FEBS Letters 587 (2013) 997–1007

journal homepage: www.FEBSLetters.org

Review

Molecular machines directly observed by high-speed atomic force
microscopy
Toshio Ando
Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japana r t i c l e

i n f oArticle history:
Received 13 November 2012
Revised 15 December 2012
Accepted 17 December 2012
Available online 11 January 2013
Edited by Wilhelm Just
Keywords:
High-speed AFM
Structure dynamic
Dynamic process
Bioimaging
Atomic force microscopy
Molecular machinea b s t r a c t
Molecular machines made of proteins are highly dynamic and carry out sophisticated biological
functions. The direct and dynamic high-resolution visualization of molecular machines in action
is considered to be the most straightforward approach to understanding how they function but thishas long been infeasible until recently. High-speed atomic force microscopy has recently been realized, making such visualization possible. The captured images of myosin V, F1-ATPase, and bacteriorhodopsin have enabled their dynamic processes and structure dynamics to be revealed in great
detail, giving unique and deep insights into their functional mechanisms.
Ó 2013 Federation of European Biochemical Societies. Published by Elsevier B.V. All rights reserved.

1. Introduction
The vital activities of cells depend on the activities of molecularmachines made of proteins with various functions. Various approaches have been taken to examine their functional mechanism.
The most frequently taken approach has been the structural analysis of proteins by X-ray crystallography, NMR, and electron microscopy. This is because the function of proteins is closely related totheir structure. More than 60,000 proteins have been clariﬁed interms of the detailed three-dimensional structure. The detailed
structure is useful in revealing local interactions between aminoacid residues but is often less useful in revealing the global interdomain and interprotein communication which plays an essential rolein the operation of molecular machines. Moreover, the obtained
structures are limited to static snapshots, while proteins in generaland molecular machines in particular are highly dynamic in nature.
To access the dynamics of proteins in action, single-molecule ﬂuorescence microscopy [1,2] and optical trap nanometry [3,4] havebeen developed and used. In single-molecule ﬂuorescence microscopy, individual protein molecules are labeled with ﬂuorophores
at a speciﬁc locus of the protein and observed under a ﬂuorescencemicroscope. Although a subset of molecular events can be indirectly
revealed for the labeled portion of the molecule, other portions arecompletely invisible. In optical trap nanometry, mechanical eventsoccurring in a molecule (displacement or force production) are
E-mail address: tando@staff.kanazawa-u.ac.jp

measured by optical detection of the displacement of a microsphereattached to the molecule or its interacting partner. Thus, most, if
not all, of the structural information is omitted in these frequently
performed measurements.
Simultaneously and directly observing both the structure anddynamics of molecular machines in action is considered to be themost straightforward approach to understanding how they operate. To this end, Ando and colleagues embarked on the development of high-speed atomic force microscopy (HS-AFM) two
decades ago. AFM is a unique imaging tool that enables the highresolution visualization of objects made of any material in any
environment (vacuum, air, and liquids) [5]. However, it takes atleast 30 s (usually minutes) to capture an image, and therefore,
moving objects cannot be imaged or can only be blurrily imaged.
Through the creation of new devices and techniques and theimprovement of the ﬁrst version of our instrument [6], HS-AFM
has now been realized [7,8] and can capture images every 40–
100 ms. Importantly, the function of fragile molecules is not disturbed by the interaction with a cantilever tip. This performance
has made it possible to visualize, for example, myosin V walking
along an actin ﬁlament [9], bacteriorhodopsin responding to light[10,11], and rotorless F1-ATPae with rotationally propagating conformational changes [12]. Unlike single-molecule ﬂuorescencemicroscopy, various dynamic events of molecules appear in an
AFM movie without their preplanned selection. Therefore, we canobserve several facets of molecular action in detail in a single
imaging experiment. Moreover, the visualized molecular action

0014-5793/$36.00 Ó 2013 Federation of European Biochemical Societies. Published by Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.febslet.2012.12.024

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T. Ando / FEBS Letters 587 (2013) 997–1007

that appears on the video can often be interpreted straightforward
without intricate analyses and interpretations, enabling convincing
conclusions to be reached. These excellent general features of
HS-AFM imaging greatly facilitate and accelerate our understanding of the functional mechanisms of molecular machines. However,a drawback of HS-AFM imaging is that the sample preparation for
HS-AFM imaging is not as easy as that for ﬂuorescence microscopy
imaging.
This review ﬁrst describes the principle of AFM, which isfollowed by an outline of the HS-AFM instrumentation as well as
its speed and low-invasiveness performance. Then, imaging studies
carried out by my group on three different molecular machines arepresented. Finally, a brief perspective is given on upcoming biological research that will be opened up by high-speed AFM.
2. Principle of AFM
Among a number of operation modes, HS-AFM employs the tapping mode [13]. In this mode, the cantilever is oscillated at its ﬁrst
resonant frequency fc. A stylus probe attached to the free end of thecantilever is intermittently brought into contact with the sample.
The mechanical response of the cantilever upon this contact(change in amplitude) is measured and then the detected amplitude is compared with a preset amplitude value (i.e., set point
amplitude). The difference between them (error signal) is input
to a proportional-integral-derivative (PID) controller, whose output is sent to a voltage ampliﬁer (piezodriver). The sample stage,
which is attached to a piezoactuator-based z-scanner, is ﬁnally
moved in the z-direction by the piezodriver output so that the error signal approaches zero. (see Fig. 1). This series of operations isrepeated many times at different points of the sample surface during lateral scanning of the sample stage. Consequently, the movement of the sample stage traces the sample surface, and therefore,
the feedback controller output at each xy position represents the
sample height at that position. Thus, the sample surface topography is constructed from the recorded feedback controller output.
In the tapping mode, the cantilever tip is vertically oscillated.
Hardly any lateral force is exerted between the tip and the sample,
provided that the lateral displacement of the sample stage is verysmall (0.1 nm) for the period of tip-sample contact. As cantilevers
are much stiffer in the lateral direction than in the vertical direction, this nearly zero load is advantageous for imaging biologicalsamples. For protein molecules, the resolution is generally 2–
3 nm in the lateral direction and 0.1–0.2 nm in the vertical direction, although depending on the sharpness of the tip. AFM imaging
relies on sequential height measurement at many different points

on the sample surface. Moreover, because AFM contains slow
mechanical devices (a cantilever and a scanner), the temporal
resolution of conventional AFM is usually on the order of minutes.
3. High-speed AFM
High-speed AFM was realized by the enhancement of the
response speed of all the devices contained in the closed feedback
loop, particularly of the cantilever and scanner, as well as bypassive and active damping of the scanner vibration. Details of
the instrumental development are described elsewhere [7]. Here,the speed performance of the major devices employed in our
HS-AFM instrument is summarized. The response speed of thecantilever was increased by its size reduction [14,15]. Small cantilevers with a highest response speed are 6 lm long, 2 lm wide,and 90 nm thick (BL-AC7DS-KU3, custom made by Olympus) andhave the following mechanical properties: fc 1.2 MHz in water(3.5 MHz in air), quality factor Qc 2 in water, and spring constant kc 0.2 N/m. Therefore, the time constant Qc/(pfc) with which
the oscillating cantilever responds to tip-sample contact is 0.53 ls.
The cantilever oscillation amplitude is detected by a fast detector
every half or one cycle of oscillation (detection time, 0.42–
0.83 ls) [6,7]. The fastest piezoactuator-based z-scanner, which
is displaceable up to 800 nm at 100 V and controlled by an active
damping technique [16], responds within 0.36 ls when driven at
100 kHz. As a result of these shortened response times andthe compensation for the time delay by the (P + D) operation of
the PID feedback controller, the overall feedback bandwidth fB
has now reached 110 kHz.
4. Highest possible imaging rate
The feedback bandwidth is deﬁned by the z-scanner displacement (i.e., feedback) frequency fB at which a phase delay of p/4
or a time delay of 1/(8fB) occurs when tracing the sample surface.
Because of this delay, the cantilever oscillation amplitude A cannot
be exactly maintained at its set point amplitude As, which results ina larger tip-tapping force (and A < As) in uphill regions of the sample and a smaller tip-tapping force (and A > As) in downhill regions[17,18]. When the tip-tapping force applied to the sample is excessive, the scan velocity Vs in the x-direction has to be reduced sothat the phase delay h becomes smaller than p/4.
The highest possible imaging rate (i.e., highest possible framerate) Rmax depends on the feedback bandwidth fB as well as on
the imaging conditions, the spatial frequency of the corrugationon the sample surface (1/k) to be imaged, and the sample fragility.

Fig. 1. Schematic of tapping-mode AFM setup (see text).

T. Ando / FEBS Letters 587 (2013) 997–1007

The imaging conditions are given by the scan range in the x-direction W and the number of scan lines N. When the x-scanner is displaced at velocity Vs, the z-scanner is driven at frequency f = Vs/k.
Thus, the imaging rate is R = Vs/(2WN) = kf/(2WN). When the sample is sufﬁciently strong for withstanding the excessive tip force
due to the p/4 phase delay, the feedback frequency f can be increased up to fB. In this case, we obtain Rmax = kfB/(2NW). In reality,
fragile biological molecules cannot withstand a large tip force.
Denoting the maximum phase delay allowable for a fragile molecule as hmax, the highest possible imaging rate is given by Rmax = 2
kfBhmax/(pNW).
According to our experiences of HS-AFM imaging of protein
molecular machines such as F1-ATPase and myosin V, the maximum possible phase delay is approximately p/9 for a cantileverfree oscillation amplitude of A0 = 1–2 nm and a set point amplitudeof As = 0.9  A0. For example, for k = 5 nm, N = 100, andW = 100 nm, Rmax is 12 fps.
5. Low invasiveness
Under the conditions of A0 = 1–2 nm, As = 0.9  A0, and h = p/9,
the maximum tapping force is estimated to be about 100 pN for
typical proteins with height h0 5 nm when small cantilevers withspring constant kc = 0.2 N/m are used. This force sounds large
because motor proteins generate a force of only a few pN. However, it should be noted that the mechanical quantity that affects
the sample (i.e., causes a change in momentum) is not the force itself but the impulsive force (the product of the force acting on thesample and the time during which the force acts). The time during
which force acts is short (100 ns or less) for small cantilevers
with a resonant frequency of 1.2 MHz in water. Thus, the magnitude of the impulsive force is small.
The effect of tip tapping on the sample can also be estimatedfrom the energy loss of an oscillating cantilever. When a cantileverwith kc = 0.2 N/m is oscillated at A0 = 1 nm, its mechanical energy
is only 24 kBT (similar to the energy of ATP hydrolysis,
20 kBT), where kB is Boltzmann’s constant and T is 300 K. Even
when all of this energy is lost to excite the sample, the energy will
be distributed into many degrees of freedom within the tapped
portion of the sample and will quickly dissipate into more degrees
of freedom including those of the surrounding water molecules(see below). In actual imaging experiments, such events ofcomplete energy loss happen only very rarely, if at all, and the cantilever oscillation amplitude is generally maintained at 0.9  A0 =0.9–1.8 nm. On average, the energy loss after every tapping is verysmall: ½kc(A02  As2)/Qc = 2.39 kBT.
A tip oscillating at a high fc in water iteratively taps the sample.
When an image with an area of Wx  Wy nm2 is captured at an
imaging rate R, a molecule with a top surface area (Atop nm2) perpendicular to the tip is tapped fc/R  Atop/(WxWy) times per frame.
For example, under the conditions (fc 1 MHz, R = 12.5 fps, andWx  Wy = 45  22 nm2) used for the non-invasive imaging ofrotorless F1-ATPase (i.e., without the c subunit) [12], one moleculeof rotorless F1-ATPase (Atop 100 nm2) is tapped 8,000 times perframe and 4 million times in total during the 40 s successive
imaging. Nonetheless, the rotorless F1-ATPase is fully functional
throughout the imaging, as shown later. This clearly indicates thatthe energy transferred from the oscillating tip to the sample does
not accumulate in the sample but dissipates quickly (much fasterthan the oscillation period of the cantilever of 1 ls).
6. HS-AFM imaging of molecular machines
Although only a few years have passed since the establishmentof HS-AFM instrumentation in 2008 [7], a number of HS-AFM

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imaging studies have already been conducted on various protein
systems. However, the number of studies that have revealed thestructure dynamics of molecular machines in action is limited:
myosin V walking on an actin ﬁlament [9], rotorless F1-ATPasewith rotationally propagating conformational changes [12], lightdriven proton pump bacteriorhodopsin responding to light[10,11], and trimeric P2X4 receptor with a dilating central pore in
ATP [19]. Other studies have focused on diffusion processes [20–23], dynamic interactions between molecules [24–28], self-assembly processes [29,30], enzymatic-reaction-coupled molecular processes [18,31–36], and dynamics occurring with DNA origamis[37–39]. In most of these studies, the structure of the molecules involved in the dynamic processes was not well resolved. Nonetheless, important results inaccessible with other approaches havebeen obtained.
For example, in the observation performed by Igarashi and collaborators on the hydrolysis of cellulose ﬁbers by cellulase [31,32],
unidirectional processive movement of the cellulase molecules
was demonstrated. Moreover, the retardation of the enzymatic
reaction with time, which has been previously observed in biochemical assays, was revealed to arise from trafﬁc congestion ofmoving molecules [32]. In the observation performed by Scheuring
and colleagues on the dynamic association and dissociation between membrane-embedded ATP-synthase c-rings with 6.5 nm
diameter [24], their association was revealed to be driven by the‘‘hydrophobic mismatch’’ [40] between the hydrophobic thicknessof the lipid bilayer and the length of the hydrophobic region of themembrane-embedded c-rings. Moreover, from the distribution of
the center-to-center distance of a c-ring dimer, the potential
energy distribution was obtained, which showed that an energy
minimum of 3.5 kBT occurred at a center-to-center distance of
10.3 nm (T = 300 K).
Below, imaging studies conducted by my group on the structure
dynamics of molecular machines in action are described in detail.
6.1. Walking myosin V
Double-headed myosin V (M5), which functions as a cargotransporter in cells, has been extensively studied (see review[41]). It moves processively along an actin ﬁlament [42,43] towards the plus end of the ﬁlament in a hand-over-hand manner,
advancing 36 nm [44–46] in every ATP hydrolysis cycle [47].
‘‘Hand-over-hand’’ means that the two heads step alternately,exchanging leading and trailing roles at each step, very much like‘‘walking’’. In spite of this well-established understanding of how
M5 steps, here are still fundamental questions to answer. How is
the chemical energy liberated by ATP hydrolysis used to generate
the forward movement? How is the powerstroke (lever-arm
swing) coupled with the ATPase cycle, if at all? If it really is coupled, which chemical transitions are involved in the generationof tension and lever-arm swing? These fundamental questions
have long been explored in muscle contraction studies and a consensus is thought to have already been reached for some of them.
Are there any inconsistencies when the consensus view is applied
to the motility of M5? Previous single-molecule measurements of
M5 stepping do not seem to have answered these questions clearly.
Therefore, we performed imaging studies on M5 to demonstrate
the power of HS-AFM [9]. Its imaging results were not only consistent with the well-established facts mentioned above but also
solved some of the fundamental questions that had previously
been difﬁcult to address.
6.1.1. Step motion of myosin V
Partially biotinylated actin ﬁlaments were immobilized on amica-supported phospholipid bilayer surface containing biotin-lipid through streptavidin with a low surface density. The positively

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T. Ando / FEBS Letters 587 (2013) 997–1007

charged lipid 1,2-dipalmitoyl-3-trimethylammonium-propane(DPTAP, 5%) was included in this lipid bilayer to facilitate observing
the characteristic sideways ﬁgure of actin-bound M5-HMM (tailtruncated M5). M5-HMM moving processively with 36 nm steps
was clearly visualized at 7 fps (Fig. 2A). As illustrated in Fig. 2B, thetwo-headed bound M5-HMM exhibited distinct structural featuresthat are consistent with previous electron microscopy observations[48,49]; the neck–motor domain junction appears smooth in theleading head but is V-shaped in the trailing head without exception, 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 features can be used to determine thepolarity of actin when the bound M5-HMM is stationary. WhenDPTAP was absent in the lipid bilayer, the average translocation
velocity was similar to that measured by ﬂuorescence microscopy
under the same buffer solution condition, indicating no effect of
the tip–sample interaction on the motor activity. However, we
could not observe the molecular behavior during a step in detailas it was completed within a frame. To slow down the step, more
streptavidin molecules were placed on the substrate surface as

moderate obstacles to the advance. This method allowed the visualization of the stepping process (Fig. 2C and D).
After trailing head detachment, the leading head appeared tospontaneously rotate from the reverse arrowhead orientation towards the arrowhead orientation (the term ‘‘arrowhead’’ originatesfrom the conﬁguration of single-headed myosin bound to an actin
ﬁlament in the rigor state). Before completing this rotation, theleading head was brieﬂy halted by colliding with a streptavidin
molecule in its natural path. In this halted state, the detached trailing head was at its furthest from the actin ﬁlament (thus, the twoheads were aligned nearly straight, pointing in opposite directions)and slightly rotated around the neck-neck junction (second framein Fig. 2C). Then, the leading head overcame the streptavidin blockade and rotated to take the arrowhead orientation. Accompaniedby this further rotation, the trailing head bound to a forward siteof the actin ﬁlament to become the leading head, completing one
step. Here, it was clearly revealed that during and before the completion of a step, the trailing head never interacts with actin andhence passively moves forwards, driven by the rotating leading
head. Thus, the ‘‘inchworm’’ mechanism (a molecule moves for-

Fig. 2. Walking M5-HMM captured by HS-AFM. (A) Successive AFM images showing processive movement of M5-HMM in 1 lM ATP. Frame rate, 7 fps. (B) Schematic of twoheaded bound M5-HMM. (C) Successive AFM images showing hand-over-hand movement in 1 lM ATP. Frame rate, 7 fps. The swinging lever is highlighted with a thin whiteline. (D) Schematic explanation of the images in C. The ‘‘+’’ marks indicate the plus ends of actin ﬁlaments. (E) A model of mechanochemical coupling in actin–myosininteraction proposed on the basis of the swinging lever-arm hypothesis for muscle contraction.

T. Ando / FEBS Letters 587 (2013) 997–1007

ward like an inchworm, in which one head always leads while theother head follows) considered for kinesin [50] as well as the‘‘biased diffusion’’ mechanism (a molecule moves forward bydirectional diffusion according to the hypothetical potential slope
created along the actin helix) proposed for M5 [51] can be ruledout. The rotation of the leading head is the swinging lever-arm motion itself initially proposed by Hugh Huxley for the powerstroke ofmuscle myosin [52].
6.1.2. Generation of tension
The seemingly spontaneous rotation of the leading head following trailing head detachment suggests that the intramolecular tension required for the advance already existed in the two-headed
bound molecule. In this bound state, the trailing head is in thearrowhead orientation which is the natural orientation, at least
for the ADP-bound or nucleotide-free head. Hence, the intramolecular tension is not produced by the bound trailing head. On theother hand, the leading head is not in the natural orientation(i.e., reverse arrowhead orientation) and hence pays an energy costto generate the intramolecular tension. In fact, the bound leadinghead is slightly curved outwards, while the trailing head is straight.
Upon trailing head detachment, the constraint that keeps thebound leading head in the unnatural orientation is removed, andhence the leading head spontaneously rotates forwards, meaningthat the bound leading head is in a strained prestroke state andthat the lever arm swing is not accompanied by a chemical transition at the leading head. This conclusion was reinforced by theobservation that the short coiled-coil tail of two-headed bound
M5-HMM sometimes unfolded and then the leading head rotatedfrom the reverse arrowhead orientation to the arrowhead orientation, which was very similar to the swinging lever arm motion observed during a step. The uncoiling is caused by intramoleculartension, not by the interaction with the oscillating cantilever tip.
This is suggested by the fact that some single-headed species arealso contained in the sample of M5-HMM puriﬁed through actin
binding and dissociation [53].
In general, during the actomyosin ATPase cycle, the strained
prestroke state has been thought to always be formed through Pi
release from an ADP–Pi-bound head weakly interacting with actin[54,55]. Thus, it may be considered that the strained prestroke
state, which is formed directly by the binding of the ADP-boundleading head to actin, does not participate in the forward step inATP. However, we observed molecular behavior in ATP indicatingthat it does participate in the step. In two-headed bound M5HMM, both of the motor domains frequently exhibited brief dissociation and reassociation on the same actin ﬁlament (or a brieftranslocation of 5 nm along the actin ﬁlament), whereas M5HMM remained at approximately the same position on the ﬁlament. We have termed this behavior ‘‘foot stomp’’. Foot stomps
were more frequently observed at the leading head than at thetrailing head. The brieﬂy detached leading head does not carrybound Pi because Pi was released from an ADP–Pi-bound M5 head
when it initially bound to actin. This rapid release of Pi is a featureof the actin-activated M5 ATPase reaction [56]. Nevertheless, thedetached leading head with only ADP bound rebinds to actin, stillin the reverse arrowhead orientation, and then swings forward following trailing head detachment. The detachment of the leadinghead during a foot stomp is not caused by ATP binding to the leading head. As described in the next section, a nucleotide-free leadinghead is often sharply bent. However, we never observed a sharply
bent leading head before a foot stomp.
6.1.3. Mechanism of hand-over-hand movement
The leading head of two-headed bound M5-HMM is straight(slightly curved outwards) in ADP or ATP, while it is often sharply
bent under the nucleotide-free condition. Therefore, simply by1001

looking at the conformation of the leading head, we can judge
whether or not the leading head contains nucleotides. To estimate
the ADP dissociation rate constant at the leading head, M5-HMM
bound to actin at various concentrations of ADP was observed,
and then the lifetime of the straight leading head was analyzed asa function of ADP concentration. The ADP dissociation rate constantat the leading head was estimated to be 0.1/s. This means that ADP
is released from the leading head every 10 s on average. However,
M5-HMM walks many steps in 10 s, meaning that ADP does not dissociate from the leading head. That is, before ADP dissociation fromthe leading head, the leading head is switched to the trailing head
during walking. ADP dissociation, the subsequent ATP binding,and the resulting detachment from actin solely occur at the trailinghead. This is the basis underlying the processive hand-over-handmovement. This mechanism was previously inferred from various
indirect experiments [47,57–61] but has now been clearly anddirectly demonstrated by high-speed AFM observation.
6.1.4. Newly raised questions
The swinging lever-arm hypothesis is no longer a hypothesis.
HS-AFM visualized its occurrence with irrefutable clarity. However, our HS-AFM observation results raise question on the mechanochemical coupling mechanism which has been modeled in thecontext of the swinging lever-arm hypothesis for muscle contraction (Fig. 2E). In this model, the structural conﬁgurations are considered to be tightly coupled to the nucleotide states of the motor
domain. The M–ADP–Pi state (M denotes the myosin head) is considered to play an essential role in head binding to actin. In thisstate, the head takes a prepowerstroke conformation resulting
from the recovery stroke of the neck domain, which facilitates its
binding to actin in the reverse arrowhead orientation. Coupled tothe dissociation of Pi after binding to actin, the myosin is considered to swing its neck only once from the reverse arrowhead orientation to the arrowhead orientation and ﬁnally take thepostpowerstroke conformation. Thus, in this model, the myosin
head must pass through the chemical state of M–ADP–Pi to generate a force and move forward. However, the observation by HSAFM of walking M5-HMM shows that the lever-arm swing spontaneously occurs after trailing-head detachment from actin. Moreover, the tension responsible for the lever-arm swing can begenerated by the ADP-bound leading head (after foot stomping),
without directly passing through an ADP–Pi-bound chemical state.
In addition, two-headed bound M5-HMM in ADP can generate sufﬁcient tension to cause the occasional unwinding of the shortcoiled-coil tail (i.e., force generation occurs spontaneously). Therefore, the recovery stroke or the prepowerstroke conformation,
which has been considered to occur uniquely in the ADP–Pi-boundstate, does not appear to be necessary for the lever-arm to swing.
Thus, the chemical energy liberated by ATP hydrolysis does not appear to be used for the recovery stroke, tension generation and thelever-arm swing. M5 can step forward once the trailing head detaches from actin without any chemical transitions.
We would, however, inevitably encounter a ‘‘perpetuum mobile
problem’’ if we supposed that neither tension generation nor leverarm swing require any energy input. Even in ADP or in the nucleotide-free condition, the foot stomp occurs at both the leading andtrailing heads of two-headed bound M5-HMM, with more frequent
occurrence at the leading head than at the trailing head [9]. Whenthe 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 moleculewould not take a backward step because the trailing head is boundto actin in a stable orientation (i.e., the arrowhead orientation).
Thus, without any energy input, the M5-HMM molecule can beconsidered to step forward many times (albeit slowly), even withthe less frequent occurrence of foot stomps at the trailing head.

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This is forbidden perpetuum motion. However, in this gedankenexperiment, the ﬂexibility of the neck-motor domain junction or
positional ﬂuctuations of the molecule bound to actin via a single
head are not considered. Taking these effects into account, we
can expect the occurrence of a backward step after leading headdetachment from actin, although its probability is low. Thus, theprinciple of detailed balance can hold between the forward stepafter trailing head detachment and the backward step after leadinghead detachment, resulting in no net movement of M5 when noenergy is input. For example, in the presence of 1 mM ADP, the proportion of foot stomp frequency observed at the leading head wasabout 0.8, while it was 0.2 at the trailing head. This ratio of 4:1 appears to be insufﬁcient for the detailed balance. However, the footstomps occur very fast compared with the imaging rate used
(7 fps) and the bound M5-HMM almost always stays at the same
position after foot stomping at the leading head. The observed ratioof 4:1 is therefore likely to be an underestimate.
From numerous X-ray crystallography and electron microscopy
studies, there is no doubt that myosin changes its conformation
upon binding to nucleotides and takes a prepowerstroke-like conformation in the ADP–Pi-bound state [49,62–67]. As mentioned
above, this prepowerstroke-like conformation certainly facilitates
the binding of the head (the leading head in the case of M5) to actinin the reverse arrowhead orientation. However, in the conventionalview of the mechanochemical coupling of myosin (Fig. 2E), the conformations of myosin have long been considered to be inﬂexible.
This is mainly due to the fact that the static three-dimensional
structures of myosin have often been used to construct models for
its mechanochemical coupling. In reality, they are ﬂexible. Otherwise, M5 could not bind to an actin ﬁlament via both heads underthe ADP-bound and nucleotide-free conditions. However, it can doso even in the absence of nucleotides, which is consistent with theresults of other studies [48,57]. Thus, myosin takes two (or multiple)
conformations even under a given nucleotide condition (or a given
biochemical state). Myosin can go back and forth between different
conformations but its dynamic equilibrium shifts to one side
depending on the nucleotide condition. Our HS-AFM observationsof M5-HMM indicated that structural ﬂuctuations indeed exist
and enable the myosin head to take the prepowerstroke conﬁguration even in the ADP-bound and nucleotide-free states. In addition,
it was also indicated that the degree of ﬂuctuation depends on the
nucleotide state since the leading head of M5-HMM was observed
to bind to actin most easily in ATP, next in ADP, and then in the absence of nucleotides. Recently, it has also become possible to study
the ﬂexible structures of myosin with the use of spectroscopy andsite-directed labeling with ﬂuorescent and spin probes (see review[68]). For example, when a spin probe is attached to the regulatory
light chain of myosin in muscle ﬁbers, the spin probe angle relative
to the ﬁber axis was observed to be distributed widely around two
peaks with nearly equal populations in ATP (in contracting and relaxed ﬁbers), while in the rigor state it was distributed around
two peaks with markedly different populations [69].
The ﬂexibility of the myosin head conformation raises the question of whether the chemical energy of ATP hydrolysis is reallyused to change the conformation. In the conventional view, themyosin head with a strained conformation in the prepowerstroke
state must retain energy that is then liberated during powerstroke.
However, the prepowerstroke and postpowerstroke conformations
are in dynamic equilibrium without markedly different populations. Therefore, the energy difference between the two conformations is very likely to be less than 3 kBT (T = 300 K). Thus, we canconclude that although the conformational change of the myosin
head caused by the recovery stroke is an effective strategy for facilitating its binding to actin in the reverse arrowhead orientation,
the energy required to produce the prepowerstroke conformation
is much less than that liberated by ATP hydrolysis.

Let us perform another gedankenexperiment. Each head of M5
is assumed to be labeled with a chromophore. Upon light absorption, the head-bound chromophore is assumed to alter the conformation of the head so that the head dissociates from actin. When alight pulse only illuminates the trailing head of an M5 molecule,the molecule spontaneously steps forward after trailing headdetachment. By repeating this directed light illumination only at
the trailing head, we can move the molecule for a long distancein one direction.
Therefore, it appears plausible that a large fraction of the chemical energy liberated by ATP hydrolysis is used elsewhere and islikely to be used to detach a nucleotide-free bound myosin headfrom actin. How much energy is required for this detachment?
The unbinding (rupture) force was previously measured to be15 pN by AFM [70]. The rupture distance was also estimated tobe 1.7–2.5 nm in this study. This rupture distance is unusually longcompared with 0.23 nm for an antigen–antibody [71], 0.15–0.3 nm
for streptavidin–biotin [72], and 0.05–0.3 nm for a-actinin–actin[73]. This long rupture distance is consistent with the fact that actin–myosin interfaces contain bonds formed between ﬂexible loops[74,75]. From the values of rupture force and rupture distance, theenergy required to rupture the actin–myosin rigor bond is 6–9 kBT,
which is 30–45% of the chemical energy of ATP hydrolysis (20
kBT). This mechanically estimated value is approximately consistent with the value of 9 kBT estimated biochemically for the free-energy change in the dissociation A–M ? A + M [76], where A denotes
actin. For the ATP-induced dissociation A–M + T ? A + M–T (T denotes ATP), the free-energy change is biochemically estimated tobe 10 kBT, which is similar to that for the mechanical dissociation
A–M ? A + M. However, one may argue as follows against the statement that a large fraction of the chemical energy liberated by ATPhydrolysis is used to detach a nucleotide-free bound myosin headfrom actin; ATP is not yet hydrolyzed in the actin–myosin dissociation step; thus, the energy of ATP hydrolysis is not yet liberated.
However, since the account balance of energy should be considered
for the whole cycle of the ATPase reaction, the ATP-binding energy
can be used as part of the energy of ATP hydrolysis.
6.2. Rotary catalysis of rotorless F1-ATPase
The a3b3c complex, which is the minimum unit of F1-ATPaseand retains the full ATPase activity, is a rotary motor (Fig. 3A)[77–79]. The coiled-coil part of the rotary shaft c is inserted in
the central cavity formed by a stator ring a3b3 where three a andthree b subunits are arranged alternately [80]. Three ATP bindingsites are located at the a–b interfaces, mainly in the b subunits.
The rotation of c occurs in the counterclockwise direction as
viewed from the exposed globular portion of c (or from the C-terminal side of a3b3). In the ATPase cycle, three b subunits take different chemical states: ATP-bound, ADP-bound, and nucleotidefree (empty) states [77,80]. Each chemical state cyclically propagates over the three b subunits. Thus, there is strong cooperativity
between the three b subunits. How is this cooperativity engendered? There is no direct contact between the b subunits and theb-c interaction is always different for the three b subunits because
c has no symmetry. Considering these facts, it was proposed thatinteractions with c control and determine the conformationaland catalytic states of the individual b subunits [81]. This idea
was reinforced by studies showing that the backward mechanicalrotation of c with an external force reverses the chemical reaction
toward ATP synthesis [82,83], whereas forced forward rotation results in accelerated ATP binding [84]. This view was, however,challenged by the ﬁnding that even when c is shortened so thatmost c–b interaction sites are removed, the short c still rotates
unidirectionally [85]. However, using single-molecule optical
microscopy, there was no means of observing the rotation after

T. Ando / FEBS Letters 587 (2013) 997–1007

1003

Fig. 3. AFM images of a3b3 subcomplex at C-terminal surface. (A) Schematic of F1-ATPase. (B) Averaged AFM image obtained under nucleotide-free condition. (C) C-terminal
surface of crystal structure of nucleotide-free a3b3 subcomplex. (D) Simulated AFM image of (C). (E) Averaged AFM image obtained in 1 mM AMP–PNP. (F) C-terminal surfaceof crystal structure of a3b3 subcomplex obtained in ATP. (G) Simulated AFM image of (F). (H) Successive images of C-terminal surface of a3b3 subcomplex in 2 lM ATP,
captured by HS-AFM at 12.5 fps. The highest pixel position in each image is marked with a red circle.

the complete removal of c. Thus, we exploited the power of
HS-AFM to resolve this issue [12].
a3b3 with Lys7-tags at the N-termini of the b subunits was covalently immobilized on an amino-reactive mica surface. An image of
the C-terminal side of a3b3 without nucleotides shows a pseudo
sixfold symmetric ring (Fig. 3B). Each subunit has an upwardly protruding portion at the inner top side of the ring but three of these
portions at alternate positions are higher. By the comparison of thisimage with a simulated AFM image (Fig. 3D) constructed from thecrystal structure of nucleotide-free a3b3 (PDB code: 1SKY) [86](Fig. 3C), the three subunits showing higher protrusions were identiﬁed as the b subunits. In 1 mM AMP–PNP, the shape of a3b3 became triangular and the central hole became obscure (Fig. 3E).
Only one subunit had a higher protrusion together with an outwardly extended distal portion, which was identiﬁed to be an
empty b by the comparison of the AFM image with a simulatedone (Fig. 3G) constructed from the crystal structure of a nucleotide-bound a3b3c subcomplex (PDB code: 1BMF) [80] from whichc was removed (Fig. 3F). In addition, the two subunits with retracted distal portions were identiﬁed to be nucleotide-bound b
subunits.
When imaged in 2–4 lM ATP at 12.5 fps, distinct conformational dynamics occurred at the b subunits (Fig. 3H). Each b exhibited a conformational transition between the outwardly extended
high state (open (O) state) and the retracted low state (closed (C)state). The following prominent features were observed: (i) only
one b assumes the O-state, and (ii) when the O-to-C transition occurs at one b, the opposite C-to-O transition occurs simultaneously
at its counterclockwise neighbor b in most cases. Thus, the O conformation propagates counterclockwise (see the highest pixels,
each marked with a red circle in Fig. 3H). The ATP hydrolysis rates
at 2, 3, and 4 lM ATP were approximately one-third of the rates ofconformational propagation at 2, 3, and 4 lM ATP, respectively.
Thus, the conformational propagation is tightly coupled with theATPase reaction. The cumulative rotation angle of the highest pixel
with discrete 120° steps increases linearly with time until the endof imaging (40 s in total). Thus, the function of a3b3 is completely
retained throughout the imaging. The a3b3 molecule is tapped withthe oscillating tip 4 million times in total during this successive
imaging.
These dynamic transitions indicate that the O-to-C transition
occurs when ATP is bound to an empty b, while the C-to-O transition occurs when an ADP-bound b releases ADP. Thus, the empty,
ADP-bound, and ATP-bound b subunits are arranged counterclockwise in this order, and therefore, the observed conformational
propagation demonstrates rotary catalysis by the a3b3 subcomplex. Thus, we reach the most important conclusion that the intrinsic cooperativity responsible for torque generation to rotate c iselicited through the b–b interplay alone and c is passively subjected to torque causing it to rotate [12].

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T. Ando / FEBS Letters 587 (2013) 997–1007

6.3. Bacteriorhodopsin responding to light
A membrane protein, bacteriorhodopsin (bR), is a light-driven
proton pump that transfers protons across the membrane fromthe cytoplasmic side to the extracellular side [87,88]. bR comprises
seven transmembrane a-helices (named A–G) surrounding the retinal chromophore covalently bound to Lys216 via a protonated
Schiff base [89,90]. In the native membrane (called the purple
membrane), bR assembles into trimers, which are arranged in ahexagonal lattice. bR has been extensively studied by optical spectroscopy techniques. Upon the absorption of light, photoisomerization from the all-trans conformation to the 13-cis conformation of
the retinal chromophore takes place. This isomerization induces
proton transfer from the Schiff base to Asp85, which triggers a cascade of changes in the bR structure. A series of intermediates designated as J–O are deﬁned, among which M (M410), which has anabsorption peak at 410 nm and the longest lifetime, is the onlyintermediate containing a deprotonated Schiff base [88]. In contrast to the spectroscopic characterization of bR during the photocycle, the overall dynamic structural change in bR upon
photoactivation has long been elusive despite the establishmentof its detailed static structure in the unphotolyzed state by X-ray
crystallography and electron microscopy [91–93]. Thus, bR appears
to be an ideal target for demonstrating the power of HS-AFM.
However, the photocycle of the wild type bR at neutral pH proceeds very fast (10 ms). To slow down the photocycle, we usedthe D96N bR mutant that has a longer photocycle (10 s at pH7) but still retains the ability of proton pumping [94]. Native purple
membranes containing D96N bR were directly placed on a baremica surface. Fig. 4 shows successive images of the D96N mutant
at the cytoplasmic surface captured at 1 fps [10]. Upon illumination with green light (532 nm, 0.5 lW), bR markedly changed its
structure (compare the images at 1 and 2 s) and returned to theground state within a few seconds when light was discontinued.
This structural change was speciﬁcally induced by green lightand not by blue or red light, which is consistent with the fact thatbR in the ground state effectively absorbs green light. The centersof mass of all activated bR monomers moved outwards by 0.7–
0.8 nm from the center of the respective trimers and rotated counterclockwise by 7–8° around the center of the respective trimers.
The movement visualized here is ascribed to the displacement of
the E–F loop, which was derived by the comparison with the atomic model of the a-helical cytoplasmic ends in the unphotolyzed
state [90]. Note that the overall position of each bR molecule does
not change because of indiscernible alterations at the extracellularsurface. The photoactivated state observed by high-speed AFM wasattributed to the spectroscopically identiﬁed MN intermediate[11,95], which exhibits outward tilt of helix F similar to that in

the N state of the wild type [96], while the Schiff-base is still
deprotonated like the precedent M state because of the absenceof the proton donor residue (D96) for the Schiff base. Thus, thephoto-induced structural change in bR, which has long been studied but remained elusive, is now clearly revealed by its direct
visualization.
As a result of the outward displacement, nearest-neighbor bR
monomers (one set is shown with a light-blue circle in the imageat 1 s in Fig. 4), each belonging to a different adjacent trimer(shown with three white circles), appear to transiently assemble(as indicated with a light-blue circle in the image at 2 s). Here,
we use a new designation, ‘‘trefoil’’, for a triad of nearest-neighbormonomers belonging to different trimers to distinguish it from theoriginal trimer. To investigate the possible effect of the bR–bR
interaction within a trefoil on the photocycle, bR was illuminatedwith green light of different intensities (0.007–0.5 lW). Under
weak illumination, only one monomer in each trefoil was activatedin most cases. When only one monomer was activated in a trefoil,
it decayed with a time constant of 7.3 ± 0.58 s. In contrast, understronger illumination, two or three monomers within a trefoil
tended to be activated together. Interestingly, the decay of eachmonomer markedly depended on the order of its activation. The
monomer that was activated most recently among the activatedmonomers in a trefoil decayed with a shorter time constant of
2.0 ± 0.16 s. On the other hand, the decay kinetics of the monomers
activated earlier did not follow a single exponential curve, and theaverage decay time lengthened to 13 s. Thus, the bR–bR interaction within a trefoil under illumination engenders both positive
and negative cooperative effects in the decay kinetics as the initialbR recovers.
Unlike previous expectations [97,98], these cooperativities do
not occur within a trimer but within a trefoil. This ﬁnding was
made possible by closely looking at individual bR molecules at ahigh spatial resolution in real space and real time. It is difﬁcult
to imagine how these cooperativities could have been found using
techniques other than HS-AFM.
7. Future prospects of HS-AFM studies on molecular machines
7.1. Dynamic imaging of isolated biomolecular systems
Even for biomolecular systems that have already been studied
in detail by other approaches, HS-AFM allows us to carry out moredetailed inquiries or answer questions that have been difﬁcult or
impossible to address by other methods. Thus, HS-AFM provides
the opportunity to make unexpected new ﬁndings. This is indeed
exempliﬁed by our studies on M5-HMM, F1-ATPase, and bR. Importantly, the HS-AFM instrument we have developed is now com-

Fig. 4. Successive images of the cytoplasmic surface of D96N bR under dark and illuminated conditions, captured by HS-AFM at 1 fps. The green bars at 2 and 3 s indicate theillumination of 532 nm green light with an intensity of 0.5 lW. A bR trimer is highlighted with a white triangle in the image at 0 s. In the image at 1 s, three adjacent trimers
are encircled with white circles, while a trefoil comprised of three nearest-neighbor monomers, each belonging to one of the encircled trimers, is indicated with a light-blue
circle. Compare the structures of the same trefoil at 2 and 3 s.

T. Ando / FEBS Letters 587 (2013) 997–1007

mercially available from the Research Institute of Biomolecule
Metrology, Inc. (Tsukuba, Japan). Small cantilevers made of silicon
nitride (9–10 lm long, 2 lm wide, and 130 nm thick; BL-AC10DS,
Olympus, Tokyo) with fc = 1.5 MHz in air and 0.6 MHz in water,
kc = 0.1 N/m, and quality factor Q 2 in water are already commercially available. Smaller cantilevers are also being commercialized
by NanoWorld. Therefore, it is expected that the expanded exploration by dynamic HS-AFM imaging will be carried out for a wide
range of biomolecular systems in the near future. Thus, a comprehensive and deeper understanding of how biological molecularmachines operate is expected to be attained and accelerated.
However, note that the successful visualization of dynamic
events in biomolecular systems often requires much more deﬁned
conditions for the substrate surface than those in static AFM imaging. The substrate surface must have an appropriate afﬁnity for themolecules. To observe dynamic interactions between different species of molecules, the selective attachment of one species of molecules to a surface is required. Otherwise, there is no chance toobserve their interaction. Moreover, the sample on a surface canonly be viewed from the direction perpendicular to the surface.
Therefore, to observe the characteristic ﬁgure of molecules, theysometimes have to be placed in a particular orientation, as in thecase of the observation of M5-HMM molecules walking on an actin
ﬁlament [9]. For the selective attachment of one species of molecules, the following ﬂat surfaces have thus far been characterized
and used: (i) mica-supported phospholipid bilayers containing
functional lipids [9,18] and (ii) two-dimensional crystals of streptavidin formed on biotin-containing lipid bilayers [18,27,99]. To
observe dynamic DNA–protein interactions, DNA origami tiles[35,38] seem useful when they are placed on a surface resistant
to non-speciﬁc binding such as a phospholipid bilayer surface. Various substrate surfaces are expected to be developed along withthe expansion of HS-AFM imaging studies.
7.2. Dynamic in situ imaging of molecules on higher-order structures
The fast scanners that we have developed are optimized for thehigh-speed imaging of isolated biomolecules; their resonant frequencies are very high but the scan range is limited to 1, 4, and1 lm in the x-, y-, and z-directions (at 100 V), respectively. Therefore, large samples such as live bacterial and mammalian cells havenot been able to be imaged by HS-AFM until recently. To overcome
this limitation, we have recently developed a wide-area scannerthat can be displaced by up to 43 lm in the x- and y-directions
and 2.5–4 lm in the z-direction. The resonant frequency of its xand y-scanners is low (1.6 kHz) but can be increased to10 kHz by an inverse compensation-based active damping technique [7]. Using this scanner and another medium-range scanner,whole live bacterial cells have recently been visualized. After taking an image of an entire magnetic bacterial cell, part of its outersurface was imaged fast, enabling the visualization of densely
packed but rapidly diffusing porin molecules [100]. This capacity
will provide new opportunities for the in situ dynamic imaging
of molecular machines on higher-order structures such as live
mitochodoria, nuclei, and neuronal spines.
Acknowledgments
I would like to thank Takayuki Uchihashi, Noriyuki Kodera,
Daisuke Yamamoto, Hayato Yamashita, Mikihiro Shibata, Ryota Iino,
Hiroyuki Noji, and Hideki Kandori for collaborations and discussions
on the topics described in this review. This work was supported by aGrant-in-Aid for Basic Research S (20221006 and 24227005) from
the Ministry of Education, Culture, Sports, Science and Technologyof Japan, by the Knowledge Cluster Initiative Project, and by the
CREST project from the Japan Science and Technology Agency.

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