynein and interpolar Ncd, respectively (Fig. 14.1A). The following model can explain these experiments quantitatively (Cytrynbaum et al., unpublished data). In this model the outward force responsible for the initial fast rate of pole separation is due to cortical dynein generating a few tens of pN pulling force on a subset of astral MTs together with a pushing force due to polymerization of interpolar MTs. As the centrosomes separate towards opposite sides of the nucleus, this outward force becomes directed almost perpendicular to the surface of the nuclear envelope, and its projection onto this surface decreases. The decreasing outward force is opposed by an increasing inward force generated by Ncd motors that cross-link MTs in the interpolar MT bundle between the poles. Assuming there are about a dozen Ncd motors per micron, Z 10 pN inward force would be generated per micron of interpolar bundle, and the total inward force would grow in proportion to the overlap between the interpolar MTs which in

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turn increases as the poles separate. At a few microns separation, the outward and inward forces equilibrate, explaining the stable steady state. One important question concerns the precision and robustness of pole separation. During the prophase steady state (as well as those occurring at other stages of mitosis), the fluctuations in the interpolar distance are of the order of only a few percent. According to probabilistic arguments, if, on the average, N MTs reach the cell cortex, then the average fluctuations in this number of MTs is of p the order of N . Therefore the relative fluctuations of the outward force and consequently the relative p fluctuations of the separation distance between the poles, would be Z 1/ N . For example, a 3 % fluctuation corresponds to N Z 1000 MTs which is far greater than the number of MTs observed. This could indicate that additional control mechanisms maintain the spindle dimensions. Further quantitative research is needed to address the problem of the precision of spindle morphogenesis. 14.6.2

Mitotic Motors and Force Generation in Prometaphase Metaphase

Following nuclear membrane fenestration there is another pause, the pro-metaphase steady state, followed by an episode of pole separation that increases pole pole spacing to distances characteristic of the metaphase anaphase A steady state. These changes have been explained qualitatively in terms of changes in the balance of bipolar kinesin, Ncd and dynein motor-generated forces, although further work is required to learn the roles of other motors and MT dynamics in these events, as well as to elucidate the details (Sharp et al., 2000a). Furthermore, following the fenestration, spindle formation and function is more complex as new players, the chromosomes and kinetochores, emerge and begin to play dominant roles. Kinetochores are specialized sites on condensed chromosomes which form a localized, high-affinity site for the capture of spindle MTs. They ensure a high fidelity of segregation and act as central players in chromosome motility by monitoring chromosome attachment to MTs and regulating the metaphase anaphase transition. During pro-metaphase, dynamically unstable MTs associated with centrosomes probe the cytoplasm in an exploratory fashion, and chromosome capture depends upon a chance attachment of the side of a kinetochore to the wall of a growing MT. The dependence of this attachment process on the stochastic phenomenon of dynamic instability seems unreliable, but the quantitative model of Holy and Leibler (1994) argues that, in fact, spindle MT dynamics seemed to be tuned to optimize this ‘search and capture process’. For example, the distance between the spindle poles and the chromosomes is d Z 10 mm, similar to the average length of growing MTs in metaphase. If MTs were much longer than this, those that ‘miss’ a kinetochore would wastefully grow too long but if they were too short, most of them would fail to reach the kinetochores. When Z 100 MTs (which is the order of magnitude of the number of MTs radiating from each pole) grow to Z 10 mm, there is


one MT fiber per few square microns of the surface on which the chromosomes are distributed. The area of a kinetochore is Z 1 mm2, so a MT ‘finds’ a kinetochore after just a few tries. Each trial takes less than a minute, because the plus ends polymerize at rates of tens of microns per minute, so over a few minutes each kinetochore is ‘captured’ by a MT fiber. This estimate is in good qualitative agreement with observations. Note, that dynamic instability provides a much more effective mechanism of ‘search and capture’, than equilibrium polymerization kinetics could. The latter would result in an effective random walk of the polymer’s tips with the steps equal to the dimer’s size d Z 8 nm. The corresponding average time for such polymers to reach d Z 10 mm length is greater than that for the unstable growing MT by the factor (d/d) Z 1000, which would make the ‘search and capture’ time equal to many hours. Once attached, the kinetochore translocates polewards along the wall of the MT at rates approaching 1 mm s 1, close to the velocity of MT gliding induced by cytoplasmic dynein (Rieder and Alexander, 1990) and within a minute the MT develops a stable plus end-contact with the kinetochore (Fig. 14.1B). Subsequently, 15 30 MTs rapidly establish end-on connections with each kinetochore and become cross-linked into a tight bundle (MT MT spacing 50 100 nm) referred to as a kinetochore (kt) fiber. MTs do not randomly ‘search for’ the kinetochore after the first attachment, but grow and connect to it in a cross-linked state, which reduces the time of kt fiber formation. The establishment of bipolar attachment involves the formation of such kinetochore fibers on each member of a pair of sister chromatids. The fascinating process of congression follows, in which position-dependent forces align the chromosomes on the metaphase plate (Kapoor and Compton, 2002). Current evidence suggests that these forces are generated locally at the kinetochores and chromosomes rather than by ‘traction fibers’ acting along the length of the kt fiber, with pulling forces on the kinetochore being antagonized by pushing forces exerted on the chromosome arms from astral ejection forces (Hays et al., 1982, Kapoor and Compton, 2002, Rieder and Salmon, 1998, Rieder et al., 1986; Fig. 14.1C). By measuring the bending of chromosome arms, Marshall et al. (2001) estimated that the magnitude of the astral ejection force exerted by one MT fiber is Z 1 pN. During congression, mono- and bi-oriented chromosomes undergo a series of oscillatory movements at rates Z 2 3 mm min 1 and amplitudes of Z 2 3 mm. These oscillations are coupled to MT polymerization at the lagging kinetochore and depolymerization at the leading one. Some observations indicate that poleward motion is driven by an action at the leading kinetochore, while the lagging one is passive and does not push. It is likely that some bi-stability based on a positive feedback between sister kinetochores underlies these oscillations, superficially similar to the cooperative bi-stability of dynein and kinesin motors observed in in vitro motility assays (Vale et al., 1992). It is likely that multiple mitotic motors cooperate to position chromosomes during chromosome capture, congression and alignment. MT polymerization can, in

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principle, exert pushing forces on kinetochores and chromosome arms, whereas depolymerization can generate pulling forces on these structures. Cytoplasmic dynein is associated with kinetochores in many systems, and may contribute to chromosome positioning by transporting kinetochores polewards. Indeed, the inhibition of cytoplasmic dynein function in Drosophila embryos interferes with congression and prevents the proper alignment of chromosomes on the metaphase equator (Sharp et al., 2000b), although the significance and generality of this observation is a matter of debate (Kapoor and Compton, 2002). MT-destabilizing KIN-I motors could depolymerize MTs at the kinetochore and they have been implicated in many aspects of kinetochore function (Maney et al., 2000). The plus-end-directed orphan kinesin motor, CENP-E may be associated with kinetochore movements towards and away from the spindle equator, being capable of both translocating kinetochores along the polymer lattice towards the plus ends of MTs and also coupling plus-end motility to the depolymerization of kt MTs (Lombillo et al., 1995b, Schaar et al., 1997, Yucel et al., 2000). The polar ejection force acting to generate antipolar forces on chromosome arms appears to depend upon a combination of MT polymerization and plus-end transport driven by chromokinesins (Brunet and Vernos, 2001). Thus both polymerization and plus end-directed motors can contribute to powering anti-poleward movement of chromosomes while depolymerization and minus-end-directed motors can drive poleward movement, while kinetochore motors can couple the translocation of kinetochores to assembly/disassembly of the MT plus-ends. Understanding exactly how these force-generating elements cooperate to position chromosomes and achieve the balance that aligns chromosomes on the spindle equator at the metaphase steady state presents a fascinating technical challenge (Section 14.4). It has been proposed that chemical gradients originating at the poles could contribute to the positioning of the chromosomes in spindles (Karsenti and Vernos, 2001), but the large fluctuations characteristic of these gradients deem this suggestion unlikely. An even greater puzzle than the mechanism of force generation by mitotic motors, is how the kinetochore machinery detects and responds to tension. During congression, high tension promotes switching to anti-poleward movement, whilst low tension tends to lead to poleward movement (Rieder and Salmon, 1994). Also, high tension stabilizes attachment of MT fibers to the kinetochore (Nicklas and Ward, 1994). Whether mitotic motors contribute to tension detection and the response at the kinetochore is another fascinating question. Finally, mitotic motors may also contribute to the poleward flux of MTs in half spindles, which is superimposed on kinetochore-localized movements (Desai et al., 1998, Mitchison, 1989, Mitchison and Sawin, 1990). During metaphase, when the spindle length is constant, there is continuous concerted depolymerization of minus ends near the spindle poles, possibly mediated by the MT severing factor, katanin, and polymerization of plus ends near the cell’s equator. These dynamic events are likely to be coupled to the motor-driven poleward translocation of the MT polymer lattice itself, but the identity of the motor(s) responsible, the factors that regulate this process, as well as its function, remain mysterious.


14.6.3

Mitotic Motors and Force Generation in Anaphase

Segregation of chromosomes at the onset of anaphase begins when all chromosomes are properly aligned on the spindle equator and the anaphase-promoting machinery exerts its effects (Section 14.9) including the degradation of chromokinesins with the consequent decrease in the plateward forces acting on chromosome arms which tips the balance of forces in favor of poleward motion (Funabiki and Murray, 2000). Multiple motors located on kinetochores could contribute to poleward chromosome movement (Fig. 14.2). How much force is required to move a chromatid at the observed rate of poleward chromosome movement (microns per minute)? The viscous drag coefficient of a chromosome in a fluid with the viscosity of cytoplasm (Alexander and Rieder, 1991, Marshall et al. 2000, Nicklas, 1983) is Z 10 pN s mm 1, which means that a small force Z (10 pN s mm 1)p(0.1 mm s 1) Z 1 pN would be sufficient to drag the chromosome poleward at the velocities observed. Nicklas (1983: 0.1 pN), Houchmandzadeh et al. (1997: 1 pN) and Alexander and Rieder (1991: 10 pN) made similar estimates. This force is comparable to the force generated by a single depolymerizing MT, or by a single molecular motor. However, the kinetochores seem to generate much greater poleward forces. The force the generated on the kinetochore in anaphase was measured using a calibrated flexible glass needle (Nicklas, 1983). Nicklas discovered that chromosome velocity was not affected until the opposing force reached approximately 100 pN, and then fell rapidly with increasing force. The opposing force that caused chromosome velocity to fall to zero the force that matched the maximum force the spindle could exert on the chromosome was of the order of 700 pN, which is several orders of magnitude greater than the calculated value of 0.1 10 pN required to overcome viscous drag. Similarly, Houchmandzadeh et al. (1997) mea-

Generation of poleward force at the kinetochore. The kinetochore-associated motor (CENP-E) walks to the plus end of depolymerizing MTs (blue) and couples MT disassembly to polar transport of the kinetochore. KIN-I motors, such as MCAK, facilitate MT disassembly as indicated by the formation of

Figure 14.2.

curved protofilaments. In addition, the minusend-directed kinetochore-associated dynein motor pulls the chromosome poleward. Its movement is governed by the MT plus end disassembly. The dynein is shown attached via dynactin to the fibrous ‘corona’ that emanates from the kinetochore (red) itself.

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sured the force exerted by the spindle on a newt chromosome at anaphase as being hundreds to thousands of pN, again supporting the conclusion that kinetochores generate far larger forces than necessary for poleward movement. Moreover, poleward chromosome movements are accompanied by depolymerization of kt MTs (Inoue and Salmon, 1995), but kinetochore motility is much slower (microns per minute) than the free depolymerization rate (tens of microns per minute). These findings suggest that some velocity governors have to exist at the kinetochores. Why are the poleward movement rates an order of magnitude slower than the free depolymerization velocity and how is the maximal force of few hundreds pN per kinetochore generated? What is the nature of the velocity governors? Finally, what is the reason for generating this excessive force, when a much smaller one would be sufficient? First of all we note that 15 20 depolymerizing kt MTs acting together could generate more than 100 pN pulling force on the kinetochore. Similarly, a few tens of cytoplasmic dynein motors associated with the kinetochore could generate Z 100 pN force so in principle such forces can be developed by either motor proteins or MT dynamics. One of the simplest possibilities for the mechanism of the kinetochore force is that dynein motors associated to the kinetochore pull it poleward (Savoian et al., 2000, Sharp et al., 2000b; Fig. 14.2). One could speculate that the release of disassembling tubulin dimers from the plus ends of the kinetochore fiber MTs is hindered sterically at the kinetochore. This would explain the slow rate of the poleward movement. This slow depolymerization would stall the action of the dynein motors, which would explain the large measured maximal force developed at the kinetochore: this force would be equal to the sum of the stall forces for all motors. Such a mechanism could provide more faithful and precise poleward movement. Indeed, the randomness (rate of growth of displacement variance) of a number of motors (including kinesin (Vischer et al., 1999) and a depolymerizing MT (Peskin and Oster, 1995)) is large at the free movement of the motor and decreases dramatically when a load force opposes this movement, reaching minimum near stall. So, it could be that the force-generating elements at the kinetochore are dynein motors. The role of slow MT depolymerization is to stall the dyneins, which makes the effective poleward movement very steady. Further experiments are necessary to put more stringent constraints on models of kinetochore movement and force generation. Separation of chromosomes consists of anaphase B spindle elongation in addition to chromatid-to-pole motion. In many systems anaphase B involves anti-parallel sliding of interpolar MTs coupled to the polymerization of overlapping MTs at their plus ends, which effectively adds to the separation distance between the chromosomes. Bipolar kinesins and dynein have been implicated in driving MT MT sliding for anaphase B in both yeast and Drosophila (Hildebrandt and Hoyt, 2000, Sharp et al., 2000a) but the detailed mechanism and its method of regulation is unclear. Recent work carried out in Drosophila suggests that the spindle poles are maintained at a constant spacing throughout the metaphase anaphase A steady state by a balance of forces involving MT flux, inward forces generated by interpolar C-terminal kinesin, Ncd, and outward forces exerted by cortical dynain and interpolar bipolar


kinesin, KLP61F. This balance appears to be tipped by a downregulation of Ncd activity and a suppression of MT depolymerization at the poles, which tips the balance of forces in favor of outward forces which drive pole pole separation and spindle elongation (I. Brust-Mascher and J. M. Scholey, unpulished data).

14.7

Does a Spindle Matrix Facilitate the Function of Mitotic Motors?

Microtubules and motors are obviously critical components of the spindle machinery, but recent findings have revived interest in the long-standing and important problem of whether the spindle contains another, unidentified mechanical component, a spindle matrix, that could serve as a stationary substrate against which MTs and motors function (Bloom, 2002, Scholey et al., 2001, Wells, 2001). One such finding is the observation made using fluorescence speckle microscopy that the bipolar kinesin, Eg5 remains relatively static in the spindle while its underlying MT tracks flux poleward (Kapoor and Mitchison, 2001). The explanation favored by the authors of this provocative study is that Eg5 is immobilized on a stationary matrix against which the MTs are translocated poleward as they polymerize at the equator and depolymerize at the poles, although other explanations for the slow dynamics of Eg5 are also possible (Wells, 2001). Additional findings that draw attention to the matrix hypothesis concern the recent discovery of two novel filamentous nuclear proteins, Skeletor in fruitfly and Fin1 in yeast, that are proposed to assemble into spindle-shaped structures independent of spindle MTs. It is proposed that these proteins could function as components of a spindle matrix that serves as a stationary platform on which MTs and motors can perform their function (van Hemert et al., 2002, Walker et al., 2000), but unfortunately, the evidence obtained so far is merely guilt by association, as no clear functional evidence is available in support of any mitotic functions for Skeletor or Fin1. Thus the reality of the spindle matrix remains an unproven but fascinating issue. If the spindle matrix is real, one of its possible functions could be to strengthen the spindle machine. The effects of MT elasticity on spindle mechanics have not been investigated thoroughly, but here we make some rough estimates. The MT’s persistence length is lmt Z 103 mm (Bray, 2001). An l 10 mm long polymer with this persistence length would buckle if a force of the order F Z 10 kBT lmt/l2 Z 1 pN was applied to the end of the polymer. In fact, defects in MT lattice would reduce the MTpersistence length, and thus the buckling force even more. It is very likely that forces of the order of tens of pN are applied to the interpolar MTs by bipolar kinesin and Ncd motors, and that forces of hundreds of pN are developed at the kinetochores, more than enough to buckle the MT. Cross-linking of a few MTs would strengthen the bundle of MTs dramatically, and this may be a primary function of motors like bipolar kinesins that cross-link MTs throughout the spindle in a way that allows the underlying MT tracks to remain dynamic. A dense bundle of N MTs would have rigidity and buckling force roughly N2 times greater than a single MTso that for example, 20 MTs cross-linked into a kinetochore fiber would buckle at Z 400 pN, which is the same

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order of magnitude, as the maximal kinetochore force. Similarly, an interpolar bundle of Z 10 cross-linked MTs during prophase would buckle at Z 100 pN, which is the same order of magnitude as the force that multiple bipolar kinesin and Ncd motors could generate. It is possible that something in addition to MT cross-linkers is required to strengthen kt and interpolar MT fibers, and a spindle matrix could provide this. Indeed, if the proposed matrix turned out to be an effective elastic medium with a Young’s modulus of Y Z 104 Pa, similar to that of a dense actin meshwork, then a MT associated with it would be buckled by only very large forces of magnitude Z 100 pN. A bundle of such MTs would be impossible to buckle by the forces characteristic of the spindle. Moreover, this buckling force would be independent of the length of the fiber (Landau and Lifshitz, 1995), unlike an equivalent fiber in an aqueous medium, so even very long MTs would be stable. Such arguments underscore the importance of resolving the fascinating question of whether the spindle matrix does indeed exist (Bloom, 2002, Scholey et al., 2001).

14.8

Mitotic Motors and Intracellular Transport Systems

The notion that vesicle transport in spindles could be significant for the mechanism of mitosis (Sawin and Scholey, 1991) has recently gained renewed interest, based on recent work suggesting that some MT-based motor proteins function during late stages of mitosis to transport both vesicles and signaling molecules along spindle MTs and that these activities are required for the completion of cytokinesis (Finger and White, 2002, Shuster and Burgess, 2002, Skop et al., 2001). This aspect of mitotic motor function in some ways resembles the actions of neuronal and intraflagellar transport motors, including for example kinesin-I, which appears to attach to transmembrane receptor proteins on its vesicular cargo via the Jun N-terminal kinase signal scaffolding proteins, JIP1/2 and JIP3/syd (Goldstein, 2001). In animal cells, cytokinesis is biphasic, involving the determination and ingression of the cleavage furrow, followed by the scission of the mid-body remnant resulting in the final separation of daughter cells (Finger and White, 2002). MT-based vesicle transport is thought to be required to provide additional membrane and thus to increase the surface area for the ingression of the cleavage furrow (at least in some systems) and to seal off the plasma membrane of the two daughter cells as they separate (Fig. 14.3). The scission events associated with vesicle transport along anti-parallel mid-body MTs resemble those involved in cell plate formation associated with the phragmoplast of dividing plant cells, which has long been understood to depend upon the MT-based transport of Golgi-derived vesicles (Lee et al., 2001). The precise roles of the signaling molecules that are associated with MT-based transport systems during these events are not well understood, but they could be involved in controlling MT dynamics in the mid-body or phragmoplast, in regulating the activity of the motors themselves, in signaling cleavage furrow positioning, ingression and scission, or they could be precursors of signaling complexes that assemble in association with new plasma membrane.


Figure 14.3. Intracellular transport by mitotic motors. New work indicates that some molecular motors in the spindle transport vesicles (green) and signaling complexes, delivering new surface membrane for the completion of cytokinesis (see text).

Which mitotic motors are involved in these aspects of cell division? The best candidates are those that associate with the anti-parallel MT arrays that constitute the animal cell mid-zone and the plant cell phragmoplast. For example, the two phragmoplast kinesins, AtPAKRP1 and AtPAKRP2 bind to MTs in a nucleotide-sensitive fashion and, by immunofluorescence microscopy, display different distributions within the phragmoplast (Lee and Liu, 2000, Lee et al., 2001). Thus, AtPAKRP1 forms a relatively tight band that may correspond to the plus ends of overlapping MTs, and is proposed to cooperate with bipolar and C-terminal kinesins to control the organization of mid-zonal MT bundles, whereas AtKRP2 localizes to a relatively broad band of punctate structures, presumably Golgi-derived vesicles, and it is proposed to deliver these vesicles to the developing cell plate (Lee et al., 2001). An additional phragmoplast kinesin, NACK1, is proposed to bind and activate the signaling molecule, NPK1, a Map kinase, and to transport the kinase to the equator of the phragmoplast where the complex is required for the outward expansion of the cell plate and the completion of cytokinesis (Nishihama et al., 2002). The precise mechanism of action of these phragmoplast motors is currently unclear, but the work shows how multiple motors can cooperate to facilitate different aspects of phragmoplast function during cytokinesis, including organizing MTs into ordered arrays that can serve as tracks for the efficient transport of vesicles to the site of abscission. A similar functional cooperation is seen among the motors associated with midzonal MTs during animal cell cytokinesis, most notably members of the MKLP1 family. For example, the C. elegans mid-zonal kinesin, ZEN-4/MKLP1, is required for the tight bundling of mid-zonal MTs into a normal mid-body, and the loss of its function gives rise to a failure of the completion of cytokinesis (Powers et al., 1998, Raich et al., 1998). This motor appears to interact functionally with components of G-protein signaling pathways because ZEN-4/MKLP1 appears to interact with a

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Rho-family GAP protein, CYK-4, forming a tight complex that cross-links MTs into bundles (Mishima et al., 2002) potentially forming organized tracks in the midzone for efficient vesicle delivery to the site of scission. Since CYK-4 binds to the neck region of ZEN-4/MKLP1 it may regulate motility and MT MT bundling by the motor complex. The transport of Golgi-derived vesicles along the bundled MTs may be mediated by another mid-zonal MKLP1-related kinesin, Rab6-KIFL, which binds the small ras-related GTPase, Rab6, and is implicated in membrane traffic associated with the Golgi apparatus. Rab6 KIFL localizes to the spindle mid-zone of mitotic vertebrate cells much like ZEN-4/MKLP1, and the perturbation of its activity disrupts cytokinesis, suggesting that Rab6 KIFL may transport Golgi-derived vesicles along MTs that have been bundled into organized tracks by ZEN-4/MKLP1-related proteins, thus providing membrane for the final stages of cell cell scission (Hill et al., 2000). This is by no means the whole story for intracellular transport motors in the spindle mid-zone however, as ZEN-4 and its Drosophila homolog, PavKLP display functional interactions with the Polo and Aurora kinase regulatory systems, respectively (Carmena et al., 1998, Severson et al., 2000). The finding that the MKLP1-related kinesin, CHO-1 has an extra actin-binding domain that is missing in other MKLP1s, and which may allow this motor to connect mid-zonal MTs to the cell cortex (Kuriyama et al., 2002), introduces additional complexity. Moreover, the kinesin KLP3A from Drosophila is not a member of the MKLP1 family yet it is also required for the proper organization of MTs in the spindle mid-zone, and loss of its function leads to failures in cytokinesis (Williams et al., 1995), further complicating the issue of how these mid-zonal motors may cooperate to ensure the successful completion of cytokinesis. There are some indications that the participation of the aforementioned mid-zone and phragmoplast motors in cytokinesis represents only the tip of the iceberg. For example, there exist motors with well-characterized intracellular transport functions in non-mitotic cells that might also be deployed to perform cell division-related functions in the spindle, although the evidence is currently less compelling. One of these is conventional kinesin-I itself, a protein that associates with vesicles in sea urchin embryonic mitotic spindles (Wright et al., 1991) and is proposed to deliver exocytotic vesicles out along astral MTs to the cell surface for resealing damaged membranes (Bi et al., 1997). The resealing of wounded membranes by Ca2 -regulated exocytosis resembles the membrane fusion events that are involved in the scission of daughter cells during cytokinesis (Finger and White, 2002) and it is plausible to think that kinesin-I-dependent vesicle transport along astral MTs could be responsible for the new membrane addition that occurs in the late telophase cleavage furrows of sea urchin embryos (Shuster and Burgess, 2002), although antibody and dominant negative inhibition experiments did not reveal a requirement for kinesin-I in cell division (Bi et al., 1997, Wright et al., 1993). Heterotrimeric kinesin-II is another candidate (Cole et al., 1993). This motor is best known for its role in intraflagellar transport and ciliogenesis (Goldstein, 2001) but it localizes to punctate detergentsensitive structures in some mitotic spindles (Henson et al., 1995) and, like Rab6 KIFL it is implicated in G-protein signaling and Golgi-associated membrane


trafficking (Le Bot et al., 1998, Shimizu et al., 1996). This suggests that kinesin-II may participate in spindle MT-based targeted secretion in association with cytokinesis, although no functional data support its mitotic role. Finally, the monomeric kinesin, UNC-104 is a well-characterized axonal pre-synaptic vesicle transport motor that has also been implicated in Golgi-associated membrane traffic (Dorner et al., 1998) raising the possibility that it might also be involved in targeted secretion during cell division, although in this case we are not aware of any evidence localizing the protein to mitotic spindles. Finally, it is possible that cytoplasmic dynein also drives vesicle transport in spindles, as it has been shown to play an important role in breaking down the nuclear envelope and transporting fragments of membrane along centrosomal MTs (Beaudouin et al., 2002). In summary, abundant evidence suggests that some intracellular transport motors are deployed in spindles where they play critical roles in cytokinesis by transporting vesicle and signaling molecules along spindle MTs. It is also clear that deciphering the precise roles and interactions of the complex network of motors that participate in these events will remain an active and fascinating research topic for some time.

14.9

Mitotic Motors and the Spindle Assembly Checkpoint

It has become clear that the force-generating and intracellular transport properties of mitotic motors are also used to regulate chromatid segregation during anaphase, by acting as components of the spindle assembly checkpoint (Shah and Cleveland, 2000). This checkpoint ensures faithful chromatid segregation by inhibiting anaphase onset until all the chromosomes in a mitotic cell are properly aligned in a bipolar configuration on the metaphase spindle equator. Prior to bipolar attachment, unattached kinetochores release a diffusible inhibitor of the cell cycle proteolysis machinery that initiates anaphase onset, but MT attachment to kinetochores and/or the development of tension across kinetochore pairs prevents the release of the soluble inhibitor, allowing the proteolysis machinery to degrade key substrates including the cohesins that ‘glue’ together sister chromatids, a chromokinesin which pushes chromosomes towards the spindle equator, and some bipolar kinesins which cross-link MTs throughout the spindle (Funabiki and Murray, 2000, Gordon and Roof, 2001, Hildebrandt and Hoyt, 2001, Shah and Cleveland, 2000). The degradation and inactivation of these substrates removes constraints on chromatid-to-pole motion allowing progression through anaphase A, and removes restraining cross-linkers thus permitting spindle elongation, disassembly and reassembly for subsequent mitosis. What is the nature of the soluble inhibitor and how is it inactivated? The outer kinetochore region contains binding sites for a complex set of proteins that are thought to participate in the spindle assembly checkpoint, including the kinesin, CENP-E, the dynein/dynactin complex, and several checkpoint proteins including MAD2, ZW10 and Rod (Shah et al., 2000). It is hypothesized that the soluble protease inhibitor is a fraction of MAD2 that binds the kinetochore where it is induced to

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oligomerize and is then released as an active diffusible oligomer capable of inhibiting the cell cycle proteolysis machinery. Mitotic motors are thought to contribute to the inactivation of this checkpoint in at least two ways. First, CENP-E is essential for the stable bi-oriented attachment of kinetochores to spindle MTs and, together with dynein (Howell et al., 2001), is involved in the development of tension across chromosomes once they are properly aligned; this CENP-E-dependent MT attachment and tension generation is proposed to block the formation of active, oligomeric MAD2 (Yao et al., 2000). Secondly, once kinetochores are attached to MTs and held under tension, the dynein/dynactin complex is proposed to deplete the checkpoint machinery from the kinetochores and thus to inactivate the checkpoint by actively transporting checkpoint proteins such as MAD2, ZW10 and Rod towards the minus ends of kinetochore MTs, from the kinetochores to the spindle poles where these proteins are observed to accumulate subsequent to metaphase (Howell et al., 2001, Wojcik et al., 2001). Thus these mitotic motors not only contribute directly to the physical alignment and segregation of chromosomes, but they also contribute to the regulation of these processes, thus ensuring the fidelity of mitosis.

14.10

Conclusions and Future Studies

Despite great progress in the field, there remain many outstanding questions about the roles of motor proteins and MT dynamics in mitosis. How are the force-generating properties of dynamic MTs and motor proteins coordinated? What is the role of the multiple redundant force-generating and velocity-governing systems? How are they integrated and regulated? What mechanisms insure the precise temporal and spatial morphogenesis of the mitotic spindle? What are the precise roles of mitotic motors and intracellular transport systems in the spindle? How do force-generating elements contribute to the spindle assembly checkpoint? Is there a spindle matrix? Both experimental and theoretical work is needed to answer these questions and dissect the process of mitosis. Visualizing spindle dynamics, inhibiting motors and imaging the effects on mitotic movements is one of the approaches currently being used, but other approaches will be needed, including better quantization of the mechanical and force-generating properties of spindles before and after genetic and/or biochemical manipulations of the spindle machinery. Additional biochemical approaches, including in vitro reconstitution from purified and characterized components will play important roles as well. Finally, as the field is maturing, complementation of these experimental approaches by theoretical modeling is becoming increasingly feasible and important.

Acknowledgements

We wish to thank Dr Ingrid Brust-Mascher for the help with the figures. Our work on mitosis is funded by NIH grant number GM 55507.


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Howard, J. 2001. Mechanics of Motor Proteins and the Cytoskeleton. Sunderland, MA: Sinauer. Howard, J., Hudspeth, A. J., and Vale, R. D. 1989. Movement of microtubules by single kinesin molecules. Nature 342: 154 158. Howell, B. J., McEwen, B. F., Canman, J. C., Hoffman, D. B., Farrar, E. M., Rieder, C. L. and Salmon, E. D. 2001. Cytoplasmic dynein/ dynactin drives kinetochore protein transport to the spindle poles and has a role in mitotic spindle checkpoint inactivation. J. Cell Biol. 155: 1159 1172. Hoyt, M. A. and Geiser, J. R. 1996. Genetic analysis of the mitotic spindle. Annu. Rev. Genet. 30: 7 33. Hunter, A. W. and Wordeman, L. 2000. How motor proteins influence microtubule polymerization dynamics. J Cell Sci. 113: 4379 4389. Inoue, S. and Salmon, E. D. 1995. Force generation by microtubule assembly/disassembly in mitosis and related movements. Mol. Biol. Cell. 6: 1619 1640. Kapoor, T. M. and Compton, D. A. 2002. Searching for the middle ground: mechanisms of chromosome alignment during mitosis. J. Cell Biol. 157: 551 556. Kapoor, T. M. and Mitchison, T. J. 2001. Eg5 is static in bipolar spindles relative to tubulin: evidence for a static spindle matrix. J. Cell Biol. 154: 1125 1133. Karabay, A. and Walker, R. A. 1999. Identification of microtubule binding sites in the Ncd tail domain. Biochemistry 38: 1838 1849. Karki, S. and Holzbaur, E. L. 1999. Cytoplasmic dynein and dynactin in cell division and intracellular transport. Curr. Opin. Cell Biol. 11: 45 53. Karsenti, E and Vernos, I. 2001. The mitotic spindle, a self-made machine. Science 294: 543 547. King, S. M. 2000. AAA domains and organization of the dynein motor unit. J. Cell Sci. 113: 2521 2526. Kuriyama, R., Gustus, C., Terada, Y., Uetake, Y. and Matuliene, J. 2002. CHO1, a mammalian kinesin-like protein interacts with F-actin and is involved in the terminal phase of cytokinesis. J. Cell Biol. 156: 783 790. Landau. L. D. and Lifshitz, E. M. 1995. Theory of Elasticity (3rd edn). Oxford: ButterworthHeinemann.

14 Mitotic Spindle Motors Le Bot, N., Antony, C. White, J. Karsenti, E. and Vernos, I. 1998. Role of Xklp3, a subunit of the Xenopus kinesin II heterotrimeric complex, in membrane transport between the endoplasmic reticulum and the Golgi appartatus. J. Cell Biol. 143: 1559 1573. Lee, Y.-R. and Liu, B. 2000. Identification of a phragmoplast-associated kinesin-related protein in higher plants. Curr. Biol. 10: 797 800. Lee Y.-R., Giang, H. M. and Liu, B. 2001. A novel plant kinesin-related protein specifically associates with the phragmoplast organelles. Plant Cell 13: 2427 2439. Lombillo, V. A., Nislow, C., Yen, T. J., Gelfand, V. I., and McIntosh, J. R. 1995b. Antibodies to the kinesin motor domain and CENP-E inhibit microtubule depolymerization-dependent motion of chromosomes in vitro. J. Cell Biol. 128: 107 115. Lombillo, V. A., Stewart, R. J., and McIntosh, J. R. 1995a. Minus-end-directed motion of kinesin-coated microspheres driven by microtubule depolymerization. Nature 373: 161 164. Ma, S., Trivinos-Lagos, L., Graf, R., and Chisholm, R. L. 1999. Dynein intermediate chain mediated dynein-dynactin interaction is required for interphase microtubule organization and centrosome replication and separation in Dictyostelium. J. Cell Biol. 147: 1261 1274. Maney, T., Ginkel, L. M., Hunter, A. W., and Wordeman, L. 2000. The kinetochore of higher eucaryotes: a molecular view. Int. Rev. Cytol. 194: 67 131. Marshall, W. F., Marko, J. F., Agard, D. A. and Sedat, J. W. 2001. Chromosome elasticity and mitotic polar ejection force measured in living Drosophila embryos by four-dimensional microscopy-based motion analysis. Curr. Biol. 11: 569 578. Merdes, A., Ramyar, K., Vechio, J. D. and Cleveland, D. W. 1996. A complex of NuMA and cytoplasmic dynein is essential for mitotic spindle assembly. Cell 87: 447 458. McDonald, H. B., Stewart, R. J. and Goldstein, L. S. 1990. The kinesin-like ncd protein of Drosophila is a minus end-directed microtubule motor. Cell 63: 1159 1165. McIntosh, J. R., Hepler, P. K. and Van Wie, D. G. 1969. Model for mitosis. Nature 224: 659 663. McIntosh, J. R. and McDonald, K. L. 1989. The mitotic spindle. Sci. Am. 261: 48 56.

Mishima, M., Kaitna, S. and Glotzer, M. 2002. Central spindle assembly and cytokinesis require a kinesin-like protein/RhoGAP complex with microtubule bundling activity. Dev. Cell. 2: 41 54. Mitchison, T. and Kirschner, M. 1984a. Dynamic instability of microtubule growth. Nature 312: 237 242. Mitchison, T. and Kirschner, M. 1984b. Microtubule assembly nucleated by isolated centrosomes. Nature 312: 232 237. Mitchison, T., Evans, L., Schulze, E. and Kirschner, M. 1986. Sites of microtubule assembly and disassembly in the mitotic spindle. Cell 23: 515 527. Mitchison, T. J. 1988. Microtubule dynamics and kinetochore function in mitosis. Annu. Rev. Cell Biol. 4: 527 549. Mitchison, T. J. 1989. Polewards microtubule flux in the mitotic spindle: evidence from photoactivation of fluorescence. J. Cell Biol. 109: 637 652. Mitchison T. J. and Sawin, K. E. 1990. MT flux in the mitotic spindle: where does it come from, where is it going? Cell Motil. Cytoskeleton 16: 93 98. Mitchison, T. J. and Salmon, E. D. 2001. Mitosis: a history of division. Nature Cell Biol. 3: E17 E21. Mogilner, A. and Oster, G. 1999. The polymerization ratchet model explains the forcevelocity relation for growing microtubules, Eur. Biophys. J. 28: 235 242. Mogilner, A., Elston, T., Wang, H.-Y., and Oster, G. 2002. Molecular motors: theory and examples. In Computational Cell Biology. Edited by C. P. Fall, E. Marland, J. Tyson and J. Wagner. New York: Springer, pp. 321 384. Nicklas, R. B. 1983. Measurements of the force produced by the mitotic spindle in anaphase. J. Cell Biol. 97: 542 548. Nicklas, R. B. and Ward, S. C. 1994. Elements of error correction in mitosis: microtubule capture, release, and tension. J. Cell Biol. 126: 1241 1253. Nishihama, R., Soyano, T., Ishikawa, M., Araki, S., Tanaka, H., Asada, T., Irie, K., Ito, M., Terada, M., Banno, H., Yamazaki, Y. and Machida, Y. 2002. Expansion of the cell plate in plant cytokinesis requires a kinesin-like protein/MAPKKK complex. Cell 109: 87 99. Nogales, E., Whittaker, M., Milligan, R. A., and Downing, K. H. 1999. High-resolution model of the microtubule. Cell 96: 79 88.

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crotubule nucleation. Curr. Opin. Cell Biol. 12: 113 118. Scholey, J. M., Rogers, G. C. and Sharp, D. J. 2001. Mitosis, microtubules and the matrix. J. Cell Biol. 154: 261 266. Schroer, T. A. 2001. Microtubules don and doff their caps: dynamic attachments at plus and minus ends. Curr. Opin. Cell Biol. 13: 92 96. Severson, A. F., Hamill D. R., Carter, J. C., Schumacher, J. and Bowerman, B. 2000. The aurora-related kinase, AIR-2 recruits ZEN-4/ CeMKLP1 to the mitotic spindle at metaphase and is required for cytokinesis. Curr. Biol. 10: 1162 1171. Shah, J. V. and Cleveland, D. W. 2000. Waiting for anaphase: Mad2 and the spindle assembly checkpoint. Cell 103: 997 1000. Sharp, D. J., McDonald, K. L., Brown, H. M., Matthies, H. J., Walczak, C., Vale, R. D., Mitchison, T. J., and Scholey, J. M. 1999. The bipolar kinesin, KLP61F, cross-links microtubules within interpolar microtubule bundles of Drosophila embryonic mitotic spindles. J. Cell Biol. 144: 125 138. Sharp, D. J., Rogers, G. C., and Scholey, J. M. 2000a. Microtubule motors in mitosis. Nature 407: 41 47. Sharp, D. J., Rogers, G. C., and Scholey, J. M. 2000b. Cytoplasmic dynein is required for poleward chromosome movement during mitosis in Drosophila embryos. Nature Cell Biol. 2: 922 930. Shimizu, K., Kawabe, H., Minami, S., Honda, T., Takaishi, K., Shirataki, H., and Takai, Y. 1996. SMAP, an Smg GDS-interacting protein having arm repeats and phosphorylated by Src tyrosine kinase. J. Biol. Chem. 271: 27013 27017. Shuster, C. B. and Burgess, D. R. 2002. Targeted new membrane addition in the cleavage furrow is a late separate event in cytokinesis. Proc Natl. Acad. Sci. USA 99: 3633 3638. Skop, A. R., Bergmann, D., Mohler, W. A. and White, J. G. 2001. Completion of cytokinesis in C. elegans requires brefeldin A-sensitive membrane accumulation at the cleavage furrow apex. Curr. Biol. 11: 735 746. Vale, R. D. and Fletterick, R. J. 1997. The design plan of kinesin motors. Annu. Rev. Cell. Dev. Biol. 13: 745 777. Vale, R. D., Malik, F. and Brown, D. 1992. Directional instability of microtubule transport in the presence of kinesin and dynein, two

14 Mitotic Spindle Motors opposite polarity motor proteins. J. Cell Biol. 119: 1589 1596. van Doorn, G. S., Tanase, C., Mulder, B. M., and Dogterom, M. 2000. On the stall force for growing microtubules. Eur. Biophys. J. 29: 2 6. van Hemert, M. J., Lamers, G. E., Klein, D. C., Oosterkamp, T. H., Steensma, H. Y., and van Heusden, G. P. 2002. The Saccharomyces cerevisiae Fin1 protein forms cell cycle-specific filaments between spindle pole bodies. Proc. Natl. Acad. Sci. USA 99: 5390 5393. Visscher, K., Schnitzer, M. J., and Block, S. M. 1999. Single kinesin molecules studied with a molecular force clamp. Nature 400: 184 189. Walker, D. L., Wang, D., Jin, Y., Rath, U., Wang, Y., Johansen, J. and Johansen, K. M. 2000. Skeletor, a novel chromosomal protein that redistributes during mitosis, provides evidence for a spindle matrix. J. Cell Biol. 151: 1401 1411. Wells, W. A. 2001. Searching for a spindle matrix. J. Cell Biol. 154: 1102 1104. Williams, B. C., Riedy, M. F., Williams, E. V., Gatti, M. and Goldberg, M. L. 1995. The Drosophila kinesin-like protein, KLP3A is a midbody component required for central spindle assembly and initiation of cytokinesis. J. Cell Biol. 129: 709 723. Wittman, T., Hyman, A., and Desai, A. 2001. The spindle, a dynamic assembly of micro-

tubules and motors. Nature Cell Biol. 3: E28 E34. Wojcik, E., Basto, R., Serr, M., Scaerou, F., Karess, R., and Hays, T. 2001. Kinetochore dynein: its dynamics and role in the transport of the Rough Deal checkpoint protein. Nature Cell Biol. 3: 1001 1007. Wright, B. D., Henson, J. H., Wedaman, K. P., Willy, P. J., Morand, J. N., and Scholey, J. M. 1991. Subcellular localization and sequence of sea urchin kinesin heavy chain: evidence for its association with membranes in the mitotic apparatus and interphase cytoplasm. J. Cell Biol. 113: 817 833. Wright, B. D., Terasaki, M. and Scholey, J. M. 1993. Roles of kinesin and kinesin-like proteins in sea urchin embryonic cell division: evaluation using antibody microinjection. J. Cell Biol. 123: 681 689. Yao, X., Abrieu, A., Zheng, Y., Sullivan, K. F., and Cleveland, D. W. 2000. CENP-E forms a link between attachment of spindle microtubules to kinetochores and the mitotic checkpoint. Nature Cell Biol. 2: 484 491. Yucel, J. K., Marszalek, J. D., McIntosh, J. R., Goldstein, L. S. B., Cleveland, D. W. and Philip, A. V. 2000. CENP-meta, an essential kinetochore kinesin required for the maintenance of metaphase chromosome alignment in Drosophila. J. Cell Biol. 150: 1 12.

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15 The Roles of Molecular Motors in Generating Developmental Asymmetry Janice A. Fischer

15.1

Introduction

Specific localization of mRNAs, proteins and organelles is a widely used mechanism for generating asymmetric developmental cues or asymmetric structures within a cell. The idea that molecular motor proteins may transport molecules or organelles critical to development is exciting because it opens up a whole world of regulatory possibilities. Localization of factors by motors could, in theory, be regulated by the differential expression or post-translational modification of motor proteins, the adaptors that link them to specific cargo, or by changes in cytoskeletal organization. In this chapter, I concentrate on a few developmental processes which clearly depend on motor proteins. In the examples presented here, the motors appear to play a wide variety of different roles in generating asymmetry, and the apparent complexity of the mechanisms and regulation of these processes is only just beginning to emerge.

15.2

Localization of a L/R Determinant by Asymmetric Flow of Extraembryonic Fluid 15.2.1

Situs Inversus in Humans

Humans, like all vertebrates, are normally L/R asymmetrical; for example, the heart, spleen, and pancreas are on the left side, while most of the liver and the gallbladder are on the right. In addition, the left lung has two lobes, while the right lung has three. What patterns the L/R axis? People with a rare disease Kartagener’s syndrome provided the first clue that the answer might have some connection with cilia that are powered by axonemal dynein. Kartagener’s syndrome is a form of primary ciliary dyskinesia (PCD), a disease whose symptoms are chronic bronchial and sinus infections, and infertility (Afze-

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15.2 Localization of a L/R Determinant by Asymmetric Flow of Extraembryonic Fluid

lius, 1976, 1999, Afzelius and Mossberg, 1995, Aylsworth, 2001, Kosaki and Casey, 1998). At the microscopic level, PCD is caused by the immotility of cilia in the respiratory passages and the immotility of sperm flagella. PCD is inherited as an autosomal recessive disease, and PCD alleles have been mapped to several different chromosomes, meaning that homozygotes for mutations in any one of a large number of genes result in PCD. Recently, two mutant genes which can cause PCD have been identified: DNAH5 encodes an axonemal dynein heavy chain gene (Olbrich et al., 2002, Omran et al., 2000), and DNAI1, an axonemal dynein intermediate chain gene (Guichardet et al., 2001, Pennarum et al., 2000, Zariwala et al., 2001). While cytoplasmic dynein moves on microtubule tracks, axonemal dynein works with microtubules to power cilia and flagella. About half of the PCD patients are described as having Kartagener’s syndrome; in addition to the other PCD symptoms, these people display inversion of L/R asymmetry, a condition known as situs inversus. Therefore, determination of which side is right and which is left is randomized in people with PCD. What is the connection between axonemal dynein and determining rightness versus leftness? 15.2.2

Mice with Mutations in kif3a, kif3b, or lrd Lead to Nodal Flow Model

Molecular genetic analysis of mutant mice with randomized L/R axes, followed by a series of elegant and technically impressive experiments with mutant embryos, has led to a model where the beating of cilia on a structure in the early mouse embryo, called the node, determines leftness and rightness. A randomized L/R axis in mice was originally found to be associated with three different genes: left right dynein (lrd), which encodes an axonemal dynein heavy chain (Hummel and Chapman, 1959, Supp et al., 1999), and kif3a and kif3b, which encode subunits of the kinesin superfamily 3 (KIF3) kinesin motor (Marszalek et al., 1999, Nonaka et al., 1998, Takeda et al., 1999). As KIF3 is required for ciliogenesis, kif3a or kif3b knock-out mice lack cilia. By contrast to people with PCD, however, lrd mutant mice have normal respiratory cilia, but lack cilia on a structure present in neurula-stage mouse embryos called the node. The node is a cup-shaped cavity at the anterior tip of the primitive streak, which gives bilateral symmetry and a midline axis to the embryo. The node is equivalent functionally to Spemann’s organizer of Xenopus, Hensen’s node in the chick, and the embryonic shield of zebrafish; these structures are called organizers because they can induce a second midline axis if grafted to a new location in the embryo. Cells on the ventral side of the mouse node each have a cilium that projects into the extraembryonic space and rotates clockwise rapidly (Z 600 r. p. m.) (Nonaka et al., 1998, Okada et al., 1999, Takeda et al., 1999). As the results of many experiments have implicated the node in L/R axis determination (Levin et al., 1995, 1997), it was hypothesized that the beating of nodal cilia could generate directional flow of a leftness or rightness determinant to one side of the embryo. To test the ‘nodal flow’ hypothesis, flow of extraembryonic fluid across the node was measured in wild-type and mutant mouse embryos using fluorescent tracer

15 The Roles of Molecular Motors in Generating Developmental Asymmetry

The nodal flow model for left/right determination in mice. At the top is a cartoon of the mouse embryonic node. The clockwise beating of cilia results in a net flow of extraembryonic fluid leftwards, which may result in the accumulation of an unidentified leftness determinant, which flows into the node symmetrically, on the left side of the node. The leftness determinant would activate the leftness pathway genes. At bottom is a diagram of single nodal cilium, which is powered by axonemal dynein. After Hamada et al., 2002.

Figure 15.1.

beads (Nonaka et al., 1998 Okada et al., 1999, Takeda et al., 1999). In wild-type embryos, the extraembryonic fluid was found to flow leftwards, while there was no nodal flow at all in lrd or kif3 mutants. These observations led to a variety of more specific versions of the nodal flow model for L/R axis determination; in all of them, the symmetry-breaking event is the clockwise beating of the nodal cilia. A simple version of the model (Nonaka et al., 1998, Okada et al., 1999) is shown in Fig. 15.1. A left-side determinant (its identity is presently unknown), released into the node uniformly, is swept leftward, where it binds to a receptor present ubiquitously in the node, and activates downstream left-side determinants. (Several genes required for left right axis determination that are expressed only on the left side of the embryo have been identified and organized into a pathway (Burdine and Schier, 2000, Capdevila et al., 2000, Hamada et al., 2002, Ramsdell and Yost, 1998)). According to this model, L/R determination is randomized in lrd and kif mutants because, in the absence of leftward flow, the leftness determinant diffuses randomly, and thus activates receptors randomly on one side or the other (which would result in normal sidedness or situs inversus), on both sides, or neither side. Leftness receptor activation on both sides or neither side would be expected to result in leftness isomerism, or rightness isomerism, respectively. These conditions, as well as the in-between condition of heterotaxia or situs ambiguous in which some organs have normal L/R positioning and others are reversed, have been observed in some people with PCD (although rarely) (Aylsworth, 2001, Kosaki and Casey, 1998), and also frequently in lrd mutant mice (Hummel and Chapman, 1959, Supp et al., 1999). In addition, alteration of the expression patterns of the

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downstream ‘leftness’ pathway genes in the mutant backgrounds are consistent with the above model. 15.2.3

Inv Mutants Challenge the Nodal Flow Model

The nodal flow hypothesis generates many questions. Notable among them is how to explain the phenotype of mouse inversion of embryonic turning (inv) mutants; mice homozygous for inv knock-out mutations are always L/R reversed (Yokoyama et al., 1993). The inv gene encodes the ankyrin repeat protein Inversin, whose function is unknown (Mochizuki et al., 1998, Morgan et al., 1998). The cilia of inv knock-out mice appear normal morphologically, but the leftward nodal flow in inv mutants is slower and more turbulent than normal (Okada et al., 1999). Okada et al. (1999) discuss several possible explanations for the inv mutant phenotype. One possibility assumes that an inactive Factor X flows into the node from both the right and left sides (as opposed to secretion by the node cells themselves), and becomes active a few seconds later. According to this model, Factor X does not normally activate the receptors on the right side, because leftward nodal flow has already swept most of it to the left before it is activated. In inv mutants, the Factor X that comes in from the left is effectively swept out of the node, but most of the Factor X that comes in from the right side remains there when Factor X is activated. A second possibility is that Inversin functions downstream of receptor activation by Factor X, and that in inv mutants, the response to receptor activation is reversed. In this case, the nodal flow defects could be caused by a secondary function of Inversin that is inconsequential for L/R determination. Alternatively, the inv phenotype could indicate that nodal cilia function is really downstream of the symmetry-breaking event; if Factor X flows in from the right, slower leftward flow in inv mutants could result in receptor activation on the right side. Another challenge to the idea that nodal flow is the symmetry-breaking event is the observation that the effects of mutants that abolish nodal flow (lrd, kif3a, and kif3b) on the expression of downstream ‘leftness’ determination genes seem not to be identical (Wagner and Yost, 2000). One plausible explanation takes into account the presence of immotile cilia in lrd mutants, as opposed to the complete absence of cilia in kif mutants, but there may even be differences in downstream gene expression between kif3a and kif3b mutants. Alternative possibilities are that background differences in the mutants, differences in experimenters, and/or the small numbers of animals analyzed contribute to the apparent discrepancies in the phenotypes. Along similar lines, it is also unclear why people with PCD, who have immotile cilia, almost always have normal L/R axes or else have a completely reversed L/R axis (situs inversus), as opposed to the range of isomerisms and heterotaxias observed in mouse lrd mutants. Perhaps the organ malformations often associated with isomerism and heterotaxia, coupled with PCD, are usually lethal conditions. Further supporting the importance of the node in L/R axis determination, mice with knock-out mutations in other genes that result in the absence of nodal cilia


(i. e. Polaris, which encodes a protein of unknown function), or the absence of the node (i. e. Hnf3; hepatocyte nuclear factor 3, which encodes a transcription factor), display L/R axis defects (Brody et al., 1999, Murcia et al., 2000). Interestingly, Hnf4 (hepatocyte nuclear factor 4) mutants have L/R defects even though nodal cilia are present (Chen et al., 1998); the function of the cilia remains to be tested. Additional experiments are clearly required to rule out the idea that nodal flow is a red herring and that kinesin and dynein are actually affecting L/R axis formation through functions within the cell cytoplasm. If nodal flow is the symmetry-breaking event, it is curious that this mechanism is unlikely to be conserved in many other vertebrates (Burdine and Schier, 2000, Capdevila et al., 2000, Wright, 2001). For example, Henson’s node in the chick appears to have sparse cilia, and the Spemann’s organizer in Xenopus has none. It would be interesting to know whether axonemal dynein is required for L/R asymmetry in these organisms; if so, this would support a cytoplasmic role for these motors. Another way to resolve this issue would be to manipulate nodal flow mechanically and look at L/R effects. Finally, identification of Factor X will also help in determining whether nodal flow is superfluous, is the symmetry-breaking event, or maintains a prior asymmetry.

15.3

Asymmetric RNA Localization

Specific mRNAs are localized within a cell as a means of localizing their protein products. Two of the best-studied examples where mRNA localization controls developmental fate patterning along the Drosophila A/P axis, and determination of mating type in Saccharomyces cerevisiae are described here. Molecular motors were presumed initially to be involved with mRNA localization because, using chemical inhibitors, it was shown that microtubule and/or actin tracks are used to transport the mRNAs from the nucleus to their destinations. More recently, motors have been shown directly to be required for mRNA localization; genetic studies have shown a requirement for particular motors in the localization of specific mRNAs, and motors have been isolated biochemically in complexes either with mRNAs or with proteins known to be required for mRNA localization. In most cases, however, it is yet unclear whether the motor connected with a particular localization process actually transports the mRNAs itself, transports materials needed to anchor the mRNA, or acts directly as an anchor for the mRNA once it reaches its destination. 15.3.1

A/P Patterning in Drosophila Oocytes and Embryos

The Drosophila oocyte is connected by cytoplasmic bridges to 15 nurse cells that supply it with mRNAs and proteins. Included among the nurse cell mRNAs that are transported into the oocyte are bicoid (bcd) and oskar (osk), which determine the anterior and posterior poles of the oocyte, respectively. Much attention has

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15.3 Asymmetric RNA Localization

Figure 15.2. A model for localization of bicoid and oskar mRNAs in the Drosophila oocyte. In the oocyte/nurse cell complex, the bcd and osk mRNAs are localized to and define the anterior and posterior poles of the oocyte, respectively. In the nurse cells, Exu-containing protein complexes, probably linked to microtubule motors, transport bcd and osk mRNAs to the oocyte. In the oocyte, an unknown sorting mechanism is thought to link an Exu and bcd-containing particle to dynein for transport to the anterior, and a Stau and osk-containing particle to kinesin for transport to the posterior pole of the oocyte. Subsequent anchoring of bcd and osk to the poles requires Stau and then Swa; Swa has been shown to bind dynein directly (Stau has not been shown to bind osk mRNA or a molecular motor directly).

been focused on how bcd and osk mRNAs are transported to their respective positions at the anterior and posterior of the oocyte (Fig. 15.2).

Anterior localization of bcd mRNA in the oocyte Localization of bcd mRNA to the anterior cortex of the oocyte can be thought of as three processes: transport of bcd from the nurse cell to the oocyte, transport to the anterior cortex within the the oocyte, and then anchoring of bcd mRNA there. Three genes have been shown to be required for bcd localization: exuperantia (exu), swallow (swa), and staufen (stau) (Berleth et al., 1988, St Johnston et al., 1989). In swa mutants, bcd mRNA enters the oocyte but fails to localize anteriorly. The exu gene is required even earlier than swa; in exu mutants, bcd mRNA moves aberrantly in the nurse cells, but manages to make it into the oocyte, where it also fails to localize to the anterior cortex. The stau gene is required later for anchoring bcd mRNA. The results of several experiments described below suggest that the three bcd localization processes may all involve dynein motors. Of the three proteins, only Stau has been shown to be an RNA-binding protein (Ferrandon et al., 1994), and only Swa has been shown to be associated with dynein directly (Schnorrer et al., 2000). 15.3.1.1


Exu protein is part of a transport complex that may link bcd mRNA to a motor

Exu-containing particles were first observed in living egg chambers of flies expressing GFP-Exu (Theurkauf and Hazelrigg, 1998). Movements of the GFP-Exu particles in nurse cells reflect the microtubule organization there, and particle movement is microtubule-dependent, both of which suggest that it is motor driven. The GFP-Exu particles are transported into the oocyte and accumulate at the oocyte anterior cortex (Cha et al., 2001). More recently, the results of experiments in which fluorescent bcd mRNA was injected into egg chambers suggest that an Exu-containing particle enables bcd mRNA to move into the oocyte, and once there, to travel specifically along the microtubules that extend from the anterior cortex (Cha et al., 2001). Fluorescent bcd mRNA injected into the middle of the oocyte is transported to the closest cortical surface in a microtubule-dependent manner. However, if the bcd mRNA is first injected into exu nurse cell cytoplasm, removed, and then injected into the middle of the oocyte, it goes to the anterior, also dependent on microtubules. What motor might be moving the Exu particles? Originally, it was thought that at the time when bcd mRNA is localized anteriorly in the oocyte, the majority of the microtubules were arranged with their minus ends anterior and their plus ends posterior. This led to a simple model where bcd mRNA would be transported by dynein to the anterior. This idea was based largely on the observations of two markers for microtubule polarity: a Nodbgalactosidase (Nod-bgal) fusion protein, and a kinesin-bgalactosidase (kinesinbgal) fusion protein (Clark et al., 1994, 1997). When expressed in the oocyte by transgenes, Nod-bgal accumulates at the anterior cortex, and kinesin-bgal at the posterior. However, using confocal imaging to visualize tubulin, Cha et al. (2001) showed that while the majority of microtubules extend from the anterior cortex in stage 9/10a oocytes, there are also many microtubules extending laterally from the oocyte cortex into the oocyte. The overall conclusion is that the bulk of microtubules in the middle of the oocyte are not asymmetric enough to specify the anterior pole (Fig. 15.2). The results of the bcd injection experiments suggest that something happens to bcd mRNA in the nurse cell cytoplasm, which is exu-dependent, and which allows bcd to be transported anteriorly on a grossly non-polar jungle of microtubules in the oocyte; bcd mRNA is confronted with this network when injected into the middle of the oocyte, and presumably also when it enters the oocyte normally from the nurse cells. The authors propose that Exu forms a complex with bcd and one or more anterior localization factors present in the nurse cells. The anterior localization factor would enable the bcd-containing particle to travel only on the microtubules emanating from the anterior cortex. This subpopulation of microtubules must somehow be functionally distinct. Presumably, the Nod-bgal protein also recognizes only this microtubule subpopulation, and the kinesin-bgal protein sees a different subpopulation of microtubules extending from the posterior. It is quite possible that the anterior localization factor, or one component of it, could be a motor. An Exu-containing particle has now been purified biochemically from ovaries; it contains at least seven other proteins and an mRNA which is not bcd, but osk

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(Wilhelm et al., 2000). One of the seven proteins is the RNA-binding protein Ypsilon Schachtel (YpS), which may bind directly to osk mRNA (see below for a discussion of osk localization). The authors propose that Exu is a core component of a large protein complex that localizes mRNAs both within nurse cells and the oocyte (Fig. 15.2). Sorting of the complexes to anterior and posterior destinations would occur at the anterior of the oocyte. The Exu complexes isolated could well be heterogeneous. It will be interesting to know whether any contain motors, and where the various proteins are located in the oocyte/nurse cell complex and where required to get at the mechanism of sorting. Swa protein may link bcd mRNA to dynein for anchoring

Swa normally localizes to the anterior pole independently of Exu or bcd mRNA, and its localization depends on the presence of microtubules, and on their polarity (Schnorrer et al., 2000). In gurken mutants, where microtubule organization is aberrant, Swa protein is found with bcd mRNA (and Nod-bgal protein) at both anterior and posterior poles. Swa has a coiled-coil domain essential to its function and anterior localization; the coiled-coil domain binds to dynein light chain (encoded by Ddlc-1) in vitro and in vivo. It is unlikely, however, that bcd uses Swa for transport to the anterior pole; in the experiments of Cha et al. (2001) described above, fluorescent bcd mRNA injected into swa oocytes localizes to the oocyte cortex. Swa is probably involved in anchoring bcd mRNA, via dynein, to the anterior cortex, prior to the anchoring function of Stau protein (Fig. 15.2). In order to understand the significance of the Swa/Ddlc-1 interaction, it will be important to determine genetically whether or not dynein is required for bcd anterior localization. These experiments are complicated by the requirement for dynein in early stages of oogenesis; in ovaries homozygous for strong loss-of-function mutations in the dynein heavy chain (Dhc64C) gene, oocytes do not differentiate (McGrail and Hays, 1997). Oocyte/nurse cell complexes labeled with antiDhc64C antibodies show that most of the Dhc64C is at the posterior pole at the time when Exu, Swa and bcd mRNA are at the anterior pole (McGrail and Hays, 1997). This result seems difficult to reconcile not only with the accumulation of kinesin-bgal at the posterior pole and Nod-bgal at the anterior (Clark et al., 1994, 1997), but also with the association between Swa and Ddlc-1 (Schnorrer et al., 2000). Perhaps there are two distinct dynein complexes at each pole, only one of which contains Dhc64C.

Posterior localization of osk mRNA within the oocyte by kinesin Kinesin has been shown definitively to be required for the localization of osk mRNA to the posterior cortex of the oocyte (Brendza et al., 2000). In female flies with ovaries homozygous for loss-of-function mutations in a kinesin heavy chain (Khc) gene, osk mRNA and Stau protein accumulate at the anterior and never migrate to the posterior pole (Stau co-localizes with and probably binds to osk in addition to bcd, and is required for osk posterior localization (Ephrussi et al., 1991, Kim-Ha et al., 1991, St Johnston et al., 1991)). This effect is unlikely to be due 15.3.1.2


to general disruption of oocyte microtubule organization in Khc ovaries; bcd mRNA is localized normally at the anterior, the oocyte nucleus migrates normally (see Section 15.4.2 below), the MTOC is normally located, and kinesin-bgal is localized posteriorly. A simple model for the role of kinesin is that it transports an osk/Stau complex to the posterior pole (Fig. 15.2). However, this is likely to be an oversimplification. First, fluorescently-labeled osk mRNA can localize to the posterior pole in colchicine-treated ooctyes, as long as it is injected near the posterior, suggesting that there is a microtubule-independent anchoring system at the posterior for osk (Glotzer et al., 1997). In addition, osk posterior localization is blocked in tropomyosin II mutants (Erdelyi et al., 1995). Non-muscle tropomyosins are actin-binding proteins that stabilize actin filaments and regulate movement of myosin motors along them. Perhaps kinesin transports osk part of the way to the posterior pole, and transfers the osk particle to a myosin motor; this idea is also supported by the observation that microtubules at the posterior do not extend all the way to the pole (Cha et al., 2001). A completely different possibility suggested by Brendza et al. (2000) is that kinesin does not directly transport osk mRNAs, but could deliver materials needed for assembly of an osk/Stau complex prior to stage 7/8, when the plus ends of microtubules are anterior.

Localization of mRNAs in blastoderm embryos Transcripts for some pair-rule genes (fushi tarazu (ftz) and runt) and also the segment polarity gene wingless (wg) are localized apically in the blastoderm embryo (Davis and Ish-Horowicz, 1991, Lall et al., 1999, Simmonds et al., 2001). Pairrule genes are expressed periodically in stripes along the A/P axis; they encode transcription factors that divide the A/P axis into segments. Wg is a diffusible morphogen that determines A/P patterning within each segment. Apical localization of these mRNAs depends on microtubules, dynein, and the 3l-UTRs of the mRNAs (Davis and Ish-Horowicz, 1991, Lall et al., 1999, Simmonds et al., 2001, Wilkie and Davis, 2001). In the case of wg, apical mRNA localization has been shown to be required for wg function (Simmonds et al., 2001). Experiments where localized mRNAs were expressed ectopically, either in the oocyte or the blastoderm embryo, show that nurse cell-to-oocyte transport and apical localization of mRNAs in the zygote occur by a common mechanism (Bullock and Ish-Horowicz, 2001). Moreover, two proteins, Egalitarian (Egl) and Bicaudal D (BicD), are required for both processes, and may link dynein to mRNAs (Bullock and Ish-Horowicz, 2001, Mach and Lehmann, 1997, Ran et al., 1994, Wilkie and Davis, 2001). 15.3.1.3

Dynein is required for apical mRNA localization in blastoderm embryos

Apical localization of mRNAs is thought to occur by active transport because nascent mRNAs for runt, ftz, and wg, which are apically localized, as well as other mRNAs that are destined to be localized basally or unlocalized, are all observed at all sides of the nuclear periphery. At some point, the mRNAs are sorted into localization complexes. Injected runt, ftz, or wg mRNAs localize apically in particles

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as do endogenous mRNAs; each particle may contain several different mRNAs which are destined for the apical membrane, but not unlocalized or basally-destined mRNAs (Wilkie and Davis, 2001). As microtubules are oriented with their minus ends apical in blastoderm embryos (Foe et al., 1993), a simple model would have dynein transport the pairrule and wg mRNAs to the apical membrane. Microtubules are required, as colcemid injected prior to mRNA injection blocks apical localization (Wilkie and Davis, 2001). Moreover, several lines of evidence indicate that dynein is required for apical mRNA localization (Wilkie and Davis, 2001). First, injection of anti-Dhc or dynamitin protein (dynamitin is a dynactin subunit; dynamitin over-expresion has a dominant negative effect) inhibits apical transport of injected runt, ftz, and wg mRNAs. Second, homozygotes for hypomorphic Dhc64C alleles survive through blastoderm and mRNAs injected into these embryos move more slowly than in wild-type embryos. Moreover, further decreasing dynein function in the hypomorphic embryos by injection of anti-Dhc inhibits apical localization of endogenous ftz. These experiments provide the first clear demonstration that dynein is required for mRNA localization. An Egl/BicD complex may link mRNAs to dynein

An Egl/BicD complex is required for oocyte differention and transport of particular mRNAs into the oocyte (Mach and Lehmann, 1997, Ran et al., 1994). Both proteins are supplied maternally to embryos, and in blastoderm, they accumulate at the minus ends of microtubules (Mach and Lehmann, 1997, Oh and Steward, 2001). When injected into blastoderm embryos, mRNAs like bcd that are normally transported from the nurse cells into the oocyte, localize apically. Conversely, injection into oocytes or ectopic expression from transgenes of pair-rule mRNAs (ftz, runt, or hairy) which are normally apically localized in blastoderm embryos, results in their transport into the oocyte. As mutant mRNAs behave identically in the two assays, oocyte-to-nurse cell transport and apical localization in blastoderm are analogous processes, both of which require Egl and BicD (Bullock and Ish-Horowicz, 2001). The similar distribution of the BicD/Egl complex and Dhc during oogenesis and in blastoderm embryos suggests that BicD/Egl links mRNAs to dynein (Swan et al., 1999), but no direct interaction between BicD or Egl and dynein has yet been detected. 15.3.2

Yeast Mating Type Switching

In yeast (S. cerevisiae), daughter cells (buds) exclusively accumulate Ash1p, which prevents mating type switching by repressing transcription of the HO endonuclease gene (Bobola et al., 1996, Jansen et al., 1996, Long et al., 1997, Munchow et al., 1999, Sil and Herskowitz, 1996, Takizawa et al., 1997). A myosin motor, called Myo4p or She1p, has been shown to transport an RNP containing Ash1 mRNA from the mother cell to the bud tip along actin filaments (Fig. 15.3). Transport of Ash1 mRNA to the bud tip is the only clear example of a myosin motor

15 The Roles of Molecular Motors in Generating Developmental Asymmetry Ash1 mRNA localization to the bud tip in yeast. Ash1 mRNA is transported to the bud tip, where it controls the mating type of the daughter cell. In the nucleus, Loc1p identifies Ash1 mRNA for binding to She2p, and She3p acts as adapter, linking the myosin motor Myo4p to She2p/Ash1. Myo4p then walks along actin cables that extend from the mother cell to the bud tip.

Figure 15.3.

transporting a localized mRNA, and also provides the very first demonstration of an association between an mRNA and a motor protein. Six proteins, She1p She5p and Loc1p, are known to be required for Ash1 mRNA localization; She1p is Myo4p, She2p binds Ash1 mRNA, She3p is an adapter that links She2p to Myo4p, She4p and She5p (a. k. a. Bni1p or Bud6p) regulate the actin cytoskeleton, and Loc1p is a nuclear protein that binds Ash1 mRNA and may direct it towards the bud tip localization machinery (Beach and Bloom, 2001, Bohl et al., 2000, Long et al., 2000, 2001, Takizawa and Vale, 2000). The She proteins were first identified genetically. To determine if the She1p/ Myo4p motor actually transports Ash1 mRNA, the movements of a fluorescent Ash1 mRNA were analyzed (Beach et al., 1999, Bertrand et al., 1998). The Ash1 mRNA was tagged with GFP by expressing two different gene fusions in yeast: one which produces a fusion protein of MS2, a bacteriophage coat protein, to GFP, and the other of which produces an Ash1 mRNA containing MS2-binding sites. Three observations suggest that Myo4p (She1p) associates with a particle that contains Ash1 mRNAs. First, Myo4p (She1p) and the GFP-tagged Ash1 co-localize at the bud tip and in the non-budding cell. Second, the GFP particles do not move in a she1 point mutant and assemble less efficiently in a she1 deletion mutant that makes no protein. Finally, the speed of the GFP particles is similar to that expected for myosin V motors. Further experiments demonstrated that Ash1 mRNA co-immunoprecipitates specifically with Myo4p (Munchow et al., 1999). Many genetic and biochemical experiments have led to a model where She3p functions as an adapter, linking Ash1 mRNA, via She2p, to Myo4p (Bohl et al., 2000, Kwon and Schnapp, 2001, Long et al., 2000, Takizawa and Vale, 2000; Fig. 15.3). GFP particle formation and the Myo4p/Ash1 mRNA association are both dependent on she2 and she3 gene functions. She2p binds specifically to the regions of Ash1 mRNA shown to be required for its localization, and mutant Ash1 mRNAs that do not localize also do not bind efficiently to She2p. There is much evidence for an adaptor function for She3p. First, She3p binds to She2p via its C-terminus and to Myo4p via its N-terminus. Second, Ash1 mRNA co-immunoprecipiates with She3p and Myo4p in wild-type, but not in she2 yeast, and while the Ash1/Myo4p interaction requires She3p, the She3p/Ash1 interaction does not require Myo4p. Finally, the requirement for She2p in localization to the bud tip of a reporter mRNA containing MS2-binding sites, can be circumvented if She3p is given an MS2 RNA-binding domain.

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15.4 Asymmetric Organelle Localization

15.4

Asymmetric Organelle Localization 15.4.1

Localization of the Fusome and Drosophila Oocyte Selection

In Drosophila, an oocyte is chosen from among 16 interconnected cells formed by incomplete cytokinesis of dividing cystoblasts; the other 15 cells become nurse cells, which provide the oocyte with mRNAs and proteins (Reichmann and Ephrussi, 2001). Oocyte selection certainly requires dynein, as Dhc64C mutants have 16 nurse cells and no oocyte (McGrail and Hays, 1997). Dynein is required for several processes in early oogenesis, and as they are all interdependent, it is unclear which one is of primary importance for oocyte selection (Theurkauf, 1997). Unequal distribution of an organelle called the fusome, is the first apparent asymmetry in the cystoblast, and fusome integrity and localization requires dynein. The fusome is a single vesicular organelle that branches through all of the cell junctions, or ring canals, in the cyst. Dynein mediates the stable association of the fusome with microtubules during cystoblast divisions (deCuevas and Spradling, 1998, McGrail and Hays, 1997, Spradling, 1993). As one pole of the mitotic spindle is anchored in the fusome, the position of the fusome determines spindle pole orientation during cystoblast divisions, and thereby determines the pattern of connections between the cystoblast cells (Deng and Lin, 1997). The fusome also organizes the microtubule network in the oocyte/nurse cell complex (Grieder et al., 2000), which may be necessary for transport of determination factors into the oocyte; dynein may also be required directly for transport of these factors. Mutations in three additional genes, BicD, egl, and DLis-1 (the Drosophila homolog of the human Lissencephaly gene (see below)), also result in 16 nurse cells and no oocyte (Lei and Warrior, 2000, Liu et al., 1999, Mach and Lehmann, 1997, Swan et al., 1999). The DLis-1 protein contains an N-terminal coiled-coil domain and seven WD40 repeats. DLis-1 appears to be a regulator of dynein; in the oocyte, dynein localization depends on DLis-1 (Swan et al., 1999). Thus, DLis-1 may, with dynein, regulate fusome integrity. BicD and Egl are required to organize the microtubule network in the cystoblast, and are also among the first proteins transported into the oocyte (Mach and Lehmann, 1997, Suter et al., 1989, Theurkauf et al., 1993, Wharton and Struhl, 1989). It is not known whether BicD and Egl work directly with dynein in this context. 15.4.2

Drosophila Oocyte Nuclear Migration

In the Drosophila oocyte, the position of the nucleus determines both the A/P and D/V embryonic axes through transcription of gurken within the oocyte nucleus (van Eeden and St Johnston, 1999). The oocyte nucleus is posterior at first, where it signals the posterior follicle cells surrounding the oocyte. This signal defines the A/P axis and initiates a major reorganization of the oocyte cytoskeleton, such that the


minus ends of the microtubules are anterior (kinesin-bgal) and the plus ends (Nodbgal) are posterior. The oocyte nucleus then migrates to a random side of the anterior cortex, where it expresses gurken again, which induces dorsal fate in the nearby follicle cells (Gonzalez-Reyes et al., 1995, Neuman-Silberg and Schupbach, 1993, Ray and Schupbach, 1996, Roth et al., 1995, 1999, Theurkauf et al., 1992). There is much evidence in favor of a simple model where the oocyte is transported anteriorly along microtubules by a minus end-directed motor. First, in mutants where the microtubules reorganize aberrantly such that the minus ends are at the anterior and posterior poles and the plus ends are central, the nucleus does not move (Gonzalez-Reyes et al., 1995, Lane and Kalderon, 1994, Micklem et al., 1997, Roth et al., 1995). If the nucleus associates with dynein, it makes sense that it would not move, as it is already located at the microtubule minus-ends. No defects in positioning of the oocyte nucleus were reported in dhc mutants, but the particular mutants tested were hypomorphic (McGrail and Hays, 1997). Second, experiments with DLis-1 mutant oocytes suggest a role for dynein in nuclear migration. As mentioned above, DLis-1 is the Drosophila homolog of the gene associated with Lissencephaly, a brain disease where neuronal migration is defective, perhaps due to defects in nuclear migration. In DLis-1 mutants, the oocyte nucleus is mispositioned, and Dhc, which normally accumulates at the nucleus (and elsewhere) is no longer localized (Swan et al., 1999). One interpretation of these observations is that DLis-1 associates with dynein, enabling it to transport the nucleus anteriorly. However, why the nucleus migrates specifically to an anterolateral corner of the oocyte is unclear. 15.4.3

Lipid Droplet Migration in Drosophila Embryos

Lipid droplets migrate from the embryo periphery at syncitial blastoderm, to the center at cellularization, and then apically again during gastrulation (Welte et al., 1998; Fig. 15.4). The droplet migrations appear to be of little consequence to the embryo, but they have provided a unique opportunity to analyze developmentally-regulated, microtubule-dependent organelle movements in vivo. The second lipid droplet migration is dynein dependent; the droplets fail to migrate back out to the periphery in embryos from Dhc64C mutant mothers (Gross et al., 2000). Consistent with the idea that the lipid droplets hitch a ride on dynein out to the apical surface of the embryo, the kinesin-bgal marker indicates that the microtubules are arranged with their minus ends at the periphery (apical) and their plus ends basal at this time (Welte et al., 1998). However, biophysical experiments performed on the lipid droplets in wild-type and klarsicht (klar) mutants, suggests that this model is an oversimplification. Klar is an apparently Drosophila-specific and unique protein (Mosley-Bishop et al., 1999), which is required for lipid droplet migration and also for nuclear migration in developing photoreceptors (see below). As in Dhc64C mutants, in embryos from klar mutant mothers, the second migration of lipid droplets out to the periphery fails to occur. Welte et al. (1998) observed that individual lipid dro-

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15.4 Asymmetric Organelle Localization

The Drosophila Klarsicht protein may regulate motor-driven migrations of lipid droplets and photoreceptor nuclei. In the embryo, lipid droplets migrate apically to the embryo periphery; each lipid droplet is powered by several dynein motors. Klar may somehow regulate the relative contributions to net movement of dyneins and kinesins bound simultaneously to a single lipid droplet. In the developing eye, perinuclear Klar is required for apical migration of photoreceptor nuclei. Dynein is known to be required for these nuclear migrations as well, but the roles of dynein and Klar in this process are uncertain.

Figure 15.4.

plets move bidirectionally in wild-type embryos, suggesting that both kinesin and dynein may be attached to a single lipid droplet. In addition, the speed of the droplets is the same in either direction, and the differences in net movements are as a result of the time spent traveling in the plus or minus direction. In klar mutant embryos, the speeds and distances traveled in the minus direction were reduced markedly. Moreover, measurements of the forces on the lipid droplets using optical tweezers indicated that in wild-type embryos, they are in increments of 1.1 pN, a reasonable estimate of the force exerted by one motor protein; this observation suggests that multiple motors normally power the movement of a single lipid droplet. In klar mutants, the forces on the droplets are reduced to one incremental unit (embryos lacking Dhc also showed reduced forces on lipid droplets). From these data, the authors propose that Klar may attach motors to lipid droplets, or coordinate the activities of kinesins and dyneins bound to the same vesicle. Dhc64C colocalizes with lipid droplets (Gross et al., 2000), and in order to evaluate the model for Klar function further, it will be important to determine whether or not Klar colocalizes with lipid droplets as well.


15.4.4

Nuclear Migration in Drosophila Photoreceptors

The Drosophila compound eye is made up of hundreds of identical facets, or ommatidia, each of which has eight photoreceptors. The eye develops within an epithelial monolayer called the eye-antennal imaginal disc, where cells are suspended between an apical and a basal membrane. Prior to differentiation, the nuclei of all cells in the disc plunge basally, and then rise to the apical surface as photoreceptor cells differentiate (Wolff and Ready, 1993). Although apical localization of the nucleus appears not to be essential for most aspects of photoreceptor determination (Fischer-Vize and Mosley, 1994), most of the photoreceptor cell cytoplasm surrounds the nucleus, and therefore nuclear position greatly affects photoreceptor cell shape. Dynein is clearly required for the apical migration of photoreceptor cell nuclei that accompanies determination; expression in the eye of dominant negative forms of Glued protein (the p150 subunit of dynactin) results in the failure of apical nuclear migration (Fan and Ready, 1997). In addition, DLis-1 and BicD, which are essential for several other dynein-dependent processes in the oocyte and embryo (see above), are also required for nuclear migration in developing photoreceptors (Swan et al., 1999). The roles of DLis-1 and BicD in eye development have not yet been elucidated. The idea that Klar is a mechanical component of the cytoskeleton is supported by the finding that Klar protein is perinuclear in the developing Drosophila eye (Mosley-Bishop et al., 1999). However, there is at present no simple model for nuclear migration that takes into account both the perinuclear location of Klar and the idea that Klar regulates motor function directly. On the basis of Nodbgal and kinesin-bgal markers, and g-tubulin localization, which show that the MTOC is associated with the nucleus, the microtubules of differentiating photoreceptors appear to be organized with their minus ends at the nucleus and their plus ends apical (Mosley-Bishop et al., 1999; Swan et al., 1999; K. Patterson and J. A. Fischer, manuscript in preparation). Given this cytoskeletal organization, and the observation that dynein is essential for apical nuclear migration, a simple model for the mechanism of nuclear migration, taken from budding yeast (Morris et al., 1998), is that the nucleus, attached to the MTOC, is ‘reeled up’ to the apical cell surface by dynein tethered to the apical membrane. According to the model for Klar function in lipid droplet transport, Klar would be expected to be located at the apical cell surface, where it might tether dynein or regulate dynein function. More experiments, including localization of Dhc in the eye disc, are required to formulate a better model for Klar function.

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References

15.5

Future Directions

Major advances in understanding the roles of motors in development have taken place during the past few years. These advances are largely due to creative imaging and biophysical techniques and imaginative manipulations of cells, combined with more traditional genetic and biochemical approaches. One interesting observation is that the same proteins are used to set up cellular asymmetries in many different contexts, implying that there may be core motor and cytoskeletal regulatory complexes that may have different accessory factors. These factors could function as adapters to enable the motors to bind different cargo, and may be involved in the process that enables motors to choose between different cytoskeletal tracks. The regulation of these two processes is among the most exciting avenues for future discovery.

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16 Motors and Membrane Trafficking Kristen J. Verhey

16.1

Introduction

One hallmark of eukaryotic cells is that they contain many functionally distinct membrane-bounded compartments, or organelles. Each organelle carries out highly specialized functions because it possesses its own unique combination of biochemical components. Organelles can additionally be identified by their characteristic organization or topology and, in some cases, by their distinct localization in cells. Although each organelle has a unique molecular composition, proteins and lipids are continuously shuttled between compartments via transport vesicles that move in the secretory and endocytic membrane trafficking pathways. Organelles, transport vesicles and macromolecular complexes in eukaryotic cells move along the underlying cytoskeleton the microtubules and actin filaments. These movements are crucial for cell division, cell polarity, cell migration, embryonic development, and the formation of specialized cellular structures such as axons and dendrites, gap and tight junctions, and cilia and flagella. Both microtubules and actin filaments are polar structures with a defined orientation of their plus(fast growing) and minus- (slow growing) ends within cells. Microtubules are generally oriented with their minus-ends anchored in the cell center at the microtubule organizing center (MTOC) and their plus-ends pointed to the cell periphery although there are exceptions (Fig. 16.1). Actin filaments do not show such general polarity but are concentrated in the cell periphery and may display localized polarity, for example at the leading edge of a migrating cell. The many organelles and transport vesicles that need to be carried to different destinations in the cell use molecular motors to move along the microtubule and actin tracks. Indeed, the study of these motor proteins has implicated them in a wide range of cellular functions, particularly in vesicular transport during interphase and in spindle formation, chromosome segregation and cytokinesis during mitosis and meiosis. Genetic and molecular analysis has demonstrated that the kinesins and myosins each comprise a large superfamily whose members exhibit extensive homology in their mechanical (motor) domains but little homology

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Schematic diagram of cytoskeletal organization and vesicle transport in animal cells. (a) In a non-polarized fibroblast cell, the microtubules (blue) have their minus-ends anchored in the microtubule organizing center (MTOC) and their plus-ends in the cell periphery. Kinesins carry transport vesicles to the plus-ends of microtubules whereas dyneins carry transport vesicles to the minus-ends of microtubules. Actin filaments (red) do not show such overall polarity but are primarily localized in the periphery of the cell. Myosin family members carry transport vesicles to both the plus- and the minus-ends of the actin

Figure 16.1.

filaments. (b) In neurons, the minus-ends of the microtubules are anchored in the cell body and the plus-ends extend to the presynaptic terminal of the axon. In the dendrites, microtubules are oriented with both plus- and the minus-ends pointed to the distal postsynaptic terminal. (c) In a vertebrate photoreceptor cell, the minus-ends of the microtubules are anchored at the base of the connecting cilium (CC) and the plus-ends extend across the CC to the outer segment (OS) as well as to the secretory pathway in the inner segment (IS). (d) In a budding yeast cell, actin cables extend from the mother cell into the bud.

in their non-motor regions. These divergent regions are thought to impart specificity to each motor protein in terms of cargo binding, regulation and oligomerization (see Chapter 1 by Kieke and Titus and Chapter 3 by Hirokawa and Takemura). In contrast, there is no superfamily of dynein motors but rather cytoplasmic dynein is a large multi-subunit complex comprised of the protein products of many genes (see Chapter 2 in this volume by King). Most members of the kinesin superfamily move to the plus-ends of microtubules and are thought to drive all long-distance anterograde (toward the cell periphery) movement of organelles and vesicles (Fig. 16.1). Multiple dyneins are also present in cells and, as they move to the minus-ends of microtubules, are presumed to transport multiple car-

16 Motors and Membrane Trafficking

goes in a retrograde direction (back to the cell center; Fig. 16.1). The numerous members of the myosin superfamily move in both directions along actin filaments and carry cargoes shorter distances in the cell periphery (Fig. 16.1). The specific focus of this chapter will be to highlight general principles that have emerged concerning the roles of the cytoskeleton and motor proteins in membrane trafficking in animal cells. This chapter will not be comprehensive concerning the cellular functions of individual members of the kinesin, dynein and myosin families of motor proteins. For such discussions, the reader is referred to the accompanying chapters by Hirokawa and Takemura (Chapter 3), Kierke and Titus (Chapter 1) and King (Chapter 2). In addition, it is clear that many aspects of membrane trafficking are different in plants which show a greater dependence on actinbased movements (see Chapter 18 by Reddy). At present, the major questions in the motor protein field deal with identifying which motors are directly involved in moving which cargoes, the molecular mechanism by which motor proteins interact with their cargo, and how motor protein transport of cargo is regulated.

16.2

The Logic and Order of Membrane Trafficking

Most of what we know about the roles of the cytoskeleton and motor proteins in membrane trafficking comes from studies in cultured fibroblasts. In animal cells, the ER network and early endosomes are spread to the periphery of the cell whereas the Golgi apparatus and late endosomes are located near the MTOC in the center of the cell. These organelles are functionally linked by the secretory and endocytic pathways. In general, the secretory pathway describes the paths taken by protein and lipid components synthesized in the peripheral ER, transported to the Golgi apparatus in the cell center, and then transported to the ER or the plasma membrane in the cell periphery (Lippincott-Schwartz et al., 2000). In the endocytic pathway, material taken in at the plasma membrane is transported first to peripheral early endosomes then to recycling endosomes or late endosomes in the cell center and finally to peripheral lysosomes (Lemmon and Traub, 2000). The movement of protein and lipid along these pathways is mediated by transport vesicles that bud from the donor organelle and fuse with the target organelle. This is a highly regulated and ordered process that occurs without compromising the fidelity of the donor and acceptor organelles. The directed transport of vesicles through the membrane trafficking pathways is governed by the inherent structural and organizational differences between microtubules and actin filaments as well as the intrinsic directionality of motor protein movement (Fig. 16.1). Work over the past 20 years has led to several general principles regarding how membrane trafficking is controlled at the molecular level (Chen and Scheller, 2001, Mellman and Warren, 2000, Pfeffer, 1999, Stephens and Pepperkok, 2001, Zerial and McBride, 2001). The budding of transport vesicles from the donor compartment membrane is aided by cytoplasmic coat proteins. These transport vesicles

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16.3 The Cytoskeleton and Motor Proteins in Membrane Trafficking

then fuse with each other to form large tubulovesicular carriers that move to the target organelle. The tubulovesicular carriers may be sorting stations as well in that, in some cases, they pinch off smaller vesicles that return to the donor organelle. The transport carrier is delivered to its cellular destination by molecular motors moving along the actin and/or microtubule cytoskeletons. Once the transport carrier has reached its target, tethering proteins collect and restrain vesicles at or near the target membrane. Finally, recognition of and subsequent fusion with only the appropriate target compartment is governed by organelle-specific members of the SNARE protein family and small GTPases of the Rab protein family.

16.3

The Cytoskeleton and Motor Proteins in Membrane Trafficking 16.3.1

Role of the Cytoskeleton and Motor Proteins in Organelle Localization

The cytoskeleton is required for proper positioning of most, if not all, of the membranous components of the cytoplasm (Cole and Lippincott-Schwartz, 1995). The spreading of the ER network to the periphery of the cell requires microtubules and a plus-end motor as depolymerization of microtubules causes the ER to retract back towards the cell center (Allan, 1996). Similarly, the normal localization of the Golgi apparatus at the MTOC in the center of the cell, maintained by the activity of cytoplasmic dynein, is disrupted by microtubule depolymerization and results in the scattering of Golgi elements throughout the cytoplasm (Allan, 1996, Burkhardt, 1998). Localization of endosomal compartments near the MTOC is also dependent on an intact microtubule cytoskeleton and possibly cytoplasmic dynein, whereas lysosomes located throughout the cell are more motile and potentially utilize both the microtubule and actin cytoskeletons (Cordonnier et al., 2001, Lebrand et al., 2002 and references therein). Organelles outside the secretory and endocytic pathways also utilize the cytoskeleton and motor proteins for their localization and movement. For example, mitochondria move bidirectionally along microtubules as well as along actin filaments (Hollenbeck, 1996). Nuclear movement in motile and in mitotic cells is also dependent on the cytoskeleton and motor proteins, particularly cytoplasmic dynein and the associated dynactin complex (Dujardin and Vallee, 2002, Lippincott-Schwartz, 2002). 16.3.2

Role of the Cytoskeleton in Membrane Trafficking Events

A role for the cytoskeleton in the secretory pathway is generally accepted although it has been difficult to demonstrate. Whereas disruption of the actin cytoskeleton has no effect on delivery of secretory proteins to the plasma membrane, reports using microtubule antagonists have provided mixed results, sometimes preventing or randomizing surface delivery while at other times showing little to no effect.


There are several explanations for these conflicting effects. First, prolonged drug exposure leads to fragmentation and dispersal of the Golgi apparatus to ER exit sites near the plasma membrane such that long-distance microtubule transport is no longer required (Bloom and Goldstein, 1998). Second, the flat nature of tissue-culture cells may allow transport vesicles to reach the plasma membrane by random diffusion when the microtubule tracks are destroyed. Third, some microtubules in cells are stable and difficult to fully depolymerize with microtubule antagonists and so continued vesicle trafficking along the secretory pathway may occur on the few remaining microtubules (Bre et al., 1987). The most striking evidence that transport carriers in the secretory pathway move along the microtubule cytoskeleton comes from the recent advent of green fluorescent protein (GFP)-fusion proteins in visualizing membrane trafficking in living cells. These studies have clearly demonstrated that GFP-fusion proteins incorporated into tubulovesicular carriers moving between the ER and Golgi complex, as well as between the Golgi complex and the plasma membrane, move down labelled microtubule tracks toward their target organelle (Lippincott-Schwartz et al., 2000). Studies aimed at determining the role of the cytoskeleton in endocytic trafficking events in animal cells have also provided variable results which may be related to differences in cell type, growth conditions, drug conditions, and so forth. Taken together, a large body of work has provided the following general concepts for endocytic trafficking. Early events in the endocytic pathway, such as internalization from the cell surface and homotypic fusion of early endosomes, are not affected by microtubule depolymerization but rather are facilitated by the actin cytoskeleton and myosin motors (Apodaca, 2001, Gruenberg, 2001, Munn, 2001, Qualmann et al., 2000). The actin cytoskeleton may also facilitate endocytic events due to the role of actin polymerization in generation of mechanical force to deform the membrane or in propelling endocytic vesicles through the cytoplasm (Jeng and Welch, 2001, Lanzetti et al., 2001, Schafer, 2002). Transport along the next step of the endocytic pathway, from peripheral early endosomes to central recycling and late endosomes, is dependent on microtubules and minus-end directed motors (Gruenberg, 2001). The transport of material from central recycling and late endosomes to plasma membrane and lysosomes, respectively, is likely to occur along microtubules. Plasma membrane receptors likely return from central recycling endosomes to the cell surface along microtubules; although this step is not affected by treatment with microtubule-disrupting reagents, such treatments result in the dispersion of the recycling compartment to the periphery of the cell and may therefore eliminate the requirement for long distance transport (Gruenberg, 2001). In addition, a kinesin family member has been implicated in this transport step (Lin et al., 2002). The final step in the endocytic pathway, transport of material from central late endosomes to lysosomes, is dependent on both microtubules and actin cytoskeletons (Gruenberg, 2001).

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16.3 The Cytoskeleton and Motor Proteins in Membrane Trafficking

16.3.3

Role of Motor Proteins in Membrane Trafficking Events

Determining the role of various motor proteins in particular membrane trafficking pathways has proved to be more difficult than expected. Genetic studies have demonstrated that individual motor proteins play a role in membrane trafficking but have failed to determine the molecular identity of the cargo (Bowman et al., 1999, Brendza et al., 2000, Harada et al., 1998, Martin et al., 1999, Seiler et al., 1999). Similarly, studies utilizing inhibitory antibodies and dominant negative proteins have clearly implicated kinesins in anterograde transport events and cytoplasmic dynein in retrograde trafficking events, but have also failed to identify the exact transport cargo (Burkhardt et al., 1997, Lafont and Simons, 1996). While a general role for myosin motor proteins in late secretion and early endocytic events is clear, the specific role of individual myosin family members, particularly in early secretion events, remains to be determined (Buss et al., 2001b, Stow et al., 1998). Particularly frustrating is that investigations into the roles of individual motor proteins in a particular membrane trafficking step have yielded contradictory results. This is perhaps most true for the kinesin I family of kinesin motor proteins. Inhibition of kinesin I in cultured cells using antisense oligonucleotides or function-blocking antibodies was found to interfere with membrane traffic from the Golgi apparatus to the ER (Feiguin et al., 1994, Ferreira et al., 1992, LippincottSchwartz et al., 1995). However, targeted disruption of the KHC subunit of ubiquitous kinesin I in mice and Drosophila, although lethal during development, has no effect on Golgi-to-ER traffic, nor does expression of mutant kinesin I molecules in cultured cells (Brendza et al., 2000, Hollenbeck and Swanson, 1990, Hurd and Saxton, 1996, Nakata and Hirokawa, 1995; Tanaka et al., 1998, Verhey et al., 2001). A similar contradiction exists for a kinesin I requirement for mitochondrial and lysosomal movement. It is possible that these apparently conflicting results are due to artifacts such as cross-reacting antibodies or depletion of molecules essential to more than one pathway. Such caveats make it difficult to determine whether an observed effect is ‘direct’ due to inhibition of the motor protein in question or whether the effect is ‘indirect’ due to inhibition of something downstream. A resolution of these conflicting results requires the identification of molecular components of the cargo and the motor cargo linker. Recent studies have suggested that a motor protein is capable of carrying multiple cargoes, and possibly different cargoes depending on cellular context, due to the ability of individual motor proteins to bind to many polypeptides that each link to a different cargo (Goldstein, 2001, Karcher et al., 2002). For example, the KLC subunit of kinesin I binds directly to several putative motor cargo linker proteins such as the JNK-interacting proteins-1 and -2 (JIP-1, -2), the JIP3/Syd protein, and the Alzheimers Precursor Protein (APP) (Bownman et al., 2000, Kamal et al., 2000, Verhey et al., 2001). Similarly, the KAP3 subunit of kinesin II has been shown to bind directly to the tumor suppressor gene adenomatous polyposis coli (APC), the small GTPase regulator SmgGDS, the mixed lineage serine/threonine kinase MLK2, and the neuronal spectrin a-fodrin (Jimbo et al., 2002, Nagata et al.,


1998, Shimizu et al., 1996, Takeda et al., 2000). In the case of cytoplasmic dynein, there is no dynein superfamily but rather subunit heterogeneity is thought to assemble many forms of dynein and, in some cases, to bind dynein to different cargoes (Chuang et al., 2001, King et al., 2002, Susalka and Pfister, 2000, Tai et al., 2001). In addition, cytoplasmic dynein is regulated by an associated complex, the dynactin complex, which is likely to contribute to the ability of dynein to bind to many different cargoes. Over-expression of the dynamitin subunit of the dynactin complex disrupts a variety of general dynein functions (Allan, 2000, Burkhardt et al., 1997,; Holleran et al., 1998, Valetti et al., 1999). Members of the myosin V family of motor proteins also appear to ferry multiple cargoes due to interactions with multiple motor cargo linker molecules. The globular tail of Myo2p, a class V myosin involved in transporting post-Golgi secretory vesicles and vacuoles in Saccharomyces cerevisiae, contains subdomains with separable cargo-binding regions whose mutation leads to defects in vacuole inheritance but not polarized growth and vice versa (Catlett et al., 2000). This result demonstrates that different domains of Myo2p are responsible for binding to different organelles and suggests that the two classes of organelles have different receptors for the motor. In addition to the idea that a motor can carry multiple cargoes, an emerging idea is that a particular organelle can bind to multiple motors, possibly simultaneously, and that each motor activity is regulated. For instance, mitochondria have been proposed to be transported by kinesin I, KIF1B and KLP647A (Nangaku et al., 1994, Pereira et al., 1997, Tanaka et al., 1998). Importantly, this observation suggests that cells employ multiple, redundant motors to transport organelles such that no single motor is indispensable. In addition, this implies that the various motors on a transport carrier must be coordinately regulated to ensure directed transport to the correct cellular destination (Section 16.4). These concepts, and the contradictory results concerning the roles of individual motor proteins, will only be resolved when the molecular identity of motor cargo linker molecules and of the cargo components are determined (Section 16.5).

16.4

Cooperation between Motors 16.4.1

Coordination of Movement along Microtubule and Actin Tracks

For many years, microtubule-based and actin-based motility were considered to be two independant systems performing separate tasks in cellular events such as cell locomotion, organelle organization and vesicle transport. However, recent studies demonstrate that these systems work together. The first suggestion that transport vesicles move on both cytoskeletal filaments came from studies in extruded squid axoplasm where individual particles were observed to move along a microtubule and then switch to moving along an actin filament (Kuznetsov et al., 1992). Since then, a large variety of organelles and transport vesicles have been demon-

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16.4 Cooperation between Motors

strated to move along both actin filaments and microtubules and, in some cases, to contain both myosin and kinesin or dynein motor proteins. The general model for coordination of motility along both cytoskeletal filaments is that, at least in animal cells, microtubule-dependent transport is used to drive long-range movements between the cell center and the cell periphery and actin-dependent transport is used to drive short-range movements primarily in the periphery (Fig. 16.1). This model is attractive as concerted action between the two systems would help ensure the precise distribution of organelles and vesicles in the cell (Allan and Schroer, 1999, Brown, 1999, Goode et al., 2000). One of the best examples of the coordination of microtubule- and actin-based transport systems has been provided by studies in fish and frog melanophores where pigment granules (melanosomes) aggregate or disperse to achieve color changes in response to external stimuli. Aggregation and dispersion of melanosomes is accomplished by bidirectional transport along microtubules, via the motor proteins kinesin II and dynein, coupled to shuttling of melanosomes along randomly-oriented actin filaments, via the motor protein myosin V (Rodionov et al., 1998, Rogers et al., 1997, Rogers and Gelfand, 1998, Tuma et al., 1998). Melanosome movement in mouse melanocytes occurs by a similar mechanism. Melanosomes are transported to the tips of the melanocyte’s dendritic extensions where they accumulate and are taken up via phagocytosis by neighboring keratinocytes for pigmentation of skin and hair. Peripheral accumulation of melanosomes is accomplished by a cooperative transport mechanism in which longrange, bidirectional transport along microtubules, driven by the motor proteins kinesin II and dynein, is coupled with myosin Va-dependent capture in the actin-rich periphery (Nascimento et al., 1997, Provance et al., 1996, Wu et al., 1997, 1998). However, it was unclear whether movement of melanosomes in either system occurs by sequential activation of microtubule- and actin-based motors or whether the motor proteins work at the same time to move melanosomes along both filaments. A recent report has indicated that both mechanisms can occur (Gross et al., 2002a). During the dispersion of melanosomes in Xenopus melanophores, myosin V and kinesin II work together as myosin V carries kinesin II-driven granules along neighboring actin filaments. In contrast, myosin V and dynein work sequentially as myosin V terminates dynein-dependent movement to the microtubule minus-ends and. During aggregation, the situation is reversed and melanosomes are driven to the center of the cell by cytoplasmic dynein. In this case, myosin V seems to be released from melanosomes, suggesting that coordination of microtubule- and actin-based systems may involve modulating not only motor activity but also the motor cargo linkage. Coordinated control of microtubule- and actin-based systems has been observed in a variety of other systems and organisms. In vertebrate photoreceptor cells, lightsensing components such as rhodopsin depend on the coordinated activity of kinesin II, cytoplasmic dynein and myosin VIIa motors for their transport from the inner segment (the dendritic equivalent) through the connecting cilium to the outer segment (Fig. 16.1; Williams, 2002). In neurons, cooperation of myosin V and kinesin family members in fast axonal transport is thought to be due to se-


quential activation of kinesins for long-range microtubule-dependent movement and then activation of myosin Va (previously carried as a passive passenger) in the distal actin-rich presynaptic terminal (Fig. 16.1). This scenario is complicated by the fact that myosin V can support long-range axonal transport in the absence of microtubules (Bridgman, 1999, Evans et al, 1998, Prekeris and Terrian, 1997). Finally, in hepatoma cells, a myosin I family member has been proposed to aid microtubule-based movement not by moving lysosomes along actin filaments but rather by transiently tethering them on actin filaments during their bidirectional movement along microtubules (Cordonnier et al., 2001). 16.4.2

Coordination of Bidirectional Movement along Microtubule Tracks

In addition to coordinating the long- and short-range movements of organelles and vesicles along microtubule and actin tracks, cells also coordinate bidirectional transport along microtubules. A variety of organelles and transport vesicles have been observed to move towards both the plus- and the minus-ends of microtubule tracks, often reversing directions within short time periods (Gross et al., 2002a,b, Ma and Chisholm, 2002, and references therein). One of the best examples of bidirectional movement along microtubules is intraflagellar transport (IFT) (Rosenbaum et al., 1999). In this process, best characterized in the green alga Chlamydomonas, large assemblies of flagellar precursor materials are carried by kinesin-II to the tips of flagella for their formation and maintenance (anterograde) and then by cytoplasmic dynein back to the basal body for recycling (retrograde) (Fig. 16.2). Mutants lacking either motor protein are defective in transport of membranous cargo and have short or missing flagella (Cole et al., 1993, Kozminski et al., 1995, Pazour et al., 1998, 1999, Porter et al., 1999, Walther et al., 1994). Bidirectional transport of IFT complexes also appears to play a role in the formation and function of internodal cilia in mammalian embryos (Hirokawa, 2000), of non-motile chemosensory neurons in mice and C. elegans (Cole et al., 1998, Orozco et al., 1999, Signor et al., 1999a,b, Wicks et al., 2000) and of photoreceptors in vertebrates (Beech et al., 1996, Marszalek et al., 2000, Muresan et al., 1997, 1999, Pazour et al., 2002). Bidirectional movement on microtubules could be due to the alternating presence of plus- and minus-end motors to a transport cargo or due to alternating activities of opposite motors bound to the same cargo. Visualization of GFP-labeled KIF1A kinesin in C. elegans and GFP-labeled dynein in Drosophila demonstrated that motors are present on structures that move along microtubules in both minus- and plus-end directions and rapidly change directions (Ma and Chisholm, 2002, Zhou et al., 2001). These results suggest that bidirectional transport on microtubules is due to the presence of dynein and kinesin motors on the same transport cargo. This is attractive because it would explain how minus-end motors required for retrograde transport could be carried to the cell periphery and how plus-end motors could be carried back to the cell center for another round of transport.

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16.4 Cooperation between Motors

Bidirectional transport of IFT particles on microtubules. Flagellar precursor materials (black) are carried on IFT rafts (gray) to the tip of the flagellum by kinesin II (purple) and then to the basal body at the base of the

Figure 16.2.

flagellum by cytoplasmic dynein (green). In addition, kinesin II and dynein can be carried as cargo and returned to their point of departure by the oppositely directed motor.

The simultaneous binding of motors which direct transport in opposite directions necessarily requires that the activities of these bound motors are regulated to achieve net transport to the cargo’s final destination in the cell. One possibility is that motor proteins on the same vesicle may be active at the same time (the tugof-war model) as suggested by the finding that the more processive plus-end motors override the activity of minus-end motors on the same vesicle (Muresan et al., 1996). An alternative possibility is that the regulation of the anterograde and retrograde motors is coordinated such that they are active sequentially (the coordination model). In support of this, several reports have demonstrated that motors can be carried as inactive cargo (Evans et al., 1997, 1998, Iomini et al., 2001, Lane and Allen, 1999, Lippincott-Schwartz et al., 1995, Ma and Chisholm, 2002, Nilsson et al., 1996, Reese and Haimo, 2000, Welte et al., 1998). In addition, genetic studies in Drosophila have demonstrated that disruption of kinesin or dynein inhibits transport in both directions and therefore that the functions of kinesin, dynein and the dynactin complex are interdependent in fast axonal transport (Bowman et al., 1999, Martin et al., 1999). Strong support for the coordination model comes from recent studies on lipid droplets in Drosophila embryos. Lipid droplets are carried bidirectionally along microtubules by cytoplasmic dynein and a kinesin family member. When droplets reverse direction, they do not pause but move in the opposite direction without delay (Gross et al., 2000). When minus-end motility is specifically impaired, plus-end motility is not improved (as expected in a tug of war) but rather is impaired in an allele-specific manner (Gross et al., 2002b). Similar conclusions were reached when bidirectional transport of melanosomes along microtubules, driven by kinesin II and cytoplasmic dynein, was examined in Xenopus melanophores (Gross et al., 2002a). Taken together, these results suggest that the activity of plus- and minus-end motors on the same droplet is coordinated to prevent simultaneous activity.


16.4.3

Molecular Mechanisms for the Coordination of Motors on the Same Transport Cargo

How could the coordination of microtubule- and actin-based motor proteins be accomplished at the molecular level? Recent discoveries suggest some interesting possibilities. Although direct protein interactions between kinesin and dynein/dynactin have not been detected, members of the myosin family have been found to interact directly with members of both the kinesin and dynein families. In most cases, the functional significance of these interactions remains to be determined (Benashski et al., 1997, Espindola et al., 2000, Prekeris and Terrian, 1997, Schroer, 1994). A direct molecular and functional link between myosin and kinesin family members has been demonstrated in the budding yeast Saccharomyces cerevisiae. Polarized growth is due to the transport of post-Golgi secretory vesicles to the bud by Myo2p, a myosin V family member (Fig. 16.1). Mutations in Myo2p can be partially suppressed by over-expression of Smy1p, a kinesin family member (Lillie and Brown, 1992). The interaction between Smy1p and Myo2p is direct; the tail of Myo2p binds to the tail of Smy1p and is responsible for the polarized distribution of Smy1p (Beningo et al., 2000, Lillie and Brown, 1998). Surprisingly, Myo2p and Smy1p localization and function require an intact actin cytoskeleton but do not require the presence of microtubules or a functional ATPase motif in Smy1p (Lillie and Brown, 1998). As a direct molecular interaction has also been demonstrated for myosin V and kinesin I in animal cells (Huang et al., 1999), it is tempting to speculate that direct interactions between myosins and kinesins may indeed exist during active transport. Whether this interaction is involved in regulating the activity of the motors or whether the motors are being carried as components of the cargo remains to be determined. It has been postulated that when budding yeast lost the hyphal aspect of polarized growth, and thus their dependence on microtubule-based transport, Smy1p lost its ability to interact with and move along microtubules. The persistence of an interaction of Smy1p with Myo2p then is thought to stabilize the formation and/or transport of secretory vesicles along actin cables (Brown, 1999, Hammer and Wu, 2002). Another mechanism that might mediate the coordination of actin- and microtubule-based vesicle transport has emerged from recent studies on proteins that localize to the plus-ends of microtubules. The cytoplasmic linker protein CLIP-170 was first discovered as a protein involved in tethering endocytic cargo to the plus-ends of microtubules. The binding of CLIP-170 to the plus-ends of microtubules is as dynamic as the microtubules themselves and GFP-CLIP-170 appears to ‘track’ the growing plus-ends of the microtubules (McNally, 2001, Schroer, 2001, Schuyler and Pellman, 2001). Cytoplasmic dynein and the dynactin complex co-localize with CLIP-170, possibly via the type I lissencephaly protein LIS1, at the plus-ends of microtubules (Coquelle et al., 2002, Tai et al., 2002, Valetti et al., 1999, Vaughan et al., 1999). In addition, a class VI myosin implicated in endocytic trafficking in Drosophila, myosin 95F, associates with and co-localizes with dynactin and a Drosophila CLIP-170 homolog, D-CLIP-190, at the plus-ends of microtubules

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16.5 Molecular Mechanisms of Motor–Cargo Linkage

(Lantz and Miller, 1998). This has led to the attractive possibility that the distal ends of microtubules represent a specialized compartment designed for track switching. In this model, vesicles delivered from the plasma membrane through the actin cortex via myosin VI are captured at microtubule plus-ends by means of the interaction between myosin VI and CLIP-170 (Buss et al., 2001a). The interaction between CLIP-170 and dynein/dynactin would allow subsequent transfer for microtubule-based transport to the cell center. Several other cytoplasmic linker proteins belonging to the CLIP-170 family have been identified and localized to various organelles (De Zeeuw et al., 1997, Perez et al., 2002, Thieman et al., 2000). An attractive possibility then is that the CLIP proteins play a general role in facilitating the interactions between transport vesicles, the cytoskeleton and motor proteins and thus in the coordination of vesicle transport along both microtubule and actin tracks.

16.5

Molecular Mechanisms of Motor–Cargo Linkage

One of the most exciting advances in the roles of motor proteins in membrane trafficking is the identification of molecules that serve as motor cargo linkers. Proteins that serve as receptors for motors are not specialized proteins whose only function is to bind to the motor. Instead, the motor cargo linker proteins are bonafide components of the cargo and have specific functions at their destination. In addition, an emerging concept is that motor proteins utilize multiple motor cargo linkages, including binding to transmembrane and peripheral membrane proteins. While this would allow individual motor proteins to transport different cargoes in different contexts (Section 16.3), it is also possible that multiple attachments may be made to the same cargo. This would allow a tight interaction between motor and cargo that would counteract the viscous drag of pulling a transport carrier through the cytoplasm (Holzworth et al., 2002). For example, it is possible that the dynactin complex mediates a generic interaction of dynein with many types of vesicles that have spectrin networks on their surfaces whereas dynein subunits mediate specific interactions with various cargoes. 16.5.1

Soluble Adaptor or Scaffolding Proteins as Motor Cargo Linkers

Recent work has suggested an attractive concept for how members of the kinesin superfamily of motor proteins link to their cargoes motors bind to soluble adaptor or scaffolding proteins that link them indirectly to their cargoes (Fig. 16.3 and Tab. 16.1; Verhey and Rapoport, 2001). In general, adaptor and scaffolding proteins are defined as molecules that contain no enzymatic activity of their own but rather contain a variety of protein protein and protein lipid interaction domains. Through these domains, scaffolding proteins bind to many different proteins at the same time and assemble large multi-protein complexes. The presence of


Motor proteins utilize two general types of motor cargo linker molecules to bind to transport vesicles. (a) Motor proteins bind to soluble scaffolding proteins which then bind to transmembrane proteins on the transport

Figure 16.3.

Table 16.1

vesicle, as well as to cytoplasmic proteins such as kinases. (b) Motor proteins bind to soluble adaptor proteins which then bind to organellespecific GTP-bound Rab proteins.

Motor proteins that utilize scaffolding or adaptor proteins as motor cargo linkers.

Motor protein

Scaffold

Reference

Kinesin I

JIP-1/-2

Verhey et al. (2001)

Kinesin I

JIP-3

Bowman et al. (2000) Verhey et al. (2001)

Kinesin II

IFT raft

Rosenbaum et al. (1999)

KIF1C

14-3-3

Dorner et al. (1999)

KIF13A

AP-1 adaptor

Nakagawa et al. (2000)

KIF17

mLin10

Setou et al. (2000)

GAKIN

hDlg

Hanada et al. (2000)

Dynein

Dynactin

Vaughan and Vallee (1995)

NINAC myosin III

INAD

Sheng and Sala (2001)

Myosin VI

AP-2 adaptor

Buss et al. (2001a)

Myosin Va

Melanophilin?

Fukuda et al. (2002), Hume et al. (2002)

389

390


both cytosolic and transmembrane proteins in this complex would link the motor to membranous vesicles and allow the motor to carry a variety of cargo molecules at the same time. Transport of cargoes down axons to the presynaptic membrane is driven, at least in part, by kinesin I. The KLC subunit of kinesin I binds three scaffolding proteins, the JIP-1, -2 and -3 proteins, that were originally identified as scaffolding proteins for the JNK (c-Jun N-terminal Kinase) signaling cascade. JIP-1 and -2 are very similar in sequence and domain structure but JIP-3 (Syd in Drosophila and UNC16 in C.elegans) is unrelated. The TPR motifs of KLC bind to the extreme C-terminal residues of JIP-1 but to internal residues in JIP-3/Syd/UNC-16, suggesting that the mechanism of motor scaffold interaction may be different (Bowman et al., 2000, Verhey et al., 2001). The JIP proteins bind in turn to multiple proteins, including components of the JNK signaling cascade, the GTPase activator RhoGEF, and the transmembrane receptor proteins ApoER2 and APP (Matsuda et al., 2001, Scheinfeld et al., 2002, Verhey and Rapoport, 2001). Binding to the transmembrane proteins presumably links kinesin I to its membrane-bound cargo. Over-expression of a dominant negative form of kinesin I or disruption of the interaction between KLC and JIP-1 abolishes the proper localization of the JIP scaffolding proteins, as well as an associated kinase, to the tip of the neurites in cultured neuronal cell lines (Verhey et al., 2001). Recent genetic studies lend strong support to the hypothesis that kinesin I and the JIP scaffolding proteins function in the same pathways in axonal transport. In Drosophila, the JIP-3/ SYD/UNC-16 mutant displays a phenotype similar to that of the kinesin I mutant in terms of larval paralysis and organelle build-up in the axons (Bowman et al., 2000). In C. elegans, loss-of-function mutations in the gene encoding JIP3/Syd/ UNC-16 results in the mislocalization of synaptic vesicle and glutamate receptor markers (Byrd et al., 2001). Taken together, these data suggest that kinesin I functions in the anterograde transport of vesicular cargo down axons and that the linkage of kinesin I to the cargo membrane is mediated by the JIP scaffolding proteins. Several other members of the kinesin family have been found to interact directly with scaffolding proteins and thus indirectly with their membrane cargoes. The kinesin motor protein KIF13A, most closely related to the KIF1 family of kinesins, binds directly to the b1-adaptin subunit of the AP-1 adaptor complex and thereby indirectly to transmembrane proteins such as the mannose-6-phosphate receptor (M6PR). The AP-1 complex functions in the formation of clathrin-coated vesicles that carry protein and membrane from the trans-Golgi network (TGN) to endosomes/lysosomes or to the plasma membrane. Over-expression of KIF13A caused mislocalization of AP-1 and M6PR from the perinuclear Golgi complex to the cell periphery (Nakagawa et al., 2000). It remains to be determined whether KIF13A carries all types of TGN-derived vesicle cargoes or is relatively specific for M6PRcontaining cargoes. In addition, the exact role of KIF13A and the AP-I adaptor complex in transport from the TGN to endosomes versus TGN to plasma membrane remains to be determined. Another member of the KIF1 kinesin family, KIF1C, binds to several isoforms of 14-3-3 proteins, a conserved family of ubiquitously expressed molecules that serve as scaffolds for a variety of signaling proteins.


KIF1C has been implicated in retrograde trafficking from the Golgi complex to the ER, but it remains to be determined how its interactions with 14-3-3 proteins relates to this cellular function (Dorner et al., 1998, 1999, Nakajima et al., 2002). The kinesin family member KIF17 binds directly to one of the PDZ domains of mLIN10, a component of the mLIN2/7/10 scaffolding complex that organizes postsynaptic signaling pathways in neuronal cells, and thus indirectly to membrane proteins that are components of the transport cargo. Indeed, immunoprecipitation of KIF17 brings down the mLIN complex as well as the NMDA receptor. In vitro, KIF17 transports a large vesicular complex containing the NMDA receptor along microtubules and this process is inhibited by the addition of a peptide corresponding to the tail of KIF17. These data suggest that KIF17 functions in the anterograde transport of vesicular cargoes down dendrites and that the linkage of KIF17 to the vesicle membrane is mediated by the mLIN2/7/10 scaffolding complex (Setou et al., 2000). The results of this study raise some interesting questions such as how the KIF17 motor complex delivers its cargo selectively to dendrites as both axons and dendrites contain microtubules oriented with their plus-ends in the distal end of the process. Although most of the evidence to date describes the binding of kinesin motor proteins to scaffolding proteins, such an interaction has also been demonstrated for several members of the myosin family of motor proteins. The myosin family member myosin VI, the only family member known to move to the minus-ends of actin filaments, has been implicated to play a role in clathrin-mediated endocytosis. Biochemical studies suggest that myosin VI interacts with the AP-2 adaptor complex that functions in clathrin-dependent transport from the plasma membrane to early endosomes (Buss et al., 2001a). This is reminiscent of the binding of the KIF13A kinesin to the AP-1 adaptor complex and lends strong support to the concept that adaptor/scaffolding proteins can function as motor cargo linkers for both actin- and microtubule-based transport. Further support comes from studies on a member of the myosin III class, NINAC, in Drosophila photoreceptor cells. NINAC binds directly to the scaffolding protein INAD which binds in turn to several proteins involved in phototransduction including rhodopsin and the TRP channel. In ninaC null flies, some components of the phototransduction cascade, such as calmodulin, were mislocalized whereas others were not. While a role for NINAC in the proper localization of signaling molecules in the photoreceptor is still an attractive possibility, the functions of NINAC and INAD in this process are still unclear (Fanning and Anderson, 1999, Sheng and Sala, 2001). More recently, the myosin family member myosin Va has been demonstrated to interact with melanophilin and thereby with Rab27a on the vesicle membrane (Fukuda et al., 2002, Hume et al., 2002, Matesic et al., 2001, Wu et al., 2002). While melanophilin does not look like a scaffolding protein based on domain analysis, it does serve as a motor cargo linker and further analysis may indicate its ability to function as an adaptor/scaffolding protein. Taken together, these studies indicate a general concept for motor-dependent vesicle trafficking in which the linkage of myosin and kinesin motors to their membrane-bound cargoes is mediated by soluble scaffolding proteins.

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Other scaffolding complexes These examples have indicated that individual scaffolding proteins can act as motor cargo linkers. In these cases, the connection between the motor and the motor cargo linker seems to be broken at the destination so that the scaffolding protein can carry out other functions. However, several examples exist of scaffolding complexes that are not released at the destination and therefore seem to function primarily to link microtubule-based motor proteins to their cargoes. One of the best examples comes from studies on intraflagellar transport (IFT) in which flagellar components are carried bidirectionally along microtubules by kinesin II and cytoplasmic dynein (Fig. 16.2; Rosenbaum et al., 1999). Kinesin II binds to a multiprotein complex, the IFT raft, which is localized between the microtubules of the axoneme and the flagellar membrane. The IFT raft binds to flagellar precursor materials to form the IFT particle that is carried to the tip of the axoneme. Upon reaching the destination, it is thought that the IFT raft links to cytoplasmic dynein for the return trip. Mutants lacking either the kinesin II motor protein or the IFT raft complex are defective in transport of membranous cargo and have short or missing flagella (Cole et al., 1993, Kozminski et al., 1995, Pazour et al., 2000). The IFT raft functions as a scaffolding complex, in that it binds to both the kinesin II motor and to the membranous cargo, thus providing the motor cargo linkage. As kinesin II and IFT rafts play a critical role in the transport of cargo material required for the formation and function of motile and non-motile cilia in a variety of cells and organisms, an attractive model is that the IFT raft functions as a scaffold upon which a variety of kinesin II-dependent cargoes can be attached depending on cellular context. Further work is needed to understand the molecular mechanisms by which the IFT raft links to both kinesin II and membrane cargo proteins as well as the role of the IFT raft in dynein-dependent retrograde transport. Another example of a scaffolding complex that can act as a motor cargo linker comes from studies on cytoplasmic dynein. Associated with cytoplasmic dynein is a complex of proteins, the dynactin complex, which is likely to play multiple roles in dynein-dependent transport (Allan, 2000, Holleran et al., 1998). Dynactin clearly functions as an activator of dynein activity and processivity (King and Schroer, 2000) which may be due at least in part to dynactin’s ability to bind to the microtubule tracks. Dynactin’s role as a scaffolding complex and motor cargo linker are due to its ability to bind multiple proteins, including binding directly to the IC subunit of the dynein motor protein as well as to cytoplasmic and membrane cargo proteins such as spectrin, a component of the membrane-associated skeleton, and BICD2, the mammalian homolog of the Drosophila gene Bicaudal-D involved in microtubule organization and transport of polarity factors during oocyte differentiation (Holleran et al., 2001, Hoogenraad et al., 2001, Kumar et al., 2001, Muresan et al., 2001). The role of dynactin as a motor cargo linker may be to mediate dynein binding to a variety of cargoes via its interaction with the spectrin network (Section 16.5.3) whereas the individual dynein subunits mediate specific interactions with distinct cargoes. Clearly, the exact role that dynactin plays as a scaffolding complex linking dynein to its cargoes requires further study. 16.5.1.1


16.5.2

Motor Cargo Linkage via Members of the Rab Family of Small G-proteins

Small monomeric guanosine triphosphate (GTP)-binding proteins play a wide range of regulatory functions in cells. Rab proteins, which represent the largest family of monomeric ras-related GTPases, play a crucial role in determining the specificity of membrane transport steps in eukaryotic cells by directing vesicle budding and membrane fusion reactions (Zerial and McBride, 2001). A general role for G-proteins in microtubule-based membrane trafficking events was first suggested by the observation that the non-hydrolyzable GTP analog GTPgS reduced both anterograde and retrograde transport in squid axoplasm (Bloom et al., 1993) or in Golgi-derived membranes (Fullerton et al., 1998). The exact role that small G-proteins play in microtubule- or actin-based transport is unclear. The Rab protein or its downstream effectors may function to regulate the activity of motor proteins bound to the same vesicle membrane. Alternatively, the Rab protein may function as part of the receptor or motor cargo linker that recruits the motor to the correct transport vesicle. Either mechanism would provide a link between vesicle formation and vesicle transport that is controlled by the nucleotide state of the GTPase switch. Although an attractive possibility, the actual sequence of events in docking of motor proteins to their cargoes and activation of motor activity, remains largely unknown (Section 16.6). Recent results have begun to shed some light on exactly which G-proteins are involved and how they regulate actin- and microtubule-based motor proteins (Fig. 16.3 and Tab. 16.2; Hammer and Wu, 2002). The best evidence of a functional role for small G-proteins in regulating motor proteins comes from studies on interactions between Rabs and the myosin family of motor proteins. In mice, altered coat color in the dilute, ashen and leaden mutants is due to defective transport of melanosomes, the pigment-containing organelle in melanocytes, to neighboring keratinocytes and eventually into coat hairs. A large body of work has pointed to a functional interaction between Rab27a, the product of the ashen locus, and myosin Va, the product of the dilute locus (Hume et al., 2001, Wilson et al., 2000, Wu et al., 2001). Recent studies have begun to delineate the molecular interactions and sequence of events involved in Rab27a-dependent recruitment of myosin Va to melanosomes in mouse melanocytes. The new results demonstrate that Rab27a does not bind directly to myosin Table 16.2

Motor proteins whose motor cargo linkage is regulated by Rab proteins.

Motor protein

Rab protein

Reference

Rabkinesin-6

Rab6

Echard et al. (1998)

Dynein LIC

Rab4

Bielli et al. (2001)

Myosin Va

Rab27a

Hume et al. (2001)

Myosin Vb

Rab11a

Lapierre et al. (2001)

393

394


Va, but rather Rab27a in its active, GTP-bound form on the melanosome surface recruits melanophilin, the product of the leaden locus, which in turn recruits myosin Va (Fukuda et al., 2002, Hume et al., 2002, Matesic et al., 2001, Wu et al., 2002). The motor cargo linker, melanophilin, can be considered to be an adaptor protein and thus fits the general concept presented above (Section 16.5.1). These results are important for two reasons: (a) they are the first to detail the molecular mechanisms by which a myosin motor binds to its cargo; and (b) they indicate that the role of the Rab proteins in controlling membrane trafficking may be not only in ensuring the specificity of membrane fusion, but also to coordinate formation of a vesicle with the recruitment of its motor. Further work is needed to determine whether this interaction is sufficient to activate myosin Va on the surface of the melanosome. In addition, it remains to be determined whether the Rab27a/melanophilin/myosin Va complex is present on melanosomes moving bidirectionally on microtubules or whether this complex is recruited to the melanosome surface only after it has reached the actin-rich periphery of the cell. However, these results provide an attractive model for how Rab proteins and their effectors function to regulate such events as budding, transport and fusion of organelles and vesicles in membrane trafficking. In addition to recruiting myosin motors indirectly, Rab proteins may also bind directly to motor proteins. Another member of the myosin V class, myosin Vb, appears to bind directly and specifically to the GTP-bound form of a different Rab protein, Rab11a, which regulates the return of membrane components from the early endosomes to the plasma membrane. Rab11a and myosin Vb co-localize to an endosomal compartment and over-expression of a dominant negative myosin Vb tail domain reduces receptor recycling (Lapierre et al., 2001). Finally, a class V myosin required for polarized growth in budding yeast, Myo2p, interacts by genetic and biochemical analyses with a Rab protein, Sec4p. Mutation of either protein causes the accumulation of vesicles in the mother cell, consistent with a role for these proteins in movement of secretory vesicles on actin cables from the mother cell to the bud (Finger and Novick, 1998, Pruyne and Bretscher, 2000, Pruyne et al., 1998, Schott et al., 1999). Rab protein regulation of members of the myosin V class of myosin motor proteins involves transport events occurring at or close to the plasma membrane. As short-range vesicular movement utilizes myosin motors on actin filaments enriched in the cell periphery, it is tempting to speculate that the regulation of myosin motor activity by Rab proteins will be a general concept for transport events occurring at the cell surface. However, several reports have indicated that Rab proteins are involved in regulating microtubule-based transport events as well. The first report of molecular mechanism linking Rabs to kinesin motor proteins came from studies on Rab6, a family member involved in directing the retrograde trafficking of protein and lipids from the Golgi complex to the ER. The active, GTPbound form of Rab6 interacts with Rabkinesin-6, a member of the kinesin superfamily, and in this way is thought to regulate the movement of Golgi-derived vesicles along microtubules (Echard et al., 1998). However, recent studies on the human homolog of Rabkinesin6, Rab6-KIFL, suggest that this kinesin family


member functions during formation of the cleavage furrow and cytokinesis and is found solely in cells during mitosis (Fontijn et al., 2001, Hill et al., 2000). Thus, the exact role of Rab6-KIFL in Golgi trafficking and/or mitosis remains to be determined. Several other studies suggest a link between small G-proteins and microtubulebased motors. Rab5 regulates not only docking and fusion events in the trafficking of early endosomes, but also stimulates the motility of early endosomes toward the minus end of microtubules. Surprisingly, this motility is thought to be driven not by cytoplasmic dynein, but by a minus end-directed kinesin family member yet to be identified (Nielsen et al., 1999). A different member of the Rab family of small G-proteins, Rab7, regulates late endosome/lysosome trafficking perhaps at least in part by regulating dynein- and kinesin-dependent motility of late endosomes. An increase in membrane-associated Rab7 results in vesicles that move towards the cell center and stay there presumably because they cannot use a kinesin family member for transport to the cell periphery (Lebrand et al., 2002). The effects of Rab7 may be mediated by the Rab7 effector protein RILP (Rab-interacting lysosomal protein) which has been demonstrated to play a role in the recruitment of dynein dynactin motor complexes, but not a kinesin motor, to endosomal membranes (Jordens et al., 2001). 16.5.3

Other Mechanisms for Linking Microtubule-based Motors to their Cargoes Attachment to the membrane cytoskeleton A membrane-associated cytoskeleton that imparts structural integrity and mechanical stability to the cell was first described for the erythrocyte plasma membrane (Bennett and Gilligan, 1993). A key component of this membrane skeleton is the protein spectrin, a multi-functional protein that binds to actin filaments as well as to membrane proteins (via ankyrin) and acidic phospholipids. Through these interactions, the spectrin skeleton restricts the lateral diffusion of proteins in the membrane and thus generates specialized membrane domains. In this sense, spectrin can be considered to be a multi-functional scaffold linking membrane proteins (Section 16.5.1), cytoplasmic proteins and the major cytoskeletal filaments systems. Recently it has become clear that a spectrin-based membrane skeleton is associated with intracellular organelles in non-erythroid cells and plays a variety of roles in organelle membrane stability and protein organization as well as in membrane trafficking (Bennett and Chen, 2001, De Matteis and Morrow, 2000). A considerable amount of evidence has accumulated implicating spectrin in linking microtubule motor proteins to membranes (Tab. 16.3; Holleran and Holzbaur, 1998, Lippincott-Schwartz, 1998). Dynein, dynactin, Golgi membranes and the Golgi-associated bIII-spectrin isoform co-purify and co-localize in a variety of systems (Beck et al., 1997, Holleran et al., 1996, 2001). Disruption of either dynein/ dynactin or spectrin inhibits vesicle transport through the Golgi complex (Burkhardt et al., 1997, Devarajan et al., 1997, Presley et al., 1997). More recently, 16.5.3.1

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396

16.5 Molecular Mechanisms of Motor–Cargo Linkage Table 16.3

Motor proteins that bind to transmembrane (TM) or spectrin proteins.

Motor protein

Binding partner (type)

Reference

Kinesin I

Kinectin (TM)

Ong et al. (2000)

Kinesin I

APP (TM)

Kamal et al. (2000)

Kinesin II

Fodrin (spectrin)

Takeda et al. (2000)

Dynein

Rhodopsin (TM)

Tai et al. (1999)

Dynein

TrkA (TM)

Yano et al. (2001)

Dynein

Spectrin (TM)

Holleran et al. (2001), Muresan et al. (2001)

Myosin Va

Synaptophysin (TM)

Prekeris and Terrian (1997)

Myosin Va

Synaptobrevin (TM)

Ohyama et al. (2001)

Myosin VIIa

Vezatin (TM)

Kussel-Andermann et al. (2000)

it has been shown that the spectrin skeleton links dynein/dynactin to the membrane. Dynein and the dynactin complex can bind to phospholipid vesicles via spectrin and ankyrin (Muresan et al., 2001). This is likely due to a direct interaction between the Arp1 subunit of dynactin and bIII spectrin (Holleran et al., 2001). Taken together, these results suggest that dynein-mediated transport of Golgi membranes may be mediated at least in part by dynactin-dependent interaction with the spectrin matrix. At least some members of the kinesin family may also use a spectrin-based membrane skeleton to link to their cargoes. The KAP3 accessory subunit of the kinesin family member KIF3 was found in a two-hybrid screen to interact with a-fodrin, a neuronal form of a-spectrin. KIF3 and fodrin co-localize to the same vesicles in neurons and are transported down axons at similar rates (Takeda et al., 2000). Whether the involvement of spectrin or other membrane cytoskeleton components contributes to the cargo binding of other kinesin family members, remains to be determined. However, it is tempting to speculate that such interactions could provide a general mechanism of linking motors to their cargoes that would work in concert with the specific protein protein interactions that link motors to particular cargoes.

Attachment via integral membrane proteins For a long time it was thought that motors would attach to their membranous cargoes via transmembrane proteins that serve as receptors for a specific motor protein. Although the general concept from recent studies suggests that motors link indirectly to their cargoes via adaptor or scaffolding proteins, several studies have indeed demonstrated a direct interaction between motors and transmembrane cargo molecules (Tab. 16.3). The KHC subunit of kinesin-I binds to kinectin, 16.5.3.2


an integral membrane protein localized to the ER, however, whether kinectin functions in kinesin-driven transport is controversial (Ong et al., 2000, Plitz and Pfeffer, 2001). The KLC subunit of kinesin-I binds to the cytoplasmic tail of the Alzheimers Precursor Protein (APP), a Type I membrane protein whose accumulation in the plaques of Alzheimer patients is thought to contribute to progression of the disease (Kamal et al., 2000). The function of APP as a motor receptor would link kinesin-I to axonal vesicles containing presenilin-1, BACE, GAP43 and TrkA and, in addition, proteolytic cleavage of APP in the vesicles could release kinesin-I from the membrane and thereby terminate vesicle transport (Kamal et al., 2001). This is an attractive model, however, it should be pointed out that APP probably does not link kinesin-I directly to the membrane, as the cytoplasmic tail of APP binds directly to the soluble JIP-1 scaffolding proteins which have been demonstrated to act as motor cargo linkers for kinesin I (Section 16.5.1; Matsuda et al., 2001, Scheinfeld et al., 2002). Cytoplasmic dynein can apparently also bind directly to transmembrane proteins. The DLC Tctex-1 subunit of cytoplasmic dynein has been shown to interact with the cytoplasmic tail of rhodopsin, an interaction presumed to be responsible for dynein-dependent transport of rhodopsin along microtubules to the outer segment of the photoreceptor cell. Surprisingly, this interaction does not require the dynactin complex (Tai et al., 1999). The Tctex-1 subunit has also recently been suggested to bind directly to the cytoplasmic tail of the nerve growth factor (NGF) receptor TrkA and thereby allow dynein to drive retrograde transport of activated NGF receptors from the axon terminal to the cell body (Yano et al., 2001). Finally, a myosin family member, myosin VII, interacts with the transmembrane protein vezatin and co-localizes with vezatin at adherens junctions (Kussel-Andermann et al., 2000). Future studies are required to demonstrate the physiological significance of these interactions and to determine whether a direct interaction with transmembrane proteins is indeed one way that motor proteins link to their cargoes.

16.6

Regulation of Motor Activity 16.6.1

Motor Proteins must be Regulated at Several Steps of their Transport Cycle

The activity of the kinesin, dynein and myosin motor proteins must be tightly regulated in cells to ensure not only the proper trafficking of organelles and vesicles, but also to prevent unnecessary ATP hydrolysis and cytoskeletal binding by cargo-less motors. Although it makes biological sense that motor activation should be coupled to cargo binding, evidence for this point of view is scant. In fact, one of the most poorly understood issues at present is the actual sequence of events in docking of motor proteins to their cargoes and activation of motor activity. Regulation must occur at several levels: inhibition of enzymatic activity until the motor is needed, connecting each motor to the correct cargo, activation of motor activity to

397

398

16.6 Regulation of Motor Activity

produce directed cargo movement, and finally inactivation of the motor at the destination. Some regulatory events, such as inhibition of cargo-less motors, may be common to many different motors whereas others are likely to be motor specific and related to cargo binding (see Chapter 17 in this volume by Haimo and recent review by Reilin et al., 2001). Motor proteins must be kept inactive until needed to prevent futile ATP hydrolysis and movement on microtubule and actin filaments. Several kinesin family members have been found to be inactive when not bound to cargo. This is due to a folded conformation in which the tail domain of the motor directly interacts with and inhibits the motor domain. Members of the myosin family of motor proteins have also been found in a folded inhibited state. It is tempting to speculate that self-inhibition is a general mechanism for keeping kinesin and myosin motors inactive until needed. Little is known about how the activity of dynein motors is regulated except that the dynactin complex is involved. Cargo attachment needs to be tightly regulated to ensure that each motor protein carries the appropriate cargo to the correct cellular destination. A variety of genetic and biochemical approaches have begun to identify the molecules involved in linking motor proteins to their cargoes (Section 16.5). How these interactions are regulated in cells such that motor cargo binding is permitted at the point of departure and subsequently terminated at the destination is unknown. One possibility is that modification of the cargo or of some of its components, such as the phosphorylation of a vesicle protein or a modification of the lipid composition of a nascent vesicle, may expose binding sites for the motor. An attractive mechanism emerging from recent studies is that binding of motor proteins to their vesicular cargoes is regulated by the nucleotide state of the Rab GTPase switch proteins (Section 16.5). Alternatively, post-translational modification and/or a shape change of the motor molecule itself may induce its binding to the cargo. Similarly unclear is how motor activity is activated once the proper motor cargo linkage has taken place. Theoretically, cells can regulate vesicle transport by regulating the binding of a motor to its cargo, by modulating the activity of a motor after it has bound to the cargo, or both. As several members of the kinesin and myosin families have been found in an inactive self-inhibited state, the simplest model would propose that cargo binding to the tail domain of the motor releases its interaction with the motor domain and thereby relieves the self-inhibition. However, the fact that cargoes contain multiple motors but move in a directed manner suggests that motor track interactions are regulated as well as motor cargo interactions (Section 16.4). Indeed, studies on myosin V and dynein show that the motors can be regulated at both levels: binding of the motor to cargo and regulation of its activity once bound (Evans et al., 1998, Lane and Allan, 1999, Roghi and Allan, 1999, Schott et al., 1999). Thus it seems most likely that cargo binding to the tail domain of myosin and kinesin family members is not sufficient to activate the motor but rather that subsequent events and/or proteins are required to activate the motor after it has bound (Verhey and Rapoport, 2001). Once the transport cargo has reached its destination, the motors used during transport are no longer needed. It is unclear how these motors are released


from the transport cargo and/or are inactivated. Likewise, the fate of the released motor is not known. One possibility is that motors are degraded at end of the track. This is supported by immunolocalization data demonstrating that kinesins and anterograde cargoes accumulate on the proximal side (near the cell body) of ligated axons but not on the distal side (near the end of the track). This suggests that the anterograde kinesin motors do not make the return trip to the cell body but are instead degraded at the axon terminal (Dahlstrom et al., 1991, Hirokawa et al., 1991, Kondo et al., 1994, Noda et al., 1995, Okada et al., 1995, Yamazaki et al., 1995, Yang and Goldstein, 1998). Another possibility is that motors become inactivated and return to the point of departure as components of cargoes carried by motor proteins moving in the opposite direction. This is supported by mutants of kinesin that accumulate at the distal ends of neurites and hyphae (Seiler et al., 2000, Verhey et al., 1998, 2001) as well as by the observations that motors can be carried in an inactive state as cargo components by other motors (Section 16.4). 16.6.2

Molecular Mechanisms for Regulating Motor Activity

A variety of mechanisms have been proposed to regulate the activity of the many motor proteins in cells (see Chapter 17 by Haimo and review by Reilin et al., 2001). One mechanism that regulates the cargo binding and enzymatic activity of myosin motor proteins is changes in calcium concentration. As kinesin I can be activated by small changes in pH, local changes in ion concentration may be one mechanism that contributes to the regulation of motor activity at the right time and place. Another possibility is that molecular chaperone proteins localized to the destination interact with the motor and release it from its cargo (Tsai et al., 2000). A mechanism that has emerged from a variety of recent studies is that the motors that carry vesicular cargoes are regulated by members of the Rab family of small GTPases (Section 16.5.2). The nucleotide state of the Rab GTPase switch proteins, known to regulate vesicle formation, tethering, docking and fusion, may logically also regulate the recruitment of motor proteins and activation of vesicle motility. In this model, at the point of departure, an organelle-specific Rab protein in its GTP-bound active state would recruit the appropriate motor protein to the vesicular cargo and then, in its GDP-bound inactive state at the destination, release the motor from the membrane. This is an attractive possibility and will be an exciting area of further research. Another mechanism for regulating motor protein activity that has been recognized for many years but not at all understood, is the role of phosphorylation. Many motor proteins are known to be phosphorylated or to bind to a kinase or phosphatase or even to contain kinase domains in their primary sequence (Thaler and Haimo, 1996, Reilin et al., 2001). In most cases, the relationship of these findings to cargo binding and motor activity is unclear. There are, however, several demonstrations of a functional relationship between phosphorylation, or at least signaling proteins, and motor proteins. In non-muscle cells, reversible phosphorylation by myosin light chain kinase (MLCK) has been demonstrated to control the

399

400


unfolding and activation of myosin II (Bresnick, 1999). In fish melanophores, hormones regulate cAMP and Ca2 levels, which in turn modulate phosphatase and kinase activities that regulate the activity of kinesin II and myosin motor proteins and thereby drive pigment granule aggregation and dispersion in response to extracellular stimuli (Haimo and Thaler, 1994). In neurons, the protein kinase GSK3 phosphorylates and releases kinesin I from membrane-bound organelles (Morfini et al., 2002). In mitotic cells, p34cdc2 kinase phosphorylates the LIC subunit of dynein, releasing it from membranes and thus inhibiting dynein-dependent motiltiy (Dell et al., 2000, Niclas et al., 1996). The recent concept that motors link to their cargoes via scaffolding proteins suggests that signaling molecules that regulate motor activity may be assembled on the motor’s cargo. Indeed, genetic evidence in C. elegans supports a role for the kinesin I motor cargo linker JIP-3/Syd/UNC-16, as well as several associated JNK signaling proteins, in the control of axonal transport (Byrd et al., 2001). During Drosophila development, a large multi-protein complex that includes the kinesin family member Costal 2, the serine/threonine kinase Fused, and the transcription factor Cubitis Interruptus binds to microtubules in the absence of the Hedgehog protein. Hedgehog signaling results in decreased binding of the complex to microtubules, suggesting that Fused or other kinases may phosphorylate Costal 2 and regulate its activity (Ascano et al., 2002, Robbins et al., 1997, Sisson et al., 1997).

16.7

Concluding Remarks

Motor proteins have been known to be involved in organelle organization and vesicle trafficking for many years. Recent exciting work has begun to reveal the molecular mechanisms by which motors link to their appropriate cargoes. Future work in this area will validate the two suggested general concepts for motor cargo linkers, scaffolding proteins and Rab proteins, and will lead to a clearer picture of which motor protein is carrying which transport cargo. Particularly exciting has been the molecular links between motor proteins and the molecules known to regulate vesicle trafficking, such as the clathrin adaptor proteins and the Rab proteins, as well as the links between motor proteins and signal transduction. In addition but outside the scope of this chapter, are the recent revelations on the contributions of motor proteins to RNA transport, mitosis and meiosis, as well as to human disease. A major focus of the future will be to determine the mechanisms that regulate motor cargo binding and motor activity. Particularly challenging will be the determination of how multiple motors on the same cargo are regulated. Answers to these questions will help us understand how the organization of animal cells is set up and maintained both temporally and spatially.


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Reilein, A. R., S. L. Rogers, M. C. Tuma, and V. I. Gelfand. 2001. Regulation of molecular motor proteins. Int. Rev. Cytol. 204: 179 238. Robbins, D. J., K. E. Nybakken, R. Kobayashi, J. C. Sisson, J. M. Bishop, and P. P. Therond. 1997. Hedgehog elicits signal transduction by means of a large complex containing the kinesin-related protein costal2. Cell 90: 225 234. Rodionov, V. I., A. J. Hope, T. M. Svitkina, and G. G. Borisy. 1998. Functional coordination of microtubule-based and actin-based motility in melanophores. Curr. Biol. 8: 165 168. Rogers, S. L. and V. I. Gelfand. 1998. Myosin cooperates with microtubule motors during organelle transport in melanophores. Curr. Biol. 8: 161 164. Rogers, S. L., I. S. Tint, P. C. Fanapour, and V. I. Gelfand. 1997. Regulated bidirectional motility of melanophore pigment granules along microtubules in vitro. Proc. Natl Acad. Sci. USA 94: 3720 3725. Roghi, C. and VJ. Allan. 1999. Dynamic association of cytoplasmic dynein heavy chain 1a with the Golgi apparatus and intermediate compartment. J. Cell Sci. 112: 4673 4685. Rosenbaum, J. L., D. G. Cole, and D. R. Diener. 1999. Intraflagellar transport: the eyes have it. J. Cell Biol. 144: 385 388. Schafer, D. A. 2002. Coupling actin dynamics and membrane dynamics during endocytosis. Curr. Opin. Cell Biol. 14: 76 81. Scheinfeld, M. H., R. Roncarati, P. Vito, P. A. Lopez, M. Abdallah, and L. D’Adamio. 2002. Jun NH2-terminal kinase (JNK) interacting protein 1 (JIP1) binds the cytoplasmic domain of the Alzheimer’s beta-amyloid precursor protein (APP). J. Biol. Chem. 277: 3767 3775. Schott, D., J. Ho, D. Pruyne, and A. Bretscher. 1999. The COOH-terminal domain of Myo2p, a yeast myosin V, has a direct role in secretory vesicle targeting. J. Cell Biol. 147: 791 808. Schroer, T. A. 1994. New insights into the interaction of cytoplasmic dynein with the actin-related protein, Arp1. J. Cell Biol. 127: 1 4. Schroer, T. A. 2001. Microtubules don and doff their caps: dynamic attachments at plus and minus ends. Curr. Opin. Cell Biol. 13: 92 96. Schuyler, S. C. and D. Pellman. 2001. Search, capture and signal: games microtubules and centrosomes play. J. Cell Sci. 114: 247 255.

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Tai, A. W., J. Z. Chuang, C. Bode, U. Wolfrum, and C. H. Sung. 1999. Rhodopsin’s carboxyterminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 97: 877 887. Tai, C. Y., D. L. Dujardin, N. E. Faulkner, and R. B. Vallee. 2002. Role of dynein, dynactin, and CLIP-170 interactions in LIS1 kinetochore function. J. Cell Biol. 156: 959 968. Takeda, S., H. Yamazaki, D. H. Seog, Y. Kanai, S. Terada, and N. Hirokawa. 2000. Kinesin superfamily protein 3 (KIF3) motor transports fodrin-associating vesicles important for neurite building. J. Cell Biol. 148: 1255 1265. Tanaka, Y., Y. Kanai, Y. Okada, S. Nonaka, S. Takeda, A. Harada, and N. Hirokawa. 1998. Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria. Cell 93: 1147 1158. Thaler, C. D. and L. T. Haimo. 1996. Microtubules and microtubule motors: mechanisms of regulation. Int. Rev. Cytol. 164: 269 327. Thiemann, M., M. Schrader, A. Volkl, E. Baumgart, and H. D. Fahimi. 2000. Interaction of peroxisomes with microtubules. In vitro studies using a novel peroxisome-microtubule binding assay. Eur. J. Biochem. 267: 6264 6275. Tsai, M. Y., G. Morfini, G. Szebenyi, and S. T. Brady. 2000. Release of kinesin from vesicles by hsc70 and regulation of fast axonal transport. Mol. Biol. Cell 11: 2161 2173. Tuma, M. C., A. Zill, N. Le Bot, I. Vernos, and V. Gelfand. 1998. Heterotrimeric kinesin II is the microtubule motor protein responsible for pigment dispersion in Xenopus melanophores. J. Cell Biol. 143: 1547 1558. Valetti, C., D. M. Wetzel, M. Schrader, M. J. Hasbani, S. R. Gill, T. E. Kreis, and T. A. Schroer. 1999. Role of dynactin in endocytic traffic: effects of dynamitin overexpression and colocalization with CLIP-170. Mol. Biol. Cell. 10: 4107 4120. Vaughan K. T. and R .B. Vallee. 1995. Cytoplasmic dynein binds dynactin through a direct interaction between the intermediate chains and p150Glued. J. Cell Biol. 131: 1507 1516. Vaughan, K. T., S. H. Tynan, N. E. Faulkner, C. J. Echeverri, and R. B. Vallee. 1999. Colocalization of cytoplasmic dynein with dynac-

16 Motors and Membrane Trafficking tin and CLIP-170 at microtubule distal ends. J. Cell Sci. 112: 1437 1447. Verhey, K. J., D. L. Lizotte, T. Abramson, L. Barenboim, B. J. Schnapp, and T. A. Rapoport. 1998. Light chain-dependent regulation of kinesin’s interaction with microtubules. J. Cell Biol. 143: 1053 1066. Verhey, K. J., D. Meyer, R. Deehan, J. Blenis, B. J. Schnapp, T. A. Rapoport, and B. Margolis. 2001. Cargo of kinesin identified as JIP scaffolding proteins and associated signaling molecules. J. Cell Biol. 152: 959 970. Verhey, K. J. and T. A. Rapoport. 2001. Kinesin carries the signal. Trends Biochem. Sci. 26: 545 550. Walther, Z., M. Vashishtha, and J. L. Hall. 1994. The Chlamydomonas FLA10 gene encodes a novel kinesin-homologous protein. J. Cell Biol. 126: 175 188. Welte, M. A., S. P. Gross, M. Postner, S. M. Block, and E. F. Wieschaus. 1998. Developmental regulation of vesicle transport in Drosophila embryos: forces and kinetics. Cell 92: 547 557. Wicks, S. R., C. J. de Vries, H. G. van Luenen, and R. H. Plasterk. 2000. CHE-3, a cytosolic dynein heavy chain, is required for sensory cilia structure and function in Caenorhabditis elegans. Dev. Biol. 221: 295 307. Williams, D. S. 2002. Transport to the photoreceptor outer segment by myosin VIIa and kinesin II. Vision Res. 42: 455 462. Wilson, S. M., R. Yip, D. A. Swing, T. N. O’Sullivan, Y. Zhang, E. K. Novak, R. T. Swank, L. B. Russell, N. G. Copeland, and N. A. Jenkins. 2000. A mutation in Rab27a causes the vesicle transport defects observed in ashen mice. Proc. Natl Acad. Sci. USA 97: 7933 7938. Wu, X., B. Bowers, Q. Wei, B. Kocher, and J. A. Hammer, III. 1997. Myosin V associates with melanosomes in mouse melanocytes: evi-

dence that myosin V is an organelle motor. J. Cell Sci. 110: 847 859. Wu, X., B. Kocher, Q. Wei, and J. A. Hammer, III. 1998. Myosin Va associates with microtubule-rich domains in both interphase and dividing cells. Cell Motil. Cytoskelet. 40: 286 303. Wu, X., K. Rao, M. B. Bowers, N. G. Copeland, N. A. Jenkins, and J. A. Hammer, III. 2001. Rab27a enables myosin Va-dependent melanosome capture by recruiting the myosin to the organelle. J. Cell Sci. 114: 1091 1100. Wu, X. S., K. Rao, H. Zhang, F. Wang, J. R. Sellers, L. E. Matesic, N. G. Copeland, N. A. Jenkins, and J. A. Hammer, III. 2002. Identification of an organelle receptor for myosinVa. Nature Cell Biol. 4: 271 278. Yamazaki, H., T. Nakata, Y. Okada, and N. Hirokawa. 1995. KIF3A/B: a heterodimeric kinesin superfamily protein that works as a microtubule plus end-directed motor for membrane organelle transport. J. Cell Biol. 130: 1387 1399. Yang, Z. and L. S. Goldstein. 1998. Characterization of the KIF3C neural kinesin-like motor from mouse. Mol. Biol. Cell 9: 249 261. Yano, H., F. S. Lee, H. Kong, J. Chuang, J. Arevalo, P. Perez, C. Sung, and M. V. Chao. 2001. Association of Trk neurotrophin receptors with components of the cytoplasmic dynein motor. J. Neurosci. 21: RC125. Zerial, M. and H. McBride. 2001. Rab proteins as membrane organizers. Nature Rev. Mol. Cell Biol. 2: 107 117. Zhou, H. M., I. Brust-Mascher, and J. M. Scholey. 2001. Direct visualization of the movement of the monomeric axonal transport motor UNC-104 along neuronal processes in living Caenorhabditis elegans. J. Neurosci. 21: 3749 3755.

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17 Regulation of Molecular Motors Leah T. Haimo

17.1

Introduction

Cells move organelles towards their cell center and away from their cell center, they activate and repress organelle movements, they move chromosomes towards the equator and then away from the equator, and they build a spindle and then destroy the spindle (see Chapter 16 by Verhey and Chapter 14 by Scholey and Mogilner). All of these events require specific molecular motors which in turn must be regulated so that they function when needed and do not interfere with other movements when not needed. This chapter will focus on the mechanisms that regulate microtubule motors. However, because microtubule motors interact with myosin V, regulation of this motor is also discussed. Two general mechanisms effect motor regulation. A regulatory event may recruit motors to membranous organelles or to specific sites in the spindle. Alternatively, a regulatory event may modify motor activity so that its ability to generate force is enhanced or inhibited.

17.2

The Role of Phosphorylation in Regulating Molecular Motors

Phosphorylation is the most widely utilized regulatory mechanism within cells. Molecular motors, or the proteins with which they interact, are substrates for kinases and phosphatases, and, in many cases, changes in phosphorylation have been correlated with changes in localization of the motor or in changes in the transport of cargo by that motor. In other cases, phosphorylation has been correlated with a change in motor activity.

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17.2 The Role of Phosphorylation in Regulating Molecular Motors

17.2.1

Phosphorylation can Control Motor Organelle or Motor Spindle Binding Interaction of dynein with organelles can be regulated by phosphorylation A number of studies suggest that attachment of motors to membranous organelles and to sites within the spindle can be regulated by phosphorylation. Serum starvation of fibroblast cells resulted in a loss of cytoplasmic dynein from lysosomes and an increase in phosphorylation of its heavy chain, suggesting that phosphorylation of dynein induces its dissociation from organelles (Lin et al., 1994). In Xenopus egg extracts, the extent of organelle transport, which appears to be primarily towards the minus ends of microtubules, is significantly dampened when that extract is converted from an interphase to a metaphase extract (Allan and Vale, 1991). Cytoplasmic dynein is the primary minus-end microtubule motor in the extract, and both dynein and dynactin dissociate from membranes in metaphase extracts. The light intermediate chain of dynein becomes phosphorylated in the metaphase extracts, suggesting that phosphorylation of this subunit of dynein may mediate the release of dynein from organelles (Niclas et al., 1996). Cdc2 kinase phosphorylates dynein light intermediate chain in vitro, depletion of Cdc2 kinase from Xenopus metaphase extracts results in a loss of dynein light intermediate chain phosphorylation, and incubation of membranes isolated from interphase egg extracts with this kinase results in the dissociation of dynein from the membranes. These studies suggest that Cdc2 kinase phosphorylates dynein light intermediate chain in vivo at metaphase when this kinase becomes activated and induces dynein dissociation from membrane-bound organelles (Addinall et al., 2001, Dell et al., 2000). Serine 197 on the light intermediate chain may be the target of Cdc2 (Dell et al., 2000). However, this site may remain phosphorylated during interphase and, instead, one of three other possible consensus sites for Cdc2 present within amino acids 379 405 may be the targeted site (Addinall et al., 2001). 17.2.1.1

Interaction of dynein with dynactin can be regulated by phosphorylation The interaction between dynein and dynactin can be regulated by phosphorylation (Vaughan et al., 2001), and, if dynactin is required for dynein membranous organelle binding (Gill et al., 1991, Karki and Holzbaur, 1999), then such phosphorylation events could regulate dynein binding to organelles and thereby regulate organelle transport. The amount of dynactin that co-immunoprecipitates with dynein from Xenopus egg extracts is greatly reduced when phosphatase inhibitors are included in the extracts, a finding which suggests that a phosphorylation event dissociates dynein from dynactin (Niclas et al., 1996). Indeed, recent studies demonstrate that phosphorylation of dynein’s intermediate chain, at serine 84, mediates this dissociation of dynein from dynactin. Further, this phosphorylation correlates inversely with the ability of dynein to transport organelles in vivo (Vaughan et al., 2001). The kinase that phosphorylates dynein intermediate chain at serine 84 was not identified in this study, but at least one kinase, casein kinase II, binds to dynein intermediate chain and phosphorylates it in vitro (Karki et al., 1997). 17.2.1.2

17 Regulation of Molecular Motors

Dynactin, as well as dynein, dissociates from membranes in Xenopus egg metaphase extracts, although only phosphorylation of dynein light intermediate chain was observed (Niclas et al., 1996). These findings would be difficult to reconcile with a model in which dynactin is required for dynein to bind to organelles because, a priori, phosphorylation of a subunit of dynein would not be expected to affect dynactin’s ability to bind to membranes. However, other studies indicate that the dynein intermediate chain and the p150 subunit of dynactin also become hyperphosphorylated during mitosis relative to interphase (Huang et al., 1999a), and it is possible, but not yet demonstrated, that phosphorylation of the p150 subunit is responsible for the dissociation of dynactin from membranes observed earlier (Niclas et al., 1996). The p150 subunit of dynactin can also become phosphorylated upon treatment of cells with activators of PKA or of PKC, and these treatments correlate with an increase in vesicle transport, both anterograde and retrograde (Farshori and Holzbaur, 1997), so it is unlikely that this phosphorylation of p150 induces its dissociation from membranes. It has yet to be determined whether p150 or other subunits of dynactin can be phosphorylated at different sites by specific kinases and, as a result, change its behavior with respect to dynein binding, microtubule binding, or membrane binding. Dynactin can also interact with a kinesin related protein, Eg5 (Blangy et al., 1997), and so phosphorylation of dynactin might also affect Eg5 function.

Interaction of kinesin with organelles can be regulated by phosphorylation Interaction of kinesin with membranes may be enhanced or inhibited by phosphorylation. The heavy chain of kinesin, present on membranes isolated from fibroblasts, is more highly phosphorylated than is the heavy chain of kinesin obtained from the soluble pool in these same cells (Lee and Hollenbeck, 1995), suggesting that phosphorylation of kinesin results in its recruitment onto membranous organelles. Similarly, stimulation of pancreatic acinar cells to induce secretion of zymogen granules correlates with a three-fold increase in the amount of kinesin on the granules and a simultaneous hyperphosphorylation of kinesin heavy chain (Marlowe et al., 1998). Together these studies suggest that phosphorylation of kinesin heavy chain induces its recruitment to organelles. In contrast, addition of PKA to crayfish axoplasm results in a decrease in transport of small vesicles towards the plus ends of microtubules. This decrease in movement is correlated with an increase in phosphorylation of kinesin heavy chain and a decrease in the amount of kinesin bound to organelles, suggesting that phosphorylation of kinesin heavy chain by PKA results in its dissociation from vesicles (Okada et al., 1995). Like PKA, glycogen synthase kinase 3 induces a reduction in vesicle transport towards the plus ends of microtubules and results in a reduced amount of kinesin on organelles, but it phosphorylates kinesin light chain rather than kinesin heavy chain (Morfini et al., 2002). It will be necessary to identify sites on kinesin heavy and light chains that are phosphorylated by specific kinases in vivo and then determine how the presence of a phosphate group at each site influences kinesin behavior. 17.2.1.3

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Interaction of kinesin family members with the spindle can be regulated by phosphorylation The kinesin related protein Eg5 is required for normal spindle formation and progression through mitosis (Blangy et al., 1995, Sawin et al., 1992). Eg5 is concentrated on centrosomes and at the spindle poles during mitosis. A Cdc2 consensus phosphorylation site is present at threonine 937 on Xenopus Eg5 (thr 927 in human Eg5). This site is phosphorylated during mitosis in vivo and by Cdc2 in vitro, and mutagenesis of this site to alanine causes the motor to lose its ability to associate with the centrosomes (Blangy et al., 1995, Sawin and Mitchison, 1995). Most interestingly, dynactin binds via its p150 subunit to Eg5, and this interaction is enhanced by Cdc2 phosphorylation of Eg5 at threonine 927 (Blangy et al., 1997). Perhaps a motor complex of dynein, dynactin and Eg5 forms which allows these two opposing motors to be regulated. Eg5 can also be phosphorylated by the aurora-like kinase, pEg2, which also localizes to centrosomes. pEg2 phosphorylates Eg5 on a serine residue in the stalk domain of the protein, but the biological importance of this phosphorylation has not yet been determined (Giet et al., 1999). CENP-E is a kinesin related protein that localizes to kinetochores and is required for proper chromosome alignment (Schaar et al., 1997). At anaphase, CENP-E relocalizes to the mid-zone. This motor contains two microtubule binding domains, one in its motor domain and a second, ATP-insensitive binding site, at its carboxy terminus. The C-terminus also contains a Cdc2 consensus site, and a C-terminal fragment of CENP-E can be phosphorylated by Cdc2 in vitro, producing an identical phosphopeptide to that obtained from CENP-E phosphorylated in vivo. Most importantly, the ATP-insensitive microtubule binding is abolished upon phosphorylation. Therefore, phosphorylation of CENP-E by Cdc2 may mask the second microtubule binding site on CENP-E until Cdc2 is inactivated at the anaphase transition (Liao et al., 1994). This regulation may permit CENP-E to express a function at anaphase distinct from its role at metaphase, although no anaphase function for CENP-E has been clearly demonstrated. 17.2.1.4

The binding of myosin V to melanosomes is regulated by phosphorylation Pigment granules, or melanosomes, utilize both actin filaments and microtubules for transport to the cell periphery. Myosin V dissociates from melanosomes incubated in a metaphase, but not interphase, Xenopus egg extract (Karcher et al., 2001, Rogers et al., 1999). A serine residue at position1650 in the tail domain of myosin V is phosphorylated in the mitotic but not the interphase extracts. When this residue is altered to alanine, a mimic of the dephosphorylated state, the myosin tail binds to melanosomes regardless of the cell cycle phase of the extract. Conversely, when this residue is altered to glutamic acid, a mimic of the phosphorylated form of the protein, the myosin V tail cannot bind to melanosomes in either interphase or mitotic extracts. Calcium/calmodulin-dependent protein kinase II phosphorylates serine1650, and inhibitors of this kinase prevent the dissociation of the myosin V from melanosomes incubated in mitotic extracts (Karcher et al., 2001). Thus, myosin V’s interaction with melanosomes appears 17.2.1.5


Model of myosin Va release from melanosomes in mammals. Myosin Va attachment can be controlled by two different mechanisms. Rab27a binds to melanosomes via its geranylgeranyl lipid groups and recruits melanophilin and myosin Va to the organelle when GTP is bound to Rab27a (Wu et al., 2002). The melanosomes, with bound myosin Va, can be captured by actin filaments. Upon GTP hydrolysis by Rab27a, melanophilin and myosin Va are released from the organelle. This mechanism would regulate the ability of the mel-

Figure 17.1.

anosomes to interact with actin filaments, and might be utilized either to control movements of the melanosomes on actin or, alternatively, might be utilized to release melanosomes from the actin meshwork so that the melanosomes can be transferred to keratinocytes. During mitosis, myosin Va is released from melanosomes by a mechanism that involves phosphorylation of serine1650 in the tail domain of the molecule and is likely mediated by Ca2/calmodulin kinase II (Karcher et al., 2001). Drawing modified from Wu et al., 2002.

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to be regulated by a phosphorylation event during the cell cycle. As discussed in Section 17.3.1.1, the interaction of myosin V with melanosomes is also regulated by the GTP-bound state of a G protein. Accordingly, a single motor may be regulated by multiple mechanisms (Fig. 17.1). 17.2.2

The Activity of Motors may be Regulated by Phosphorylation Phosphorylation of axonemal dynein inhibits its activity The ability of dynein family or kinesin family members to bind to and/or to generate a force along microtubules may be regulated by phosphorylation. The best characterized example of regulation utilizing such a mechanism derives from studies of axonemal dynein in Chlamydomonas flagella (Habermacher and Sale, 1996, 1997, Howard et al., 1994, Yang and Sale, 2000). Phosphorylation of the intermediate chain (IC138) of inner arm dynein 1 (I1) inhibits microtubule sliding (Habermacher and Sale, 1997). Casein kinase I may be the responsible kinase, as casein kinase I inhibitors block phosphorylation of IC138 and also block the resulting inhibition of microtubule sliding. Casein kinase I appears to be anchored on the outer doublet microtubules, likely in a position where it can directly interact with and phosphorylate I1 (Yang and Sale, 2000). Protein kinase A (PKA) also inhibits dynein activity (Howard et al., 1994), and a PKA-anchoring protein (AKAP) is localized on the radial spokes (Gaillard et al., 2001). Moreover, protein phosphatase 1 is required to reverse the kinase-mediated inhibition of dynein activity (Habermacher and Sale, 1996). PP1 localizes to the central pair and may function via a signaling pathway that includes PP2A, positioned on the outer doublets (Yang et al., 2000). The axoneme, therefore, represents a highly ordered and regulated motile apparatus in which the kinases and phosphatases that regulate dynein activity are anchored together with their substrates. 17.2.2.1

Phosphorylation can activate or inhibit cytoplasmic dynein The activity of cytoplasmic dynein may also be regulated by phosphorylation. In vivo studies using brain tissue suggested that the heavy chain of cytoplasmic dynein, present in an anterograde compartment, is hypophosphorylated compared to that of dynein present in the entire cytoplasmic pool. Because dynein in the anterograde compartment might be less active than dynein in a general pool would be, these studies suggest that phosphorylation of dynein heavy chain activates dynein (Dillman and Pfister, 1994). Similarly, interphase Xenopus egg extracts form an extensive network of membrane tubules that move along microtubules towards their minus ends. Okadaic acid, which inhibits phosphatase activity, enhances these membrane movements but does not alter the amount of dynein associated with the membranes (Allan, 1995). These studies suggest that phosphorylation activates dynein activity, although it is not known if dynein is the target of this phosphorylation nor has the responsible kinase been identified. 17.2.2.2


Dynein obtained from N. crassa mutants, lacking the p150 subunit of dynactin, has reduced ATPase activity and reduced ability to bind to microtubules relative to dynein obtained from wild-type cells. The dynein from these mutants is more highly phosphorylated, likely on its light chains, than is dynein from wild-type cells. l Phosphatase dephosphorylates this dynein and enhances its ATPase activity significantly (Kumar et al., 2000). It is possible that interaction of dynein with dynactin in vivo prevents phosphorylation of dynein light chains and thereby maintains dynein in its active state. If so, then dynein not associated with dynactin might be phosphorylated to keep the dynein in an inactive state so that it cannot interact non-productively with microtubules. Dynein isolated from insulin-treated adipocytes exhibits decreased microtubule binding activity relative to dynein isolated from untreated cells. Treatment of cells with a PI3 kinase inhibitor blocks the insulin-induced inhibition in dynein activity (Huang et al., 2001), but it is not known if dynein is a target of an insulin-induced phosphorylation cascade.

Phosphorylation can activate or inhibit the activity of members of the kinesin family Kinesin activity may be inhibited by phosphorylation of kinesin light chains. Treatment of cells with tumor necrosis factor (TNF) induces mitochondria to cluster perinuclearly (De Vos et al., 1998), a finding similar to the results obtained in cells which lack kinesin (Tanaka et al., 1998). The amount of kinesin present on mitochondria isolated from control and from TNF-treated cells does not differ, suggesting that the clustering of mitochondria at the minus ends of microtubules does not occur because kinesin has dissociated from these organelles. However, the ability of mitochondria to move on microtubules in vitro in the presence of cytosol is greatly reduced when the cytosol is obtained from TNF-treated cells. Moreover, the light chains of kinesin become hyperphosphorylated when incubated in cytosol from TNF-treated cells versus untreated cells. A MAP kinase is likely involved in the signaling cascade from TNF to kinesin phosphorylation, as an inhibitor of this kinase prevents mitochondrial perinuclear clustering and kinesin light chain phosphorylation in TNF-treated cells (De Vos et al., 2000). These studies suggest that kinesin activity can be inhibited when its light chains become phosphorylated. Similar inhibition of activity is observed when Costal2 becomes phosphorylated. Hedgehog (Hh) is a signaling molecule required for normal Drosophila development. Signal transduction in the Hedgehog pathway is mediated via a protein complex that consists of Costal2 (Cos2), a member of the kinesin superfamily, Fused (Fu), a serine/threonine protein kinase, and Cubitis interruptus (Ci), a transcription factor. In the absence of Hh signaling, this complex binds to microtubules. In the presence of Hh signaling, this complex exhibits weak binding to microtubules, and both Fu and Cos2 are hyperphosphorylated (Robbins et al., 1997). Fu phosphorylates Cos2 at serines residues 572 and 931 in vitro, and these phosphorylation events appear to mimic those induced by Hh signaling in vivo (Nybakken et al., 2002). Thus, phosphorylation of Cos2 at sites distant to the motor/microtubule binding domain can affect the interaction of this motor with microtubules. 17.2.2.3

417

Inhibition of microtubule sliding

Fu2 CaMKII

Ser592/Ser931 Ser 1650

Intermediate chain (IC138) Kinesin heavy chain Kinesin heavy chain Kinesin light chain Kinesin light chain Tail domain Tail domain Stalk domain/ Tail domain Tail domain

Axonemal dynein (Chlamydomas)

Kinesin (chick fibroblasts/rat pancreas)

Kinesin (crayfish)

Kinesin (rat brain)

Kinesin (cultured mammalian cells)

Eg5 (Xenopus, human)

CENP-E (human)

Cos2 (Drosophila)

Myosin V (Xenopus)

Thr 937 (Xenopus) Thr 927 (human)

Cdc2

Dissociation from melanosomes

Decreased microtubule binding

Decreased ATP-insensitive microtubule binding

Karcher et al. (2001)

Robbins et al. (1997), Nybakken et al. (2002)

Liao et al. (1994)

Sawin and Mitchison (1995), Blangy et al. (1995, 1997) Enhanced centrosome association, enhanced dynactin binding

Cdc2

Morfini et al. (2002)

Okada et al. (1995)

Lee and Hollenbeck (1995), Marlowe et al. (1998)

Habermacher and Sale (1997)

Kumar et al. (2000)

Vaughan et al. (2001)

De Vos et al. (2000)

Decreased membrane binding

Decreased vesicle binding

Enhanced membrane binding

Dissociation from dynactin

Dell et al. (2000), Addinall et al. (2001)

Reference

MAP kinase mitochondrial perinuclear clustering

GSK3

PKA

Casein kinase I

Reduced ATPase activity

Light chains?

Cytoplasmic dynein (N. crassa)

Ser 84

Intermediate chain

Cytoplasmic dynein (rat liver)

Dissociation from membranes

Cdc2

Ser 197 OR Cdc2 site in aa 379-405

Light intermediate chain

Cytoplasmic dynein (Xenopus)

Effect

Kinase

Site

Domain or subunit phosphorylated

Motor (source)

Table 17.1. Effect of phosphorylation on motor behavior. Summary of the changes that occur in motor behavior upon phosphorylation. Those motors targeted for phosphorylation and where the change in phosphorylation has been correlated with a change in motor localization or activity, are included. No entry indicates that the information is currently lacking.

418



The light chains of kinesin bind Ca2/calmodulin, and this binding reduces kinesin’s ATPase activity, but phosphorylation of kinesin light chains by PKA appears to protect kinesin activity from this inhibitory effect (Matthies et al., 1993), suggesting that kinases modulate kinesin activity in conjunction with other regulatory molecules. In plants, a member of the kinesin superfamily, kinesin calmodulin binding protein (KCBP), contains a calmodulin binding site within its heavy chain, and binding of Ca2/calmoldulin to this site prevents KCBP from binding to microtubules (Kao et al., 2000, Narasimhulu and Reddy, 1998). While it is not known if KCBP is phosphorylated and if such phosphorylation might overcome the inhibitory effects of calmodulin, this kinesin binds directly to a protein kinase (Day et al., 2000). Table 17.1 summarizes the known effects of phosphorylation on motors. Included in the table are those cases in which phosphorylation of a motor subunit has been correlated with a change in its attachment to cargo or the spindle or with a change in its activity.

17.3

The Role of G Proteins in Regulating Molecular Motors

Studies examining the effects of GTPgS, a non-hydrolyzable GTP analog, on vesicle movements provided the first suggestions that small G proteins might exert regulatory control on motors (Bloom et al., 1993). Small GTP binding proteins bind and hydrolyze GTP and thereby exist in two forms, a GTP-bound form and a GDPbound form. Consequently, G proteins can function as switches, activating a downstream molecule or process when the G protein is in its GTP-bound state and inactivating that molecule or process when it is in its GDP-bound state. Small G proteins comprise a superfamily of five related subfamilies: Ras, Rho, Rab, Sar1/Arf and Ran (reviewed in Takai et al., 2001). A growing body of evidence suggests that members of the Rab and Ran subfamilies affect molecular motors, either by recruiting motors to organelles or by altering motor activity. In a number of cases, G proteins, including members of the Rab, Sar1/Arf and Ras subfamilies, have been found to bind directly to microtubule motors, but a function for most of these interactions has not yet been elucidated. In addition, the Rho family has been shown to regulate the actin and microtubule cytoskeletons (reviewed in Ridley, 2001, Wittmann and Waterman-Storer, 2001). 17.3.1

G Proteins Mediate Motor Cargo Interactions Rab27a recruits myosin Va to melanosomes Recent studies suggest that the Rab family of G proteins may control organelle distribution by regulating motor attachment to that organelle. G proteins may facilitate motor organelle binding when the G protein is in its GTP-bound state and may induce motor release from the organelle when the G protein is in its GDP17.3.1.1

419

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17.3 The Role of G Proteins in Regulating Molecular Motors

bound state. At present, the strongest evidence in support of such a mechanism of regulation is provided by studies on myosin Va binding to melanosomes. Myosin Va is present on melanosomes (pigment granules) in mammalian melanocytes (Wu et al., 1997). Normally, melanosomes disperse along microtubules and accumulate in the dendritic extensions of the melanocytes where they are retained at the cell periphery for their subsequent transfer to skin and hair follicle keratinocytes. This retention does not occur in dilute mutants in mouse (Provance et al., 1996, Wei et al., 1997). Instead, the melanosomes are transported back to the cell center along microtubules where they accumulate perinuclearly (Wu et al., 1998). The dilute mutation and some forms of Griscelli disease in humans, both of which are characterized by a lack of pigment in the skin and hair, are caused by a defect in myosin Va (Mercer et al., 1991, Pastural et al., 1997). Myosin Va normally interacts with actin filaments at the cell cortex and allows the pigment granules to be transferred from the microtubule array onto the actin meshwork (Wu et al., 1998). In the absence of myosin Va, the melanosomes cannot be captured by actin and the granules are returned along microtubules to the cell center. Defects in mice harboring the ashen mutation are indistinguishable from those harboring the dilute mutation (cited from Wu et al., 2001) and, interestingly, the ashen mutation and some forms of Griscelli’s syndrome are caused by defects in a member of the Rab family, Rab27a (Menashe et al., 2000, Wilson et al., 2000). Like myosin Va, Rab 27a localizes to melanosomes (Bahadoran et al., 2001, Hume et al., 2001, Wu et al., 2001), and is essential for myosin Va binding to melanosomes (Hume et al., 2001). Rab27a must be in its GTP-bound state in order for myosin Va to bind to melanosomes. Introduction of DNA encoding wild-type or a GTP-bound mutant of Rab27a into melanocytes can rescue the ashen phenotype whereas introduction of DNA encoding a GDP-bound form of Rab27a cannot. Moreover, the latter DNA induces an ashen-like phenotype in wild-type melanocytes (Wu et al., 2001). Similarly, over-expression of a dominant-negative form of Rab27a, in which the Rab27a is effectively retained in its GDP-bound form, results in a redistribution of pigment granules to the cell center, mimicking the ashen and dilute phenotypes (Hume et al., 2001). Together, these studies suggest that Rab27a may control myosin Va attachment to pigmented organelles as a function of its state of GTP hydrolysis. Rab27a with bound GTP would be predicted to bind myosin Va to the organelle. Hydrolysis of GTP to GDP by Rab 27a should result in the dissociation of myosin Va from the organelle. Myosin Va and Rab27a co-immunoprecipitate (Hume et al., 2001), but their interaction is indirect and is mediated by a third protein, melanophilin (Wu et al., 2002), the product of the leaden mutation in mouse (Matesic et al., 2001). Leaden mice have a color coat defect indistinguishable from the coat defects of ashen and dilute mice (Wu et al., 2002). Melanophilin binds to both Rab27a and to myosin Va and appears to do so via different domains; the N terminus of melanophilin binds to the GTP-bound form, but not the GDP-bound form, of Rab27a, while the C terminus of melanophilin binds to the tail domain of the melanocyte– but not the brain–isoform of myosin V (Wu et al., 2002).


These various studies suggest a model in which myosin Va is recruited to melanosomes and thereby affect their transport on actin filaments in a process that depends on the GTP-bound state of Rab27 (Fig. 17.1). Myosin Va and melanophilin appear to be present on melanosomes regardless of their location in the cell (Wu et al., 2002), so it seems unlikely that Rab27a cycles into its GTP-bound state, and thereby binds melanophilin and myosin Va, only when the melanosomes reach the cell periphery where capture by actin filaments is proposed to occur. An alternative hypothesis, coupling regulation of myosin Va binding to these organelles to the transport process, is that hydrolysis, either stochastic or enhanced by a GAP, might be required to dissociate myosin Va from the organelles, thereby releasing the organelles from the actin filament meshwork at the cell cortex in anticipation of their intercellular transfer to keratinocytes.

G proteins may recruit microtubule motors to organelles Microtubule motors, like myosin Va, may also be recruited to specific sites within the cell, either to organelles or to the spindle, by G proteins, although the data in support of such a model are currently much less definitive than those discussed above for myosin Va melanosome binding. Various members of the Rab family are present on specific organelles. One such member, Rab7, is present on late endosomes and lysosomes (Bucci et al., 2000, Chavrier et al., 1990). The GTP-bound, but not GDP-bound, form of Rab7 interacts with a protein named RILP i. e. Rab-interacting lysosomal protein (Cantalupo et al., 2001). Over-expression of RILP causes lysosomes to cluster at the MTOC, the minus ends of microtubules. Immunofluorescent analysis of the lysosomes reveals that dynein and dynactin, but not kinesin, are recruited to these organelles when RILP is over-expressed, and membrane preparations prepared from these cells are significantly enriched in the p50 dynamitin subunit of dynactin relative to the amount on membranes prepared from control cells (Jordens et al., 2001). At present, there is no information concerning how Rab7 and RILP recruit dynein and dynactin to lysosomes, as the motor complex apparently does not bind directly to either RILP or Rab7 (Jordens et al., 2001). Moreover, it has not yet been demonstrated that Rab7 and RILP are required for dynein dynactin binding to lysosomes in normal cells. If Rab7 is required for dynein to bind to lysosomes, it will be of great interest to determine whether motor association with these organelles can be controlled by the GTP-bound status of Rab7. The G protein Ran alters motor activity in egg extracts in a GTP-dependent fashion and may do so by recruiting Eg5 to asters, although the results of these studies are also compatible with a model in which Ran GTP activates Eg5. Short microtubules are translocated along astral microtubules formed in Xenopus egg extracts. In the absence of Ran GTP, the short microtubules are transported primarily towards the minus ends of the astral microtubules, with only 20 % of the microtubules transported to the plus ends of microtubules. In the presence of Ran GTP, the percentage of microtubules transported to the plus ends of microtubules increases two-fold. Inhibition of Eg5 results in a loss of these plus-end movements 17.3.1.2

421

422

17.3 The Role of G Proteins in Regulating Molecular Motors

(Wilde et al., 2001). It is not clear from these studies whether more Eg5 becomes active and can thereby participate in microtubule transport, or if more Eg5 is recruited to the asters by the GTP-bound form of Ran. Nonetheless, because Eg5 is essential for spindle formation (Sawin et al., 1992), Ran GTP may be required to regulate Eg5 function in organizing the spindle microtubules. 17.3.2

G Proteins may Activate Motors

G proteins may affect motors only by recruiting them to specific locations where they are needed to induce movements. However, at least one study suggests that dynein activity rather than its localization may be affected by its interaction with a G protein. In unstimulated adipocytes, the insulin-sensitive glucose receptors GLUT4 are localized to the Golgi and early endosomes. Upon insulin stimulation, these receptors are delivered via vesicles to the cell surface. Upon insulin withdrawal, the receptors are again internalized. Antibodies to either Rab5 or to dynein stimulate transport of GLUT4 to the plasma membrane in the absence of insulin and inhibit their normal removal from the plasma membrane upon insulin withdrawal. Dynein co-immunoprecipitates with Rab5 regardless of its GTP- versus GDP-bound state, so it is unlikely that Rab5 recruits dynein to GLUT4 -membranes as a function of the GTP hydrolysis state of Rab5. Insulin appears to lock Rab5 into its GDP-bound state, and an inhibitor of phosphatidylinositol-3-OH-kinase (PI3 Kinase) overcomes this inhibitory effect. Insulin also inhibits dynein’s microtubule binding activity, and the same PI3 kinase inhibitor restores dynein’s ability to bind microtubules. Because dynein and Rab5 bind to each other, because inhibitory antibodies against each deliver GLUT4 to the plasma membrane and block GLUT4 re-internalization, because insulin blocks Rab5 from binding GTP and also blocks dynein activity, and because the same kinase inhibitor restores Rab5 GTP binding and dynein activity, dynein activity may be coupled to the state of Rab5 activation (Huang et al., 2001). 17.3.3

Motors may Bind Directly to G Proteins but the Function of these Interactions Remains Unclear

There are a number of examples in which a motor binds directly to a G protein, but no known function has been elucidated for these interactions. The light intermediate chain of cytoplasmic dynein was identified as a binding partner for Rab4 in a yeast two-hybrid screen. This interaction requires that Rab4 be in its GTP-bound state; interaction with the dynein subunit occurs only when either wild-type Rab4 or a Rab4 mutant which lacks GTPase activity (and is thereby ‘locked’ into its GTP-bound state) are utilized in the screen. A Rab4 mutant which cannot bind GTP and a second Rab4 mutant with a defect in its effector binding domain, do not interact with the dynein subunit. Dynein and Rab4 partially co-localize to a population of vesicles that reside in the perinuclear region of the cell (Bielli et al.,


2001). The direct interaction between dynein and active Rab4 and their co-localization in vivo suggest that Rab4 may recruit dynein to those organelles to which Rab4 binds, but studies demonstrating that Rab4 is necessary for dynein localization and/or that it affects dynein motor activity on these organelles are lacking. Similar studies using the Rab6 gene in a yeast two-hybrid screen detected a kinesin family member, Rabkinesin-6, as a Rab6-interacting partner. Rabkinesin-6 interacts with the wild-type and GTP-locked mutant of Rab6, but not with GDPlocked Rab6 or with Rab6 possessing a mutation in its effector domain. In addition, Rab6 and Rabkinesin-6 co-immunoprecipitate, and Rabkinesin-6 localizes to Golgi membranes, the site of Rab6 localization and function (Echard et al., 1998). More recent findings reveal that Rabkinesin-6 accumulates during mitosis and suggest that this motor is required for cleavage furrow formation and cytokinesis (Hill et al., 2000), but no additional studies have been undertaken to elucidate the importance of the relationship between this kinesin and Rab6. Another member of the kinesin family, MKLP1, interacts with the GTP-bound form of Arf3. This interaction has been mapped to the tail domain of the MKLP1 (Boman et al., 1999). The significance of the interaction of this kinesinlike protein with a G protein is not known, but MKLP1, like Rabkinesin-6, has been implicated in cleavage furrow formation (Adams et al., 1998). Finally, one additional kinesin family member, KIF9, was identified in a yeast two-hybrid screen for interaction partners of Gem. Gem is a member of the ras subfamily, and over-expression of Gem causes cells to form long, dendritic extensions. Unlike most of the studies discussed above, KIF9 interaction with GEM does not require that Gem be in its GTP-bound state (Piddini et al., 2001). 17.3.4

A Light Chain of Dynein may be Involved in Regulating G Protein GTPase Activity

Recent analysis of relatedness among bacterial, archaea and eukaryotic gene families has raised the interesting specter that the dynein light chains belonging to the Roadblock/LC7 family may be related to an ancient family of proteins, called the MglB family, which may function to regulate the GTPase activity of small G proteins (Koonin and Aravind, 2000). In Drosophila, mutations in the Roadblock gene lead to defects in axonal transport as well as in mitosis (Bowman et al., 1999). It is possible that the Roadblock/LC7 family, like the MglB family, is a regulator of nucleoside triphosphatatase activity and mediates its effect by regulating the ATPase activity of the heavy chain of dynein. Another possibility, though, is that the Roadblock/LC7 family functions to regulate the GTPase activity of G proteins that interact with and regulate dynein.

423

424

17.4 Other Mechanisms of Regulation

17.4

Other Mechanisms of Regulation

In addition to motor regulation by phosphorylation and motor regulation by interaction with G proteins, there are additional mechanisms that can regulate motors. 17.4.1

Kinesin Folding

The activity of kinesin may be regulated by conformational changes that it undergoes upon binding to cargo. Soluble kinesin has low microtubule-stimulated ATPase activity relative to a fragment of kinesin that lacks its C-terminus (Kuznetsov et al., 1989). Kinesin can undergo a change in conformation from a compact to an extended form; the compact form requires the presence of the C terminus which is also required for inhibition of microtubule-stimulated ATPase activity (Stock et al., 1999). Binding to cargo may de-repress kinesin, as full-length kinesin bound to artificial cargo exhibits high microtubule-stimulated ATPase activity relative to soluble motor. Removal of the tail domain results in activation of kinesin without the need for cargo binding, and re-addition of the tail domain peptide results in renewed inhibition (Coy et al., 1999). Removal of hinge 2 from kinesin, a domain which allows kinesin to fold and thereby the head and tail to interact, results in a kinesin molecule with motile and ATPase properties similar to those of kinesin lacking its carboxy tail (Friedman and Vale, 1999). These studies suggest that, in a folded conformation, the tail domain interacts with the motor domain to repress its activity. Binding of kinesin to cargo may unfold the molecule so that repression is alleviated. In vivo studies in Neurospora crassa suggest that regulation mediated by the tail domain of kinesin may be essential for regulated organelle transport. Point mutations in a conserved domain, RIAKPLR, near the C terminus of kinesin affect neither folding of the molecule nor cargo binding. However, this mutant tail domain is unable to repress the ATPase activity of the kinesin motor domain as does the wild-type tail and molecules carrying these mutations accumulate abnormally at the hyphal tips (Seiler et al., 2000). This domain of kinesin has been shown to prevent ADP release from the motor domain (Hackney and Stock, 2000). Together, these studies suggest that a domain within the C-terminus of kinesin specifically interacts with the motor domain of kinesin in folded molecules to repress kinesin activity. Such a mechanism would help prevent non-productive interactions of kinesin, unattached to cargo, with microtubules. Upon kinesin binding to cargo, kinesin would unfold and repression would be alleviated so that kinesin can move that cargo on microtubules. In the absence of repression, as exhibited by the mutants in N. crassa, kinesin that was not attached to cargo could nonetheless move along microtubules and would thereby accumulate abnormally at the plus ends of microtubules, potentially negatively affecting cell growth (Seiler et al., 2000).


17.4.2

Lis 1 Interaction with Cytoplasmic Dynein

Mutations in the LIS1 gene cause lissencephaly in humans, in which neurons fail to migrate properly in the developing brain (Vallee et al., 2001; see Chapter 20 by Vallee and Tai). That Lis1 might interact with dynein was first recognized in studies of nuclear migration in Aspergillus nidulans. A mutation in the NudF gene results in a defect in transport of nuclei into the fungal hyphae, a phenotype that resembles defects in the NudA gene, which encodes the heavy chain of dynein. The NudF protein is 42 % identical to the mammalian Lis1 protein, and mutations in the Nud A gene can suppress mutations in the NudF gene, suggesting that dynein heavy chain and Lis1 interact and function in the same pathway (Willins et al., 1997, Xiang et al., 1994, 1995). Moreover, NudF and NudA co-localize at the plus ends of microtubules (Han et al., 2001). Similarly, mutations in the Lis1 gene in Drosophila result in numerous defects, including axonal transport, cell division, and abnormal nuclear positioning in oocytes and in photoreceptor cells (Lei and Warrior, 2000, Liu et al., 1999, 2000, Swan et al., 1999). Dynein normally accumulates at the posterior pole of the oocyte, but does not do so in Lis1 mutants, suggesting that Lis1 may be required for normal dynein localization or activity (Lei and Warrior, 2000, Swan et al., 1999). Mammalian cytoplasmic dynein and Lis1 protein physically interact, as assayed by co-immunoprecipitation studies (Faulkner et al., 2000, Smith et al., 2000), and enhanced Lis1 expression in Cos-7 cells or in embryonic fibroblasts cells results in a more compact Golgi complex at the minus ends of microtubules, suggesting that Lis1 activates dynein/dynactin function (Smith et al., 2000). Over-expression of Lis1 can be detrimental and disrupt normal mitotic progression (Faulkner et al., 2000). Both Lis1 and dynein co-localize at the kinetochores and at the cell cortex (Faulkner et al, 2000). NudF and Lis1 bind directly to the heavy chain of dynein and, moreover, bind to its first P-loop, the site of its motor activity (Hoffmann et al., 2001, Tai et al., 2002). These studies suggest that Lis1 may affect dynein activity by interacting directly with its motor domain. At present, it is the only known protein that can bind directly to the motor domain within the dynein heavy chain. 17.4.3

Motor Protein Regulation during the Cell Cycle

Phosphorylation events, tied to the cell cycle, may regulate the behavior of molecular motors. In addition to regulation by phosphorylation, the synthesis and degradation of motors, coupled to the cell cycle, may be a mechanism used to regulate motors. The kinesin-related proteins, CENP-E, HINKEL, Cin8p, and chromokinesin, appear to cycle with the cell cycle (Brown et al., 1994, Funabiki and Murray, 2000, Hildebrant and Hoyt, 2001, Strompen et al., 2002). Cin8p, like Eg5 in vertebrates, performs an essential function in spindle assembly. The levels of Cin8p decrease at the end of mitosis, and this loss is the result of its destruction via APCinduced ubiquitination. Over-expression of a non-degradable form of Cin8p is

425

426

17.4 Other Mechanisms of Regulation

lethal to cells, whereas expression of normal amounts of the non-degradable form of the protein causes spindle defects (Hildebrant and Hoyt, 2001), suggesting that loss of this kinesin is an essential regulatory event. Chromokinesin is a kinesin-related protein that binds to chromatin and generates the polar ejection forces that push the chromosome arms towards the equator (Levesque and Compton, 2001, Wang and Adler, 1995). Normally, chromokinesin, like Cin8p, is degraded by a ubiquitin pathway at anaphase, but, if additional chromokinesin is introduced into cells at anaphase, chromosome movement towards the poles is inhibited (Funabiki and Murray, 2000). These studies suggest that degradation of motors may be an essential mechanism to ensure that a motor cannot interfere either with microtubules or with its cargo when the motor is no longer needed. 17.4.4

Motor Complexes and Coordination

Motor molecules might interact with each other, either cooperatively or antagonistically, and these interactions might regulate motor activity. Lipid droplets in Drosophila embryos exhibit bi-directional transport, and, in wild-type cells, the transport in one direction appears to be unimpeded by the motor that drives transport in the opposite direction, suggesting that the motors are regulated so that they do not antagonize each other (Gross et al., 2002a). Impedance between the motors occurs in klar mutants, and it is possible that the klar protein participates in the coordination of motor activity (Welte et al., 1998). In yeast, secretion defects caused by a mutation in the myo2 gene, which encodes a myosin V, can be suppressed by Smy1p, a kinesin-related protein (Lillie and Brown 1992). These two proteins bind directly to each other in a yeast two-hybrid screen and co-localize at the bud tip and at the mother- daughter neck. The suppression by Smy1 does not require microtubules, and Smy1p localization depends on Myo2p, whereas Myo2p localization does not depend on Smy1p (Beningo et al., 2000, Lillie and Brown, 1994, 1998). These findings suggest that suppression does not occur when cargo, normally transported by Myo2p on actin filaments, is instead transported on microtubules by Smy1p. Instead, suppression may occur because Smy1p activates Myo2p by binding directly to it (Beningo et al., 2000). Myosin V also binds directly to the heavy chain of conventional kinesin (Huang et al., 1999b), and it is possible that these motor interactions are utilized to coordinate motors. In Xenopus melanophores, myosin V, kinesin II and dynein are present on the pigment granules (Rogers et al., 1997, Rogers and Gelfand, 1998) and both myosin V and kinesin II are needed to induce pigment dispersion (Rogers and Gelfand, 1998, Tuma et al., 1998). Myosin V may contribute to kinesin II-driven movements by restricting the ability of dynein to drive pigment granule transport back towards the cell center during pigment dispersion but not during pigment aggregation (Gross et al., 2002b). Thus, regulation between dynein-driven and kinesin-driven transport may be mediated by myosin V interference affecting only one of the two microtubule motors.


17.5

Summary

The mechanisms that cells utilize to regulate their molecular motors are varied, and a single motor may be regulated by more than one mechanism, the effect of which would allow the cell to control that motor under different physiological circumstances. The sites on motors or interacting proteins that mediate regulation are now being elucidated, and it is likely that different sites on a single motor can differentially regulate the motor. Phosphorylation is a predominant mechanism regulating motors, but G proteins may also exert significant influence on motor localization or behavior. In addition, intramolecular or intermolecular interactions of motor proteins may influence their behavior. Motors serve numerous essential roles for cells, and motor regulation is a fundamental component of their proper functioning. Continued progress in this field will be rewarding and exciting.

Acknowledgements

The author acknowledges support from the National Science Foundation and the University of California Academic Senate.

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17 Regulation of Molecular Motors neracts with the novel kinesin-like protein KIF9. EMBO J. 20: 4076 4087. Provance, D. W. J., W. Wei, V. Ipe, and J. A. Mercer. 1996. Cultured melanocytes from dilute mutant mice exhibit dendritic morphology and altered melanosome distribution. Proc. Natl. Acad. Sci. USA 93: 14554 14558. Ridley, A. J. 2001. Rho proteins: Linking signaling with membrane trafficking. Traffic 2: 303 310. Robbins D. J., K. E. Nybakken, R. Kobayashi, J. C. Sisson, J. M. Bishop, and P. P. Therond.1997. Hedgehog elicits signal transduction by means of a large complex containing the kinesin-related protein costal2. Cell 90: 225 234. Rogers, S. L. and V. I. Gelfand. 1998. Myosin cooperates with microtubule motors during organelle transport in melanophores. Curr. Biol. 8: 161 164. Rogers, S. L., I. S. Tint, P. Fanapour, and V. I. Gelfand. 1997. Regulated bi-directional motility of melanophore pigment granules along microtubules in vitro. Proc. Natl. Acad. Sci. USA 94: 3720 3725. Rogers, S. L., R. L. Karcher, J. T. Roland, A. A. Minin, W. Steffen, and V. I. Gelfand. 1999. Regulation of melanosome movement in the cell cycle by reversible association with myosin V. J. Cell Biol. 146: 1265 1275. Sawin, K. E. and T. J. Mitchison. 1995. Mutations in the kinesin-like protein Eg5 disrupting localization to the mitotic spindle. Proc. Natl. Acad. Sci. USA 92: 4289 4293. Sawin, K. E., K. Leguellec, M. Philippe, and T. J. Mitchison. 1992. Mitotic spindle organization by a plus-end directed microtubule motor. Nature 39: 540 543. Schaar, B. T., G. K. T. Chan, P. Maddox, E. D. Salmon, and T. J. Yen. 1997. CENP-E function at kinetochores is essential for chromosome alignment. J. Cell Biol. 139: 1373 1382. Seiler, S., J. Kirchner, C. Horn, A. Kallipolitou, G. Woehlke, and M. Schliwa. 2000. Cargo binding and regulatory sites in the tail of fungal conventional kinesin. Nature Cell Biol. 2: 333 337. Smith, D. S., M. Niethammer, R. Ayala, Y. Zhou, M. J. Gambello, A. Wynshaw-Boris, and L.-H. Tsai. 2000. Regulation of cytoplasmic dynein behaviour and microtubule organization by mammalian Lis1. Nature Cell Biol. 2: 767 775.

Stock, M. F., J. Guerrero, B. Cobb, C. T. Eggers, T.-G. Huang, X. Li, and D. D. Hackney. 1999. Formation of the compact confomer of kinesin requires a COOH-terminal heavy chain domain and inhibits microtubule-stimulated ATPase activity. J. Biol. Chem. 274: 14617 23. Strompen, G., F. El Kasmi, S. Richter, W. Lukowitz, F. F. Assaad, G. Jurgens, and U. Mayer. 2002. The Arabidopsis HINKEL gene encodes a kinesin-related protein involved in cytokinesis and is express in a cell cycle-dependent manner. Curr. Biol. 12: 153 158. Swan, A., T. Nguyen, and B. Suter. 1999. Drosophila Lissencephaly-1 functions with BicD and dynein in oocyte determination and nuclear positioning. Nature Cell Biol. 1: 444 449. Tai, C. Y., D. L. Dujardin, N. E. Faulkner, and R. B. Vallee. 2002. Role of dynein, dynactin, and CLIP-170 interactions in LIS1 kinetochore function. J. Cell Biol. 156: 959 968. Tanaka, Y., Y. Kanai, Y. Okada, S. Nonaka, S. Takeda, A. Harada, and N. Hirokawa. 1998. Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal perinuclear clustering of mitochondria. Cell 93: 1147 1158. Takai, Y., T. Sasaki, and T. Matozaki. 2001. Small GTP-binding proteins. Physiol. Rev. 81: 153 208. Tuma, M., A. Zill, N. Le Bot, I. Vernos, and V. Gelfand. 1998. Heterotrimeric kinesin II is the microtubule motor protein responsible for pigment dispersion in Xenopus melanophores. J. Cell Biol. 143: 1547 1558. Vallee, R. B., C. Tai, and N. E. Faulkner. 2001. LIS1: cellular function of a disease-causing gene. Trends Cell Biol. 11: 155 160. Vaughan, P. S., J. D. Leszyk, and K. T. Vaughan. 2001. Cytoplasmic dynein intermediate chain phosphorylation regulates binding to dynactin. J. Biol. Chem. 276: 26171 26179. Wang, S. Z. and R. Adler. 1995. Chromokinesin: a DNA-binding, kinesin-like nuclear protein. J. Cell Biol. 128: 761 768. Wei, Q., X. Wu, and J. A Hammer III. 1997. The predominant defect in dilute melanocytes is in melanosome distribution and not cell shape, supporting a role for myosin V in melanosome transport. J. Muscle Res. Cell Motil. 18: 517 527. Welte, M. A., S. P. Gross, M. Postner, S. M. Block, and E. F. Wieschaus. 1998. Developmental regulation of vesicle transport in

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18 Molecular Motors in Plant Cells A. S. N. Reddy

18.1

Introduction

Molecular motors regulate diverse cellular functions including the organization and dynamics of microtubule (MT) and actin cytoskeleton, cytoplasmic streaming, cell polarity, cell growth, morphogenesis, chromosome segregation and transport of vesicles, organelles and macromolecular complexes. In eukaryotes, there are three broad families of molecular motors: the kinesins, the dyneins and the myosins. All three types of motors utilize energy from the hydrolysis of ATP to move along filamentous structures. Kinesins and dyneins move on MTs whereas the myosins translocate on actin filaments. Using biochemical, cell biological, molecular, and genetic approaches several molecular motors have been identified in plants and functions of some of these are beginning to be understood. Recent completion of genome sequence of several eukaryotes ranging from yeast, a simple unicellular eukaryote, to highly evolved multicellular organisms including humans and flowering plants has permitted comparative analysis of three families of motors in various species. Such analysis across phylogenetically divergent species has yielded some interesting insights into evolutionary and functional relationships among motor proteins from various organisms. Systematic analyses of the recently completed Arabidopsis genome sequence with the conserved motor domain of kinesins, myosins and dyneins revealed the presence of 78 molecular motors (61 kinesin-like proteins and17 myosins) in this organism. Of the two families of MT-based motors, dyneins are absent in flowering plants. Surprisingly, Arabidopsis has the largest number of kinesin-like proteins (KLPs) as compared to other multicellular organisms including humans, suggesting that the kinesin superfamily is expanded considerably in plants. Also, all plant myosins belong to two novel classes. Although the identification of molecular motors in plants is progressing rapidly due to genome and EST sequencing projects, only a few plant motors have been characterized in any detail and the functions of many these motors are not known. Nevertheless, it is becoming obvious that plants contain novel families of KLPs and myosins with novel functions and regulatory me-

434

18.2 Microtubule-based Motors

chanisms. In this article I focused primarily on plant molecular motors, their function and regulation with emphasis on the model plant, Arabidopsis. For other aspects of plant motors that are not covered here the reader is directed to other reviews (Asada and Collings, 1997, Liu and Lee, 2001, Reddy, 2001b, Yamamoto et al., 1999).

18.2

Microtubule-based Motors

The motors that move on MTs belong to two different families: kinesins and KLPs that move either toward the plus-end or minus-end of the MTs, and the dyneins, which move toward the minus-end of MTs. 18.2.1

Kinesin-like Proteins

Members of the kinesin superfamily, which consists of conventional kinesin and kinesin-like proteins (KLPs), are known or implicated to play important roles in many fundamental cellular and developmental processes including intracellular transport of vesicles and organelles, mitotic and meiotic spindle formation and elongation, chromosome segregation, germplasm aggregation, MT organization and dynamics and intraflagellar transport (Endow, 1999, Goldstein and Philip, 199, Hirokawa, 1998, Reddy, 2001b). A large number of KLPs have been identified in plants and animals (Berg et al., 2001, Lawrence et al., 2002, Reddy and Day, 2001a, 2001b, Siddiqui, 2002, Yamashita et al., 2000; Tab. 18.1). All members of the kinesin superfamily have a highly conserved catalytic region of about 350 amino acid residues, known as the motor domain, with ATP- and MT-binding sites. The motor domain in KLPs is located either in the N terminus, C terminus or in the middle of the protein. In addition to the motor domain, most KLPs have a stalk region that forms an alpha-helical coiled-coil region which aids in dimerization, and a highly variable tail, which is thought to interact with a specific cargo. Outside the motor domain KLPs share no sequence similarity between the members of kinesin superfamily. Although molecular motors have been implicated in a variety of cellular processes in plants including spindle function, cytokinesis, cell polarity, morphogenTable 18.1

The total number of KLPs and myosins in the completely sequenced organisms. S. cerevisiase

S. pombe

C. elegans

D. melanogater

H. sapiens A. thalaiana

KLPs

6a

9a

21b

24a

45c

61a

Myosins

5d

5d

17d

13d

40d

17d,e

Total

11

36

37

a

14 b

c

85 d

78 e

Day and Reddy, 2001b; Siddiqui, 2002; Miki et al., 2001; Berg et al., 2001; Day and Reddy, 2001a

18 Molecular Motors in Plant Cells

esis, organelle and vesicle transport, and cytoplasmic streaming (Asada and Collings, 1997, Lloyd, 1991, Reddy, 2001b, Williamson, 1993), until recently very little was known about their molecular identity. Using antibodies to animal kinesins, KLPs have been identified in pollen tubes of tobacco and other plants (Cai et al., 1993,; Liu et al., 1994, Tiezzi et al., 1992). Recently, Cai et al. isolated an MTbased motor (90 kD) from tobacco pollen tubes which induced MT gliding in motility assays (Cai et al., 2000). Immunolocalization studies have indicated that this motor binds to organelles associated with MTs in the cortical regions of the pollen tube, suggesting its involvement in organelle transport. Two KLPs (125 and 120 kDa) that showed plus-end motor activity in in vitro motility assays and MT-dependent ATPase and GTPase activity, were isolated from tobacco phragmoplasts (Asada and Shibaoka, 1994, Asada et al., 1991). Using various approaches cDNAs for several KLPs have been characterized from plants. Ten of these are from Arabidopsis (KatA, KatB, KatC, KatD, AtKCBP, AtPAKRP1, AtPAKRP2, AtMKRP1, AtMKRP2, AtNCAK1) (Itoh et al., 2001, Lee and Liu, 2000, Lee et al., 2001, Mitsui et al., 1996, Mitsui et al., 1994, 1993; Nishihama et al., 2002, Reddy et al., 1996b, Tamura et al., 1999), 15 from tobacco (TKRP125, KCBP, NCAK1, NCAK2, and TBK1 to TBK11) (Asada et al., 1997, Matsui et al., 2001, Nishihama et al., 2002, Wang et al., 1996), two from carrot (KRP120-1 and KRP120-2 (Barroso et al., 2000) and one from potato (KCBP). A kat gene family (katA, katB, katC, katD, and katE) encoding KLPs in Arabidopsis thaliana was characterized using primers corresponding to conserved regions of the kinesin motor domain (Mitsui et al., 1993, 1994, 1996, Tamura et al., 1999). Using a similar approach, 12 different KLPs that belong to seven subfamilies have been isolated from tobacco BY-2 cells (Matsui et al., 2001). It is interesting that in tobacco BY-2 cells alone, 15 different KLPs that belong to different subfamilies are expressed (Asada et al., 1997, Bowser and Reddy, 1997, Matsui et al., 2001, Nishihama et al., 2002). Using an indirect assay, KatA (89 kD) protein has been shown to be a minus-end-directed motor and a similar protein has been found in carrot and tobacco (Liu et al., 1996). The central region of KatD shares sequence similarity to the motor domain of kinesin and is followed by about 240 residues at the C-terminus. Phylogenetic analysis with the KatD motor domain indicates that it belongs to the C-terminal family despite the fact that it has a 240-amino acid stretch following the motor domain (Reddy and Day, 2001b, Tamura et al., 1999). The amino-terminal region of KatD showed sequence similarity to calponin homology (CH) domain. KatD is a flower-specific motor protein suggesting that it may function in transport or cytoskeleton organization in pollen. Asada et al. (1997) isolated a cDNA encoding the 125-kDa polypeptide (TKRP125, tobacco kinesin-related polypeptide of 125 kDa). It is an N-terminal KLP with strong similarity to members of the BimC (blocked in mitosis) subfamily. Two other members of BimC class KLPs (DcKRP120-1 and DcKRP120-2) were isolated from carrot suspension cells (Barroso et al., 2000). DcKRP120-1 is a homolog of TKRP125. In Arabidopsis there seems to be at least three TKRP-like proteins. Recently, two phragmoplast-associated kinesin-related proteins (AtPAKRP1 and AtPAKRP2) have been isolated from Arabidopsis and characterized (Lee and Liu, 2000, Lee et al., 2001).

435

436


AtPAKRP1 is an N-terminal KLP and is most closely related to XKlp2, an ungrouped KLP from Xenopus laevis. AtPAKRP2 is also an N-terminal motor with no known homologs in non-plant systems. In phylogenetic analysis, AtPAKRP2 does not group with any of the existing families of KLPs. These motors have distinct functions in cytokinesis (Lee et al., 2001). KCBP (kinesin-like calmodulin-binding protein), a novel KLP with a C-terminal motor domain was isolated from a number of plants as a calmodulin-binding protein (Abdel-Ghany et al., 2000, Abdel-Ghany and Reddy, 2000, Reddy et al., 1996a, 1996b, Wang et al., 1996). Although KCBP, like other kinesin-like proteins, contains three distinct regions (a motor domain, a coiled-coil stalk and a tail), it has two features that make it unique among members of the kinesin superfamily in eukaryotes. These include (i) a calmodulin-binding domain adjacent to the motor domain at the C-terminus, and (ii) a myosin tail homology and talin-like region in the N-terminal tail (Reddy and Reddy, 1999). KCBP binds calmodulin in a calcium-dependent manner at physiological calcium concentrations (Reddy et al., 1996b, 1999). KCBP is a minus-end directed MT motor (Song et al., 1997) and has two MT binding domains, one located at the C terminus and the second one located at the N terminus (Narasimhulu and Reddy, 1998). Unlike the MT binding domain in the C terminus, the N-terminal region of KCBP binds MTs both in the presence and absence of ATP, indicating that the MT binding domain in the N terminus is insensitive to ATP. KCBP is highly conserved in phylogenetically divergent plant species including dicots, monocots, gymnosperms and algae (S. E. Abdel-Ghany and A. S. N. Reddy, unpublished data). Recently, a calmodulin-binding C-terminal kinesin (kinesin C) was cloned from sea urchin (Rogers et al., 1999). The calmodulin-binding domain of kinesin C shared 35 % sequence identity with the calmodulin-binding domain in KCBP. The existence of calmodulin-binding KLPs in both plants and sea urchin suggests that the origin of this group of KLPs pre-dates the divergence of plants and animals from a common ancestor which is believed to have occurred about 1.5 billion years ago (Wang et al., 1999). If this was the case, there must have been insertion of some domains such as MyTH4 and talin-like regions in the tail of KCBP (or deletion of these domains in kinesin C) to acquire functional specialization of these KLPs. Alternatively, calmodulin-binding KLPs may have evolved independently in plant and animals after they diverged from a common ancestor. The amino-terminal tail and stalk regions of KCBPs from different plant systems are highly conserved and contain myosin tail homology (MyTH4) and talin-like regions that are not present in kinesin C. In phylogenetic analysis of KLPs KCBPs and kinesin C are grouped with other known C-terminal KLPs. However, Arabidopsis KCBP together with its orthologs from other plants constitute a distinct group within the C-terminal subfamily of motors. Recently, two Arabidopsis KLPs (AtMKRP1 and AtMKRP2) with mitochondrial targeting signals were found to be targeted to mitochondria, suggesting that these motors might function in this organelle (Itoh et al., 2001). AtMKRP1 and AtMKRP2 belong to ungrouped KLPs (Reddy and Day, 2001b). What functions these MKRPs perform in mitochondria remains to be seen. There are other Arabi-


dopsis KLPs with targeting signals to other organelles such as chloroplasts. It would be interesting to see if any other Arabidopsis KLPs are targeted to organelles. KLPs that are targeted to organelles have not been reported previously in the literature. Mutations in the HINKEL (HIK) gene in Arabidopsis result in defective cytokinesis. These mutants have high frequency of incomplete cell walls and multinucleate cells. The HIK gene encodes a plant-specific N-terminal KLP that plays an important role in cytokinesis (Strompen et al., 2002). In a recent report, Nishihama et al. (2002) have shown that two tobacco KLPs (NACK1 and NACK2) interact with NPK1, a mitogen-activated kinase kinase kinase, and stimulate the activity of the kinase. Orthologs of NACK1 and NACK2 have been identified in Arabidopsis and AtNACK1 was found to be identical to HIK. These KLPs are highly diverged N-terminal motors and do not group with any of the known KLP subfamilies (Reddy and Day, 2001b). The closest non-plant KLP that shares significant similarity with the motor domain of NCAKs is CENP-E. At least eight Arabidopsis KLPs are known to function in some aspect of cell division (see Section 18.4.1 on cell division). Analysis of the recently completed Arabidopsis genome sequence with the conserved motor domain of KLPs has resulted in identification of 61 KLPs in the model plant (Reddy and Day, 2001b). A corresponding cDNA or EST (Expressed Sequence Tag) was found for 37 of these KLPs (Tab. 18.2), suggesting that most KLPs in Arabidopsis are expressed and not likely to be pseudogenes. The genes encoding KLPs are distributed throughout the genome. Chromosome 3 has the highest number (18) of KLP genes. In comparison, S. cerevisiae, S. pombe, C. elegans, Drosophila, and Homo sapiens have 6, 9, 21, 24 and 45 KLPs respectively (Tab. 18.1). Surprisingly, the Arabidopsis genome contains the largest number of KLPs among all eukaryote genomes that have been sequenced. Arabidopsis has the highest percentage (0.24 %) of the total number of genes as compared to S. cerevisiae and S. pombe with 0.1 and 0.17 % respectively, C. elegans with 0.11 % and Drosophila with 0.18 %. Based on the number of KLPs in Arabidopsis, the number of cellular processes that are regulated by KLPs in plants is likely to match or exceed the number of processes controlled by animal KLPs. Only 10 of the 61 Arabidopsis KLPs have been reported in the literature (Lee and Liu, 2000, Lee et al., 2001, Itoh, 2001, Liu et al., 1996, Mitsui et al., 1993, 1994, Nishihama et al., 2002, Reddy et al., 1996b, Tamura et al., 1999) whereas the three AtKRP125 KLPs (AtKRP125a,b, and c; Tab. 18.2) show sequence similarity to a tobacco kinesin (NtKRP125) that was isolated from phragmoplasts of tobacco (Asada et al., 1997). AtKRP125b has 68 % identity with NtKRP125 over the 1000 residues they have in common. The number of known and predicted introns in KLPs range from 3 to 34 (Tab. 18.2). In the Arabidopsis genome, the number of introns ranges from 0 to 77 with an average of about five (Reddy, 2001c). More than 85 % of Arabidopsis genes have 10 or less introns while the Arabidopsis kinesin genes have an average of 16.4 with only 10 genes having 10 or less introns. The number and locations of the introns need to be verified experimentally. It is likely that the predicted sizes of

437

N-terminal N-terminal

C-terminal C-terminal C-terminal

At2g37420 (AtKRP125a)

At2g36200 (AtKRP125b)

At2g28620 (AtKRP125c)

At3g45850

At4g05190

At4g21270 (AtKatA)

At4g27180 (AtKatB)

At5g54670 (AtKatC)

At5g27550

At2g22610

At1g72250

At3g10310

At1g18410

At1g73860

At1g63640

At5g41310

1

2

3

4

5

6

7

8

9

10

11

12

13

14

15

16 Internal

Internal

Internal

Internal

Internal

Internalc

c

c

c

c

c

Internalc

b

N-terminal

C-terminal

N-terminal

N-terminal

Motor location

EST/ cDNA

Kinesin-like proteins in Arabidopsis.

Gene code (published name)

Table 18.2

967

1056

1050

1162

*

897

1195

1068

425

754

*

745

*

793

777

1058

1076

1056

1022

#aa

19

19

17

17

16

17

18

6

15

15

8

17

21

18

22

17

# of introns

a

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

Minus

ND

ND

ND

ND

ND

Motility

CC

CC

CC

CC

CC

Other Domains

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

CC, CH

CC, CH

CC

CC

CC, CH

CC

CC

CC

CC

CC

Mitotic MT arrays CC

ND

ND

ND

ND

ND

Cellular localization

Mitsui et al, 1994.

Mitsui et al, 1994.

Mitsui et al., 1993; Liu et al., 1992.

Asada et al., 1997.

Asada et al., 1997.

Asada et al., 1997; Barroso et al., 2000.

Ref

438


N-terminal N-terminal N-terminal N-terminal

At5g10470

At5g65460

At5g27950

At1g55550

At5g65930 (AtKCBP)

At1g20060

At3g10180

At1g59540

At5g06670

At3g12020

At1g21730 (MKRP1)

At4g39050 (MKRP2)

At2g21380

21

22

23

24

25

26

27

28

29

30

31

32

33

767

At3g44730

861

c

N-terminal

N-terminal

N-terminal

N-terminal

C-terminal

857

1121 (1055)

909* (890)

956

997

823

459

18

22

22

22

22

17

11

21

9

8

923*

887

N-terminal

N-terminal

22

20

640

b

N-terminal

22

14

14

18

17

1259

1264

b

1273

b

N-terminal

b

Internal

Internal

20

At2g47500

c

19

1032*

Internalc

At1g09170

987

Internalc

18

At5g27000 (AtKatD)

17

ND

ND

ND

ND

ND

ND

ND

ND

Minus

ND

ND

ND

ND

ND

ND

ND

ND

ND

Mitochondria

Mitochondria

ND

ND

ND

ND

ND

PPB, spindle, Spindle poles, Phragmoplast

ND

ND

ND

ND

ND

ND

ND

ND

CC

CC

CC

CC

CC

CC

CC

CC, MyTH4 Talin-like, CBD, PEST

CC

CC

CC

CC

CC

CC, CH

CC, CH

CC, CH

Itoh et al., 2001.

Itoh et al., 2001.

Reddy et al., 1996.

Tamura et al., 1999.

18 Molecular Motors in Plant Cells 439

N-terminal

At1g18370 (HINKEL/AtNACK1)

At3g43210 (ATNACK2)

At4g38950

At2g21300

At3g51150

At5g66310

At5g42490

At4g24170

At3g16630

At3g16060

At5g02370

At1g18550

At3g49650

At5g23910

At3g50240

At5g47820

34

35

36

37

38

39

40

41

42

43

44

45

46

47

48

49

N-terminal N-terminal N-terminal N-terminal

N-terminal

1032

1075

665

813

22

21

17

14

3

703

10

*

12

12

14

9

13

13

9

11

13

11

# of introns

664

706

Internal N-terminal

799

Internal

1037

1263

N-terminal

968

N-terminal

N-terminal

581

1087

N-terminal

834

932

1003* (974)

#aa

N-terminal

N-terminal

N-terminal

Motor location

EST/ cDNA

(continued).

Gene code (published name)

Table 18.2

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

Motility

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

Interzonal MTs Phragmoplast

Phragmoplast equator

Cellular localization

CC

CC

HhH1

CC

CC

CC

CC

CC

CC

CC

CC

CC

CC,

CC

Other Domains

Nishihama et al., 2002.

Strompen et al., 2002; Nishihama et al., 2002.

Ref

440


At3g23670

59

60

61

N-terminal

16

30

*

959 (869) 9

24

1292 (1294)

14

I

16

1103* 1268*

20

1229

2158

34

2756

1070

895 20

18

*

885

11

*

24

439

1335*

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

CC,

CC

CC

CC,

CC,

CC, ARM

CC, ARM

CC, ARM,

CC

CC

Punctate staining CC of Phragmoplast

Spindle midzone, CC Phragmoplast

ND

ND

ND

ND

ND

ND

ND

ND

ND

ND

Lee et al., 2001

Lee and Liu, 2000

by indirect assay; balthough the motor is N-terminal, it groups into C-terminal family; calthough the motor is internal, it groups into C-terminal family; iNCBI predicted protein has 1662 amino acids; *slight discrepancy in #aa between AtDB and MIPS predicitions; The number of amino acids in parenthesis are based on the deduced number from cDNA. EST, expressed sequence tag, CC, coiled coil; CH, calponin homology domain; MyTH4, domain present in the tail region of some myosins; Talinlike, talin-like domain found in some myosins and band 4.1 superfamily; CBD, calmodulin binding domain; PEST, motif rich in proline, glutamine, serine and threonine residues; ARM, Armadillo/beta-catenin-like repeats; HhH1, helix-hairpin-helix. (Modified from Reddy and Day, 2001a)

a

At3g20150

58

N-terminal

N-terminal

N-terminal

At3g44050

57

At3g17360

56

N-terminal

At4g14330 (AtPAKRP2)

At3g19050

55

N-terminal

N-terminal

At3g54870

54

N-terminal

At1g12430

53

N-terminal

At4g14150 (AtPAKRP1)

At1g01950

52

N-terminal

N-terminal

At3g63480

51

N-terminal

At5g60930

50

18 Molecular Motors in Plant Cells 441

442


the proteins in Tab. 18.2 and 18.3 may change somewhat as corresponding cDNAs are characterized. In a few cases where the cDNAs were isolated recently there are some differences between the predicted and actual size of the protein due to inaccurate predictions of introns and exons (Itoh et al., 2001, Lee and Liu, 2000, Lee et al., 2001). All Arabidopsis KLPs except three have a coiled-coil region, suggesting that they may function as homo or heterodimers (Fig. 18.1 and Tab. 18.2). Interestingly, some Arabidopsis KLPs have predicted domains that are not present non-plant Table 18.3

Myosins in Arabidopsis.

Name

# of aa

Gene code

EST/ cDNA

Class Domains

Reference

1. At ATM

1166

AT3g19960 (ATM1)a

VIII

MD, CC, IQ

(Knight and Kendric-Jone, 1993)

2. At ATM2

1111 1101b

AT5g54280 ATM2/AtMYOS1a

VIII

MD, CC, IQ

(Kinkema et al., 1994)

3. At VIIIA

1085

AT1g50360

VIII

MD, CC, IQ

4. At VIIIB

1126

AT4g27370

VIII

MD, CC, IQ

5. At MYA1

1520 1599c

AT1g17580 (AtMYA1)a

XI

MD, CC, IQ

6. At MYA2

1505c 1515

AT5g43900 (AtMYA2)a

XI

MD, IQ

(Kinkema and Schiefelbein, 1994)

7. At XIA

1730

AT1g04600

XI

MD, CC, IQ


8. At XIB

1519

AT1g04160

XI

MD, IQ

9. At XIC

1572

AT1g08730

XI

MD, CC, IQ

10. At XID

1611

AT2g33240

XI

MD, CC, IQ

11. At XIE

1529

XI

MD, CC, IQ

XI

MD, IQ

XI

MD, CC, IQ

12. At XIF

1556

13. At XIG

1502

AT1g54560 d

AT2g31900

AT2g20290 AT4g28710

XI

MD, CC, IQ

1374

AT4g33200

XI

MD, CC, IQ

16. At XIJ

1242 963b 998b

AT3g58160 AtMYOS3)a (AtMYA3)a

XI

MD, CC, IQ

17. At XIK

1544

AT5g20490

IX

MD, CC, IQ

14. At XIH

1452

15. At XI-I

a

d

b


Name as reported in the literature; Number of amino acids previously reported for partial sequence; Number of amino acids predicted by NCBI; dEdited for full-length sequence; MD, motor domain; CC, Coiled-coil region; IQ, putative calmodulin-binding motif (Modified from Reddy and Day, 2001a) c

U

Structural features of Arabidopsis KLPs. The deduced amino acid sequence of KLPs in the Arabidopsis database was analyzed for the presence of various domains (Reddy and Day, 2001b). The KLPs are arranged according to their grouping in the phylogenetic tree in Fig. 18.3. Characterized KLPs are indicated in

Figure 18.1.

MCAK/KIF2

Unc104/KIF1

Sp

Chromo/KIF4

Sc

BimC

Ce

KRP85/95

Dm

KHC

At

MKLP1

10 8 6 4 2 0

C. Term

26 24 22 20 18

Kip3

Number of KLPs


bold. ARM, Armadillo/beta-catenin-like repeat; CBD, calmodulin-binding domain; CC, coiledcoil region; CH, Calponin homology domain; MD, motor domain; MyTH4, myosin tail homology region; Talin-like, talin-like domain; HhH1, helix-hairpin-helix domain. Bar 100 aa. Modified from Reddy and Day, 2001b.

KLPs (Fig. 18.1, Tab. 18.2). Six of the Arabidopsis KLPs have a calponin homology (CH) domain, which is an actin-binding domain present in the N-termini of spectrin-like proteins. The CH domain is a protein module of approximately 110 residues found in cytoskeletal and signal transduction proteins either as two domains in tandem or as a single copy (Banuelos et al., 1998). Proteins with a tandem pair of CH domains cross-link F-actin, bundle actin or connect intermediate filaments to cytoskeleton. Proteins with a single copy are involved in signal transduction (Banuelos et al., 1998, Leinweber et al., 1999). Perhaps the KLPs containing a CH domain bind actin and are involved in signal transduction or linking of actin and MTs. Some domains that are involved in protein protein or protein DNA interactions are also present in some KLPs (Fig. 18.1 and Tab. 18.2). These include armadillo repeats, a helix-hairpin-helix DNA-binding domain. KCBP has MyTH4 and talin-like domains present in some myosins, suggesting that it has domains of both MT- and actin-based motors. Such motors may be involved in cross talk between MT and actin cytoskeleton (Reddy, 2001b). KCBP also has been shown to have a calmodulin-binding domain (Reddy et al., 1996b).

Phylogenetic analysis In non-plant systems, nine subfamilies of KLPs with some ungrouped KLPs have been identified by phylogenetic analysis using the conserved motor domain (Kim and Endow, 2000). Three of the subfamilies (KHC, KRP85/95, and Unc104/KIF1) are involved in transport (Goldstein and Philip, 1999). Members 18.2.1.1

443

444


from the other subfamilies (C-terminal, Kip3, MKLP1, BimC, chromokinesin/ KIF4, and MCAK/KIF4) function in various processes associated with cell division (Goldstein and Philip, 1999). A phylogenetic analysis of the 61 Arabidopsis KLPs motor domain sequences with 113 other motor domain sequences from nonplant systems has revealed that seven of the nine families are represented in Arabidopsis (Figs 18.2 and 18.3). However, several Arabidopsis KLPs do not fall into any of the nine subfamilies and are likely to represent additional subfamilies that are unique to plants (Figs 18.2 and 18.3). Most of the Arabidopsis KLPs are more closely related to another Arabidopsis or another plant kinesin than to any other kinesin used in the comparison. The subfamilies that are involved in transport are under-represented in Arabidopsis (Fig. 18.2). There are no members of the KRP85/95 or Unc104/KIF1 subfamilies in Arabidopsis (Figs 18.2 and 18.3). No KHCs have been reported in plants. The phylogenetic tree indicates that there is possibly one KHC-type kinesin in Arabidopsis. At3g63480 falls into the KHC group with a closer relationship to KHCs found in fungi and, like fungal KHC, lacks a binding site for kinesin light chain proteins (Diefenbach et al., 1998; Reddy and Day, 2001b). One Arabidopsis kinesin groups with the MKLP1 subfamily (Fig. 18.3). Two internal motor Arabidopsis KLPs grouped with the MCAK/KIF2 subfamily (also called the internal family) whose members are involved in vesicle transport, chromosome movement and MT catastrophe (Goldstein and Philip, 1999). Three Arabidopsis KLPs fall into a group with Kip3 subfamily members in which ScKip3 is involved in nuclear movement (DeZwaan et al., 1997). Three Arabidopsis KLPs form a branch off of the chromokinesin/KIF4 subfamily members, some of which are involved in vesicle transport (HsKIF) and spindle organization and chromosome positioning (Xlklp1) (Goldstein and Philip, 1999). Arabidopsis has many KLPs (24 out of 61) that do not fall into any of the known groups (Figs 18.2 and 18.3). Members of BimC subfamily, which are N-terminal plus-end motors, are present in all five eukaryotic organisms that have been sequenced. The three NtKRP125-like Arabidopsis KLPs (AtKRP125a, b, and c) are grouped with the tobacco homolog in the BimC subfamily which are involved in cross-linking and anti-parallel sliding of MTs (Goldstein and Philip, 1999). Twenty-one Arabidopsis KLPs were grouped into the C-terminal subfamily. This is an unusually large number compared to the other organisms. It is also unusual in that 11 KLPs in this group have internal motors and five have N-terminal motors. The internal KLPs have a motor domain that is closer to the C-terminus than the N-terminus but each has some sequence C-terminal to the motor domain and could be called internal depending on the parameters used to define an internal motor. It will be interesting to find out the direction of movement of these KLPs whose motor domains are either N-terminal or internal but are most closely related to the C-terminal subfamily. KLPs in the C-terminal subfamily translocate toward the minus-end of MTs (Woehlke and Schliwa, 2000) and have a conserved sequence at the neck/motor core junction (Endow and Waligora, 1998). It was reported that the residues G and N residues at the neck/motor core junction are necessary for minus-end directed movement (Endow and Waligora, 1998). Analysis of the neck/motor core junction of the 21 C-terminal class Arabidopsis KLPs showed con-

18 Molecular Motors in Plant Cells Strict

BimC C-Terminal MKLP1

AnBIMC ScCIN8 ScKIP1 SpoCUT7 BmKRP DmKLP61F CeF23B12.8 HsKSP XlEg5 XlEg52 NTKRP125 AtKRP125b At3g45850 AtKRP125a AtKRP125c AnKLPA SpoKLP2 SpoPKL1 ScKAR3 DmNCD CgCHO2 MmKIFC1 HsCHO2 XlCTK2 At4g05190 AtKATA AtKATB AtKATC CeKLP HsKIFC3 MsFKIF2 MmKIFC2 XlCTK1 At5g27550 At2g22610 At1g72250 At3g10310 At1g18410 At1g73860 At1g63640 At5g41310 AtKATD At1g09170 At2g47500 At3g44730 CeC41G7.2 CeMO1E11 CeWO2B12 At5g10470 At5g65460 At5g27950 At1g55550 NtKCBP StKCBP AtKCBP SpKinesinC CeMO3D4.1 CgCHO1 HsMKLP1 DmPACKLP MmKlp174 At1g20060 CrKLP1 ScKIP2 HsCENPE At3g10180 At1g59540 UmKin1 At5g06670 At3g12020 AtMKRP1 AtMKRP2 At2g21380

AtHINKEL/NACK1

The number of KLPs in each family in different organisms. C.Term, C-Terminal motor; Chromo/KIF4, chromokinesin/KIF4; U, ungrouped. At, Arabidopsis thaliana, Dm, Dro-

Figure 18.2.

KRP85/95

AtNACK2 At4g38950 At2g21300 At3g51150 At5g66310 At5g42490 At4g24170 CeC06G3.2 CeLC33H5 CeF20C5 HsKIF3B MmKIF3B XlKlp3

sophila melanogaster; Ce, Caenorhabditis elegans; Sc, Saccharomyces cerevisiae; Sp, Schizosaccharomyces pombe. Adapted from Reddy and Day, 2001b.

servation of these residues in most of the C-terminal Arabidopsis KLPs (Reddy and Day, 2001b). Several Arabidopsis KLPs show some similarity to other ungrouped KLPs (Figs 18.2 and 18.3). The ungrouped centromeric proteins (HsCENPE and UmKin1) cluster with seven Arabidopsis KLPs. CENP-E binds to the kinetochore throughout mitosis and to MTs of the spindle mid-zone during late stages of mitosis (Goldstein and Philip, 1999). One Arabidopsis kinesin is paired with HsKid,

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Unc104/KIF1 MCAK/KIF2 KIP3 Chromo/KIF4 KHC

Phylogenetic tree of all Arabidopsis KLPs along with113 non-plant KLPs. The tree was generated using the motor domain of KLPs as described (Reddy and Day 2001a). Vertical dashes indicate ungrouped KLPs (light dashes, those grouped with other KLPs; bold dashes, exclusively Arabidopsis KLPs). The Arabidopsis KLPs are in bold. KLPs from the following organisms were used: An, Aspergillus nidulans; Bm, Bombyx mori; Ce, Caenorhabditis elegans; Cf, Cylindrotheca fusiformis; Cg, Cricetulus griseus; Cr, Chlamydomonas rheinhardtii; Dd,

Figure 18.3.

KRP85/95

HsKIF3C MmKIF3C RnKIF3C SpKIN95 CrFLA10 HsKIF3 MmKIF3A SpKIN2A DmKlp68D CeOSM3 CeF56E3 DmKlp73 CeUNC104 HsATSV MmKIF1A MmKIF1B RnKIF1B HsbKIF1C RnKIF1D CeLR144 Dm38B HsCMKRP CeK11d9.C CgMCAK RnKRP2 HsMCAK XlKCM1 HsKin2 MmKIF2 MmKIF2C XlKCM2 LmKIN At3g16630 At3g16060 CfDSK1 HsKid At5g02370 DmKlp67A At1g18550 ScKIP3 ScVII607 SpoBC2F12.13 SpoBC649.01C At3g49650 At5g23910 CeTO1G1 Ggchrkin HsKIF4 MmKIF4 Xlklp1 At3g50240 At5g47820 At5g60930 Dmklp3A CeY43F4B CeKHC DmKHC LpKHC HsKHC MmKHCS MmKHCx HsnKHC HsxKHC MmKIF5c MmKHC SpKHC NcKHC NhKin1 UmKin2 SyKin1 At3g63480 DdK7 CeLF22F4 DmNOD At1g01950 At1g12430 At3g54870 LcKIN Xlklp2 At3g19050 At3g17360 At3g44050 At3g20150 At3g23670 AtPAKRP1 AtPAKRP2 ScSMY1

Dictyostelium discoideum; Dm, Drosophila melanogaster; Gg, Gallus gallus; Hs, Homo sapiens; Lc, Leishmania chagasi; Lm, Leishmania major; Lp, Loligo pealii; Mm, Mus musculus; Ms, Morone saxatilis; Nc, Neurospora crassa; Nh, Nectria haematococca; N, Nicotiana tabacum; St, Solanum tuberosum; Rn, Rattus norvegicus; Sc, Saccharomyces cerevisiae; Sp, Strongylocentrotus purpuratus; Spo, Schizosaccharomyces pombe; Sr, Syncephalastrum racemosum; Um, Ustilago maydis; Xl, Xenopus laevis. Modified from Reddy and Day 2001b.


an ungrouped kinesin. HsKid is a kinesin-like DNA-binding protein that is involved in spindle formation and the movements of chromosomes during mitosis and meiosis (Goldstein and Philip, 1999). Arabidopsis PAKRP1 along with five other Arabidopsis KLPs are grouped with XlKlp2. XlKlp2 is required for centrosome separation and maintenance of spindle bipolarity (Boleti et al., 1996) whereas PAKRP1 associates with the phragmoplast (Lee and Liu, 2000) and is expected to function in cytokinesis. Three Arabidopsis KLPs form a subgroup separate from any other kinesin but share a branch with CeLF22F4 and DmNOD (Fig. 18.3). Eight other Arabidopsis KLPs form a subgroup separate from KLPs of any other organism. In many cases a group of Arabidopsis KLPs forms a separate branch within the major subgroup in which they fall. The large number of KLPs in plants is expected to control many diverse cellular processes including some plant-specific functions. There are many MT-associated processes that are unique to plants (Lloyd and Hussey, 2001, Reddy, 2001b). For example, during cell division in plants several plant-specific MT arrays such as the preprophase band and the phragmoplast are formed that are important in determining the future location of the cell wall and cell wall formation, respectively (Lloyd and Hussey, 2001). These unique processes are likely to require additional plant-specific motors. In addition, centrosomes play an important role in MT organization in animals whereas plants have no well-defined centrosomes. Hence, MT organization and dynamics in plants may also require additional MT motors. The presence of unique organelles such as chloroplasts and MT-dependent processes such as cell wall synthesis may also warrant additional motors. Also, cell to cell transport of macromolecules such as RNA and viruses through plasmodesmata may require MT-based motors (Reddy, 2001b). It is very likely that plant KLPs may participate in functions other than transport, and MT dynamics and organization. Recently, it has been shown that two plant KLPs function as activators of a protein kinase (Nishihama et al., 2002). It appears that flowering plants do not have dyneins. Hence, some KLPs may be performing functions that are carried out by dyneins in other systems. 18.2.2

Dyneins

Dyneins move along MTs toward the minus ends. The motor domain in dyneins (Z 1000 amino acids) is much longer than in kinesins (Z 340 amino acids) and does not share sequence similarity with the kinesin motor domain. Cytoplasmic dynein is a large multi-subunit complex consisting of two heavy chains (Z 530 kDa each), three intermediate chains (74 kDa), and four light intermediate chains (Z 55 kDa) (Hirokawa, 1998). The central and C-terminal regions of dynein heavy chain, which are predicted to form a globular structure, interact with MTs and contain motor activity whereas the N-terminal region is thought to bind cargo. The intermediate and light intermediate chains of cytoplasmic dynein associate with a protein complex called dynactin which is composed of 10 subunits (Hirokawa, 1998). Although there is some immunological and biochemical data in-

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18.3 Actin-based Motors

dicating the presence of dynein heavy chains in plants (Moscatelli et al., 1995), sequences that are similar to dynein and dynactin subunits have not been found in the Arabidopsis genome or plant EST databases (Lawrence et al., 2001), suggesting that they may be absent in flowering plants. The functions performed by dyneins in animals may be carried out by the expanded family members of the kinesin superfamily. Alternatively, it is possible that plant dyneins have diverged considerably, hence are not recognizable by the current search programs.

18.3

Actin-based Motors 18.3.1

Myosins

Myosins, a diverse group of actin-based molecular motors, perform a broad array of cellular functions. A large number of myosins have been described in many eukaryotes (Berg et al., 2001, Reddy and Day, 2001a). Most of these are identified based on sequence similarity with the myosin motor domain. All myosin heavy chains have three common domains: a highly conserved motor domain (Z 850 amino acids) located at the amino-terminus in almost all myosins, which is followed by a neck region that binds to calmodulin or calmodulin-related proteins and a C-terminal tail domain. The calmodulin-binding domain ‘IQ motif’ in these myosins has a consensus sequence ‘IQXXXRGXXXR’ (I, isoleucine; Q, glutamine; R, arginine; G, glycine) (Wolenski, 1995). This motif is present in one to seven repeats depending on the type of myosin (Mermall et al., 1998). The only myosin that does not contain an ‘IQ motif’ is myosin XIV. The tail region of different myosins varies considerably in length and structure. The tail is characteristic to each myosin with interesting domains that are also found in non-motor proteins. The number of myosins in an organism varies among organisms. However, not all types of myosins have been identified in a single organism (Berg et al., 2001, Reddy and Day, 2001a). Although there is compelling indirect evidence for the role of actin and actinbased motors in various transport processes in plants, little is known about specific plant myosins that perform these functions. Myosins have been identified in plants both biochemically (Heslop-Harrison and Heslop-Harrison, 1989, Miller et al., 1995, Yokota et al., 1995, 1999b, Yokota and Shimmen, 1994, 1999) and at the molecular level (Kinkema and Schiefelbein, 1994, Kinkema et al., 1994, Knight and Kendrick-Jones, 1993). Immunological detection of myosins using animal myosin antibodies identified proteins of various sizes from different plants (Parke et al., 1996, Qiao et al., 1989, Tang et al., 1989). Immunofluorescence studies localized myosin to the surface of organelles, and the vegetative nuclei and generative cells in pollen grains and tubes (Heslop-Harrison and Heslop-Harrison, 1989), to the active streaming lanes and cortical surface in pollen tubes (Miller et al., 1995) and more recently to plasmodesmata in root tissues (Radford and White, 1998, Reichelt et al., 1999). Motility assays (Kohno et al., 1991) and ATPase assays


(Kohno et al., 1991, Ma and Yen, 1989; Vahey et al., 1982) using myosin-like proteins isolated from plants have also demonstrated the presence of myosins in plants. Using PCR-based approaches, a few myosins have been cloned from Arabidopsis and other plants (Kashiyama et al., 2000, Kinkema and Schiefelbein, 1994, Kinkema et al., 1994, Knight and Kendrick-Jones, 1993, Liu et al., 2001, Moepps et al., 1993). A 170-kDa myosin has been purified from lily pollen and the antibodies against this myosin stained particles of various sizes in the apical cytoplasm of lily and tobacco pollen tubes (Kohno et al., 1990, 1991, 1992, Yokota and Shimmen, 1994). These reports indicate that several myosins are likely to be present in pollen tubes. The 170- and 175-kDa myosins that translocate F-actin at a velocity of about 9 and 3 4 mm s 1, respectively, were found to be associated with calmodulin (Yokota et al., 1999a, 1999b). A myosin II-like protein was identified in Nitella by Kato et al. (Kato and Tonomura, 1977) and a myosin was purified from Chara (Yamamoto et al., 1994, 1995). In in vitro motility assays, the characean myosin translocated actin at 50 mm s 1. Rotary-shadowed images of the purified myosin showed two heads that are comparable to heads of myosin II in shape and size (Yamamoto et al., 1995). In Arabidopsis, the expression of several myosins in a given cell type implies that actin-based motors are involved in a wide range of cellular functions. By analyzing the Arabidopsis genome sequence with the conserved motor domain of myosins, a total of 17 myosins were identified in Arabidopsis (Reddy and Day, 2001b). Table 18.3 lists the myosins by name as given in the phylogenetic tree by Hodge and Cope (2000) and as assigned by Reddy and Day (2001a). In comparison, S. cerevisiae, Schizosaacharomyces pombe, C. elegans, D. melanogaster and H. sapiens have 5, 5, 17,13 and 40 myosins respectively. Five of the 17 Arabidopsis myosins have been reported in the literature (Kashiyama et al., 2000, Kinkema and Schiefelbein, 1994, Kinkema et al., 1994, Knight and Kendrick-Jones, 1993). All Arabidopsis myosins have three to six putative calmodulin-binding ‘IQ’ motifs (Fig. 18.4). The IQ domains usually follow right after the motor domain but are separated slightly from the motor domain in At XID, At XI-I, and At XIK. There are three or four IQ domains in class VIII myosins and five or six in class XI except for At XIK which has only four (Fig. 18.4). There are coiled-coil domains in all the myosins that differ in length and number. They often follow directly after the IQ domains but in some cases there is intervening sequence. Based on the presence of the coiled-coil domains, it is likely that the Arabidopsis myosins either form dimers or interact with other proteins (Cope et al., 2000). The class XI myosins are much longer than the class VIII myosins with the difference being in the length of the Cterminal region following the conserved domains found in myosins. Myosins containing IQ domains are typically calmodulin-sensitive. However, the interaction with, or regulation by calmodulin has not yet been demonstrated for Arabidopsis myosins. The information on myosin light chains is limited in plants. Recently, Yokota and his colleagues (Yokota et al., 1999a, Yokota and Shimmen, 1994, 1999) have purified two myosins from plants and demonstrated that calmodulin associates with the purified myosin and regulates its motor activity. Since the genes encoding these proteins are not cloned the structural features of these myosins are

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18.3 Actin-based Motors 1. At ATM 3. At VIIIA 2. At ATM2 4. At VIIIB 13. At XIG 14. At XIH 6. At MYA2 8. At XIB 10. At XID 7. At XIA 12. At XIF 9. At XIC 11. At XIE 16. At XIJ

5. At MYA2 15. At XI-I 17. At XIK

Motor Domain

IQ Motif

Schematic diagram showing various predicted domains in Arabidopsis myosins. The Arabidopsis genome database was searched with the conserved motor domain of a myosin to identify Arabidopsis myosins. The deduced amino acid sequence of Arabidopsis myosins

Figure 18.4.

Coiled-coil Domain

was analyzed for the presence of various domains (Reddy and Day, 2001a). The numbers refer to the numbers in Tab. 18.1. The first four myosins are in class VIII and the following 13 are in class XI. The bar represents 100 amino acids. Adapted from Reddy and Day, 2001a.

not known. Besides the motor, IQ and coiled-coil domains, several other domains have been identified in myosins from classes other than the plant classes VIII and XI. These include SH3 domains (Src homology 3 domains), ankyrin repeats, MYTH4 (domain of unknown function found in a few classes of myosins), zincbinding domain, plecstrin homology, FERM/talin (band 4.1/ezrin/radixin/moesin), GPA-rich domains and a protein kinase domain (Berg et al., 2001). Plant myosins do not have any of these other domains.


18.3.2

Phylogenetic analysis

Phylogenetic analysis using the conserved motor domain grouped all known myosins into 18 distinct classes (Burge and Rogers, 2000). Phylogenetic analysis of the Arabidopsis myosins’ motor domain with non-plant and plant myosins revealed that all the Arabidopsis myosins and other plant myosins fall into only two groups, class VIII and class XI (Fig. 18.5). Four Arabidopsis myosins are in class VIII and 13 in class XI. These groups contain exclusively plant or algal myosins with no animal or fungal myosins, suggesting that plants have a unique set of myosins (Fig. 18.5). The motor domain in all cases is in the N-terminal region (Fig. 18.4). The motor domain starts at about 50 55 residues for the class XI myosins while the class VIII myosins have a longer N-terminal region prior to the motor domain (99 159 residues). An algal (Chara corallina) myosin, Cc ccm, does group with the plant class XI myosins but is on a separate branch from any other vlass XI myosin (Fig. 18.5). A phylogenetic tree that was constructed using the full-length sequences of Arabidopsis and other plant myosins showed similar grouping of plant myosins (Reddy and Day, 2001a). Among the class XI myosins the similarity ranges from 40 85 % (full length) and 61 91 %(motor domain). The similarity between the class VIII myosins ranges from 50 83 % (full length) and 64 92 % (motor domain). When class VIII myosins are compared to class XI myosins the similarity only ranges from 22 29 % (full-length) and 35 42 % (motor domain). Myosins have 131 highly conserved residues spread throughout the motor domain that define a core consensus sequence (Cope et al., 2000). Comparison of an alignment of Arabidopsis myosin motor domains to these conserved sequences shows a great deal of conservation among them (Reddy and Day, 2001a), suggesting that they are capable of motor function.

18.4

Cellular Roles of Motors 18.4.1

Cell Division

Although many of the dynamic processes of cell division such as formation of the spindle, congression of chromosomes at the equatorial region and migration of chromatids during cell division are common to both animals and plants, there are significant differences between plant and animal cell division. For example, the dividing plant cells organize their cytoskeletal elements into unique structures that are not found in non-plant systems. Just prior to prophase, plant cortical MTs rearrange to form a band of MTs called the pre-prophase band (PPB) (Goddard et al., 1994, Gunning and Wick, 1985), which accurately predicts the future location of the cell plate. Another distinctive feature of plant mitosis is the formation of a bipolar spindle in the absence of centrosomes (Franklin and Cande, 1999, Smir-

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Phylogenetic analysis of Arabidopsis myosins. A phylogenetic tree was generated as described earlier (Reddy and Day, 2001a). The myosin groups, as defined by Hodge and Cope (2000) and Yamashita et al (2000), are identified on the left in roman numerals. The Arabidopsis myosins are in bold. Myosins from the following organisms were used: Ac, Acanthamoeba castellani; Acl, Acetabularia cliftoni; Cc, Chara corallina, Ha, Helianthus annuus;

Figure 18.5.

At ATM At VIIIA Ha hamy1 Zm ZMM3 At ATM2 At VIIIB Ha hamy3 At MYA1 At XIC At XIE At XIJ AT XIK Ha hamy2 Ha hamy5 Zm MYO1 At MYA2 At XIB Ha hamy4 At XIG At XIH At XIA At XID At XIF At XI-I Cc ccm Acl myo1 Acl myo2 Ac HMWMI Bt X Mm X Ce HUM-2 Mm Dilute Rn myr6 Sc Myo2 Ce HUM-3 Dm 95F Mm Waltzer Ce HUM4 En csmA Pg csm1 Ce HUM-6 Dm 35B Hs UsherIb Ce HUM-7 Hs IXa Mm IXb Rn myrV Ce IA Dm IB Hs Ib Sc MYO3 Dd IA Ce IIB Dm IIB Dd II Hs nmIIA Sc MYOI Dm PDZA Hs MysPDZ Dd MyoI Dd MyoJ Dd myoM Dm NinaC Lp III Hs XV Mm XV Pf PfM-A Tg myoA Rn XVI

VIII

XI

XIII IV X V VI XII XVII VII IX

I

II XVIII

III XV XIV XVI

Zm, Zea mays; Bt, Bos taurus; Mm, Mus musculus; Ce, Caenorhabditis elegans; Dm, Drosophila melanogaster; Rn, Rattus norvegicus; Sc, Saccharomyces cerevisiae; Hs, Homo sapiens; Dd, Dictyostelium discoideum; Lp, Limulus polyphemus; En, Emericella nidulans; Pg, Pyricularia grisea; Pf, Plasmodium falciparum; and Tg, Toxoplasma gondii. The numbers at the branches indicate the number of times the dichotomy was supported out of 100 bootstrap tries.


nova and Bajer, 1992). The structures involved in the organization of MTs are the centrosomes in animal cells and the spindle pole body (SPB) in fungal cells, which are lacking in plant cells (Smirnova and Bajer, 1992). Finally, cytokinesis, the process that produces two daughter cells following the completion of nuclear division, in plants is accomplished quite differently from animals. Cytokinesis in plant cells occurs via the formation of a polysaccharide cell plate, which expands from the center to the periphery (Staehelin and Hepler, 1996, Sylvester, 2000). The phragmoplast, a structure composed of two disks of parallel MTs with their plus ends toward the equatorial zone and actin filaments (Euteneur et al., 1982, Hepler and Jackson, 1968, Staehelin and Hepler, 1996, Wick, 1991), is involved in cell plate formation. Vesicles carrying the cell plate materials are transported to the equatorial zone of the phragmoplast (Samuels et al., 1995, Staehelin and Hepler, 1996, Yasuhara et al., 1993). The association of vesicles with phragmoplast MTs indicate that MT-based motors are likely to be involved in transporting the vesicles to the cell plate (Otegui and Staehelin, 2000, Otegui et al., 2001, Samuels et al., 1995). The fusion of vesicles in the cell plate is mediated by phragmoplastin, a dynamin homolog, in plants (Gu and Verma, 1996, 1997, Samuels et al., 1995). The plant spindle, which is formed in the absence of a well-defined centrosome, and several unique plant-specific MT arrays such as the pre-prophase band and the phragmoplast (Lloyd and Hussey, 2001) suggest that plants are likely to contain additional plant-specific KLPs that are not present in animal cells. In animals, six subfamilies of KLPs have been shown to be involved in some aspect of cell division (Goldstein and Philip, 1999, Reddy, 2001b). Several recent reports on KLPs expression and immuunolocalization indicate that at least seven plant KLPs (Kat A, KatB/C, TKRP125, AtKCBP, AtPAKRP1, AtPAKRP2, DcKRP120-2, HINKEL/AtNACK1) that belong to different families have mitotic function (Asada, 1996, Barroso et al., 2000, Bowser and Reddy, 1997, Lee and Liu, 2000, Lee et al., 2001, Liu et al., 1996, Nishihama et al., 2002, Smirnova et al., 1998, Strompen et al., 2002). Some KLPs are expressed in a cell cycle-dependent manner. TKRP125, a member of the BimC subfamily and NCAK from tobacco and two KLPs from Arabidopsis (KCBP, KatB/C) show a high level of expression in M-phase of the cell cycle (Asada, 1996, Bowser and Reddy, 1997, Mitsui et al., 1996, Nishihama et al., 2002). DcKRP120-1 and DcKRP120-2 are not detectable in non-dividing cells (Barroso et al., 2000). TKRP125 localizes to the cortical MTs in the S phase, along MTs of pre-prophase band (PPB) and perinuclear MTs in prophase, the equatorial plane of spindle MTs in metaphase and anaphase, and the phragmoplast MTs in telophase and cytokinesis. A carrot homolog of TKRP125 (DcKRP120-1) also localizes to the cortical MTs, the pre-prophase band, spindle and the phragmoplast (Barroso et al., 2000). Antibodies to another member of the BimC subfamily (DcKRP120-2) stained the spindle and phragmoplast, predominantly in the mid-line of the phragmoplast where plus ends of the MTs overlap (Barroso et al., 2000). AtPAKRP1, a non-member of BimC, which resembles an ungrouped Xklp2 from Xenopus, also localizes to the mid-line of the phragmoplast. KatA, a C-terminal motor, localizes near the mid-zone at metaphase and anaphase and the phragmoplast in Arabidopsis and tobacco BY-2 suspension

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cells (Liu et al., 1996). Anti-KCBP antibodies stain the pre-prophase band, the spindle and the phragmoplast (Bowser and Reddy, 1997). The KCBP antibody did not stain the cell plate region of the phragmoplast but MT bundles on either side of the cell plate are strongly labeled (Fig. 18.6). Localization of KCBP in Haemanthus also showed localization to mitotic MTs arrays. In Haemanthus anti-KCBP staining is seen almost exclusively at the spindle poles in late anaphase (Smirnova et al., 1998). The assembly of the acentriolar spindle in plants may involve convergence of MT minus-ends leading to the formation of spindle poles (Franklin and Cande, 1999, Smirnova and Bajer, 1992). The fact that KCBP is a minus-end-directed motor (Song et al., 1997) and localizes to the spindle poles suggests that it may be involved in acentriolar spindle formation in plants. Between early and late anaphase, the localization of KCBP shifts toward the spindle pole, supporting a role for KCBP in the formation of a converging bipolar spindle (Smirnova et al., 1998, Smirnova and Bajer, 1992). Localization of minus-end (KatA and KCBP) and plus-end (TKRP125) motors in the spindle suggests their involvement in counterbalancing forces generated by plus- and minus-end motors to stabilize the spindle. Constitutive activation of KCBP during late prophase caused premature breakdown of the nuclear envelope and arrest of cells at pro-metaphase whereas activation of KCBP at late metaphase or anaphase did not effect the progression of anaphase but caused aberrant phragmoplasts formation and delayed cytokinesis (Vos et al., 2000). This study suggests that KCBP is differentially active during the various phases of cell division. Its activity is downregulated in metaphase and telophase and upregulated in anaphase most likely by changes in cytosolic calcium level (Vos et al., 2000). Co-localization of some of the KLPs with mitotic MT arrays suggests that they may have a role in forming these arrays by bundling MTs. The motor domain of KCBP has been shown to bundle MTs (Kao et al., 2000). Numerous forces are at play in the development, maintenance, and function of the phragmoplast. Golgi-derived vesicles are transported to the forming cell plate where the plus-ends of phragmoplast MTs interdigitate. It is likely that MT motors are involved in forming the phragmoplast and powering the movement of the vesicles along the MTs. Seven plant KLPs (Kat A, TKRP125, AtKCBP, AtPAKRP1, AtPAKRP2, DcKRP120-2, HINKEL/AtNACK1) including plus- and minus-end motors and some plant-specific KLPs are associated with the phragmoplast and so are expected to function in cytokinesis (Asada, 1996, Barroso et al., 2000, Bowser and Reddy, 1997, Lee and Liu, 2000, Lee et al., 2001, Liu et al., 1996, Nishihama et al., 2002, Smirnova et al., 1998, Strompen et al., 2002). The localization patterns with various KLPs differed considerably. Immunolocalization studies as well as phragmoplast MT gliding assay in the presence of TKRP antibodies indicates its involvement in the organization of MT arrays especially in phragmoplast function (Asada et al., 1997). The polarity of MTs and microfilaments in the phragmoplast (Euteneur et al., 1982, Staehelin and Hepler, 1996) indicates that plus-end motors are likely to be involved in the transport of vesicles to the cell plate. Of the seven KLPs that localize to the phragmoplast, five are N-terminal motors (TKRP125, PAKRP1, DcKRP120-2, PAKRP2, HIK/NCAK) and two are C-terminal (KatA and KCBB). TKRP125 is a plus-end motor, and the other four N-terminal


DAPI

MT

Triple localization of nucleus, MTs and KCBP in a cytokinetic cell of tobacco. Nuclei stained with DAPI (DAPI), MTs (MT) are localized with a monoclonal antibody to b-tubulin and KCBP (KCBP) was localized with affinitypurified anti-KCBP antibody. Secondary antibodies conjugated to FITC and Cy3 were used to detect MTs and KCBP, respectively. From Bowser and Reddy, 1997.

Figure 18.6.

KCBP

KLPs are likely plus-end motors. One or more of these are likely to be involved in vesicle transport to the cell plate. The punctate staining pattern of PAKRP2 and its association with vesicles, suggest that it is involved in transporting vesicles to the cell plate (Lee et al., 2001). It is unlikely that KCBP and KatA play a role in vesicle transport to the cell plate as these are known to be minus-end motors and the MTs are oriented with their plus ends facing the cell plate (Euteneur et al., 1982, Liu et al., 1996, Song et al., 1997). The possible functions of minus-end motors are stabilization of the phragmoplast and/or recycling of Golgi vesicle membranes from the expanding cell plate. Functional analysis of TKRP125 indicates its role in crosslinking and sliding of anti-parallel MTs in the phragmoplast (Asada et al., 1997, Asada and Shibaoka, 1994). Immunofluorescence with AtPAKRP1 antibodies showed staining of interzonal MTs at late anaphase and later on its localization is restricted to the plus ends of interdigitating MTs in the phragmoplast (Lee and Liu, 2000). Functional studies with this KLP indicates its involvement in maintaining the integrity of phragmoplast MTs (Lee and Liu, 2000). In order for the growth of the phragmoplast to be achieved, a rapid turnover of phragmoplast MTs must occur (Hepler and Hush, 1996, Hush et al., 1994). The expansion of

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the phragmoplast from the center to the periphery involves disassembly of the MTs inside and assembly of MTs at the outside. Blocking of disassembly in the center by MT stabilizing drugs causes cessation of phragmoplast expansion and formation of incomplete cell walls (Nishihama and Machida, 2001, Yasuhara et al., 1993). Phragmoplast-associated KLPs may play a role in the MT assembly and disassembly associated with its growth. The loss of MTs from the center of the phragmoplast as it grows does not occur in hik mutants, suggesting that it controls directly or indirectly the dynamics of MTs in the phragmoplast (Strompen et al., 2002). HIK does not appear to be involved in either vesicle transport or organization of MTs in the phragmoplast. Studies with tobacco NCAK1, a homolog of HINKEL, showed its localization to the equatorial zone of the phragmoplast but not in the inner regions of the phragmoplast where the cell plate matures (Nishihama et al., 2002). It appears that NCAK through a protein kinase, NPK1, regulates depolymerization of MTs in the center of the phragmoplast as it expands (Nishihama et al., 2001, 2002). There is some evidence for the presence of a myosin in the phragmoplast (Parke et al., 1996), suggesting that actin-based motors may also be involved in the transport of vesicles on the phragmoplast. During cell division the anti-class VIII myosin staining remains confined to the transverse cell walls and is strongest in the newly formed cell wall (Reichelt et al., 1999). Immunogold electron microscopy showed labeling of class VIII myosin associated with the plasma membrane and plasmodesmata. These studies suggest that class VIII myosins may be involved in new cell wall formation and transport in the plasmodesmata. 18.4.2

Cell Polarity and Morphogenesis

Both actin and MT cytoskeleton have been implicated in cell polarity and morphogenesis (Mathur and Chua, 2000, Mathur et al., 1999, Reddy and Day, 2000, Szymanski et al., 1999, Vidali and Hepler, 2001). The first indication that a KLP might play a role in morphogenesis came from studies with the zwichel mutant. In wild-type Arabidopsis, trichomes are unicellular with a stalk and three branches (Oppenheimer, 1998). In zwichel (zwi) mutants trichomes are abnormal with a short stalk and one or two branches depending on the severity of the allele (Fig. 18.7). Cloning of ZWICHEL has indicated that it is identical to KCBP, a calmodulin-binding KLP (Oppenheimer et al., 1997), suggesting the requirement of this motor for expansion of the stalk and branching. Using paclitaxel, a known MT stabilizer, zwichel mutants were induced to form similar growth points indicative of branch formation in normal trichomes (Mathur and Chua, 2000). Transient stabilization of MTs could compensate for KCBP/ZWI activity suggesting that it may be involved in stabilization of MTs. The two MT-binding sites of KCBP (Narasimhulu and Reddy, 1998) could be involved in stabilization or possibly a KCBP/ ZWI-interacting protein(s) could be responsible for the stabilization. Several alleles of zwi have been characterized. All zwi mutants grew normally with no apparent defects in cell division except that they contain abnormal trichomes (Krishnakumar and Oppenheimer, 1999, Oppenheimer et al., 1997). The lack of a phenotype in


Scanning electron micrographs of Arabidopsis trichomes of wild-type and KLP (zwichel) mutants. (a) Wild-type; (b) zwi; (c) zwi w2; (d) zwi9311-11. In zwi w2 most trichomes have two branches but the length of the second branch is varied. In the zwi9311-11 mutant

Figure 18.7.

about 40 % of trichomes are unbranched and the rest have two branches. Seeds of zwichel mutants were kindly provided by Dr David Oppenheimer and Dr Martin Hülskamp. Scale bar 100 mm. From Reddy and Day, 2000.

other tissues suggests that either KCBP is non-essential or another motor with overlapping functions may substitute for ZWI function in other tissues. In Arabidopsis there are 61 KLPs including several C-terminal motors. Three extragenic suppressors (suppressor of zwichel-3; suz1, suz2 and suz3) that rescued the trichome branch number defect in a zwichel mutant have been isolated (Krishnakumar and Oppenheimer, 1999). All three suppressors were found to be allele specific, indicating direct interactions between these proteins. The suz1 zwi-3 double mutants are male sterile due to a defect in pollen germination and pollen tube growth, suggesting a role for these genes in pollen germination and tube growth (Krishnakumar and Oppenheimer, 1999). At least two proteins, a plant-specific protein kinase and a protein required for trichome branching (ANGUISTIFOLIA), have been shown to interact with KCBP/ZWI (Day et al., 2000, Folkers et al., 2002). 18.4.3

Cytoplasmic Streaming

Cytoplasmic streaming in plant cells is important in intracellular transport. The force that drives the cytoplasmic streaming appears to be dependent on actin and actin-based motors (Shimmen and Yokota, 1994, Staiger and Schilwa, 1987, Williamson, 1976). Insight into the function of plant myosins in cytoplasmic streaming has been gained primarily by studies carried out with algae and pollen tubes. Characean cells exhibit a very rapid cytoplasmic streaming (Z 70 mm s 1). Also in pollen tubes, vesicles flow rapidly from the shank to the tip of the pollen tube where they fuse to support extensive tip growth of these cells (Emons et al., 1991, Heslop-Harrison and Heslop-Harrison, 1990). In addition to the bi-directional movement of vesicles, the generative cell and the nucleus move unidirectionally toward the tip (Mascarenhas, 1990, 1993). In Chara, an increase in Ca2 concentration causes cytoplasmic streaming to stop (Hayama et al., 1979). A myosin isolated from the alga Chara corallina was shown to be responsible for cytoplasmic

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streaming (Yamamoto et al., 1994, 1995, 1999). The myosin was cloned and found to be a class XI myosin related to the Arabidopsis MYA myosins (Kashiyama et al., 2000). Using immunofluorescence, myosin was localized to vesicles, organelles and generative cells and vegetative nuclei in grass pollen tubes (Heslop-Harrison and Heslop-Harrison, 1989). A myosin isolated from lily pollen has been shown to be responsible for cytoplasmic streaming in pollen tubes and two myosins identified in tobacco cell cultures are also thought to participate in cytoplasmic streaming (Yokota and Shimmen, 1994, Yokota et al., 1999b). Myosin has been localized to vesicles, chloroplasts and other organelles, indicating their involvement in transport of these organelles (Heslop-Harrison and Heslop-Harrison, 1989, Miller et al., 1995). The stop-and-go movements of Golgi stacks in plants also appear to be dependent on myosin (Nebenfuhr et al., 1999). However, specific myosin(s) responsible for the movement of these organelles have not been identified. Biochemical properties and the localization data of two lily pollen myosins (175 and 170 kDa) suggest that these myosins are at least partially responsible for cytoplasmic streaming, organelle transport and calcium sensitivity of cytoplasmic streaming (Yokota et al., 1999a,1999b, Yokota and Shimmen, 1999). It has been shown that the directional movement of mitochondria is dependent on the F-actin and myosin system (Van Gestel et al., 2002). Unlike in animals, the movement of peroxisomes in plants is actin based and actin polymerization appears to drive the movement of these organelles (Mathur et al., 2002). 18.4.4

Microtubule Dynamics and Organization

Microtubules are highly dynamic structures with a half-life of an individual MT of about 10 min. Drugs that disrupt the dynamics of MTs have been shown to severely hamper the MT-dependent process (Reddy, 2001b). Treatment of dividing cells with colchicine leads to disappearance of the mitotic spindle, suggesting that MTs in the spindle undergo continuous polymerization and deploymerization. Taxol, in contrast to colchicine, binds to MTs and stabilizes them. Addition of taxol to dividing cells leads to arrest of dividing cells in mitosis. The dynamics of the MT cytoskeleton during cell division in plant cells have been studied extensively (Baskin and Cande, 1990, Hepler and Hush, 1996). The rapid turnover of plant MTs suggests that the transition between MT arrays and change in orientations may involve depolymerization/re-polymerization of MTs and/or movement of MTs. Dynamic instability of MTs is important for their normal function. For example, stabilization of MTs in elongating root hairs causes loss of growth directionality and promotes branching (Bibikova et al., 1999). The MTs of the phragmoplast have a t of 60 s (Hush et al., 1994). However, the mechanisms that regulate the formation and dynamic instability of MT arrays in plants are not understood. Several aspects of MT dynamics such as cross-linking, zippering, bundling, sliding, and stability of MTs is regulated by motor proteins (Goldstein and Philip, 1999). KCBP like Ncd in Drosophila can cross-link and zipper MTs to focus MTs to form spindle poles (Chandra et al., 1993, Kao et al., 2000, Matthies et al., 1996, Smirnova et al., 1998). Two dis-


tinct MT binding sites and minus-motor activity enable these proteins to perform such a function. In addition C-terminal motors have been shown to bundle MTs (Chandra et al., 1993, Kao et al., 2000). Some motors are implicated in polymerization and depolymerization of cytoskeletal elements. There is at least one example where a KLP has no motor activity but controls MT dynamics (Desai et al., 1999). Plant KLPs such as NCAK1 appear to be involved in MT dynamics (Nishihama et al., 2002, Strompen et al., 2002). Hence, the paradigm that motors bind cargo and move along cytoskeletal tracks may not explain the functions of all the motors. 18.4.5

Intercellular Transport

Plant cells are connected through plasmodesmata to form a continuous symplastic network. Recent studies show that macromolecules such as RNA and protein move from cell to cell through plasmodesmata (Lazarowitz and Beachy, 1999, Lucas et al., 1995, McLean et al., 1995, 1997). The cytoskeleton and molecular motors have been implicated in this regulated transport of macromolecules through plasmodesmata (Blackman and Overall, 1998, McLean et al., 1995, White et al., 1994). The movement protein of viruses which enables the movement of viruses from cell to cell colocalizes to MTs, suggesting a role for cytoskeleton and possibly molecular motors in the transport of protein from cell to cell through plasmodesmata (Heinlein et al., 1995, Lazarowitz and Beachy, 1999, Mas and Beachy, 1999, 2000, McLean et al., 1995). These studies hint at an interesting possibility that motors may play a role in transporting macromolecules from cell to cell and influence cell-to-cell communication. Immunolocalization studies have also detected myosin associated with plasmodesmata. A recent study using an antibody to a cloned class VIII Arabidopsis myosin ATM1 (At ATM) localized this myosin to the plasmodesmata and the plasma membrane regions involved in the assembly of new cell walls (Reichelt et al., 1999). Earlier work suggested that actin was involved in regulation of plasmodesmal transport (Ding et al., 1996). Other studies using antibodies to animal myosins in root tissues of Allium cepa, Zea mays and Hordeum vulagare have also indicated the presence of myosin in the plasmodesmata (Radford and White, 1998). 18.4.6

Other functions

It appears that certain plant motors may perform unexpected novel functions. For example, two plant KLPs (NCAK1 and NCAK2) have been shown to interact with NPK1, a MAP kinase kinase kinase, and stimulate its activity (Nishihama et al., 2002). Motors in plants may also be involved in regulating cell polarity, development and differentiation by transporting the proteins or mRNA to localized regions within the cell (Arn and MacDonald, 1998). There is substantial indirect evidence indicating that asymmetric localization of mRNA (e. g. ASH1 mRNA in yeast), protein (e. g. calmodulin in Drosophila photoreceptors, a MAPKKK in

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18.5 Regulation of Motors

tobacco BY2 cells) and RNA protein complexes in some eukaryotic cells is dependent on cytoskeletal motors (Bertrand et al., 1998, Carson et al., 1997, 1998, Long et al., 1997, Munchow et al., 1999, Nishihama et al., 2002, Oleynikov and Singer, 1998, Perhonen et al., 1998, Porter et al., 1993, 1995; Takizawa et al., 1997).

18.5

Regulation of Motors

Although a large number of KLPs have been characterized from diverse organisms (www.blocks.fhcrc.org/kinesin/), the mechanisms that regulate the activity/function of MT-dependent motors is limited both in plants and animals (Reddy, 2001b, Reilein et al., 2001). 18.5.1

Calcium/Calmodulin

Calcium is a key messenger in transducing many hormonal and environmental signals in plants (Reddy, 2001a). Calmodulin, a ubiquitous calcium-binding protein in eukaryotes, is highly conserved in eukaryotes and regulates many diverse cellular functions by modulating the activity of the proteins that interact with it. Upon binding calcium, calmodulin becomes activated via a conformational change and is able to regulate the activity of its target proteins (Reddy, 2001a). Cytoplasmic streaming of characean cells is regulated by the cytosolic calcium concentration (Nagai, 1993, Shimmen and Yokota, 1994, Williamson and Ashley, 1982). Elevated levels of calcium inhibit cytoplasmic streaming and the movement of organelles (Hayama et al., 1979, Kikuyama and Tazawa, 1982, Kohno and Shimmen, 1987, 1988a, 1988b, Takagi and Nagai, 1986, Tominaga et al., 1983, Williamson and Ashley, 1982). Movement of pollen tube organelles along actin cables is inhibited by calcium, suggesting that the activity of actin-based motors is sensitive to calcium levels. The mechanisms by which calcium regulates the activity of these motors is just beginning to emerge. All myosins in Arabidopsis possess two or more putative calmodulin-binding motifs, suggesting that myosins in plants are likely to be regulated by calmodulin in response to changes in intracellular calcium. Two myosin heavy chains (170 and 175 kDa) in plants associate with calmodulin, indicating that calmodulin serves as a light chain for these myosins. Yokota et al. (1999a,b) using highly purified plant myosins have provided the first evidence that calcium inhibits the myosin motor activity as well as myosin-activated ATPase activity in plants. The inhibition of myosin activity in the presence of calcium appears to be due to dissociation of calmodulin from the heavy chain since calcium inhibition of motor activity can be restored by exogenous addition of calmodulin. KCBP and recently discovered kinesin C are the only KLPs among the kinesin superfamily that bind calmodulin (Reddy et al., 1996b, Rogers et al., 1999). The binding of activated calmodulin to KCBP inhibits its interaction with MTs via the calmodulin-binding domain resulting in the inhibition of MT-dependent


ATPase activity and motility (Deavours et al., 1998, Narasimhulu et al., 1997, Narasimhulu and Reddy, 1998, Song et al., 1997). These studies strongly suggest that the binding of calmodulin to KCBP affects MT binding regions resulting in inhibition of KCBP binding to MTs and dissociation of the KCBP/MT complex. Structural studies with KCBP in free and calmodulin-bound forms are needed to verify these speculations. The calmodulin effect on KCBP suggests that activated calmodulin can act as a molecular switch to downregulate the activity of KCBP. Based on in vitro studies with KCBP it is reasonable to speculate that spatial and temporal changes in free cytosolic calcium levels in response to signals are likely to regulate KCBP activity in the cell. 18.5.2

Protein Phosphorylation

The members of the BimC family have a conserved ‘BimC Box’ in which a threonine residue is phosphorylated by cdc2-kinase during cell division. Targeting of the KLP to the spindle is regulated by phosphorylation (Blangy et al., 1995, Sawin and Mitchison, 1995). TKRP125, all three members of Arabidopsis TKRP-like proteins (AtKRPa, b and c) and DcKRP120-1 and DcKRP120-2 that belong to BimC family have the conserved BimC box, suggesting that phosphorylation of these KLPs may be involved in targeting them to the right cellular location. Recently it was found that the tail region of KCBP also interacts with a protein kinase (KIPK, KCBP interacting protein kinase) in the yeast two-hybrid system (Day et al., 2000). The association of KIPK with KCBP suggests regulation of KCBP or KCBP-associated proteins by phosphorylation and/or KCBP is involved in targeting KIPK to its proper cellular location. The tobacco NPK1, a mitogen-activated protein kinase kinase kinase, NPK1, interacts with NACKs. This interaction results in activation and autophosphorylation of NPK1 and phosphorylation of NACK1 (Nishihama et al., 2002). Although the significance of NACK1 phosphorylation is not known, NPK1 phosphorylation targets it to the equatorial zone of the phragmoplasts (Nishihama et al., 2002).

18.6

Concluding Remarks

The complete sequencing of several phylogenetically diverse model organisms ranging from a unicellular eukaryote to highly complex multicellular animals and a plant has allowed comparative analysis of molecular motors in eukaryotes. Plants contain a large number of motors. About 0.4 % of the predicted genes in Arabidopsis encode molecular motors. The comparative analysis provides a framework for future functional studies with plant KLPs. The number of myosins in Arabidopsis is comparable to other multicellular organisms of similar genome size. However, plant myosins are unlike myosins from any other organism except algae. Nonplant myosins contain a variety of other known domains that are lacking in plants.

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References

Furthermore, Arabidopsis has a surprisingly large number of KLPs among the completed eukaryotic genomes. The role of molecular motors in many plant cellular processes is beginning to be unraveled. Although the functions of most of the plant motors remain to be determined, phylogenetic analysis and known domains in these proteins are providing clues to their function which can be tested empirically. Many Arabidopsis KLPs do not fall into any known subfamilies of KLPs and several Arabidopsis KLPs are not present in other organisms and are likely to represent new plant-specific subfamilies. Analysis of expression of each motor and cellular localization, using reporter fusions coupled with studies using the loss-of-function and gain-of-function mutants, should help to elucidate the functions of these motors. Identification of the proteins that are associated with motors is also important to our understanding of their roles in plants. Knockout mutants for almost all Arabidopsis motors are already available from SIGnAL (http://signal.salk.edu/) or TMRI (http://www.tmri.org). Because of the large number of motors and likely functional redundancy or overlap with other motors it will be necessary to create not only single mutants but also double or triple mutants. Analysis of the function and regulation of plant motors will be an exciting area of research in plant cell biology and is bound to produce many surprises.

Acknowledgments

I thank members of my laboratory and all my collaborators for their contributions. I wish to thank Irene Day for preparing the figures and critically reading the manuscript. Research on motors in my laboratory is supported by grants from the National Science Foundation (MCB-9630782 and MCB-0079938).

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Part 4 Motors in Disease

19 Myosin Myopathies John P. Konhilas and Leslie A. Leinwand

19.1

Introduction: Inherited Myosin Myopathy

Muscle contraction is ultimately dependent on the coordinated interaction of thick and thin filaments contained within each sarcomere. Myosin is the central component of the muscle thick filament. The myosin head, or cross-bridge, extends outward from the thick filament backbone and contains the catalytic site and the actinbinding domain. The binding to actin and subsequent ATP-dependent conformational change in cross-bridge orientation forces the interdigitated thick and thin filaments to slide past one another and is responsible for generating the power stroke and transmitting the force necessary for muscle contraction. Over the past decade, numerous mutations in the myosin molecule have been linked to muscle myopathies. Myosin mutations are most common in the heart giving rise to either dilated or hypertrophic cardiomyopathies. The best characterized of these inherited diseases is Familial Hypertrophic Cardiomyopathy (FHC). FHC is an autosomal dominant disease characterized by heterogeneous anatomic and histologic features including left ventricular hypertrophy, myofibrillar disarray, and increased interstitial collagen. Patients suffering from FHC have a variable clinical outcome usually associated with contractile dysfunction that can range from being benign and relatively symptom free to sudden death. Although mutations in many sarcomeric components can lead to FHC, the majority are missense mutations in the b-myosin heavy chain (MyHC) genes (Seidman and Seidman, 2001). To date, at least 60 mutations in MyHC have been described and are associated with the clinical manifestation of FHC in humans. Although the precise mechanisms responsible for the development of myosin myopathy remain unclear, it most certainly involves alterations in force production as a result of these mutations. In support of this idea, analysis of the crystallographic structure of the myosin head reveals that most, but not all mutations, including the skeletal myosin mutation (Darin et al., 1998), map to the head and neck region of the myosin molecule in close proximity to the functional domains of the motor (see Fig. 19.1; Rayment et al., 1993, 1995). The locations of these mu-

474

19.1 Introduction: Inherited Myosin Myopathy

Figure 1.

A

ATP binding pocket

Myosin S1 ELC

actin binding domain

RLC

B S2

LMM

S1

C

866

1 NH2

S1 globular head

1938

S2 neck/hinge

Myosin molecule structure and location of FHC mutations. (A) Crystal structure of chicken skeletal S1 showing the actin-binding domain, ATP binding pocket, and placement of the essential (ELC) and regulatory light chains (RLC). (B) Schematic of a single myosin mole-

LMM

COOH

myosin rod

Figure 19.1

cule. The locations of some of the FHC mutations are indicated by the inverted arrows (b). (C) Linear representation of the full length rat MHC with the relative location of each subunit.

tations suggest an interference with either the catalytic activity or actin myosin interaction leading to an alteration in the generation of power necessary for muscle contraction. From this model of pathogenesis, however, it is difficult to reconcile two distinct phenotypes such as the development of hypertrophic cardiomyopathy (HCM) or dilated cardiomyopathy (DCM) from mutations holding similar positions in the myosin head (Kamisago et al., 2000, Seidman and Seidman, 2001). In addition, this model does not readily explain FHC induced by mutations in the myosin rod (Blair et al., 2002). It seems logical to conclude that mechanism by which these rod mutations initiate the FHC disease process appear to involve aspects of myosin function independent of myosin motor activity. Much of our understanding of how mutations in the sarcomere can result in a condition such as FHC, DCM or skeletal myopathy has emerged from a wide range of approaches including analysis of the enzymatic and motility properties of mutated myosin molecules and the phenotypic and genetic characterization of transgenic mice expressing these mutant alleles. In this chapter, we will examine the functional characteristics of specific mutant myosins and transgenic models harboring these same mutations. We will further describe the molecular basis of FHC with a discussion of mutations in myosin-interacting proteins. Finally, although the majority of this discussion focuses on cardiac-based myopathies because of the extensive analysis of this disease, we also discuss skeletal muscle myopathies resulting from mutations in myosin and myosin-interacting proteins.

19 Myosin Myopathies

19.2

Cardiac Myosin Heavy Chains 19.2.1

MyHC Structure and Function

Striated muscle MyHC isoforms exhibit striking sequence homology. Thus, in considering the mechanisms whereby mutations in myosin cause myopathy, it is important to review what is known about the structure and functional domains of the cardiac myosins. Conventional myosin or myosin II is the main component of the muscle thick filament and accounts for 30 % of the total myofibrillar proteins. Myosin is a hexameric molecule consisting of two heavy chains and two non-identical light chains (MLC). The myosin rod is an a-helical coiled coil, termed light meromyosin (LMM), arranged in parallel along the axis of the thick filament (Fig. 19.1). It is linked via a hinge region termed subfragment-2 (S2) to a globular head, or subfragment-1 (S1). Myosin S1 is at the N-terminus of the heavy chain and contains the catalytic site and the actin binding domain. Whereas the S1 subunit of myosin hydrolyzes ATP and can bind actin, LMM mediates filament assembly (McLachlan and Karn, 1982) and provides binding sites for myosin-associated proteins such as myosin binding protein C and titin (Okagaki et al., 1993, Soteriou et al., 1993). In the mammalian heart, two isoforms (a-MyHC and b-MyHC) with 93 % amino acid identity are expressed (McNally et al., 1989). Three distinct myosins (V1, V2, and V3) have been described in the adult heart. V1 and V3 are homodimers of a-MyHC and b-MyHC, respectively. V2 myosin is the a/b heterodimer. In mammals, the relative proportion of each polypeptide depends on the species, developmental stage and physiological or pathophysiological status of the myocardium. Whereas b-MyHC predominates in the human heart, small rodents express 90 100 % a-MyHC in the adult myocardium. Intermediate between the rodent and human myocardium, the adult rabbit heart is comprised of approximately 70 % b-MyHC (VanBuren et al., 1995). In all mammals, b-MyHC is the dominant isoform during the embryonic stages. However, a-MyHC expression is essential during mouse development as demonstrated by the lethality of the a-MyHC null mutation (Jones et al., 1996). Around birth in small rodents, there is shift to the a-MyHC with complete silencing of b-MyHC expression (Lyons et al., 1990). Shifts of the cardiac MyHC isoforms have been demonstrated in response to hormonal changes, aging, exercise and pathology. For example, hypothyroidism induces a shift to b-MyHC in both rabbits and rats (de Tombe and ter Keurs, 1991, VanBuren et al., 1995). Similarly, pressure overload brought about by aortic banding, myocardial infarction, or other pathological stressors significantly increases b-MyHC content in the myocardium (Nadal-Ginard and Mahdavi, 1989). Transgenic mouse hearts with mutations in sarcomeric proteins demonstrate a MyHC isoform switch to predominantly b-MyHC (Freeman et al., 2001a, Tardiff et al., 1999). MyHC isoform shifts were thought to be of no significance in humans since the normal heart was previously believed to be composed entirely of b-MyHC

475

476

19.2 Cardiac Myosin Heavy Chains

(Bouvagnet et al., 1989, Schiaffino et al., 1984). However, evidence has been presented indicating that a-MyHC comprises Z 10 % of total MyHC in non-failing hearts and becomes virtually undetectable in the failing myocardium (Lowes et al., 1997, Miyata et al., 2000). Interestingly, in contrast to the induction of b-MyHC by pathologic stimulation, exercise induces expression of a-MyHC in rats (Schaible and Scheuer, 1985). Despite significant structural and sequence homology, the two isoforms exhibit distinct functional characteristics. Early studies illustrated that myosin composed of a-MyHC exhibits two to three times the actin-activated and calcium-stimulated myosin ATPase activity when compared to myosin composed of primarily b-MyHC (Harris et al., 1994, Litten et al., 1982, VanBuren et al., 1995). Moreover, actin filament velocities of each isoform in an in vitro motility assay show parallel differences (Harris et al., 1994, VanBuren et al., 1995). These functional differences are achieved independent of alterations in the Ca2 sensitivity of tension and maximum tension-generating capability (Fitzsimons et al., 1998). The results with isolated proteins are consistent with observations that intact cardiac muscle preparations with a-MyHC have increased unloaded shortening velocities compared to b-MyHC in both papillary muscles from thyroxine treated (hyperthyroid) rabbits (Pagani and Julian, 1984) and intact cardiac trabeculae from PTU-treated (hypothyroid) rats (de Tombe and ter Keurs, 1991). Skinned cardiac myocytes expressing a-MyHC have augmented loaded shortening and power output when compared to those composed of primarily b-MyHC (Herron et al., 2001). Consistent with these observations, the rates of submaximal and maximal tension development are lower (60 %) coupled with an increase in the half-time for relaxation in skinned myocardium expressing b-MyHC from that of controls (Fitzsimons et al., 1998). Furthermore, the isozyme characteristics are not affected by b-adrenergic treatment suggesting that these properties are due to the intrinsic differences in cross-bridge kinetics between a-MyHC and b-MyHC (de Tombe and ter Keurs, 1991, Fitzsimons et al., 1998). The increased kinetics of a-MyHC compared to b-MyHC are not, however, without an energetic cost. Warshaw and colleagues illustrated in the in vitro motility assay that b-MyHC has greater average isometric force-generating capacity than a-MyHC (VanBuren et al., 1995). Coupled with a slowed actin filament velocity and reduced myosin ATPase, b-MyHC undergoes a slower transition from the force-generating to the non-force-generating state once strongly bound. Thus, the prolonged detachment rate of b-MyHC suggests a greater economy of force production, a similar conclusion reached by the same group using an in vitro motility assay (Harris et al., 1994). Consistent with this idea, Holubarsch et al. (1985) showed that left ventricular papillary muscles from PTU-treated rats produce less heat, relative to controls, when contractile heat was normalized to force (Holubarsch et al., 1985). A direct measurement of force-dependent ATP consumption in skinned rat cardiac preparations yielded similar results as demonstrated by a linear decrease in tension cost with increasing b-MyHC isoform content relative to a-MyHC (Rundell et al., 2002).


Taken together, these mechanical data predict that myocardium composed primarily of a-MyHC would develop tension, or pressure, at a greater rate (due to enhanced myosin ATPase activity and increased power output) than myocardium expressing predominantly b-MyHC. Furthermore, it could be argued that the extended half-time for relaxation in myocardial preparations of b-MyHC (Fitzsimons et al., 1998) should similarly slow the rate of relaxation in an intact ventricle. Indeed, this has been demonstrated in several studies in which hypothyroidism in the mouse results in a decrease in the time derivative of both pressure development ( dP/dt) and relaxation (– dP/dt) in isolated work-performing hearts (Brittsan et al., 1999, Kiss et al., 1998) and intact closed-chest preparations (Lorenz and Robbins, 1997). Additional effects independent of MyHC expression confound the experimental models of hyperthyroidism and, similarly, hypothyroidism. To address the specific role of b-MyHC in the intact myocardium without the accompanying hormonal effects of thyroid hormone manipulation, a transgenic murine model was generated that expresses 12 % b-MyHC in adult ventricular tissue (Tardiff et al., 2000). The depression in myofibrillar ATPase activity determined from transgenic hearts correlates with the significant decrease in systolic function assessed by a Langendorff preparation (Tardiff et al., 2000). Thus, it appears that MyHC isoform expression in the intact heart does have functional consequences even in the absence of additional alterations. An important observation from these studies is that expression of b-MyHC in the heart does not result in an overt cardiac pathology (Tardiff et al., 2000). Therefore, the implication is that the isoform switch to b-MyHC, in and of itself, does not contribute to the pathogenesis of cardiomyopathy in general and that HCM is not caused purely by a reduction in the ATPase activity of the myosin molecule. Instead, in light of an enhanced economy of force production associated with b-MyHC, an increase in b-MyHC content is, at least initially, an adaptive response to the energetic demands in the context of a compromised myocardium. 19.2.2

Cardiac Muscle Regulation and Disease

Cardiac function is ultimately dependent on the ability of the myofilaments to generate force, which, in turn, is dependent on the amount of cytosolic Ca2, phosphorylation state, and isoform content of the cardiac myocyte (de Tombe and Solaro, 2000, Solaro and Rarick, 1998). Thus, in an attempt to understand the development of myosin myopathies resulting from mutations, it is important to elucidate the relationship between myosin function and these additional factors involved in the regulation of muscle contraction. As detailed above, the mechanical properties of the cardiac heavy chain isoforms have a direct impact on cardiac function independent of other signals. On the other hand, factors such as thyroid hormone are involved in both the regulation of MyHC isoform content and cardiac function (Lorenz and Robbins, 1997).

477

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19.3 Cardiac MyHC Myopathy

In general, an increase in physiologic load forces heart cells to adapt and match the intensity and dynamics of their mechanical activity to the prevailing hemodynamic demands. The beat-by-beat regulation of cardiac output is accomplished through intrinsic and extrinsic feedback regulation that acts in a relatively instantaneous manner. Changes in end-diastolic volume can alter the contractile state of the ventricle through an intrinsic mechanism known as the Frank Starling law of the heart (Starling, 1918). Myocardial function can also be tuned to the hemodynamic load via neurohumoral factors such as adrenergic stimulation and subsequent activation of intracellular signaling pathways (Solaro and Rarick, 1998). However, the inability of the myocyte to match cardiac output to the physiological demand that occurs with a chronic hemodynamic load such as hypertension or altered contractile function such as mutant sarcomeric proteins initiates a remodeling process. Central to this remodeling process is myocyte enlargement (hypertrophy) since the adult cardiac myocyte is terminally differentiated and unable to proliferate. Clearly, the development of FHC is dependent on the coordinated integration of the multiple signals to counteract the contractile dysfunction. Very often, however, these signals lead to maladaptations that can exacerbate the myocardial dysfunction (de Tombe and Solaro, 2000, Vikstrom et al., 1998). Ultimately, the heart is unable to respond to changes in hemodynamic load resulting in the clinical syndrome of congestive heart failure (CHF) (Francis and Cohn, 1990). Clearly, hypertrophy is an adaptive, compensatory mechanism which attempts to match cardiac output to the increase in physiological demand. Yet, hypertrophy resulting from pathological stimuli such as a mutant protein or hypertension is initially adaptive but can become maladaptive and is a leading predictor of CHF. Although it is unclear how myofilament dysfunction resulting from these mutant proteins translates into the complex phenotypes associated with FHC, it is most certainly related to a functional impairment of the contractile apparatus and subsequent integration of the hypertrophic signals arising from this dysfunction as discussed below.

19.3

Cardiac MyHC Myopathy 19.3.1

Functional Characterization of MyHC Motor Domain Mutations

To date, the vast majority of mutations in the b-MyHC described in human disease are located in the motor domain of the myosin molecule, i. e. the actin-binding region, the nucleotide binding pocket and the hinge region (Rayment et al., 1995). The functional impact of several single amino acid substitutions has been assessed in assays such as actin-activated ATPase and in vitro motility. In most instances, the severity of the disease correlates with the functional defects in motor activity as measured by these assays. For example, patients with an arginine substituted by a glutamine at residue 403 (R403Q) generally have a poor clinical prognosis (Geis-

19 Myosin Myopathies Table 19.1

In vitro functional analysis of myosin mutations. ATPase

Study

Source

T124I

Human, HMM/S1

q

Cuda et al. (1997)

Y162C

Human

qq

Cuda et al. (1997)

R249Q

HMM/S1

q

Sata and Ikebe (1996), Cuda et al. (1997)

G256E

Human, HMM/S1

m, q

m

Sweeney et al. (1994), Lankford et al. (1995), Cuda et al. (1997), Palmiter et al. (2000)

R403Q

Human, HMM/S1, q Mouse

qq, o

q, o

Sweeney et al. (1994), Lankford et al. (1995), Sata and Ikebe (1996), Cuda et al. (1997), Roopnarine and Leinwand (1998), Blanchard et al. (1999), Palmiter et al. (2000)

R453C

HMM/S1

q

q

Sata and Ikebe (1996)

V606M

Human, HMM/S1

q, m

q

Cuda et al. (1997), Roopnarine and Leinwand (1998)

G741R

Human

q

R870H

Human

q

Cuda et al. (1997)

L908V

Human

qq, o

Cuda et al. (1997), Palmiter et al. (2000)

q

filament velocity

Ca2 sensitivity

Mutation

q

Lankford et al. (1995)

HMM, heavy meromyosin; S1, myosin (S1) head domain.

terfer-Lowrance et al., 1990). Accordingly, actin-activated ATPase activity of the R403Q mutation expressed in baculovirus or mammalian cells is greatly reduced (Roopnarine and Leinwand, 1998, Sata and Ikebe, 1996, Sweeney et al., 1994). In addition, this mutation results in a velocity as measured by the in vitro motility assay that is nearly 5-fold less than normal myosin (Sweeney et al., 1994). A reduction in this parameter was also found in myosin purified from cardiac and skeletal muscle biopsies with this same mutation as well as other myosin mutations (T124I, Y162C, G256E, R870H, and L908V) (Cuda et al., 1997, Lankford et al., 1995). Moreover, skinned fibers from patients with the R403Q mutation exhibit lowered force/stiffness ratio and depressed velocity of shortening and power output (Lankford et al., 1995). A summary of the mechanical and/or kinetic properties of additional mutations is presented in Table 19.1.

479

480

19.3 Cardiac MyHC Myopathy

The biochemical properties of additional myosin mutations located in the myosin S1 were also studied and found to correspond to the clinical prognosis (Roopnarine and Leinwand, 1998, Sata and Ikebe, 1996). The V606M mutation lies in the actin-binding domain and the R249Q and R453C mutations are located at the base of the ATP-binding pocket (Rayment et al., 1995). The V606M mutation is typically associated with a benign form of FHC (Watkins et al., 1992) although an unfavorable prognosis has been reported in several families with the V606M (Fananapazir and Epstein, 1994, Havndrup et al., 2001). Patients with the R453C mutation present with a severe form of FHC. In contrast, the R249Q mutation has a moderate phenotype with respect to incidence and age of sudden death. These mutations (R249Q, R453C, and V606M) result in a reduction in sliding velocities of actin filaments (Sata and Ikebe, 1996). In the ATPase assay, the V606M mutation shows the mildest decrease and R453C mutation has the most significant decrease of the three (Roopnarine and Leinwand, 1998). In contrast, Sata and Ikebe (1996) found no difference in the actin-activated ATPase between the V606M mutation and wild-type controls. However, these functional differences may be due to the use of human b-MyHC S1/S2 fragment (Sata and Ikebe, 1996) versus myosin S1 of the rat a-MyHC (Roopnarine and Leinwand, 1998). Thus, in the majority of cases, the degree of impairment in the mutant myosin motors correlates with clinical prognosis. However, the phenotypes of certain mutations depend largely on additional genetic and non-genetic factors. Support of this is found in a recent study demonstrating that a polymorphic modifier gene can affect the hypertrophic response in two inbred mouse strains expressing a mutant allele (R403Q; Semsarian et al., 2001). More apparent genetic factors such as ethnic background also contribute to the development of the disease. For example, Korean kindred with the R403Q mutation had predominantly left ventricular outflow obstruction and no sudden cardiac death while Caucasian pedigrees had non-obstructive hypertrophic cardiomyopathy with a high incidence of sudden cardiac death (Fananapazir and Epstein, 1994). Thus, the mechanical factors only partially predict the penetrative nature of this disease, a notion consistent with the clinical heterogeneity. 19.3.2

Transgenic Models of Myosin-based FHC

The pathogenesis of myosin-based FHC has been studied using transgenic murine models with the missense mutation (R403Q) in the mouse a-MyHC corresponding to the human b-MyHC mutation (Geisterfer-Lowrance et al., 1996, Vikstrom et al., 1996). In addition, a transgenic rabbit model of the R403Q mutation in the bMyHC gene has been described (Marian et al., 1999). The murine model described by Geisterfer-Lowrance et al. (1996) placed the mutation into the endogenous locus while that described by Vikstrom et al. (1996) used a transgenic approach. The latter model contains an additional deletion of amino acids 468 527 bridged by nine non-myosin amino acids (Vikstrom et al., 1996). Although each mouse model recapitulates aspects of the myocardial pathology associated with FHC, they exhibit distinct phenotypes.


In both mouse models, cardiac histopathology characterized by myocellular disarray and interstitial fibrosis was evident by 3 4 months of age. As early as 5 weeks of age, the mice described by Geisterfer-Lowrance et al. (1996) had altered LV diastolic kinetics with delayed pressure relaxation and chamber filling accompanied by decreased cardiac output (Geisterfer-Lowrance et al., 1996, McConnell et al., 2001). They also observed enlarged atria in the absence of ventricular hypertrophy (Geisterfer-Lowrance et al., 1996). The diminished actin-activated ATPase activity and enhanced dynamic stiffness in cardiac muscle strips from these hearts is consistent with diastolic dysfunction (Blanchard et al., 1999). Interestingly, in this particular model the cardiac contractile dysfunction at 5 weeks of age preceded any histopathology or morphological abnormalities. Lending further support for the mechanical basis of this disease was the observation that the decay of the intracellular Ca2 transient was markedly delayed in isolated myocytes from the transgenic mice independent of alterations in the amount of sarcoplasmic reticulum Ca2 ATPase (Kim et al., 1999). This suggests that altered mechanical activity of the sarcomere not only leads to aberrant ventricular function but also induces the cellular pathology associated with FHC. In contrast to the mice described above, the transgenic mice showed significant left ventricular hypertrophy at 4 months of age (Freeman et al., 2001a, Vikstrom et al., 1996). These mice exhibited diastolic dysfunction at both 4 months and 8 10 months of age with either normal or increased systolic function in females. Unlike the female mice, hemodynamic function worsened with age in males (Freeman et al., 2001a, Olsson et al., 2001). At an older age (8 months), male mice developed progressive left ventricular dilation similar to cases reported in humans (ten Cate and Roelandt, 1979, Vikstrom et al., 1996). Interestingly, augmentation of catecholamine reactivity in the mutant myocardium by high level over-expression of the b2 -adrenergic receptor hastened the progression to cardiac failure. In contrast, the male CHF phenotype was prevented by genetic manipulations that blunted the adrenergic signaling pathway (Freeman et al., 2001b). Thus, the underlying cause of the myocellular pathology can be attributed to the mutated protein, yet it appears that the progression of the disease is influenced by additional factors, such as gender, associated with the hypertrophic response. Recently, a transgenic rabbit model of FHC was described (Marian et al., 1999). The hearts in these rabbits carry the human b-MyHC with the R403Q mutation. A significant increase in ventricular septal wall thickness and left ventricular mass was seen in the transgenic hearts compared to hearts expressing the wild-type human transgene. Despite normal ventricular dimensions and systolic function (measured by echocardiography), transgenic mutant hearts displayed significant myocyte disarray and increased interstitial collagen. Premature death was also more common in mutant than in wild-type transgenic animals. The interesting feature of the transgenic rabbit model is that, similar to humans, rabbits express predominantly b-MyHC. Thus, trangenic rabbits may prove a more ideal model over existing transgenic mouse models to study the pathogenesis of human FHC. In one of the transgenic mouse lines with minimal expression of the mutant transgene (0.6 2.5 %), the appearance of histopathology preceded the onset of ven-

481

482

19.4 MyHC Interacting Proteins and FHC

tricular hypertrophy suggesting that the myocellular degeneration triggers the hypertrophy (Vikstrom et al., 1996). Thus, considering each model, the time course of events is such that, sarcomeric dysfunction results in altered ventricular function triggering remodeling that can also feedback and contribute to this dysfunction. In order to compensate for decreased cardiac output, the hypertrophic response is initiated starting a vicious cycle of positive feedback into this aberrant, maladaptive process. Although transgenic models provide a direct link between altered contractile function and the development of cardiomyopathies, there is no complete model describing the integration of the multiple specific pathways induced by the myofilament dysfunction and leading to the pathologies associated with FHC. Given the impact of additional factors on the progression of this disease (Freeman et al., 2001b, McConnell et al., 2001), it seems obvious that development of FHC depends on the coordinated interaction of multiple signaling pathways. For example, the identification of two mutations in the myosin rod (A1379T and S1776G) that lead to hypertrophic cardiomyopathy cannot be readily explained by a deficit in force transmission (Blair et al., 2002). These missense mutations may disrupt the coiled-coil structure and/or thick filament assembly that may lead to ultrastructural disorder, despite the presumably normal force-generating capability of the myosin head (Blair et al., 2002). Yet, it is not known how a potential impairment of filament structure or assembly translates into the complications associated with FHC. Potential paradigms in which to identify and distinguish these pathways may reside in the observation that there are marked differences between female and male animals (Geisterfer-Lowrance et al., 1996, Olsson et al., 2001, Vikstrom et al., 1996). Moreover, the identification of myosin mutations (S532P and F764L) in the actin-binding domain and hinge region that result in a dilated cardiomyopathy unaccompanied by hypertrophy, also points to a distinct signaling response (Kamisago et al., 2000). Thus, it appears that the performance of the contractile machinery not only initiates this disease process but is central to the integration of the signals that perpetuate and promote this response.

19.4

MyHC Interacting Proteins and FHC 19.4.1

The Essential and Regulatory Light chains

The structure of the myosin rod (LMM) allows the specific myosin myosin interactions necessary for thick filament assembly. In addition, other myosin-interacting proteins are necessary for normal muscle functioning. The carboxyl terminus of the S1 region forms a single a-helix. Structural and functional data suggest that this region acts as the lever arm, such that the catalytic domain of the myosin head swings relative to this neck domain (Uyeda et al., 1996). Thus, this motion accompanied with the strong acto myosin interaction during muscle activation


is responsible for the displacement between the thick and thin filament necessary for contraction. Each of the two light chains, the essential light chain (ELC) and regulatory light chain (RLC), wrap around this a-helix, encompassing the majority of this region, and may play a role in providing structural support for this segment of the myosin rod (Rayment et al., 1993). In addition to providing structural support for myosin, the light chains may modulate myofilament contractile properties. The RLC of myosin contains a divalent cation-binding site similar to other Ca2 binding ‘EF-hand’ proteins like troponin C and calmodulin (Collins, 1976). Removal of the light chains reduced the velocity of actin filament movement in an in vitro motility assay (Lowey et al., 1993) whereas only removal of the ELC reduced isometric force (VanBuren et al., 1994). Two adjacent serine residues in the N-terminal domain of the RLC are targeted by the Ca2/calmodulin-dependent myosin light chain kinase (MLCK). The phosphorylation of these sites by MLCK in smooth and non-muscle myosins plays an integral role in the activation of myosin ATPase and subsequent contractile strength (Bresnick, 1999). However, in striated muscle, the primary regulation of muscle contraction is not dependent on RLC phosphorylation. Nevertheless, phosphorylation of the RLC may act to modulate cross-bridge cycling kinetics in contracting muscle. In support of this, RLC phosphorylation results in an increase in Ca2 -sensitivity of tension development and the rate of force redevelopment in intact and skinned skeletal muscle fibers (Metzger and Moss, 1992, Sweeney and Stull, 1990, Sweeney et al., 1993, Szczesna et al., 2002). Szczesna et al. (2002) also demonstrated that the increase in Ca2 -sensitivity was accompanied by an increase in maximal force development and higher Ca2 -dependent ATPase activity in skinned muscle fibers reconstituted with phosphorylated RLC. However, because RLC phosphorylation significantly increased the rate constant for isometric force redevelopment (Metzger and Moss, 1992, Sweeney and Stull, 1990) without a concomitant increase in the rate of isometric force-dependent ATPase consumption (Sweeney and Stull, 1990), it has been proposed that RLC phosphorylation enhances the rate of transition from non-force-generating cross-bridges to force-generating states. The structural basis for these physiological findings may reside in an increase in the mobility of the myosin heads following RLC phosphorylation as demonstrated by disordering of the myosin head array (Levine et al., 1998, Yang et al., 1998b). Although these studies were performed using skeletal muscle preparations, experimental evidence has suggested an important role of RLC phosphorylation in the stretch activation response of cardiac papillary muscles (Vemuri et al., 1999). The high level of RLC phosphorylation in the epicardium will increase tension and decrease the stretch-activation response while resulting in the opposite effect in the endocardium, allowing different muscle mechanics across the ventricular wall (Davis et al., 2001, Vemuri et al., 1999).The stretch-activation response coupled with the spatial gradient of phosphorylated RLC may facilitate the torsional contraction observed in the intact ventricle (Davis et al., 2001). Moreover, the presence of mutations within the a-helix (the light chain-binding domain) and in both ELC and RLC underlie distinct phenotypes of FHC.

483

484


19.4.2

Myosin Light Chain-based FHC

The first identified mutations in the RLC (A13T, E22K, and P94R) and the ELC (M149V) were associated with a form of FHC characterized by mid-ventricular cavity obstruction due to papillary muscle hypertrophy (Poetter et al., 1996). However, a more typical phenotype of hypertrophic cardiomyopathy dominated by increased left ventricular mass and abnormal contractile function was subsequently found in patients with the RLC mutations F18L and R58Q (Flavigny et al., 1998). Interestingly, these mutations were not associated with cases of sudden death (Flavigny et al., 1998, Poetter et al., 1996). The mechanism by which the light chain mutations produce the FHC phenotype is unclear but most likely involves an interference of the interaction between the light chains and myosin thereby affecting force generation. Skinned fibers with the E22K mutation had an increased Ca2 sensitivity of tension and enhanced disordering of the myosin heads (Levine et al., 1998). Furthermore, RLC mutations (A13T, F18L, E22K, R58Q, and P95A) reduced Ca2 -binding with the largest reduction observed in the F18L mutation and complete obliteration in the R58Q mutation (Szczesna et al., 2001). Moreover, phosphorylation of the RLC altered the Ca2 affinity to varying degrees in wild-type RLC and the RLC mutants (Szczesna et al., 2001). In the context of the intact sarcomere, a transgenic mouse in which the human ELC mutation (M149V) was inserted into the murine genome recapitulated FHC (Vemuri et al., 1999). However, if the homologous mutation was made in the context of the murine gene locus, there was histopathology consistent with FHC but without the characteristic hypertrophic response (Sanbe et al., 2000). Instead, the hearts from these mice were smaller as measured by heart weight-to-body weight ratios. This occurred in conjunction with an increased myofibrillar ATPase activity and the Ca2 -sensitivity of tension and a concomitant decrease in maximum power output (Sanbe et al., 2000). Sanbe et al. (2000) also reported that unlike patients with the E22K RLC mutation as described above, transgenic mice carrying the same mutation exhibited no hypertrophy and no pathology. To determine the functional impact of myosin mutations that disrupt the interaction of the light chains and myosin, a transgenic mouse was generated with a deletion in the light chain-binding domain of the a-cardiac MyHC (Welikson et al., 1999). Skinned myocytes and multicellular preparations from transgenic hearts exhibited decreased Ca2 -sensitivity of tension and rates of relaxation (Welikson et al., 1999). These hearts displayed an increase in anterior wall thickness whicht translated into an overall increase in heart mass. Histopathological features were consistent with FHC including small vessel coronary disease, myocellular disarray, and interstitial fibrosis (Welikson et al., 1999). In addition, these mice had a significant valvular pathology seen in a subset of patients with FHC (Welikson et al., 1999). Similar to the MyHC mutations, the in vitro functional characteristics of light chain mutations do not necessarily predict the myocardial phenotype harboring the corresponding mutations. Moreover, light chain mutations in transgenic mod-


els do not always recapitulate human FHC indicative of the phenotypic diversity seen in the clinical realm. Again, the implication is that the pathogenesis of FHC involves the integrative response to multiple signaling pathways and modifying factors. 19.4.3

Myosin Binding Protein C-Based FHC

Another major myosin accessory protein present in nearly all vertebrate striated muscle is myosin-binding protein C (MyBP-C). MyBP-C is relatively large (Z 130 kDa) and belongs to the intracellular immunoglobulin superfamily. It is composed of repeated Ig and fibronectin domains (Einheber and Fischman, 1990). The C-terminal region of MyBP-C also interacts with the LMM and with titin, presumably anchoring the protein to the thick filament complex (Alyonycheva et al., 1997, Okagaki et al., 1993). The N-terminal portion binds to the myosin S2 segment in close proximity to the lever arm of the myosin head (Gruen and Gautel, 1999). Despite an undefined, yet necessary role of MyBP-C in thick filament assembly (Koretz, 1979, Maw and Rowe, 1986, Winegrad, 1999), a role for MyBP-C as a physiological regulator of cardiac contractility is emerging in the literature (Flavigny et al., 1999, Kunst et al., 2000, Winegrad, 2000, Yang et al., 1999). The influence of MyBP-C on actomyosin ATPase is dependent on ionic strength, the isoform of myosin, and the presence of the light chains (Hartzell, 1985, Moos and Feng, 1980, Winegrad, 1999). Another important issue apart from the effect on actomyosin ATPase is the physiological role of MyBP-C phosphorylation. Within the N-terminal region that interacts with the myosin S2 segment are phosphorylation sites targeted by cAMPdependent protein kinase (PKA) and Ca2/calmodulin-dependent kinase (CaMK) (Garvey et al., 1988, Hartzell and Glass, 1984). Introduction of non-phosphorylated recombinant cardiac MyBP-C into skinned muscle fiber preparations reduced Ca2 activated maximal force and increased the Ca2 -sensitivity of force development compared to MyBP-C in the phosphorylated state (Kunst et al., 2000). However, recent studies demonstrated that PKA treatment of skinned myocardium from transgenic mice that undergo PKA-dependent phosphorylation of MyBP-C in the absence of TnI phosphorylation had no effect on the Ca2 -sensitivity of force development or Ca2 -activated maximal force (Fentzke et al., 1999, Kentish et al., 2001). Despite an undetermined functional role, phosphorylation of MyBP-C has also been shown to have a structural effect on both the myosin heads (Weisberg and Winegrad, 1998) and myofilament lattice structure (Konhilas et al., 2000). FHC has also been linked to mutations in cardiac MyBP-C (Niimura et al., 1998). In general, patients with mutations in MyBP-C have later onset, lower penetration, a better prognosis and longer life expectancies compared to most MyHC mutations and very often remain asymptomatic. A knock-in mouse model that carries a shortened MyBP-C at the N-terminus lacking the mutant site identified in FHC (Niimura et al., 1998) shows no pathology despite an increase in Ca2 -sensitivity of force (Witt et al., 2001). The majority of these mutations, however, result in trunca-

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486


tions of the MyBP-C molecule at the C-terminus (Niimura et al., 1998) which is the region that interacts with the myosin rod and with titin (Alyonycheva et al., 1997, Freiburg and Gautel, 1996). Given the interactions of the C-terminus with myosin and titin, the mutated complexes could theoretically disrupt sarcomeric structure and interfere with normal contractile function of the heart potentially leading to a hypertrophic response. This idea is supported by transgenic models in which a truncated MyBP-C expressed in the heart elicited aspects of FHC (McConnell et al., 1999, Yang et al., 1998a, 1999). Similar to the human form of MyBP-C-based FHC, the pathology associated with these mutations was quite subtle. Despite an increase in the Ca2 sensitivity of force and a reduction in the maximum power output in both permeabilized cardiac fibers or myocytes, hypertrophy was not evident in the hearts of transgenic mice expressing a mutant MyBP-C that lacks both the myosin- and titin-binding domains (Yang et al., 1998a). A similar mechanical effect accompanied by cardiac hypertrophy was found in hearts that expressed a mutant MyBPC that lacks the myosin-binding domain but retains the titin-binding domain (Yang et al., 1999). In both models, the mutant peptide was incorporated into the sarcomere, albeit in an aberrant fashion. Interestingly, there were no changes in myocyte and interstitial morphology yet striking structural abnormalities such as deficits in sarcomere organization and arrangement, were exhibited (Yang et al., 1998a, 1999). When the mutant protein was expressed in a homozygous fashion, there was profound pathology characterized by a dilated cardiomyopathy at birth that progressed into significant cardiac hypertrophy by 8 12 weeks of age (McConnell et al., 1999). Despite well-organized sarcomeres, the hearts had depressed contractile function and a prominent histopathology including myocyte hypertrophy, myofibrillar disarray, and fibrosis consistent with the hypertrophic response (McConnell et al., 1999). Similarly, inactivation of the MyBP-C gene and thus elimination from the cardiac sarcomere resulted in significant hypertrophy, depressed contractile function, and a characteristic hypertrophic histopathology (Harris et al., 2002). In the MyBP-C null hearts, sarcomere striation patterns were visible although the Z-lines appeared out of register (Harris et al., 2002). Although the functional impact of MyBP-C-based mutations cannot be ignored, based on these studies, the notion of MyBP-C as an essential component in sarcomere assembly must be modified, at a minimum, to a modulatory or regulatory role. In support of this, FHCassociated mutations in the MyBP-C-binding domain of the MyHC S2 segment drastically reduced MyBP-binding without affecting the coiled-coil structure of myosin or myofibrillar integrity (Gruen and Gautel, 1999). Again, this suggests that the contractile dysfunction associated with MyBP-C-based FHC is associated with a functional impairment rather than an assembly deficit.


19.4.4

Titin-based Familial Hypertrophic Cardiomyopathy

In striated muscle, titin (also known as connectin) spans from the Z-line to the Mline of the sarcomere and binds tightly with the thick filament (Obermann et al., 1997, Soteriou et al., 1993). Titin has been targeted as an essential contributor to the passive elasticity of striated muscle, and the variable extensibility of titin isoforms may explain, in part, the differences in passive stress between cardiac and skeletal muscle (Granzier and Irving, 1995, Linke et al., 1998, Trombitas et al., 2000, Wang et al., 1991). Although a multi-domain polypeptide, only the I-band region is functionally extensile (Granzier and Irving, 1995, Labeit et al., 1997, Linke et al., 1998). Within this region is a segment rich in proline (P), glutamate (E), valine (V), and lysine (K) residues (the so-called PEVK segment) that is serially flanked by immunoglobulin-like (Ig) domains (Labeit and Kolmerer, 1995). In addition, two differentially spliced extensile regions, the N2A/N2B elements that join the tandem Ig domains with the PEVK element, are included in this region and may explain the variation in passive stiffness of cardiac myocytes (Trombitas et al., 2000). During passive stretch, these regions do not stretch in parallel but instead tandem Ig and PEVK segments extend sequentially with the Ig domain associated with small stretch and low passive force while the PEVK domain associated with greater stretch and passive force increases (Granzier et al., 1996, Linke et al., 1998). The contribution of titin to passive tension arises from the extensibility properties of these specific domains. Likewise, titin acts as scaffolding for thick filament assembly as suggested by its interactions with several sarcomeric components. The carboxyl terminus of titin extends into the A-band and binds near the end of the LMM region of the myosin tail forming a stiff structure relative to the I-band segment (Houmeida et al., 1995, Soteriou et al., 1993). This region of titin contains domain patterns that reflect myosin head repeats, suggesting titin may have a role in determining thick filament structure and arrangement of the myosin heads (Houmeida et al., 1995). Consistent with this structural role, titin interacts with an M-band protein, myomesin, an association that is regulated by the phosphorylation state of a serine residue within this region of myomesin (Obermann et al., 1997). Also within the carboxy terminal region, titin binds to MyBP-C, a protein that has been implicated in the structural organization of the thick filament as mentioned above (Freiburg and Gautel, 1996, Yang et al., 1998a). Interestingly, titin can associate with actin and this interaction is limited to the Z-line. This presumably maintains high stiffness of the relaxed cardiac myofibril (Linke et al., 1997, Soteriou et al., 1993), suggesting both a structural and physiological role for titin. Within the Z-line, titin forms a ternary complex with actin and a-actinin, an important Z-line component (Sorimachi et al., 1997). Although the functional consequence of this association remains to be elucidated, specific titin mutations in this region linked to the development of FHC may provide a testable framework to study this interaction (Satoh et al., 1999). Recently, a mutation in titin was identified in patients with a typical hypertrophic cardiomyopathy who had no known

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mutations in other genes known to cause FHC (Satoh et al., 1999). This mutation (R740L) is within the binding domain of a-actinin in the Z-line region. The hypertrophic response in these patients may be empirically related to the observation that this titin mutation increased binding affinity to a-actinin (Satoh et al., 1999). However, Satoh et al. (2002) also mapped mutations to the Z-line binding region of the titin gene in patients presenting with a familial form of dilated cardiomyopathy (DCM). The two mutations (V54M and A743V) decreased the affinity of titin to a-actinin and another Z-line protein, T-cap/telethonin as measured by a yeast two-hybrid assay (Itoh-Satoh et al., 2002). Two additional mutations (Q4053ter and S4465D) found in FHC patients were localized to N2A extensible domains (Itoh-Satoh et al., 2002). Although the mechanical properties of the mutated titins have yet to be defined, it is likely that both the HCM and the DCM have arisen from a functional impairment in force transmission from Z-line to Z-line mediated through the titin molecule.

19.5

Myosin-based Myopathies in Skeletal Muscle

As indicated by the discussion above, the relationship between mutations in sarcomeric components of cardiac muscle and FHC are well defined. Mutations in sarcomeric proteins of the skeletal contractile apparatus can also lead to a severe myopathic phenotype. A majority of skeletal muscle myopathies are recessive although some actin and myosin mutations display a dominant inheritance (Darin et al., 1998, Nowak et al., 1999). There have also been a number of de novo mutations identified. Skeletal myopathies are generally characterized by structural abnormalities, hypotonia and severe muscle weakness that can result in early death (within the first year of life) usually due to respiratory failure. Also, skeletal muscle mutations typically lead to muscle degeneration rather than the myocellular hypertrophy which is usually observed in FHC (Askanas and Engel, 1998, Burton and Davies, 2002). These congenital myopathies typically display one of two distinct phenotypes: nemaline myopathy, which is identified by the appearance of intranuclear and/or sarcoplasmic rod structures, termed nemaline bodies, consisting of proteins from the Z-line and thin filament, and actin myopathy, which is distinguished from nemaline myopathies by the presence of large sub-sarcolemmal accumulations of thin filaments composed primarily of actin (Marston and Hodgkinson, 2001, Nowak et al., 1999). Whereas mutations in actin can lead to both actin and nemaline myopathies (Nowak et al., 1999), patients with mutations in tropomyosin (Laing et al., 1995), troponin T (Johnston et al., 2000), and nebulin (Pelin et al., 1999) exhibit a pure nemaline myopathy. Another group of inheritable skeletal myopathies are classified as hereditary inclusion-body myopathies (HIBM). In general, muscle fibers from patients with HIBM have morphological features dominated by rimmed vacuoles and filamentous inclusion bodies (Askanas and Engel, 1998). A novel mutation in the skeletal


MyHC-IIa gene has been identified in patients with HIBM and is inherited in an autosomal dominant fashion (Darin et al., 1998, Martinsson et al., 2000). The histopathological findings are heterogeneous depending on age, with adolescents having variability in fiber size, central nuclei and focal disorganization; older patients show progressive muscle weakness, dystrophic changes and inclusion bodies (Darin et al., 1998). This mutation is a missense mutation leading to an amino acid substitution at residue 706 (E to K) and is located in the highly conserved motor domain of the myosin head. The manifestation of the pathology strongly suggests a severe deficit in mechanical capacity of these myofilaments as has been demonstrated by the incorporation of point mutations in skeletal MyHC genes in Drosophila, C. elegans, and Dictyostelium or by targeted disruption of fast MyHC IIb or IId in mice (Ruppel and Spudich, 1996). A muscular dystrophy described in mice has been linked to a mutation in the region that links the Ig with the PEVK domain in the skeletal isoform of titin (Garvey et al., 2002). While the heterozygote mice are unaffected by this mutation, homozygotes develop progressive muscle degeneration and severe structural abnormalities (Garvey et al., 2002). As detailed above, each of these domains is important in the passive properties of both skeletal and cardiac muscle (Granzier et al., 1996, Linke et al., 1998). Given that the Ig and PEVK domains are stretched sequentially and not in parallel, a mutation in the linker region may disrupt this transition in passive stretch. In addition, this mutation is in the calpain binding domain of titin. Calpain is a highly autolytic, skeletal muscle-specific protease that is presumably stabilized by its binding to titin. The significance of this interaction is supported by mutations in the calpain gene found in patients exhibiting human limb-girdle muscular dystrophy characterized by pelvic girdle weakness, atrophic pattern of muscle involvement, lax abdominal muscles and mobility difficulties (Bushby, 1999). Thus, the etiology of this mutation as a muscular dystrophy may be due to the autolytic activity of calpain, abnormal force transmission, or both.

19.6

Conclusions

The discovery of sarcomeric mutations underlying human cardiac and skeletal myopathies has greatly accelerated our understanding of the how myofilament contractile dysfunction can induce a diseased state. The ability to recapitulate many aspects of the disease state in transgenic models has hastened this learning process. One of the more confounding issues is the fact that the same mutations can induce a clinically heterogeneous phenotype in humans. Similarly, the same mutation inserted into the murine genome does not exactly mimic the human disease. For one thing, murine physiology is very different from that of larger mammals. In addition, our ability to study cardiac physiology although improving, is limited when compared to the study of larger mammals (Robbins, 2000). Clearly, the context and physiological environment play a critical role in determining the

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phenotype. Nevertheless, the transgenic mouse model, including the many other tools described in this chapter, play a critical role in describing the molecular pathogenesis of the disease. The elucidation of the signaling pathways that participate in the remodeling process associated with these myopathies will provide the necessary link between sarcomeric mutations, muscle dysfunction, and the manifestation of the disease.

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Trombitas, K., et al. 2000. Extensibility of isoforms of cardiac titin: Variation in contour length of molecular subsegments provides a basis for cellular passive stiffness diversity [in process citation]. Biophys. J. 79: 3226 3234. Uyeda, T. Q., et al. 1996. The neck region of the myosin motor domain acts as a lever arm to generate movement. Proc. Natl Acad. Sci. USA 93: 4459 4464. VanBuren, P., et al. 1995. Cardiac v1 and v3 myosins differ in their hydrolytic and mechanical activities in vitro. Circ. Res. 77: 439 444. VanBuren, P., et al. 1994. The essential light chain is required for full force production by skeletal muscle myosin. Proc. Natl Acad. Sci. USA 91: 12403 12407. Vemuri, R., et al. 1999. The stretch-activation Res.ponse may be critical to the proper functioning of the mammalian heart. Proc. Natl Acad. Sci. USA 96: 1048 1053. Vikstrom, K. L., et al. 1998. Hypertrophy, pathology, and molecular markers of cardiac pathogenesis. Circ. Res. 82: 773 778. Vikstrom, K. L., et al. 1996. Mice expressing mutant myosin heavy chains are a model for familial hypertrophic cardiomyopathy. Mol. Med. 2: 556 567. Wang, K., et al. 1991. Regulation of skeletal muscle stiffness and elasticity by titin isoforms: A test of the segmental extension model of Res.ting tension. Proc. Natl Acad. Sci. USA 88: 7101 7105. Watkins, H., et al. 1992. Characteristics and prognostic implications of myosin missense mutations in familial hypertrophic cardiomyopathy. N. Engl. J. Med. 326: 1108 1114. Weisberg, A. and Winegrad, S. 1998. Relation between cross-bridge structure and actomyosin atpase activity in rat heart. Circ. Res. 83: 60 72. Welikson, R. E., et al. 1999. Cardiac myosin heavy chains lacking the light chain binding domain cause hypertrophic cardiomyopathy in mice. Am. J. Physiol. 276: H2148 H2158. Winegrad, S. 1999. Cardiac myosin binding protein c. Circ. Res. 84: 1117 1126. Winegrad, S. 2000. Myosin binding protein c, a potential regulator of cardiac contractility [editorial; comment]. Circ. Res. 86: 6 7. Witt, C. C., et al. 2001. Hypercontractile properties of cardiac muscle fibers in a knock-in mouse model of cardiac myosin-binding protein-c. J. Biol. Chem.. 276: 5353 5359.

19 Myosin Myopathies Yang, Q., et al. 1998a. A mouse model of myosin binding protein c human familial hypertrophic cardiomyopathy. J. Clin. Invest. 102: 1292 1300. Yang, Q., et al. 1999. In vivo modeling of myosin binding protein c familial hyper-

trophic cardiomyopathy. Circ. Res. 85: 841 847. Yang, Z., et al. 1998b. Changes in interfilament spacing mimic the effects of myosin regulatory light chain phosphorylation in rabbit psoas fibers. J. Struct. Biol. 122: 139 148.

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20 The Role of Dynein in Disease Richard B. Vallee and Chin-Yin Tai

20.1

Dynein Functional and Structural Classes

Dynein is a minus end-directed microtubule motor involved in a very wide range of physiological activities, from ciliary and flagellar motility to a broad range of cytoplasmic functions. Most of the dozen or more forms of dynein found in vertebrates are axonemal, a term which refers to their association with the bundle of 20 microtubules plus accessory proteins that comprises the working core of both cilia and flagella. Cytoplasmic dyneins, in contrast, participate in diverse activities. The major form of cytoplasmic dynein, also referred to as MAP 1C or dynein 1, is present in all cell types, and is involved in mitosis, vesicular and nuclear transport, organizing and orienting the cytoplasmic microtubule network, and other functions. A second, minor form of cytoplasmic dynein, also referred to as DHC2b or dynein 2, has been implicated both in Golgi organization (Vaisberg et al., 1996) and in a recently discovered form of motility within cilia and flagella required for their growth and maintenance (Rosenbaum et al., 1999). This form of motility, referred to as intraflagellar transport (IFT), affects motile cilia and flagella, but also immotile primary cilia and the modified connecting cilia found in photoreceptor cells and other sensory neurons. All forms of dynein contain at least one, usually two, and in some organisms three motor domains (Fig. 20.1). Each motor domain represents the C-terminal two-thirds (Z 350 380 kDa) of a heavy chain (HC) subunit (Fig. 20.4). The Nterminal third of each HC also forms part of the base of the molecule. The motor domain region of the HC is highly conserved among dynein forms and throughout evolution. The N-terminal portion of the axonemal and cytoplasmic dynein HCs is considerably more divergent. The dyneins all have accessory subunits, which associate primarily with the Nterminus of the heavy chain and participate in cargo binding (Figs. 20.1 and 20.4). Intermediate chains (ICs), typically in the 70-kDa size range, are found in both cytoplasmic and axonemal dyneins. Several classes of light chain (LC) are also common to the two forms of dynein, whereas light intermediate chains

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20.2 Diseases Associated with Axonemal Defects

Stalk

Motor Domain HC C-terminus

Cargo Binding HC N-ter., ICs, LICs, LCs

Electron micrograph of dynein molecule. Cytoplasmic dynein (MAP 1C/dynein1) purified from brain tissue and visualized by quick-freeze rotary shadow electron microscopy. The large globular motor domains, each corresponding to the C-terminal two-thirds of a heavy chain (HC) polypeptide, are at the top, with their extended microtubule-binding stalks.

Figure 20.1.

The base of the dynein molecule is morphologically less well defined. This region consists of the N-terminal third of the HCs, plus the intermediate chains (ICs), light intermediate chains (LICs), and light chains (LCs), each of which has been implicated in cargo binding. From Gee et al., 1997.

(LICs) have so far been found only in the cytoplasmic dyneins. Recent work has revealed a novel form of LIC, at present the only known accessory subunit for dynein 2 (Grissom et al., 2002, Mikami et al., 2002). The dyneins have been implicated in a growing number of diseases, evidence for which will be reviewed here.

20.2

Diseases Associated with Axonemal Defects

Structural defects in the axoneme have long been associated with human ciliary and and flagellar dysfunction, also referred to as primary ciliary dyskinesia (PCD; Afzelius, 1976). Common features of this syndrome include male infertility resulting from sperm immotility, and recurrent respiratory tract infections resulting from ciliary immotility. Hydrocephalus may also be seen, which is thought to result from disruption in the flow of cerebrospinal fluid. A more mysterious feature of PCD found in the subset of patients with Kartagener’s syndrome is situs inversus, the reversed orientation of the visceral organs, including the heart, which may also exhibit developmental abnormalities. How defects in axonemal function might contribute to this condition has been uncertain. However, recent work has revealed the existence of a bundle of ‘nodal cilia’ in the gastrula stage embryo, which have been proposed to produce a directional flow of extracellular fluid at the time of axis determination (Nonaka et al., 1998). Mutations in a dynein HC expressed in the nodal cells and limited additional regions have been correlated with altered left right asymmetry in mice (Supp et al., 1999).

20 The Role of Dynein in Disease

This result and the absence of axonemal dynein arms in many PCD cases suggest that axonemal immotility syndromes can be caused by mutations in dynein itself. Indeed, mutations in an axonemal dynein outer arm HC have recently been associated with primary ciliary dyskinesia in humans (Olbrich et al., 2002), a result which has also been replicated in a mouse model (Fig. 20.2; Ibanez-Tallon et al., 2002). Mutations in an axonemal dynein outer arm intermediate chain have also been associated with PCD (Guichard et al., 2001, Pennarun et al., 1999). In view of the substantial number of dynein forms associated with a given cilium or flagellum (e. g. Piperno et al., 1990), and the large number of dynein HC and accessory subunit genes in mammals (Tanaka et al., 1995, Vaughan et al., 1996), it seems reasonable to expect that the number of dynein loci in humans affected by axonemal dysfunction syndromes may prove to be substantial. Dynein 2, the minor form of cytoplasmic dynein, could also potentially be involved in ciliary disease, although there is no direct evidence for this possibility at present. Dynein 2 has been implicated in retrograde IFT, so far within flagella (Pazour et al., 1999, Porter et al., 1999) and within the connecting cilia of sensory neurons in C. elegans (Signor et al., 1999). Transport involves large proteinaceous rafts of particles of defined polypeptide composition which move bidirectionally beneath the ciliary or flagellar membrane and may serve as platforms for structural components of the axoneme (Cole et al., 1998). A mutation in a mouse gene encoding a subunit of the IFT particles was recently found to result in a fatal form of polycystic kidney disease in mice, associated with shorter primary cilia in the kidney (Pazour et al., 2000), and with defects in retinal connecting cilia, associated with retinal degeneration (Pazour et al., 2002). Whether mutations in dynein 2 will prove to produce related effects, and whether mutations in any of the genes involved in IFT occur in the human population remains to be determined.

Effect of dynein heavy chain mutation on ciliary structure. (A) Schematic diagram of a cilium in cross-section. The microtubules are arranged in a 9 2 fashion and are connected through dynein arms, nexin links and

Figure 20.2.

radial spokes. (B) Electron microscopic images of ciliary cross-sections from the nasal mucosa of a wild-type and an Mdnah5 /--mouse, which lacks the outer dynein outer arms. From IbanezTallon et al. 2002.

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20.3 Role of a Cytoplasmic Dynein Light Chain in Retinitis Pigmentosa

20.3

Role of a Cytoplasmic Dynein Light Chain in Retinitis Pigmentosa

Retinitis pigmentosa (RP) is a degenerative disease of the retina which typically manifests itself between 10 and 30 years of age. A number of mutations in the rhodopsin gene have been associated with inherited RP, several located within the cytoplasmic tail of the protein (Sung et al., 1991, Macke et al., 1993). Yeast two-hybrid analysis identified a dynein LC, TcTex-1, as a binding partner for the rhodopsin cytoplasmic domain (Tai et al., 1999). Of considerable interest are four distinct missense mutations associated with RP which interfered with this interaction. Together these results suggested that TcTex-1 served as a linker between cytoplasmic dynein and rhodopsin, a conclusion which was also supported by biochemical cofractionation of cytoplasmic dynein and the photopigment. Rhodopsin requires continuous replenishment in the outer segment of the rod cell. After biosynthesis, it is transported within the inner segment, and then through a non-motile connecting cilium which links the inner to the outer segment (Fig. 20.3).

Figure 20.3. Organization of vertebrate rod photoreceptor cell. Rhodopsin is synthesized in the inner segment and is transported to the stacks of membrane disks located within the outer segment. Cytoplasmic microtubules are shown extending throughout the inner segment from the basal body region. The connecting cilium also extends from a basal body to link the inner and outer segments. From Tai el al., 1999.


The transport pathway has not been fully elucidated, but the evidence for a role for cytoplasmic dynein appears to provide part of the answer. Microtubules within the inner segment emanate from their minus ends from a pair of basal bodies at the base of the connecting cilium . Therefore, cytoplasmic dynein-dependent transport should direct rhodopsin toward the base of this structure. TcTex-1 was found along with rhodopsin in the inner segment (Tai et al., 1999), where dynein 1 HC is also present (Baker et al., 2000), consistent with this possibility. The state of rhodopsin during transport is unknown, but it is likely to be associated either with membranous vesicles or the plasma membrane. Whether it is subsequently transported through the connecting cilium is unknown. Dynein 2 is associated with this structure, and at a lower concentration within the inner segment (Mikami et al., 2002). However, TcTex-1 has not been detected in dynein 2 preparations (Grissom et al., 2002, Mikami et al., 2002) or within the connecting cilium, suggesting that it is a defect specifically in dynein 1-mediated transport that is responsible for retinitis in the cases studied. Evidence has been obtained that the mutant rhodopsin is, indeed, mislocalized in a manner consistent with a defect in dynein-mediated transport. Immunohistochemical analysis of mice heterozygous for a C-terminal mutant form of rhodopsin revealed the mutant protein to accumulate in the plasma membrane of the rod cell body and inner segment, in contrast to the normal targeting of the wild-type protein (Sung et al., 1994). The role of the TcTex-1 subunit of dynein in rhodopsin targeting was examined in polarized MDCK kidney epithelial cells (Tai et al., 2001). TcTex-1 expression was found to be dramatically depressed by over-expression of a related LC, RP3, which, itself, does not bind rhodopsin, allowing for comparison between TcTex-1-expressing and non-expressing cells. Heterologously expressed rhodopsin was specifically targeted to the apical plasma membrane in cells expressing TcTex-1, but the photopigment was found uniformally distributed at the apical and basolateral cell surfaces in its absence, consistent with a complete loss of dynein-mediated transport.

20.4

Role of Cytoplasmic Dynein in the Smooth Brain Disease Lissencephaly

Lissencephaly (smooth brain) is a human brain developmental disease in which part or most of the brain surface lacks its normal convolutions. Neuronal differentiation appears to be normal, but the neurons, especially within the cerebral cortex, are broadly dispersed relative to the well defined layers normally seen. Based on these observations, it is thought that these defects result from failure in the outward migration of neurons from the ventricular and subventricular zones where they are generated. Presumably, this problem is only partial, as neurons do leave the dividing cell layers but then fail to reach their destination. The type I form of the disease is caused by sporadic mutations in one of the two alleles of the LIS1 gene (Reiner et al., 1993), resulting in a decreased level of LIS1 expression. Mice hemizygous for LIS1 exhibit a comparable neuronal misdistribu-

501

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20.4 Role of Cytoplasmic Dynein in the Smooth Brain Disease Lissencephaly

tion phenotype to that observed in humans (Hirotsune et al., 1998). Homozygous null mutations in LIS1 are unknown in humans, but homozygous LIS1 mutant mice exhibit pre-implantation embryonic lethality, a far more severe phenotype than expected if LIS1 were involved only in neuronal migration. Apparent orthologs of LIS1 have been found to function in the cytoplasmic dynein pathway in a number of lower eukaryotic organisms. Initially, defects in the LIS1-related NudF gene of Aspergillus nidulans were found to interfere with nuclear migration within the hyphae of this filamentous fungus. This phenotype is identical to that produced by mutations in the cytoplasmic dynein 1 HC, which is encoded by the NudA gene (Xiang et al., 1995). In S. cerevisiae, the PAC1 gene is involved in nuclear orientation during cell division, and mutations in PAC1 produce a common phenotype with those in genes encoding dynein subunits (Geiser et al., 1997). In Drosophila the dLIS1 phenotype is more complex, but defects in nuclear positioning in oocyte and eye development are observed, again similar to the effects of dynein mutations (Swan et al., 1999). Together, these results suggested that LIS1 might function in the dynein pathway in vertebrates. It also seemed reasonable that defects in a motor protein might affect neuronal motility. In fact, impaired nuclear migration was reported within the processes of cerebellar granule cells obtained from mice hemizygous for LIS1 (Hirotsune et al., 1998). Complicating matters is the finding that, in vertebrates, LIS1 co-purifies stoichiometrically with the catalytic subunits of platelet activating factor acetylhydrolase 1b (PAF acetylhydrolase 1b) (Hattori et al., 1994). PAF acetylhydrolases inactivate platelet activating factor (PAF), a lipid messenger involved in the inflammatory response and possibly other physiological functions. Application of PAF has been reported to arrest the migration of nuclei within the processes of cultured cerebellar granule cells. How this effect of PAF, the levels of which are controlled by PAF acetylhydrolase, relates to a possible role for cytoplasmic dynein in neuronal motility has been uncertain. Evidence from work in vertebrates has since indicated that LIS1 interacts physically with cytoplasmic dynein 1 and its companion regulatory complex dynactin (Faulkner et al., 2000, Smith et al., 2000, Sasaki et al., 2000). Unlike the interaction between LIS1 and the catalytic subunits of PAFAH, which is strong and stoichiometric (Hattori et al., 1993), only a small fraction of dynein, dynactin, and LIS1 coimmunoprecipitate with each other. However, extensive phenotypic and localization data in favor of a common site of action in mammalian cultured cells has been obtained. Over-expressed full-length LIS1 (Faulkner et al., 2000) or N- and C-terminal LIS1 fragments (Tai et al., 2002) were each found to produce a pronounced accumulation of cells in mitosis. Application of antisense oligonucleotides or antibody microinjection also interfered with mitosis. LIS1 was localized to two sites within the mitotic cell, the cortex and the kinetochores by immunofluorescence microscopy of endogenous (Faulkner et al., 2000) or heterologously expressed protein (Tai et al., 2002). Cytoplasmic dynein itself had previously been identified at the cortex of dividing cells, where it was proposed to orient the spindle (Busson et al., 1998). It is also prominent at kinetochores, where it may play a role in microtubule


capture (Rieder and Alexander, 1990), poleward chromosome movement (Savoian et al., 2000, Sharp et al., 2000), and the removal of checkpoint proteins from this site (Howell et al., 2001, Scaerou et al., 2001). In polarized epithelial cells, in which mitotic spindles typically lie within the epithelial plane, LIS1 over-expression randomized spindle orientation, with concomitant disruption of the cortical association of dynein and dynactin (Faulkner et al., 2000). Injection of anti-LIS1 antibody into dividing cells caused pronounced delays in the alignment of individual chromosomes at the metaphase plate, as was also observed with antibody to dynein 1 IC. The extent of LIS1 participation in non-mitotic dynein functions is not fully clear. LIS1 does seem to participate in at least one form of nuclear migration in vertebrates as well as lower eukaryotes, as noted above (Hirotsune et al., 1998). However, over-expression of LIS1 or its N- and C-terminal fragments had no effect on the distribution of other membranous organelles, including the Golgi apparatus, endosomes and lysosomes in one set of studies (Faulkner et al., 2000, Tai et al., 2002). In another study full-length LIS1 over-expression in normal cells or cells from a LIS1 hemizygous mouse was reported to compact the Golgi apparatus (Smith et al., 2000), and the basis for the different results is not yet certain. The Aspergillus NudF protein has been reported to localize to the ends of growing interphase microtubules, where dynein HC was also detected (Han et al., 2001), and mutations in either protein altered microtubule assembly dynamics. LIS1 over-expressed in mammalian cells associates with growing microtubule ends (Coquelle et al., 2002, N. E. Faulkner, et al., unpublished data) where dynein and dynactin have also been localized (Vaughan et al., 1999) and displaces the two complexes (Faulkner et al., 2000). However, endogenous LIS1 has not been observed at these sites in mammalian cells, and the physiological function of this fraction is not yet certain. The kinetochore has provided a useful site for investigating the hierarchy of LIS1 interactions at the cellular level. Endogenous LIS1 could be displaced from this site by over-expression of an N-terminal LIS1 fragment, but there was no detectable effect on the association of dynein or dynactin with the kinetochore (Tai et al., 2002). Conversely, LIS1 was displaced along with dynein and dynactin by over-expression of the dynamitin subunit of dynactin (Tai et al., 2002, Coquelle et al., 2002). These results suggest an intimate connection between LIS1 and the dynein and dynactin complexes. Evidence has, in fact, been obtained for a direct interaction between LIS1 and the dynein 1 HC (Sasaki et al., 2000, Tai et al., 2002, Hoffmann et al., 2002), as well as with the dynein ICs and the dynamitin subunit of dynactin (Tai et al., 2002) by use of either the yeast-two hybrid method or by co-immunoprecipitation of co-expressed polypeptides. The specific role of LIS1 in dynein regulation remains an important question. The interaction of LIS1 with multiple subunits of dynein and dynactin suggests thats LIS1 could serve to modulate the interaction between the two complexes. Of additional interest is the mode of interaction between LIS1 and the dynein HC. LIS1 was found to bind to two distinct sites located some 1000 amino acids apart within the HC (Fig. 20.4). One site lies near the N-terminus of the HC and overlaps sites for IC and LIC binding (Tai et al., 2002). Thus, it is located

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20.4 Role of Cytoplasmic Dynein in the Smooth Brain Disease Lissencephaly

within the region thought to be located at the base of the dynein complex responsible for cargo binding and subcellular targeting. The other site for LIS1 binding lies within the dynein motor domain (Sasaki et al., 2000, Hoffmann et al., 2001, Tai et al., 2002). This portion of the dynein HC consists of a series of six AAA ATPase repeats (Neuwald et al., 1999) probably organized into a ring (Samso et al., 1998). Protruding between the fourth and fifth AAA module is a 10 12-nm stalk, with the microtubule binding site at its tip (Gee et al., 1997). LIS1 interacts with fragments which contain the first AAA repeat (Sasaki et al., 2002, Hoffmann et al., 2001, Tai et al., 2002), and it can, in fact, interact with a fragment corresponding precisely to this structural unit (Tai et al., 2002). This region of the HC has been most clearly implicated in motor function. Vanadate-mediated photocleavage splits the HC in this region, destroying most of its ATPase and motor activity (Gibbons et al., 1987). Conversely, replacement of the lysine residue within the P-loop element of the first AAA domain blocked vanadate photocleavage and induced a rigor-like microtubule binding state (Gee et al., 1997). The role of the other AAA modules remains uncertain, although they may also bind ATP (Mocz and Gibbons, 1996) and participate in or indirectly affect hydrolysis (Gee et al., 1997). Thus, LIS1 interacts with the region of the HC most clearly implicated in motor activity, suggesting a role in regulating motor function. Type I lissencephaly could, therefore, represent the first disease involving a defect in cytoplasmic dynein motor activity. Whether this will, indeed, prove to be the case, and whether LIS1 promotes or inhibits motor activity remains to be determined. Cargo Binding Subunit Interactions

Motor Domain AAA

AAA

AAA

AAA Stalk

(AAA)

(AAA)

LIS1 binding sites HC-HC IC/LC LIC Figure 20.4. Domain organization of dynein

heavy chain (HC) showing sites of LIS1 interaction. Rat cytoplasmic dynein HC of 532 kDa is shown (Mikami et al., 1993, Zhang et al., 1993). The N-terminal third of the polypeptide has been implicated in self-association and binding of intermediate chain light chain complexes (IC/LC) and light intermediate chains (LICs) (Habura et al., 1999, Tynan et al., 2000). The Cterminal two-thirds of the HC is composed of a series of six predicted AAA ATPase domains (Neuwald et al., 1999) with an extending microtubule-binding stalk between the fourth and fifth AAA unit (Gee et al., 1997). Based on a

yeast two-hybrid library screen (Sasaki et al., 2000) and analysis of interactions between coexpressed LIS1 and HC fragments(Tai et al., 2002), two sites for LIS1 binding were deduced. One site overlaps the sites for dynein heavy chain dimerization, IC, and LIC binding. The other site corresponds to the first AAA repeat sequence. LIS1 was also found to interact with the ICs and the p150Glued subunit of the dynactin complex. These data suggest a role for LIS1 in linking subunits between dynein and dynactin, and, possibly in the regulation of dynein motor activity. From Tai et al., 2002.


Despite the evidence for a role for LIS1 in dynein regulation, it remains unresolved whether type I lissencephaly is caused by a defect in dynein or PAFAH1b function. Efforts to inhibit dynein and PAFAH independently during brain development may be necessary to shed light on this issue. It is also possible that LIS1 not only participates in both pathways, but serves to link them. However, existing evidence based on co-immunoprecipitation of each of the two complexes and affinity chromatography with the PAFAH catalytic subunits suggest that the LIS1 interactions are biochemically independent (Smith et al., 2000, Tai et al., 2002). The catalytic subunits of PAFAH 1b were also undetectable in those vertebrate cells in which a dynein-like distribution and phenotype for LIS1 were observed (Tai et al., 2002), and they appear to be absent from the genome of S. cerevisiae in which mutations in the LIS1-related gene PAC1 produce a dynein-like phenotype (Geiser et al., 1997). If type I lissencephaly proves to result partially, or completely, from a defect in dynein function, what aspects of dynein function lead to altered neuronal distribution? Defects in nuclear migration could certainly be envisioned to contribute to altered migration of progenitor cells. Some evidence does exist for nuclear translocation within the processes of migrating neuronal/glial progenitor cells in brain slices, although how defects in this aspect of progenitor cell motility affect the overall migration process is unknown. Cerebellar granule cells, in which decreased LIS1 expression interferes with nuclear migration (Hirotsune et al., 1998), migrate by a somewhat different mechanism and their distribution is relatively unaffected in lissencephaly. In view of the pronounced mitotic effects observed in LIS1 over-expressing, under-expressing, or antibody-injected cells, alteration in the timing, extent, or orientation of cell division could also be expected to affect neuronal distribution. Decreased brain mass can, in fact, be observed in lissencephalic children (see e. g. images in Sweeney et al., 2000), and homozygous LIS1 mutations clearly affect cell proliferation in Drosophila (Lei and Warrior, 2000, Liu et al., 1999, 2000) and mouse (Hirotsune et al., 1998). Of additional interest, the orientation of the cell division plane has been correlated with the timing of neurogenesis in mammals, which, in turn, determines the subsequent migration distance (Chenn and McConnell, 1995, McConnell and Kaznowski, 1991). Conceivably, therefore, cell division defects could alter neuronal distribution (Faulkner et al., 2000, Vallee et al., 2000, 2001). To what extent the dramatically altered pattern seen in type I lissencephaly results from mitotic errors remains an important unanswered question. Cytoplasmic dynein could well have an additional direct role in neuronal migration. Recent work has found that cytoplasmic dynein participates in the directed migration of fibroblasts during the wound-healing process (Etienne-Manneville, 2001, Palazzo et al., 2001). Over-expression of the dynamitin subunit of dynactin or injection of anti-dynein antibody each arrested the rotation of the nuclear and microtubule network that precedes migration into the wound region. Conceivably this new function for dynein could be a general one, and dynein could play a related role in other types of directed movement, such as that of glial/neuronal progenitor cells. Whether this is the case, and whether LIS1 regulates this important process, remain additional questions for future investigation.

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20 The Role of Dynein in Disease and gamma-tubulin. J. Biol. Chem. 276: 38877 38884. Howell, B. J., B. F. McEwen, J. C. Canman, D. B. Hoffman, E. M. Farrar, C. L. Rieder, and E. D. Salmon. 2001. Cytoplasmic dynein/ dynactin drives kinetochore protein transport to the spindle poles and has a role in mitotic spindle checkpoint inactivation. J. Cell Biol. 155: 1159 1172. Ibanez-Tallon, I., S. Gorokhova, and N. Heintz. 2002. Loss of function of axonemal dynein Mdnah5 causes primary ciliary dyskinesia and hydrocephalus. Hum. Mol. Genet. 11: 715 721. Kakita, A. and J. E. Goldman. 1999. Patterns and dynamics of SVZ cell migration in the postnatal forebrain: monitoring living progenitors in slice preparations. Neuron 23: 461 472. Lei, Y. and R. Warrior. 2000. The drosophila lissencephaly1 (DLis1) gene is required for nuclear migration [In Process Citation]. Dev. Biol. 226: 57 72. Liu, Z., R. Steward, and L. Luo. 2000. Drosophila Lis1 is required for neuroblast proliferation, dendritic elaboration and axonal transport. Nature Cell Biol. 2: 776 783. Liu, Z., T. Xie, and R. Steward. 1999. Lis1, the Drosophila homolog of a human lissencephaly disease gene, is required for germline cell division and oocyte differentiation. Development 126: 4477 4488. Macke, J. P., C. M. Davenport, S. G. Jacobson, J. C. Hennessey, F. Gonzalez-Fernandez, B. P. Conway, J. Heckenlively, R. Palmer, I. H. Maumenee, P. Sieving, and et al., 1993. Identification of novel rhodopsin mutations responsible for retinitis pigmentosa: implications for the structure and function of rhodopsin. Am. J. Hum. Genet. 53: 80 89. McConnell, S. K., and C. E. Kaznowski. 1991. Cell cycle dependence of laminar determination in developing neocortex. Science 254: 282 285. Mikami, A., B. M. Paschal, M. Mazumdar, and R. B. Vallee. 1993. Molecular cloning of the retrograde transport motor cytoplasmic dynein (MAP 1C). Neuron 10: 787 796. Mocz, G., and I. R. Gibbons. 1996. Phase partition analysis of nucleotide binding to axonemal dynein. Biochemistry 35: 9204 9211. Neuwald, A. F., L. Aravind, J. L. Spouge, and E. V. Koonin. 1999. AAA: A class of chaperone-like ATPases associated with the as-

sembly, operation, and disassembly of protein complexes. Genome Res. 9: 27 43. Nonaka, S., Y. Tanaka, Y. Okada, S. Takeda, A. Harada, Y. Kanai, M. Kido, and N. Hirokawa. 1998. Randomization of left-right asymmetry due to loss of nodal cilia generating leftward flow of extraembryonic fluid in mice lacking KIF3B motor protein. Cell 95: 829 837. Olbrich, H., K. Haffner, A. Kispert, A. Volkel, A. Volz, G. Sasmaz, R. Reinhardt, S. Hennig, H. Lehrach, N. Konietzko, M. Zariwala, P. G. Noone, M. Knowles, H. M. Mitchison, M. Meeks, E. M. Chung, F. Hildebrandt, R. Sudbrak, and H. Omran. 2002. Mutations in DNAH5 cause primary ciliary dyskinesia and randomization of left-right asymmetry. Nature Genet. 30: 143 144. Palazzo, A. F., H. L. Joseph, Y. J. Chen, D. L. Dujardin, A. S. Alberts, K. K. Pfister, R. B. Vallee, and G. G. Gundersen. 2001. Cdc42, dynein, and dynactin regulate MTOC reorientation independent of Rho-regulated microtubule stabilization. Curr. Biol. 11: 1536 1541. Paschal, B. M., and R. B. Vallee. 1987. Retrograde transport by the microtubule associated protein MAP 1C. Nature 330: 181 183. Pazour, G. J., S. A. Baker, J. A. Deane, D. G. Cole, B. L. Dickert, J. L. Rosenbaum, G. B. Witman, and J. C. Besharse. 2002. The intraflagellar transport protein, IFT88, is essential for vertebrate photoreceptor assembly and maintenance. J. Cell Biol. 157: 103 113. Pazour, G. J., B. L. Dickert, Y. Vucica, E. S. Seeley, J. L. Rosenbaum, G. B. Witman, and D. G. Cole. 2000. Chlamydomonas IFT88 and its mouse homologue, polycystic kidney disease gene tg737, are required for assembly of cilia and flagella. J. Cell Biol. 151: 709 718. Pazour, G. J., B. L. Dickert, and G. B. Witman. 1999. The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. J. Cell Biol. 144: 473 481. Pennarun, G., E. Escudier, C. Chapelin, A. M. Bridoux, V. Cacheux, G. Roger, A. Clement, M. Goossens, S. Amselem, and B. Duriez. 1999. Loss-of-function mutations in a human gene related to Chlamydomonas reinhardtii dynein IC78 result in primary ciliary dyskinesia. Am. J. Hum. Genet. 65: 1508 1519. Piperno, G., Z. Ramanis, E. F. Smith, and W. S. Sale. 1990. Three distinct inner dynein arms in Chlamydomonas flagella: Molecular com-

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and L. H. Tsai. 2000. Regulation of cytoplasmic dynein behaviour and microtubule organization by mammalian Lis1. Nature Cell Biol. 2: 767 775. Sung, C. H., C. M. Davenport, J. C. Hennessey, I. H. Maumenee, S. G. Jacobson, J. R. Heckenlively, R. Nowakowski, G. Fishman, P. Gouras, and J. Nathans. 1991. Rhodopsin mutations in autosomal dominant retinitis pigmentosa. Proc. Natl Acad. Sci. USA 88: 6481 6485. Sung, C. H., C. Makino, D. Baylor, and J. Nathans. 1994. A rhodopsin gene mutation responsible for autosomal dominant retinitis pigmentosa results in a protein that is defective in localization to the photoreceptor outer segment. J. Neurosci. 14: 5818 5833. Supp, D. M., M. Brueckner, M. R. Kuehn, D. P. Witte, L. A. Lowe, J. McGrath, J. Corrales, and S. S. Potter. 1999. Targeted deletion of the ATP binding domain of left-right dynein confirms its role in specifying development of left-right asymmetries. Development 126: 5495 5504. Swan, A., T. Nguyen, and B. Suter. 1999. Drosophila Lissencephaly-1 functions with Bic-D and dynein in oocyte determination and nuclear positioning. Nature Cell Biol. 1: 444 449. Sweeney, K. J., G. D. Clark, A. Prokscha, W. B. Dobyns, and G. Eichele. 2000. Lissencephaly associated mutations suggest a requirement for the PAFAH1B heterotrimeric complex in brain development. Mech. Dev. 92: 263 271. Tai, A. W., J. Z. Chuang, C. Bode, U. Wolfrum, and C. H. Sung. 1999. Rhodopsin’s carboxyterminal cytoplasmic tail acts as a membrane receptor for cytoplasmic dynein by binding to the dynein light chain Tctex-1. Cell 97: 877 887. Tai, A. W., J. Z. Chuang, and C. H. Sung. 2001. Cytoplasmic dynein regulation by subunit heterogeneity and its role in apical transport. J. Cell Biol. 153: 1499 1509. Tai, C. Y., D. L. Dujardin, N. E. Faulkner, and R. B. Vallee. 2002. Role of dynein, dynactin, and CLIP-170 interactions in LIS1 kinetochore function. J. Cell Biol. 156: 959 968. Tanaka, Y., Z. Zhang, and N. Hirokawa. 1995. Identification and molecular evolution of new dynein-like protein sequences in rat brain. J. Cell Sci. 108: 1883 1893.

20 The Role of Dynein in Disease Troutt, L. L., and B. Burnside. 1988. Microtucalization of dynactin and cytoplasmic dynein bule polarity and distribution in teleost with CLIP-170 at microtubule distal ends. J. photoreceptors. J. Neurosci. 8: 2371 2380. Cell Sci. 112: 1437 1447. Tynan, S. H., M. A. Gee, and R. B. Vallee. 2000. Vaughan, K. T., A. Mikami, B. M. Paschal, Distinct but overlapping sites within the cyE. L. F. Holzbaur, S. M. Hughes, C. J. Echetoplasmic dynein heavy chain for dimerizaverri, K. Moore, D. J. Gilbert, N. G. Copeland, tion and for intermediate chain and light in- N. A. Jenkins, and R. B. Vallee. 1996. Identitermediate chain binding [In Process Citafication of multiple genomic loci involved in tion]. J. Biol. Chem. 275: 32769 32774. dynein-based motility. Genomics 36: 29 38. Vaisberg, E. A., P. M. Grissom, and J. R. McIn- Xiang, X., A. H. Osmani, S. A. Osmani, M. Xin, tosh. 1996. Mammalian cells express three and N. R. Morris. 1995. NudF, a nuclear midistinct dynein heavy chains that are gration gene in Aspergillus nidulans, is localized to different cytoplasmic organelles. similar to the human LIS-1 gene required for J. Cell Biol. 133: 831 842. neuronal migration. Mol. Biol. Cell 6: Vallee, R. B., N. E. Faulkner, and C. Tai. 2000. 297 310. The role of cytoplasmic dynein in the human Zhang, Z., Y. Tanaka, S. Nonaka, H. Aizawa, brain developmental disease lissencephaly. H. Kawasaki, T. Nakata, and N. Hirokawa. Biochim. Biophys. Acta 1496: 89 98. 1993. The primary structure of rat brain (cyVallee, R. B., C. Tai, and N. E. Faulkner. 2001. toplasmic) dynein heavy chain, a cytoplasmic LIS1: cellular function of a disease-causing motor enzyme. Proc. Natl. Acad. Sci. USA 90: gene. Trends Cell Biol. 11: 155 160. 7928 7932. Vaughan, K. T., S. H. Hughes, C. J. Echeverri, N. F. Faulkner, and R. B. Vallee. 1999. Co-lo-

509

21 Molecular Motors in Sensory Defects Karen B. Avraham

21.1

Introduction

Mutations in a multitude of molecular motors cause sensory defects. The loss of these senses, in particular vision and hearing, is not life-threatening, but it can severely affect one’s quality of life. The myosin mutations associated with hearing loss affect the hair cells of the cochlea, and the mutations associated with visual impairment affect the retinal pigment epithelial (RPE) cells and photoreceptors of the retina in a disease termed retinitis pigmentosa (RP) (Fig. 21.1). Mutations in five myosins, myosin IIIA, myosin VI, myosin VIIA, myosin XV, and the non-muscle-myosin heavy-chain MYH9 are known to date to lead to hearing loss, while myosin VIIA mutations also cause RP and hearing loss in Usher syndrome type IB. This chapter will describe these genes, the proteins they encode, and the pathology caused by mutations in these genes.

21.2

Development of the Visual and Auditory Sensory Systems

The three major sensory organs of the head, namely the eye, the ear and the nose, develop from interactions between neural crest cells and epidermal thickenings, the cranial ectodermal placodes. The organs of the inner ear develop from the otic placode (Goeringer, 1998). The human inner ear is composed of the cochlea, the organ for hearing, and the vestibular apparatus, formed from five organs, the saccule, utricle, and the three semicircular canals. Beginning at or about embryonic day 22 in humans, a thickening of the surface ectoderm occurs on either side of the rhombencephalon. Through a series of signaling mechanisms, interaction, and incorporation of other embryonic tissues, the otic placode develops into the otocyst or otic vesicle. Cells from the medial aspect of each otocyst, along with neural crest cells, develop into both the cochlear and vestibular statoacoustic ganglion. The fluid-filled portion of the otocyst grows into the endolymphatic sacs, the utricle

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21.2 Development of the Visual and Auditory Sensory Systems

The inner ear and eye structures affected by mutations in myosins. (A) The inner ear is composed of the cochlea, the organ responsible for hearing, and the vestibular apparatus, made up of five organs (three semicircular canals, utricle, and saccule) responsible for the position and movement of our head in

Figure 21.1.

space. Degeneration of the inner (IHC) and outer hair cells (OHC) of the organ of Corti often leads to hearing loss. (B) The retina of the eye contains photoreceptors, rods and cones, which along with the pigment epithelium, degenerate to cause retinitis pigmentosa.

and the saccule. The vestibular and cochlear components separate as the cochlear duct is formed in the fifth week. Differentiation of cochlear cells begins in the seventh week, as well as outgrowths from the utricle. Programmed cell death of these outgrowths results in the formation of the three semicircular canals. Ganglion cells from the VIIIth cranial nerve migrate along the cochlear coils to form the spiral cochlear ganglion, whose processes terminate on the organ of Corti hair cells. The otic capsule is formed beginning in the ninth week. The cochlear duct begins to take on the shape it has as in the adult as fluid-filled compartments are formed. Ossification of the otic capsule continues, until the inner ear, with an adult size and configuration, is complete at about week 22.

21 Molecular Motors in Sensory Defects

The human eye develops from the optic vesicles that begin as two bulges from the lateral walls of the diencephalon and the overlying surface ectoderm at embryonic day 22 (Gilbert, 1994). As the vesicles contact the surface ectoderm, this region thickens to form the lens placode that will differentiate into the lens. The optic vesicles invaginate to form the two layers of the optic cup, which differentiates to form the pigmented retina (the outer layer) and the neural retina (the inner layer), which is composed of glia, ganglion neurons, interneurons and light-sensitive photoreceptor neurons. Differentiation of the neural retina, the lens, and the cornea continues throughout embryogenesis.

21.3

Visual Impairment

Visual impairment can result from a number of eye abnormalities, including myopia, glaucoma, heterochromia irides, lens opacities, RP and optic atrophy and macular degeneration, to name a few (Schoem and Grundfast, 1998). The leading cause of vision impairment and blindness is age-related eye diseases, including age-related macular degeneration, cataract, glaucoma and diabetic retinopathy. RP is a disease of the rod and cones characterized by night blindness, visual field constriction, acuity loss and abnormal retinograms (ERGs). The loss can involve only peripheral vision, or can lead to total blindness. RP is a genetically heterogeneous disease that involves mutations in numerous genes and is an isolated clinical phenotype or is associated with several syndromes (http://www.sph.uth. tmc.edu/RetNet/; reviewed in Farrar et al., 2002). Genes with mutations leading to RP includes rhodopsin and 35 other genes; the proteins they encode have functions ranging from retinal development, protein trafficking and photoreceptor structure, to visual transduction. The chromosomal locations of an additional 24 RPs are known, although their causative genes remain to be discovered. The syndromes associated with RP include Usher syndrome (for example, OMIM 276900; http://www.ncbi.nlm.nih.gov/Omim), Bardet-Biedl syndrome (OMIM 209900) and Leber congenital amaurosis (LCA; OMIM 204000). Half of the RP cases are sporadic, with the remaining having a dominant, recessive, or X-linked mode of inheritance. The cells most affected by RP are the retinal rod photoreceptors, one class of neurons in the mammalian retina. RP is a progressive disease, often beginning with loss of night vision and leading to complete blindness. One of the predominant forms of blindness is Usher syndrome (USH), a disease involving both hearing loss and blindness, first described by Usher in 1913 (Usher, 1914). Several clinical types have been described based on the age of onset of RP, the cause of blindness; the age of onset and severity of hearing loss; and the presence or absence of vestibular dysfunction. The most severe form, USH type I (USH1), involves profound congenital sensorineural hearing loss with vestibular dsyfunction and an onset of RP at puberty. USH1 loci have been found on seven chromosomes, including 14q32 (USH1A) (Kaplan et al., 1992), 11q13 (USH1B), 11p15 (USH1C) (Smith et al., 1992), 10q (USH1D) (Wayne et al., 1996), 21q

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21.4 Hearing Impairment

(USH1E), 10q21–22 (USH1F), and 17q24–25 (USH1G). The genes for four of these loci have been identified. The USH1B gene is myosin VII (MYO7A) (Weil et al., 1995), the USH1C gene encodes a PDZ-containing protein named USH1C (Bitner-Glindzicz et al., 2000, Verpy et al., 2000), the USH1D gene is caderin 23 (Bolz et al., 2001, Bork et al., 2001), and the USH1F gene is protocadherin 15 (Ahmed et al., 2001, Alagramam et al., 2001). USH2 is characterized by mild to severe hearing loss, normal vestibular response, and RP. The three forms of USH2 are localized on chromosome 1q41 (USH2A) (Kimberling et al., 1990), on 3p23–24.2 (USH2B) (Hmani et al., 1999) and on chromosome 5q14.3–q21.3 (USH2C) (Pieke-Dahl et al., 2000). The gene for USH2A encodes a novel protein with laminin epidermal growth factor and fibronectin type III motifs (Eudy et al., 1998). To date, there is only one form known for USH3, characterized by post-lingual progressive hearing loss, progressive visual loss due to RP, and variable presence of vestibular dysfunction (Sankila et al., 1995). The gene encodes a protein named clarin-1, a novel protein with homology to stargazin (Aadato et al., 2002, Joensuu et al., 2001). Of all the USH genes, only one, myosin VIIA, encodes a molecular motor. This gene was isolated with the help of a mouse model for human deafness, shaker 1 (sh1). The locus for this mouse maps to chromosome 7 in a region homologous with human chromosome 11. Mutations in the myosin VIIA gene were found concurrently in both the sh1 mouse (Gibson et al., 1995) and USH1B patients (Weil et al., 1995). Two years later, myosin VIIA mutations were found in both dominant and recessive forms of human deafness (see Section 21.5.1.4).

21.4

Hearing Impairment

Hearing loss is considered to be the most common sensory loss, affecting 1 in 1000 newborns and up to half of the population at the age of 80 years. Approximately 60 % of hearing loss is due to genetic mutations. Hearing loss is genetically heterogeneous, since it is caused by mutations in many different genes (reviewed in Petit et al., 2001). Furthermore, hearing loss is most often monogenic in nature; namely, mutations in only one gene cause deafness in any given individual (or family). Genetic deafness occurs both in association with other signs in the form of syndromic hearing loss (SHL), or as an isolated finding, non-syndromic hearing loss (NSHL). It is estimated that 30 % of genetic hearing loss is associated with syndromes. There are over 500 clinically defined syndromes that include hearing loss. Hearing loss has been found associated with diabetes, peripheral neuropathy, craniofacial abnormalities, and dwarfism, to name just a few. Hearing loss is a major feature in a number of prevalent syndromes, including Pendred syndrome (with goitre), Waardenburg syndrome (WS, with pigmentary anomalies and widely-spaced eyes), Alport syndrome (with kidney defects), branchio oto renal (BOR) syndrome (with craniofacial and kidney defects), and Usher syndrome (with retinitis pigmentosa). The genes for some of these forms of SHL have been identified, in-


cluding SLC26A4 (Everett et al., 1997) (Pendred syndrome), PAX3 (Hoth et al., 1993) (WS type I and III), MITF (Tassabehji et al., 1994) (WS type II), EDNRB (Attie et al., 1995), EDN3 (Edery et al., 1996), and SOX10 (Pingault et al., 1998) (WS type IV), EYA1 (Abdelhak et al., 1997) (BOR syndrome), COL4A5 (Barker et al., 1990), COL4A3, and COL4A4 (Mochizuki et al., 1994) (X-linked and autosomal Alport syndrome). Approximately 70 % of genetic hearing loss is non-syndromic in nature. The largest proportion is inherited in an autosomal recessive mode (Z 80 %; defined as DFNB loci), Z 18 % is inherited in an autosomal dominant mode (defined as DFNA loci), and Z 2 % is X-linked (defined as DFN loci). Mitochondrial/maternal inheritance also contributes to a small (1 %) proportion of NSHL. The chromosomal locations for 70 loci have already been found for NSHL (updated regularly in the Hereditary Hearing Loss Homepage, http://dnalab-www.uia.ac.be/dnalab/hhh/). A large variety of genes are associated with NSHL. These include myosins (see Section 21.5), transcription factors, connexins, ion channels, and more. The most prevalent causative gene for NSHL is connexin 26 (GJB2) (Kelsell et al., 1997), which codes for a gap junction protein. Connexin 26 mutations account for 30 50 % of inherited hearing loss in many parts of the world (Denoyelle et al., 1997, Estivill et al., 1998, Kelley et al., 1998, Shahin et al., 2002, Sobe et al., 2000). Another three connexins, connexin 30 (GJB6) (del Castillo et al., 2002, Grifa et al., 1999, Lerer et al., 2001), connexin 31 (GJB3) (Xia et al., 1998), and connexin 43 (GJA1) (Liu et al., 2001), are also associated with human hearing loss, but they occur with much less frequency.

21.5

Myosins Involved In Sensory Defects

To date, there are five myosins associated with human and mouse hearing loss. Myosins were among the first genes cloned for NSHL in mice. Myosin VIIA mutations were discovered in sh1 mice (see Section 21.5.1.2) and in USH1B simultaneously, followed by the discovery of myosin VI mutations in Snell’s waltzer mice (sv) (Avraham et al., 1995; see Section 21.5.2.2). Five years later, a myosin VI mutation was found in a family with dominant hearing loss (Melchionda et al., 2001; see Section 21.5.2.3). Mutations in myosin XV were discovered simultaneously in mice and humans (Probst et al., 1998, Wang et al., 1998; see Section 21.5.3.2 21.5.3.3). A mutation in MYH9 was identified in a family with autosomal dominant hearing loss (Lalwani et al., 2000; see Section 21.5.4.2). And most recently, three mutations in the myosin IIIA gene were found in a family with autosomal recessive hearing loss (Walsh et al., 2002; see Section 21.5.5.3). Of all these molecular motors, only myosin VIIA is currently known to be associated with a defect in the visual system, although myosin IIIA is also expressed in the retina.

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21.5 Myosins Involved In Sensory Defects

21.5.1

Myosin VIIA

Myosin VII (OMIM 276903) was the seventh myosin to be discovered among the ‘unconventional’ myosins in a polymerase chain reaction (PCR) amplification designed to identify new myosin proteins (Bement et al., 1994). Two isoforms of myosin VII are known, myosin VIIA and myosin VIIB (Chen et al., 2001). The first myosin VIIA was identified in a human epithelial cell line and human leukocytes (Bement et al., 1994). Since then, myosin VIIA has been discovered and characterized in mouse (Gibson et al., 1995), zebrafish (Ernest et al., 2000), C. elegans (Kelleher et al., 2000), and in the Dictyostelium discoideum amoeba (DdMVII) (Titus, 1999). Mutations in myosin VIIA are associated with Usher syndrome type IB and NSHL in humans, NSHL in mice, impaired phagocytosis in Dictyostelium, impaired spermatogenesis in C. elegans, and abnormal mechanotransduction in zebrafish.

Structure, function, and expression of myosin VIIA The human myosin VIIA (locus designation MYO7A) gene consists of 48 exons, spanning a genomic region of over 100 kb (Chen et al., 1996, Weil et al., 1996). The myosin VIIA protein consists of 2215 amino acids and has a predicted molecular weight of 254 kDa. The head/motor domain is 729 amino acids long, the neck is 126 amino acids in length, and the tail is 1360 amino acids long (Fig. 21.2). The unique myosin VIIA tail contains some distinguishing features, including a coiledcoil domain and two repeats of a set of a MyTH4 (myosin tail homology 4) domain and a FERM membrane-binding domain. The repeats are connected by an SH3 domain. Recently, several proteins have been shown to interact with myosin VIIA, revealed through the use of yeast two-hybrid screens. MyRIP is a novel Rab effector, and its interaction with myosin VIIA and Rab27A has led to the speculation that a complex composed of all three proteins bridges retinal melanosomes to the actin cytoskeleton and mediates the trafficking of these melanosomes (El-Amraoui et al., 1996). Keap1, a human homolog of the Drosophila ring canal protein, kelch, interacts with myosin VIIA through the SH3 domain. While the interaction of myosin VIIA and keap1 was identified in a testes library, these two proteins are also co-expressed in the inner ear (Velichkova et al., 2002). Myosin VIIA has also been shown to interact with the ype I alpha regulatory subunit (RI alpha) of protein 21.5.1.1

Figure 21.2.

Myosin VIIA protein domain structure.


kinase A (Kussel-Andermann et al., 2000a). The FERM domain is the region to which this A-kinase-anchoring protein binds. A retina yeast two-hybrid library was used to identify these interacting proteins and they were also found to both be expressed in the outer hair cells of the cochlea and rod photoreceptor cells of the retina. To date, a fourth myosin VIIA-interacting protein has been identified in the yeast two-hybrid library retina library, vezatin, a novel transmembrane protein that bridges myosin VIIA to the cadherin catenins complex (Kussel-Andermann et al., 2000b). Using antibodies specific against myosin VIIA, it has been shown that this motor is expressed in the inner ear, retina, testes, lung, and kidney (El-Amraoui et al., 1996, Hasson et al., 1995; Fig. 21.3). In frog, mouse, rat, and guinea pig inner ears, myosin VIIA is expressed in both the cochlea and vestibular epithelium (Hasson et al., 1997). The specific expression is in the cell bodies, the cuticular plate (the region immediately under the stereocilia) and along the length of the stereocilia, suggesting that this protein might anchor connectors between stereocilia and fulfill a structural role (Fig. 21.3). In the eye, myosin VIIA is expressed in the pigmented epithelium and the photoreceptors of the retina (El-Amraoui et al., 1996, Hasson et al., 1997, Liu et al., 1997c). Myosin VIIA may act as a vesicle transporter in the eye, and be involved in transport of melanosomes or opsin to required regions of the RPE cells that lack these molecules (Liu et al., 1998). Based on its potential structural role, myosin VIIA may also be involved in actin organization, affecting melanosome distribution. Most intriguing, a role for myosin VIIA in auditory transduction has recently been described (Kros et al., 2002). The mechanical sound wave normally enters the fluidfilled cochlea and is transduced into an electrochemical signal. The sound wave causes deflection of the hair bundles that project from the apical portion of the hair cells, composed of actin-rich structures named stereocilia (Fig. 21.1). Transduction channels between adjacent stereocilia open and close, depending on the tension of gating springs. A resting tension is maintained by an adaptation motor, allowing channels to be open even in the absence of stimuli. This element of auditory transduction allows for the incredible sensitivity and speed in the system. In shaker 1 mice

Figure 21.3. Myosin VIIA expression in the ear and eye (el-Amraoui et al. 1996). (A) Immunolabelling of myosin VIIA in the inner (IHC) and outer hair cells (OHC) of a mouse inner ear.

(B) Immunolabelling of myosin VIIa in the pigmented epithelium (E) and (C) photoreceptor (P) cells of the retina of a macaque eye.

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lacking myosin VIIA (see Section 21.5.1.2), the resting tension is absent, requiring non-physiologically large bundle deflections to open the transduction channels. A structural role for myosin VIIA is supported since in this case it appears to participate in anchoring membrane-bound elements to the actin core of the stereocilia.

Shaker 1 Mice and Other Models The shaker 1 (sh1) mouse exhibits circling and deafness and is one of a large group of deaf mouse mutants (see Mouse Mutants with Hearing or Balance Defects, http://www.ihr.mrc.ac.uk/hereditary/mousemutants.htm). Ten recessive sh1 alleles have been discovered over the years, having arisen spontaneously or been generated by radiation. The sh1 locus lies on mouse chromosome 7, in the region homologous to human chromosome 11q13.5 (for regions of homology between human and mouse chromosomes, see Human-Mouse Homology Maps, http:// www.ncbi.nlm.nih.gov/Homology/). During a positional cloning effort to clone the gene for sh1, a Myo7a exon was found in the sh1 region and since another unconventional myosin had been found in stereociliary bundles (Gillespie et al., 1993), myosin VIIA appeared to be a very good candidate for deafness in these mice. This was a key discovery in the identification of the USH1B gene (see Section 21.5.1.3). Since then, a spectrum of Myo7a mutations have been found in sh1 mice (Libby and Steel, 2001, Self et al., 1998). Overtly, sh1 mice appear to have NSHL, along with vestibular dysfunction, as there is no evidence of visual problems. An analysis of sh1 hair cells has revealed a progressive disorganization of the hair bundles and degeneration (Fig. 21.4; Self et al., 1998). The question of why sh1 mice do not develop RP has been an ongoing one. Although sh1 mice do not appear to have any retinal degeneration, they do have defective melanosome distribution in the RPE (Liu et al., 1998). Furthermore, a number of alleles do have electroretinographic (ERG) anomalies, revealed by an observation of de21.5.1.2

An analysis of sh1 hair cells has revealed a progressive disorganization of the hair bundles. SEM of the surface of outer hair cells from 20-day old (A) control mice and

Figure 21.4.

(B) sh1 mutant homozygotes (Myo7a6J allele). Note disorganization of hair bundles. Adapted with permission from Self et al. (1998), Company of Biologists Ltd.


Stereocilia of the myosin VIIa mariner zebrafish mutant shows a splayed pattern. Five-day old larvae were fixed and labelled with Oregon green phalloidin and observed by confocal microscopy. Hair bundles

Figure 21.5.

from (A) control zebrafish are conical in shape, while (B) mutant mariner (the tn3540 allele) larvae show splaying of the stereociliary hair bundles. Adapted with permission from Ernest et al. (2000), by Oxford University Press.

creased ERG amplitudes with normal thresholds (Libby and Steel, 2001). Myosin VIIA has also been shown to be essential for aminoglycoside accumulation in cochlear hair cells and hence may be involved in ototoxicity caused by these antibiotics (Richardson et al., 1997). Unique to the myosins involved in deafness is that myosin VIIA mutations have also been found in zebrafish (Ernest et al., 2000). Zebrafish have two mechanosensory organs, the inner ear and the lateral line organ, both of which contain sensory cells. The particular advantage of studying these lower vertebrates is the visibility of hair cell development. The mariner mutant is a circling zebrafish with inner ear hair bundle defects, manifested as splaying of the stereocilia (Fig. 21.5). The Myo7a missense and nonsense mutations in mariner larvae lead to defects in mechanotransduction and inhibition of apical hair cell endocytosis (Seiler and Nicolson, 1999). The comparative analysis of zebrafish, mouse, and human myosin VIIA reveals striking similarities between their domains and the pathogenesis seen in sensory cells from zebrafish and mice help to address the pathology in humans.

Usher Syndrome Type 1B Mutations in myosin VIIA underlie the pathogenesis of Usher’s syndrome type 1B. First described in 1858 (von Graefe, 1858), this disease was named after Charles Usher, who reported the combination of RP and deafness in Britain (Usher, 1914). In USH1B, retinal (photoreceptor cells) and cochlear and vestibular sensory cells (hair cells) are defective. As a result, these children are born with congenital deafness, usually profound, and with vestibular dysfunction, seen as motor development delay. During childhood, the retinopathy begins as night blindness and progresses to complete blindness, usually by puberty. The USH1B locus (OMIM 276903) was first identified in 1992 to chromosome 11q13.5 (Kimberling et al., 1992, Smith et al., 1992) and it took only three more 21.5.1.3

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years to discover the molecular basis of this disease. When the Myo7a exon was discovered in the sh1 region homologous to the USH1B-containing region, mutations in the human gene were found as well (Weil et al., 1995). Since then, 81 mutations in the myosin VIIA gene have been discovered in USH1B (reviewed in Petit et al., 2001). Many of the mutations identified are localized to the 17 exons of the head/motor domain, although there are also mutations in the neck (three exons) and tail region (28 exons). The mutations are varied in nature, including nucleotide substitutions (missense, nonsense, and splicing mutations), small deletions and insertions, and rarer, larger deletions and rearrangements (one case for each) (summarized in Human Gene Mutation Database, http://archive.uwcm.ac.uk/ uwcm/mg/hgmd0.html).

DFNB2 and DFNA11 There are a number of cases where mutations in the same gene lead to both recessive and dominant deafness (http://dnalab-www.uia.ac.be/dnalab/hhh), and such is the case for myosin VIIA as well. Two non-syndromic deafness loci, DFNB2 (OMIM 600060) (Liu et al., 1997a, Weil et al., 1997), and DFNA11 (OMIM 601317) (Liu et al., 1997b), are associated with MYO7A mutations. DFNB2 is an autosomal recessive form of non-syndromic deafness, identified in Chinese and Tunisian families. The Tunisian family is a highly consanguineous one, with 22 affected members exhibiting profound hearing loss. The mutation in this family is a G1797A missense, changing a methionine to isoleucine at the end of exon 15, leading to a decrease in splicing efficiency. Affected DFNA11 family members from Japan have moderate progressive sensorineural hearing loss, with no evidence of RP, as revealed by ophthalmological testing (Tamagawa et al., 2002). Sequencing of DNA from affected DFNA11 family members revealed an in-frame 9-base pair (bp) deletion that results in the loss of three amino acids. This deletion occurs in the coiled-coiled region of the myosin VIIA tail (Fig. 21.2), and thus is predicted to prevent dimerization of myosin VII, leading to a dominant-negative effect. 21.5.1.4

21.5.2

Myosin VI

Myosin VI (OMIM 600970) was the sixth myosin to be discovered among the ‘unconventional’ myosins. The first myosin VI was identified in Drosophila, named 95F MHC (Kellerman and Miller, 1992). Since then, myosin VI has been discovered in C. elegans (Kelleher et al., 2000), pig (Hasson and Mooseker, 1994), bullfrog (Solc et al., 1994), chicken (Buss et al., 1998) and mouse (Avraham et al., 1995). Mutations in myosin VI are associated with NSHL in both humans and mice.

Structure, function, and expression of myosin VI The human myosin VI (MYO6) gene is composed of 32 coding exons and spans a genomic region of approximately 70 kb. The phenotype of the sesv allele clearly sug21.5.2.1


gests that there are additional upstream regulatory sequences to be identified in mice, and they may be present in human as well (see Section 21.5.2.2). The coding region of myosin VI produces a 4-kb cDNA, although by Northern analysis, transcripts of 6 and 8 kb are observed (Avraham et al., 1997). The human myosin VI protein is predicted to encode a 1263-amino acid protein, composed of a head/ motor domain (776 amino acids), a neck domain, and a tail that is made up of a coiled-coil tail (192 amino acids) and a globular region (232 amino acids) (Fig. 21.6). Myosin VI is ubiquitously expressed, with the highest expression in the kidney and brain (Avraham et al., 1997). In the inner ear, myosin VI is localized to the inner and outer hair cells of the cochlea and the vestibular organs (Avraham et al., 1995, Hasson et al., 1997) (Fig. 21.7). Myosin VI is localized in the cell body, and in particular, in the cuticular plate, the actin-rich structure underlying the stereocilia. Expression has not been observed in the stereocilia. In Drosophila, inhibition of myosin VI by antibodies demonstrates that this protein is required for proper organization of the syncitial blastoderm (Mermall and Miller, 1995). Porcine myosin VI was cloned in an effort to study myosins in the proximal tubule of the kidney, and was the source of the first mammalian myosin VI clone. In the kidney, myosin VI is expressed at the base of the brush border in proximal tubule cells (Biemesderfer et al., 2002). In the striped bass, two isoforms of myosin VI (FMVIA and FMVIB) were found in a retinal cDNA library (Breckler et al., 2000). In the retina, these two isoforms are expressed in the photoreceptors, horizontal cells and Muller cells in both fish and primate retinas.

Figure 21.6.

Myosin VI protein domain

structure.

Figure 21.7. Protein localization of myosin VI in guinea pig organ of Corti. (A) Section at level of the cuticular plate, the actin-rich region below the stereocilia, where myosin VI is enriched. (B) Section at the level of the cell bodies

of the inner (IHC) and outer hair cells (OHC). MVI, myosin VI. Adapted with permission from Hasson et al. (1997), by copyright permission of The Rockefeller University Press.

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Myosin VI has been studied both from the perspective of its inner ear pathology and as a protein involved in membrane traffic in polarized cells. Although myosin VI is ubiquitously expressed, its localization in different cell types has implicated this protein in receptor-mediated endocytosis. Myosin VI is localized to the Golgi complex and the leading, ruffling edge of fibroblasts (Buss et al., 1998). In the kidney, myosin VI is localized in the coated-pit region of the brush border and is associated with clathrin-associated proteins (Biemesderfer et al., 2002). Along with its role in Drosophila embryos (see Section 21.5.2.2), myosin VI appears to be a general membrane transport motor and have an essential role in membrane trafficking. Thus far, interacting proteins for myosin VI have been identified from brain and kidney libraries. In a search for proteins that interact with the glucose transporter binding protein GLUT1CBP, myosin VI was found to interact, suggesting a role for both in vesicle trafficking or a mechanism for linking GLUT1CBP to the actin cytoskeleton (Bunn et al., 1999). In experiments designed to find myosin VI-interacting proteins, DOC-2/DAB2 was identified. This Ras cascade signaling molecule interacts with myosin VI at the coiled-coil and globular tail region (Inoue et al., 2002, Morris et al., 2002). Thus myosin VI may transport DOC-2/DAB2 from the cell surface to its specific cellular targets. Myosin VI has several unique features. It has been shown to translocate toward the pointed (–) end of actin, contrary to the direction taken for all the other myosins characterized (Wells et al., 1999). A myosin VI construct was bound to actin, and using cryo-electron microscopy and image analysis, an ADP-mediated conformational change was observed in the domain distal to the motor in an opposite orientation to other myosins. This domain corresponds to the converter/lever arm, containing a unique 53-amino acid insertion. In addition, myosin VI has been shown to be a processive motor with an unusually large step size (Rock et al., 2001).

Snell’s waltzer mice and other models The Snell’s waltzer (sv) mouse is a recessive mutation that arose spontaneously at the Jackson Laboratory in the 1960s (Deol and Green, 1966). These mice are profoundly deaf and exhibit vestibular dysfunction, observed by circling. During a radiation screen at Oak Ridge National Laboratories, another sv allele, sesv, was created (Russell, 1971). A positional cloning endeavor led to the isolation of the svMyo6 gene, using the sesv allele to gain access to the sv locus (Avraham et al., 1995). Exon-trapping (a technique used to identify protein-encoding sequences in genomic clones; Church et al., 1994) led to the identification of an exon with 89 % homology to the porcine myosin VI gene. Subsequent analysis of the sv allele showed that there is a 2-kb genomic deletion in the mouse myosin VI gene, which leads to a 130-bp deletion in the coding region. As a consequence, a stop codon is formed in the neck region and no myosin VI protein is produced in any the sv tissues examined. In the sesv allele, a 2-cM inversion leads to the putative loss of upstream regulatory sequences. As a result, there is reduced expression of myosin VI in all sesv tissues examined. 21.5.2.2


Snell’s waltzer mice never appear to hear and sound stimulation experiments at 30 days after birth reveal only minimal reponses of a summating potential (SP) at 125 dB SPL, a high intensity stimulus that is similar to the noise of a jet engine. There was no compound action potential (CAP) or cochlear microphonics (CM) response at any frequency or intensity of stimulus used (Self et al., 1999). Although vestibular function has not been measured, the sv mice circle continuously in either direction. The hair cells of sv mice show a unique pattern due to the loss of myosin VI, as observed by scanning electron microscopy (SEM) (Self et al., 1999). Already by 3 days after birth, a disorganized pattern is apparent that includes fusion of stereocilia with one another (Fig. 21.8). By 20 days after birth, giant stereocilia are seen at the top of the hair cells. Transmission EM has demonstrated that the fusion begins at the base of the stereocilia simultaneously, hair cells degenerate, and by 6 weeks of age, there are no hair cells left and the entire organ of Corti is lost (Avraham et al., 1995, Self et al., 1999). Although endocytosis was shown to function normally in these hair cells, technical limitations may have prevented abnormal processes from being detected. Studies in lower organisms also implicate myosin VI in membrane trafficking. In Drosophila, myosin VI is known to be crucial for proper blastoderm formation (Mermall and Miller, 1995), and in both C. elegans and Drosophila, for proper spermatogenesis. C. elegans deletion mutants do not properly segregate cell compo-

Scanning electron microscopy of Snell’s waltzer (sv) inner and outer hair cells of (A) newborn, (B) 3, (C) 7, and (D) 12-day old mutant mice. Note fusion of stereocilia already

Figure 21.8.

beginning at three days after birth and general disorganization of cells. Adapted with permission from Self et al. (1999), with permission from Elsevier Science.

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nents, resulting in aberrant spermatogenesis. Mitochondria, endoplasmic reticulum/Golgi-derived fibrous-body membranous organelle complexes, actin filaments, and microtubules are not properly transported to or from spermatids (Hicks et al., 1999, Kelleher et al., 2000). During fertilization, each spermatid must go through a process called ‘individualization’, whereby it is enclosed in its own membrane. Two Drosophila mutants are known to have defects in spermatid individualization. Thus the 95F motor is required for transport and reorganization of membranes in the individualization complex within developing spermatids (Hicks et al., 1999).

DFNA22 Efforts to find a human myosin VI mutation failed for several years, despite this gene’s involvement in mouse deafness (Ahituv et al., 2000). However, in 2001, a genome scan on DNA derived from an Italian family with progressive dominant NSHL revealed linkage to chromosome 6q13, the region containing the human MYO6 gene (Melchionda et al., 2001). A missense mutation was discovered in exon 12, corresponding to the head/motor domain, between the ATP-binding and actin-binding sites (Fig. 21.6). The cysteine to tyrosine amino acid change may destablilize the protein, and thus compromise myosin VI function. This 21.5.2.3

A ribbon respresentation of a three-dimensional model of the human myosin VI motor domain, showing the mutated human myosin VI cysteine residue as a ball in the middle. The model was built using the automatic homology modeling facility, Swiss Model (http://www.expasy.ch/swissmod/ SWISS-MODEL.html) and the three-dimensional structures of several myosin motor domains. Adapted with permission from Melchionda et al. (2001), from the University of Chicago.

Figure 21.9.


cysteine is conserved in myosin VI in human, mouse, chicken, pig, striped bass and sea urchin, and in C. elegans and Drosophila, the cysteine is replaced with a similar hydrophilic residue, serine. In a comparison with 143 other myosins, no tyrosine residue appears at this position (see Myosin Motor Domain Sequence Alignment; http://www.mrc-lmb.cam.ac.uk/myosin/trees/txalign.html). It does not appear to form disulfide bonds, which are often formed by cysteines. Given the segregation of this MYO6 mutation with the affected individuals in this family, the previous association of myosin VI with deafness, and the conservation of the mutated residue, this missense mutation appears to lead to deafness in this human family. Although the protein structure of myosin VI has not been elucidated, the threedimensional structure of the head portion of myosin II is known (Rayment et al., 1993). Using homology modeling techniques, a model for myosin VI was made based on the amino acid sequence homology between these two myosins (Fig. 21.9). According to the model, the mutated cysteine that leads to deafness is partly buried in the protein core, suggesting that replacement of this small amino acid with a bulky tyrosine may destabilize the protein (Melchionda et al., 2001). 21.5.3

Myosin XVA

Myosin XVA (OMIM 602666) was discovered in a search for the gene associated with the mouse deaf mutant shaker 2 (sh2) (Probst et al., 1998) and the human NSHL locus DFNB3 (Wang et al., 1998).

Structure, function, and expression of myosin XVA The human (locus designation, MYO15A) and mouse myosin XVA genes both contain 66 exons and cover 71 and 60 kb of genomic DNA, respectively (Liang et al., 1999). A full-length isoform of myosin XVA is predicted to encode a 3530-amino acid protein with a molecular weight of 395 kDa. Like most other myosins, myosin XVA contains a motor domain, two IQ motifs, and a tail domain (Fig. 21.10). Myosin XVA also has a 1200-amino acid domain that precedes the motor at the N-terminal end. The tail domain of myosin XVA is composed of two MyTH4, two FERM-like domains, and an SH3 domain, similar to myosin VIIA (see Section 21.5.1.1). 21.5.3.1

Figure 21.10.

Myosin XVA protein domain structure.

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Figure 21.11. Myosin XVa is expressed in the inner (IHC) and outer hair cells (OHC) of the mouse. Myosin XVa is detected by indirect immunofluorescence using myosin XVa antibodies

on whole-mount preparations of organ of Corti in (A) low magnification and (B) high magnification. Adapted with permission from Liang et al. (1999), with permission from Elsevier Science.

Myosin XV has a more restricted expression pattern than myosin VI. Northern and dot-blot analysis indicates that the human gene is expressed in fetal and adult brain, ovary, testis, kidney and pituitary gland. In the mouse inner ear, as demonstrated by in situ hybridization, Myo15A is expressed in the hair cells of the cochlea and the vestibular system, including the saccule, utricle, and cristea ampularis (Liang et al., 1999) (Fig. 21.11). Labeling with antibodies showed a more restricted pattern in the hair cells, namely in the cuticular plate (the actin-rich region in the apical portion of the cell) and stereocilia.

Shaker 2 The original sh2 allele arose in an X-ray irradiation screen (Dobrovolskaia-Zavasdkaia, 1928) and was mapped to mouse chromosome 11 (Snell and Law, 1939), to a region homologous to human chromosome 17. Like sh1 and sv, these mice are deaf and exhibit the characteristic circling behavior of waltzer mice. The stereocilia of sh2 hair cells have a unique pattern in that they are extremely short, although the overall array pattern of the hair cells appears normal (Probst et al., 1998; Fig. 21.12). The sh2 mouse was proposed as a model for the human DFNB3 locus due to the mapping of homologous genes in the sh2 and DFNB3 regions (Friedman et al., 21.5.3.2

Figure 21.12. Scanning electron microscopy of the surface of shaker2 (sh2) organ of Corti. Note shortened stereocilia of outer hair cells (OHC) and inner hair cells (IHC).


1995, Liang et al., 1999). To clone the gene, a physical map of the sh2 region was constructed. Since there were many compelling candidate genes in the region, an in vivo complementation approach was taken in order to reduce the number of candidates to screen. Bacterial artificial chromosomes (BAC) from the critical region were injected into fertilized eggs derived from sh2/sh2 matings. One of the transgenic progeny born contained one of the BACs integrated in its genome. This mouse, named Sebastian after Johann Sebastian Bach, did not circle and was hearing, unexpected from a shaker 2 homozygous cross. This suggested that the gene responsible for the sh2 phenotype was contained within this 140 kb BAC. Sequencing and subsequent examination of the data generated revealed a novel unconventional myosin, named myosin XV, since it was the 15th to be identified. Since the subsequent discovery of a myosin XV, pseudogene, the myosin XV deafness gene has been renamed XVA (Boger et al., 2001). Initially, a mutation was detected in one sh2 allele within the motor domain. A G p A transition converts a cysteine to tyrosine. Comparison with 82 other myosins revealed either a cysteine or conservative change to threonine or leucine at this site, demonstrating the conservation of this residue. This change is likely to prevent myosin XVA binding to actin.

DFNB3 The DFNB3 locus (OMIM 600316) was the third recessive locus to be identified when it was found in a significant proportion of villagers from Bengkala, Bali (Friedman et al., 1995). Deafness in this community has been documented for at least seven generations, and as a result, the villagers, both hearing and deaf, developed their own sign language for effective communication (Winata et al., 1995). Additional families were found in India and Pakistan (Liburd et al., 2001). Deafness was mapped to the centromeric portion of chromosome 17 to a 3-cM region. Once the shaker 2 gene was identified, primers based on homology to the mousepredicted exons were designed and used to amplify human genomic DNA. Three mutations were identified that segregate with deafness. These include an isoleucine to phenylalanine substitution (I892F) within the MyTH4 domain, an aspartic acid to tyrosine substitution within the same domain (N890Y), and a nonsense mutation in exon 39, predicted to result in a truncated or null protein. All three mutations compromise myosin XVA protein function, an essential component of the auditory pathway. Since then, three additional mutations have been found in myosin XVA in families with non-syndromic deafness. Smith Magenis syndrome (OMIM 182290) patients have an interstitial deletion [del(17)p11.2] that includes MYO15A, leading to brachycephaly, midface hypoplasia, prognathism, hoarse voice, speech delay with or without hearing loss, psychomotor and growth retardation, and behavioral problems. Recently, the remaining MYO15A allele in one family was found to contain a substitution of a well-conserved threonine downstream of the first MyTH4 domain, which may be responsible for the moderate severe hearing loss in this individual (Liburd et al., 2001). 21.5.3.3

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21.5.4

MYH9

MYH9 (OMIM 160775), also known as non-muscle myosin IIA (NMIIA), is a nonmuscle myosin heavy-chain gene that maps to human chromosome 22q13.1. This myosin is involved in a number of diseases, including autosomal dominant giantplatelet disorders May Hegglin anomaly, Sebastian syndrome, Fechtner syndrome and Epstein syndrome. Of these, Fechtner syndrome and Epstein patients exhibit hearing loss.

Structure, function, and expression of MYH9 MYH9 was the first mammalian non-muscle myosin heavy chain gene to be identified and at the time was shown to be expressed in fibroblasts, endothelial cells, and macrophages (Saez et al., 1990). The MYH9 gene is expressed in a large number of tissues, as revealed by its presence in cDNA libraries and expressed sequence tags (ESTs) derived from stomach, uterus, pancreas, testis, and more (UniGene Cluster Hs.146550; http://www.ncbi.nlm.nih.gov/UniGene). In the rat cochlea, MYH9 protein is expressed in the outer hair cells of the organ of Corti, the spiral ligament, and Reissner’s membrane. Homologs have been identified in mouse, rat, Arabidopsis, Drosophila, C. elegans and yeast. Class II myosins consist of a pair of heavy chains, a pair of light chains, and a pair of regulatory light chains that together form a hexameric myosin molecule (Sellars, 1999). MYH9 encodes an amino-terminal head domain that contains the actin and nucleotide binding sites and a carboxy-terminal coiled-coil a-helical rod structure, separated by an IQ motif. The 40 coding exons of MYH9 encompass 5886 bp of coding sequence. The protein is predicted to encode 1960 amino acids with a molecular weight of 226 kDa. 21.5.4.1

DFNA17 The DFNA17 locus (OMIM 603622) was identified on 22q12.2 22q13.3 when linkage analysis of a five-generation non-consanguinous American family was carried out (Lalwani et al., 1997). DFNA17 is inherited in an autosomal dominant manner. Affected members of this family suffer from progressive hearing loss, beginning with the high frequencies at the age of 10 years, and by the age of 30 years the condition manifests itself as moderate to severe HL. Histopathological analysis was performed on the temporal bones of a deceased male with profound hearing loss, demonstrating classical Scheibe degeneration or cochleosaccular degeneration. In this condition, there is a loss of hair cells and supporting cells in the cochlea and saccule (an organ of the vestibular apparatus), with an accompanying loss of neurons, as well as degeneration of the stria vascularis (Fig. 21.13). Sequence analysis of MYH9 revealed a G p A heterozygote mutation in exon 16, leading to a substitution of arginine by histidine in codon 705 (Lalwani et al., 2000). The R705H mutation occurs in a highly conserved 16-amino acid linker re21.5.4.2


Histopathological analysis of the temporal bones from a hearing individual (A) and a (B) deceased male with profound hearing loss, demonstrating classical Scheibe degen-

Figure 21.13.

eration or cochleosaccular degeneration. This phenotype is characteristic of DFNA17/MYH9 hearing loss. Adapted with permission from Lalwani et al. (1997).

gion that contains two free thiol groups SH1 and SH2. Since this region is believed to play a major role in myosin head conformational changes, the DFNA17 mutation may disrupt mechanical function of the motor domain. 21.5.5

Myosin IIIA

The myosin IIIA (gene locus, MYO3A) (OMIM 606808) was first identified in Drosophila (Montell and Rubin, 1988) and the horseshoe crab Limulus polyphemus (Battelle et al., 1998), where the gene is expressed exclusively in eye photoreceptor cells. Mutations in the Drosophila myosin IIIA, NINAC, are associated with abnormal photoreceptor electrophysiology. The human MYO3A gene was isolated with the intention of following its retinal expression (Dose and Burnside, 2000), with the expectation that this gene would be involved in a human eye disease. Therefore it was surprising that the first human mutations in MYO3A are associated with NSHL only (Walsh et al., 2002).

Structure, function, and expression of myosin IIIA The human myosin IIIA gene (locus designation, MYO3A) has an open reading frame of 4848 bp that codes for a predicted 1616-amino acid protein (Dose and Burnside, 2000). This alternatively spliced 35-exon gene spans 308 kb of genomic sequence. Myosin IIIA is composed of a motor domain, a neck and a tail domain (Fig. 21.14). There are two IQ motifs in the neck, and unique to this protein, an IQ motif in the tail. There is no evidence of a coiled-coil domain, suggesting that myosin IIIA functions as a monomeric myosin. The most unusual feature of myosin IIIA is the N-terminal kinase domain that most resembles a serine/threonine kinase. This kinase is also conserved in the Limulus and Drosophila myosin IIIA proteins, although they do not possess a similar tail motif. The mouse myosin IIIA 21.5.5.1

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Figure 21.14.

Myosin IIIA protein domain structure.

Figure 21.15. Myosin IIIa is expressed in the inner (IHC) and outer hair cells (OHC) of the mouse inner ear. mRNA in situ hybridization was performed to demonstrate Myo3a expression in a newborn mouse cochlea. (A) Entire cochlea showing hair cells, with line indicating orientation of cross sections. (B) Cross section

of cochlear duct showing specific labelling of IHC and OHC with Myo3a antisense probe. (C) Cross section of cochlear duct with Myo3a sense probe, indicating that expression in (B) is specific. Adapted with permission from Walsh et al. (2002), c 2002 National Academy of Sciences, U.S.A.

has recently been identified and shows high similarity to the human gene and protein (Walsh et al., 2002). While the Limulus and Drosophila myosin IIIA is expressed solely in the photoreceptor cells, human myosin IIIA has a slightly more broad expression. Myosin IIIA mRNA is found in human and monkey retina, as well as in human pancreas but not in the heart, brain, placenta, lung, liver, skeletal muscle or kidney (Dose and Burnside, 2000). In the mouse cochlea, MYO3A is expressed specifically in the inner and outer hair cells, as observed by in situ hybridization (Walsh et al., 2002; Fig. 21.15). Antibodies against the fish myosin IIIA demonstrate expression specifically in the photoreceptor cells of the retina (http://mcb.berkeley.edu/labs/ burnside/myosin.html). The human MYO3A gene was localized by radiation hybrid mapping to the human chromosome 10p11.2 region (Dose and Burnside, 2000). Although there are at least six retinal diseases that map to chromosome 10 (http:// www.sph.uth.tmc.edu/RetNet/), no retinal diseases map to the MYO3A region. Recently, an NSHL locus, DFNB30, was localized to a 13-cM region on chromosome 10, corresponding to 10 Mb (Walsh et al., 2002). Although many genes are present in this region, myosin IIIA appeared to be a particularly promising candidate due to the known involvement of myosins in the auditory pathway and the role of NINAC in the visual pathway.


Myosin IIIA mutants A mouse mutant with myosin IIIA mutations has not yet been produced. Thus far, the only animal model produced exhibits abnormalities in the visual transduction pathway. The ninaC Drosophila mutant was recovered in an electrophysiological screen to identify altered photoresponse (Montell and Rubin, 1988). The ninaC gene codes for two splice variants, p132 and p174. They are identical in their kinase and motor domain, but diverge after the first IQ motif. p132 expression is restricted to the photoreceptor cell bodies, while p174 is expressed in the rhabdomeres. Removal of the kinase domain of both p132 and p174 lead to altered ERGs, but no retinal degeneration. Removal of the myosin motor domain leads to both abnormal ERGs and retinal degeneration. Additional mutagenesis studies led to the conclusion that the motor domain is required for photoreceptor maintenance but not for initiation of the phototransduction cascade (Porter and Montell, 1993). 21.5.5.2

DFNB30 The DFNB30 locus is represented by an Israeli family originating in Mosul, Iraq (Walsh et al., 2002). The hearing loss in this family is progressive, beginning in the second or third decade of life. During the initial characterization of this family, it was unclear whether the inheritance pattern was recessive or dominant. Until now, all families with progressive hearing loss have shown a dominant pattern of inheritance. A genome scan revealed age-dependent recessive inheritance, although homozygosity was restricted to only a part of the critical region. Therefore it was not surprising to discover several mutations in the MYO3A gene, corresponding to the haplotypes segregating in the family. The first mutation is a splice mutation that leads to an unstable message, 732 (-2) A p G (Fig. 21.14). The second mutation is a splice mutation at 1777 (-12) G p A, which leads to the deletion of exon 18 and a stop at codon 668. The third mutation is a nonsense mutation, 3126 T p G, which leads to a stop at codon 1043. Most interesting, there is a genotype phenotype correlation seen in affected family members (Fig. 21.16). Individuals who were homozygous for the nonsense mutation had an earlier onset of hearing loss. Between the ages of 25 and 50 years, individuals homozygous for the nonsense mutation had significantly poorer hearing thresholds than those heterozgyote for the nonsense and either of the missense mutations. Although hearing impaired individuals have never complained of visual problems, detailed ophthamological tests will determine whether the retina is affected, as myosin IIIA is expressed in the retina and ninaC mutations lead to retinal degeneration (see Section 21.5.5.2). 21.5.5.3

531

532


Figure 21.16. A phenotype-genotype correlation is present in myosin IIIA mutations in the DFNB30 family. Three mutations segregated in the hearing impaired individuals from this family. Individuals with the nonsense (NN) mutation on both MYO3A alleles had a more severe hearing loss that began in the early 20s. Those who are compound heterozygote for a nonsense and either splice mutation (NS)

began to notice a change in their hearing in their 30s, and their hearing loss is not as severe. The figure shows the hearing thresholds in decibels (dB) at ages 25–39 years and 40–49 years, measured by pure-tone audiometry. Adapted with permission from Walsh et al. (2002), c 2002 National Academy of Sciences, U.S.A.

21.6

Concluding Remarks

Myosins associated with sensory defects perform many functions, ranging from structural, developmental and physiological roles. To date, five myosins are associated with hearing and visual defects in organisms ranging from Drosophila to humans. Additional myosins clearly play a role in the auditory and visual pathways, as they are expressed in cells of the cochlea and retina and perform essential functions in these pathways. For example, myosin Ic mediates auditory adaptation and is expressed in the stereocilia of mammalian hair cells (Holt et al., 2002). At least an additional 40 deafness loci and 35 RP loci have been mapped, so that additional myosins may be associated with these disorders. Thus far, we have learned a great deal about the function of the molecular motors associated with sensory defects in humans, and availability of mouse models will undoubtedly lead to yet more discoveries.


Acknowledgements

K. B. A. is supported by grants from the European Commission, the Israel Ministry of Health, the Israel Academy of Sciences, the Israel Ministry of Science, Culture and Sports, and the National Institutes of Health Fogarty International Center Grant. I would like to thank Yehoash Raphael, Lisa Beyer, Anil Lalwani, Teresa Nicolson, Thomas Friedman, Aziz El-Amraoui, and Christine Petit for contributing figures and the many collaborators who have worked on myosins with me over the years: Tama Hasson, Karen Steel, Corne Kros, Paolo Gasparini, Mary-Claire King, Nancy Jenkins and Neal Copeland. I would also like to thank my laboratory members who have worked on myosins: Tama Sobe, Nadav Ahituv, Orit BenDavid, and Sarah Vreugde.

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21 Molecular Motors in Sensory Defects Hicks, J. L., W. M. Deng, A. D. Rogat, K. G. Miller and M. Bownes. 1999. Class VI unconventional myosin is required for spermatogenesis in Drosophila. Mol. Biol. Cell 10: 4341 4353. Hmani, M., et al., 1999. A novel locus for Usher syndrome type II, USH2B, maps to chromosome 3 at p23-24.2. Eur. J. Hum. Genet. 7: 363 367. Holt, J. R., et al., 2002. A chemical-genetic strategy implicates myosin-1c in adaptation by hair cells. Cell 108: 371 381. Hoth, C. F., et al., 1993. Mutations in the paired domain of the human PAX3 gene cause Klein-Waardenburg syndrome (WS-III) as well as Waardenburg syndrome type I (WS-I). Am. J. Hum. Genet. 52: 455 462. Inoue, A., O. Sato, K. Homma and M. Ikebe. 2002. DOC-2/DAB2 is the binding partner of myosin VI. Biochem. Biophys. Res. Comm. 292: 300 307. Joensuu et al., 2001. Mutations in a novel gene with transmembrane domains underlie Usher syndrome type 3. Am. J. Hum. Genet. 69:.673 684. Kaplan, J., et al., 1992. A gene for Usher syndrome type I (USH1A) maps to chromosome 14q. Genomics 14: 979 987. Kelleher, J. F., et al., 2000. Myosin VI is required for asymmetric segregation of cellular components during C. elegans spermatogenesis. Curr. Biol. 10: 1489 1496. Kellerman, K. A. and K. G. Miller. 1992. An unconventional myosin heavy chain gene from Drosophila melanogaster. J. Cell Biol. 119: 823 834. Kelley, P. M., et al., 1998. Novel mutations in the connexin 26 gene (GJB2) that cause autosomal recessive (DFNB1) hearing loss. Am. J. Hum. Genet. 62: 792 799. Kelsell, D., et al., 1997. Connexin 26 mutations in hereditary non-syndromic sensorineural deafness. Nature 387: 80 83. Kimberling, W. J., et al., 1992. Linkage of Usher syndrome type I gene (USH1B) to the long arm of chromosome 11. Genomics 14: 988 994. Kimberling, W. J., et al., 1990. Localization of Usher syndrome type II to chromosome 1q. Genomics 7: 245 249. Kros, C. J., et al., 2002. Reduced climbing and increased slipping adaptation in cochlear hair cells of mice with Myo7a mutations. Nature Neurosci. 5: 41 47.

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Russell, L. B. 1971. Definition of functional units in a small chromosomal segment of the mouse and its use in interpreting the nature of radiation-induced mutations. Mutat. Res. 11: 107 123. Saez, C. G., J. C. Myers, T. B. Shows and L. A. Leinwand. 1990. Human nonmuscle myosin heavy chain mRNA: generation of diversity through alternative polyadenylylation. Proc. Natl Acad. Sci. USA 87: 1164 1168. Sankila, E. M., et al., 1995. Assignment of an Usher syndrome type III (USH3) gene to chromosome 3q. Hum. Mol. Genet. 4: 93 98. Schoem, S. R. and K. M. Grundfast (1998). Oculoauditory syndromes. In: Pediatric Otology and Neurotology. Edited by A. K. Lalwani and K. M. Grundfast. New York: Lippincott Raven Publishers, pp. 365 374. Seiler, C. and T. Nicolson. 1999. Defective calmodulin-dependent rapid apical endocytosis in zebrafish sensory hair cell mutants. J. Neurobiol. 41: 424 434. Self, T., et al., 1998. Shaker-1 mutations reveal roles for myosin VIIA in both development and function of cochlear hair cells. Development 125: 557 566. Self, T., et al., 1999. Role of myosin VI in the differentiation of cochlear hair cells. Dev. Biol. 214: 331 341. Sellars, J. R. (1999). Myosins. Oxford: Oxford University Press. Shahin, H., et al., 2002. Genetics of congenital deafness in the Palestinian population: multiple connexin 26 alleles with shared origins in the Middle East. Hum. Genet. 110: 284 289. Smith, R. J., et al., 1992. Localization of two genes for Usher syndrome type I to chromosome 11. Genomics 14: 995 1002. Snell, G. D. and L. W. Law. 1939. A linkage between shaker-2 and wavy-2 in the house mouse. J. Hered. 30: 447. Sobe, T., et al., 2000. The prevalence and expression of inherited connexin 26 mutations associated with nonsyndromic hearing loss in the Israeli population. Hum. Genet. 106: 50 57. Solc, C. F., B. H. Derfler, G. M. Duyk and D. P. Corey. 1994. Molecular cloning of myosins from the bullfrog saccular macula: a candidate for the hair cell adaptation motor. Auditory Neurosci. 1: 63 75. Tamagawa, Y., et al., 2002. Phenotype of DFNA11: a nonsyndromic hearing loss

21 Molecular Motors in Sensory Defects caused by a myosin VIIA mutation. Laryngoscope 112: 292 297. Tassabehji, M., V. E. Newton and A. P. Read. 1994. Waardenburg syndrome type 2 caused by mutations in the human microphthalmia (MITF) gene. Nature Genet. 8: 251 255. Titus, M. A. 1999. A class VII unconventional myosin is required for phagocytosis. Curr. Biol. 9: 1297 1303. Usher, C. H. 1914. On the inheritance of retinitis pigmentosa, with notes of cases. R. Lond. Ophthalmol. Hosp. Rep. 19: 130 236. Velichkova, M., et al., 2002. A human homologue of Drosophila kelch associates with myosin-VIIa in specialized adhesion junctions. Cell Motil. Cytoskelet. 51: 147 164. Verpy, E., et al., 2000. A defect in harmonin, a PDZ domain-containing protein expressed in the inner ear sensory hair cells, underlies Usher syndrome type 1C. Nature Genet. 26: 51 55. von Graefe, A. 1858. Vereinzelte Beobachtungen und Bemerkungen. Exceptionelle Verhalten des Gesichtsfeldes bei Pigmentenartung des Netzhaut. Albrecht Graefes Arch. Klin. Ophthalmol. 4: 250 253. Walsh, T., et al., 2002. From flies’ eyes to our ears: Mutations in a human class III myosin cause progressive nonsyndromic hearing loss DFNB30. Proc. Natl Acad. Sci. USA 99: 7518 7523

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Part 5 Beyond Biological Applications

22 Systematized Engineering of Biomotor-powered Hybrid Devices Jacob J. Schmidt and Carlo D. Montemagno

22.1

Introduction

There has been much recent interest and excitement surrounding molecular motors due to their incredible properties; these nanometer-sized molecular assemblies can move objects thousands of times their weight at high speeds while not losing a single atom in the process. The functioning of these motors is awe-inspiring and immediately raises the question: Can they be useful in artificial devices as well? Devices having these properties would promise performance far beyond that which can be attained with current technologies and possibly unattainable using any other technology. Unfortunately, however compelling these motors are, there is a huge gulf between appreciating their potential as components in devices and making the devices that incorporate them a reality. To bridge this gulf, a newborn engineering discipline has made its first steps towards providing solutions to the problems of incorporating molecular motors into useful devices. Early attempts to incorporate molecular motors into devices have suffered from low yields, crude control mechanisms, short lifetimes, dependence on chemical fuel, and high sensitivity to environmental conditions. These limitations are not endemic to these previous efforts specifically, but rather are common to all hybrid biomotor-powered systems made with current technology. Rather than focus on improvement of one aspect of one device, making an incremental advance, it is critical for the field to address the problems common to all engineered biomolecular devices. Finding solutions to these general problems will enable the creation of entire classes of devices and will allow the field to move forward rapidly, contributing useful solutions to outstanding technological problems. Ultimately, all engineered hybrid biomotor systems, regardless of composition, function, or implementation, have several fundamental characteristics in common: they must utilize energy; they must be controllable; they must be made in large quantities in desired locations; they must be capable of interacting with their external environment; and they must be capable of functioning predictably for periods of time sufficiently long to be useful. These are all aspects of core technologies, each

542

22.1 Introduction

of which represents a single building block that collectively form the foundation of a hybrid biomechanical system. The core technologies concern all aspects of hybrid device construction and operation: manufacture, assembly, energy, control, and function. Each of these core technologies is founded upon a number of basic science and engineering capabilities that collectively address universal technological challenges associated with the engineering of hybrid biomolecular devices. The core technologies comprise the following: x

x

x

x

x

Nanoscale-directed assembly. Diffusion-based assembly breaks down for large component parts, and some motile protein complexes require orientation in addition to deposition. Manufacture of hybrid organic/inorganic devices will require improvements on diffusive transport as well as other assembly techniques. Molecular energy transduction. Customization of the energy type and mechanism of delivery to the biomotors is a very powerful ability and expands the possible environments and applications as devices are freed from solely relying upon the chemical fuel ATP. Control mechanisms. Regulation of function and operation of biomotor-powered devices is crucial to fully realize their potential. Development of a library of controls directly acting on motor proteins, their fuel, the components interfaced with them, or a combination of all of these are necessary. Multimedia device construction. Biological and inorganic hybrid nano-machines combine two classes of materials with very different processing and handling requirements. Construction and combination of these components as well as development of novel fabrication techniques and materials are required to explore fully the range of possible devices. Engineering issues. The performance characteristics for assembled devices depend sensitively on loads, force tolerances and lifetimes of the bonds holding the structure together, as well as wear, corrosion or other adverse environmental effects. Extension of device lifetimes, enhancement of force tolerances, and improvement of device performance are conventional engineering pursuits that can also be fruitfully applied to biomotor-powered nanodevices.

Different combinations of these technological building blocks are required for different devices. For example, devices intended as autonomous drug delivery mechanisms may require a regenerating chemical energy source, chemical controls, and specialized packaging. Likewise, substrate-bound devices may require electrical power, electrical control and a direct interface with a rigid inorganic substrate. The design, construction, and operation of these devices are by no means isolated processes. Construction pathways inform component choices; low yields of functioning devices can help to improve construction processes, and so on. Even the core technologies themselves are interdependent. The input of each influences the device assembly strategy, which may in turn influence the device design, closing a feedback loop. If the manufacture of biomotor-powered hybrid devices is ever to progress beyond clever one-off ‘parlor trick’ types of devices, a generalized technological foundation must be established. This requires the development of basic scientific and

22 Systematized Engineering of Biomotor-powered Hybrid Devices

engineering elements that are focused on the establishment of clearly defined core technologies, which in turn support the realization of useful devices. As one would expect, the bricks and mortar associated with the engineering of hybrid biomolecular devices are extraordinarily diverse and require interdisciplinary integration to an unprecedented degree.

22.2

The Core Technologies 22.2.1

Nanoscale Directed Assembly

A critical step in the construction of hybrid nanodevices using biomolecular motors is to extract the motors from their natural environment and insert them into an artificial one while retaining their natural function. While careful scrutiny has been given to the natural function and the unique ways in which Nature has designed various motors, the assembly processes through which proteins can be manipulated and attached has received relatively little attention. As the individual elements are too small and numerous to be manipulated individually, bulk parallel device synthesis is necessary. Diffusion-based assembly breaks down for large component parts, and some motile protein complexes (e. g. myosin and actin) require orientation as well as deposition. Manufacture of hybrid organic/inorganic devices will require the use of and improvement on diffusive transport as well as other assembly techniques. The core technology of Nanoscale Directed Assembly concerns the mating of the biological and inorganic components of a hybrid device: the rational design, construction, and assembly. It represents the selection of bonding strategies; inorganic materials; fabrication techniques; assembly methods; coordination of the assembled parts; increasing the attachment efficiency; and exploration of new methods and practices for joining the components. The number of bonds necessary for device assembly may be large; an enlarged view of a device powered by the biomotor F1-ATPase is shown in Fig. 22.1 (Soong et al., 2000). The inorganic components were created using electron beam lithography, metal evaporation, wet etching, and reactive ion etching. The organic components were designed using recombinant DNA technology, modifying the motor so that a chemically accessible sulfhydryl group is present on the rotor and nickelbinding histidine tags are on the base, followed by standard biochemistry. The rotor is bonded to a nickel rod through two covalent bonds, two ligand/receptor bonds, and His tags. Once the bonding strategy for the device is chosen, the parts must be assembled. Biomolecular hybrid devices produced by self-assembly alone rely on diffusion of the component parts, a process that works exceptionally well for biological and chemical systems. However, when certain component parts are relatively few in number, have small diffusion constants, or the solution volume is large, the time necessary to construct the devices becomes prohibitively great. Further, diffu-

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22.2 The Core Technologies Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His Cys-His

Biotin Streptavidin Biotin

Ni rod Cys-GGSGGS-6xHis Biotin Streptavidin Biotin Cys on γ subunit β subunit-10xHis Ni cap SiO2 post

Figure 22.1.

Enlarged depiction of a hybrid nanodevice highlighting each connecting bond.

sion constants decrease as molecules leave the bulk solution and approach surfaces (Schuster et al., 2000). In addition, it may be desirable to attach proteins to certain areas while preventing them from attaching to others. Finally, some devices may function only when their incorporated proteins are oriented properly. For these reasons, unassisted diffusion-based self-assembly will not suffice for devices with parts having poor diffusion characteristics or requiring subsequent manipulation. It is not enough that some proteins collide with their intended targets they must be oriented and bonded specifically to them and no other. Simple diffusion processes must be augmented and issues of orientation addressed; this should increase device yield, allow for structures made from a larger variety of components, and enable more complex devices. Although collision rates can be improved through increasing the number of particles or decreasing the assembly chamber size, for rare parts or unusual device geometries, this will be insufficient. One possible approach is to use electrophoresis. Although this is a widespread chromatographic technique for sizebased separation of DNA and proteins, it has also been used to immobilize DNA (Edman et al., 1997) and microscopic fabricated components (Edman et al., 2000) at specific locations on fabricated substrates. For uncharged proteins and fabricated components in ionic buffers, AC dielectrophoresis (DEP) can be successfully substituted for DC electrophoresis. The substrate and electrode geometries can be computationally simulated (Hughes et al., 1996) and, once selected, they can be constructed using electron beam lithography and semiconductor processing


Figure 22.2.

Scanning electron micrograph of quadrupole electrodes for dielectrophoretic trapping of proteins and particles on a central post (diameter I 50 nm). Scale bar is 500 nm.

techniques (Fig. 22.2). DEP has been used to trap protein structures like the 280-nm long tobacco mosaic virus and the 250-nm diameter Herpes simplex virus (Morgan et al., 1999), as well as latex spheres down to 14 nm in diameter (Muller et al., 1996) in well-defined surface locations. In addition, traveling-wave dielectrophoresis (TWDEP) can be used to move trapped particles between phased electrodes, enabling controllable collection and manipulation of particles (Hughes et al., 1996). Enhanced docking rates may be thus be achieved by immobilizing the proteins in specific locations using DEP and controllably introducing other components to the proteins with TWDEP, repeating the process until the devices are fully assembled. Assembled devices can be removed using TWDEP and the process repeated. Protein orientation is essential to properly assemble hybrid devices made from molecular motors and other proteins. While the spatial asymmetry existing at the boundary between a substrate and the surrounding solution can be used to orient the proteins vertically (cf. the histidine tags on the base of the motor in Fig. 22.1), orientation within the plane is more challenging. Orientation should be distinguished from patterned deposition. Through patterning of electrodes or chemical functionalities on the surface, it is relatively straightforward to deposit proteins in selected regions using either specific or non-specific binding. However, depositing a microtubule, for example, in a specific location is not sufficient to fully direct the motion of the kinesin along it, as the kinesin travels only toward the plus end. Therefore, the orientation of the plus end of the microtubule must also be directed, which is more complicated. Orientation has been achieved through a variety of methods: the substrate may be engineered directly and the proteins orient themselves through their motion on the substrate (Hiratsuka et al., 2001); they may be oriented through their motion and fluid flow (Bohm et al., 2001, Stracke et al., 2000); or one end is tethered to the surfaces and alignment

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is achieved through fluid flow (Limberis et al., 2001). Additional possibilities may center on plus- and minus-end capping proteins which themselves may be engineered to attach in specific locations. In addition to hybrid assemblies formed in solution, directed assembly is also applicable to the manufacture of membrane-bound protein devices. The same calculation, simulation, and experimental techniques used to attract and dock proteins with fabricated structures in solution can be used to orient proteins in membranes, crucial for the proper functioning of transport proteins (as discussed in the Section 22.2.2). Membrane proteins are somewhat more straightforward to orient; due to their solubility in the lipid bilayer, they only have two stable configurations, parallel and anti-parallel to the membrane normal. Most membrane proteins have a transport function and thus are asymmetric. This asymmetry can be probed and exploited for orientation using a variety of methods such as electrostatic interactions (He et al., 1998) or electric and magnetic fields (Der et al., 1995). 22.2.2

Molecular Energy Transduction

The vast majority of molecular motors known operate using the ubiquitous biochemical fuel adenosine triphosphate (ATP). This distinguishes them from conventional MEMS devices, which are powered by electricity. Diffusion-delivered chemical fuel allows arbitrary device motion, as it is not associated with a substrate or localized power source. However, in other circumstances, such as immobilized surface-bound devices or devices with electronic interfaces, electrical power is the ideal choice. Incorporation of biochemically-powered motors in such devices places a big hurdle before device construction and application. For example, devices with closed packaging would need to have their ATP supplies continuously refreshed. Conversely, other forms of energy utilization have weaknesses as well; electronic medical monitors implanted within tissue are limited by battery capacity and lifetime. The ability to specify the energy type and mechanism of delivery to the biomotors is very powerful and will greatly expand the possible environments and applications. Although this is a radical idea from a MEMS perspective, it is commonplace in biological systems. Different enzymes transducing different forms of energy help organisms from bacteria to mammals best utilize that which their environments offer. Proteins capable of converting energy between different forms (optical to electro-osmotic, electrical to chemical, etc.) are also able to function in tandem (e. g. conversion of optical energy to electro-osmotic energy and electro-osmotic energy to chemical energy result in optical to chemical energy conversion). The aim of the core technology of energy transduction is simply to replicate these natural energy conversion chains. Although there are large numbers of proteins that perform these functions naturally, three serve as good examples to illustrate the principles of this core technology (pictured in Fig. 22.3): ATP synthase, bacteriorhodopsin (BR), and cytochrome oxidase (COX). ATP synthase is a ubiquitous enzyme that synthesizes ATP from ADP and Pi in the presence of a proton gradient. Bacteriorhodopsin


, e se as ha id n t Ox Sy e P om AT c h r to

Natural energy transduction processes.

Figure 22.3.

Optical Energy

Cy

Ba c AT ter P ior Sy ho nt do ha ps se in ,

Chemical Energy (ATP)

Bacteriorhodopsin , Cytochrome Oxidase

Electrical Energy

is a natural optically active proton pump that transports protons across the bacterial membrane upon absorption of a photon of green light. As protons are pumped out of the cell, a gradient of charge and pH forms across the cell membrane, creating an electro-osmotic potential which enables ATP synthase to produce ATP. COX is an electron- and proton-transporting protein, the final enzyme through which respiration occurs. In respiration, high-energy electrons are used to transport protons into the mitochondrial space, creating an electrochemical proton gradient used by ATP synthase to generate ATP. BR and COX are very similar in their resultant production of a proton gradient: that of BR is enacted by light, that of COX by electronic energy. Indeed, both BR and COX are used in Halobacterium salinarium and for the same purpose: BR is used when there is not enough oxygen for respiration and COX to be useful to the organism. The possibility of generating renewable sources of ATP to power bionanomachines is very attractive. The natural biological ATP-regeneration system of BR and ATP synthase has been replicated in the laboratory (Pitard et al., 1996, Richard and Graber, 1992, Richard et al., 1995) through the insertion of BR and ATP synthase in a lipid vesicle. Substitution of an artificial photosynthetic complex for BR in this ATP-regeneration system has also been successfully demonstrated (Steinberg-Yfrach et al., 1998), an achievement that may allow the capture and conversion of wavelengths of light outside the sensitivity of BR and circumvent limitations inherent in the use of proteins. Hydrolysis of ATP within these vesicles is a source of ADP and Pi, which can be used to make more ATP through the action of BR (or the synthetic optical proton pump) and ATP synthase. Incorporation of an ATP-dependent biomotor within this lipid vesicle would make a closed opticallypowered system capable of operation indefinitely (see Fig. 22.4). Because these fuel-generating systems are self-contained, they can be added as a modular fuel supply component to a device. The high efficiency of biological proteins also enables reversible operation. Partial backward electron flow in COX was observed in mitochondria following a reverse proton electro-osmotic pressure created by the addition of ATP (Wikstrom,

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Figure 22.4. Depiction of hybrid F1-ATPase nanodevices fueled by a light-powered BR/ATP synthase ATP-regeneration system.

1981). The prospect of enzyme reversibility allows new energy transduction pathways: operation of BR and COX in reverse results in a light-powered electrical system, a biosolar cell. Similarly, ATP synthase combined with COX results in an ATPpowered electrical system, useful for electrical devices implanted in vivo. Additional energy-transducing proteins may further expand the palette available for device design. Because BR, ATP synthase, and COX have a proton gradient in common, each can be used with the other in a ‘plug-and-play’ type of architecture: BR and ATP synthase together make the light-powered ATP production system mentioned previously. COX and ATP synthase compose the electron-powered ATP production system found naturally. This reversibility also allows combination of any two of the above energy conversion pathways in Fig. 22.3 to yield another; such combinations are commonly found in natural biological systems. The flexibility of energy sources is a particular advantage to the device designer, as the geometry and components of the system often dictate the forms of energy input to or extracted from the designed system. For example, a mobile device may be best powered by optical or chemical energy, while a device immobilized on a substrate may be best powered by electrical energy. Using the same components for each kind of device would be problematic but for the existence of the energy conversion pathways listed above. This ability to transform energy places the biological nanodevice at a distinct advantage to wholly inorganic systems, which are presently able to be powered using electricity only.


22.2.3

Control Mechanisms

Control of the hybrid devices at the molecular or device level is crucial. Motor activity must be directed for device manipulation, efficient use of fuel, and navigation. In natural biological systems such as the cell, thousands of proteins work interdependently, regulating themselves and others to ensure that the cell can adapt to changing external conditions, wasting no resources and ensuring that essential functions are carried out. These natural biological control mechanisms of proteins and protein complexes regulate their function in response to a variety of external stimuli: chemical, thermal and mechanical, among others. The control interface and mechanism must be considered for each device. The advantages and disadvantages of a single energy source discussed above also apply to control mechanisms. Chemical or optical controls might best serve mobile devices. Chemically-controlled motors could result in devices that respond to environmental variables such as pH, ionic concentration, and temperature. Immobile substrate-bound devices may be best served by electrical contacts integrated into the surface; this allows for differential control, in which some devices are operational while others are not. These contacts have the advantages of easy and configurable connections to the outside world and rapid conveyance of signals from the controller to the device. Each family of control mechanisms has unique capabilities that may suit some applications better than others, providing the needed flexibility in device design. There are several different strategies for the implementation of these controls into hybrid biomotor devices. In contrast to natural systems, primitive engineered hybrid nanosystems have contained only a few proteins removed from their natural environments and therefore also removed from their natural control mechanisms. As a consequence, much regulation of the function of motor proteins is often possible only through regulation of the fuel supply. Fuel-based control techniques are able only to activate motors, with the motors functioning until the fuel is exhausted. There are significant shortcomings with these approaches: other ATP-utilizing molecules will also be affected, there is no differential motor control, and fine control of operational parameters such as speed is not possible. Full control over hybrid nanodevices will be necessary for all but a few instances. Recent work in chemical control mechanisms has utilized light-activated ‘caged’ ATP a chemical derivative of ATP in a non-hydrolyzable form, which is able to release a defined amount of ATP related to the intensity, duration, and location of optical illumination (Dantzig et al., 1998). This technique has controlled the operation of kinesin motors for proscribed periods of time as they proceeded along microtubule tracks (Hess et al., 2001). Alternative mechanisms of fuel-based control include myosin mutants with engineered binding sites having sensitivities for analogs of ADP with bulky side-chains such as N6 -(2-methylbutyl) ADP, allowing differential control between specific motor types (Gillespie et al., 1999). Chemical control of the motors apart from fuel regulation may be effected through chemical modification of the motor itself. By engineering peptides which bind directly to each other, the motor can be stopped in the presence of

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the appropriate chemical reagents, as in reversible disulfide cross-linking of kinesin (Tomishige and Vale, 2000). Alternatively, enzyme activity may be altered by engineering metal ion-binding sites (e. g. the addition or substitution of histidine, aspartate, or lysine residues (Kasianowicz et al., 1999)) into appropriate regions of the protein. The presence of these metal-binding sites can fix adjacent regions of the protein to each other by introduction of metal ions into these binding sites. This induces strong bonding between the ions and adjacent residues. Since ions can bind several amino acids simultaneously, it is possible to attach two separate, independent regions to the ion in effect altering the local elasticity of these subunits by fixing them to each other. If movement of these regions is necessary for protein activity, this activity can be impaired or entirely halted. This type of control has been achieved in our laboratory through the engineering of zinc ion binding sites into the a and b subunits of F1-ATPase (H. Liu et al., 2002). Amino acids close together that exhibited large relative motion during catalysis were identified. After selecting those which were separate from the binding sites, residues of serine and glycine were substituted with histidine using sitedirected mutagenesis (Fig. 22.5). The locations of these binding pockets were chosen so that the zinc ions would bind both to the moving and stationary portions, in effect binding them to each other and immobilizing the protein. Bulk enzyme activity assays show that the mutant’s ATPase activity differs little from that of the wild-type. However, in the presence of Zn, the ATPase activity of the mutant is completely inhibited, while the wild-type is unaffected. Addition of a chelator to the reaction mixture pulls the Zn out of the binding sites and restores enzymatic activity. Single molecule studies confirm this behavior: observations of fluorescent actin filaments attached to the g subunit show rotation upon ATP addition, cessation of motion in the presence of Zn, and restoration of activity upon its removal. The wild-type shows no change. This activity is separate from the ATP binding, and so the ATP activity of unmodified ATPases as well as other ATP-dependent molecules is unaffected. While release of these chemical agents would affect all such sensitive systems (i. e. each motor is not individually addressable), the agents can freely diffuse throughout the environment and thus the devices are not constrained to a specific location to be controlled. Electrical control of protein activity is also possible. Externally applied electric fields can cause some proteins to deform sufficiently that their operation is impaired. Voltage-induced conformational changes are commonly-occurring natural phenomena. These shape changes can alter the catalytic or hydrolytic activity of the protein, slow it down, or affect its interactions with other proteins. The advantages of engineering this kind of response in motor protein-based hybrid devices draw upon the capability of MEMS and NEMS fabrication to place arrays of electrodes at arbitrary positions on a substrate surface. Electric fields produced by such electrodes would give the power to individually address single motors. Electrical control has been shown to exist naturally in the family of porins. Porins are pore proteins that transport water and other solutes across cell membranes, controlling concentration-dependent properties such as osmolarity and acidity. Some porins have been shown to open and close physically in response to an ap-


A model of F1-ATPase showing the engineered binding sites (red) with bound Zn ions (yellow). The motor activity stops when Zn is added and returns when it is removed. Figure 22.5.

plied voltage, a shape change that activates and deactivates the proteins’ transporting ability (Bainbridge et al., 1998, Durell et al., 1998, Muller and Engel, 1999). For example, the pore protein OmpF changes conformation at voltages i 140 mV and in certain ranges of pH gradients or ionic strengths. These shape changes are a result of charged or polar groups in the protein responding to an external electric field and, on a small scale, occur in every ligand receptor binding event. It may be possible to engineer similar responses on other proteins, using polar or charged residues to enhance the response of the protein to an applied electric field. Proteins not containing the necessary functional groups to respond to the applied fields can be engineered to contain the groups or to further tune their response, much in the same way as the chemical binding sites discussed above were created. Electrical control mechanisms have the promise of addressability, control of activity, and easy interfacing with external instrumentation. Motor activity can also be controlled without directly affecting the protein. Attachment of cargo presenting variable loads resulting from viscous, electric, or magnetic forces, will change the motor speed, as discussed above. Although this will not be possible in every case, it is a powerful and flexible method of motor control. Any moving part attached to the motor, if susceptible to an external field, may be used to increase controllably the drag force experienced by the motor. Work by Noji and coworkers have used this principle to study the rotation characteristics of F1-ATPase (Noji et al., 2001). In these experiments, magnetic beads were attached to F1-ATPase motors lying on a substrate while observed by optical microscopy. As the rotary shaft of the motor turned, the bead turned with it. An external magnetic field with controllable direction and magnitude was applied to this system, creating a counterforce opposing the rotation of the motor. Fields large enough to slow and

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stall the motor were used, with the result that the rotary energy potential of the motor was determined. Each type of control mechanism has unique advantages and disadvantages relative to the intended application. Chemical control mechanisms are not tied to any specific location, although all engineered motors will be affected. Electrical control is only possible on a membrane or substrate, although individual motor control is possible. Some proteins, as we have seen in OmpF, can be electrically and chemically controlled. This flexibility in choices allows the design and application requirements to dictate the components and not vice versa. Exploration and development of artificial protein binding sites, determination of optimal electrode configurations, electric field magnitudes and frequencies, and buffer compositions utilize computational calculations and simulations. These techniques are necessary for screening large numbers of variables and exploring regions of parameter space prohibitive from a cost and time perspective. Establishment of this core technology is a step toward a wider array of controllable hybrid bionanodevices. 22.2.4

Multimedia Device Construction

The two main pillars on which the emerging discipline of hybrid biomolecular engineering stand, nanofabrication and molecular biology, have become two of the fastest growing and most impressive fields of technology in recent years. Construction of machines made from biological and inorganic components combine two classes of materials with very different processing and handling requirements. To bridge this compatibility gap, special fabrication techniques and unconventional materials must be developed. Nanofabrication evolved directly or indirectly from the science and technology of semiconductor electronics, which by itself has driven fabrication limits toward the nanoscale range as demand increases for more complex and powerful electronic systems. Basic sciences have contributed new techniques and methods that have expanded the available list of possible fabrication processes. A huge demand for materials other than silicon, the predominant substrate material in the semiconductor industry, has pushed equipment manufacturers and process engineers towards development of new process strategies, radically departing from their original semiconductor roots. Many fabrication techniques that are of little or no interest to semiconductor researchers (due to contamination and electronic reliability issues) can be revisited from a biotechnology perspective. Nanoscale structures intended for hybrid integration have to be physically and chemically compatible with biological molecules. Biocompatibility dictates the choice of materials, which also must be integratable at the nanoscale level while preserving their properties. Factors such as microstructure (grain size, distribution, and orientation, contaminants, stress, etc.) and the presence of native oxides can drastically change the suitability of some materials for integration. Mechanical properties of the fabricated constituent components, such as spring constants and drag forces, have to be taken into account at the design stage.


Besides developing inorganic fabrication processes and molecular biology protocols for engineering of biomolecular motors, interfacial protocols must be developed that will allow seamless merging of the nanofabrication and biological process flows. In general, the fabrication environment is not amenable to biological materials and molecular biology procedures and materials are detrimental to semiconductor fabrication equipment. As a result the final integration is often performed in isolation of each of the originating environments and the protocols used must conciliate the conflicting requirements of each. A good example is the choice of liquid media. Deionized water is the universal solvent in nanofabrication but will denature biological molecules. Conversely, phosphate buffer is a standard biochemical medium but is problematic in the fabrication laboratory (it contains sodium and potassium, the worst contaminants in cleanrooms). Therefore one has to render the nanofabricated parts amenable to biological use while preserving their characteristics, such as preventing the oxidation of metal film surfaces. These protocols are unique and so far have been developed on a case-by-case basis; at present there is no such thing as a well-established arsenal of integration procedures. Once the protocols and process flows have been worked out for a specific biomotor-powered hybrid device, the device can be constructed utilizing the different strengths of each fabrication process: inorganic lithography excels at the precise placement of prescribed features on two-dimensional surfaces. Biological synthesis is able to make three-dimensional mechanical structures nanometers in size which diffuse and self-assemble. Combination of these two enables self-assembling biological nanostructures located at arbitrary desired positions on substrate surfaces. Our group has successfully demonstrated the integration of the F1-ATPase motor with nanofabricated parts, shown schematically in Fig. 22.1 (Soong et al., 2000). The mutagenesis, bacterial production, and biochemical modification of the motor were all standard established techniques. However, considerable innovation and effort was required to tailor nanofabrication techniques and materials to build compatible structures and combine the dissimilar worlds of nanofabrication and molecular biology for the final system integration. The key linkage joining the biological and inorganic components was (His)6 -Nickel. Such protein/inorganic interfaces are virtually unknown, and the work of Belcher and Brown described below in Section 22.2.5 are important initial steps in expanding and establishing a library of linkers between proteins and inorganic materials. Processing of the inorganic components to interface or bond with proteins or biomolecules often includes chemical functionalization of the surfaces. The most common surfaces used are silicon, glass, and gold. Covalent attachment of self-assembled monolayers to these surfaces effectively changes the chemical group displayed (Allara, 1996). Gold binds specifically and strongly to sulfhydryl groups. The functional group displayed by silicon, glass, and most other oxidized surfaces is the -OH group, which is relatively unreactive toward most proteins and biomolecules. Change of the surface group is then easily brought about through the use of reactive heterobifunctional molecules. These molecules are thiols and silanes, for gold and oxide surface modification respectively, and the functional group on the oppo-

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site end of the molecule determines the final functionality of the surface. In this way, the glass surface may be transformed into an amine surface, a thiol surface, etc. Once the surface functionality is as desired, the proteins can be joined directly to the presented group, or linkers can be employed to target specific functional groups on the protein or increase reaction yield (Hermanson, 1996). Changing the surface chemistry can also alter non-specific protein adhesion properties. For example, use of the silane diethylenetriaminopropyltrimethoxysilane attaches six amino groups to each -OH group bonded to a silane. At physiological pH, this can present a rather large positive charge and thus attract or repel charged proteins, and has been used to attract negatively charged microtubules to specific regions on a fabricated substrate (Turner et al., 1995). Membrane-bound proteins incorporated into engineered devices must be incorporated into biocompatible polymers. Use of polymer membranes is desirable for the following reasons: they have a longer lifetime than lipid membranes, they are more rugged, and properties such as electronic and ionic conductivity and permeability can be tailored to suit each application. The interiors of these membranes must be hydrophobic and elastic so that the natural protein environment can be simulated as closely as possible. A large number of materials have demonstrated biocompatibility and can incorporate proteins while maintaining their functionality, such as gelatin, hydrated polymers, sol-gels, poly(vinyl alcohol), and poly(acrylamide) (Birge et al., 1999). For devices in which the protein environment must replicate the two-dimensional nature of natural membranes, amphiphilic polymers can used, such as the triblock copolymer poly(2-methyloxazoline)-poly(dimethylsiloxane)-poly(2-methyloxazoline) (PMOXA-PDMS-PMOXA). The PMOXA blocks are hydrophilic, while the PDMS block is hydrophobic. The copolymer spontaneously forms a rich variety of phases from laminar to vesicular, similar to natural lipid systems. The porin OmpF has been shown to retain its natural functionality in this polymer (Nardin et al., 2001). 22.2.5

Engineering Issues

Assembled hybrid devices have performance characteristics which derive directly from the combination of the fabricated inorganic components with biological molecules. The interactions between the constituent components determine the device performance, and the linkages between them determine the device lifetimes and force tolerances. Environmental parameters such as temperature, pH, and ionic strength are also important, as chemical bonds are strongly dependent on these variables, and the proteins themselves may have optimal ranges of operation (e. g. non-thermophilic proteins function better at low temperatures). Although at a vastly smaller size, standard macroscopic engineering issues still apply at the micro-and nano- scales. Structural analysis, study and improvement of the chemical linkers attaching the organic to the inorganic components, and design considerations maximizing the lifetime, speed, or reliability of the constructed devices are necessary for each new application.


As an example, consider a device powered by a linear molecular motor; many factors go into the selection of the specific motor. The operating temperature may dictate a motor derived from a thermophilic organism. Further, the kind of motor, subtype (e. g. myosin II or myosin V) and the design of the device must be cognizant of stalling forces. If the load to be moved exceeds this value, multiple myosins in parallel must be employed. Large numbers of motors in parallel may require that the motors be non-processive. The force or torque output of the motors can also regulate component size. For example, F1-ATPase has been shown to rotate with a constant torque of Z 40 pN nm 1 (Yasuda et al., 2001). The drag torque experienced by the motor resulting from an object attached to its rotor is proportional to the rotational velocity. Therefore, an increase in the drag torque (resulting from an increase in the object’s size, for example) will decrease the rotation speed. As discussed in the Section 22.2.3, this can be used to regulate the motor speed. If a certain speed is required for a particular application, the motor or the load can be chosen to accommodate this need. After the choice of motor proteins has been made, the following steps occur: the components are designed and constructed, the construction strategy is determined and implemented, and the devices are assembled. Once assembled these devices operate until failure. The devices are held together by a number of biological/inorganic bonds; at each peptide/inorganic interface these bonds must be sufficiently strong to withstand thermal and viscous forces over the lifetime of the device. As with much larger devices, typical engineering pursuits include the identification of failure modes and effects of the environment on device function, lifetime, and the exploration of other operational defects and weak points. Just as the device assembly strategy is dictated by the type and location of each bond composing the device in Fig. 22.1, the choice of the bonds themselves is dictated by considerations of the strength and lifetime of these bonds. Although proteins can bond non-specifically to various inorganic surfaces through hydrophilic, hydrophobic, electrostatic, and van der Waals forces, specific bonds are crucial for rational device design and construction. Peptide bonds to inorganic surfaces are virtually unknown, but have significantly increased in number due to recent work discovering short peptide sequences which adhere specifically to gold and chromium (Brown, 1997), iron oxide (Brown, 1992), and various semiconductors (Si(100), InP(100), GaAs(100), GaAs(111)A, and GaAs(111)B) (Whaley et al., 2000). Techniques of tethering proteins to solid inorganic substrates have only begun to be exploited, and have already been used to assemble microscopic particles of different materials (Brown, 2001). Site-directed or cassette mutagenesis can be used to engineer these peptide sequences in desired locations on proteins. The metals and semiconductors forming the other component of the bond can be fabricated and patterned using optical and electron beam lithography, allowing spatial localization of the attached proteins. Device performance, yield and reliability can be estimated through measurements of bond strength using dynamic force spectroscopy. Dynamic force spectroscopy experiments can sensitively measure the sizes and strengths of various bonds between biological ligands and their receptors as well as peptide inorganic

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interfacial bonds (Merkel et al., 1999). These measurements can indicate ‘weak links’ in device construction and allow calculation of bond lifetimes (Evans, 2001), which are crucial in determining device performance and failure rates. These experiments can be performed using many of the same instruments and techniques used to measure the biophysical properties of the constituent motors. Optical tweezers, atomic force microscopy, and the biomembrane force probe are some of the apparatus capable of sensitively measuring force at the pN level and displacement at the sub-nm level necessary to determine bond strengths. These studies are essential to the development of mass-produced robust hybrid bionanomechanical devices.

22.3

The Core Technologies as a Whole

This assemblage of core technologies represents what we believe are the essential building blocks necessary for realizing truly useful hybrid nanobiomechanical devices. A parallel hierarchical technological architecture is not unique; such architectures are integral to both the electronics and the pharmaceutical industries. As in these industries, each core technology member is composed of a number of scientific and engineering elements that together establish a suite of enabling technologies for the design and construction of hybrid nanodevices. Development of the core technologies may enable the large-scale production of practical hybrid devices using molecular motors. Establishment of these technologies will also create a conceptual framework for device construction and, by simplifying complex systems into assemblies of the core technologies, give the workers in this field a common language. When selected and packaged to meet desired performance objectives, they facilitate the design of application-driven systems with minimal re-engineering. By defining and developing units of functionality we are certain that redundant ‘discoveries’ will be significantly reduced and the accessibility of the required technologies will be greatly enhanced so that the transition of promising concepts to reality will be significantly accelerated. In summary, the core technologies build on a foundation of fundamental science and engineering principles that will address the shortcomings of current state-ofthe-art of hybrid nanodevice construction. Consequently, the endpoint is not just the demonstration of revolutionary nanosystems but also the establishment of a technological foundation that enables these hybrid biomolecular devices to achieve their promise. Furthermore, new applications for hybrid devices may directly result from the core technologies as the capabilities of the technologies and the motors themselves are made plain. Until they are in place, efforts at device construction may continue to result in devices made in small numbers with limited functionality having little possibility of widespread use.


References Allara, D. L. 1996. Nanoscale structures engineered by molecular self-assembly of functionalized monolayers. In: Nanofabrication aAnd Biosystems: Integrating Materials Science, Engineering, and Biology. Edited by H. C. Hoch, L. W. Jelinski and H. G. Craighead. New York: Cambridge University Press. Bainbridge, G., I. Gokce, et al., (1998). Voltage gating is a fundamental feature of porin and toxin b-barrel membrane channels. FEBS Lett. 431: 305 308. Birge, R., N. Gillespie, et al. 1999. J. Phys. Chem. B 103: 10746 10766. Bohm, K. J., R. Stracke, et al. 2001. Motor protein-driven unidirectional transport of micrometer-sized cargoes across isopolar microtubule arrays. Nanotechnology 12: 238 244. Brown, S. 1992. Engineered iron oxide-adhesion mutants of the Escherichia coli phage l receptor. Proc. Natl Acad. Sci. 89: 8651 8655. Brown, S. 1997. Metal-recognition by repeating polypeptides. Nature Biotechnol. 15: 269 272. Brown, S. 2001. Protein-mediated particle assembly. Nanoletters 1: 391 394. Dantzig, J. A., H. Higuchi, et al., 1998. Studies of molecular motors using caged compounds. Methods Enzymol., Caged Comp. 291: 307 348. Der, A., R. Toth-Boconadi, et al., 1995. FEBS Lett. 377: 419. Durell, S. R., Y. Hao, et al., 1998. Structural models of the transmembrane region of voltage-gated and other K channels in open, closed, and inactivated conformations. J. Struct. Biol. 121: 263 284. Edman, C. F., D. E. Raymond, et al., 1997. Electric field directed nucleic acid hybridization on microchips. Nucl. Acids Res. 25: 4907 4914. Edman, C. F., R. B. Swint, et al., 2000. Electric field directed assembly of an InGaAs LED onto silicon circuitry. IEEE Photon. Technol. Lett. 12: 1198 1200. Evans, E. 2001. Probing the relation between force Lifetime and chemistry in single molecular bonds. Ann. Rev. Biophys. Biomol. Struct. 30: 105 128. Gillespie, P. G., S. K. H. Gillespie, et al., 1999. Engineering of the Myosin-Ib Nucleotidebinding Pocket to Create Sensitivity to N6 -

modified ADP Analogs. J. Biol. Chem. 274: 31373 31381. He, J., L. Samuelson, et al., 1998. Oriented Bacteriorhodopsin/Polycation Multilayers by Electrostatic Layer-by-Layer Assembly. Langmuir 14: 1674. Hermanson, G. 1996. Bioconjugate Techniques. San Diego: Academic Press. Hess, H., J. Clemmens, et al., 2001. Lightcontrolled molecular shuttles made from motor proteins carrying cargo on engineered surfaces. Nanoletters 1: 235 239. Hiratsuka, Y., T. Tada, et al., 2001. Controlling the direction of kinesin-driven microtubule movements along microlithographic tracks. Biophys. J. 81: 1555 1561. Hughes, M. P., R. Pethig, et al., 1996. Forces on particles in travelling electric fields: computer-aided simulations. J. Phys. D: Appl. Phys. 29: 474 482. Kasianowicz, J., D. Burden, et al., 1999. Genetically engineered metal ion binding sites on the outside of a channel’s transmembrane b-barrel. Biophys. J. 76: 837 845. Limberis, L., J. Magda, et al., 2001. Polarized alignment and surface immobilization of microtubules for kinesin-powered nanodevices. Nanoletters 1: 277 280. Liu, H., J. Schmidt, et al., 2002. Control of a biomolecular motor-powered nanodevice with an engineered chemical switch. Nature Materials 1: 173–177. Merkel, R., P. Nassoy, et al., 1999. Energy landscapes of receptor ligand bonds explored with dynamic force spectroscopy. Nature 397: 50 53. Morgan, H., M. P. Hughes, et al., 1999. Separation of submicron particles by dielectrophoresis. Biophys. J. 77: 516 525. Muller, D. and E. Engel 1999. Voltage and pHinduced channel closure of porin ompf visualized by atomic force microscopy. J. Mol. Biol. 285: 1347 1351. Muller, T., A. Gerardino, et al., 1996. Trapping of micrometre and sub-micrometre particles by high-frequency electric fields and hydrodynamic forces. J. Phys. D: Appl. Phys. 29: 340 349. Nardin, C., J. Widmer, et al., 2001. Amphiphilic block copolymer nanocontainers as bioreactors. Eur. Phys. J. E 4: 403 410.

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F0F1-ATP synthase in an artificial photosynthetic membrane. Nature 392: 479 481. Stracke, R., K. J. Bohm, et al., 2000. Physical and technical parameters determining the functioning of a kinesin-based cell-free motor system. Nanotechnology 11: 52 56. Tomishige, M. and R. D. Vale 2000. Controlling kinesin by reversible disulfide cross-linking: identifying the motility-producting conformational change. J. Cell Biol. 151: 1081 1092. Turner, D. C., C. Chang, et al., 1995. Selective adhesion of functional microtubules to patterned silane surfaces. Biophys. J. 69: 2782 2789. Whaley, S., D. English, et al., 2000. Selection of peptides with semiconductor binding specificity for directed nanocrystal assembly. Nature 405: 665-668. Wikstrom, M. 1981. Energy-Dependent Reversal of the Cytochrome Oxidase Reaction. Proc. Natl. Acad. Sci. USA 78: 4051 4054. Yasuda, R., H. Noji, et al., 2001. Resolution of distinct rotational substeps by submillisecond kinetic analysis of F1-ATPase. Nature 410: 898 904.

23 Synthetic Molecular Motors Richard A. van Delden, Matthijs K. J. ter Wiel, Nagatoshi Koumura and Ben L. Feringa

23.1

Introduction

The fascinating molecular motors ubiquitous in biological systems offer a great source of inspiration for the design of artificial motors and in the quest to achieve controlled movement at the molecular level. Synthetic chemists are involved in the challenging endeavor to exploit or mimic the properties of these intriguing molecules found in nature. Movement in biological systems can be divided into linear motion, as found in, for example, kinesin and myosin (Vale and Milligan, 2000) or RNA polymerase (Yin et al., 1995), and rotary motion found in bacterial flagella (Berg and Anderson, 1973) or the F1-ATPase motor (Boyer, 1998, Walker, 1998). Nature’s sophisticated examples of (supra)molecular motors and especially the stunning direct observation of rotation for F1-ATPase (Noji et al., 1997) are a great stimulus for the synthetic chemist trying to mimic these systems. The major objective in designing a molecular motor is to generate controlled motion which is different from Brownian motion. In a molecular motor consumption of energy should result in controlled translational or rotary motion. Such motion could eventually enable a device to perform mechanical work, a particularly challenging goal in the context of future nanotechnology (Whitesides and Love, 2001). So far, only relatively simple artificial synthetic motors have been constructed. Also in these cases, one can roughly distinguish between linear molecular motors showing (intra)molecular translation and rotary molecular motors showing (unidirectional) (intra)molecular rotation. Translational and rotary motors will be treated separately in this chapter.

23.2

Translational Synthetic Molecular Motors

The most prominent examples of controlled molecular movement in linear molecular motor systems involve molecular shuttles based on linear pseudo-rotaxane or

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23.2 Translational Synthetic Molecular Motors

rotaxane systems, which have been extensively studied and reviewed (Ballardini et al. 2001, Balzani et al., 2000, Collin et al., 2001, Pease et al., 2001, Schalley et al., 2001). For a detailed discussion on the various molecular shuttles that have been constructed, the reader is referred to different reviews and references therein (Raymo and Stoddart, 2001, Sauvage and Amendola, 2001). The basic principle is schematically depicted in Scheme 23.1. A macrocycle, the train, can travel along a molecular chain, the rail between two stations. The position of the train is dependent on the exact electronic nature of the two stations as well as the train itself.

Scheme 23.1

Schematic representation of a switchable [2]rotaxane.

The translocation of the macrocycle relative to the chain has been achieved either chemically or by redox or acid/base stimuli, but photochemical translocation is also achieved. A typical example is given in Scheme 23.2 (Bissell et al., 1994). In the chain of compound 1 two biaryl stations are present. An electron-poor macrocycle prefers the relatively electron-rich diamine substituted biaryl station in the neutral equilibrium state. Under the influence of trifluoroacetic acid, the two amine groups in the molecular chain are protonated (a) and the interaction between

Scheme 23.2

Acid/base-controlled translocation in a [2]rotaxane system 1.

23 Synthetic Molecular Motors

the macrocycle and this first station becomes repulsive. As a result, a translocation of the macrocycle from the first protonated station towards the second electronically neutral biaryl station takes place (b). After deprotonation (c) the system will relax back to the initial state upon reverse translocation of the macrocycle (d). In this motor system the movement of the macrocycle relative to the molecular chain is controlled by the location of the macrocycle in the initial state and driven by protonation. Protonation of the initial state will considerably increase the energy of this configuration and part of this energy is released by a translocation of the macrocycle. This process is driven by the repulsive interaction between the protonated station and the train and translocation results in a minimum energy state for the diprotonated [2]rotaxane system. Upon deprotonation this configuration is no longer the energetic minimum and in a slower step the macrocycle moves back to the initial location. This results again in the initial-minimum energy state for the unprotonated system. Redox-driven systems function via a very similar mechanism (as discussed in detail for the related catenane systems below). Rotaxanes based on cyclodextrins are a special class of rotaxanes. The ability of cyclodextrins to bind organic guest molecules has been exploited in the synthesis of a variety of systems. A molecular shuttle based on one of these systems 2 is depicted in Scheme 23.3 (Murakami et al., 1997). An a-cyclodextrin (CD) moiety serves as the train on a molecular chain containing a photosensitive azobenzene moiety, which adopts a trans or cis geometry. The cyclodextrin moiety prefers the

Scheme 23.3

Light-controllable shuttling of a cyclodextrin-based [2]rotaxane 2.

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23.2 Translational Synthetic Molecular Motors

trans azobenzene station. Upon irradiation (360 nm) a trans cis isomerization is induced thereby changing the geometry of the azobenzene station (a). As a result the cyclodextrin shuttles away from the azobenzene towards one of the two spacers (b). Upon subsequent 430 nm irradiation the azobenzene is switched back to its trans-state (c) and the cyclodextrin shuttles back to its original location (d). This process can very elegantly be followed by circular dichroism (CD) spectroscopy, where the induced circular dichroism of the achiral azobenzene moiety is highly indicative for the presence of the chiral cyclodextrin in the vicinity of the chromophore. Upon shuttling, this CD absorption decreases whereas the reverse shuttling leads to an increase of the CD signal. An elegant example of a translational motor that makes use of both redox and photon stimuli was recently reported (Brouwer et al., 2001). In the system 3 depicted in Scheme 23.4, the macrocycle and the molecular chain interact via hydrogen-bonding rather than electrostatic interactions. In the initial state there is a preference for the macrocyclic train for the succinamide station (left). Upon photoexcitation of the second, naphthalimide, station followed by an electron injection from an external electron-donor (D), (a) the macrocycle is translocated towards the naphthalimide station due to preferred electronic interactions (b). Upon reduction of the naphthalimide station (c) the macrocycle shuttles back to the original succinamide station (d). This shuttling process was fully elucidated by a variety of photophysical techniques giving time-dependent data on the shuttling process. At room temperature in acetonitrile, the photoinduced translocation of the macrocycle (steps a and b) proceeds in about 1 ms. After charge recombination (step c) the reverse translocation (step d) proceeds considerably slower in about 100 ms. The functioning of these molecular shuttles can be compared with the functioning of a macroscopic piston. Energy input results in a movement of the macrocycle to an energetically unfavored location and after the initial input is replaced by a second (opposite) stimulus, full relaxation to the initial state involves the reverse movement of the macrocycle. It is important to note that the developed catenane- and rotaxane-based systems are close to real application in nanotechnology i. e. as a molecular electronic switching element (Collier et al., 2000). Although they exhibit controlled movement, these systems function as molecular switches rather than molecular motors. Two closely related systems that have recently been reported and should be mentioned here, are based on pseudorotaxanes which can function as solid-state supramolecular machines (Chia et al., 2001). One of the systems is physically trapped in a rigid nanoporous optically-transparent matrix while the other is tethered onto the surface of a silica film, while preserving their ability to undergo translational motion. Applications in molecular transistors as part of electronic circuitry and as digital elements for nanocomputing are readily envisioned. A remarkable recent finding (Jiménez et al., 2000) involves the contraction and stretching of a linear rotaxane dimer, mimicking a natural muscle at work (Scheme 23.5) (Feringa, 2000). Analogous to real muscles this molecular assembly was designed in order to allow the two filaments to glide along one another resulting in an extended and a contracted situation.


Scheme 23.4

A photo-responsive, multiple H-bondassembled, molecular shuttle.

Scheme 23.5

Schematic representation of the molecular muscle at work.

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23.3 Synthetic Rotary Molecular Motors

The system consists of two intertwined rotaxane units, each containing both a macrocycle and a molecular chain. The macrocycle contains a bidentate ligand moiety, so called because it has two atoms for coordinating to a metal center. The molecular chain carries both a bidentate and a tridentate moiety (which has three points of coordination) at distinct locations (comparable to the two stations of the rotaxanes described above). The process is driven by the difference in coordination behavior of Cu(I) and Zn(II) ions. In the presence of copper ions, which prefer binding to two bidentate ligands (four-coordination), the molecule shows a stretched geometry. Upon replacing copper for zinc ions, which prefer binding to one bidentate and one tridentate ligand (five-coordination), the two intertwined rotaxanes move along each other leading to a contracted geometry. This process is fully reversible. The length of the full structure (approximately 8.3 nm in the stretched form) changes by about 27 %, which, purely coincidentally, is very similar to the change in length of natural muscles. Although this stretching contracting motion is fully established, a real muscle function remains to be demonstrated. The main question that remains is whether the system can function against an applied force.

23.3

Synthetic Rotary Molecular Motors

A key feature in any macroscopic motor is the consumption of energy in the process of controlling motion in order to perform mechanical work. If control of the direction of a full 360h rotary motion in a molecular type motor can be realized, a basic requirement for the construction of functioning molecular machines might be fulfilled. In some primitive examples, two rotational motions within a molecule are coupled due to steric hindrance (Akkerman, 1970, Akkerman and Coops, 1967). The concept of such so-called molecular propellers was further exploited to develop the first molecular gear (Cozzi et al., 1981) and later elaborated towards more advanced geared systems (Clayden and Pink, 1998, Stevens and Richards, 1997) and a molecular turnstile (Bedard and Moore, 1995). In these systems, no control over the speed or direction of rotation is exerted, however. Rather, these are examples of conformational interconversions that result from hindered rotation around a single bond. Catenanes are a class of compounds closely resembling rotaxanes (Scheme 23.6). These molecular systems consist of two interlocked rings. One of the rings, analogous to the macrocycle in the rotaxanes (vide supra), can be considered to be the train. When the other ring is functionalized with two distinct stations (0 and 1), repeated shuttling between the two stations will result in a rotation of the train around the second ring. Two important aspects have to be emphasized here. First, since the direction of movement in these (achiral) systems as in the rotaxane-based systems, is controlled only by the distinct starting point and finish of the translocation of the train, there is no unidirectional rotation in these systems. Continuous shuttling merely results


Schematic representation of a catenane-based molecular rotary motor.

Scheme 23.6

in an oscillation of the train between the two stations, which statistically involves full rotation in only 50 % of all cases. The second point to be made is that the movement in this molecular system, although cyclic in nature, should be considered a cyclic linear movement restrained by the geometry of the ring system rather than a true rotation. Because of this aspect catenanes only differ in their functioning from rotaxanes by their geometry. Therefore, again various types of stimuli, trains and stations, can be used and a large variety of rotaxane systems are known (Sauvage and DietrichBuchecker, 1999). One illustrative example, based on redox-stimuli is depicted in Scheme 23.7 (Cárdenas et al., 1996). The concept is closely related to the molecular muscle discussed above. Two rings bearing both a bidentate (phenanthroline) and a tridentate (terpyridine) complexation unit are intertwined in 4. In the initial state Cu(I)-ions are present which have already been shown to prefer binding to two bidentate moieties. Oxidation (step a) of the metal ions to Cu(II)-ions, which prefer binding to two tridentate moieties, results in rotation of both rings consecutively (steps b and c). Upon reduction (step d) of the Cu-ions to Cu(I) again, rotation of each of the rings occurs separately (step e and f) and the system returns to the initial state. A net full rotation (or cyclic translation) of both rings has taken place whereas the direction of rotation is merely a statistical distribution of oscillation and rotation.

Scheme 23.7

Redox-controllable rotary motion in a [2]catenate system 4.

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23.3 Synthetic Rotary Molecular Motors

In the catenane systems reported so far, induced rotation driven by chemical or photochemical means is possible. The next important objective in the development of true molecular motor systems is to control rotary motion. This implies both control of the speed of rotation i. e. to produce motors which are capable of functioning at more than one speed and control over the direction of rotation, which is essential for a true motor. Without directional control, rotary motion and oscillatory motion will always be in competition. Tashiro et al. (2000) have developed a bisporphyrinate double-decker complex, depicted in Scheme 23.8, in which the rotary speed can be controlled. In the complex a metal center (either cerium or zinconium) functioning as a ball-bearing is sandwiched between two porphyrin ligands (rotating rings) that can rotate relative to each other. By virtue of the bulky substituents on both porphyrin ligands the metal complexes are chiral and two mirror image conformations (rotamers) exist. The enantiomeric (mirror-image) complexes show rotation of one porphyrin ligand relative to the other in opposite directions. The change in chirality can be used to study the rotary motion after chiral separation of the two forms. The direction of the rotary motion is controlled by the chirality of the initial state.

Scheme 23.8 Schematic

representation of the rotation behavior of a chiral metal bisporphyrinate double-decker complex.

It should also be noted that rotation results in racemization of the system where any control of the direction of rotation is fully lost. It was shown for the cerium complex that rotation of the porphyrin ligand is accelerated by a factor larger than 300 upon reduction of the metal center. Analogously, for the zirconium complex the rotation is decelerated upon oxidation of the metal center. These effects are attributed to the change in distance between the porphyrin moieties when the oxidation state of the central metal is altered. In the first case, reduction of the cerium center increases the ion radius thereby increasing the distance between the two


porphyrin rings. The increased distance results in decreased steric hindrance for rotation and the rotary motion is thus accelerated. The oxidation of the metal center in the case of the zirconium complex has the opposite effect, hence the deceleration of the rotary motion. Although the use of these systems in true nanotechnological machinery remains to be demonstrated, the results show that the speed of rotation can be controlled in a single molecular motor system by redox stimuli. Perhaps of even greater significance is the control of the direction of rotation. Two types of molecular motors, which show unidirectional rotation, have been developed. The first type is driven by chemical energy and the second by light (Davies, 1999, Freemantle, 1999).

23.4

Chemically Driven Unidirectional Molecular Motor

Kelly et al. (1999, 2000) demonstrated that unidirectional rotary motion is possible on a molecular scale by sequential chemical conversions. The design of the system is based on efforts to develop a molecular ratchet (Kelly et al., 1997, 1998), where the rotation of a trypticene-based wheel is sterically hindered by a bend pawl (Scheme 23.9). When the trypticene wheel is functionalized with an amino-substituent and a [4]helicene, in which a pending alcohol functionality serves as a pawl, a unidirectional 120h rotation is possible. The system is fueled by phosgene and controlled by the intrinsic chirality of compound 5 (Kelly, 2001). This rotation involves five consecutive steps: (a) phosgene-fueled isocyanate formation, (b) slight rotation, (c) urethane formation, (d) rotation involving an energy barrier in an irreversible manner and (e) hydrolysis of the urethane bond, as depicted in Scheme 23.9.

Scheme 23.9

A chemically driven unidirectional motor 5 allowing 120h rotation.

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23.5 Light-driven Unidirectional Molecular Motors

A chemical reaction between the initial amine and phosgene results in a highly reactive isocyanate moiety. Due to random movement of one half of the molecule relative to the other this reactive moiety may come into the proximity of the hydroxy group in the other half of the molecule leading to a fast intramolecular reaction locking the energetically unfavored, sterically demanding conformation. The energy is then released by a controlled rotation about the molecular axis. Subsequent hydrolysis completes the chemically-driven unidirectional rotation over 120 h. Although this is the first and up to now the only example of a chemically-driven unidirectional molecular motor, there are some major limitations to this system. First of all, the rotary motion still only leads to one-third of a full unidirectional rotation. Second, two separate stimuli are needed to continue the process and even if extrapolation to full rotation was possible, then six stimuli would be needed, which cannot be introduced simultaneously due to their competing reactivity. A third drawback for any real application is the use of the extremely poisonous phosgene gas as a fuel, although these drawbacks might be overcome in the future. This system and the next one (vide infra) prove the principle that unidirectional rotation can be accomplished in a molecular system solely by controlling its chirality.

23.5

Light-driven Unidirectional Molecular Motors

In our group, sterically-overcrowded alkenes were shown to be able to function as chiroptical molecular switches (Feringa et al., 2000, 2001). In these systems molecular chirality can be controlled by light. A prominent example is shown in Scheme 23.10 (Jager et al., 1995). Due to steric hindrance around the central olefinic bond these types of alkenes are forced to adopt a helical structure and hence show intrinsic chirality. The chirality is denoted P for a positive helical structure and M for a negative helical structure. Upon photoinduced cis trans isomerization the helicity of the molecular switch is inverted. The two states of the bistable switch system show near mirror-image behavior yet are not true enantiomers, but rather pseudoenantiomers. The selectivity of these light-driven molecular switches is dependent on the differences in UV absorption of the respective cis and trans pseudoenantiomers. For 6 which is shown below in Scheme 23.10, the asymmetric donor acceptor substitution of the lower half of the molecule results in relatively large absorption differences leading to a highly selective molecular switch. In nhexane solution irradiation at 365 nm results in a photostationary state consisting of a 70 : 30 ratio of isomers in favor of the (P)-trans-isomer. Irradiation at 435 nm results in a second photostationary state consisting of a 90 : 10 ratio in favor of the (M)-cis-isomer. This process is fully reversible. The exact properties of the switch can readily be tuned by changing the substitution pattern of the molecular skeleton by synthetic modification. The use of these sterically-overcrowded alkenes as chiroptical molecular switches has been studied in detail. For example, it was demonstrated that, by employing


Scheme 23.10

A donor-acceptor substituted chiroptical molecular switch 6.

liquid crystals as host compounds, the macroscopic chirality of a liquid crystalline film can be controlled using chiroptical molecular switches as guest compounds (Huck et al., 1995). These photobistable systems might have potential in future nanotechnological applications such as optical data storage units or as true switching devices in optical data processing. The developed systems as such also show the basic features of unidirectional rotary motion. The actual switching process already involves a near 180 h-rotation around the central olefinic bond. This rotation itself is unidirectional since irradiation results in a preferred movement of one half of the molecule relative to the other. The direction is fully controlled by the chiral helical shape of the initial state of the molecule (and the wavelength of irradiation). It is evident that this does not comprise a full rotary motion but it shows the possibility of (at least partly) unidirectional rotation driven by light and controlled by the intrinsic chirality of the sterically-overcrowded alkene. Extending the rotation process to a full 360 h-rotation can be done thermally but results in racemization of the system and as a consequence control over the direction of rotation is lost. In order to induce full rotation in a sterically-overcrowded alkene an additional chiral influence is required (vide infra). One approach towards controlled molecular rotation in a sterically overcrowded alkene system involves the rotation around a single bond in photoswitchable molecule 7 modified with a biaryl type rotor (Scheme 23.11) (Schoevaars et al., 1997). Here, the sterically overcrowded alkene moiety, which already proved to be efficient in switching between P and M helical structures, was functionalized with a xylylbased rotor moiety.

Scheme 23.11

Controlled intramolecular rotation in a chiroptical molecular switch 7.

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570


Photoisomerization between the (M)-cis and (P)-trans forms should cause a distinct difference in rotation rate for the biaryl rotation because steric hindrance in the vicinity of the rotor is completely different for the two isomers. Dynamic NMR studies revealed barriers for the biaryl rotation of DG 79.5 and 82.5 kJ mol 1 for the cis- and trans-isomers, respectively. In contrast to expectation, but in agreement with semi-empirical calculations, the barrier for the trans-compound was higher than that for the cis-compound. The remarkable rotary behavior was attributed to distinct differences in the chiral conformations and steric effects associated with folding in the molecules. In particular, the methyl groups of the xylyl rotor meet severe steric hindrance from the CH2 groups of the upper half in the trans isomer, whereas the nearly planar naphthalene moiety, in case of the cis-isomer, simply bends away during the rotary process. This rotor system, however, suffers from a small difference in energy barriers and inefficient photoswitching. Furthermore, there is no control over the direction of rotation, a conditio sine qua non for a molecular motor, and the system can therefore better be described as a molecular gear. Employing another class of sterically-overcrowded alkenes and based on a concept closely related to the chiroptical molecular switches discussed above, a true light-driven unidirectionally rotating molecular motor was developed. Following extensive studies on the thermal and photochemical isomerization processes of biphenanthrylidenes (Harada et al., 1997a, Zijlstra et al., 1999), it was demonstrated that the intrinsic chirality associated with chiroptical molecular switches can be used to accomplish unidirectional rotary motion (Feringa, 2001). In this case light energy is used as a fuel since the process involves photoisomerization steps, similar to those discussed above. With the sterically-overcrowded alkene (3R,3lR)-(P,P)-trans-1,1l,2,2l,3,3l,4,4l-octahydro-3,3l-dimethyl-4,4l-biphenanthrylidene, 8 (Scheme 23.12), where the two methyl substituents adopt the energetically-favored axial orientation due to steric effects, it is possible to achieve unidirectional rotation by combining two energetically uphill, photochemically-induced isomerization steps with two energetically downhill thermal helix inversion steps. In this way, a full light-driven 360 h-rotation of one (rotor) half of the molecule relative to the other (stator) half in a unidirectional fashion is induced (Koumura et al., 1999). Starting from the energetically favorable (P,P)-trans state, with diaxial methyl substituents, a trans cis isomerization is induced by irradiation (i 280 nm). This results after a nearly 120 h-rotation of one half of the molecule relative to the other in the energetically highly unfavorable (M,M)-cis state. The instability is caused by the fact that this isomerization forces both methyl substituents to adopt an unfavorable equatorial orientation. At room temperature the (M,M)-cis state already rapidly releases its internal energy by helix inversion to form the (P,P)-cis isomer in which the methyl substituents again adopt the energetically favorable axial orientation. A second isomerization step using the same wavelength of light results in the energetically unfavorable (M,M)-trans state, with diequatorial methyl substituents. Energetically downhill helix inversion of (M,M)-trans after heating to 60 hC, results in the initial (P,P)-trans state completing a full unidirectional 360 h-rotation.


Scheme 23.12

Light-driven unidirectional molecular motor 8.

The control elements that govern the unidirectional rotation include the helicity of the overcrowded alkene, the absolute configuration at the stereogenic centers and the conformational flexibility of the rings in the vicinity the central olefinic bond. The driving forces in this rotary process are the photoinduced isomerizations. In both cases the methyl substituents are forced to adopt an energetically unfavorable equatorial orientation. The subsequent helix inversions release the energy in a unidirectional process to form the stable isomers with axial methyl groups again. The direction of this rotation is controlled solely by the configuration at the stereogenic centers. This first example of a unidirectional molecular motor driven by light allows controlled and repetitive 360h rotation. When the system is constantly fueled with photon energy (i 280 nm irradiation) under appropriate thermal conditions (i 60 hC) all four steps involved in the rotation process are simultaneously induced, resulting in a continuous rotation process of one half of the motor molecule relative to the other, as is shown schematically in Fig. 23.1. Recently, it was shown that upon rotation this molecular motor can induce changes in a cholesteric liquid crystalline (LC) environment resulting in full color control of a liquid crystal matrix (van Delden et al., 2002). These findings demonstrate that the light-fueled unidi-

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572


Schematic representation of the unidirectional rotation of the first light-driven molecular motor.

Figure 23.1.

rectional rotation of this molecular motor can perform work, in this case by inducing the reorganization of mesogenic molecules, which leads to an observable macroscopic effect (color change of an LC film). This conversion of controlled kinetic energy into work is an essential feature for further application of these systems as molecular motors in nanotechnological applications. Although with this system it was shown for the first time that full unidirectional rotation can be accomplished, there is still considerable room for improvement. The first challenge is to reduce the temperature requirements of the system where, although photon-energy is the driving force, heating to about 60 hC is necessary to continue the rotary motion due to the high energy barrier of the thermal inversion step of the less stable trans-isomer. A design problem immediately arises when possible improvements are envisioned. The system, as presented, has little opportunity for structural variation, which is a serious drawback not only in attempts to decrease the rotation barrier but also when other structural modifications of the system are envisioned, for example, covalent attachment to polymer systems or organization on surfaces. Therefore a so-called second-generation motor concept was developed. The first-generation motor is based on a biphenanthrylidene system, which consists of two identical halves, and effectively, the two identical halves in the molecule both perform the same function. An essential question in approaches towards a more suitable motor system is whether the presence of only one of these rotor parts would suffice to induce unidirectional rotation. The other half of the molecule, in accordance with the chiroptical molecular switches, could then be used for adjusting the molecular properties by synthetic modifications (Huck et al.,

Second-generation molecular motor: combining the structural features and properties of the first generation molecular motor with those of the chiroptical molecular switches.

Figure 23.2.


1995). The goal in the studies on this second-generation motor was to combine versatility of the design of our chiroptical molecular switches with the unique rotational behavior of the first-generation molecular motor. A single chiral 2-methyl2,3-dihydrothiopyran upper part would induce a similar rotary behavior. These design features are schematically depicted in Fig. 23.2. The first requirement to be fulfilled with this second-generation motor is to establish that a single stereogenic center does indeed suffice to induce light-driven unidirectional rotation. For the parent compound 9 the photochemical and thermal-induced isomerization steps are shown in Scheme 23.13 (Koumura et al., 2000). A strong preference for an axial conformation of the methyl group at the stereogenic center in both (M)-trans-9 and (M)-cis-9 was established. This stereochemical feature was also essential for the functioning of the first-generation motor.

First example of unidirectional rotation controlled by a single stereogenic center; the prototype of the second generation motor.

Scheme 23.13

For this new type of motor, with distinct upper and lower halves, starting from the energetically stable (2lR)-(M)-trans isomer, a trans to cis isomerization was induced upon irradiation with 365 nm light. This resulted in the corresponding energetically unstable (2lR)-(P)-cis isomer; a process completely analogous to the energetically uphill photoisomerization found for the first-generation molecular motor. On heating to 60 hC, the unstable cis form, (2lR)-(P)-cis, epimerizes to the stable (2lR)-(M)-cis form. A second energetically up-hill photoisomerization step yields the unstable

573

574


(2lR)-(P)-trans form which upon heating reverts to the stable (2lR)-(M)-trans initial state, completing a full 360h rotation in a counterclockwise sense. This prototype of the second-generation motor shows that unidirectional rotation can be controlled by a single stereogenic center. The second objective of this new generation of motors was to be able to tune the properties. The different structures of the two halves, the upper half being the rotor and the lower thioxanthene half, the so-called stator, allow synthetic modifications comparable to those in the chiroptical molecular switches. In the lower half of this second-generation molecular motor, functionalities can be introduced and in this way a number of distinct motors can be made. A first objective of the research was to decrease the energy barrier for the thermal helix inversion steps to allow fast rotation at room temperature. Extensive research has already been carried out on the chiroptical molecular switches to examine the effect of the nature of the heteroatoms in both upper and lower halves of the molecule on the racemization barrier i. e. the thermal helix inversion process. The main objective in the case of the switches is to prevent this racemization under normal condition, while for the motors the lowering of the barrier for helix inversion to increase the speed of the motor can be considered to be exactly the opposite objective. Based on the experience with molecular switches among others, the compounds depicted in Fig. 23.3 were synthesized. Due to the decreased dimensions of O and CH2 compared to S, helix inversion was expected to be facilitated (Koumura et al., 2002). Indeed, the Gibbs energy of activation (DG0) for the thermal steps (at room temperature in n-hexane solution) decreased with decreasing (hetero)atom size from 105.7 kJ mol 1 (10) to 100.6 kJ mol 1 (11) and 91.6 kJ mol 1 (12). Changing the sulfur atom for an oxygen atom in the lower half resulted in a decrease in the half-life at room temperature by a factor of about 8, whereas the corresponding change from a sulfur to a carbon atom in the upper half decreased the half-life by a factor of about 320. This change resulted in motor 12 with a half-life for thermal helix inversion of about 2400 s at room temperature, which implies a drastic acceleration of the rotary speed. These examples clearly illustrate the potential for functionalization and tuning of the properties for this second-generation system. Currently more advanced motors including a system driven by visible light and sophisticated geared systems, are being developed.

Second-generation motor: unidirectional rotation controlled by a single stereogenic center and structural modifications that allow control of rotary behavior.

Figure 23.3.


23.6

Conclusion and Prospects

The design and construction of molecular and supramolecular systems in which controlled translational and rotary motion is achieved, is without doubt one of the most ambitious endeavors in contemporary synthetic chemistry. By combining molecular beauty with elegant functions in rotaxanes, catenanes, helical-shaped alkenes and a trypticene-type propeller, the first primitive examples of linear and rotary motors are now a reality. Despite the fact that the field of molecular motors and nanomachines is still in its infancy, the developments are rapidly gaining momentum. As we discover that controlled motion at the molecular level is indeed possible and that various methods for precise (self-)assembly at the nanoscale level are becoming available, the diversity of functional molecular architectures appears to be unlimited. The coupling of linear or rotary motion to other functions is just one of the challenges ahead. The use of molecular motors to perform mechanical work has yet to be established and the organization of motors on surfaces and connection to nanoobjects or incorporation into multifunctional machines without interfering with the motor functions, is yet to be accomplished. ‘Fighting the Brownian motion’, so elegantly done by Nature’s molecular motors, is a tantalizing goal.

References Akkerman, O. S. 1970. Hindered rotation in diphenylmethane derivatives. Part III A comparison of NMR kinetic studies with the results of racemization measurement. Rec. Trav. Chim. Pays-Bas 89: 673 680. Akkerman, O. S. and J. Coops. 1967. Optical activity of symmetrical substituted acetic acids. Part 1. The stability against racemization of some ortho-alkyl-substituted diphenylacetic acids. Recl. Trav. Chim. Pays-Bas 86: 755 761. Ballardini, R., V. Balzani, A. Credi, M. T. Gandolfi and M. Venturi. 2001. Artificial molecular-level machines: Which energy to make them work? Acc. Chem. Res. 34: 445 455. Balzani, V., A. Credi, M. Raymo and J. F. Stoddart. 2000. Artificial molecular machines. Angew. Chem. Int. Ed. 39: 3349 3391. Bedard, T. C. and J. S. Moore. 1995. Design and synthesis of a ‘molecular turnstile’. J. Am. Chem. Soc. 117: 10662 10671. Berg, H. C. and R. A. Anderson. 1973. Bacteria swim by rotating their flagellar filaments. Nature 245: 380 382.

Bissell, R. A., E. Córdova, A. E. Kaifer and J. F. Stoddart. 1994. A chemically and electrochemically switchable molecular shuttle. Nature 369: 133 137. Boyer, P. D. 1998. Energy, life, and ATP. Angew. Chem. Int. Ed. 37: 2297 2307. Brouwer, A. M., C. Frochot, F. G. Gatti, D. A. Leigh, L. Mottier, F. Paolucci, S. Roffia and G. W. H. Wurpel. 2001. Photoinduction of fast, reversible translational motion in a hydrogen-bonded molecular shuttle. Science 291: 2124 2128. Cárdenas, D., A. Livoreil and J.-P. Sauvage. 1996. Redox control of the ring-gliding motion in a Cu-complexed catenane: A process involving three distinct geometries. J. Am. Chem. Soc. 118: 11980 11981. Chia, S., J. Cao, J. F. Stoddart and J. I. Zink. 2001. Working supramolecular machines trapped in glass and mounted on a film surface. Angew. Chem. Int. Ed. 40: 2447 2451. Clayden, J. and J. H. Pink, 1998. Concerted rotation in a tertiary aromatic amide: Towards a simple molecular gear. Angew. Chem. Int. Ed. 37: 1937 1939.

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Huck, N. P. M. and B. L. Feringa. 1995. Dualmode photoswitching of luminescence. J. Chem. Soc., Chem. Commun. 1095-1096. Huck, N. P. M., B. L. Feringa and H. van Doren. 1995. Chiroptical switching between liquid crystalline phases. J. Am. Chem. Soc. 117: 9929 9930. Jager, W. F., J. C. de Jong, B. de Lange, N. P. M. Huck, A. Meetsma and B. L. Feringa. 1995. A highly stereoselective optical switching process based on donor-acceptor-substituted dissymmetric alkenes. Angew. Chem. Int. Ed. Engl. 34: 348 350. Jiménez, M. C., C. Dietrich-Buchecker and J. P. Sauvage. 2000. Towards synthetic molecular muscles: Contraction and stretching of a linear rotaxane dimer. Angew. Chem. Int. Ed. 39: 3284 3487. Kelly, T. R. 2001. Progress toward a rationally designed molecular motor. Acc. Chem. Res. 34: 514 522. Kelly, T. R., J. P. Sestelo and I. Tellitu. 1998. New molecular devices: In search of a molecular ratchet. J. Org. Chem. 63: 3655 3665. Kelly, T. R., H. de Silva and R. A. Silva. 1999. Unidirectional rotary motion in a molecular system. Nature 401: 150 152. Kelly, T. R., R. A. Silva, H. de Silva, S. Jasmin and Y. Zhao. 2000. A rationally designed prototype of a molecular motor. J. Am. Chem. Soc. 122: 6935 6949. Kelly, T. R., I. Tellitu and J. P. Sestelo. 1997. In search of molecular ratchets. Angew. Chem. Int. Ed. Engl. 36: 1866 1868. Koumura, N., E. M. Geertsema, A. Meetsma and B. L. Feringa. 2000. Light-driven molecular rotor: Unidirectional rotation controlled by a single stereogenic center. J. Am. Chem. Soc. 122: 12005 12006. Koumura, N., E. M. Geertsema, M. B. van Gelder, A. Meetsma and B. L. Feringa. 2002. Second generation of light-driven molecular motors. Unidirectional rotation controlled by a single stereogenic center with near perfect photoequilibria and acceleration of the speed of rotation by structural modification. J. Am. Chem. Soc. 124: 5037 5051. Koumura, N., R. W. J. Zijlstra, R. A. van Delden, N. Harada and B. L. Feringa. 1999. Light-driven monodirectional molecular rotor. Nature 401: 152 155.

23 Synthetic Molecular Motors Murakami, H., A. Kawabuchi, K. Kotoo, M. Kunitake and N. Nakashima. 1997. A lightdriven molecular shuttle based on a rotaxane. J. Am. Chem. Soc. 119: 7605 7606. Noji, H., R. Yasuda, M. Yoshida and K. Kinosita Jr. 1997. Direct observation of the rotation of F-1-ATPase. Nature 386: 299 302. Pease, A. R., J. O. Jeppesen, J. F. Stoddart, Y. Luo, C. P. Collier and J. R. Heath. 2001. Switching devices based on interlocked molecules. Acc. Chem. Res. 34: 433 444. Raymo, F. M. and J. F. Stoddart. 2001. Switchable catenanes and molecular shuttles. In: Molecular Switches. Edited by B. L. Feringa. Weinheim: Wiley-VCH, pp. 219 248. Sauvage, J.-P and V. Amendola. (Eds). 2001. Molecular Machines and Motors. Structure and Bonding Vol. 99. Berlin: Springer. Sauvage, J.-P. and C. Dietrich-Buchecker. (Eds). 1999. Molecular Catenanes, Rotaxanes and Knots. Weinheim: Wiley-VCH. Schalley, C. A., K. Beizai and F. Vögtle. 2001. On the way to rotaxane-based molecular motors: Studies in molecular mobility and topological chirality. Acc. Chem. Res. 34: 465 476. Schoevaars, A. M., W. Kruizinga, R. W. J. Zijlstra, N. Veldman, A. L. Spek and B. L. Feringa. 1997. Toward a switchable molecular rotor. Unexpected dynamic behavior of functionalized overcrowded alkenes. J. Org. Chem. 62: 4943 4948. Stevens, A. M. and C. J. Richards. 1997. A metallocene molecular gear. Tetrahedron Lett. 38: 7805 7808.

Tashiro, K., K. Konishi and T. Aida. 2000. Metal bisporphyrinate double-decker complexes as redox-responsive rotating modules. Studies on ligand rotation activities of the reduced and oxidized forms using chirality as a probe. J. Am. Chem. Soc. 122: 7921 7926. Vale, R. D. and R. A. Milligan. 2000. The way things move: looking under the hood of molecular motor proteins. Science 288: 88 95 and references therein. van Delden, R. A., N. Koumura, N. Harada and B. L. Feringa. 2002. Unidirectional rotary motion in a liquid crystalline environment: Color tuning by a molecular motor. Proc. Natl Acad. Sci. 99: 4945 4949. Walker, J. E. 1998. ATP synthesis by rotary catalysis (Nobel Lecture). Angew. Chem. Int. Ed. 37: 2300 2319. Whitesides, G. M. and J. C. Love. 2001. The art of building small – Researchers are discovering cheap, efficient ways to make structures only a few billionths of a meter across. Sci. Amer. 285: 38 47. Yin, H., M. D. Wang, K. Svoboda, R. Landick, S. M. Block and J. Gelles. 1995. Transcription against an applied force. Science 270: 1653 1657. Zijlstra, R. W. J., W. F. Jager, B. de Lange, P. T. van Duijnen, B. L. Feringa, H. Goto, A. Saito, N. Koumura and N. Harada. 1999. Chemistry of unique chiral olefins. 4. Theoretical studies of the racemization mechanism of transand cis-1,1l,2,2l,3,3l,4,4l-octahydro-4,4l-biphenanthrylidenes. J. Org. Chem. 64: 1667 1674.

577

Index a AAA ATPase 48 ff., 216, 335, 504 Acanthamoeba 5, 15, 31 Actin cables 272, 387 Actin ribbons 275 AMP-PNP 144 Amyloid precursor protein 90, 98, 99, 382, 390, 397 Arp 2/3 15 Ash1p 17, 366 ff., 459 ATP hydrolysis 250 ff., 254, ff., 278 ATP synthase 141 ff., 208 f., 546 ATPase activity – of kinesin 246, 247, 248, 262 – of myosin V 280

d

Deafness 29, 30, 511 f., 514 f. Dictyostelium 5, 9, 14, 15, 22, 27, 489, 516 Dilated cardiomyopathy 487 dilute 28, 393, 420 Directionality – of kinesin 100, 217 – of myosin 7 ff., 217 DNA polymerase 153 ff. DNA rotation 161 Duty ratio 8 Dynactin 388, 392, 395, 412, 414, 503, 505 Dynamic instability 332, 458 Dynein heavy chain 46, 48 ff., 237 f., 340, 364 ff., 371, 383, 384, 395 ff., 412, 425, 498 b Dynein light intermediate chain 46, 56 f., Bead assay 245, 272 498 bicoid mRNA 362 ff. Dynein intermediate chain 46, 53 ff., 498 Brownian ratchet 166, 190, 195, 207, 212, Dynein light chain 46, 57 ff., 498 223, 320, 331 – calmodulin-related 63 f. – I2/3 65 c – LC1 62 f. Caenorhabditis 21, 86, 87, 263, 347, 385, 400, – LC7/roadblock 61 ff., 423 489, 516, 523 – p29 64 Cardiac myopathy 478 ff. – Tctex 59 ff., 69, 397, 500 Cargo binding 98 ff. Dynein light chain- LC8 57 ff., 68 Cdc2 kinase 412 e CENP-E 95, 336, 342, 349, 414, 418, 425 Efficiency of motors 218 Chemo-mechanical coupling 312 ff. E-hook 262 Chlamydomonas 51, 52, 55, 57, 60, 61, 63, Endocytosis 25, 381, 387 65, 67, 385, 416 CHO1/KIF23 95 f Chromosome movement 327, 341, 343 F0 -motor 141, 212 CLIP-170 387 F1-motor 141ff., 154, 208, 543, 550, Congestive heart failure 478 553 Converter domain 3, 234, 237 – crystal structure 143, 146 ff. Coordination – rotation 144 ff., 148 ff. – of bidirectional movement 385 ff. – g subunit 142 – of kinesin heads 248 Familial hypertrophic cardiomyopathy 473 ff. Cytoplasmic streaming 458 f.

580

Index Force – during mitosis 331 ff., 344 – of DNA polymerase 154, 164 – of dynein 66, 335 – of kinesin 316, 335 – of myosin 307 – of RNA polymerase 154, 164 Fusome 368

g Golgi apparatus 25, 68, 380, 381, 394, 422, 503, 522 G-protein 293, 411 ff. GRIP1 90, 100 Griscelli syndrome 28

h

KIF21 94, 100 KIF26 96 KIFC1 97 KIFC2/C3 97 KIFC3 98 Kinesin chimera 231, 289 Kinesin folding 398, 424 Kinesin heavy chain 87 ff., 100, 154, 364 Kinesin hinge 292 Kinesin I 335, 342, 382, 390, 396 Kinesin light chains 89, 90, 99 Kinesin neck 233, 288, 292 Kinesin necklinker 233, 288, 292 Kinesin substeps 265 Kinesin superfamily proteins 81 ff. Kinesin, tetrameric 91 Kinetochores 340, 343, 503 f. klarsicht 369 ff., 426 K-loop 262

Helicase – crystal structure 184 ff. – efficiency of 191 l – translocation of 182, 188 ff. Hereditary inclusion body myopathies 488 f. Left-right axis 357 ff. Left-right dynein 358, 360 i Lissencephaly 69, 368, 369, 425, 501 ff. Intraflagellar transport (IFT) 385 f., 392, 499 Load 257 ff. Inversion 360 m IQ motif 3, 10, 234, 280, 282, 448 MAP 1C 497 j Melanophilin 19, 20, 394, 420 JNK interacting proteins 89, 91, 99, 346, Melanososmes 11, 18, 384, 393 382 f., 390 Membrane traffic 379, ff., 522 Microneedle 307 k Minus-end movement 230, 234, 317, Kar3 97, 289, 295, 298, 299 366 Kartagener’s syndrome 357, 488 Mitotic spindle Katanin 342 – microtubules 327 ff. Kid 95 – morphogenesis 327, 394 KIF1 91 f., 99, 262 , 289, 295, 298, 300, 383, MKLP1 347 f., 423 385, 390 Motor-cargo linkage 382, 388 ff., 395, 398, KIF2 96, 99 412 ff. KIF3 93 f., 98, 358, 360, 396 mRNA localization 361 ff., 459 KIF4 94 MTOC 371, 378, 380, 421 KIF5 87 ff., 100 Myocardium 476 ff. KIF6 98 Myosin KIF9 98, 423 – cardiac 475 ff. KIF11 91 – ankyrin repeat 4 KIF12 96 – coiled-coil 4, 9 KIF13 99, 390, 391 – FERM domain 4, 8, 516 KIF13 92 – lever arm 273 KIF15 96 – light chain 10 f., 481 ff. KIF16 92 – MyTH4 domain 4, 10, 516 KIF17 94, 391 – N-terminal extension 4, 8 KIF18 96 – PEST site 4, 9 KIF20 95 – pleckstrin homology domain 4, 8

Index – proline-rich region 4 – protein kinase domain 4, 8 – regulation 10 ff. – rho-GAP domain 4, 9, 27 – S1 273, 274, 319, 475 ff. – src homology domain 4, 9 – zinc-binding domain 4, 9 Myosin binding protein 485 Myosin chimera 235 Myosin I 5, 11, 21, 24, 25, 26, 27 Myosin II 14, 277, 291, 475 ff. Myosin III 529 ff. Myosin V 7, 8, 10, 11, 16, 17, 18, 20, 21, 22, 25, 28, 57, 235, 278 ff., 317 ff., 367, 383, 384, 387, 393, 396, 414, 418, 420, 426 Myosin VI 20, 21, 22, 24, 25, 234, 235, 277, 280, 317, 387, 391, 515, 520 ff. Myosin VII 20, 21, 24, 29, 384, 396, 514 ff. Myosin VIII 448 ff., 459 Myosin IX 234, 317, 515, 528 f. Myosin XI 449 ff. Myosin XV 525 ff.

P-loop 48, 50, 291, 294 Plus-end movement 230 Power stroke 207 ff., 214, 221 - of helicases 195 - of RNA polymerase 164 Pre-prophase band 452 Primary ciliary dyskinesia 357 ff., 498 Processivity – of DNA polymerase 155 – of dynein 66 – of helicases 197 – of kinesin 244 ff., 249, 314 – of myosin 7 ff., 317, 378 – of RNA polymerase 155

n

s

Nanoscale directed assembly 543 ff. Ncd 97, 230, 233, 259 ff., 289, 299 Neuron 79 NinaC 21, 26, 391, 530 Nodal flow 358 ff., 498 NTP polymerization 155 ff. Nuclear migration 371, 380

r Rab 27a 19, 20, 28, 391, 394, 419 f., 516 Rab6 kinesin 95, 394, 423 Retinitis pigmentosa 60, 500, 511 Reynolds number 329 Rhodopsin 60, 500 RNA polymerase 153 ff. RNAi 31, 32

Saccharomyces 5, 16, 57, 87, 179, 361, 366 ff., 387 Schizosaccharomyces 5, 14 Secretion 281 ff. Shaker 1 24, 29, 30, 518 f. Shaker 2 526 f. She proteins 366 ff. Single molecule techniques o – F1 motor 144 ff. Optical tweezers 159, 163, 245, 247, 254, 275, – kinesin 245 ff. , 314 ff. 276, 306f. – myosin 272 ff., 306 ff. oskar mRNA 362 ff. – RNA and DNA polymerase 158 ff. Osm3/KIF17 94, 99 Situs inversus 68, 360 Smy 1 16, 387, 426 p SNARE proteins 380 p150glued 55, 371, 413 Snell’s waltzer 22, 24, 30, 515, 522 ff. PAF acetylhydrolase 502, 505 Spindle assembly checkpoint 349 Phosphorylation Spindle matrix 345 – dynactin 412 Spindle pole body 453 – dynein 52, 400, 412, 416, 418 Step size – kinesin 400, 413 f., 418, 461 – of myosin V 277, 278 ff., 318 – myosin 11, 399, 414, 418, 485 – of DNA polymerase 154 Phragmoplast 454 ff. – of helicases 192 Plant dyneins 447 f. – of kinesin 246, 316 Plant kinesins 434 ff. – of myosin 275, 278, 280, 309 f., 318 Plant myosins 448 ff. – of RNA polymerase 154 Plasmodesmata 459 Stereocilia 21, 22 Plasmodium 5, 15 Switch I and II 273, 291, 292, 297, 299

581

582

Index

t Tetratrico peptide repeat motifs 89 Thioredoxin 64 Titin 487 Total internal reflection microscopy 311 Toxoplasma 5, 15

– of dynein 66 – of helicases 197 – of kinesin 154 – of myosin 154, 479 – of RNA polymerase 154 Viscosity 329 Visual impairment 513 ff.

u Unc104/KIF1 91 f., 99, 262 Usher syndrome 513, 519 f.

x

v

z

Velocity – of DNA polymerase 154

Z-line 487

X-ray diffraction 271

Molecular Motors

Molecular Machines & Motors

Molecular Machines & Motors (Structure and Bonding)

Myosins - A Superfamily of Molecular Motors

Myosins: A Superfamily of Molecular Motors

Myosins: A Superfamily of Molecular Motors (Proteins and Cell Regulation)

Fibrous Proteins: Muscle and Molecular Motors (Advances in Protein Chemistry)

ENERGY-EFFICIENT ELECTRIC MOTORS

Electric Motors Drives

AC Motors(en)(92s)

Energy-efficient electric motors