The Enzymes VOLUME XXIII
ENERGY COUPLING AND MOLECULAR MOTORS Third Edition
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THE ENZYMES Edited by David D. Hackney
Fuyuhiko Tamanoi
Department of Biological Sciences Carnegie Mellon University 4400 Fifth Ave. Pittsburgh, PA 15213, USA
Department of Microbiology, Immunology and Molecular Genetics and Molecular Biology Institute University of California Los Angeles, CA 90095, USA
Volume XXIII ENERGY COUPLING AND MOLECULAR MOTORS
THIRD EDITION
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Oxford
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Dedication This volume is dedicated to Dr. David S. Sigman, Professor of Biological Chemistry at the University of California, Los Angeles. Dr. Sigman served as a series editor for The Enzymes and is responsible for the publication of volumes such as ‘‘The Mechanisms of Catalysis.’’ His enthusiasm for science in general and enzyme mechanisms in particular was an inspiration. He had the foresight to bring together a volume on the diverse field of molecular motors, and he initiated this project and contributed greatly to its development. Sadly, Dr. Sigman passed away on November 11, 2001 without seeing the fruit of his effort. We greatly miss his originality and leadership.
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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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1. Muscle Contraction YALE E. GOLDMAN I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII. XIII.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sarcomere Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Distribution of Myosin Superfamily Members and Contractile Proteins . Myosin Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Working Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Actomyosin ATPase Cycle in Solution . . . . . . . . . . . . . . . . . . . . . . . . Comparison of ATPase Kinetics Between a Protein Suspension and the Sarcomeric Filament Lattice . . . . . . . . . . . . . . . . . Myofibrillar ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Muscle Fiber Mechanics and Energetics . . . . . . . . . . . . . . . . . . . . . . . Biochemical Rate Constants in Muscle Fibers . . . . . . . . . . . . . . . . . . . Structural Changes Leading to Force Generation and Filament Sliding . Why Does Myosin Have Two Heads? . . . . . . . . . . . . . . . . . . . . . . . . . Summary, Uncertainties and Future Directions . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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2. Mechanics of Unconventional Myosins RONALD S. ROCK, THOMAS J. PURCELL, JAMES A. SPUDICH
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I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Single-Molecule Analysis Revealed a Unitary Small Step in Motion as Myosin Interacts with Actin. . . . . . . . . . . . . . . . . . . . . . . . . . . III. Molecular Genetic Approaches have Indicated Roles of Various Domains and Specific Residues of the Myosin Motor. . . . . . . . . . . IV. The Unconventional Myosins V and VI are Adapted for Cellular Transport Roles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Requirements of Processive Motors . . . . . . . . . . . . . . . . . . . . . . .
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CONTENTS
VI. Conclusions and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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3. Motor Proteins of the Kinesin Superfamily DAVID D. HACKNEY I. II. III. IV. V. VI. VII. VIII.
Introduction . . . . . . . . . . . . . . Structure . . . . . . . . . . . . . . . . Characterization of Motility. . . ATPase Mechanism. . . . . . . . . MT Decoration. . . . . . . . . . . . Generation of Motility. . . . . . . Regulation and Cargo Binding . Perspectives . . . . . . . . . . . . . . References . . . . . . . . . . . . . . .
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4. The Bacterial Rotary Motor HOWARD C. BERG I. II. III. IV.
Introduction . . . . . . Bacterial Behavior . . The Flagellar Motor Future Work . . . . . References . . . . . . .
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5. The ATP Synthase: Parts and Properties of a Rotary Motor THOMAS M. DUNCAN I. II. III. IV. V. VI.
Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conservation of General Structure and Function in FOF1 . . . . . . . . . The Binding-change Mechanism for ATP Synthesis and Hydrolysis . . F1’s Structural compatibility with a Cooperative, Rotary Mechanism Demonstration and Analysis of Subunit Rotation in F1 and in FOF1 . Further Characteristics of FO and F1 Subunits as Components of the Rotor or Stator. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Remaining Puzzles for Rotational Catalysis. . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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6. Bacteriophage T7 Gene 4 Protein: A Hexameric DNA Helicase DONALD J. CRAMPTON
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CHARLES C. RICHARDSON
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Isolation and Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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CONTENTS III. Structural and Biochemical Properties. IV. Models for Energy Transduction . . . . V. Future Directions . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . .
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7. DNA Helicases, Motors that Move Along Nucleic Acids: Lessons from the SF1Helicase Superfamily TIMOTHY M. LOHMAN, JOHN HSIEH, NASIB K. MALUF, WEI CHENG, AARON L. LUCIUS, CHRISTOPHER J. FISCHER, KATHERINE M. BRENDZA, SERGEY KOROLEV, AND GABRIEL WAKSMAN I. II. III. IV. V. VI. VII. VIII. IX. X. XI. XII.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phenomenological Features of DNA Unwinding . . . . . . . . . . . . . . . . . Structural Features of SF1 DNA Helicases . . . . . . . . . . . . . . . . . . . . . Protein Oligomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . DNA Binding by E. coli Rep . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanisms of Nucleotide Binding and ATP Hydrolysis by E. coli Rep Single-Stranded DNA Translocation by Monomers of SF1 Helicases. . . Presteady-State, Single-Turnover DNA Unwinding Studies . . . . . . . . . . DNA Unwinding by E. coli Rep and UvrD Helicases. . . . . . . . . . . . . . Helicase Activity of SF1 Monomers . . . . . . . . . . . . . . . . . . . . . . . . . . E. coli RecBCD Helicase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Proposed Mechanisms for DNA Unwinding and Translocation by SF1 Helicases . . . . . . . . . . . . . . . . . . . . . . . . . . XIII. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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8. Type II DNA Topoisomerases: Coupling Directional DNA Transport to ATP Hydrolysis JANET E. LINDSLEY I. II. III. IV.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Basic Biochemical and Structural Information about Type The Mechanism of Strand Passage . . . . . . . . . . . . . . . . . Concluding Thoughts . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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I. The Work Carried out by Chaperonins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. ATP Action in Driving Chaperonin-Assisted Protein Folding – Structural States and the Overall Chaperonin Cycle . . . . . . . . . . . . . . . . . . . . .
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9. The Role of ATP in Directing Chaperonin-Mediated Polypeptide Folding ARTHUR L. HORWICH
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WAYNE A. FENTON
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CONTENTS III. Mechanistic Studies of the Nucleotide Cycle IV. Polypeptide and the Nucleotide Cycle . . . . . V. Cooperativity and Allostery . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .
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Author Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Preface Molecular motors are remarkable cellular machines that convert chemical energy generated by the hydrolysis of nucleotides or by ion gradients into mechanical work. These tiny machines catalyze critical cellular functions such as muscle contraction, transportation of vesicles along cytoskeletal filaments, ATP synthesis, cell locomotion and nucleic acid transactions. Recent enzymatic, structural as well as single-molecule studies have led to deeper understanding of how these molecular machines function. This volume is intended to capture these dramatic developments by bringing together works on diverse molecular machines and to provide an in-depth review of common principles, as well as differences, among molecular machines that couple chemical energy to mechanical work. The information should provide valuable guide to biochemists and biophysicists studying molecular machines. Defects in molecular motors have been implicated in a number of human diseases. For example, myosin defects are involved in myopathies and hearing loss. Understanding how molecular motors function should be of importance for investigating human diseases involving molecular motors. Molecular motors have also captured imagination of chemists who are building artificial motors and machines of nanosize dimensions that achieve linear and rotary motion. Attempts are being made to drive these ‘‘nanoscale’’ machines by chemical, electrochemical or photochemical forces. Basic guiding principles for building such artificial machines may be gleaned from this volume. The first three chapters discuss structure and function of cytoplasmic molecular motors that move along cytoskeletal filaments. These proteins couple ATP binding and hydrolysis to conformational changes that are amplified and converted into movement. Chapters 1 and 2 discuss myosins that move along actin filaments, with emphasis on how myosins generate movement in organized muscle fibers and at the single molecule level, respectively. Kinesins are motor proteins that move along microtubules and they are discussed in Chapter 3. The core motor domains of myosin and kinesin are structurally homologous, but their mechanisms for generation of movement are not as similar. Myosin has a lever arm that can swing to cause large displacements, whereas kinesin does not, and kinesin likely produces xi
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PREFACE
movement at least in part through conformational changes in the neck regions adjacent to the motor domain. Another type of molecular motor is a rotary machine that converts energy generated from ion gradients into rotary motions. Elucidation of the structure and function of F0F1 ATP synthase represents one of the major milestones in the study of molecular motors. This remarkable machine utilizes the energy generated by the flow of protons down the gradient through F0 to catalyze ATP synthesis on F1 through physical rotation of its subunits. Rotary motors are also used for cell motility as seen in the case of bacterial flagellar motors that use proton motive force to rotate the helical flagellar filament driving cell motility. How these highly efficient rotary motors function and how they respond to chemical gradients are discussed in Chapters 4 and 5. Molecular motors also play critical functions during DNA metabolism. Chapters 6 and 7 feature DNA helicases, linear molecular motors that translocate along the DNA lattice and perform unwinding of DNA duplexes. This is accomplished by coupling the energy generated from the binding and hydrolysis of nucleoside triphosphates to their mechanical function. The generation of single-stranded DNA is critical for DNA metabolism such as DNA replication, repair and recombination. A hexameric helicase, T7 gene 4 protein, as well as superfamily 1 DNA helicases are discussed. One of the major issues in understanding molecular motors is how the energy generated by ATP hydrolysis is coupled to mechanical work. This theme continues in Chapters 8 and 9. Chapter 8 discusses type II topoisomerases, enzymes that utilize ATP binding and hydrolysis to catalyze directional transport of one duplex DNA segment through a transient break in another DNA duplex. The resulting topological changes of DNA are critical for DNA metabolism such as unlinking replicated chromosomes. Recent studies on protein folding led to the realization that the folding of some proteins occurs within a hollow, barrel-like structure called chaperonin. In the case of the prokaryotic GroEL machine, 14 identical subunits are arranged in two stacked rings. ATP binding and hydrolysis is critical for the function of this machine, as ATP binding and hydrolysis influences the ring to shift between an open polypeptide-accepting state and a closed folding-active state. Chapter 9 discusses structure and function of this machine. A list of molecular motors continues to grow, as novel classes of molecular motors emerge. For example, a family of proteins catalyzing chromatin remodeling has been identified. These proteins called ACF
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(ATP-dependent chromatin-assembly factor) use the energy of ATP hydrolysis to deposit histones into nucleosome arrays. SMC (structural maintenance of chromosomes) proteins form cohesion and condensin complexes that play critical roles during sister chromatid cohesion and mitotic chromosome condensation. Also, mechanisms used by the AAA ATPases such as the motor protein dynein are just starting to be elucidated in detail. These and other novel molecular motors may be the subjects of a future volume. We greatly appreciate the contributors for providing thorough and original reviews. This project could not have taken off without the effort of Dr. David S. Sigman. We would like to dedicate this volume in recognition of his contributions. We are particularly indebted to Dr. Paul D. Boyer whose work on F0F1 ATP synthase gave inspiration to put together this volume. He provided critical guidance throughout the preparation of this volume. We greatly value his insight and advice. We thank Shirley Light of Academic Press who assisted us during the initial stage of the preparation of this volume. We also thank Mica Haley of Academic Press/Elsevier for her assistance. Fuyuhiko Tamanoi David D. Hackney
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1
Muscle Contraction YALE E. GOLDMAN Department of Physiology and Pennsylvania Muscle Institute School of Medicine University of Pennsylvania Philadelphia, PA 19104-6083, USA
I. II. III. IV. V. VI. VII. VIII. IX. X.
XI.
XII. XIII.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sarcomere Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Distribution of Myosin Superfamily Members and Contractile Proteins . Myosin Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Working Hypothesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Actomyosin ATPase Cycle in Solution . . . . . . . . . . . . . . . . . . . . . . . . Comparison of ATPase Kinetics Between a Protein Suspension and the Sarcomeric Filament Lattice . . . . . . . . . . . . . . . . . Myofibrillar ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Muscle Fiber Mechanics and Energetics . . . . . . . . . . . . . . . . . . . . . . . Biochemical Rate Constants in Muscle Fibers . . . . . . . . . . . . . . . . . . . A. Steady-State ATPase Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. ATP-Induced Actomyosin Dissociation . . . . . . . . . . . . . . . . . . . . . C. ATP Hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Pi Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. ADP Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structural Changes Leading to Force Generation and Filament Sliding . A. Tilting of the Light-Chain Domain . . . . . . . . . . . . . . . . . . . . . . . . B. Tilting of the Motor Domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . Why Does Myosin Have Two Heads? . . . . . . . . . . . . . . . . . . . . . . . . . Summary, Uncertainties and Future Directions . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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1 THE ENZYMES, Vol. XXIII Copyright ß 2003 by Academic Press All rights of reproduction in any form reserved.
2
YALE E. GOLDMAN
I. Introduction Animal locomotion is the most obvious consequence of any biological energy transducer. The functional outputs of muscle contraction – force and shortening – are easy to observe and to quantify at high time resolution. These characteristics enabled investigation of the mechanics and energetics of the contraction mechanism starting in the mid 1800s (1), and in great detail by A. V. Hill and colleagues beginning in the 1910s (2). Because the contractile proteins, myosin and actin, account for over half of the protein in a muscle cell, they were among the first macromolecules to be isolated (3), associated with adenosine triphosphatase (ATPase) activity (4), and purified (5, 6). Detailed studies of transient kinetics (7, 8) were also facilitated by the ready availability of purified myosin and actin. The assembly of the muscle contractile apparatus into a nearly crystalline periodic array enabled detailed structural studies by light microscopy [reviewed by A. F. Huxley (9)], electron microscopy (10), and low-angle X-ray diffraction (11) [reviewed by Squire (12)]. By the 1970s, mechanical, structural, and biochemical studies had led to the hypotheses of sliding filaments, a cyclic interaction between actin and myosin, and tilting of the myosin heads. Protein filaments composed of actin and myosin interdigitate in overlap zones within the sarcomere, the contractile organelle. When a muscle shortens, the two filaments do not appreciably change length, but instead, they slide relative to each other, increasing the overlap (13, 14). Sliding motions generated in all of the sarcomeres spaced sequentially along a muscle cell sum to produce macroscopic shortening of the whole muscle. Thus, the problem of understanding generation of force and shortening of a muscle is reduced to the molecular interactions between the two filaments. Myosin and actin are thought to undergo a cyclic association and dissociation that leads to production of force but still allows sliding to occur (15–17). This crossbridge cycle is a sequence of enzymatic reaction steps that are coupled to the binding and splitting of ATP and release of the hydrolysis products, orthophosphate (Pi) and ADP. The immediate effect of ATP binding to actomyosin is to dissociate the two proteins (18). A structural change, possibly tilting of the myosin head while it is attached to actin, is the direct cause of the filament sliding, pulling the actin toward the center of the sarcomere (19, 20). Biochemical and structural dynamics of the actomyosin mechanism in working muscle fibers were established during the 1980s (21–24). Recapitulation of motility in vitro from purified components was accomplished (25, 26) and enhanced using laser tweezers (27) and molecular biology (28).
1. MUSCLE CONTRACTION
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The crystal structure of myosin (29–31) refined the tilting theory by showing structural changes within the myosin head. During force production, the whole myosin head probably does not rotate, but a domain containing light chains serves as a lever arm that rotates to exert the sliding force between the two filaments. A motor domain, containing the ATP- and actin-binding sites stays rigidly attached to actin during the lever arm rotation (30, 32–34). Surprisingly, close structural homology between the nucleotide-binding fold of myosin and other motor proteins, such as kinesin, the GTP-binding signal transducing proteins (G-proteins), and nucleotide-binding metabolic enzymes, makes it likely that all of these proteins share some common mechanistic features (35, 36). The present chapter summarizes the structural, mechanical, and biochemical data that relate actomyosin ATPase activity to force production and shortening of striated muscle. The enzymatic cycle is described for purified components and in fully functioning preparations of muscle. Investigation of certain aspects of the mechanism, especially stress–strain mechanics and control of biochemical kinetics by mechanical loads, requires the intact filament array. The description of studies on organized muscle preparations complements the chapter in this compendium which emphasizes myosin function in vitro (37). Other reviews of actomyosinbased motility and muscle contraction have appeared recently (33, 38–40). Regulation of contraction has also been reviewed elsewhere (41, 42).
II. Sarcomere Structure In striated muscle cells (heart and skeletal muscle), the contractile machinery is packaged in myofibrils, long 1 m-diameter cylindrical organelles (Fig. 1A). There is no membrane diffusion barrier between the cytoplasm and interior of the myofibril. Each myofibril is a column of sarcomeres (Fig. 1B), the basic contractile units, which are approximately 2.2 m in length and delimited by the electron-dense Z lines. The contractile and structural proteins within each sarcomere self-assemble to form a highly ordered, nearly crystalline lattice of interdigitating thick (myosin) and thin (actin) myofilaments. The sarcomeres and their internal filament arrays are remarkably uniform in both length and lateral registration, giving rise to the cross-striated histological appearance of skeletal and cardiac muscles (43). Myosin cross-bridges between these two sets of filaments produce the sliding force that causes muscle shortening. Muscle myosin (Fig. 1C) is a highly asymmetric 470-kDa protein composed of two heavy chains and four myosin light chains. Each heavy chain has a long ( 156 nm) C-terminal -helical coiled-coil tail region
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FIG. 1. Structure of the contractile apparatus. (A) The myofibril. Bands are due to the axial alignment of adjacent thick and thin filaments. The myofibril is about 1 m in diameter. One sarcomere extends 2.2 m along the myofibril between two Z lines. A sarcomere contains a centrally located A band flanked by half of an I band at each end. (B) Expanded view of the sarcomere. Three filament systems are shown: thick myosin-containing filaments, thin actincontaining filaments, and titin filaments. (C) Whole myosin molecule. Each molecule contains six peptides: two heavy chains, two regulatory light chains, and two essential light chains. The heavy chains extend from the heads (S1) at the left into the coiled-coil rod, making up subfragment-2 (S2) and light meromyosin (LMM). (D) Structure of the thin filament. Each actin monomer in this cartoon is represented by a peanut-shaped density. Tropomyosin winds
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[subfragment-2 (S2) and light meromyosin (LMM) in Fig. 1C] and a 16-nm globular subfragment-1 head (S1, Fig. 1E), which constitutes the crossbridge. In muscle, the LMM regions of approximately 300 myosin molecules are polymerized to form the backbone of each 1.6 m-long thick filament (Fig. 1B, dimensions applicable to mammalian skeletal muscle). The S1 cross-bridges, protruding from the filament shaft contain a motor domain having the ATPase and actin-binding sites responsible for the conversion of chemical energy into mechanical work. S1 also includes the regulatory light chain (RLC, Fig. 1E) and the essential light chain (ELC), which, along with an underlying heavy-chain helix, is termed the light-chain domain (LCD, Fig. 1E). Three myosin molecules (six S1 heads) are spaced every 14.3 nm along the filament axis (44). An antiparallel arrangement of the myosin tails associated at the center of the thick filaments makes the filament bipolar (the tails point oppositely on each side) and produces a central area ( 0.15 m long), termed the bare or pseudo-H zone, that does not contain S1 heads. C-protein is located at discrete intervals along each half of the thick filament, fulfilling structural and modulatory roles (45). Thick filaments are located in the center of the sarcomere in the optically anisotropic A band (Figs. 1A and B). They are organized into a hexagonal lattice stabilized at the M line (Fig. 1B) by M-protein, myomesin (46–48), and muscle-specific creatine kinase [MM-CK, (49–51)]. Thin filaments (Fig. 1D) are helical polymers of actin that extend 1.1 m from each side of the Z line occupying the optically isotropic I band (Fig. 1A) and extending into the A band to interdigitate with the thick filaments. Monomers of actin are 45 kDa globular proteins, termed G-actin. Actin polymerizes into a double-stranded helical filament, 8 nm in diameter. The pitch of the helix is 74 nm, and so in longitudinal electron microscopic (EM) images, the two strands cross over each other every 36–40 nm (Figs. 1B and D). In the thin filament, the monomers can also be considered to be wound in a tighter coil, termed the genetic helix, with left-handed 5.9 nm and right-handed 5.1 nm pitches. The overall spacing of monomers along the filament axis is 2.7 nm. Each thin filament projecting from the Z line contains approximately 360 actin monomers.
in a double-helical coiled-coil polymer along each groove between the actin protofilaments. Each tropomyosin molecule is associated with a troponin complex containing troponin I (TnI), troponin C (TnC), and troponin T (TnT). Troponin and tropomyosin regulate contraction in vertebrate striated muscle. (E) The crystal structure of S1. The heavy chain winds through the motor domain (MD) and extends into an -helix that forms the backbone of the light-chain domain (LCD). The regulatory light chain (RLC) and essential light chain (ELC) bind to this helix. Panel E is adapted from (104).
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Actin filaments are polarized with slower- and faster-polymerizing ends termed the pointed and barbed ends, respectively. The pointed ends, located away from the Z lines, are capped by tropomodulin (52, 53). The barbed ends associate with CapZ protein (54, 55) and insert into the Z line which contains the densely packed structural protein -actinin. The activity of vertebrate striated muscle is switched on and off by regulatory proteins on the thin filaments. The regulatory complex contains one tropomyosin molecule and three troponin subunits (TnC, TnT, and TnI) associated with each successive group of seven actin monomers along the thin filament [Fig. 1D, (42, 56)]. Calcium binding to TnC initiates each contraction. Two additional sarcomeric proteins, titin and nebulin, are among the largest individual peptides identified throughout cell biology. They have been postulated to serve as ‘‘molecular rulers’’ defining the length and position of the thick and thin filaments during sarcomere assembly and maintenance (55, 57). Titin (Fig. 1B) is associated with the thick filament and -actinin. Individual titin molecules ( 3 MDa) extend all the way from the M line to the Z line (47, 58), a distance of 1–2 m. Titin contains repeating fibronectin, immunoglobulin, and unusual proline-rich domains that confer mechanical elasticity on the resting sarcomere and probably help position the thick filaments in the center of the sarcomere (59). Nebulin ( 800 kDa) is associated with the Z line and thin filaments (57). It may help to define the length of actin filaments in skeletal muscle and modulate contraction by associations with actin, myosin, and tropomyosin (57, 60). Cross-sections of the myofibril in the regions of overlap show that the filaments are arrayed into a double-hexagonal lattice. The thin filaments occupy so-called trigonal positions, equidistant from three thick filaments (Figs. 10C and D). Each thick filament is surrounded by six thin filaments. Both sets of filaments are structurally polarized. Relative to the tails, the myosin heads are positioned away from the center of the sarcomere. The pointed ends of the actin filaments are oriented away from the Z line. Thus, the two halves of the sarcomere are related by a two-fold rotational symmetry. During contraction of an active muscle, the filaments do not appreciably change length. Instead, the enzymatically driven interaction between the thick and thin filaments within each half sarcomere produces a sliding force that translates the thin filaments toward the M line. The overall sarcomere shortening is the sum of sliding motions generated within each of the two overlap zones, so the amount of filament sliding is equivalent to the shortening per half sarcomere. Owing to their sequential connection along each myofibril, the shortening of the sarcomeres sums to produce macroscopic shortening of the whole muscle.
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III. Distribution of Myosin Superfamily Members and Contractile Proteins Muscle fibers adapt to their specific functional role and the demand for activity by balancing speed of contraction against resistance to fatigue. In mammals, three main fiber types display stereotyped patterns of contractile protein isoforms, mechanical properties, balance between aerobic and glycolytic metabolism, fiber size, and content of mitochondria and sarcoplasmic reticulum (61–63). For instance, muscles used for postural maintenance express an isoform of myosin with slower ATPase rate than muscles designed for rapid bursts of intense activity. There are also more specialized fiber types with corresponding isoforms (64, 65) and intermediate fiber types. Ventricular myocardium shares isoforms of the contractile proteins with the slower fiber types of skeletal muscle (66). The light chains and regulatory proteins also have specific fiber-type isoform expression patterns (61, 62). Smooth muscle does not display cross-striations, which contributed to doubt in the early 1900s about relevance of the sarcomeres to the mechanism of contraction in heart and skeletal muscle (9). Although less regularly organized, actin, myosin, and dense bodies containing -actinin are arranged serially within smooth muscle cells to generate force and transmit it to the cell exterior as in striated muscle. According to conservation of their amino acid sequences, smooth muscle myosin is more closely related to conventional myosins found in vertebrate nonmuscle cells than to striated muscle myosin (67). Smooth muscle contraction is regulated by specific phosphorylation and dephosphorylation of myosin RLC (68, 69), although regulatory proteins on the thin filaments also modulate contraction (70). Myosin is also common in nonmuscle cells. The first nonmuscle isoform was discovered in the microorganism Acanthamoeba castellani (71) and termed myosin I because it contains only one head in contrast to ‘‘conventional’’ two-headed myosin from muscle (myosin II). Members of the myosin I class are now recognized widely in phylogeny (72) and take part in chemotaxis, endocytosis, and other functions (73). Further members of the myosin superfamily were classified in order of their identification and the group has expanded to at least 18 classes (39, 67, 74, 75). They exhibit sequence homology in the head regions but have variable LC composition and tails which determine their cellular location and cargo specificity. These proteins exhibit highly diverse functional attributes according to their myriad roles in cell motility such as chemotaxis, cytokinesis, pinocytosis, targeted vesicle transport, organelle assembly, modulation of sensory systems, and signal transduction (39, 75). They are typically regulated by phosphorylation of the heavy chain or by binding of calmodulin to the ‘‘neck’’ region between the head and tail (76).
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Class V myosins associate with actin over a much larger proportion of their ATPase cycle than muscle myosin (77). This characteristic probably enables single molecules or very small groups to transport vesicular and granular cargoes along actin filaments (78, 79), possibly in cooperation with microtubule-based motors (75, 80). Class VI myosin, located with actin filaments in the golgi apparatus and at the leading edge of ruffling cells (81), translates toward the pointed end of actin, opposite to the direction of other myosins (82). This feature is consistent with centrally directed cargo transport from the cell membrane. Myosin III participates in phototransduction in Drosophila retina (83, 84). Myosins I, VI, VII, and XV have been located in hair cells, the mechanoelectrical transducers of the inner ear (39, 83, 85, 86). Mutations of these classes can lead to inherited neurological diseases (87, 88). Myosin IX contains a GTPase-activating domain for rho, a G-protein that controls actin filament dynamics (89). Functions of many of the other classes are still obscure. Actin is ubiquitous in the cytoskeleton of eukaryotic cells and fulfills many roles in determining cell shape, locomotion, endocytosis, targeted vesicle transport, and cytokinesis (90, 91). Actin and -actinin are present in stress fibers and terminate at attachment plaques in nonmuscle cells attached to substrates (92). Titin antigenic epitopes have been found in association with mitotic chromosomes suggesting that titin may provide mechanical elasticity during chromosomal condensation (93).
IV. Myosin Structure Muscle myosin contains two 200-kDa heavy chains and four 17–23-kDa light chains. The C-terminal tail portion of the heavy chain is almost entirely an -helical coiled-coil rod, LMM forming the backbone of the thick filaments and S2 linking the globular head domains to the backbone (Fig. 1C). Within each globular, N-terminal subfragment-1 (S1), a ‘‘motor domain’’ (MD) contains the ATP- and actin-binding sites [(29), Fig. 1E]. A ‘‘neck region,’’ also termed the ‘‘light-chain domain’’ (LCD) and the ‘‘lever arm,’’ connects the MD to the tail. The LCD contains a 9-nm long heavy-chain -helix and the two light chains (Fig. 1E). The ELC and RLC, which are related to calmodulins, provide structural integrity to the lever arm and are involved in regulation of force production in some muscles. A flexible hinge at the S1–S2 junction allows the S1 to adopt a wide range of angles relative to the tail. About 60 nm from the S1, between S2 and LMM, is a second hinge (Fig. 1C). Trypsin cleaves the S1 heavy chain into three fragments that define regions of the MD (Fig. 2A). A 25-kDa N-terminal domain, participates
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FIG. 2. Crystal structures and cartoons of three conformations of myosin S1. The light chains have been removed for clarity. (A) Near-rigor structure in the absence of nucleotide. (B) Prepower-stroke structure crystallized in the presence of ADP AlF4. (C) Detached structure, crystal form obtained in the presence of ADP. In cartoons of B and C, right side, rotations of the lower 50 kDa and converter domains relative to the structure in A are shown as curved arrows around their axes of rotation (straight lines or a dot). Switch II, the SH1 helix, and the relay serve as joints linking the various subdomains to each other. In the detached configuration (C) the lower 50 kDa domain and the converter domain are more loosely associated with the N-terminal segments suggesting that these domains would be mobile in solution. Adapted from (94). (See color plate.)
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in binding ATP. The 50-kDa central fragment forms the remainder of the nucleotide-binding site and the actin-binding interface. The 50-kDa segment is divided by a prominent cleft extending from the actin binding to the ATPbinding regions, defining ‘‘upper 50 kDa’’ and ‘‘lower 50 kDa’’ domains. A C-terminal 20-kDa segment completes the motor domain and extends through the lever arm as a 10-nm long single -helix. The portion of the 20 kDa segment within the MD has been termed the ‘‘converter domain’’ (94). Small changes of structure at the active site move the converter through a large angle which it transmits to the lever arm (31, 95, 96). The core of the motor domain contains a seven-stranded -sheet, six -helices and several short nucleotide-binding sequences that show astonishing tertiary structural resemblance to many other nucleotidases, including GTP-binding proteins, kinesins, and the mitochondrial F1-ATP synthase (35, 36). Although there is very little overall sequence homology between these enzyme families, short loops in S1 contain consensus sequences that are shared with other nucleotide-binding enzymes. The P-loop, switch I, and switch II by homology with the corresponding motifs in GTP-binding proteins (35, 36, 97) provide liganding for the Mg2 þ ion and water molecules essential for hydrolysis of ATP. Switch II also senses the absence of the product phosphate following its release. A fourth consensus sequence [termed 3 in G-proteins (97)], contributes to defining the nucleotide base specificity. The structures of S1 from several isoforms have been solved by X-ray crystallography. Three markedly different conformations, bound to various ligands, have been identified in these studies (Fig. 2), providing a structural basis for postulating the lever arm mechanism of filament sliding. Chicken skeletal muscle myosin S1, without any nucleotide bound (Fig. 2A), was solved by Rayment and colleagues (29, 30) using protein purified from muscle tissue and methylated to enable crystallization. This structure of S1 is virtually identical to that of a Dictyostelium discoideum myosin II construct with bound ligands that resemble ATP such as ADP and ADPberyllium fluoride [ADP BeFx (95)]. The structure of myosin complexed to actin has not been solved to atomic resolution, but it has been determined at lower (2–5 nm) resolution by cryoelectron microscopy (30, 98–101). In the absence of ATP myosin binds tightly to actin in a conformation, the rigor complex, thought to represent the end of the mechanical working stroke. The atomic model of the nucleotide-free chicken structure (Figs. 1E and 2A) fits well into the cryoEM density maps of actin filaments decorated with nucleotide-free S1 (30), suggesting that this crystal structure is similar to the structure of S1 at the end of the power stroke. Thus, it is termed the ‘‘near-rigor’’ structure. The orientation of the actin filament axis when S1 is docked into the cryo-EM
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density map is shown in Fig. 2A to indicate the expected direction of force generation. During contraction, the thick filament is pulled toward the barbed end of the actin filament (at the Z line in muscle) downward and to the left in Fig. 2. Truncated Dictyostelium myosin II bound to ADP-aluminum fluoride [ADP AlF4 (95)] and ADP-vanadate [ADP VO4 (96)], as well as chicken smooth muscle myosin constructs in both the ADP VO and ADP BeFx states, are in the configuration depicted in Fig. 2B (31). This state most likely represents the transition state during cleavage of the – phosphoryl bond of ATP. It exhibits a major structural change relative to the near-rigor state involving rotations between the subdomains of S1. The lever arm is pointed away from the barbed end of actin, a position compatible with the beginning of the working stroke, so this state is termed the ‘‘prepowerstroke’’ conformation. The -phosphate sensor, switch II, is located close to the substrate (Fig. 2B) and conserved switch II residues (Gly457 and Glu459 in Dictyostelium myosin II) form hydrogen bonds with an essential water molecule and position it for in-line attack of the -phosphate. Switch II is positioned farther from the active site in the near-rigor state and crystals of truncated Dictyostelium S1 trapped in this conformation are incapable of hydrolyzing ATP, demonstrating that interactions between switch II and the -phosphate are essential for ATP hydrolysis (102). Motion of switch II toward the -phosphate of ATP in the prepower-stroke state closes the large cleft within the 50 kDa domain, allowing formation of hydrogen bonds and salt bridges between the upper and lower 50 kDa segments. This closure traps product phosphate (Pi) within the protein after hydrolysis, which explains the otherwise puzzling finding that Pi can rotate in the active site much faster (>100 s1) than it dissociates [50% overall efficiency accomplished by a whole muscle. Mechanical and X-ray diffraction studies that seemed to indicate high values of F (150, 151, 154) were then difficult to rationalize with the energetics. A provocative idea that could explain these results was the possibility that cross-bridge mechanical events are not tightly coupled to the splitting of ATP. Then several attachments, working strokes, and detachments, each liberating 10–20 1021 J, could occur during each ATPase cycle. In fact,
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several studies of sliding distance in vitro (168, 169) and in muscle fibers (166, 170) suggested such loose coupling. The interpretation of these muscle fiber studies depended on a high value of F (>0.5) and they could be reinterpreted without loose coupling if F is less than 0.3 (155). Thus, mechanical studies on muscle fibers imply molecular events during contraction that are compatible with the dimensions of the myosin head and the lever arm hypothesis. The uncertainty of F during contraction, however, prevents precise estimation of the force and energy transduction during individual actomyosin interactions. The long step sizes with muscle myosin and loose coupling implied by some in vitro experiments (168, 169) are still controversial (37). Based on the steady-state mechanical and energetic properties of contracting muscle, A. F. Huxley (16) postulated a quantitative model for the cross-bridge cycle that accounted for many of the mechanical and energetic properties of muscle contraction. In this model, the rate constants for attachment and detachment are functions of mechanical strain in the individual cross-bridges. In a shortening muscle, the myosin heads are postulated to attach to actin at positive strain, exert sliding force immediately upon attachment, and then detach. At high velocities, the attached cross-bridges are carried past the position giving zero force by the sliding generated by other cross-bridges. They are thus dragged into a region of negative strain where they detach rapidly. The negatively strained condition was mentioned earlier regarding the apparent distortion of the myosin S1 crystal structure of Fig. 2C. The maximum velocity of shortening in the Huxley 1957 model is determined by a balance of positively and negatively strained attachments. Two further mechanical effects determine the shape of the force–velocity curve: at high velocities the number of cross-bridges diminishes, and the force produced immediately upon crossbridge attachment decreases. These concepts are still applicable to explain the steady isometric force, the force–velocity curve, and the amount of energy liberated including the increased heat produced during shortening. This type of model is termed a ‘‘thermal ratchet’’ (114) because random diffusional motions of the detached cross-bridge are captured and rectified to produce directional motion and work by the strain dependence of the attachment and detachment rate constants. Later experiments, particularly mechanical transients [(160), Fig. 6 here], the crystal structures suggesting internal flexion in S1 (Fig. 2), and in vitro studies showing velocities and step sizes proportional to the length of the LCD (171, 172), suggested that attachment and force generation are two separate processes. Power-stroke models, such as the lever arm hypothesis do not rule out contributions from thermal ratchet motions of the head as discussed further below.
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X. Biochemical Rate Constants in Muscle Fibers A. STEADY-STATE ATPASE ACTIVITY The average rate of ATP hydrolysis in relaxed skeletal muscle fibers is QR ¼ 0.002 s1 per myosin head [intact frog muscle at 0–4 C (173)] to 0.1 s1 per myosin head [skinned rabbit fiber at 20 C (174)]. Intact muscles tend to exhibit lower resting ATPase activity than skinned fibers, presumably because of damage or loss of a regulatory factor during preparation of the skinned fiber. When the muscle is activated at fixed length, the average ATPase rate increases to QA ¼ 0.6–3.1 s1 [intact or skinned frog fibers at 0–15 C or rabbit skinned fibers at 13–25 C (reviewed in 23, 167 )]. The actomyosin ATPase rates of intact and skinned fibers are more similar during activity than in relaxation. If the number of myosin heads participating and the proportion of the ATPase cycle they spend attached to actin are known, then the attachment and detachment rates can be calculated. During active contractions only one head of each myosin molecule probably attaches to actin (131, 175, 176) and due to the mismatched periodicities of the two sets of filaments in vertebrate-striated muscle, it is possible that some of the myosin heads do not interact with actin at all. Ignoring the latter possibility for simplicity, take the overall ATPase rate to be 3 s1 per head [rabbit skinned fiber at 20 C, 200 mM ionic strength (113)]. Let F ¼ 0.25 of the cross-bridges be attached during a maximally activated isometric contraction. Then the attachment rate constant for a detached myosin head participating in the cross-bridge cycle is QA/(1F) ¼ 4 s1 and the detachment rate constant is QA/F ¼ 12 s1. If F ¼ 0.75, then the detachment rate is 4 s1 and the attachment rate for cycling heads is 12 s1. During shortening, the ATPase rate increases mainly due to accelerated detachment rather than attachment because the mechanical stiffness and X-ray diffraction data indicate that fewer cross-bridges are attached during shortening than in isometric contraction (139, 177, 178). B. ATP-INDUCED ACTOMYOSIN DISSOCIATION In the absence of ATP, the affinity of myosin for actin is very high, so virtually all of the myosin heads attach to actin forming rigor complexes. ATP can be rapidly liberated within a rigor muscle fiber by laser photolysis of the photolabile molecule, caged ATP (179). Changes of tension and stiffness (180, 181), as well as structural signals such as X-ray diffraction (182, 183), birefringence (184), and polarization of fluorescence from extrinsic probes (185, 186) all indicate that ATP binds to the rigor cross-bridges rapidly and detaches them from actin. The second-order rate constant
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for ATP-induced dissociation of myosin from actin is 105–106 M1 s1 (180, 181). Binding of caged ATP to actomyosin before photolysis should be taken into account in calculating this rate constant (187). The ATP dependence of the elastic modulus of muscle fibers, determined by sinusoidal analysis, also implied ATP binding and detachment are fast (188). The rigor state represents the minimum free energy in the actomyosin cycle and almost half of the free energy available from the ATPase reaction is used to exit from this state and dissociate myosin from actin (189). The remainder of the free energy is associated with release of Pi and ADP from the actin products complex, AM ADP Pi. C. ATP HYDROLYSIS The elementary step of ATP cleavage to protein-bound ADP and Pi in muscle fibers is rapid (190) and readily reversible (174). When ATP was liberated from caged ATP, a burst of ADP production was detected by rapid freezing and analysis of the nucleotide content (‘‘flash and smash’’ technique). The amplitude of this burst corresponded to approximately one ADP molecule per myosin head. The rate of ADP formation indicated that k2d þ k2d (Fig. 4) ¼ 60 s1 [12 C, 200 mM ionic strength (191)]. As discussed above for the case of S1 in solution, exchange of oxygen atoms between the solvent and ATP bound to myosin indicates that the hydrolysis step reverses repeatedly before Pi release. Reversibility implies that the free energy of M* ATP and M** ADP Pi are similar [G ¼ kBT ln(k/k þ )]. The steady-state ATPase rate is much slower than ATP binding and hydrolysis. Thus, the rate-limiting step for ATP turnover follows the hydrolysis step in both relaxed and isometrically contracting muscle fibers. D. Pi RELEASE Product release accounts for more than half of the free-energy liberation in the actomyosin ATPase cycle. Most of this free-energy change is accounted for by Pi release from AM ADP Pi. Thus in solution, the dissociation constant for Pi (steps k þ 4a and k4a in Fig. 4) is 100 mM or greater. Yet Pi binds to AM0 ADP much more readily in actively forcegenerating muscle fibers than in isolated actomyosin (174, 188, 192) and it rephosphorylates ADP to generate ATP in the active site (174, 193). The reversibility of Pi release in an actively force-generating fiber indicates that this step is much closer to equilibrium than for actomyosin in solution. The rate of Pi release is slower in the fiber and binding of Pi to AM0 ADP is faster, presumably due to mechanical strain as discussed above in the section comparing a protein suspension with the filament lattice.
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Addition of Pi to the medium of a contracting skinned muscle fiber reduces tension production without reducing shortening velocity and with only a minor reduction of the ATPase rate (113, 149, 194–199). In most of these studies, the dependence of force on Pi concentration saturates at approximately 10 mM Pi. Added Pi suppresses the ATPase activity in slow muscle fibers more than in fast muscle (200). Enzymatic depletion of Pi from within a skinned muscle fiber increases the active force (201). These studies provisionally suggest a linkage between Pi release and the transition to the force actomyosin-generating state. When a skinned muscle fiber is activated by photolysis of caged Ca2 þ or by caged ATP in the presence of Ca2 þ , tension increases to a plateau force with rate constant 50–100 s1 (181, 202, 203). Addition of Pi to the medium suppresses the steady-state tension, and markedly accelerates the approach to the steady state (149). Considering the reaction scheme of Fig. 4 and its kinetics, the observed rate constant for force development in this type of experiment is given approximately by kobs ffi k þ 9 k þ 3a/(k þ 9 þ k9) þ k3a k4a [Pi]/(k þ 4a þ k4a [Pi]) if steps 4 and 9 are rapid (108). If Pi binds to the actively force-generating intermediate, AM0 ADP, kobs is expected to increase. Tension declines as [Pi] is increased because leftward shift of the equilibrium between the force-generating states AM0 ADP Pi $ AM0 ADP þ Pi (step 4 in Fig. 4) reverses the previous transition (step 3) by mass action. The resultant state, AM ADP Pi, is weakly bound and does not produce force. When the fiber is initially in the rigor state with caged ATP in the absence of Ca2 þ , photoliberation of ATP causes the tension and stiffness to decline. This relaxation is also accelerated by Pi, supporting the idea that binding of Pi to the cross-bridges reverses steps 4 (Pi release) and 3 (transition leading to the force-generating state). Thus, the acceleration of muscle fiber relaxation and activation, and the decrease in steady force strongly suggest that Pi release and force generation are closely linked. A sudden increase of [Pi] produced within a contracting fiber by photolysis of caged Pi reduces force with a rate constant that increases as the Pi concentration is increased (20–80 s1 at 10 C and 80–170 s1 at 20 C) and saturates at 10–20 mM (108, 199, 204–207). This experiment indicates a close kinetic linkage between Pi release and force production. This relationship between the kinetics of the Pi release step and generation of mechanical force has also been investigated using sinusoidal analysis (188, 208), temperature jump (209, 210), and pressure jump (211) perturbations over a range of Pi concentrations. These studies lead to a model in which the weakly bound AM ADP Pi state (Fig. 4) isomerizes into a strongly bound, force-generating state (AM0 ADP Pi) which is stabilized by release of Pi to form AM0 ADP. This sequence is also depicted in the scheme of Fig. 3,
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states 3 ! 4 ! 5, and in Fig. 5 as weakly bound A–M ADP Pi ! forcegenerating A M ADP Pi ! force-generating A M ADP. A direct spectroscopic measurement of Pi release from actomyosin is provided by a fluorescently labeled mutant of Escherichia coli phosphatebinding protein [PBP (212)]. Fluorescence of the labeled PBP analog increases 5-fold on specific binding of Pi. Muscle fibers were activated by caged ATP photolysis and the time course of Pi release from the crossbridges was detected in real time from the fluorescence increase. Surprisingly, the initial rate of Pi release within skinned muscle fibers was about 10-fold higher than the steady-state ATPase rate per myosin head for 2–3 turnovers (213). These results seemed to contradict the flash and smash experiments mentioned earlier that identified an approximately stoichiometric burst of ADP production. More recent experiments have revealed that the Pi release above one molecule per myosin head is probably linked to initial shortening of the muscle fiber following activation (214). The PBP technique has been applied to many enzyme systems including isolated actomyosin (212), myofibrils (143), and GTP-binding proteins (215). Orthovanadate (Vi) acts as a chemical analog of Pi, binding tightly to M ADP or AM ADP and forming a stable M ADP Vi complex both in solution (216) and in muscle fibers (217–220). Based on its crystal structure, the myosin M ADP Vi complex probably represents the transition state between M ATP and M ADP Pi (96), with the lever arm ‘‘primed’’ in the prepower-stroke position [Fig. 2B here and (31)]. In skinned fiber experiments, Vi bound during cross-bridge cycling, but not in relaxation, rigor, or rigor with added ADP (218). Thus, Vi binds to a relatively longlived intermediate (AM0 ADP in Fig. 4 or A M ADP in Fig. 5) present only during active force generation. These results also confirm that Pi dissociates before ADP as in solution. The M ADP Vi complex binds weakly to actin and suppresses shortening velocity much less than ATPase activity (219–222). Vi release from M ADP Vi is accelerated by actin, as Pi release is from M ADP Pi (218). The experiments described in this section provide considerable evidence supporting the hypothesis that Pi release from AM0 ADP Pi is coupled to the structural change leading to generation of force. Reversibility of the Pi release step also provides a straightforward explanation for fatigue of highly active muscles. If the rate of Pi production by actomyosin, coupled to phosphoryl transfer by creatine phosphokinase, exceeds the rate of oxidative phosphorylation, the Pi concentration rises from its normal 1–3 mM value up to 10–20 mM (223–226). Pi binds to the force-generating AM0 ADP or A M ADP intermediate increasing the population of low force AM ADP Pi states (Figs. 3–5). Although other metabolic alterations [e.g., reduced pH (164, 226)] contribute to fatigue in various circumstances,
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the accumulation of Pi is a significant factor in the rapid reduction of force-generating capacity in highly active skeletal muscle and ischemic myocardium (225, 226). E. ADP RELEASE For a wide variety of muscle types, the shortening velocity is proportional to the rate of ADP release from actomyosin in solution (227), suggesting that the lifetime of the attached cross-bridge is determined by this step. In rabbit skinned fibers, elevation of MgADP concentration slows active shortening and sinusoidal response with apparent half-saturation for inhibition (KI) at 200–300 M (195, 228). Isometric force increases with added ADP due to a prolonged attachment lifetime. As expected from the structure of myosin with a single nucleotide-binding site per head, elevated MgATP counteracts the effects of MgADP (195, 229, 230), consistent with competitive inhibition. These observations suggest that ADP release may be the biochemical step in the reaction pathway that limits the isometric ATPase rate and is accelerated during shortening as expected from the energetics of muscle contraction described earlier. Evidence for strain dependence of the ADP release rate was provided in a study of activation and relaxation of muscle fibers by photolysis of caged ATP in the presence of ADP (109). When ADP was present in the medium, ATP-induced detachment of rigor cross-bridges bearing positive strain was much slower (13–45 s1) than from those put under negative strain (160–400 s1). This detachment was partly limited by the rate of ADP dissociation from the cross-bridges, suggesting a strong effect of the force on the ADP release rate. The kinetics of activation in the presence of Ca2 þ could also be explained by such a strain-dependent ADP dissociation rate (109). However, X-ray diffraction experiments did not detect slow detachment limited by slow ADP release (182). An alternative interpretation of the strain dependence in the caged ATP experiments is rapid generation of positive force after ATP is photoliberated, rather than detachment of negatively strained cross-bridges (231). In smooth muscle myosin, tilting of the light-chain domain upon ADP release (discussed further below), higher affinity of ADP for actomyosin, and less weakening of the myosin-ADP affinity by actin compared to skeletal muscle myosin suggest that the strain-dependent mechanism of ADP release is enhanced in smooth muscles (110). Strain-dependent ADP release has also been suggested to play a role in processive motility of unconventional myosins (37, 232, 233). In the model of Fig. 3, ADP release from state 5 is slow unless the filaments slide, thereby relieving force in the cross-bridge. ADP release is faster from state 6. This mechanical effect
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on the biochemical rate constant may correspond to control of ADP release by an isomerization between AM0 ADP and AM ADP (Fig. 4) or in A M ADP (Fig. 5). Cross-bridge lifetime is controlled by the effective rate of ADP release because following this step, ATP binding rapidly detaches the nucleotide-free AM head. In skeletal muscle fibers in rigor, addition of ADP causes a small diminution of force, compatible with less than 0.2 nm of motion at the myosin head–rod junction (109, 229, 234–237). X-ray diffraction (238–240), birefringence (241), EPR spectroscopy (242), and fluorescence polarization (185) experiments have also indicated that tilting of skeletal muscle myosin heads on binding of ADP is slight. These experiments suggest that the ADP release step does not contribute actively to force generation or filament sliding. Corresponding experiments in smooth muscle and some nonmuscle myosins reveal larger tilting motions of the light-chain domain on ADP dissociation from AM ADP as described later. In intact muscle subjected to intense activity, the myoplasmic ADP concentration increases, raising the possibility that it contributes to fatigue. Even at the highest rates of contractile activation, though, the high phosphorylation potential of creatine phosphate (CP) and the enzymatic activity of creatine phosphokinase (CPK) within the sarcomeres and mitochondria normally maintain the concentration of ATP above 3 mM and the concentration of ADP below 200 M. As the two nucleotides compete for the same site on myosin, this level of myoplasmic MgADP is probably not sufficient to contribute substantially to decreased work output (195, 225). The concomitant elevation of Pi and H þ concentrations are more important in causing recoverable fatigue of the muscle. In pathological or ischemic conditions, though, CP and CPK activity may not be sufficient to maintain the ATP to ADP ratio.
XI. Structural Changes Leading to Force Generation and Filament Sliding Early models of the cross-bridge cycle postulated that a structural change of the myosin head while attached to actin caused the filaments to slide (16, 20). Electron micrographs and X-ray diffraction patterns of insect flight muscle showed that the myosin heads were mostly perpendicular to the fiber axis in relaxation and tilted toward the Z line in rigor (19). This finding suggested that detached heads, oriented perpendicular to the fiber, attached to the thin filament during active contraction and then tilted toward the Z line to produce force or sliding. This attractive hypothesis has stimulated
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many studies to identify the expected motion and to determine which parts of the cross-bridge tilt. But conclusive evidence for tilting motions of the magnitude (30–60 ) required to slide the filaments by 5–10 nm was difficult to find. Recently, however, evidence for appropriate tilting motions of both the light-chain domain and the catalytic domain have been obtained by several methods. Crystal structures of S1 (Figs. 1E and 2) indicate that the catalytic domain and the light-chain domain can tilt relative to each other. In the various conformations detected, the LCD and the C-terminal portion of the MD (the converter domain) tend to move together, but not with absolute rigidity (31). Independent mobility of the MD and LCD has also been detected using EPR spectroscopy (243) and electron microscopy (244). Thus, structural studies in the organized contractile apparatus have focused on motions of the LCD and MD during force generation. A. TILTING
OF THE
LIGHT-CHAIN DOMAIN
Fluorescent and EPR probes bound to the LCD are disordered in relaxed muscle indicating high mobility of the myosin heads (245–247). The probes become more ordered in rigor (in the absence of ATP) due to attachment of the heads to actin (185, 186, 245). During contraction, it might be anticipated that a new orientation would emerge, corresponding to the position of the heads before the transition into the force-generating state. However, angular distributions of the probes during steady contraction are quite similar to those in relaxed muscle indicating highly mobile or disordered heads (248, 249). Although the overall distribution of probes cannot be described solely by the sum of relaxed and rigor components (247), only a small fraction of heads (30 ) compatible with the imposed filament sliding and a 9–10 nm lever arm (186, 252). Comparably small signal deflections were also obtained with bifunctional rhodamine (253) bound to pairs of cysteine residues placed into the RLC sequence (254). The average orientation of the bifunctional probe is the same as the line joining the two engineered cysteine residues, allowing insertion of the probe into the RLC at predetermined local angles. By combining data from several such probes in separate experiments, the bifunctional probe technique enables determination of both tilt and twist of the protein domain within the frame of the muscle fiber (254, 255). With S1s from smooth muscle and some nonmuscle myosins bound to actin, the LCD rotates 20–30 when ADP binds or dissociates (100, 256, 257).
1. MUSCLE CONTRACTION
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However, as discussed in Section X, ADP release does not seem to contribute a major component of the power stroke. The angle changes detected with skeletal S1 bound to actin (257) or in RLC exchanged into the endogenous myosin heads of skeletal muscle fibers [(185); Fig. 7B here] are very small (5 mM (72). When the mant-ADP complex is preformed in the absence of Mg2 þ and chased with ADP and excess EDTA, the ADP is released within the time required for manual mixing (72). More recent stopped-flow results indicate that the rate of release of mant-ADP is 10 s1 (unpublished observations). These results indicate that Mg2 þ release is reversible with a dissociation rate of 0.04 s1. In the absence of free Mg2 þ , the release of Mg2 þ from the E ADP complex at 0.04 s1 is followed by rapid ADP release at 10 s1. In the presence of free Mg2 þ , the rebinding of Mg2 þ will reform E MgADP and suppress the net rate of ADP release. The limiting rate of 0.003 s1 at saturating Mg2 þ could represent the rate of conversion of the closed conformation to an open conformation in which both Mg2 þ and ADP are released rapidly. An alternative explanation considers the likely two-step nature of Mg2 þ rebinding with initial formation of a weak complex followed by an isomerization to a tight complex. For this model the rate of 0.003 s1 at saturating Mg2 þ would be determined by the partitioning of this weakly bound intermediate between ADP release and conversion to the tight complex. In either case the net rate of release of ADP is strongly accelerated by over 3000-fold (from 7 m apart. By anchoring 33 complexes to well-separated nickel sites that could likely accommodate only one or two 33 per site (diameters: 33, 10 nm, nickel dot 30 nm), they could observe multiple beads rotating simultaneously within a single field for long time periods [also see related commentary in Science (152)]. Compared to previous filament assays, in which only a few percent of filaments rotated, this approach appeared to avoid problems of immobile complexes caused by attachment of a filament to multiple 33 motors or by collision of closely spaced filaments. In a subsequent study this group used an array of nickel-capped SiO2 posts (200 nm high, 80 nm average diameter, >2.5 m spacing) and, instead of a bead, attached a nanofabricated metal rod (150 nm diameter, 750 or 1400 nm long) to the -Cys of each immobilized 33 complex2 (153). The height of the posts was intended to minimize contributions of surface proximity to the viscous drag on rotating rods, so that a more accurate estimate could be made for the torque and thus thermodynamic efficiency of ATPase-driven rotation.
Although the nickel-capped posts were spaced >2.5 m apart, the authors noted an unexplained low yield ( 1%) or rotating rods, with 80% of rods apparently anchored at more than one point (i.e., lack of Brownian rotary fluctuations). The immobile rods were probably anchored to the subunit of multiple 33 complexes bound per post. The diameter of the posts (50–120 nm, 80 nm average) were large enough to bind multiple 33 complexes (diameter 10 nm) per post, and the coverslip arrayed with posts was incubated with excess 33 (1 mg ml1, 3 M). 2
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Of course, considering the site of attachment to (see * in Fig. 3) and other uncertainties in the geometry of the immobilized complex, some rods were probably not oriented parallel to the coverslip’s surface, as assumed (153), and the ends of some rods still could have approached the surface. Nevertheless, for distinct rods of the same length (and relative point of attachment along the rod), the observed rotational rates were very similar ( 5% for 750 nm rods), indicating little variation due to surface drag effects. Rotation of short and long rods yielded a consistent estimate of 20 pN nm for the torque, and the authors estimated an upper limit of 80% for the thermodynamic efficiency of the motor, compared to estimates closer to 100% efficiency from most actin filament assays discussed above. The collaborating groups of Kinosita and Yoshida have also continued to improve upon the dynamic microscopy assays for subunit rotation that they pioneered. Their most fruitful approach thus far was designed to avoid the large viscous drag imposed by micron-sized filaments or beads in order to focus on the kinetics of unimpeded stepwise rotation (154). Thermophilic 33 complexes were anchored by -His10 tags to a nickel-coated coverslip as before, and the attachment of streptavidin to involved two engineered, biotinylated cysteines in ’s protruding stalk region in order to achieve a more fixed, oblique attachment of the rotary probe. The rotary probe consisted of a colloidal gold bead, 40 nm in diameter, coated with biotinylated BSA for attachment to through streptavidin. Gold beads were observed by laser dark-field microscopy: light scattered by each bead produced a diffraction-limited spot ( 0.3 m) and, due to the high intensity and signal/ noise ratio, movement of the spots could be recorded at up to 8000 frames per second. As noted for the earlier bead assays, rotation of within immobilized 33 will cause an obliquely attached bead to precess around a central point. Following the centroid (bright center) of each bead’s image, observed rotational paths traced diameters of 25–55 nm, consistent with a maximum possible diameter of 60 nm, as estimated from the height of immobilized 33 and the dimensions of the linker proteins between and the bead (154). Compared to assays with 1 m filaments or beads, the 40 nm probe should reduce the viscous drag on ’s rotation by 3–4 orders of magnitude, and assays with a range of bead sizes (0.04–0.6 m) demonstrated that beads of 0.1 m diameter do not impede ’s rate of rotation at saturating or subKM concentrations of MgATP. Further, use of larger, rate-limiting beads yielded smooth rotation whereas rotation of 40 nm beads was stepwise even with saturating MgATP. Thus, with a 40 nm bead fixed to , observed rotation rates correlated closely with one-third of the ATPase rate (for beadfree 33 in solution) over a nM–mM range of [ATP], and the data fit to simple Michaelis–Menten behavior (Vmax 130 rps, KM 15 M MgATP). These results support the contentions that one ATP is hydrolyzed per 120
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rotational step and that bisite catalysis is sufficient to drive rotation (i.e., loading of all three sites with MgATP is not required for significant rates of cooperative hydrolysis and subunit rotation). With [ATP] KM, only three 120 steps could be observed per rotary cycle, even at the fastest recording rate. However, with [ATP] near or below KM, many 120 steps actually occurred as substeps of 90 and 30 , and there are several indications that these substeps are not artifacts of the assay [see discussion in (154)]. Of course, the apparent 90 and 30 substeps may reflect rigid rotation of primarily, but may include some contributions from conformational twisting and/or tilting of . The average dwell time before 90 substeps was longer with lower [ATP], indicating that the largest rotary substep is driven by binding of MgATP to a (second) catalytic site. The dwell time of a few ms before each 30 substep was independent of [ATP], and so probably correlates with the limiting step(s) of product release from the (firstloaded) catalytic site. Under both conditions, the dwell periods accounted for most of the time of each rotary cycle, while each rotary substep required only 0.25 ms. The distributions of dwell times (>12,000 total counted) were analyzed for rotating complexes at various [ATP] and a minimum of three rate constants were needed to fit the combined data: an ATP-binding rate of 3 107 M1 s1 that limits the 90 step, and two intrinsic reactions of 1 ms each that precede the 30 step. The apparent ATP-binding rate is consistent with the Michaelis–Menten kinetic parameters noted above for this study and is similar to the rate measured for ATP binding to a single catalytic site of MF1 (155). The latter two reactions noted above could represent dissociation steps for both products, and the similar rates would be consistent with the apparent random order of dissociation for ADP and Pi (156). Alternatively, the last reaction could correlate with release of both products, and the intermediate reaction could relate to the hydrolytic step or, perhaps more specifically, to a putative commitment step needed to convert the ‘‘tight’’ site (catalyzing rapidly reversible hydrolysis) to a state that ensures specific release of products in the final step (5). A subsequent rotational study by Yoshida and Kinosita’s groups further suggests that the 30 rotary substep coincides with or follows dissociation of ADP (157). This study documented the correlation of long pauses (>10 s) between rotational periods (long pauses were omitted from analysis in most previous studies) with an inhibited state of the enzyme that involves MgADP tightly bound to a catalytic site in the absence of bound Pi [reviewed in (5, 40)]. The results suggest that the MgADP-inhibited state can form stochastically after the 90 rotary substep (157). Thus, most likely in a case in which Pi dissociates first by chance (158), bound MgADP may reorient or collapse to the inhibited state rather than dissociate, and thus temporarily block the 30 rotary substep.
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3. Complementary Spectroscopic Approaches Despite the many strengths of the filament/bead assays in ‘‘visually’’ characterizing general features of subunit rotation in FOF1, those assays are not amenable to use with FOF1 in native membranes or reconstituted in liposomes, which would be necessary for studies of rotation during ATP synthesis. Further, filament/bead assays monitor rotation of one rotor subunit relative to an anchored stator complex, and so cannot provide more detailed data about the relative movements between specific rotor and stator subunits. Such limitations indicate the need for complementary spectroscopic/microscopic assays that can be more easily adapted to studies of membrane-bound FOF1 and provide dynamic probes of relative motion for specific sites on rotor and/or stator subunits. Thus far, such spectroscopic studies discussed below have focused on attachment of a small fluorescent group to a single rotor subunit of F1 or FOF1, surface immobilization of labeled complexes, and measurements of fluorescence anisotropy (159) at the single-molecule level (160) to monitor rotation of the labeled subunit. This type of fluorescence anisotropy approach still monitors movement of only one labeled rotary subunit and cannot directly determine the direction of rotation, but avoids the problem of viscous drag caused by most of the larger bead/filament probes, and it can be applied to membrane-bound FOF1. Recently, successful use of fluorescence resonance energy transfer (FRET) in single-particle assays has been demonstrated in dynamic studies of site–site interactions in a number of protein/enzyme systems (160–162). The use of FRET for assays of subunit rotation in F1 and FOF1 should be forthcoming, and FRET offers the ability to monitor dynamic movements and relative rotation between specific pairs of rotor/stator sites on the enzyme. In the first single-particle study of subunit rotation in which the probe was smaller than F1 itself, CF1(, ") complexes were labeled on C322 (next to ’s C-terminus) with tetramethylrhodamine-5-maleimide, and labeled complexes were nonspecifically adsorbed to a glass coverslip at low density (163). Fluorescence of the probe on was then measured for individual immobilized CF1(, ") complexes, using a photon counting confocal microscope. Due to the linear polarization of the exciting laser, the probability of exciting the fluorophore and hence the intensity of its fluorescence emission depended on the orientation of the fluorophore relative to the polarization plane of the exciting laser. Rotation of would reorient the fluorophore and alter the fluorescence intensity observed. In the presence of mM ATP but not ADP, stepped transitions in fluorescence intensity were observed for labeled, immobilized CF1(, "), and the frequency of transitions was consistent with the reduced ATPase turnover rate of immobilized CF1(, "). Observed stepping transitions between three
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distinct emission levels were rare (3 of 200 trials), but this is not surprising since CF1(, ") complexes were nonspecifically adsorbed to the glass slide: each immobilized complex could have a different orientation of the label on relative to the plane of the exciting light, and some possible orientations would not give significant differences in emission intensity when rotated between distinct subunits. Although the three-stepped pattern likely represents rotation of between three distinctly oriented catalytic sites of the immobilized CF1(, ") complex, the undefined orientation of immobilized complexes makes it difficult to develop a more explicit correlation of the fluorescence changes with subunit rotation. For example, the apparent rotation of C322 in CF1 (163) seems to be at odds with a more recent study by the same group indicating that several of ’s C-terminal residues may not rotate with the rest of in EcF1 [(97), see Section V, B, 1]; however, the studies would agree if the CF1 complexes that displayed stepped fluorescence changes were nonspecifically anchored to the coverslip through ’s protruding stalk rather than through 33. This study was also plagued by a high rate of photobleaching of the fluorophore in the confocal system, such that each individual complex could only be monitored for 5 s. Despite these problems, the assay was also used to test for rotation of fluorescently labeled and " after reconstitution with CF1(, "). Results indicated rotation for ", although no three-stepped transitions were seen, and there were no indications for rotation of . Single-fluorophore fluorescence anisotropy measurements for ’s rotation were more thoroughly developed in a study with the thermophilic 33 system originally used for filament rotational assays (164). As before, His10-tagged subunits were used to anchor individual 33 complexes, in a defined orientation, on a Ni2 þ /chelate-coated coverslip. To optimize the response of fluorescence anisotropy to rotation of labeled , several -cysteine/fluorophore pairs were tested to identify one that exhibited the highest anisotropy value for labeled 33 in solution. An anisotropy value of 0.32 (vs. 0.4 theoretical maximum) was found for 33(I210C) labeled with Cy3-maleimide, indicating the probe is nearly immobile in its attachment to (at least in the nanosecond lifetime range of Cy3’s excited state). A wide-field epifluorescence microscope (modified to reduce background fluorescence) was used for anisotropy assays for ’s rotation. Compared to the confocal system used in the CF1 study described above, this setup allows simpler monitoring of many separate 33 complexes, and photobleaching problems were reduced >10-fold, so that fluorescence of a single 33(I210C-Cy3) complex could be monitored for at least 1 min. Also, two distinct methods were used to monitor fluorescence anisotropy in separate experiments. In the first, the polarization axis of the exciting light is rotated continuously in the sample plane, and total fluorescence emission intensity
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of each complex is measured (no polarizer on detector). This is analogous to the assay used in the earlier study with CF1; the efficiency of light absorbtion by the I210C-Cy3 probe, and thus the intensity of its fluorescence emission, will be highest when the exciting light is polarized parallel to the probe’s absorption transition moment. Thus, the fluorescence intensity of an immobile probe would oscillate with exactly the same period as the rotating excitation polarizer, whereas rotation of I210C-Cy3 within the immobilized 33 complex would shift the pattern. In the second method, the exciting light is essentially unpolarized, and the probe’s fluorescence emission is observed with horizontally and vertically polarized detectors, so that the relative orientation of the probe’s emission dipole is monitored directly as labeled rotates. The two methods gave consistent and complimentary results. Although the available time resolution limited assays to use of low [ATP], the assays confirmed that stepwise rotation of occurs in the absence of a viscous load on , and an estimate for the limiting rate of ATP binding was consistent with values obtained from filament and gold bead assays discussed earlier. One nagging technical problem with anisotropy approaches is that one must assume unidirectional rotation, so that an occasional, reverse step, as observed in filament assays (131, 132), may be misinterpreted as two faster forward steps. The first application of single-fluorophore anisotropy to monitor rotation in intact ATP synthase has now been published (165), using Na þ transporting, hybrid FOF1 complexes. FO was reconstituted from isolated P. modestum subunits; Cy3-labeled cD2C subunit was combined with 15–25 molar excess of wild-type c so that most FO complexes formed would contain no more than one Cy3-labeled c subunit. F1 was expressed and isolated with E. coli subunits except that was from P. modestum. His10-tagged subunits were used to anchor solubilized FOF1 or liposomereconstituted FOF1 ( 1 FOF1 per liposome) to the Ni2 þ -coated coverslip. Rotation of Cy3-labeled c subunit was monitored with a confocal microscope, using two polarized detectors as in the second method described above. As noted for the earlier confocal studies with CF1, photobleaching and signal/noise limitations were significant, so that most observations were20 A˚ from the closest D380 in DP of the docked MF1 structure, whereas the -carbons of cysteines must be
