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Annu. Rev. Biochem. 1999. 68:687–728
Copyright © 1999 by Annual Reviews. All rights reserved.

STRUCTURAL MECHANISMOF MUSCLE CONTRACTION
M. A. Geeves
Annu. Rev. Biochem. 1999.68:687-728. Downloaded from www.annualreviews.org
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Department of Biosciences, University of Kent, Canterbury, CT2 7NJ, United Kingdom;
e-mail: m.a.geeves@ukc.ac.uk

K.C. Holmes
Max Planck Institute for Medical Research, 69120 Heidelberg, Germany; e-mail:holmes@mpimf-heidelberg.mpg.de

Key Words

actin, myosin, structure, muscle contraction, swinging cross bridge,
molecular mechanism, kinetics, mutations■ Abstract X-ray crystallography shows the myosin cross-bridge to exist in two
conformations, the beginning and end of the “power stroke.” A long lever-arm undergoes a 60° to 70° rotation between the two states. This rotation is coupled withchanges in the active site (OPEN to CLOSED) and phosphate release. Actin binding
mediates the transition from CLOSED to OPEN. Kinetics shows that the binding of
myosin to actin is a two-step process which affects ATP and ADP affinity. The structural basis of these effects is not explained by the presently known conformers ofmyosin. Therefore, other states of the myosin cross-bridge must exist. Moreover,cryoelectronmicroscopy has revealed other angles of the cross-bridge lever arm
induced by ADP binding. These structural states are presently being characterized bysite-directed mutagenesis coupled with kinetic analysis.

CONTENTS
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688
Scope of Review . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688
The Swinging Cross-Bridge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 689
Structures of Actin and Myosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 691
Atomic Structure of Actin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 691
Atomic Structure of Myosin S1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 692
Actomyosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 699
Truncated S1—a Minimal Motor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 700
Crystallography of Truncated S1 Shows Two Conformations:
OPEN and CLOSED . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 700
Switch 2 Should Close to Enable ATP-Hydrolysis . . . . . . . . . . . . . . . . . . . . . . . 701

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and CLOSED States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 702
The Switch-2 Region Should Open for Phosphate Release . . . . . . . . . . . . . . . . 703
The Lever Arm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 703
Smooth-Muscle Myosin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 704
Molecular Cogs and Gears . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 706
Other Evidence for a Swinging Lever Arm . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 709
Effects of ADP on the Structure of Decorated Actin . . . . . . . . . . . . . . . . . . . . . 709
Use of Fluorescent Probes to Measure Lever Movement . . . . . . . . . . . . . . . . . . 709
Making the Lever Arm Longer or Shorter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 710
Myosin S1 Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 712
Allocation of States to Kinetic Schemes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 712
Sensitivity to g-Phosphate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 713
Present Structures Do Not Illuminate the Mechanismof Actin-Induced ADP Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 714
The Actin-Myosin Interaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 714
Kinetic Data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 714
Correspondence with Structural Data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 716
A Third State Is Required . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 716
A Possible Mechanism for ADP Release . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 717
Effects of Strain on Phosphate and ADP Affinity . . . . . . . . . . . . . . . . . . . . . . . . . . 718
Mutational Analysis and Sequence Comparisons . . . . . . . . . . . . . . . . . . . . . . . . . 719
General . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 719
Alternate Splicing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 719
Loops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 720
The Hinge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 722
Switch 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 723
The R245–E468 Salt Bridge . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 723
Point Mutations of G466 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 723
Cardiomyopathies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 724
Mutations Should Aid Functional Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 724

INTRODUCTION
Scope of Review
Movement in muscle is generated by the myosin cross-bridges, which interact
cyclically with thin (actin) filaments and transport them past the myosin thick filaments. During the process ATP is hydrolyzed. In the last five years protein crystallography of the myosin cross-bridge has given new insight into theseprocesses. Kinetics, spectroscopy, genetic analysis, in vitro motility assays, andchemical cross-linking have also played important roles (see 1–15).
The myosin cross-bridge is a molecular machine with communicating functional units: the actin-binding site, the nucleotide-binding site/hydrolysis, and theconverter/lever arm, which appears to amplify the small changes at the active
site into the large changes needed to transport actin past myosin. These function-

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alities were recognized and explored experimentally long before the atomic structures were known: Atomic structures now embody such concepts with machinelike linkages fabricated from polypeptide chains.
We first summarize the structural results. Then we explore the behavior of theindividual functional units in structural terms (so far as this is presently possible)and how these functionalities appear to be linked. Mutational studies begin toshow how such pathways of communication can be disrupted (16). Moreover,the natural variants of myosin—smooth/fast/slow/heart/nonmuscle myosin II andother myosins—provide new insights into both structure and function. For example, alternate splicing in Drosophila myosins identifies areas that are importantfor defining function (17).
The myosin cross-bridge may be cleaved off the myosin molecule as a soluble
molecule (myosin S1, ~120,000 Da) that contains three polypeptide chains, one
heavy and two light. Myosin S1 is a fully competent actin-activated ATPase thattransports actin in vitro motility assays. Moreover, in the absence of nucleotide,
myosin S1 forms a tight (rigor) bond to actin filaments. Biochemical and structural investigations have therefore been concentrated on myosin S1 and its interactions with actin as a minimal model for muscle. However, since it is no longer
attached to the myosin thick filament, myosin S1 cannot be an adequate model
for a strained cross-bridge. Thus data from muscle fibers (e.g. the dependence ofphosphate affinity on strain) must also be considered.
There is a rich literature on the kinetics of myosin S1 ATPase. Changes in the
intrinsic protein fluorescence play a central role in signaling conformational statesof myosin and linking these to catalytic events. To make full use of these results
we need to establish the correspondence between the crystallographic states andthe kinetic states. Moreover, the structural data urge a reinterpretation of thesekinetic schemes and oblige one to see these different processes in a wider context.
Clearly, the interaction of S1 with the actin filament is of central importance.
Unfortunately, here the structural data are of limited precision. However, by consideration of the kinetics of the actomyosin interaction, one is led to the conclusion that there must be more myosin states than the two which have so far been
revealed by protein crystallography. Finally, one can hypothesize about the natureof the missing state and the way the cross-bridge states might interconvert in
active muscle.

The Swinging Cross-Bridge
The swinging–cross-bridge theory for muscle contraction envisages that the
myosin cross-bridge binds to the actin filament in an initial conformation andthen undergoes a swinging motion that “rows” the actin filament along (10).
Myosin is a product-inhibited ATPase with an active site and mechanism similar
to that of the G-proteins (18). The ATPase is strongly stimulated by binding toactin, which is a nucleotide exchange factor for myosin. In the absence of
nucleotide the myosin cross-bridge binds tightly to the actin filament to form the“strong” or “rigor” complex (Figure 1). The binding of ATP to the ATPase site on

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Figure 1 The Lymn-Taylor cycle (19). The myosin cross-bridge is bound to actin in
the rigor, 45° “down” position (state 1). ATP binds, which leads to very fast dissociation from actin (state 2). The hydrolysis of ATP to ADP and Pi leads to a return of the
myosin cross-bridge to the 90° “up” position, whereupon it rebinds to actin (state 4).
This leads to release of the products and return to state 1. In the last transition, actinis “rowed” past myosin.

the myosin cross-bridge rapidly dissociates the actomyosin complex; myosinthen hydrolyzes ATP and forms a stable myosin-products complex (ADP.Pi);
actin recombines with this complex and dissociates the products, thereby forming the original actin-myosin complex (19). After recombining with actin, thecross-bridge undergoes a conformational change allowing the products of hydrolysis to be released, which also brings about the rowing-like stroke (this elemental event is referred to as the “power stroke”).
Although the swinging–cross-bridge hypothesis of muscle contraction features in most textbooks, it has in fact proved remarkably difficult to visualize abridge during the swing. Finally, after many years of effort involving the development of new X-ray sources (see 20 for historical review), time-resolved X-ray

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fiber diagrams from contracting frog muscle provided evidence of cross-bridge
movement (21, 22). However, such experiments do not have sufficient resolution
to define the cross-bridge movement in any detail. Owing to a number of spectroscopic and structural observations (see 23), the swinging cross-bridge wasmodified into a swinging-lever arm hypothesis in which the bulk of the crossbridge was envisaged to bind to actin without rolling on the surface during thepower stroke (as had initially been suggested); large movements were envisaged
as coming from the distal (C-terminal) part of the myosin cross-bridge moving as
a lever arm (Figure 2).
A swinging lever arm also helped to explain why substantial changes in the
cross-bridge orientation were difficult to see: Only a small fraction of the crossbridge mass moves much during the power stroke. Furthermore, it graduallybecame clear that the proportion of cross-bridges taking part in a contraction at
any one time was only a small fraction of the total (see e.g. 24), which makes theregistration of active cross-bridge movement doubly difficult.

STRUCTURES OF ACTIN AND MYOSIN
Atomic Structure of Actin
Thin filaments (f-actin) are helical polymers which have 13 actin molecules (42kDa) arranged on six left-handed turns repeating every 36 nm. The rise per subunitis 2.75 nm. The morphology of the actin helix is, rather, two intertwined, steep
right-handed helices. Along each of the morphological helices the actin monomers
are spaced by 5.5 nm. The structure of the monomer (g-actin) was solved by protein crystallography as a complex with DNase I (25) and has since been solved in
two other complexes (26, 27). The structure shows actin to consist of two similar
domains each of which contains a 5-stranded b-sheet and associated a-helices. The
phosphate moiety of a nucleotide (ATP or ADP), together with Mg2+ or Ca2+
(Mg2+ is physiological), is bound between the two b-sheet domains. Each of thedomains carries a subdomain; one is involved in actin-actin interactions, and theother in addition forms the top of the nucleotide-binding pocket.
Orientated gels of filamentous actin yield X-ray fiber diagrams to about 0.6nm resolution. Fiber diffraction patterns were calculated from models producedby placing the g-actin atomic structure in the helix for all possible orientations. A
computer search compared the calculated and observed X-ray fiber diagram tofind the best fit (28). The structure was refined to allow for the conformationalchanges taking place in actin on polymerization (29; Figure 3). The most notable
movement is that the subdomains move 0.3- to 0.4-nm closer together in f-actin,
closing off the ATP/ADP-binding site, which leads to a very low nucleotide
exchange rate (after polymerization, the nucleotide in f-actin plays a structural
rather than a metabolic role). A structure showing the reverse movement (i.e. an
opening of the ADP/ATP-binding site), which may be essential for nucleotide

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Figure 2 The major change in the cross-bridge is confined to the distal part, whichmoves as a lever arm (23).

exchange, has been reported (30). The contacts along the two long-pitch helices
are substantial in area (>30 nm2), in accord with the fact that the total pull on thefilament (~1000 pN) is transmitted through this contact.

Atomic Structure of Myosin S1
In this section we refer to the chicken skeletal myosin sequence. Where
ambiguity might occur, we preface the numbering with Sk. In later sections wealso refer to Dictyostelium myosin by Dy and chicken smooth muscle by Sm(see Table 1).

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MUSCLE CONTRACTION

Figure 3 Structure of F-actin (28, 122). F-actin has 13 actin molecules arranged on
six left-handed turns repeating every 36 nm. The rise per subunit is 2.75 nm. The
morphology of the actin helix is two intertwined, long-pitch, right-handed helices.
Along each of the morphological helices, the actin monomers are spaced by 5.5 nm.
The structure of the monomer (g-actin) (25) shows actin to consist of two similar domains, each of which contains a five-stranded b-sheet and associated a-helices. [Figure prepared by using Bobscript and Raster3D (123, 124).]

AQKPLSDDE KFLFVDKNFV NNPLA.QADW SAKKLVWVP. SEKH..GFEA ASIKEEKGDEEMAAFGEAA PYLRZSEKER IEAQ..NZPF DAZSSVFVV. HPKE..SFVZ GTIQSZEGGZIHDRTSDYH KYLKVKQGDS DLFKLTVSDK RY...IWYNP DPDERDSYEC GEIVSETSDSc

Sm297 ASEQMRNDLL LEGFN NYTF LSNGHVPIPA QQDDEMFQET LEAMTIMGFT EEEQTSILRV
Sk295 ZZPELIDMLL ITTNPYDYHY VSEGEITVPS IDDQEELMAT DSAIDILGFS ADEZTAIYZL
Dy288 ATAEEKKALH LAGPESFNYL NQSGCVDIKG VSDEDEFKIT RQAMDIVGFS QEEQMSIFKI

SW-I
*
c
c
Sm237 AKTVKNDNSS RFGKFIRINF DVTGYIVGAN IETYLLEKSR AIRQAKDERT FHIFYYLIAG
Sk235 AZTVRNDNSS RFGZFIRIHF GATGKLASAD IETYLLEZSR VTFQLPAERS YHIFYQIMSN
Dy228 AKTTRNNNSS RFGKFIEIQF NNAGFISGAS IQSYLLEKSR VVFQSETERN YHIFYQLLAG

P-loopLoop 1
Sm177 GESGAGKTEN TKKVIQYLAV VASSHKGKKD TSITQGPSFS YGELEKQLLQ ANPILEAFGN
Sk179 GESGAGKTVN TZRVIQYFAT IAASGEKKKE EQSGKMQGT. ...LEDQIIS ANPLLEAFGN
Dy179 GESGAGKTEN TKKVIQYLAS VAGRNQANGS GVLEQQILQ. .......... ANPILEAFGN

2:32 PM

Sm117 SGLFCVVINP YKQLPIYSEK IIDMYKGKKR HEMPPHIYAI ADTAYRSMLQ DREDQSILCTSk119 SGLFCVTVNP YZWLPVYNPZ VVLAYRGKKR QEAPPHIFSI SDNAYQFMLT DRENQSILIT
Dy119 SGLFLVAVNP FKRIPIYTQE MVDIFKGRRR NEVAPHIFAI SDVAYRSMLD DRQNQSLLIT

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SH3 ←
Sm 57 VTVELQENGK KVTLSKDDIQ KMNPPKFSKV EDMAELTCLN EASVLHNLRE RYFSGLIYTY
SK 60 VTVZTEG.GE TLTVKEDQVF SMNPPZYDZI EDMAMMTHLH EPAVLYNLZE RYAAWMIYTY
Dy 60 FTFKTVDGQD R.QVKKDDAN QRNPIKFDGV EDMSELSYLN EPAVFHNLRV RYNQDLIYTY

2
6
4

Sequence comparisons.

694

Sm
Sk
Dy

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GEEVES & HOLMEScontinued

Loop 2*c
Sm593 NASAWLTKNM DPLNDNVTSL LNQSSDKFVA DLWKDVDRIV GLDQMAKMTE SSLPSASKTKSk591 NISGWLEZNZ DPLNETVIGL YQZSSVZTLA LLFATYGGEA EGGGGKKGGK KKGSS.....
Dy580 EIQDWLEKNK DPLQQDLELC FKDSSDNVVT KLFNDPNIAS RAKKGAN... ..........

2:32 PMc
Sm536 GVLALLDEEC WFPKATDTSF VEKLIQEQGN HAK.FQKSKQ LKDKTE.FC. ILHYAGKVTY
Sk531 GIFSILEEEC MFPKATDTSF ZNZLYDEHLG KSNNFQKPKP AKGKAEAHFS LVHYAGTVDY
Dy523 GILALLDEQS VFPNATDNTL ITKLHSHFSK KNAKYEE..P RFSKTEFGV. .THYAGQVMY

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SWII-helix
Loop
$
§
c
Sm476 SFEQLCINYT NEKLQQLFNH TMFILEQEEY QREGIEWNFI DFGLDLQPCI ELIERPTNPP
Sk474 SFEQLCINFT NEZLQQFFNH HMFVLEQEEY ZZEGIEWEFI DFGMDLAACI ELIEZPM...
Dy465 SFEQLCINYT NEKLQQFFNH HMFKVEQEEY LKEKINWTFI DFGLDSQATI DLIDGRQ PP

50kD Linker
*
c
$ *
$
Sm416 KAQTKEQADF AIEALAKAKF ERLFRWILTR VNKALDKTKR QGASFLGILD IAGFEIFEIN
Sk415 ZGETVSQVHN SVGALAZAVY EZMFLWMVIR INQQLD.TKQ PRQYFIGVLD IAGFEIFDFN
Dy407 QHLNVEKSSS SRDALVKALY GRLFLWLVKK INNVL..CSE RAAYFIGVLD ISGFEIFKVN

c
Sm356 VSSVLQLGNI VFKKERNTDQ ASMPDNTAAQ KVCHLMGINV TDFTRSILTP RIKVGRDVVQ
Sk355 TGAVMHYGNL KFZQZQREEQ AEPDGTEVAD ZAAYLMGLNS AELLKALCYP RVGVGNEAVT
Dy348 IAGILHLGNI KFEKGAGEGA VLKDKTALNA ASTVFGVNPS VLEKALME.P RILAGRDLVA

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Comparison of the myosin head sequences for the chicken skeletal (Sk), Chicken smooth (Sm) and the Dictyostelium
cytoplasmic myosin II (Dy). The residue number for the first residue in each line is given. The parts of the molecule
discussed in the text are highlighted in color and/or labeled above.

Regulatory LC* *Sm832 YLKLRNWQWW RLFTKVKPLL Q
Sk823 FMNVZHWPWM ZLFFZIKPLL K
Dy802 YIDFKSWPWW KLFSKARPLL K

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Essential LC←→←→
Sm772 SKIFFRTGVL AHLEEERDLK ITDVIIAFQA QCRGYLARKA FAKRQQQLTA MKVIQRNCAA
Sk763 TZVFFZAGLL GLLEEMRDDZ LAEIITRTQA RCRGFLMRVE YRRMVERRES IFCIQYNVRS
Dy742 TKIFFRAGQL ARIEEAREQR ISEIIKAIQA ATRGWIARKV YKQAREHTVA ARIIQQNLRA

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SH1
Converter Domain# # →cc
Sm713 GIRICRQGFP NRIVFQEFRQ RYEILAANAI PKG.FMDGKQ ACILMIKALE LDPNLYRIGQ
Sk703 GIRICRKGFP SRVLYADFKQ RYRVLNASAI PEGQFMDSKQ ASEKLLGGGD VDHTQYAFGH
Dy684 GIRITRKGFP NRIIYADFVK RY.YLLAPNV PRD.AEDSQK ATD.VLKHLN IDPEQYRFGI

Fig 6a helix

continued

SH2
 #Sm653 KGMFRTVGQL YKEQLTKLMT TLRNTNPNFV RCIIPNHEKR AGKLDAHLVL EQLRCNGVLE
Sk646 ...FQTVSAL FRENLNZLMA NLRSTHPHFV RCIIPNETZT PGAMEHELVL HQLRCNGVLE
Dy627 ...FITVAAQ YKEQLASLMA TLETTNPHFV RCIIPNNKQL PAKLEDKVVL DQLRCNGVLETABLE 1Annu. Rev. Biochem. 1999.68:687-728. Downloaded from www.annualreviews.org
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GEEVES & HOLMES5:31 PMabove the residue indicates a Cardiomyopathy point mutations.
above the Sk-R245 and Sk-E468 indicates the salt bridge.

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KEY
Color codes for residues
W Tryptophans,* above the residue indicates it is conserved in all 3 myosins,§ above indicates the important Trp 510
C SH1 and SH2 essential thiols
blue Point mutations in Dictyostelium or smooth muscle MII$ is above the non-hydrolyser mutations# is above the Glycines in the essential SH/hinge region
blue underlined Termination sites for crystallized proteins Dy 754 and 759, Sm 790 and 819
cyan P-loop, helix shown in Fig 6a, Loop following SWII, link between upper and lower 50kD domain,
Loop following Sk-W510
red Loop 1, Loop 2 and converter domain
magenta SWI and SWII loops

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GEEVES & HOLMESThe proteolytic fragment S1 comprises the first 843 residues of the heavy
chain together with the two light chains. It is the morphological cross-bridge andcontains all the enzymatic activity of myosin (31, 32). Further limited proteolysis
breaks the S1 into three fragments named after their apparent molecular masses—
25K (N terminal), 50K (middle), and 20K (C terminal) (33). These fragments
were thought to represent subdomains of the S1. The structure of S1 (see below)
shows them rather to mark the positions of flexible loops (loop 1 and loop 2) in
the S1 (for a discussion of the function of the loops, see below; see also 14).
The first X-ray crystal structure (34) was of S1 from chicken muscle without nucleotide but with a sulfate ion in place of the b-phosphate in the activesite. The structure shows the S1 to be tadpolelike in form (Figure 4), with anelongated head consisting of a 7-stranded b-sheet and a C-terminal tail. All
three fragments (25K, 50K, and 20K) contribute to the 7-stranded b-sheet.
Numerous a-helices that surround the b-sheet form a deep cleft extending fromthe nucleotide-binding site to the actin-binding site. The C-terminal tail, whichin the intact myosin molecule is connected to the thick filament, forms an
extended a-helix that binds the two calmodulin-like “light chains.” In Figure 4,
the proteolytic fragments are color coded as follows: 25K (N terminal), green;
50K, red; and 20K (C terminal), blue. The 50K fragment actually spans twodomains, which Rayment et al (34) have called the 50K upper domain and the
50K lower domain or actin-binding domain. The actin-binding domain has beencolored grey. The N terminus lies near the start of the tail, and the first 80
residues form a protruding SH3-like b-barrel domain of unknown function [notpresent in all myosins (3)]. The rest of the 25K fragment, together with the
50K upper fragment (residues 81–486) form one large domain that accounts
for 6 of the 7 strands of the b-sheet and constitutes the bulk of the molecule.
The ATP-binding site is in this large domain near the 25K–50K fragment
boundary and contains a characteristic P-loop similar to that found in someother ATPases and G-proteins (18). The ATP-binding site is about 4.0 nm fromthe actin-binding site. The 50K lower fragment (nucleotides 487–600) actuallyforms a well-defined domain that constitutes the major part the actin-bindingsite. A large, positively charged disordered loop (625–647) follows, which isalso involved in actin binding. The first part of the ensuing 20K domain(648–689) is an integral part of the 25K–50K domain and consists of a longhelix running from the actin-binding site to a seventh strand of the b-sheet.
This is followed by a turn and a broken helix containing two reactive thiols(SH1707 and SH2697). Comparison with other structures (see below) showsthat the end of the SH1 helix forms the hinge for the ensuing lever. There follows a small compact domain (711–781) which has been termed the “converter” domain (35, 35a). This functions as a socket for the C-terminal a-helicaltail, which has been called the regulatory domain or “neck” and which carries
the two light chains. The main function of the neck, however, appears to be as
a lever arm to amplify rotational movements experienced by the converterdomain during ATP hydrolysis (see discussion below).

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Upper 50K Domain

Regulatory Light Chain

C
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N
Actin Binding Domain

Converter Domain

Essential Light Chain

Figure 4 Myosin S1 (34). Myosin S1 has an elongated head consisting of a sevenstranded b-sheet and a C-terminal tail or “neck” which carries two calmodulin-likelight chains—the regulatory light chain (magenta) and the essential light chain (yellow). The proteolytic fragments are color coded as follows: 25K (N terminal), green;
50K, red; and 20K (C terminal), blue. The 50K fragment spans two domains: the 50K
upper domain and the 50K lower domain or actin-binding domain. The actin-binding
domain has been colored grey. All three fragments (25K, 50K, and 20K) contributeto the seven-stranded b-sheet. Numerous a-helices that surround the b-sheet form adeep cleft extending to the actin-binding site. [Figure prepared by using Bobscript
and Raster3D (123, 124).]

Actomyosin
An atomic model of the actin myosin complex (Figure 5; 36, 37) was obtainedby fitting the atomic structures of f-actin and S1 into three-dimensional cryoelectron microscope reconstructions of “decorated actin.” Decorated actin is
produced by incubating f-actin with S1 in the absence of nucleotides. One
cross-bridge (S1) binds to each actin monomer. Decorated actin is taken as amodel of the rigor complex between actin and myosin. The atomic model ofchicken muscle S1 was without nucleotide and therefore was thought to beclose to the rigor configuration. This view may need some modification. The
S1 binds to the lower side of one domain of actin but with a considerable contact to the subdomain of the next actin molecule below. The actin-binding sites
and nucleotide-binding sites are on opposite sides of the sheet and are separated by 4–5 nm. The cleft in myosin extends from the ATP-binding site to theactin-binding site so that movements in this cleft could provide a physical link

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between the ATP site and the actin-binding site. The very extended C-terminal
a-helix of S1 lies distal to the actin helix and is ideally situated and orientated
to function as a lever arm.

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Truncated S1—a Minimal Motor
Genetic manipulation is not only an essential tool for analyzing function but isalso a powerful adjunct to structural studies, because it allows the production ofwell-defined units or subdomains. Unfortunately, the expression of myosin isrestricted to eukaryotic hosts because it cannot be successfully expressed inprokaryotes. Two expression systems have been used: Dictyostelium discoideum(cellular slime mold), using a plasmid to express myosin II constructs in cells
deficient in myosin II, and SF9 insect cells (Spodoptera frugperda), using baculovirus as a vector to express chicken gizzard (smooth-muscle) myosin.
Expression makes it possible to design a minimal model of the myosin motorby truncating the polypeptide chain. The truncations used mostly eliminate theneck and the associated light chains but leave the converter domain intact. The
neck makes the myosin head very asymmetric: “Neckless” myosin heads are
more globular and easier to crystallize. The expressed fragments correspond
more or less with the myosin core, which has been identified by sequence comparisons to be common to all myosins—many myosins have quite different
necks to skeletal muscle myosin, but all have very similar cores (3). Proteolytic
cleavage of chicken myosin close to the core boundary produces a kineticallynormal “motor” (38), whereas experiments with shorter constructs of expressed
Dictyostelium myosin II S1 show that damage to the converter domain leads to
S1 constructs with modified ATPases. However, the form truncated at Dy759
appears kinetically normal (39). Rayment and co-workers (40–42) initiallystudied a crystalline truncated fragment of Dictyostelium myosin II S1. Morerecently, studies from Carolyn Cohen’s laboratory, of truncated chickensmooth-muscle constructs, have yielded a wealth of new structural data (seebelow) (47).

Crystallography of Truncated S1 Shows Two Conformations:
OPEN and CLOSEDThe crystal structures of the Dictyostelium myosin truncated at Dy759 (i.e. without the neck or lever-arm) have been determined with a number of ATP analogs,
particularly ADP.BeFx, ADP.AlF4 (40), and ADP.vanadate (41) (because thecoordination geometry of the vanadate ion is bipyramidal, ADP.vanadate complexes are used as analogs of the transition state, and BeF3 is close to tetrahedral
so that ADP.BeF3 is expected to be an analog of the ATP state). Although the
ADP.BeF3 state looks similar to chicken muscle without nucleotides, as do complexes with ADP and other ATP analogs (42), the ADP.vanadate structure shows
large changes in the S1 structure. These changes are also apparent with ADP.

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Figure 5 The structure of the acto-myosin complex (36, 37) shown are (right) fiveactin molecules in an actin helix and (left) a myosin cross-bridge (S1). Note that thecross-bridge makes contact with two adjacent actin monomers. [Figure prepared byusing Bobscript and Raster3D (123, 124).]

AlF4. The g-phosphate–binding pocket closes, causing a partial closing of the
50K upper/lower-domain cleft, and there are large movements in the C-terminal
region. The 50K upper domain and the actin-binding domain rotate a few degrees
toward each other with the helix Sk648–666 as a fulcrum in a way that closes thenucleotide-binding pocket (Figure 6)—a movement of some 0.5 nm. At the sametime the outer end of the long helix (residues Sk475–509, which, by analogy with
the G-proteins, are called the switch-2 helix) and its associated loop (Sk509–519)
bend out 24°. The switch-2 loop suffers extensive reorganization. We refer to thisas the CLOSED form.

Switch 2 Should Close to Enable ATP-Hydrolysis
Smith & Rayment (41) note the similarity of the active site of CLOSED myosinwith those of Ras p21 and other G-proteins. The differences between the OPEN
and CLOSED forms in the neighborhood of the active site reside almost
entirely in the conformation of the linker region (Sk465–470), which joins the
50K upper and lower domains. The authors point out that this region is

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structurally equivalent to the switch-2 region in the protein product Ras p21,
with which it also has a very strong sequence homology. The mutual rotationand closing of the 50k upper/lower-domain cleft causes movements of about
0.5 nm around the g-phosphate–binding pocket. In the chicken crystal structure (OPEN form), which has no bound nucleotide (but with a bound sulfate in
the g-phosphate position) and which may be close to the end-of-power-stroke
conformation, the switch-2 region is not part of the nucleotide-binding pocket.
A similar movement of the switch-2 region that depends on trinucleotide beingbound is also found in the G-proteins (43). A similar but even larger movementof the switch-2 region is found in the empty site of F1-ATPase, compared with
the ATP or ADP sites (44). Only in CLOSED (e.g. in S1.ADP.vanadate) can thehydrogen bond (invariant for G-proteins, kinesins, and myosin but not present
in F1 ATPase) between the amide of SkG466 and the g-phosphate be formed.
Moreover, in CLOSED the g-phosphate is also coordinated by the Mg2+ ion,
the invariant P-loop lysine, and S181 (45). Furthermore, through the formationof a salt bridge with SkR236, SkE468 comes close enough to the putative
attacking water to stabilize and polarize this molecule. In OPEN, SkE468 takes
on a different orientation. It is difficult to see how hydrolysis can proceed in
OPEN, which would therefore appear not to be an Mg.ATPase: The closing
would appear to be essential for enabling hydrolysis. Smith & Rayment (41)proposed that the CLOSED form is brought about by the formation of the transition state complex with the g-phosphate stabilized in the pentacovalent intermediate conformation. However, ADP.BeF3 complexes can also produce thisstate (see below).

ADP.BeF3, an ATP Analog, Can Produce Both OPEN
and CLOSED States
ADP.BeF3 is thought to be an analog for ATP. Fisher et al (40) solved the structure of Dictyostelium S1 with ADP.BeF3 bound in the active site and found it tobe remarkably similar to skeletal chicken S1 without nucleotides. This result
appears to show that the structure of the ATP state is OPEN, which is puzzling
because it would not be able to hydrolyze the ATP. Moreover, the attitude of theconverter domain (and hence inferred position of the neck or lever arm) is nearto what we anticipate should be the end of the power stroke, which is also unexpected for the ATP state. More recently investigators (I Schlichting, A Becker, D
Manstein & KC Holmes, manuscript in preparation) have solved the structure ofan ADP.BeF3 complex of truncated Dictyostelium S1 and found it to be essentially identical to the ADP.vanadate complex. The g-phosphate–binding site is
CLOSED, and the converter domain is in the rotated configuration. The constructused in this case was 5 residues shorter than that used by Fisher et al (40). This
results in a tighter binding of ADP (46). It may be that the difference in binding
energy of ADP.BeF3 is adequate to favor the CLOSED structure: The transition
between the two forms of myosin therefore appears to be sensitively poised.

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Crystallographic studies on truncated forms of smooth-muscle myosin (47;see below) show the closed conformation with both ADP.BeF3 and ADP.vanadate, so that it seems rather clear that the CLOSED form can accommodate ATPboth in its normal and pentacovalent forms. It therefore seems reasonable toassume that the CLOSED form can also accommodate ADP.Pi without further
substantial alterations of the protein conformation.

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The Switch-2 Region Should Open for Phosphate Release
The CLOSED structure appears to generate a tight coordination pattern for theoxygens of the g-phosphate both before and after cleavage, which would explain
the high phosphate affinity. This interaction in turn is important for stabilizing
the closed form when g-phosphate is present. Opening the switch-2 region
destroys the g-phosphate–binding pocket and the SkE468–R245 salt bridge.
These changes would appear to facilitate g-phosphate release [a “back door
enzyme” (48, 49)]. It seems very likely that actin binding favors the OPEN form
and thereby facilitates phosphate release. Unfortunately, the structural basis of
this important mechanism has yet to be revealed.

The Lever Arm
The movement of switch 2 in the CLOSED form has other more far-ranging consequences, namely the rotation of the converter domain through about 60°.
Holmes (8, 50) used the coordinates of the chicken structure to provide the missing lever arm and proposed that the new orientation of the lever arm was indeed
the beginning of the power stroke. A model of this new state is shown in Figure 7(cf Figure 8). We show the corresponding diagram for the open state [truncatedS1 with ADP.BeF3 bound in the active site (40)], which may mimic an ADP state[i.e. the state after release of the g-phosphate; cf the structure of chicken skeletal
myosin with no nucleotide bound (Figure 6)]. The end of the lever arm has moved
through 11 nm along the actin helix axis between OPEN and CLOSED, which isabout the expected magnitude of the power stroke. This large change is driven
through molecular cogs and gears by a small (0.5-nm) change in the active site.
Therefore, it now seems rather likely that the myosin power stroke works byswitching between these two conformations.
Smith & Rayment (41) were cautious of this interpretation because they
thought that the truncation of the light chains might cause artifacts and also
because the converter domain was apparently rather mobile. Moreover, thechanges did not explain the chemical cross-linking effects (51). The subsequent
structure of truncated Dictyostelium myosin with ADP.BeF3 showed the converter domain in the second orientation very clearly (I Schlichting, A Becker, D
Manstein & KC Holmes, manuscript in preparation), but still doubts about the
integrity of such truncated constructs remained. Such doubts have been allayedby the publication of the structure of smooth-muscle myosin from Carolyn
Cohen’s laboratory.

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Figure 6a A view of the ATP-binding site looking out from the actin helix. Shownare the P-loop (green), an Mg.ATP molecule with the base at the back and the threephosphate groups in the front (carbon, yellow; nitrogen, blue; phosphate, light blue;
oxygen, red; magnesium, green); parts of the 50K upper domain (red) including theso-called “switch-1” region (right); the switch-2 element in the open “ADP” (white)and closed “ATP” (gray) conformations—the conserved glycine (Sk466) is shown ingray or white (note that this residue moves about 0.5 nm between the two conformations); and the helix (Sk648–666), which acts as fulcrum for the relative rotation of
the 50K upper and lower domains (blue).

Smooth-Muscle Myosin
Chicken smooth-muscle myosin truncated at Sm791 and Sm820 has been
expressed in insect cells by using the baculovirus vector. The shorter construct
stops at the end of the motor domain “core” (3) and the longer construct encompasses the essential light-chain binding site, that is, the first half of the lever arm,
and is complete with the essential light chain. The structures of both constructs
have been solved as complexes with ADP.vanadate and ADP.BeF3 (47). All show
the myosin cross-bridge in the CLOSED form (the putative pre-power-stroke orientation) with the converter domain in the rotated position essentially identical tothat obtained in the two Dictyostelium constructs discussed above. There are no
substantial differences arising from ADP.AlF4 as compared with ADP.BeF3,
apparently showing that the nature of the ligand (ATP analog or transition stateanalog) does not control the protein conformation. Since all the smooth-muscle
crystals display extensive noncrystallographic symmetry, CLOSED has now beenobtained 16 times in a large variety of different environments. It seems, there-8505_AR_23

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705

Figure 6b A view at right angles to the above with CLOSED at the top and OPEN
below, showing the large changes in the orientation of the converter domain (left) resulting from the inward movement of the switch-2 element (right), which results in abending and twisting of the switch-2 helix and the associated switch-2 loop, which
carries the highly conserved W510. [Figures produced by using GRASP (125) fromdata sets described in the text.]

fore, that one can rule out the possibility that the CLOSED form is an artifact oftruncation or crystalization.
An unexpected new insight into a possible control mechanism is provided by
the essential light-chain construct (Figure 9). Here the converter domain is in the
CLOSED (i.e. rotated) orientation, which leads to the lever arm helix starting out
in the predicted manner. However at residue Sm792 there is a break in the helix
so that the ensuing helix heads off at an angle about 8° inclined to the initial helix

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Figure 7 A reconstruction of the “pre-power-stroke” state from the crystallographic
data on the Dictyostelium construct truncated at 759 and complexed with ADP.vanadate (41). Note the 70° rotation of the converter domain. The missing “neck” or leverarm has been modeled from chicken S1 data by superimposing the converter domains.
To establish the orientation in relation to the actin helix (right), the 50K upper andlower domains have been superimposed on the corresponding domains in the chicken
structure shown in Figure 5. Note the rotation of the converter domain compared withthat in Figure 5. [Figure prepared by using Bobscript and Raster3D (123, 124).]

direction. This break is caused by the essential light chain forming a bond withthe top of the 25K domain rather close to the 25K–50K junction (loop 1). The
position of the lever arm inferred from this structure is very strongly angled tothe muscle filament axis. It is possibly a parking position brought about by a regulatory function of the light chains.

Molecular Cogs and Gears
The smooth-muscle structures provide a wealth of new data. Moreover, now thecomplete coordinates of the moving parts are available for almost identical

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MUSCLE CONTRACTION

707

Figure 8 The “post-power-stroke” state: modeled from the crystallographic data on
the Dictyostelium myosin motor domain truncated at residue 759 and complexed with
ADP.BeF3 (40). Details are as in Figure 7. The end of the lever arm moves about 11
nm between the two states. [Figure prepared with Bobscript and Raster3D (123, 124).]

sequences in both the OPEN (34; Brookhaven Protein Data Bank accession
number 2MYS—chicken skeletal muscle with sulfate in the active site) and the
CLOSED form (47; Brookhaven Protein Data Bank accession numbers 1BR1,
1BR2, and 1BR4—chicken smooth muscle with ADP.BeF3 and ADP.AlF4 in
the active site). This allows a rather detailed analysis of the conformationalchanges between the OPEN and CLOSED forms, which are limited to theregions Sk460–520 (switch 2 and switch-2 helix) and the SH2-SH1 hinge(Sk694–716). Nearly all the changes take place by rigid-body rotations of secondary and tertiary structure elements. The basic movement that closes the gphosphate–binding site is a rotation of the actin-binding domain in relation to
the 50K upper domain about the helix Sk648–666. This forces the SH1 helix
against the switch-2 helix, which bends out at residue Sk497. As this happens
the end of the switch-2 helix twists through about 60° (Figure 6b). This in turncauses a rotation of the converter domain (Sk711–781) with which the end of
the switch-2 helix interacts by a strong hydrophobic bond. The fulcrum for theconverter domain is provided by residues at the distal end of the SH1 helix(Sk707–711), each of which undergoes rather moderate changes in its f, c

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Figure 9 Structure of smooth-muscle myosin truncated at 819, carrying the essential light chain and with ADP.AlF4 or ADP.BeF3 bound in the active site (47). The
molecule has been orientated in relation to the actin helix by superimposing the 50K
domain with the 50K domain of chicken S1. Note the similarity to Figure 7. The
structure is CLOSED (i.e. pre-power-stroke), and the converter domain is in the rotated position. There is a break in the lever arm helix at residue Sm792; the ensuing
helix heads off at an angle of about 8° to the initial helix direction. The essential lightchain contacts the top of the 25K domain rather close to Loop 1. [Figure prepared byusing Bobscript and Raster3D (123, 124).]angles. The angle between the SH1 helix and the SH2 helix alters by a rotationthat mostly occurs at SkG699. There is no evidence of a melting or disorderingof the SH1–SH2 helices, as has been suggested by chemical cross-linking experiments (51). In fact the SH2 helix hardly moves. The physiological significanceof the cross-linking experiments remains to be explained.
This comparison shows up a remarkable fact: The transition from OPEN to
CLOSED is achieved by substantial movements in only a very few residues(Table 2). The largest f, c changes are limited to the switch-2 loop (i.e. residues
Sk509–519), which includes the invariant W510 (the side chain moves from asolvent exposed environment to a shielded environment) and the following highly8505_AR_23

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709

conserved sequence EFIDFGM where, in particular, G516 suffers a peptide flip.
In the Dictyostelium OPEN form, which we equate with the ADP form (seebelow), the switch-2 loop is in fact disordered.
At this preliminary level of analysis the changes are not reminiscent of a statechange (e.g. the R-T transformation of hemoglobin, with helices clicking from one
position to another with a high activation energy) but rather appear like the meshing of cogs and gears. Apart from changes in the g-phosphate–binding pocket itself,
which includes the formation of a salt bridge, it appears that the transformation
might occur without an appreciable activation energy. The myosin structure might
be able to flop easily between the two states to take up any intermediate state (52)in response to external forces (such as the effects of actin binding and tension on
the molecule). This would be a very desirable property for a molecular machine.

OTHER EVIDENCE FOR A SWINGING LEVER ARM
Effects of ADP on the Structure of Decorated Actin
The ideal experiment for observing the power stroke (i.e. the transition from
OPEN to CLOSED while bound to actin) would be to add ATP to decorated actin
and observe the change in the myosin cross-bridge orientation by electron
microscopy. This experiment is difficult to perform since the ATP causes rapid
dissociation of the actin-myosin complex. Two experiments using time-resolved
methods (53, 54) have produced apparently inconsistent answers. A more robust
approach is to look for the effects of adding ADP, which does not lower the actinmyosin affinity enough to cause dissociation in an electron microscope experiment. Although with skeletal muscle S1 there is no observable effect on binding
ADP, for smooth muscle and some other myosins the addition of ADP to decorated actin caused quite large changes in the angle of the lever arm (55, 56),
which can be clearly observed by cryoelectron microscopy. This effect seems to
be limited to actomyosins with a relatively high affinity for ADP. Although
clearly this is not an effect that can be related to phosphate release, these experiments are noteworthy for providing the first incontrovertible evidence of leverarm movement in response to nucleotide binding.

Use of Fluorescent Probes to Measure Lever Movement
Fluorescence polarization measurements provide a convenient method of measuring lever arm orientation if the probes can be attached in a way that hinders
probe rotation and does not inhibit myosin function. Because the atomic structures of the light chains are known (34, 35), genetically engineered light chains
can be manufactured with SH groups in suitable positions to allow difunctional
chromophores to be securely attached. Furthermore, the chemically modified
light chains can be diffused into muscle fiber and exchanged for endogenous
light chains. This approach has been used by Goldman and collaborators to show

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OPEN

CLOSED

Sk

f

c

f

c

Sm

M495

277.0

232.0

288.4

243.1

M497

Z496

273.3

240.7

2114.8

252.2

F498

V497

261.2

250.6

255.9

270.6

I499

L498

256.8

254.6

250.5

255.0

L500

E499

248.6

260.6

249.7

245.4

E501

Z500

250.4

218.6

272.3

227.2

Q502

E501

277.0

241.5

259.7

250.3

E503

E502

259.0

260.6

260.8

229.2

E504

Y503

252.5

212.5

264.9

241.3

Y505

Z504

292.7

238.6

264.6

259.6

Q506

Z505

266.3

252.0

240.6

263.5

R507

E506

271.5

245.4

261.8

217.5

E508

G507

111.0

16.5

68.0

50.6

G509

I508

277.1

160.2

290.9

160.7

I510

E509

2113.2

85.9

265.7

91.9

E511

W510

2126.1122.0

2150.6

141.4

W512

E511

2109.8138.62114.1

144.3

N513

F512

2160.4

149.2

263.6

161.3

F514

I513

279.1

147.3

2143.6

121.7

I515

D514

247.4

138.0

292.8

102.7

D516

F515

283.6

213.4

278.5

22.0

F517

G516

263.2

249.0

48.5

65.0

G518

M517

232.3

233.5

2140.9

89.3

L519

D518

262.2

227.8

2144.0138.0

D520
continuedthat in contracting muscle fibers the light chains swing through angles that arequite consistent with the molecular models for the swinging lever arm proposed
above, if one assumes that only a small proportion of the cross-bridges are activeat any one time (24, 57).

Making the Lever Arm Longer or Shorter
The length of the lever arm can be altered by inserting or deleting lightchain–binding sites (58, 59). Alternatively an artificial lever arm fabricated froma-actinin repeating units can be added to the myosin core (60). In both experiments modified-Dictyostelium–expressed myosin was used for in vitro motility

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continued
OPEN

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711

CLOSED

Sk

f

c

f

c

Sm

L519

293.5

215.8

2128.4

11.0

L521

A520

249.6

232.5

244.2

253.2

Q522

R696

277.8

242.9

290.5

257.0

R706

C697

280.9

210.4

271.9

29.2

C707

N698

276.6

246.3

296.3

215.6

N708

G699

118.3

213.8

78.1

27.6

G709

V700

257.4

246.3

256.7

265.2

V710

L701

276.3

217.8

244.0

260.0

L711

E702

288.5

222.7

254.2

245.5

E712

G703

278.3

226.8

252.8

216.2

G713

I704

274.6

241.1

2104.3

232.7

I714

R705

253.8

260.4

251.8

251.6

R715

I706

256.1

225.5

261.8

234.4

I716

C707

275.2

249.9

262.9

232.7

C717

R708

268.0

8.3

91.3

244.5

R718

K709

2104.2

230.8

264.5

262.7

K719

G710

289.1

2129.9284.5

2162.5G720

F711

2119.788.5

2122.1

81.9

F721

P712

249.3

291.2

264.2

2.1

P722

S713

257.7

128.8

2125.9

134.1

S723

R714

2133.1144.9

2143.4

151.2

R724

V715

294.1

133.2

2151.9

135.6

V725

L716

278.8

135.7

272.4

140.8

L726

assays in which the speed of transport of actin filaments across a lawn of myosin
heads was measured. It was shown that the ATPase rates of the constructs were
unaffected by the alterations to the lever arm, in which case the speed of transport would be expected to be proportional to the length of the lever arm. In bothcases it was found that the speed of actin transport was indeed proportional to
the length of the lever arm. Moreover, in the experiments of Uyeda et al (59) an
extrapolation could be made to the point of zero velocity. This came out with aradial position behind the converter domain very close to the SH1 hinge discussed above. In all these studies only the motility was measured; it will be ofinterest to determine whether the force produced also scales with the length ofthe lever arm.

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Allocation of States to Kinetic Schemes

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The biochemical mechanism of ATP hydrolysis by myosin was defined in the
1970s and became known as the Bagshaw-Trentham scheme (61), and although
there have been minor modifications, the scheme outlined below has remainedlargely unchanged. All myosins studied to date follow this same basic mechanism with modifications in both the overall ATPase rate and the relative rates ofindividual steps (62–65).

The best characterized myosin is the subfragment 1 from rabbit fast muscle.
In this myosin ATP binds in a two-step process, a diffusion-limited reaction (step1) followed by a fast almost irreversible conformational change, which results in
an increase in the intrinsic protein fluorescence (step 2). Reversible ATP hydrolysis then occurs at a rate of >100 s-1 accompanied by a further increase in fluorescence with an equilibrium constant of ~10 (step 3). The products remain tightly
bound to the protein and a slow (0.05 s-1) protein isomerization is required (step4) before Pi is rapidly released (step 5). A fluorescence decrease accompanies
step 4. After Pi release, ADP dissociates in a two-step reaction that is essentiallythe reverse of steps 1 and 2 and is associated with a further fall in fluorescence.
In terms of the above scheme, the state OPEN would most simply
be assigned to the M*N state because it is observed with crystals containing ADP,
AMP.PNP, ATPgS (an unhydrolyzable ATP) bound to the nucleotide site. The
fluorescence change accompanying nucleotide binding is most likely the resultof a perturbation of SkW113 and or SkW131 in the nucleotide-binding pocket,
because myosins lacking these tryptophan residues show little fluorescencechange on nucleotide binding [e.g. Dictyostelium and Scallop (62, 66)].
The state CLOSED therefore would correspond to M**ADP.Pi or the transition state of the hydrolysis reaction. Dictyostelium and both chicken myosins show
a fluorescence change accompanying the hydrolysis step. Of the three tryptophan
residues in the motor domain common to Dictyostelium and chickens, the local
environments of SkW440 (Dy432) and SkW595 (Dy584) do not alter appreciably
between OPEN and CLOSED forms. Therefore they are not strong candidates forthe intrinsic fluorescence signal. However SkW510 (Dy501) sits close to the junction between the switch-2 helix and the converter domain and twists from asolvent-exposed to solvent-shielded environment between OPEN and CLOSED.
Therefore it is a priori a good candidate for the signal (see also 13 and 67). Thus
the protein fluorescence change accompanying hydrolysis would arise from theperturbation of SkW510, which signals the movement of switch 2 and the lever8505_AR_23

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arm. However, as noted above, state OPEN appears to be noncompetent forhydrolysis. Thus the conformational change must precede hydrolysis. We therefore need to consider step 3 as comprising two events: Step 3a is the conformational change, and step 3b is the hydrolysis step. Unusual in this scheme is thenotion that hydrolysis should be fast and not associated with a large protein isomerization. On the other hand, 3a is a very significant isomerization because theconverter domain and lever arm move a large distance as the pocket shuts.

In all measurements made to date the second component of the fluorescencechange and the hydrolysis step occur at the same observed rate. Therefore, hydrolysis must very quickly follow the conformational change; otherwise a significant
lag of the hydrolysis behind the fluorescence change would have been observed.
The hydrolysis step is also readily reversible (K3 = 10), that is, it occurs with a verysmall free-energy change and has fast forward and reverse rate constants (k +3 + k-3 100s-1) (68). The simplest assignment of the rate constants to the two components ofstep 3 is therefore that k+3a100 s-1and k-3b10 s-1with k+3b, k-3a >> 100 s-1. Thus
M**ATP is an unstable state, breaks down rapidly in either the forward or reverse
direction, and as such has many characteristics of a transition state. This should not
to be confused with the chemical transition state for the hydrolysis step, which isstabilized by the protein and may correspond to M**ADP.Pi.
The small free-energy change for step 3 is known to be composed of large
compensatory changes in enthalpy and entropy so that although there is only asmall net change in ∆G, energy is being converted/stored in this step (69, 70).
The evidence for the fast rates of reversal of the hydrolysis reaction comelargely from 18O isotope exchange studies (71, 72), whereas the assignment of
an equilibrium constant of 10 comes from analysis of the breakdown products onacid quenching the complex (68). The possibility that the M**ADP.Pi complex isa transition complex is not ruled out by these studies. Particularly the elegant
work of Dale & Hackney (73) has demonstrated that, although the oxygens on
the g-Pi are rapidly exchanged with solvent, the position of oxygen isotopes on
the b-Pi do not change. Thus it remains possible that the b-g bond is not broken
on forming M**ADP.Pi and that the g-Pi is pentacovalent; that is, the attack bywater has occurred but the ATP b-gPi bond is not broken. Step 3 could therefore
represent the formation of the pentacovalent gPi complex (step 3b), which occurs
concomitant with the OPEN to CLOSED conformational change (step 3a).

Sensitivity to g-Phosphate
The OPEN form appears to be favored by initial ATP binding and by ADP; the
CLOSED form is favored by ADP.vanadate and some other transition state analogs.
One important difference between these forms appears to be the positionof SkGly466 in switch 2, which corresponds to Gly60 in ras p21. In ras the8505_AR_23

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sensitivity of switch 2 to the presence of either GTP or GDP in the active site is aconsequence of the coordination of the g-phosphate of GTP by the amide nitrogenof G60. This coordination is also an important part of the hydrolytic machinery. In
the OPEN form of myosin S1, this hydrogen bond cannot be made; in the presenceof the g-phosphate, the switch-2 region moves towards the binding pocket by about
0.5 nm to allow this H-bond to form. (see Point Mutations of G466 below).
Note that in the switch-2 CLOSED form the phosphate is apparently coordinated by three strong bonds in addition to the covalent bond. There is no reason
to believe it would rotate quickly after hydrolysis. The crystal structure therefore
gives no indication of how the three g-Pi oxygens can be rapidly exchanged withwater oxygens.
The two crystal structures therefore are consistent with a major change inmyosin conformation being associated with (or perhaps just before) the hydrolysis step and with this conformational change being largely a reversal (or a priming) of the conformational change of the power stroke driven by actin binding.
This is essentially the scheme proposed by Lymn & Taylor (19) in 1971 and is
the basis of Figure 1. At the time there was no direct evidence for this conformational change on the hydrolysis step, but it is a logical consequence of the mechanism they proposed.
To understand the complete ATPase cycle requires more detailed structural
knowledge of the way in which actin reverses the CLOSED-OPEN conformational change.

Present Structures Do Not Illuminate the Mechanismof Actin-Induced ADP Release
Both OPEN and CLOSED have relatively high affinity for nucleotides, so that
actin-mediated release of nucleotides must involve more structural change than
we presently see. The effect of actin would not be expected to be simply a reversal of the OPEN-CLOSED change because one expects a vectorial mechanism todrive the cycle, which is provided by the requirement for P i release to precede
ADP release. The missing part of the cycle is the mechanism whereby actin binds
to the CLOSED form and induces a series of conformational changes that lead to
Pi release followed by ADP release. Whereas the transition CLOSED to OPEN
would lead to Pi release, the structures give no indication of how actin induces
ADP release and then ATP binding induces actin dissociation. Solution biochemical studies have provided several insights into the docking of actin with myosin
and its effect on nucleotide binding to myosin.

THE ACTIN-MYOSIN INTERACTION
Kinetic Data
The current low-resolution structural view of actin binding to myosin suggeststhat no major rearrangement of the myosin takes place at the actin-myosin

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MUSCLE CONTRACTION

715

interface but that large-scale changes occur on myosin distal to the actin-binding site. However this does not mean that no structural changes take place at
the interface. In the interaction of any two proteins the docking is likely to
involve initial diffusion-limited complex formation followed by a series ofstructural adjustments (induced fit) as the stereospecific interaction site isformed, along with induced conformational changes in the proteins. Kinetically the docking of myosin onto an actin filament can be resolved into at least
three events (4). Initial complex formation, largely involving charge-chargeinteractions, is followed by two changes in conformation of the complex.
The first may involve the formation of stereospecific hydrophobic interactions(4), and the second involves a major rearrangement of the actin-S1 complex
because fluorescence probes on both actin and the nucleotide in the myosinpocket report the change simultaneously (74, 75). The second isomerization
also appears to involve a large-volume increase of the complex, which isnormally assigned to displacement of a large amount of water from thecomplex (76, 77).

The three events are best documented for rabbit myosin in the absence of
nucleotide. As we apparently do not have a high-resolution structure of thenucleotide-free myosin S1, it is not simple to predict what actin binding will doto the S1 structure. However, the three events appear similar (but with marked
changes in rate and equilibrium constants) for M.ADP (78, 79), and it is suggested that M.ADP.Pi binds in the same three steps, but the equilibrium constantfor the last step is small (~1, weak binding to actin) until Pi is displaced from themyosin. The last of the conformational changes has been suggested to be closely
coupled to the power stroke of the ATPase cycle (80–82).
The key feature of this mechanism is that the isomerization of step 2 results ina major strengthening of the actin binding to myosin and simultaneously a weakening of the nucleotide binding to actin. The equilibrium constant of this step
varies for different nucleotides. Thus, in the absence of nucleotide, K2 is large(>100), in the presence of ATP it is small (K2 << 1), and ATP displaces actin. For
other nucleotides and nucleotide analogs, K2 is intermediate, and which form
predominates, the ternary complex or the binary complex with either nucleotide
or actin, depends on the concentrations used.
For fast rabbit myosin, the formation of the R-state reduces the affinity of the
myosin head for ADP by more than 100-fold (79, 83), and the rate of ADP dissociation is accelerated more than 500-fold. These factors vary widely for different myosin types. For chicken smooth muscle myosin, for example, the ADP
affinity is reduced only fivefold, whereas the dissociation rate is accelerated tenfold by actin (84). Actin also induces acceleration of Pi dissociation, typicallymore than 200-fold.

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Correspondence with Structural Data
We can now address the question of the correspondence between the conformational changes detected by fluorescence probes in the kinetic measurements
and the structures seen in the crystals. The current evidence suggests that theformation of the A-state (probably involving stereospecific interactions in theactomyosin interface) has little influence on the nucleotide-binding pocket,
because the labels on the nucleotide do not detect formation of the A-state. Nordoes the formation of the A-state significantly weaken the affinity of nucleotide
for myosin. The formation of the A-state therefore appears to be limited to local
changes at the actomyosin interface. Therefore the binding of either the CLOSEDor OPEN form to actin to make an A-state may be expected to be similar in eachcase. The isomerization to the R-state seems to require an empty Pi site because
no R-state has been detected for myosin complexed with ATP, ATPgS, or ADP
and the Pi analogs (85). The R-state can be formed in the presence of either ADP(K2 = 10) or pyrophosphate (K2 = 2). Thus the A-state can be formed with any
myosin complex, but the isomerization to the R-state is inhibited by a tightly
bound Pi. Formation of the R-state is likely to involve opening of the back door
(allowing Pi to dissociate if not covalently bound) and then weakening of thenucleotide affinity. The order is important. Thus the A to R isomerization of the
CLOSED form/A-state occurs with opening of the backdoor and loss of Pi. Isomerization of the OPEN form depends on the state of the Pi site. With ATP, ATPgS,
or ADP.BeF3 bound, the formation of the R-state will not be allowed. With nonucleotide or ADP bound, the isomerization to the R-state is highly favored.
In summary the crystallographic data provide evidence of two conformationsof the myosin in the absence of actin, which we call CLOSED and OPEN. Theirproperties are summarized in Table 3. The CLOSED form found when myosin iscomplexed with ADP.Vi and ADP.AlF4 is the form with the tail in the up (prepower-stroke) position with switch 2 IN, poised for catalysis and therefore withthe back door for Pi release closed. The OPEN form favored by ATPgS,AMPPNP ADP, and ATP (I Rayment, unpublished data) and sometimes by
ADP.BeF3 (e.g. Dy-truncated at M759) has switch 2 OUT and the backdoor open,
and therefore catalysis cannot take place. The tail is in the down position or postpower-stroke conformation.

A Third State Is Required
We suggest that both CLOSED and OPEN forms with the gPi occupied can interact only weakly with actin to form the A-state and that we need at least one more
conformation to complete the cycle. Initially, actin must bind to the CLOSED
form to give a weakly bound actin (the A-state). Then the complex should proceed to a third form, which should have the following properties (see Table 3):
The third state should bind strongly to actin (R-state) with high pyrene-actin fluorescence; the backdoor should be open and with very weak Pi affinity and weak
nucleotide affinity (low mant-nucleotide fluorescence); the switch 2 should be out
and the converter domain and lever arm in the post-power-stroke down position.

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MUSCLE CONTRACTIONTABLE 3717

Properties of states

Property

CLOSED

OPEN?3rd state?

Nucleotide

Vi/A1F/Bef

ATP/BeF/gS/

Actin only

Switch II

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PNP/ADPOUT

Further out?

Converter/tail

UP

DOWN

DOWN

Backdoor

CLOSED

OPEN

OPEN

Catalysis

Competent/TS

Noncompetent

Noncompetent

Pi affinity

Very tight

Only covalent as
g Pi

Very weak

Fluorescence(W510)High (M**ADP.Pi)

Low (M*ATP)Low

Actin binding

Weak A-state

Weak A-state

Strong R-state

Pyr-actin Fl

High

High

Low

Nucleotide binding

StrongStrongWeakmant.ATP/ADP F1

High

High

Low

The difficult question is how the transition between the three states, CLOSED(M.ADP.Pi) to R-state (A.M or A.M.D) to OPEN (M*ATP) to CLOSED, occurs.
The crystal structures allow us to propose that, for OPEN-to-CLOSED transition, the trigger is the moving in of switch 2. At present we have no indications
as to how the other two transitions might function. However, solution biochemistry helps set the ground rules. Actin can bind to the CLOSED M**ADP.Pi toform the A-state with no major changes in conformation. The actin then produces
a conformational change in the complex which gives a stronger actin binding (Rstate sensed by pyrene on actin) and causes the switch 2 to move out (backdoor
to open), Pi dissociation, and the converter/tail to move down. Subsequent to thischange actin causes a change in the nucleotide site to promote ADP release. Thissecond change can be seen in some slower myosins as an additional tail movement on ADP release (56, 65, 86, 87). ATP binding to AM to return to the
M*ATP- OPEN state reverses the A-to-R transition but does relatively little tothe converter/tail or to the back door (or switch 2).

A Possible Mechanism for ADP Release
Such speculations allow one to construct a working scheme, but without highresolution structures of actomyosin complexes they remain conjecture. However,
because actin is a nucleotide exchange factor, we could look to other systems
involving nucleotide exchange for ideas. A simple structural scenario for the reciprocal relationship between actin displacing ADP and ATP displacing actin is provided by analogy with the G-proteins. For example, the binding of Sos to Ras p21

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is accompanied by disruption of the switch-2 region. The Mg2+ is forced to leaveby the insertion of a methyl group in its place (88). For Tu-Ts the mechanism isdifferent, but one aspect is the same—the binding site for the Mg2+ is destroyed(89). In the empty site of F1 ATPase, the Mg2+ coordination is also destroyed (44).
Thus nucleotide release in actomyosin could work through weakening the coordination of Mg2+. In myosin, ATP binds with a tightly bound Mg2+, which is thencoordinated via bonds with SkD463 and the b- and g-phosphate groups. Afterhydrolysis and Pi release, the Mg2+ coordination is weakened. It is possible that,after Pi dissociation, actin moves switch 2 and D463 further out, thereby further
weakening the Mg2+ coordination and resulting in a loss of Mg2+ and then ADP.
As ATP rebinds with tightly coordinated Mg2+, this would restore switch 2 backthrough its OPEN to CLOSED position, thereby weakening actin affinity.

EFFECTS OF STRAIN ON PHOSPHATE
AND ADP AFFINITY
The above scenario is based on the events in a solution or in an unloaded system.
It is of interest to consider the sequence for a loaded motor. Starting from state
CLOSED (M**ADP.Pi weak actin binding), actin induces backdoor opening andallows Pi release. Most consider that the major conformation change accompanying force generation occurs before Pi release (81, 90, 91), so we assume that
actin binds strongly to the myosin, opening the backdoor and swinging the converter domain, which then allows Pi to dissociate. In the absence of load thisgoes to the completion of the power stroke and then ADP is lost. If the system isheld isometric then either the swing of the tail is arrested or the strain involved in
the swing is stored in the head. This has the effect of allowing the Pi to bindreversibly to the myosin head. The open back door and presence of the Pi, however, prevent ADP release. Apparently the swing of the tail (or the dissipation of
the strain in the head) is required before actin can cause disruption of the Mg
coordination and allow ADP dissociation.
There has long been recognition of the necessity for a strain-dependent ADP
release mechanism in actomyosin contraction. This goes back to the Fenn effectand the observations that, without such a mechanism, the observed fast ADP
release followed by ATP binding would very rapidly eliminate any strained or
unstrained heads in fast skeletal muscles. Smith & Geeves (92) speculated that the
ADP pocket might be positioned on the head in such a way that any load on thehead could act to close the nucleotide pocket. Based on the analysis of fast skeletal muscles, they further argued that the fast release of ADP would suggest thatany pocket opening required for nucleotide release (or ATP binding) would probably be a transition state. Thus strain would affect the rate of nucleotide binding
with little effect on the equilibrium constant. More recently the electron
microscopy work of Whittaker et al (56) and Jontes et al (86) has shown that, forthe smooth muscle myosin and brush border myosin I, a structural change doestake place on ADP binding to actomyosin such that ADP release gives an extra

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MUSCLE CONTRACTION

719

kick to the tail. From a detailed kinetic analysis Cremo & Geeves (84) argued that
thermodynamically the ADP release would be unlikely to contribute to force generation, particularly because at cellular concentrations work would have to be put
into the system to release ADP (in much the same way that work has to be done torelease ATP from myosin and the F1 ATPase). The arrangement would, however,provide a very good strain-limited ADP release mechanism, particularly for amyosin that is designed for high forces and slow contractions such as the smoothmyosin. A similar situation appears to hold for the 130-kDa myosin I from rat
muscle. This myosin, like brush border myosin I, has been shown to have a double kick in the laser trap experiment (87) and also has a very high affinity for ADP
when complexed to actin. It therefore appears to be a myosin designed for load
bearing rather than movement (L Coluccio & M Geeves, in press)

MUTATIONAL ANALYSIS AND
SEQUENCE COMPARISONS
GeneralFurther insights into the structure-function relationship for myosin come fromstudies of changes in the sequence of myosin both from engineered mutations
and the study of naturally occurring variants. As with the classical work on hemoglobin, the study of the naturally occurring variants of the myosin molecule provide a wealth of information about which parts of the structure are essential for
function and which parts are tolerant. A combination of such structural and functional studies provides information on how the protein is tuned to particular physiological functions.
There are currently 15 known members of the myosin super family (3, 93, 94),
all related by a conserved motor domain. Few of these myosins have been examined by anything but sequence analysis or genetic knockouts. Because the cellularfunction of many myosins is unknown, we cannot anticipate how sequence differences result in changes in biological activity. However analysis of sequence comparisons both between and within families has produced insights into the essential
and variable regions of the myosin structure (3, 93, 94). The most detailed analysis has been done on the members of the myosin II family, which show a large
general homology but variation in local areas particularly between classes (e.g.
fast-, slow-, and smooth-muscle myosins and nonmuscle myosins). Analysis of
the variations between species and between classes should be quite revealing.

Alternate Splicing
Variations in myosin II structures expressed within a single species often involve
multiple genes and alternate splicing of the myosin heavy chain. Bernstein &Milligan (17) analyzed the sequences of the 12 known Drosophila myosins,
which are products of a single myosin heavy chain (MHC) gene. These 12

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GEEVES & HOLMES

sequences are produced by alternate splices at exons 3, 7, 9, and 11, which have2, 4, 3, and 5 alternate exons, respectively. Exon 7 (Sk301–335) is near the
nucleotide-binding pocket, includes Loop 1, and links with the switch-1 region.
Exons 3 (Sk69–116) and 9 [Sk472–528 (include W510 conserved throughout)]
flank the essential thiol helices in the 3-dimensional structure. Exon 3 includes
the N-terminal b-barrel, which is absent in myosin I and has been suggested to
be involved in head-head contacts in heavy meromyosin. Residues 82–101 lie
close to the SH1 and the hinge regions. Exon 9 is the loop that links switch 2 tothe converter domain and includes the W510, which reports the major OPEN-toCLOSED conformational change. Exon 11 (Sk724–764) is the major part of theconverter domain. To date there has been no structural or functional analysis ofthese alternate splices, but such studies can be expected to help reveal how thefunction of the myosin is tuned to specific cellular needs.
Spudich (14) pointed out the potential of sequence comparisons. He notedthe variation in the number of light-chain–binding motives (IQ motives) andconcluded that different myosins might bind different numbers of light chains.
This led him and others to test the lever arm hypothesis by deleting and inserting light-chain–binding sites (discussed above). He also identified the surface
loops connecting the 25-to 50- (Loop 1) and 50- to 20- (Loop 2) kDa domains
as being different between groups of myosin within the myosin II family. He
proposed that, although the head structure is generally highly conserved, variations in Loop 2, located in an actin-binding site, can modulate the interaction
with actin and hence the actin-activated ATPase. In contrast variations in Loop1, near the entrance to the nucleotide-binding pocket, can modulate nucleotide
affinity and the rate of ADP release and thereby the shortening velocity.
Although it was noted that this assignment of the roles of the two loops was
simplistic, as there is probably cross talk between the two loops, it remains a
useful hypothesis in our search for the mechanisms of ADP release and actin
binding. These ideas have been widely tested by constructing chimeras based oneither the Dictyostelium myosin II S1 (95–97) or the chicken smooth myosin
head (98, 99). Kinetic data show the hypothesis to be broadly correct although asimple separation of the roles’ effects is not always maintained, particularly forthe smooth-muscle myosin.

Loops
The chimeric constructs of Loops 1 and 2 demonstrated that large loop changes
can modulate the properties of myosin. However, these studies have not revealed
which properties of the loop are important for determining the properties of the
myosin. Two studies have attempted a more systematic approach.
Sweeney et al (99) used the smooth-muscle head expressed in baculovirus to
study nine variants of Loop 1 taken from a variety of muscle and nonmuscle
myosins. These showed that loop substitutions could produce up to a ninefold
inhibition of the rate of ADP release and a 2.7-fold and twofold, respectively,

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MUSCLE CONTRACTION

721

inhibition of in vitro velocities and ATPases. Although there was in general acorrelation between the rate of ADP release and the in vitro motility, as anticipated, the changes were not simply related to the properties of the myosins fromwhich the loops originated. Although changes in the charge and length of theloops did cause changes in the behavior of the expressed protein, the correlation
was not simple. The group therefore proposed that the primary property of theloop causing the inhibition of ADP release was the reduced flexibility of the loop.
Support for this idea came from several other properties of the myosin heads(either S1 or heavy meromyosin were changed by modifications to the loop),
including the rate of ATP binding and the ATP hydrolysis step. The loop links
two helices, one connecting to the P-loop and the second to the switch-1 region.
Thus Sweeney et al argue that changes in the flexibility of the loop could be communicated to the nucleotide-binding site and might modulate ADP release by amechanism similar to those discussed above.
Support for the role of Loop 1 controlling ADP release from actomyosin alsocame from studies of the catch and striated-muscle myosins of the scallop (66).
These myosins are coded from a single gene and the only significant change inthe myosin S1 is an alternate splice at exons 5 and 6, which includes the Loop-1
region (100). Isolation of the two myosins and the S1 molecules allowed
detailed biochemical analysis. The loop changes produced changes in in vitro
motility (fivefold) and in the basal and actin-activated ATPase rates (threefold).
Detailed transient kinetic analysis revealed that the only events modified by theexon change were a tenfold change in the affinity of ADP for acto-S1 (thought
to limit in vitro velocity) and the rate of Pi release. Thus at one level the loop
behaves as predicted by Spudich. However, in addition the changes affect the
key Pi release step, which is compatible with the idea that the loop communicates with the Pi-binding site. The change in sequence is EEEADQKK for the
striated to EPVPNLRA for catch, which would give a change in flexibility as
proposed by Sweeney.
Furch et al (97) made a series of systematic changes to the Loop 2 of Dictyostelium myosin-truncated S1. They increased the length of the loop by 4 to 20
residues and changed the net charge from -1 to +12. The results show that thebinding of ATP to the acto-S1 construct was unaffected by any of the changes,
consistent with it having no effect on the nucleotide-binding pocket. Similarly,
basal ATPase activity was also unaffected. In contrast, the affinity of the S1 fragment for actin was largely unaffected by length changes but increases in the number of positive charged residues dramatically increased the affinity of actin for
the nucleotide-free head (100-fold) and increased the apparent Km of actin (25fold) in ATPase measurements. This project resulted in the production of amyosin head which has high affinity (~10 µM in 20 mM KCl) for actin even in
the presence of ATP and may therefore prove useful for structural studies of the
weakly bound cross-bridge. This myosin may prove particularly useful whencombined with point mutations, which greatly reduce the rate of the ATP cleavage (16, 101; see below).

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The Hinge
The idea of a lever arm mechanism led naturally to an attempt to locate the site of
the fulcrum or pivot point for the lever arm. Both the crystal structure and thedependence of velocity, using in vitro motility assays, on the length of the leverarm indicated the two short helices (Sk691–714) carrying the reactive cysteines at
Sk697 and Sk707. The two helices are also of interest because they contain threeglycines (Sk699, 703, and 710) often used as hinge points in proteins. In particular SkG699 is at the bend between the two helices, and Huston et al (51) proposedthat substitution of SkG699 or SkG710 with bulkier residues might inhibit theconformational changes in this region. Kinose et al (102) prepared the equivalentof an SkG699A mutation in chicken embryonic myosin and showed that it had
impaired motility properties. The small yields limited the assays that could be performed to motility assays and actin-activated ATPase assays. Both were reduced(ten- and fivefold, respectively). Kinose et al (102) argued that the impaired motility (which was dominant, i.e. 2% mutant myosin in the presence of 98% wild typegave 50% inhibition of velocity) involved an increase in the residence time boundto actin. This probably represents a longer-lived AM.ADP or AM complex.
Using a random mutagenesis of the Dictyostelium myosin II, Patterson et al(103) isolated two myosins with mutations at DyG680V and DyG691C (homologous to Sk699 and Sk710). The expression and isolation of these myosins (andG680A, G691A, and S1-like fragments) allowed a wider range of assays. Mutations at either position reduced velocity in motility assays to 10% and, as in thechicken system, the DyG680V/A mutation was dominant when mixed with wild
type, whereas the DyG691C mutant was not. DyG680V had inhibited basal(without actin) ATPase at low temperature yet could be activated by actin,
whereas DyG691 had an elevated basal ATPase, which was not activated by actin.
The lack of actin activation of the ATPase is sufficient to account for the
reduced motility. Moreover, the nondominant effect on wild-type motility of
DyG691C suggests that the mutation accelerates the rate-limiting Pi release stepand, in doing so, decouples the P i release from the power stroke. For G680V,
Patterson et al (103) were able to show that the affinity for actin either in thepresence of ADP or during ATP turnover was enhanced and therefore concluded
that the main effect of the mutation was to produce a longer-lived AMD complex. The longer-lived AMD state can result from a reduced rate of ATP binding
to AM, increased binding of ADP to A.M (and slower release of ADP from
AM.ADP), or creation of a novel strongly bound actin state.
In a recent study, Batra et al (103a) have constructed the three mutations,
DyG680A, DyG684A, and DyG691A, in the M761-2R head fragment of Dictyostelium myosin II and have completed a detailed study of the interaction of
the mutants with actin and nucleotide. DyG684A is near normal in all of its properties. The ATPase data of G680A and G691A agree with the earlier data. For
G691 all actin- and nucleotide-binding properties are normal except the rate of Pi
release from myosin, which is accelerated. For G680A many major changes inproperties were observed, including 20-fold reduction in the rate of nucleotide

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binding to M but not AM, a 10-fold-higher affinity of ADP for M, and 100-foldhigher affinity for AM. As predicted by Patterson et al (103), the G680A mutant
appears to have a very long-lived AM.ADP state which will account for its ability to inhibit motility even in the presence of wild-type protein.

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Switch 1
The functional role of the residues in the switch-1 region of Dictyostelium myosin
II was examined by alanine scanning (104). The residues in the region
Dy233–240 have the sequence NXVSSRFG: Each residue was replaced one at atime by alanine. For each construct the ATPase, the in vitro motility, andnucleotide-binding properties were examined. The results showed that N233 andS237 were essential for nucleotide binding, whereas R238 was essential for thehydrolysis of ATP. The equivalent mutations R238C and R238H have been foundin the nonmuscle myosin VII in mice, which results in Ushers syndrome (105).
Other mutations produced myosins with moderate impairment of properties but
are all functional.

The R245–E468 Salt Bridge
The DyR238 (SkR245 SmR247) residue is of interest as it forms a conserved salt
bridge with DyE459 (SkE468 SmE470). Mutation of either residue 238 or 459
produces a myosin incapable of hydrolyzing ATP. In an elegant experiment,
Onishi et al (106) made the double mutant SmE470R/R247E in chicken smoothmyosin and showed that, although either single mutation resulted in loss ofATPase activity, the ability to hydrolyze ATP was recovered in the double mutant,although motor activity was somewhat impaired.
A mutation at E459 in Dictyostelium myosin II was found by Ruppel & Spudich (107) in their screening of random mutations in the 454–486 region of Dictyostelium myosin and was classified as a nonhydrolyzer. Three nonhydrolyzer
mutations were identified in the screen, E459V, N464K, and E476K. Two of these(E459V and E476K) were expressed in the myosin S1 fragment and characterized in detail (101). The E459V construct was particularly interesting because
the rate of ATP binding to M was slowed only fivefold and the ATP affinity
remained very high (dissociation rate constant 10-5 s-1), whereas the rate of the
ATP cleavage step was slowed from 30 s-1 to 5 3 10-5 s-1; that is, the lifetime of
the M*.ATP state was of the order of 5 hours. Such constructs could prove usefulin structural studies in the presence of actin.

Point Mutations of G466
A nearby residue was also examined in chicken smooth muscle SmG468A(equivalent to SkG466 DyG457) (108). This residue was thought to be involvedin a rotation to allow the formation of the salt bridge (SkR245, SkE468) and to8505_AR_23

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make an amide hydrogen bond to the gPi of ATP. The mutation produced amyosin with greatly reduced ATPase activity and no Pi burst (indicative of areduced rate of the ATP hydrolysis step), yet it bound actin in an ATP-sensitive
manner. It also failed to induce any increase in protein fluorescence on adding
ATP to the actin-free myosin. This result demonstrates the central role of thisresidue in bringing about the transition from OPEN to CLOSED and indicates,
furthermore, that the control of actin binding by ATP may proceed by a mechanism not closely correlated with this conformational change.

Cardiomyopathies
Another area that has identified mutations in the myosin head with important
physiological consequences is the study of familial cardiomyophathies (for a
recent review, see 109). The MYH7 gene, which codes for the myosin heavy
chain found in cardiac and slow-twitch muscle, has been extensively characterized, and 29 mutations have been located in myosin, of which 24 are in the headregion. The head mutations are clustered in four key sites: the actin-binding site,
the nucleotide-binding site, the area close to the essential thiol hinge region, andat the head–light-chain interface (110). Few of these mutations have been characterized in detail, and in most cases an interpretation of the effects on the headfunction is not possible.
Many of the mutations result in dysfunctional myosin and disrupted sarcomeric structures. For such mutations the myosin cannot be easily isolated,
although equivalent mutations can be expressed in mouse cell lines (111, 112), in
myotubes (113–116), or in Dictyostelium myosin. The first mutation ever analyzed, R403 (sk405), which is an actin-binding loop, has been expressed in theequivalent mouse gene (111, 112), R403 (sk405), and R247 (skR249) in cardiomyotubes or quail myotubes (113, 114, 116). The following mutations havebeen characterized in isolated fibers or in vitro motility assays, and all show
reduced motility: T124I (skT125), Y162C (skY163), R247Q (skR249), G256E(skG258), R403Q (R405), R453C (skR445), and V606M (skV607) (117, 118);
R719Q (skK721) shows increased motility (119). G741R and R403Q give
reduced velocity and lower force (120). The locations of the mutations aremarked on Table 1. When expressed in the equivalent positions in Dictyostelium,
R403Q, F513C, G584R, G716Q, and R719W (121) all show decreased function
in solution.

Mutations Should Aid Functional Analysis
The study of point mutations in the myosin molecule is really only just beginning. Although few mutations have yet been characterized in enough detail tointerpret their structural and functional significance, such studies already provide
clues as to which are important areas of the myosin molecule. Expression of theequivalent mutations in amenable hosts will allow a more thorough examinationof the solution properties. Moreover, in the near future one expects that the8505_AR_23

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mechanical properties will become available from single molecule assays. Only
very recently has our understanding of the function of this molecule reached alevel where one can begin to integrate the effects of mutations. However, withincreasing awareness one can expect that point mutations, both natural and synthetic, will add greatly to our understanding of the role of the different parts ofthe structure and in particular help define the pathways of information transfer
between the different functional regions.

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