Biophys Rev (2014) 6:237–260
DOI 10.1007/s12551-014-0141-7

REVIEW ARTICLE

The path to visualization of walking myosin
V by high-speed atomic force microscopy
Noriyuki Kodera & Toshio Ando

Received: 27 December 2013 / Accepted: 7 May 2014 / Published online: 18 June 2014
# The Author(s) 2014. This article is published with open access at Springerlink.com

Abstract The quest for understanding the mechanism of
myosin-based motility started with studies on muscle contraction. From numerous studies, the basic frameworks for thismechanism were constructed and brilliant hypotheses were
put forward. However, the argument about the most crucial
issue of how the actin–myosin interaction generates contractile force and shortening has not been definitive. To increase
the “directness of measurement”, in vitro motility assays andsingle-molecule optical techniques were created and used.
Consequently, detailed knowledge of the motility of muscle
myosin evolved, which resulted in provoking more arguments
to a higher level. In parallel with technical progress, advancesin cell biology led to the discovery of many classes of myosins. Myosin V was discovered to be a processive motor,
unlike myosin II. The processivity reduced experimental difficulties because it allowed continuous tracing of the motor
action of single myosin V molecules. Extensive studies ofmyosin V were expected to resolve arguments and build aconsensus but did not necessarily do so. The directness ofmeasurement was further enhanced by the recent advent ofhigh-speed atomic force microscopy capable of directly visualizing biological molecules in action at high spatiotemporal
N. Kodera : T. Ando
Bio-AFM Frontier Research Center, Kanazawa University,
Kanazawa 920-1192, Japan
T. Ando (*)
Department of Physics, College of Science and Engineering,
Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japane-mail: tando@staff.kanazawa-u.ac.jp
N. Kodera
PREST, Japan Science and Technology Agency, 4-1-8 Honcho,
Kawaguchi 332-0012, Japan
T. Ando
CREST, Japan Science and Technology Agency, 4-1-8 Honcho,
Kawaguchi 332-0012, Japan

resolution. This microscopy clearly visualized myosin V molecules walking on actin filaments and at last provided irrefutable evidence for the swinging lever-arm motion propellingthe molecules. However, a peculiar foot stomp behavior also
appeared in the AFM movie, raising new questions of thechemo-mechanical coupling in this motor and myosin motors
in general. This article reviews these changes in the researchof myosin motility and proposes new ideas to resolve the
newly raised questions.
Keywords Myosin . Actin . Muscle . Motor proteins .
High-speed AFM . Imaging

Introduction
Motions in biological systems are vital phenomena. They
inspire our curiosity to understand how motion is made possible. Skeletal muscles exhibit striking physiological action(i.e. contraction) that is visible to the naked eye and is dailyexperienced by ourselves. It was, therefore, natural that research into biological motility started with the analysis ofmuscles. Biochemical studies of muscle proteins revealed thatthe molecular entities responsible for generating contractile
force and shortening comprise just three: myosin, actin, andATP. Each head of the double-headed myosin hydrolyzes ATP
into ADP and inorganic phosphate (Pi). The ATPase rate isvery low when myosin is alone but is markedly accelerated by
its interaction with actin, where the chemical energy liberated
by ATP hydrolysis is converted into mechanical work. This
energy conversion has most attracted our curiosity and thus
been extensively studied using physiological, biophysical,
biochemical, structural biological, and theoretical methods.
It is noteworthy that several experimental techniques havebeen applied or created to study muscle contraction, and later
used in other fields of biological science. Results obtained in238each discipline have built important frameworks and hypotheses for the mechanism of muscle contraction. For example:(1) the relationship between load, speed of shortening, and therate of heat production in muscle contraction (Hill 1938,1964); (2) the filament sliding mechanism in which muscle
shortens as a result of the mutual sliding of thin filaments
containing actin relative to thick filaments containing myosin(Huxley 1953; Huxley and Hanson 1954; Huxley andNiedergerke 1954); (3) the generation of force by myosincross-bridges that extend from the myosin filaments to theadjacent actin filaments in the overlap zone in the sarcomere(Huxley 1957a, b); (4) the hypothesis of swinging lever-arm
as powerstroke (Huxley 1969); (5) the concept of a working
stroke of ∼10 nm derived from the swinging lever-arm hypothesis and from the observation of tension recovery following a quick release applied to contracting muscle fibers(Huxley and Simmons 1971); and (6) the kinetic scheme of
the actomyosin ATPase reaction (Lymn and Taylor 1971).
These ideas were integrated into a general consensus on themuscle contraction mechanism. Nonetheless, a clue to greater
clarification of the chemo-mechanical reaction in actomyosin
unexpectedly emerged from the discovery of non-muscle
myosins together with the creation of single-molecule biophysical techniques.
Currently, there are 35 (or more) distinct classes of myosins
designated I to XXXV (Berg et al. 2001; Thompson andLangford 2002; Richards and Cavalier-Smith 2005; Fothet al. 2006; Odronitz and Kollmar 2007). They constitute thesuperfamily of actin-based motors that play crucial roles indynamic cellular processes. In the initial attempt to find nonmuscle myosins, first a single-headed myosin (Pollard andKorn 1973) and subsequently a traditional double-headed
myosin (Maruta and Korn 1977) were isolated fromAcanthamoeba. Later, the former was called “myosin I” andthe latter “myosin II”. Thereafter, the Roman numerals havebeen given according to the chronological order of discovery.
Among the 35 classes of myosins, myosin V (Mehta et al.
1999; Sakamoto et al. 2000), VI (Rock et al. 2001; Nishikawa
et al. 2002), VII (Yang et al. 2006), X (Nagy et al. 2008; Ricca
and Rock 2010; Sun et al. 2010), and XI (Tominaga et al.
2003; Hachikubo et al. 2007) are known to be processive
motors, i.e. a single molecule undergoes multiple catalytic
cycles and mechanical advances before dissociation from an
actin filament. This processivity promoted the study of myosin motility because it enabled us to continuously trace themotor action of individual molecules. Note that not all isoforms in each of these classes are processive. For example, theyeast myosin V, Myo2p (Reck-Peterson et al. 2001), and theChara myosin XI (Kimura et al. 2003) are not processive,
although Myo2p is switched to a processive motor when
tropomyosin is bound to actin filaments (Hodges et al. 2012).
In parallel with the progress of non-muscle myosin studies,
fluorescence microscopy (see review by Joo et al. 2008) and

Biophys Rev (2014) 6:237–260

optical trap nanometry (see review by Neuman and Block2004) were developed to observe the dynamic action of proteins at the single molecule level. These techniques assisted
the discovery of several properties of myosin V (hereafterreferred to as M5) including three-to-five successive stepadvances under load with the ∼36-nm step size (Mehta et al.
1999), processive runs of several micrometers under no load
(Sakamoto et al. 2000), load-dependent stepping kinetics(Veigel et al. 2002), the hand-over-hand manner of this movement (Forkey et al. 2003; Yildiz et al. 2003), and the tight
relationship between the catalytic cycle and the 36-nm advance (Sakamoto et al. 2008). These discoveries led to acomprehensive view for the action of myosin motors. Nonetheless, a complete consensus has not been reached and moredetails of the motor action and the underlying chemomechanical coupling have remained elusive.
Looking back on the long history of myosin motility studies, we notice that the gap between experimental data andconclusions derived therefrom has been slowly reducing. Thisis because the level of “directness of measurement” has been
increasing. Before the single-molecule era, we had to infer theaction of individual molecules from experimental data
ensemble-averaged over many molecules. Several details of
the action of myosin were hidden in the ensemble-averagemeasurements on bulk samples. However, even when singlemolecules were observed, their level of directness was not
perfect because the subset of observed molecular events depends on the site where an optical probe is placed while theentire molecular structure remained invisible. Myosin has
been studied by X-ray crystallography, NMR, and electron
microscopy, but these structures are limited to static snapshots
and the simultaneous assessment of structure and dynamics
seemed impossible.
High-speed atomic force microscopy (HS-AFM), whichstarted to be developed around 1993 to overcome this limitation, is now sufficiently advanced (Ando et al. 2008) to allow
us to video-record the structure and dynamics of functioning
biomolecules at single-nanometer resolution, without
disturbing their function. This new microscopy was applied
to M5 (Kodera et al. 2010). The obtained image data revealed
details of its molecular action, leading not only to the corroboration of known or inferred behavior of the motor but also to
new discoveries (Kodera et al. 2010). In particular, it was asurprise to discover that the tension responsible for forward
movement can be generated without transitioning through an
ADP–Pi-bound state, meaning that no chemical energy input
is required for the tension generation. Moreover, the lever-arm
swing (powerstroke) by the leading head spontaneously occurs when the trailing head detaches, thus demonstrating thatno chemical energy input is required for the lever-arm swing
either. These findings appear inconsistent with a popular
scheme put forward for the chemo-mechanical reaction inmuscle contraction (e.g., Geeves and Holmes 1999; Goldman

Biophys Rev (2014) 6:237–260

2004). Thus, we now need to develop a unified motor mechanism that can be applied to all classes of myosin motors.
This review does not cover the entire range of myosin
motors but focuses on studies of the motor mechanism, particularly those performed with the use of single-molecule
techniques and our recent HS-AFM study of M5 motility.
Readers are referred to reviews on muscle contraction(Geeves and Holmes 1999; Huxley 2000, 2004; Goldman2004), on structure–function relationships of myosins(Ruppel and Spudich 1996; Sweeney and Houdusse 2010a),and on the cellular functions of various myosins (Hartman
et al. 2011; Coluccio 2008).

General aspects of myosin
The myosin superfamily is largely categorized into the monomeric and dimeric types (Pecham 2011), although some myosins can adopt both forms depending on the environment(Park et al. 2006; Yu et al. 2009; Tokuo et al. 2007; Umeki
et al. 2011) and both types of isoforms sometimes exist in thesame class (e.g., monomeric yeast Myo4p and dimeric yeast
Myo2p belonging to class V; Bookwalter et al. 2009). In stable
dimeric myosins, the N-terminal head is followed by a coiledcoil tail domain for dimerization, while in monomeric myosins it is followed by a tail domain or a stable, single α-helical(SAH) domain (Knight et al. 2005) and then by a tail domain.
The C-terminal tail end generally serves as a cargo-binding
domain (Fig. 1). In this regard, dimeric myosin II is rather
unique. It does not possess a cargo-binding domain and assembles into bipolar thick filaments mediated by a longcoiled-coil tail. The tail and cargo-binding domains havediverse amino acid sequences, depending on the intracellularfunction of myosin (e.g., Sellers 2000), while the head region
comprising a motor domain and a neck domain is generallycommon. The motor domain contains two functional sites, theactin-binding and nucleotide-binding sites which communicate with each other. The neck domain comprises a variable
number of IQ motifs that bind calmodulin or myosin lightchains. The neck is connected to the so-called converter region
in the head, and these two as a single entity is called the leverarm. When the SAH domain follows the neck, it is also
included in the lever-arm because it functionally extends thelength of the canonical lever-arm (Baboolal et al. 2009). The
heavy chain in the head comprises three parts (25, 50, and 20–
23 kDa) aligned in this order from the N-terminus that wererevealed by limited tryptic cleavage of subfragment-1 (S-1) ofskeletal myosin II (Mornet et al. 1979; Mornet et al. 1981).
These three parts connected to two surface loops are arranged
as shown in Fig. 2 in the three-dimensional (3D) structure of
the head (Rayment et al. 1993a; Houdusse et al. 2000). This
3D arrangement of the three parts is largely common to allclasses of myosin. The most prominent feature is the presence239

of a large cleft dividing the 50-K subdomain into lower
and upper 50-K subdomains. The outer cleft regions ofboth lower and upper 50-K subdomains bind to actin
mainly through loops. The ATP-binding site is locatedat the inner cleft region, around which all the three parts
face each other. Interestingly, the 3D structure of thecatalytic core is very similar to the entire motor domain
structure of kinesin despite complete dissimilarity in theamino acid sequence, suggesting that they share a similar
force-generating strategy (Kull et al. 1996).
Single-headed myosin S-1 binds to actin at a given
angle relative to the actin filament. The bound single head
(both motor and neck domains) often extends towards theplus end of actin at least in the nucleotide-free and ADPbound conditions (Rayment et al. 1993b; Jontes et al.
1995). This orientation towards the plus end of actin iscalled “arrowhead orientation” (Huxley 1963). Among all
members of the myosin superfamily, only myosin VI isknown to move towards the minus (barbed) end of actin(Wells et al. 1999). The motor domain of a single-headed
myosin VI construct also binds to actin in the arrowheadorientation. However, its neck domain is largely kinked
relative to the motor domain, so that the neck domain
orients slightly towards the minus end of actin (Wells
et al. 1999). The swinging lever-arm hypothesis presumes
that the lever-arm rotates so that a minute change in the
motor domain is amplified to a large displacement of thelever-arm’s distal end. When this presumption is applied tomyosin VI, its reversal movement must be achieved by therotation of the lever-arm in the direction opposite to that of
the other myosins (Wells et al. 1999). This is consistent
with the kinked neck domain of myosin VI and also withthe reversal movement of a myosin I construct with anartificial neck which is kinked towards the reversearrowhead orientation (Tsiavaliaris et al. 2004).
The actomyosin ATPase reaction proceeds as shown in
Fig. 3. Immediately after binding to ATP, the actin-boundhead detaches from actin, followed by quick hydrolysis of
the bound ATP to ADP–Pi. The ADP–Pi-bound head has a
low affinity for actin (yet a somewhat stronger affinity than the
ATP-bound head). When the ADP–Pi-bound head is attached
to actin, the bound Pi dissociates from the head, followed by
the formation of a strongly-bound complex of A–M–ADP (Aand M denote actin and myosin head, respectively), and thenby ADP dissociation. The nucleotide-free head also has astrong affinity for actin. These complexes with strong affinities are called “rigor complexes”. Thus, a myosin head has
two states with respect to its affinity for actin, the weak- andstrong-binding states. The main role of actin in the ATPasereaction is to accelerate the otherwise very slow Pi and ADP
dissociation from a myosin head. The degree of acceleration
for Pi dissociation and ADP dissociation varies among theclasses of myosin. The processivity occurs in dymeric240

Biophys Rev (2014) 6:237–260

Fig. 1 Schematic showing domain organization of myosin molecule.
The tail part is variable and contains an α-helical coiled-coil, a SAH
domain, or a chain segment with a variable amino-acid sequence,

depending on the myosin class or isoform. When a SAH domain follows
the neck, it is included in the lever-arm because it functionally extends
the canonical lever-arm

myosins when the lifetime of A–M–ADP predominates over
the other states (De La Cruz et al. 1999).
In the prevailing view on myosin motility, the chemomechanical coupling is considered to occur as follows (seeFig. 3). The myosin head undergoes conformational changes
upon ATP binding and hydrolysis to ADP–Pi. Myosin thenswings back the lever-arm (“recovery stroke”) (Geeves andHolmes 1999). The angle change between the motor and neckdomains caused by this recovery stroke facilitates the bindingof the head to actin in the reverse arrowhead orientation (notethat the motor domain still binds to actin in the arrowheadorientation). The A–M–ADP–Pi state, often termed as the“pre-powerstroke state”, is considered to be a higher-energy
state. Upon Pi release, tension for driving the lever-arm swing
is generated, the lever-arm actually swings to execute a
powerstroke, and then a lower-energy state of A–M–ADP(the post-powerstroke state) is formed. Thus, in this view,the chemical energy liberated by ATP hydrolysis is usedmainly in the Pi-release step.

Progress of motility assays

Fig. 2 Crystal structure ofskeletal myosin II S-1. The heavy
chain of myosin is shown as aribbon diagram, while two lightchains [i.e. essential light chain(ELC) and regulatory light chain(RLC)] are shown as a spherediagram. The 25-, 50-, and 20kDa regions are colored green,
red, and blue, respectively. The
converter domain in the 20-kDa
region is additionally coloredwith orange, after which IQ
motifs follow

The situation before the single-molecule era
Almost the entire enzymatic kinetics of actomyosin has beenobtained from muscle myosin. In most of kinetics measurements, soluble forms of myosin, S-1 and heavy meromyosin(HMM), were used to simplify interpretations of the data.
Then, the enzymatic kinetics and the structural characteristics
of myosin were combined to understand how the myosin
motor might work. Although the overall feature of the enzymatic kinetics was successfully revealed, no mechanical constraints were imposed to myosin molecules interacting withactin filaments in vitro. Therefore, it was difficult to deducethe most important characteristic of the mechano-enzyme,
namely how does mechanical strain imposed on myosin affectthe ATPase reaction? Moreover, the whole cycle of theATPase reaction could not be tracked down in one measurement because the initial synchrony among molecules becomesBiophys Rev (2014) 6:237–260

241

Fig. 3 Scheme of muscle actomyosin ATPase reaction including structural model. + indicates the plus end of actin. Boxes with solid lines andbroken lines represent the strong- and weak-affinity states, respectively.
The powerstroke is considered to occur mainly in the transition from the

A–M–ADP–Pi state to the A–M–ADP state. The recovery stroke isconsidered to occur in the transition from the M–ATP state to the M–
ADP–Pi stateobscured by the stochastic nature of the reaction. Therefore,
many kinetic constants derived for the entire reaction scheme
were obtained in separate experiments where the reactions
were started with different initial states. In some cases, it
was difficult to prepare adequate initial states. As such, aquantitative reaction scheme was obtained by “pasting” many
results together that were not necessarily obtained under similar solution conditions.
The mechanical behavior of myosin interacting with actin
filaments could not be observed in solution experiments. To
acquire this information, muscle fibers and myofibrils were
subjected to X-ray diffraction (Huxley et al. 1981, 1983),
fluorescence polarization (Nihei et al. 1974; Borejdo andPutnam 1977), electron paramagnetic resonance (EPR) spectroscopy (Thomas and Cooke 1980), and force analyses (Fordet al. 1977). At a very early stage intrinsic tryptophan fluorescence or fluorescent ATP analogues were used for fluorescence polarization studies (dos Remedios et al. 1972a, b).
Later, fluorescence and spin polarization probes were introduced to a specific cysteine, SH1, in the motor domain. Thiswas possible because SH1 is most reactive of all myosin
cysteines (Sekine and Kielly 1964) and indeed of all proteinsin skinned muscle fibers (Borejdo and Putnam 1977), andtherefore could be selectively labeled. Unfortunately, it later
became known that the SH1-labeled myosin does not function
as a motor (Root and Reisler 1992; Marriott and Heidecker
1996; Bobkov et al. 1997). Extensive muscle fiber X-ray
diffraction studies were performed but no clear evidence forcross-bridge movement was obtained. In the light of numerous spectroscopic and structural studies, the massive part of an
actin-bound myosin head proximal to actin was suggested toremain unchanged during the powerstroke, whereas the distalneck region with smaller mass than the proximal one wassuggested to change its orientation (Cooke 1986). In this caseit would be difficult to distinctly detect this change in the fiber

diffraction patterns. Moreover, the time in which a myosin
head interacts with an actin is very short in the ATPase cycle,
making it difficult to record the dynamics of cross-bridges.
The molecular structures of myosin heads in different
nucleotide conditions were successfully revealed at
submolecular level using electron microscopy (e.g., Walker
and Trinick 1988; Katayama 1989; Pollard et al. 1993; Takács
et al. 2010) and at the atomic level using X-ray crystallography (e.g., Rayment et al. 1993a; Fisher et al. 1995;
Dominguez et al. 1998; Houdusse et al. 2000). However, asa matter of course, these structures were static snapshots.
Moreover, even with the current state of technology, it is still
difficult to obtain the atomic structure of actin-bound myosin
head, particularly the structure in the power-stroking state(Rayment et al. 1993b; Lorenz and Holmes 2010).
In vitro motility assays
To overcome some of the above adversities, experimental
techniques called in vitro motility assays were developed in
the early 1980s, as advanced versions of previous methods
developed in the 1970s (Oplatka and Tirosh 1973; Kuroda andKamiya 1975). Dark-field microscopy was performed that
detected the movement of fibrils that were squeezed out of
the giant alga Nitella cell, washed with buffer, and mixed with
ATP (Higashi-Fujime 1980). Next, fluorescence microscopy
was conducted using a more purified system where skeletal
myosin-coated fluorescent beads were observed to slide on asubstratum of polar arrays of actin cables derived from giant
alga, Nitella (Fig. 4a) (Sheetz and Spudich 1983). The movement was ATP-dependent and blocked by inactivation of the
myosin heads using N-ethylmaleimide. This study revealed
that the bipolar thick filament structure is not necessary for
generating force and movement. Remarkably, this assay system was much simpler and more robust than previous in vitro

242

motility assay systems (Oplatka and Tirosh 1973; Kuroda andKamiya 1975; Higashi-Fujime 1980; Yano et al. 1982), andthis facilitated quantitative analyses of the myosin motility.
Using fluorescently labeled single actin filaments, their thermal bending motion was observed to increase in the presenceof HMM and ATP (Yanagida et al. 1984). Also was demonstrated that the sliding movement of individual actin filamentson myosin-coated surfaces occurred at velocities similar to
those recorded in muscles (Fig. 4b) (Spudich et al. 1985).
Moreover, actin filaments were observed to slide on oneheaded myosin filaments, indicating that a two-headed structure is not needed for motility (Harada et al. 1987). This result
was reinforced by the observation of actin filaments
gliding over myosin S1-coated surfaces (Toyoshima
et al. 1987). Thus, motor activity inherent in the head
was established, which ruled out some models of muscle
contraction.

In vitro motility studies on chemo-mechanical relationship
Understanding how actomyosin ATPase kinetics determines
the mechanical performance of this motor system (magnitude
of generated force and sliding velocity) became a central focus
of muscle research. Even in the in vitro motility assays, single
actin filament sliding and the translational motion of myosincoated beads are propelled by multiple myosin heads. Therefore, the chemo-mechanical relationship was not straightforward. Nonetheless, the in vitro motility assays made theanalysis more precise and more extensive than previous assays where muscle fibers were studied. The in vitro motility
assays allowed us to investigate actomyosin motility underwide varieties of solution conditions with myosins and actinsfrom diverse species and cell types. The velocity of actin
filament sliding over myosin-coated surfaces and the velocity
of myosin-coated beads along actin filaments were shown tobe analogous to the speed of unloaded shortening of muscle
fibers (Sheetz et al. 1984; Spudich et al. 1985; Kron andSpudich 1986; Homsher et al. 1992). Despite the reported
analogy, many studies using different types of myosin II foundno direct correlation between the steady-state actomyosin
ATPase activity and the rate of movement measured by thein vitro motility assays (Umemoto et al. 1989; Lowey et al.
1993; Uyeda et al. 1994; Vale et al. 1984; Higashi-Fujime1991). Nor was direct correlation observed using different
nucleotide triphosphates (NTPs) condition (Shimizu et al.
1991; Higashi-Fujime and Hozumi 1996). In these studies,
one concentration of actin and one concentration of NTPs
were used in the measurements of the enzymatic activities.
In similar studies where the actin concentration was varied
and a fixed concentration of NTPs was used, a moderatecorrelation was shown between the motor activity and thesubstrate turnover rate (Pate et al. 1993a; Regnier et al. 1998).

Biophys Rev (2014) 6:237–260

A more extensive study was conducted for rabbit skeletal
acto-HMM using six naturally occurring NTPs with Mg2+,
different divalent cations with ATP, and Mg2+-ATP at various
ionic strengths (Amitani et al. 2001). The maximum substrate2+
turnover rates (Vmax), Km for actin (KAm), and Km for metal NTPs (KNm) were obtained while the velocity of actin filamentgliding on HMM-coated surfaces was measured under thesame buffer and substrate conditions used for the measurements of substrate turnover rate. A large dataset acquired
revealed a direct linear relationship, except when different
divalent cations were used with ATP. The maximum sliding
velocity (Vmaxs ) extrapolated to the infinite concentration of
1/2
substrates was directly proportional to (VmaxKA(Fig. 5).
m)
This experimental correlation was analyzed based on a force–balance model, where the average sliding force (active force)
produced by a productive head, <fs>, is balanced with theaverage resistive force produced by an actin-bound nonproductive head, <fr>, which is directly proportional to thegliding velocity of the actin filament. At infinite substrateconcentration, <fs> equals pTp/Tc, where p is the magnitude
of powerstroke (force spike), Tp is the duration of single
powerstroke, and Tc is the substrate turnover time. The powerstroke speed (d/Tp; where d is actin filament displacement by
each powerstroke) should equal the maximum sliding velocity(Vmaxs ) (Uyeda et al. 1991). Thus, the following relationship isderived:< f s >¼ pdV max =V max
s :

ð1Þ

At infinite substrate concentration, the resistive force averaged over the substrate turnover time <fr> is expressed as:< f r >¼

1
λΓ V maxs =k – ;2

ð2Þ

where λ is the fraction of the actin-bound non-productive stateover all states, Γ is an elastic constant of the resistive head, and
k– is the dissociation rate of the resistive head. Supposing thatthe resistive state is the weakly bound state, k– approximately
equals k+KAm, where k+ represents the rate constant in which a
detached head in the weak-binding state binds to actin. Thus,
from Eqs. (1) to (2), the following relationship is derived:
V max¼ 2k þ pd K Asm V max =λΓ

1=2

:

ð3Þ

As the fraction of the resistive state λ is likely to be
proportional to k+, Vmax
becomes directly proportional tos
1/2(VmaxKAm) , which agrees well with the experimentally observed relationship (Fig. 5). The lifetime of the weakly boundstate is very short but this state occurs many times within aBiophys Rev (2014) 6:237–260

243

Fig. 4 Schematics showing
in vitro motility assay systems foractomyosin. a Myosin-coatedbead assay. The myosin-coatedfluorescent beads are subjected tothe polar arrays of actin cables
naturally formed on chloroplastdof the alga Nitella, and movementof the beads are observed under aflorescent microscope. b Actin
filament gliding assay. Myosin
molecules are attached to thesurface of a nitrocellulose-coatedcoverslip and gliding motion of
the fluorescently labeled actin
filaments are observed under afluorescence microscope

substrate turnover cycle. Thus, the weakly bound state becomes a predominant resistive state. This force balance model
could also be extended to the case where actin filaments
moved at various substrate concentrations by including theresistive force produced by rigor heads; thus, it could derive
the dependence of sliding velocity on the substrate concentration. Its derivation showed that the dependence of the sliding
velocity on the substrate concentration [S] followed a modinN
fied Michaelis–Menten equation of Vs =Vmaxs /{1+(Km/[S]) }(here, 1 < n < 2), which again agreed well with theexperimentally obtained dependence under various conditions(Amitani et al. 2001).
Step size
The next fundamental question to tackle experimentally
was the translocation distance that is propelled by actin—
myosin interaction during an ATPase cycle (i.e. step size).
The swinging lever-arm hypothesis predicts the size to be∼10 nm. However, as mentioned above, the in vitro actin
gliding or myosin-bead gliding assay is not a singlemolecule assay. Moreover, the duration of the working
stroke that occurs in an ATPase cycle has to be measured
to estimate the step size but cannot be measured directly.
Consequently, various values were reported for the step
size: ∼8 nm (Toyoshima et al. 1990), ∼10 nm (Pate et al.
1993b), 10–28 nm (Uyeda et al. 1991), and >60 nm(Yanagida et al. 1985; Harada et al. 1990). To resolve thislarge discrepancy, the step size had to be directly measured
at nanometer and millisecond spatiotemporal resolution.
This requirement gave birth to the so-called “single-molecule measurement assays” capable of observing and manipulating single biological molecules in action.
Single-molecule measurements

Fig. 5 Relationships between the maximum sliding velocity (Vmaxs ) andthe actomyosin NTPase kinetics parameters, Km for actin (i.e. KAm) andmaximum NTPase rate Vmax. The nucleotide substrates used are indicatedby the side of the corresponding data plots

Optical tweezers are typically used in single molecule
measurements. A micrometer-sized dielectric particle
such as a polystyrene bead is trapped by a focused laser
beam (Ashkin and Dziedzic 1985; Neuman and Block2442004) and a trapped particle can be freely moved bychanging the focal position of the laser beam. The position of the trapped particle in the XY plane perpendicular
to the laser beam can be measured with ±<1 nm precision. For the trapped particle, the focused laser beamworks as a 3D spring with a spring constant of 0.01–
10 pN/nm. When the particle is displaced by an external
force in the XY plane, the magnitude of the external
force applied can therefore be measured at high precision. This optical trap nanometry is now routinely used
to study biological molecules (Svoboda and Block 1994;
Mehta et al. 1998; Wang 1999; Bustamante et al. 2000;
Kuo 2001). Step size and the force produced by a single
myosin head were directly measured using a dual-beamlaser trap (Finer et al. 1994; Molloy et al. 1995; Guilford
et al. 1997; Tanaka et al. 1998; Veigel et al. 1999). Eachend of an actin filament was attached to a particle. The
two particles are caught in the dual-beam laser trap, andthen the actin filament is allowed to interact with asingle myosin molecule immobilized on a surface(Fig. 6a). The measured step size ranged between 3.5
and 15 nm but the extent of discrepancy between laboratories was much smaller than previous estimates madeusing actin gliding assays. It was pointed out by
Yanagida’s group that the measured step size tended tobe affected by the interaction angle between myosin andactin. With careful design, the measured step size wasshown to be dependent on the interaction angle (Tanaka
et al. 1998). When an actin filament was aligned parallelwith a myosin filament with a low myosin head density,
a relatively large step size of 10–15 nm was obtained(Tanaka et al. 1998).
A very fine glass micro-needle was also used by
Yanagida’s group as a force transducer for single moleculemechanical measurements. A single actin filament was
attached to the free end of a micro-needle and brought into
contact with an immobilized single myosin molecule. The
displacement of the micro-needle caused by the myosin–
actin interaction was measured (Kishino and Yanagida
1988; Ishijima et al. 1991; Ishijima et al. 1996). Using a
low needle stiffness (0.09 pN nm−1) and near zero load, theaverage of single displacement spikes was ∼20 nm(Ishijima et al. 1996). However, the spatiotemporal resolution was not high enough to precisely detect the displacement spikes. Then, an experimental system was further
developed (Fig. 6b); a ZnO crystal whisker with a tip radiusof ∼15 nm was attached to a fine glass needle mounted on athree-dimensional piezoelectric actuator. A single myosin
S-1 fluorescently labeled at the regulatory light chain was
attached to the end of the ZnO whisker. The molecule wasbrought into contact with actin bundles formed by chicken
gizzard α-actinin, where the actin filaments were aligned inanti-parallel (Meyer and Aebi 1990). Using a glass needle

Biophys Rev (2014) 6:237–260

with very small stiffness (0.01–0.03 pN nm−1), the spatial
resolution of 2.0 nm and the temporal resolution of <0.2 ms
were obtained (Kitamura et al. 1999, 2005). Using thistechnique, an average step size of ∼13 nm was measured,
but surprisingly several (2–5) 5.5-nm successive substeps
were also detected within each step. This size, 5.5 nm,
coincides with the distance between adjacent actin monomers in a filament. More surprisingly, the dwell between
substeps was independent of the ATP concentration (0.1
and 1 μM). Therefore, it was concluded that the multiple
substeps produced by a single myosin head occurred during
just one cycle of ATP hydrolysis. This conclusion was
reinforced by theoretical studies (Terada et al. 2002;
Takano et al. 2010). However, the experimental result has
never been reported by other groups.
Total internal reflection fluorescence microscopy(TIRFM) has also been used for single-molecule measurements (Park et al. 2007; Murcia et al. 2007; Joo et al. 2008).
When an incident laser light is totally reflected at the glass–water interface, an evanescent field is generated that decays
exponentially from the interface, penetrating to a depth of∼100 nm into the solution. Since only fluorophores locatedin the evanescent field are excited, the background lightlevel is very low, allowing the visualization of single
fluorophores (Fig. 6c). Using a Cy3-labeled ATP, the dynamic events of its binding to and dissociation from single
myosin S-1 molecules were visualized (Funatsu et al.
1995). The dissociation rate estimated from the lifetime of
the bound Cy3-nucleotide agreed with the biochemicallymeasured rate of Cy3-ATP turnover by S-1. This success
opened up a new opportunity to simultaneously measure
the chemical and mechanical events in single myosin molecules. Using one-headed skeletal muscle myosin assembled with myosin rod and a setup where TIRFM and a dualbeam laser trap were combined, mechanical events of actin–myosin dissociation/force generation (displacement)
and chemical events of binding/dissociation of Cy3-ATP(or Cy3-ADP) were simultaneously observed (Ishijima
et al. 1998). The timing of Cy3-ATP binding well coincidedwith the timing of myosin head dissociation from actin.
However, in more than 50 % of 85 detected events, theactin filament was displaced 0.3–1.8 s after the dissociationof Cy3-nucleotide from the myosin head. This surprising
result suggested that myosin has a hysteresis or memory
state in which the chemical energy liberated by ATP hydrolysis is stored even after ADP dissociation. However,photobleaching of bound Cy3-nucleotide also looks like it
is dissociated from the myosin head. Therefore, their interpretation could not be statistically verified. The step size
measured in this study (∼15 nm) was consistent with theirpreviously reported values.
Single-molecule measurements quantified other mechanicalproperties of actin, myosin, and their interactions. The tensile

Biophys Rev (2014) 6:237–260

245

Fig. 6 Schematics showing typical single-molecule measurement systems for studying myosin–actin interactions. a Dual-beam laser trap
assay. Each end of an actin filament is attached to a plastic bead, andthe beads are trapped by the focused laser beams. The trapped actin
filament is brought to a single myosin molecule attached to a silica bead.
b Mechanical measurement system with the use of a thin glass microneedle as a force sensor. A ZnO crystal whisker is attached to the end of a
glass micro-needle (not shown in this schematic). Fluorescently labeled
single myosin S-1 is attached to the apex of the ZnO crystal whisker via a
specific binding of biotin-streptavidin. The tip is brought onto actin
bundles formed by α-actinin. The displacements of the S-1 resulting fromthe interaction with actin are monitored by the detection of glass microneedle deflection. c TIRFM experiment for visualizing individual ATP
turnovers by single myosin S-1. The light chain of S-1 molecule is labeledwith Cy5 (indicated by the red star) for the confirmation of whether or not
the observed ATP turnovers are those by a single S-1 molecule. Once
Cy3-ATP binds to the S-1, the large Brownian motion of the Cy3-ATP
stops and therefore its fluorescent spot becomes visible. d Single beam
assay for the measurement of unbinding force of a single myosin–actinbond. e AFM measurement of rupture force and rupture distance of asingle actin–myosin bondand torsional rigidity of a single actin filament was determined using a glass micro-needle technique (Kishino andYanagida 1988) and optical tweezers (Tsuda et al. 1996),
respectively. The unbinding force between an actin filamentand a single myosin head in the nucleotide-free conditionwas ∼9 pN using an optical tweezers method where an actin
filament was pulled approximately along its length(Fig. 6d) (Nishizaka et al. 1995). It was also measured tobe ∼15 pN with AFM, where a single myosin head was
pulled in the direction perpendicular to the actin filament(Fig. 6e) (Nakajima et al. 1997).
As described above, no complete consensus was reachedamong research groups regarding the myosin step size taken
for each ATP hydrolysis, and hence also as to the swinging

lever hypothesis. The discrepancy was due partly to a relatively small step size occurring in a short time that wastechnically difficult to measure.

Myosin V
The technical difficulty mentioned above was significantly
removed by the discovery of M5 that possesses unique structural and biochemical features. M5 is involved in organelle
and mRNA transports as well as in membrane trafficking. Its
cellular functions and regulations are well described elsewhere (Reck-Peterson et al. 2000; Taylor 2007; Sellers and246

Knight 2007; Sellers and Weisman 2008), so here we focus on
its motor mechanism.
Structural and biochemical features of M5
The most prominent structural feature of two-headed M5 is its
neck domain (∼24 nm) (Cheney et al. 1993) which is approximately three-fold longer than that of myosin II (∼8 nm). If M5
really follows the swinging lever-arm hypothesis, a step size
markedly larger than that of muscle myosin II should beobserved. M5 was biochemically shown to dominate in a highactin-affinity state (ADP-bound state) during its ATPase cycle(De La Cruz et al. 1999), in stark contrast with muscle myosin
II where a weak affinity sate (ADP–Pi-bound state)
dominates.
Processive movement
A dual-beam optical trap was used to show that a suspended
actin filament successively moved 3–6 times when it
interacted with a single M5 molecule attached onto a bead
(Mehta et al. 1999). The dwell time depended on the ATPconcentration, and therefore, successive step movements differed from the sub-stepping observed previously with myosin
II (Kitamura et al. 1999, 2005). This was the first evidence that
M5 is a processive motor. Importantly, the step size was∼36 nm, consistent with the swinging lever-arm hypothesis.
A much longer processive run (over several microns) of single
M5 molecules along surface-immobilized actin filaments wasdemonstrated with TIRFM (Sakamoto et al. 2000). In a singlebeam optical trap, a trapped bead coated with M5 at low
density moved processively with ∼36-nm steps along asurface-immobilized actin filament (Rief et al. 2000). The∼36-nm step size corresponds to the head-head distanceshown in electron micrographs of M5 with two heads boundto the same actin filament (Walker et al. 2000). From dwell
time analyses of the data, the stepping kinetics was shown tobe limited by ADP release. In this study, a hand-over-handmodel was first proposed, i.e. the two heads alternately switch
the leading and trailing positions in every ATPase cycle.
The processivity is consistent with the cellular functions ofM5 (e.g., organelle transports). The motility of kinesin, acargo-carrying microtubule-based processive motor, has been
well characterized although its step size is relatively small(∼8 nm) (Vale et al. 1985; Block 2007; Hirokawa et al.
2009). M5 possesses both processivity and a large step size,
which were of great advantage in studying details of how M5
moves.
Inhibitory state of M5
In one of the above studies on the full length of M5 (Sakamoto
et al. 2000), the ATP turnover rate was measured

Biophys Rev (2014) 6:237–260

biochemically under the same buffer solution conditions(without Ca2+) used for the observation of long processive
movement. From the maximum actin-activated ATPase ratemeasured (∼1.2 s−1/head) and the maximum velocity of M5
movement observed with TRIFM (1 μm s−1), M5 appeared totravel ∼400 nm per ATPase cycle. This long travel distance
implied that a large fraction of M5 (more than 90 %) would bein a catalytically inhibited or low-activity state. Any M5 not inthis state would have a much higher ATPase activity (14 s−1/head). Interestingly, in previous studies an ATPase rate as highas 14 s−1/head was reported for single-headed M5 constructs
in the absence of Ca2+ (De La Cruz et al. 1999; Trybus et al.
1999) as well as for the full length of M5 in the presence ofμM Ca2+ (Nascimento et al. 1996). Later, this mystery of thefast movement (1 μm s−1) with a low ATPase activity(∼1.2 s−1/head) was solved by a finding that M5 often takes
a folded conformational state in the absence of Ca2+, in which
the C-terminal cargo-binding domains bind to the heads,
resulting in the inhibition of the ATPase and motor activities(Wang et al. 2004; Krementsov et al. 2004; Li et al. 2006; Liuet al. 2006; Thirumurugan et al. 2006; Olivares et al. 2006).
Also, several other myosins such as myosin VII and X are
now known to be regulated by head–tail interactions (Yanget al. 2009; Umeki et al. 2009, 2011). Intramolecular folding
appears to be a common mechanism to inhibit the activity of
motor proteins as a similar conformational transition wasshown previously with smooth muscle myosin (Trybus et al.
1982) and kinesin (Hackney et al. 1992). However, a recent
study of smooth muscle myosin suggests that an intramolecular interaction between the heads would be a key event in theregulation, rather than a head–tail interaction (Baumann et al.
2012).
Processive hand-over-hand movement
The ∼36-nm step size of M5 corresponds to the long pitch
pseudo-repeat of the actin helix, suggesting that M5 moves ona plane along an actin filament. This inference was certainly
proven to be the case by the observation of the movement ofan M5-attached bead on a suspended actin filament. The beadadvanced in a left-handed spiral trajectory with one revolution
per 2.3 μm, i.e. only 5.6° rotation per step (Ali et al. 2002).
The next study to be done was to confirm the tight relationship between the neck length and the step size, using M5
constructs with different neck lengths. It was actually confirmed that the step size is directly proportional to the necklength (Purcell et al. 2002; Sakamoto et al. 2003; Moore et al.
2004; Sakamoto et al. 2005). Thus, the swinging lever-arm
hypothesis was nearly secured.
Next, many studies focused on the experimental verification and close inspection of the hand-over-hand movement
previously proposed (Rief et al. 2000). To this end, various
light microscopy techniques or fluorescent probes wereBiophys Rev (2014) 6:237–260

introduced to TIRFM: fluorescence imaging with one nanometer accuracy (FIONA) (Yildiz et al. 2003), fluorescencepolarization (Forkey et al. 2003), two-color imaging with Qdots (Warshaw et al. 2005), single-molecule high-resolution
colocalization of fluorescent dyes (Churchman et al. 2005), a
combination of FIONA and fluorescence polarization (Syedet al. 2006), and defocused orientation and position imaging(Toprak et al. 2006). All these studies revealed that in fact M5
walks in a hand-over-hand manner. Not only translational
position changes of the two heads but also angular changesof the lever-arm relative to actin were observed where only
one of the two lever-arms was labeled with a fluorescentprobe. The value of the angle change (∼60°) was also consistent with a hand-over-hand movement (Forkey et al. 2003;
Syed et al. 2006; Toprak et al. 2006).
These studies supported the hand-over-hand model but
only the dwell states were observed when both heads werebound to actin. Therefore, little was known about the fleetingintermediate between the dwell states because the time resolution was low (∼33 ms or longer). To increase the time
resolution to 1 ms or less, the following imaging methods
were employed: dark field imaging of a 40-nm gold nanoparticle attached to one of the lever-arms (Dunn and Spudich2007) and recording of light-scattering from a 200-nm M5coated latex bead traveling in an interference fringe pattern(Cappello et al. 2007). Alternatively, the action of walking M5
was slowed down by attaching a large fluorescent microtubule
to one of the lever-arms and observed by conventional fluorescence microscopy (Shiroguchi and Kinosita 2007). All
three studies revealed that the detached head underwent extensive rotational Brownian motion before landing on a siteahead of the leading head, suggesting that the detached head isunlikely to interact with actin during the Brownian motion.
Moreover, it was suggested that the Brownian motion is likely
to be biased forward by the orientational change (swing) of theactin-bound leading head, by which the detached head eventually finds an actin ahead of the leading head to become a
new leading head. Thus, a clearer picture of the processive
hand-over-hand movement was obtained.
Asymmetry of enzymatic kinetics between two heads
The processive hand-over-hand movement suggests that only
the trailing head can detach from actin. If so, there are two
possibilities: (Case 1) ADP dissociation followed by ATPbinding predominantly occurs at the trailing head, so that the
ADP dissociation would be accelerated at the trailing head or
retarded at the leading head; (Case 2) the ATP-binding ratewould be enhanced at the trailing head or suppressed at theleading head. Between these possibilities, Case 1 appeared
more likely as the ATPase rate is limited at the ADP releasestep. In either case, the kinetics asymmetry between the twoheads would be engendered by head-to-head communications

247

through intramolecular strain. This kinetics asymmetry wasaddressed by solution kinetics studies and optical-trap
nanometry studies. In the solution kinetics studies various
kinetics constants were compared for S-1 and HMM constructs of M5, although the two heads of M5-HMM do notnecessarily interact with the same actin filament in solution(Rosenfeld and Sweeney 2004; Forgacs et al. 2008). Thesestudies suggested retardation of ADP release from the leadinghead by ∼45-fold (Rosenfeld and Sweeney 2004) or up to
250-fold (Forgacs et al. 2008). The latter study used new
fluorescent ADP and ATP analogues (deac-aminoADP and
deac-aminoATP) (Webb et al. 2004) whose emission intensity
increases by 20-fold upon binding to the active site of M5(Forgacs et al. 2006). In the dual-beam trap experiments, theactin filament that was interacting with a surface-attachedsingle-headed M5 molecule was pulled forward or backward
to apply a force to the actin-bound head (Purcell et al. 2005;
Veigel et al. 2002, 2005). The forward and backward forceswere hypothesized to simulate forces experienced by the
trailing or the leading head, respectively. Different designsof optical trap nanometry were also used to study the coordination between actin-bound two heads (Uemura et al. 2004;
Clemen et al. 2005; Gebhardt et al. 2006; Oguchi et al. 2008).
Most of these studies suggested that the intramolecular strain
would coordinate two heads by decelerating ADP releasefrom the leading head, whereas some studies suggested accelerated ADP release at the trailing head (Veigel et al. 2002,
2005), as proposed by a study of processive run of M5
perturbed by varying the nucleotide content (Baker et al.
2004).
In two-headed bound M5, the neck of the trailing headtakes the arrowhead orientation, whereas the neck of theleading head takes the reverse arrowhead orientation. The
leading head conformation must be mechanically distorted
because single-headed M5 binds to an actin filament in anarrowhead orientation. In fact, electron micrographs of twoheaded bound M5 showed that the leading neck was curved
slightly outwards whereas the trailing neck was straight(Walker et al. 2000; Burgess et al. 2002; Oke et al. 2010).
This conformational distortion seems consistent with the suggestion that the coordination of M5 stepping is performed bystrain-mediated inhibition of ADP release from the leadinghead.
Finally, using two-color TIRFM, Sakamoto et al. (2008)
simultaneously observed the stepping motion of fluorescently labeled M5 and the binding and dissociation of thefluorescent nucleotide (deac-aminoADP or deacaminoATP). This observation directly revealed the preferential ADP dissociation from the trailing head, which was
followed by a ∼36-nm step triggered by ATP binding. It
also revealed that every M5 molecule always keeps at least
one nucleotide (ADP in the leading head) during its
stepping, even at low ATP concentrations.

248

Discrepancies and unsolved questions in myosin V motility
The above description suggests there is now a consensus on
how M5 steps. In every ATP hydrolysis cycle, the leadinghead swings to propel M5 forwards by ∼36 nm (Sakamoto
et al. 2008). This large stride is realized by the long lever-armsof M5 (Purcell et al. 2002; Sakamoto et al. 2003; Moore et al.
2004; Sakamoto et al. 2005). The two heads alternately switch
between the leading and trailing roles so that M5 walks handover-hand (Yildiz et al. 2003; Forkey et al. 2003; Warshaw
et al. 2005; Churchman et al. 2005; Syed et al. 2006; Toprak
et al. 2006). This style of movement is made possible by the
intramolecular strain-mediated retardation of ADP releasefrom the leading head (Rosenfeld and Sweeney 2004;
Purcell et al. 2005; Forgacs et al. 2008; Oguchi et al. 2008;
Sakamoto et al. 2008).
Nonetheless, several observations that are difficult to reconcile with this generally agreed view have been reported,
mostly from the Yanagida lab. A recombinant two-headed M5
with a single IQ motif on each neck (∼4 nm long) was reported
to move processively with ∼36-nm steps (Tanaka et al. 2002).
It was claimed that the long-neck domain was not essential forboth large step size and processivity of M5 but the motor
domain alone determined the processivity and the large step
size. It was also reported that single-headed M5 constructs
underwent multiple successive large (∼32-nm) steps on an
actin filament suspended by a dual-beam optical trap
(Watanabe et al. 2004). They also underwent directional diffusion on surface-immobilized actin bundles with ∼5.5-nm
substeps to develop an average displacement of ∼20 nm,
which was independent of the neck length (2IQ and 6IQ
motifs) (Okada et al. 2007), as previously observed withsingle-headed myosin II (Kitamura et al. 1999). Likewise,
myosin VI with its short neck similar to myosin II, wasreported to move with ∼36-nm steps (Rock et al. 2001;
Nishikawa et al. 2002). However, among these reported irreconcilable conclusions, only the large step size of myosin VI
was resolved. The SAHs were found in the proximal taildomain of myosin VI, which contribute to the lever-arm
length (Baboolal et al. 2009; Spudich and Sivaramakrishnan
2010; Sweeney and Houdusse 2010b). The other discrepant
results are still incomprehensible (Cyranoski 2000; Geeves
2002).
Even if the above general view of how M5 steps iscompletely right, there remain fundamental questions to solve.
How is the chemical energy liberated by ATP hydrolysis used
to generate the tension for forward movement? How are thetension generation and lever-arm swing coupled to the ATPase
cycle? If they are coupled, which chemical transitions are
involved in these mechanical events? These fundamental
questions have long been explored in muscle contraction
studies and some are considered to be resolved. Are there
any inconsistencies when this consensus view is applied to

Biophys Rev (2014) 6:237–260

the motility of M5? Numerous single-molecule measurementsof M5 stepping have not answered these important questions.
The discrepancies mentioned above, as well as the remaining unsolved fundamental questions, have certainly proventhat the level of “directness of measurement” is not yet highenough even with single-molecule optical measurements. Proteins molecules themselves are indeed invisible in these measurements, which can never be overcome even withdiffraction-breaking super-resolution fluorescence microscopy (Egner and Hell 2005; Huang et al. 2009).

HS-AFM
HS-AFM has overcome this technical limitation. Biologicalmolecules can be directly observed at sub-molecular spatialand sub-100 ms temporal resolution without disturbing theirfunction. In this section, we briefly summarize the principle of
AFM and the current state of HS-AFM performance (Fig. 7).
Techniques involved in the HS-AFM instrument are welldescribed elsewhere (Ando et al. 2001; Ando et al. 2008;
Ando 2012). See other reviews for protocols of HS-AFM
imaging of protein molecules (Uchihashi et al. 2012) andvarious application studies of HS-AFM (Ando et al. 2014).
Principles of AFM imaging
In AFM (Binnig et al. 1986), a sharp tip attached to the free
end of a cantilever traces the sample to acquire its surface
topography while the sample stage is scanned laterally. AFM
can observe any object on a substrate at single-nanometer (orhigher) resolution under a wide range of environments (vacuum, air, and liquids). Topography images of biological samples can be acquired under physiological buffer conditions,
without fixing, staining and labeling the sample (Engel andMüller 2000; Müller and Dufrêne 2008). Among various
imaging modes, the tapping mode (Zhong et al. 1993) is mostsuited for biological samples. In this mode, the cantilever is
excited to oscillate in the Z-direction at or near its first resonant frequency. Because of this oscillation, the cantilever tip isbrought intermittently into contact with the sample surface,which can eliminate lateral friction force and hence minimize
deformation of fragile samples. The reduction of cantilever
oscillation amplitude upon tip-sample contact is measured andthen the sample stage is finally moved in the Z-direction to
restore the amplitude to its set point using a feedback control.
This series of operations is repeated many times for different
sample surface points during lateral scanning of the sample
stage. Consequently, the 3D movement of the sample stage
approximately traces the sample surface, and hence, a topography image of the sample can be constructed from the electric
signal used to drive the scanner in the Z-direction.

Biophys Rev (2014) 6:237–260

249

Fig. 7 Schematic of HS-AFM system. The system includes an original
inverted optical microscope. The objective lens with a long working
distance that is a part of the OBD detector is also used to view thecantilever and sample stage via a digital camera. The glass slide, to which
the cantilever holder and the liquid cell are attached, is placed on the
optical microscope stage. A cantilever chip is held in the holder so that its
tip points upward (opposite to the way in conventional AFM). The
sample stage, attached to the Z-scanner and facing downward, is placed
over the cantilever. An incident laser beam passing through the objective
lens is focused onto the small cantilever, and the light reflected back fromthe cantilever is collected and collimated by the same objective lens andguided to the quadrant-cell Si PIN photodiode. The incident and reflected
laser beams are separated using the quarter-wavelength (λ/4) plate and thepolarization beam splitter. A cantilever immersed in a buffer solution isoscillated with small amplitude (1–2 nm) by the excitation with an
excitation piezo at the first resonant frequency of the cantilever (0.6–
1.2 MHz in water). The cantilever oscillation amplitude is maintained
constant by feedback control. A counterbalancing method is employed to
the Z-scanner to minimize unwanted vibrations; two peazoactuators areattached to the supporting base at its counter sides and simultaneously
displaced in the same distance, in the counter directions. The scan signals
for the X- and Y-scanner are output from the computer through the DA
converters. The output from the active Q-control damper is recorded as
the sample height through the AD converter. The active Q-control damperis constructed with an LRC circuit whose transfer function is very similar
to that of the Z-scanner. Its electrical output behaves in a way similar to
the Z-scanner displacement

Current state of HS-AFM

disturbing their biological function. The spatial resolution of
HS-AFM is now comparable with that of conventional slow
AFM even with high-bandwidth detection of the cantilever
oscillation amplitude and fast scanning of the sample stage(∼2 nm for XY and ∼0.1 nm for Z in the best case). HS-AFM
has already visualized dynamic biomolecular phenomena (seereviews in Ando 2012; Ando et al. 2013, 2014) and even
dynamic phenomena in live cells (Yamashita et al. 2012;
Watanabe et al. 2013).

With conventional AFM instruments it takes at least 30 s tocapture an image. This slow rate is due mainly to the slow
response of the mechanical devices (i.e. cantilevers and Zscanner), whereas many interesting biological phenomena
occur much faster. Various efforts have been made to markedly increase the imaging rate of tapping-mode AFM over the
past 20 years. Small cantilevers, fast scanners, and fast amplitude detectors were developed to build the first prototypic
instruments (Viani et al. 2000; Ando et al. 2001). The improvement of these devices and creation of new techniques forvibration damping and fast feedback control produced thecurrent HS-AFM (Ando et al. 2008). The highest possible
imaging rate is a function of the speed of an AFM instrument
as well as the imaging conditions and the sample fragility(Uchihashi et al. 2012; Ando et al. 2013). For 50- to 200-nm
scan ranges, sufficient to image biological molecules, HSAFM can capture an image within 30–80 ms without

Walking M5 filmed by high-speed AFM
Substrate surface and observation of unidirectional movement
M5-HMM was directly imaged walking along actin filaments
using HS-AFM (Kodera et al. 2010). Here, partially biotinylated actin filaments were immobilized on a surface where

250

streptavidin was dispersed at low density on mica-supported
planar lipid bilayer containing electrically neutral phospholipids and a biotinylated lipid (Fig. 8a). M5-HMM was neverbound directly to the lipid bilayer surface and only interactedwith the immobilized actin filaments and moved thereon. The
moving M5 was frequently oriented perpendicular to thesubstrate surface so that its characteristic lateral topography
was only occasionally observed. Adding a positively charged
lipid to the bilayer solved this problem.
In the presence of 1–2 μM ATP, successive AFM images
captured at 7 fps clearly showed M5-HMM moving
processively with discrete ∼36-nm steps (Fig. 8b). The twoheaded bound M5-HMM exhibited unique structural features(Fig. 8c). The junction of the neck with the motor domain
appears smooth in the leading head but is always V-shaped in
the trailing head because the neck regions emerge from different parts of the motor domain. The short coiled-coil tail was
mostly tilted towards the minus end of actin. These structuralfeatures, which are totally consistent with an electron microscopic observation (Burgess et al. 2002), can be used todetermine the actin polarity when bound M5-HMM is stationary. In addition, the leading head always assumed a straightconformation (slightly curved outwards), which agreed withthe straight-neck model proposed for walking M5 (Forkey
et al. 2003; Syed et al. 2006; Toprak et al. 2006) but disagreed
with the bent-neck model (Walker et al. 2000; Burgess et al.
2002; Snyder et al. 2004; Oke et al. 2010).
The positively charged lipid in the lipid bilayer slightly
retarded the translocation velocities. When it was absent, thevelocities at various ATP concentrations were similar to thosemeasured by fluorescence microscopy under the same buffer
solution condition (Forkey et al. 2003; Baker et al. 2004),
indicating negligible effects of the tip-sample interaction on
the motor activity. However, molecular behavior during a step
could not be resolved as it completed very fast.
Visualization of stepping behavior
To slow the step, more streptavidin molecules were placed on
the lipid bilayer surface as moderate obstacles to the walkingof M5-HMM (Fig. 9a), which resolved the stepping process(Fig. 9b). Upon detachment of the trailing head from actin, theleading head appeared to spontaneously rotate from the reverse arrowhead orientation towards the arrowhead orientation. Before completing this rotation, the leading head was
halted for a moment by colliding with a streptavidin molecule
placed on the way of its natural path. In this halt state, thedetached trailing head was displaced forward, positioned most
distant from the actin filament (thus, the two heads werealigned nearly straight, pointing opposite directions), andslightly rotated around the neck-neck junction (the second
frame in Fig. 9b). Then, the leading head overcame the obstacle and further rotated forwards. Immediately after this the

Biophys Rev (2014) 6:237–260

trailing head bound to a forward site of the actin filament tobecome a new leading head, completing one step. Thus,
dynamic processes in the forward step were directly visualized. The observed rotation of the leading head is exactly theswinging lever-arm motion itself initially proposed for the
powerstroke of muscle myosin (Huxley 1969). Note that
before completing a step the detached trailing head neverinteracted with actin but passively moved forwards driven
by the rotating leading head. This ruled out some models of
M5 motility such as the ‘inchworm’-like model considered forkinesin (Hua et al. 2002) and the “biased diffusion” model
proposed for single-headed myosin II (Kitamura et al. 1999)
and single-headed M5 (Okada et al. 2007). M5 strictly followed the hand-over-hand mechanism. In contrast, the otherprocessive myosins VI and X, which function not only as acargo transporter but also as a structural anchor in cells, werereported to move irregularly, namely inchworm-like stepping,backward stepping, and forward hand-over-hand stepping(Yildiz et al. 2004; Sun et al. 2007; Nishikawa et al. 2010;
Ricca and Rock 2010; Sun et al. 2010).

Foot stomp and unwinding of coiled-coil tailInterestingly, in the two-headed bound state during the dwell,
the motor domain of the leading head frequently exhibited
brief dissociation and reassociation on the same actin filament,
while the molecule remained at approximately the same position on the filament (Fig. 10a). Similarly, the motor domain of
the trailing head exhibited a brief translocation by ∼±5 nm
along the actin filament. We termed these behaviors “footstomp”. The foot stomp was more frequently observed at theleading head than at the trailing head (by a ratio of approximately 3:1). Although not well documented, a foot-stomp-likebehavior was previously suggested in fluorescence microscopy observations of walking M5 molecules (Syed et al. 2006;
Shiroguchi and Kinosita 2007). More recently, the foot stomp
was further confirmed by the observation of walking M5 byhigh-speed single-molecule polarized fluorescence microscopy (Beausang et al. 2013). Thus, the foot stomp is an inherent
behavior of this motor.
The foot stomp at the leading head raises an importantquestion as to the chemo-mechanical coupling in M5. The
briefly detached leading head does not carry bound Pi because
Pi has already been released from the leading head following
its initial attachment to actin (De La Cruz et al. 1999; Olivares
et al. 2006). Nevertheless, the detached leading head with its
bound ADP reattached to actin while still in the reversearrowhead orientation. It then swung forward following
trailing head detachment. This fact was important because it
indicated that tension generation for forward movement canoccur without transitioning through the ADP–Pi bound state,i.e. it can occur directly in the ADP–bound state. Thus, theBiophys Rev (2014) 6:237–260
Fig. 8 HS-AFM imaging ofwalking M5-HMM. a Schematic
showing assay system used for
the HS-AFM imaging. b
Successive HS-AFM images
showing unidirectional
processive movement of M5HMM observed in 1 μM ATP.
Frame rate, 7 fps; Scan area,
130×65 nm2 with 80×40 pixels;
scale bar 30 nm. The verticaldashed lines show the centers ofmass of the motor domains, andthe plus sign indicates the plus
end of actin. c Schematic showing
two-headed bound M5-HMM toactin. The corresponding AFM
movies can be seen at thefollowing web site: http://www.
nature.com/nature/journal/v468/
n7320/extref/nature09450-s2.
mov

Fig. 9 Stepping behavior of M5HMM visualized by HS-AFM. a
Schematic explaining the HSAFM images shown in (b). b
Successive HS-AFM images that
resolved the stepping behavior of
M5-HMM in 1 μM ATP. Frame
rate, 7 fps; scan area, 150×75 nm2
with 80×40 pixels; scale bar
50 nm. The swinging lever-arm ishighlighted with the thin whitelines. The vertical dashed lines in(a) and (b) represent the centers ofmass of the motor domains, andthe plus sign indicates the plus
end of actin. The corresponding
AFM movies can be seen at thefollowing web site: http://www.
nature.com/nature/journal/v468/
n7320/extref/nature09450-s3.
mov

251

252

Biophys Rev (2014) 6:237–260

Fig. 10 Unique molecular behaviors of M5-HMM visualized by HSAFM. a Successive HS-AFM images showing a foot stomp event that
occurred at the leading-head (ATP, 1 μM; frame rate, 7 fps; scan area,
150×75 nm2 with 80×40 pixels; scale bar 50 nm). The leading-headdetachment is marked with the light-blue arrowheads. The corresponding
AFM movie can be seen at the following website: http://www.nature.
com/nature/journal/v468/n7320/extref/nature09450-s3.mov b HS-AFM
images before (upper panel) and after (lower panel) unwinding of
coiled-coil tail observed in 50 μM ADP. The corresponding AFM
movie can be seen at the following website: http://www.nature.com/
nature/journal/v468/n7320/extref/nature09450-s5.mov c HS-AFM

image showing the leading head with a sharply bent conformation
observed in the nucleotide-free condition. The corresponding AFM
movie can be seen at the following website: http://www.nature.com/
nature/journal/v468/n7320/extref/nature09450-s6.mov d HS-AFM
image showing nucleotide-free M5-HMM with heads bound to adjacentactin subunits (upper panel) and its illustration (lower panel). Imaging
conditions for (b), (c) and (d) are as followings: frame rate, 3 fps; scanarea, 100×100 nm2 with 80×80 pixels; scale bar 30 nm. The verticaldashed lines represent the centers of mass of the motor domains, and theplus signs indicate the plus ends of actin

tension generation for forward movement does not seem torequire chemical energy be supplied by ATP hydrolysis.
The asymmetry of foot stomp frequency between the twoheads suggested their actin-affinity difference, consistent witha biochemically measured result (Olivares et al. 2006). The
leading head binds to actin in the unnatural orientation (i.e.
reverse arrowhead orientation), and hence, pays an energy
cost by distorting the neck conformation, resulting in a lower
affinity for actin. The distortion of the leading head is likely tobe the source for tension generation for forward movement.
This was reinforced by the following observation; when the
two-heads were bound to actin in 1 mM ADP, the short coiledcoil tail was sometimes unwound, immediately after which themonomerized leading head rotated towards the arrowheadorientation, similar to the swinging lever-arm motion observed when M5-HMM was walking forwards (Fig. 10b).

1 mM ADP. This bent conformation suggested that the neckmotor domain junction is less flexible in the nucleotide-freehead than in the nucleotide-bound head. To examine thisissue, the arrowhead orientation angle of single-headed M5
relative to the actin filament was measured under the
nucleotide-free and ADP-bound conditions. In nucleotidefree solution, the angle was ∼34°, whereas in ADP the angle
was distributed widely, peaked at ∼29° and ∼51° in the proportion of approximately 6 to 1 (supplementary data in
Kodera et al. 2010), which may be relevant to the two different
ADP-bound states in equilibrium as reported for myosin II(Geeves 1989; Geeves and Holmes 1999), M5 (Robblee et al.
2005; Hannemann et al. 2005; Oguchi et al. 2008; Jacobs et al.
2011), and myosin VI (Robblee et al. 2005). Therefore, ADP
binding to the head makes the hinge of the neck–motor
domain junction flexible. Because of the rigid hinge of thenucleotide-free head, the leading neck in the reverse arrowhead orientation tends to be sharply bent to release the large
strain accumulated therein. This rigid hinge was also supported by the observation that both heads are rarely bound toadjacent actin subunits but only in the nucleotide-free condition (Fig. 10d). In the electron micrographs of actomyosin II

Flexibility of neck-motor domain junction
Under nucleotide-free conditions, the leading head frequently
exhibited a sharply bent conformation (Fig. 10c) that wasnever observed in the presence of ATP or in the presence ofBiophys Rev (2014) 6:237–260

and actomyosin VI complexes, this conformation was frequently seen, suggesting they have rigid hinges (Craig et al.
1980; Nishikawa et al. 2002).
The flexibility of the neck-motor domain junction observed
for the actin-bound M5 head with ADP does not seem sufficient to account for the two-headed binding in ADP because
the leading head binding in the reverse-arrowhead orientation
requires the junction to bend by at least ∼60°, even when thecontribution of the neck domain’s flexibility is considered.
This apparently restricted flexibility suggests that the actinunbound head with ADP is more flexible than the actin-boundone. In the other words, actin binding may make the neckmotor domain junction less flexible. This plausible flexibilitychange seems an excellent strategy for facilitating both actin
binding and the generation of enough tension to execute a
powerstroke. Very recently, such an actin-binding effect on theflexibility was proposed for muscle myosin (Billington et al.
2014).
Asymmetry of ADP dissociation rate
The straight and sharply bent conformations of the leadinghead depend on the presence or absence of bound ADP,
respectively, meaning that one can judge whether or not theleading head contains ADP by just looking at its conformation. From the proportion and lifetime of the straight conformation as a function of ADP concentration, the rate constantsof ADP binding/dissociation kinetics at the leading head were
estimated. The ADP dissociation rate constant was 0.1 s−1, i.e.
one ADP is released from the leading head every 10 s, on
average. However, M5 walks many steps during 10 s. Thus,
from the structural point of view, it was confirmed that thesequential events of ADP release, the subsequent ATP binding
and the resulting head dissociation take place solely at thetrailing head, which is the basis for the processive hand-overhand movement (Rosenfeld and Sweeney 2004; Purcell et al.
2005; Veigel et al. 2005; Forgacs et al. 2008; Oguchi et al.
2008; Sakamoto et al. 2008).

New questions
The swinging lever-arm hypothesis is no longer a hypothesis.
HS-AFM visualized its occurrence with irrefutable clearness.
However, from HS-AFM observations of M5-HMM
interacting with actin, questions were raised about thechemo-mechanical coupling in this motor as well as myosin
motors in general. The prevailing coupling mechanism had
been modeled in the context of swinging lever-arm motion(Fig. 3). In this model, the structural states are putatively
tightly coupled to the nucleotide states of the motor domain(Fig. 3). In the M–ADP–Pi state, the head takes the pre-253

powerstroke conformation caused by recovery stroke of thelever-arm, which facilitates binding of the head to actin in thereverse arrowhead orientation. Coupled to Pi release from theactin-bound head, the intramolecular tension is considered tobe generated just once by the pre-powerstroke to postpowerstroke transition. However, the observation with HSAFM of walking M5-HMM showed that the tension responsible for the lever-arm swing can be generated directly by the
ADP-bound leading head (after foot stomping). In addition, it
was shown that two-headed bound M5-HMM with ADP
alone can generate sufficient tension to cause occasional unwinding of the short coiled-coil tail, after which the lever-arm
swings. Thus, the recovery stroke or the pre-powerstroke
conformation thought to occur uniquely in the ADP–Pi-boundhead does not seem necessary for its binding to actin andlever-arm swing. More seriously, the chemical energy liberated by ATP hydrolysis does not seem to be used for theprocesses of recovery stroke, tension generation, and leverarm swing, suggesting that M5 would step forward withoutchemical energy input once the trailing head detaches fromactin.
This new idea of “no chemical energy usage” in theseprocesses would, however, inevitably lead us to encounter a“perpetuum mobile problem”. Even in ADP or in the
nucleotide-free condition, foot stomp occurs at both leading
and trailing heads of two-headed bound M5-HMM, with ahigher frequency in the leading head than in the trailing head.
When the trailing head detaches from actin, the molecule takes
a forward step by the spontaneous swing of the leading head.
On the other hand, when the leading head detaches from actin,
the molecule cannot take a backward step because the trailinghead is bound to actin in a stable orientation (i.e. the arrowhead orientation). Thus, using only thermal energy, M5 would
step forward many times (albeit very slowly), even with lessfrequent occurrence of foot stomps at the trailing head. This is
a forbidden perpetuum motion. However, in thisgedankenexperiment, positional fluctuations of thesingle-headed bound molecule due to the flexible neckmotor domain junction are not considered. Taking this
into account, we can expect a backward step after detachment of the leading head from actin, albeit with a low
probability. The ratio of foot stomp occurrence between
the leading and trailing heads was observed to be 4:1 inthe presence of 1 mM ADP. However, the foot stomp
events at the trailing head mostly occurred as a brief ±5nm translocation. Therefore, when this brief translocationis omitted and only the events of brief detachment fromactin are counted as foot stomp, the ratio of foot stomp
occurrence between the two heads becomes much larger.
Thus, the principle of detailed balance holds between theforward step after trailing head detachment and the backward step after leading head detachment, resulting in nonet movement of the molecule.

254

From numerous X-ray crystallography and electron microscopy studies, there is no doubt that myosin changes its
conformation upon binding nucleotides and takes a prepowerstroke-like conformation in the ADP–Pi-bound state(Fisher et al. 1995; Smith and Rayment 1996; Dominguez
et al. 1998; Houdusse et al. 2000; Burgess et al. 2002;
Coureux et al. 2004; Volkmann et al. 2005). These static
structures have been used to construct the prevailing modelof the chemo-mechanical coupling (Fig. 3) assuming there is atight one-to-one relationship between the chemical and conformational states. In reality, the relationship is not so tight.
Myosin takes two (or multiple) conformations even under agiven nucleotide condition. They can go back and forth but the
dynamic equilibrium is biased to one side depending on the
nucleotide condition. The HS-AFM observations of M5HMM indicates that structural fluctuations indeed exist andtherefore two-headed binding occurs even in the ADP-boundand nucleotide free conditions, consistent with other studies(Walker et al. 2000; Rosenfeld and Sweeney 2004; Olivares
et al. 2006).
The flexibility of myosin head conformation raises a question whether the chemical energy of ATP hydrolysis is reallyused to change the conformation. In the prevailing view, themyosin head with a strained (tension-generating) conformation in the pre-powerstroke state is achieved using the chemical energy of ATP, which is finally released by the swinging
lever arm motion. However, the pre-powerstroke and postpowerstroke conformations are in dynamic equilibrium withno marked energy barrier. Thus, one can conclude as follows:
although the conformational change of myosin head causedby recovery stroke is an effective strategy for facilitating its
binding to actin in the reverse arrowhead orientation, theenergy required for producing a pre-powerstroke conformation and the tension-generating state is much less than that to
be liberated by ATP hydrolysis. Therefore, the conformationalchange can occur thermally without chemical energy input. At
first glance this new view seems inconsistent with the recent
report (Shiroguchi et al. 2011) in which the energy barrier
between the pre-powerstroke state conformation (formed immediately after ATP addition) and post-powerstroke conformation (formed in the absence of nucleotides) was estimatedto be 5.2 kBT (kB, Bolzmann constant; T, room temperature in
Kelvin). For this estimation, they monitored the angular distributions of a large bead duplex attached to the motor domainof a single-headed M5 construct, while the molecule wasimmobilized to a surface of a 50-nm bead by the neck region.
This estimated energy barrier seems too large to be achieved
thermally. However, the angle distribution measurement did
not include the contribution of the flexible neck region to theangle fluctuations, because the neck region was attached to asurface. If it was included, the energy barrier could be smaller.
Moreover, the observed angular distributions only reflect projections in the image plane. Therefore, the distributions are

Biophys Rev (2014) 6:237–260

underestimated and hence the energy barrier is overestimated.
In fact, from solution kinetics measurements of muscle myosin ATPase reaction, the free energy change in the recovery
stroke-involved transition from M–ATP to M–ADP–Pi is
estimated to be 2 kBT (Howard 2001). Moreover, the attachment of a large bead duplex to the head would significantly
impede the otherwise frequent thermal transitions between the
pre-powerstroke and post-powerstroke states.
Let us perform another gedankenexperiment in the presence of ADP alone. Suppose each head of M5 is labeled with achromophore. Upon light absorption, the chromophore on thebound head will alter its conformation so that it dissociatesfrom actin. When a light pulse is illuminated only at thetrailing head of the bound M5, it spontaneously steps forward
after detachment of the trailing head. By repeated light illumination only at the trailing head, the molecule would movefor a long distance unidirectionally.
It seems plausible that a large fraction of the chemicalenergy liberated by ATP hydrolysis is used elsewhere andcould be used to detach the actin-bound nucleotide-freehead from actin. How much energy is required? As mentioned in “Single-molecule measurements”, the unbinding(rupture) force was previously measured to be ∼15 pN by
AFM (Nakajima et al. 1997). The rupture distance was also
estimated to be 1.7–2.5 nm in this study. This distance isunusually long compared to 0.23 nm for antigen–antibody(Hinterdorfer et al. 1996), 0.15–0.3 nm for streptavidin–
biotin (Yuan et al. 2000), and 0.05–0.3 nm for α-actinin–
actin (Miyata et al. 1996). However, it is consistent withthe fact that actin–myosin interfaces contain several bonds
formed by flexible loops (Milligan 1996; Kabsch et al.
1990). From the values of rupture force and rupture distance, the energy required to rupture the actin–myosin
rigor bond is 6–9kBT, which is 30–45 % of the chemicalenergy of ATP hydrolysis (∼20 kBT). This mechanicallyestimated value is approximately consistent with the valuesof 9 and 10kBT estimated biochemically for the free energychanges in the dissociation A–M → A + M and the ATPinduced dissociation A–M + T → A + M–T, respectively(Howard 2001). One may argue against this statement. For
instance, ATP is not yet hydrolyzed in the actin–myosin
dissociation step, and hence the energy of ATP is not yet
liberated in this step. However, the account balance ofenergy should be considered for the whole cycle of theATPase reaction. Therefore, unlike chemical cleavage reactions, in the enzyme-catalyzed reaction cycle the ATPbinding energy can be used as a large part of the energy
liberated by ATP hydrolysis. The role of the sequential
ATPase reaction process after ATP binding and head dissociation from actin would lie in ensuring the unidirectional progression of the mechanical process. For this assurance, only a small fraction of ATP energy, comparable tothermal energy, is sufficient.

Biophys Rev (2014) 6:237–260

Conclusions
We have shown how our understanding of the myosin motor
mechanism has advanced with increasing level of directnessof measurement. When the level of directness is low, arguments can continue without resolution. For example, in theefforts to provide evidence for or against the swinging leverarm hypothesis, experimental data on the structure and dynamic behavior of myosin were obtained separately. Detailed
but static structural data have often given the impression that
there is a tight relationship between structural and chemical
states. The single-molecule biophysical studies of the dynamic
behavior of myosin, on the other hand, have revealed a subsetof dynamic molecular events that depends on where theoptical probe is placed, and hence other subsets are missed.
Therefore, constructing models for the myosin motor mechanism on this basis is reminiscent of a jigsaw puzzle withmissing pieces that have to be filled by speculation andhypotheses. Directly observing the structural dynamics anddynamic processes of M5-HMM by HS-AFM has significantly reduced the missing parts and revealed a picture that appears similar to but is largely different from that envisaged
previously. Surprisingly, it revealed that generation of tension
responsible for forward movement and the following leverarm swing can occur without through the ADP–Pi bound state
and therefore the mechanical events do not require the input ofchemical energy. Thus, the chemical energy of ATP is mainly
used for the detachment of the trailing head from actin. Once
detached, the forward swing of the leading head and theresulting forward step of the molecule occur spontaneously
without chemical energy input. Of course, this issue of energy
usage by M5 should be further corroborated in more direct
ways. Moreover, this new principle of energy usage needs tobe confirmed for other myosin motors as well as for othermechano-enzymes. This important task remains to beaccomplished.

Acknowledgments We thank Prof. Cristobal dos Remedios for reading
our manuscript and giving valuable comments. This work was supported
by JST/CREST (to TA), JST/PRESTO (to NK), and KAKENHI from the
Ministry of Education, Culture, Sports, Science and Technology, Japan(#20221006 and #24227005 to TA).
Compliance with Ethical Guidelines The authors declare that ourexperiments described in this manuscript comply with the current lawsof Japan where they were performed.
Conflict of interest Noriyuki Kodera and Toshio Ando declare thatthey have no conflict of interest.
Human and Animal Studies This article does not contain any studies
with human or animal subjects performed by the any of the authors.
Open Access This article is distributed under the terms of the Creative
Commons Attribution License which permits any use, distribution, and255
reproduction in any medium, provided the original author(s) and thesource are credited.

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