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High-speed atomic force microscopy coming of age

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2012 Nanotechnology 23 062001(http://iopscience.iop.org/0957-4484/23/6/062001)
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IOP PUBLISHINGNANOTECHNOLOGY

Nanotechnology 23 (2012) 062001 (27pp)

doi:10.1088/0957-4484/23/6/062001

TOPICAL REVIEW

High-speed atomic force microscopy
coming of age
Toshio Ando
Department of Physics and Bio-AFM Frontier Research Center, Kanazawa University, Kakuma-machi,
Kanazawa 920-1192, Japan

Received 18 August 2011, in final form 9 November 2011
Published 17 January 2012
Online at stacks.iop.org/Nano/23/062001
Abstract
High-speed atomic force microscopy (HS-AFM) is now materialized. It allows direct
visualization of dynamic structural changes and dynamic processes of functioning biologicalmolecules in physiological solutions, at high spatiotemporal resolution. Dynamic molecular
events unselectively appear in detail in an AFM movie, facilitating our understanding of how
biological molecules operate to function. This review describes a historical overview oftechnical development towards HS-AFM, summarizes elementary devices and techniques usedin the current HS-AFM, and then highlights recent imaging studies. Finally, future challengesof HS-AFM studies are briefly discussed.
S Online supplementary data available from stacks.iop.org/Nano/23/062001/mmedia(Some figures may appear in colour only in the online journal)

1. Introduction

long-standing problem prevailing protein research and make itpossible to simultaneously record the structure and dynamics
of biological molecules, high-speed AFM has been developedin the last two decades [5, 6] and is now coming of age [7].
After the advent of AFM in 1986 [8], its potential
capabilities as a novel tool in biological research have been
actively explored in pioneering works [9–11]. In life sciences,
AFM is now routinely used as a nanotool for various measurements under physiological solution environments [12, 13];high-resolution imaging of biological samples [14–16],
recognition and localization of specific molecules [17, 18]
as well as force measurements to estimate the strengthof intra- and intermolecular bonds at the single-molecule
level [19–21] and the elasticity of biological surfaces [22, 23].
This remarkable progress has been basically supported
by reinforcement of the initial configuration of an AFM
apparatus with newly developed devices and techniquessuch as microfabricated cantilevers [24], the optical levermethod for cantilever deflection detection [25] and the tapping
mode [26]. The microfabricated cantilevers and the opticallever method simplified the AFM apparatus and its operation(figure 1). In the tapping mode, the cantilever tip is verticallyoscillated at the first resonant frequency of the cantilever

A vast array of studies have long been performed byvarious approaches to understand how proteins work. The
recent technical innovation in protein studies is brought
about by single-molecule fluorescence microscopy createdin the early 1990s [1, 2]. The dynamic behavior ofindividual protein molecules at work can be examined by
the observation of the dynamic behavior of fluorescent spots,
each being emitted from a fluorophore attached to a selectedlocus of the molecule. By this innovation, the depth of
our understanding of how proteins function has become
incomparable with the past. Nevertheless, an insufficiency
in our understanding is obvious. This is mainly because we
still have technical limitations. Protein molecules themselves
are invisible in the fluorescence observations even whensuper-resolution fluorescence microscopy [3] is used. The
structure of biological molecules has been studied using x-ray
crystallography, NMR, electron microscopy and atomic force
microscopy (AFM) but the obtained structures are essentially
static. Thus, the concomitant assessment of structure anddynamics is infeasible. Even the emerging x-ray free-electron
laser [4] will not break this limitation. To overcome this0957-4484/12/062001+27$33.00

1

c 2012 IOP Publishing Ltd Printed in the UK &the USA

Nanotechnology 23 (2012) 062001

Topical Review

Figure 1. Schematic presentation of tapping mode AFM system.

and brought into periodic contact with the sample surface.
Hence, the tapping mode significantly reduces the frictional
force exerting from the cantilever tip to the sample during theraster scanning of the sample. In the previous contact mode,
biological samples weakly attached to a substrate surface areeasily swept away by the cantilever tip.
In the early stage when pioneering studies of bio-AFM
imaging were being carried out, several groups attempted toobserve dynamic biological events. Hansma and colleagues
observed at 1 min intervals the clotting process by fibrin
which was initiated by the digestion of fibrinogen withthrombin [11]. A group including one of the inventors of
AFM (i.e. Binnig) also attempted to image the dynamic
processes of live cells being infected by virus [27] and bindingof antibody to the S-layer [28]. After the introduction of
the tapping mode, DNA-involved dynamic processes werestudied by Hansma’s group and Bustamante’s group; DNAdigestion by DNAase [29] binding of RNA polymerase
to DNA [30], transcription process [31] and diffusion of
RNA polymerase along DNA strands [32]. Other types ofdynamic processes studied were digestion of collagen I bycollagenase [33] and closing of nuclear pores upon exposure
to 5% CO2 [34]. The imaging rate used in these studies wasvery low as a matter of course. Even in the fastest imaging,it was ∼2 frames min−1 . Therefore, no clear images wereacquired for rapidly moving molecules and their time-lapse
images could not well resolve the dynamic processes. To
circumvent the low imaging rate, Radmacher measured heightfluctuations (∼1 nm) at a top-surface locus of lysozyme
which specifically occurred when the enzyme was digestingoligoglycosides [35]. A similar study was carried out withurease and ∼1 nm height fluctuations of 100–200 Hz weredetected [36].
Looking back these early studies, it is obvious that AFM
was expected to play a role in visualizing dynamic biologicalprocesses at high spatial resolution because AFM is only
the method capable of directly visualizing vital biologicalsamples in aqueous solutions. However, this expectation could

not be met by the low imaging rate. As mentioned above,rather than AFM, single-molecule fluorescence microscopy
created in the early 1990s has been much more successful
in observing dynamic biomolecular processes, although theobservation is performed in an indirect way.
As will be described later, efforts towards increasing
the imaging rate of AFM have been carried out since
the early 1990s. Through steady efforts for two decades,
high-speed AFM has now materialized. In fact, several
biomolecular processes have recently been captured clearly,
and the number of publications on high-speed AFM imaging
has rapidly increased in the last 2 years. In this review,a historical overview of technical developments towards
high-speed AFM is first described, which is followedby theoretical considerations for the imaging rate anddescriptions of elementary devices and techniques specialized
for high-speed AFM. Then, recent studies of high-speed AFM
imaging are highlighted. This review is concluded by the
discussion of future challenges to be met in order to expand
the scope of high-speed AFM applications in biologicalresearch. More comprehensive descriptions of high-speed
AFM instrumentation are given elsewhere [7]. Details of thehistorical overviews of bio-AFM development and application
are also given elsewhere [37, 38].

2. History of high-speed AFM development
2.1. Initial stage
In the early 1990s, only a few years after the invention of
AFM, Calvin Quate and colleagues initiated the developmentof high-speed AFM [39]. Quate, one of the inventors of
AFM, aimed at fast inspection and lithography of a wide
area of hard material surfaces using AFM. To this end, they
developed cantilevers and cantilever arrays with integrated
sensors and/or actuators using MEMS technology [40–44].
The self-sensing and self-actuation capabilities significantly
simplified the configuration of AFM set-up. However, the2Nanotechnology 23 (2012) 062001

Topical Review

fabrication of these cantilevers was only possible in relatively
large dimensions, resulting in low resonant frequencies andlarge spring constants. Insulation coating of the integrated
cantilevers for the use in liquids further lowered the resonant
frequency [45]. This approach, therefore, did not extend tohigh-speed AFM for biological research. Nevertheless, theirintention of increasing the scan speed of AFM was taken overby Hansma and Ando.
Hansma’s group and my group employed the same
approach, i.e. increasing the feedback bandwidth by developing fast-response devices. Both groups developed smallcantilevers [6, 46, 47] and optical beam deflection (OBD)
detectors for small cantilevers [6, 48]. My group additionallydeveloped a fast scanner and a fast amplitude detector [6].
Hansma’s group reported imaging of DNA at 0.6 frames s−1(fps) [49] and dynamic association/dissociation between
GroES and GroEL [5]. However, the feedback bandwidth
was not high enough and therefore the molecular events wererecorded by scanning the sample stage only in the x and z
directions. A more complete high-speed AFM system wasreported by my group in 2001 and 2002 [6, 50], whichenabled filming of the Brownian motion of myosin V on mica
surfaces at 12.5 fps [6]. However, the feedback bandwidth
was not high enough to have a capability of low invasivenessto fragile proteins. The period from the early 1990s through2002 is considered to be the initial stage of high-speed AFM
development.

an ambient environment or to very stable non-biologicalsamples in liquids. It is notable that, in this developmental
stage, researchers of control engineering participated indeveloping fast positioning techniques for AFM using moresophisticated control techniques. However, the complicatedcontrol techniques can only be implemented with slow digital
circuits. Schitter, one of the previous collaborators of Hansma,
is now trying to implement sophisticated control schemes
into emerging field programmable analog array (FPAA)
circuits [59].
2.3. Current stage
After the second stage, the number of users of thehigh-speed AFM apparatus designed by my group is
rapidly increasing. Consequently, a relatively wide range ofapplication studies are now being carried out. Regarding
the technical development of high-speed AFM, a wide-areaand high-speed scan capability is being pursued. However,the feedback bandwidth seems to have already reachedthe maximum possible level (∼100 kHz) as far as theconfiguration of the AFM set-up similar to previous ones
is used (i.e. tapping mode, piezoactuator-based sample-stage
scanner, small cantilevers and OBD detection of cantileverdeflection). Rather than attempting to further increase the scan
speed, functional augmentation to high-speed AFM is now
being pursued and bringing a new modality of bio-imaging
which will be briefly described in section 6.

2.2. Second stage

3. Theoretical basis for feedback bandwidth andimaging rate

In the second stage from 2003–2007, further efforts werecarried out. A notable improvement was achieved by
the invention of an active damping technique for thez-scanner [51] and a dynamic proportional–integral-derivative
controller [52]. Further smaller cantilevers with a resonant
frequency of 1.2 MHz in water and a small spring constantof ∼0.2 N m−1 were developed by Olympus. As a result,the feedback bandwidth was significantly increased up to∼100 kHz. Fast phase imaging was also achieved by thedevelopment of a fast phase detector capable of detecting
the phase signal every oscillation cycle of a cantilever [53].
In 2008, my group finalized the development of high-speed
AFM suitable for observing dynamic structural changes of
biological molecules [7]. Since then, my group has beenshifting our efforts towards applying the finalized high-speed
AFM instrument to the study of proteins and developing thenext generation of high-speed AFM applicable to live cells.
Hansma’s group developed fast data acquisition systems [54,55] and high-speed scanners with a relatively large scan
range (6–15 µm in x and y directions and 6 µm in thez direction) [55, 56]. Miles’ group developed a resonant
x-scanner using a tuning fork with resonant frequencies of
30–100 kHz [57] and a fast digital processing system for
acquiring cantilever deflection data [58], and demonstrated
ultrafast imaging (1300 fps) of collagen fibers in an ambientenvironment [58]. This remarkable speed was, however,achieved in the constant-height mode (without feedback scanof the z-scanner). Their high-speed AFM is therefore only
applicable to samples firmly attached to a substratum inHere, factors limiting the scan speed of tapping mode AFM
are quantitatively described. It will be shown that the highest
possible imaging rate Rmax is determined by the bandwidth of
feedback control to maintain the tapping force constant during
scanning as well as by scan parameters and the sample itself.
3.1. Highest possible imaging rate
When a sample is scanned over an area of W × W withscan velocity Vs in the x direction and N scan lines in they direction, the image acquisition rate R is given by R =Vs /(2WN). Supposing that the sample surface corrugationis characterized with a single spatial frequency ν in the x
direction, then the feedback scan is executed in the z direction
with frequency f = νVs to trace the sample surface.
The bandwidth fB of the closed feedback loop should beequal to or higher than f , and thus we obtain the relationshipR ≤ fB /(2νWN). The feedback bandwidth fB is usually
defined by a feedback frequency f at which a closed-loop
phase delay of π/4 occurs in tracing the sample surface.
The phase delay in the closed loop (ϕclosed ) is approximately
twice that in the open loop (ϕopen ), provided the feedback
gain is maintained at ∼1 [7]. The phase delay is causedby the limited response speed of devices contained in the
feedback loop and by the time delay due to a ‘parachuting’effect mentioned below. By denoting the total time delay by1τtotal , ϕopen is given by ϕopen = 2πf 1τtotal . The phase delay
3

Nanotechnology 23 (2012) 062001

Topical Review

Figure 2. Feedback loop of tapping mode AFM and main timedelays. The input to the feedback loop is the variation of sample
surface height and the output is the displacement of the z-scanner.
The time delay in the loop is mainly produced by the cantileverresponse time (τc ), the parachuting time (τp ), the time required tomeasure the cantilever oscillation amplitude (τm ), the integral timeof error signal at the PID controller (τI ) and the z-scanner responsetime (τz ).
Figure 3. Small cantilever and EBD tip. (a) SEM image of a smallcantilever. (b)–(d) SEM images of EBD tips. The apex radius of thetip formed by EBD is not small (c) but can be reduced by nitrogenor oxygen plasma etching to 4–5 nm (d).

can be compensated by a factor of α using the (P + D)
operation of the PID feedback controller. fB is thus given
by fB = α/(161τtotal ). Hence, the highest possible imaging
rate is given by Rmax = α/(32νWN1τtotal ), when the sample
fragility is not taken into account. To materialize, for example,
Rmax = 25 fps for α = 2.5, ν = 0.1 nm−1 , W = 200 nm
and N = 100, we have to achieve fB = 100 kHz and thus
1τtotal = 1.6 µs.

z-scanner is driven stepwise [7] but this assumption is notpractical because the z-scanner is driven by the output
signal from an analog feedback controller through thez-piezodriver. The integral time of error signal with a feedback
controller τI cannot be expressed explicitly but was previously
given through a theoretical consideration and experimental
measurements [7]. It is approximately expressed by τI =4h0 sin(ϕclosed /2)/(A0 fc ), where h0 is the sample height andA0 is the free oscillation amplitude of the cantilever.
When the cantilever encounters a steeply inclined regionof the sample, the tip completely detaches from the sample
surface and time elapses until it lands on the surface again(i.e. parachuting). The parachuting time is prolonged when
the amplitude set point As to be maintained constant during
scanning is set close to the free oscillation amplitude A0 .
This is because the error signal (A0 − As ) to be integrated
by a feedback controller is saturated at a small value during
parachuting [7, 52]. An approximate analytical expression
for the parachuting time τp was previously given to be τp =[(tan β)/β − 1]/fc , where β is given by β = cos−1 [(A0 −As )/{5h0 sin(ϕclosed /2)}] [7].

3.2. Time delay
In the closed feedback loop, there are several devices withcertain time delays to response (figure 2). The main timedelays are as follows; (1) the response time of an oscillating
cantilever, (2) the time required for measuring the cantilever
oscillation amplitude, (3) the time of integrating the error
signal with a feedback controller, (4) the response time of thez-scanner and (5) the ‘parachuting’ time mentioned below.
When a cantilever tip taps a sample surface, a brief
stepwise force is exerted on the cantilever. The cantileverresponds to this stepwise force with a response time given byτc = Qc /(πfc ), where Qc and fc are the cantilever’s quality
factor and the first resonant frequency in an aqueous solution,
respectively. The minimum time required for measuring
the cantilever oscillation amplitude τm is given by τm =1/(2fc ). The z-scanner is driven at a feedback frequency
f . The response of the z-scanner with the first resonant
frequency fz and a quality factor Qz is characterized witha second-order transfer function. The phase delay ϕz at the
feedback frequency f is estimated from the phase–frequency
relationship (i.e. the Bode phase plot) of the transfer function.
The time delay τz is given by τz = ϕz /(2π f ). A previous
argument as to τz is based on an assumption that the

4. Instrumentation of high-speed AFM
In the high-speed AFM apparatus we developed, all thetime delays mentioned above are minimized. This apparatus
is now commercially available from Research Instituteof Biomolecule Metrology (RIBM), Co., Ltd (Tsukuba,
4

Nanotechnology 23 (2012) 062001

Topical Review

Japan). Here, devices with minimized time delays are briefly
described.
4.1. Small cantilevers, OBD detector and amplitude detector
Several types of small rectangular cantilevers made ofsilicon nitride have been developed by Olympus collaboratingwith my group [6, 60]. The highest end cantilevers havedimensions of 6 µm long, 2 µm wide and 90 nm thick(figure 3(a)). The resonant frequencies are 3.5 MHz in air
and 1.2 MHz in water. The spring constant is 0.2 N m−1
and the quality factor in water is ∼2. Therefore, the responsetime in water is ∼0.5 µs. A slightly larger type (BioLeverfast BL-AC10DS-A2, Olympus; 9 µm long, 2 µm wide
and 130 nm thick; resonant frequencies 1.5 MHz in air
and 0.6 MHz in water; spring constant ∼0.1 N m−1 )is commercially available from Olympus (Tokyo, Japan),
Atomic Force F&E GmbH (Mannheim, Germany), Bruker(CA, USA) and Asylum Research (CA, USA). Although
not yet commercialized, small cantilevers will soon also be
available from NanoWorld AG (Neuchâtel, Switzerland). Thechange in the angle 1ϕ of a cantilever’s free end is given by
31z/2L, where 1z is the displacement of the free end and
L is the length of the cantilever. Therefore, for the deflection
detection of small cantilevers, angle-based OBD detectors areadvantageous compared with distance-based detectors. The
shorter cantilevers give 10–20 times higher sensitivity to an
OBD detector than conventional cantilevers.
The tip of the small cantilevers manufactured by Olympus
has a bird-beak-like shape with an apex radius of 15–24 nm.
Small cantilevers with a sharper tip made by electron beam
deposition (EBD) are also available as an option. When ascanning electron microscope (SEM; a low vacuum type isideal) is available, EBD tips can be easily grown under an
atmosphere of phenol gas sublimated from phenol crystals(figures 3(b) and (c)). They grow at ∼1 µm min−1 under alow vacuum condition. The tip can be sharpened by nitrogenor oxygen plasma etching (figure 3(d)). The apex radius canbe reduced usually to ∼4 and to ∼0.5 nm in the best case.
In the OBD detector (figure 4), a 20× objective lenswith a long working distance of 24 mm (SLWD 20×, Nikon)
is used to focus a laser beam onto a small cantilever [6].
This lens is also used to collect a laser reflected back fromthe cantilever and to view the cantilever and sample stage
under an optical microscope. The incident and reflected laser
beams are separated using a quarter-wavelength plate and apolarization beamsplitter.
The oscillation amplitude of a cantilever is measured bya peak-hold method at every half-cycle of oscillation (i.e.
τm = 1/(2fc )) [6] or by a Fourier method at every cycleof oscillation (i.e. τm = 1/fc ) [7]. In the former method,
the peak and bottom voltages of a sinusoidal signal output
from the split photosensor of the OBD detector are heldby sample/hold circuits, and their difference is output as apeak-to-peak amplitude signal. Since only two-point voltages
are used, it is susceptible to the effect of thermal fluctuationsof a cantilever. The latter method uses almost all the signals
within one oscillation cycle and hence is much less susceptible
to the thermal effect.

Figure 4. Schematic presentation of objective lens type of OBD
detection system. The collimated laser beam is reflected up by thedichroic mirror and incident on the objective lens. The beamreflected back from the cantilever is collimated by the objective lens,separated from the incident beam by the polarization beamsplitterand λ/4-wave plate, and reflected onto the split photodiode. The
objective lens is also used to view the cantilever, the focused laser
spot and the sample stage through the CCD camera.

4.2. Sample-stage scanner and active damping of z-scanner
To design a sample-stage scanner to be driven at highfrequencies, we have to consider various factors; not only
the scanner’s resonant frequency and unwanted vibrations
but also hydrodynamic pressure. Quick displacement of asample stage in the z-direction produces high hydrodynamic
pressure, which tends to move the cantilever chip as well as
the cantilever holder since they are located close to the sample
stage [50]. This movement results in a considerable time delayin the cantilever response. Therefore, we use a small sample
stage 1–2 mm in diameter. In addition, the laterally overlapped
region of the sample stage and the cantilever is minimized.
This means that only areas near the edge region of the sample
stage are imaged [61].
Among the x-, y- and z-scanners, the z-scanner isdisplaced at the highest frequency. This high-frequency
displacement produces a large impulsive force to its
supporting mechanism, which is apt to cause unwanted
vibrations of the z-scanner. To counteract the impulsive force
and thereby minimize vibrations, two identical piezoactuators
are placed at a supporting base in the opposite direction anddisplaced simultaneously with the same length [6, 7, 50, 62].
An alternative method we have recently employed is to holda piezoactuator at its four rims parallel to the displacement
direction so that the center of mass of the piezoactuator is not
displaced [63]. In the former method, the resonant frequency
and the maximum displacement are ∼170 kHz and 2 µm,
respectively, while in the latter method they are ∼370 kHz
and 1 µm, respectively. In the x-scanner with the maximum
displacement of ∼2 µm, a piezoactuator is held at both ends
with flexures so that its center of mass is unchanged during
scanning [7]. The slowest y-scanner displaces a block of the5Nanotechnology 23 (2012) 062001

Topical Review

Figure 5. High-speed scanner and active damping. (a) Sketch of high-speed scanner. A sample stage (glass rod) is glued on the top surfaceof the z-piezoactuator using nail enamel (a dummy stage attached to the bottom surface of the z-piezoactuator is hidden). The
z-piezoactuator is held at the four rims parallel to the displacement direction. The dimensions (W × L × H) of the z-piezoactuator are
3 × 3 × 2 mm3 . The gaps in the supporting base are filled with an elastomer for passive damping. (b) Active damping of z-scanner
vibrations. The LRC circuit (mock z-scanner) is constructed so that its transfer function is similar to that of the real z-scanner.

x- and z-scanners, the x-scanner displaces a block of thez-scanner and the z-scanner displaces a glass-rod sample stage(figure 5(a)).
To damp z-scanner vibrations, we developed an active
damping technique based on a Q-control method with a ‘mockz-scanner’ [51]. In a conventional Q-control method [64, 65],
the displacement or velocity (Vz ) of the z-scanner has to bemeasured and a signal proportional to −Vz is added to thesignal to be input to the z-piezodriver (i.e. output from a
feedback controller). However, it is difficult to measure thedisplacement or velocity. In our method, the mock z-scanner(an LRC circuit) characterized with a transfer function similar
to that of the real z-scanner is used (figure 5(b)). An input
signal to the z-piezodriver is also input to the mock z-scannerand the output from the mock z-scanner is used for Q-control(i.e. its minus derivative is added to the output signal froma feedback controller). As a result, the quality factor Qz is
reduced to 1–2. Although it is possible to further reduce Qz
down to 0.5, we do not do so because the smaller Qz results
in a larger phase delay and also because the first resonanceat 370 kHz would not be excited by a feedback signal witha frequency of ∼100 kHz. The time delay of the z-scanner is∼0.36 µs for 100 kHz driving.

4.3. Feedback controller and drift compensator
It is difficult to make high-speed imaging compatible with low
invasive imaging. In order for the oscillating cantilever tip
to tap the sample surface with a small force, the amplitude
set point has to be set very close to the free oscillation
amplitude. As already mentioned, this condition promotes
tip-parachuting and prolongs its duration. We solved thisproblem by devising a new PID controller (named a dynamic
PID controller) with a capability to automatically change the
PID’s gain parameters, depending on the cantilever oscillation
amplitude [52]. When the amplitude exceeds a threshold level(set at the amplitude set point or slightly above), the gains are
increased. The oscillation amplitude thereby quickly returns
towards the set point before the error signal is saturated.
Consequently, the feedback bandwidth becomes independentof the set point as long as the set point is smaller than
0.9–0.95.
When a piezoactuator is driven continuously for a longtime, its temperature is increased. As a result, the extension
coefficient of the piezoactuator is usually lowered to someextent. In the tapping mode, a cantilever is continuously
excited by a sinusoidally oscillated piezoactuator placed at
the cantilever holder. The cantilever free oscillation amplitude
6

Nanotechnology 23 (2012) 062001

Topical Review

A0 gradually declines by the thermal effect. When A0 is set
at 1 nm and the set point As is set at 0.9A0 (these values
are routinely used for imaging delicate biological samples),their difference is only 0.1 nm. Under this condition, thedownward drift of A0 easily leads to complete dissociationof the cantilever tip from the sample surface because thefeedback system misinterprets the change in the cantilever
oscillation amplitude and hence withdraws the sample stage
from the cantilever tip. Thus, to achieve stable imaging undera small tapping force, A0 has to be kept constant precisely.
However, there is no way to measure A0 during successive
imaging. Instead of measuring A0 , we measure the second
harmonic oscillation amplitude A2nd which is sensitive to thetip–sample interaction [52, 66]. A2nd is maintained constantby changing the cantilever excitation power using a slow
feedback control [52]. This compensation method allows very
stable imaging under a small tapping force.

captured but fragile samples such as microtubules were
often broken during imaging. Torimitsu’s group imaged
the complex of biotinylated DNA and streptavidin but
the complex was disrupted during imaging [75]. Using a
feedback-control-free high-speed AFM apparatus developedby Miles’ group [57], Hobbs’ group imaged growing
polyethylene oxide films at 14 fps but the resolution was
low [68]. Takeyasu’s group successfully observed the opening
and closing of the apical region of GroEL during the ATPasereaction as well as the dynamic interaction between GroEL
and GroES [70]. However, in this study (as well as in theprevious study mentioned above [62]), GroEL was placed ona mica surface in an end-up orientation, so that one of thetwo rings was invisible and inaccessible to GroES. Therefore,
the postulated [120] but controversial [121] asymmetricconformational changes in the double-ring structure andthe resulting asymmetric binding of GroES could not be
examined. This experimental restriction was removed later by
the preparation of an appropriate substrate surface [82, 93].
Among various imaging studies carried out in thisstage, the dynamic visualization of EcoP15I–DNA complex
performed by Takeyasu and collaborators [74] is notable.
EcoP15I is a type III restriction–modification enzyme with a
low ATPase activity. The establishment of the endonucleaseactivity requires complex formation by the two molecules
bound to unmethylated and inversely oriented specific nonpalindromic recognition sites (i.e. 50 -CAGCAG—CTGCTG)
which are distantly separated (up to 3.5 kb) on thesame DNA molecule. Thus, it has been an interestingquestion how the two enzyme molecules efficiently encounter
each other to overcome the distance and to cleave the
DNA double strand [122]. A simple model accounting
for the efficient encounter could be that the ATPasereaction by the enzyme molecules bound to the recognition
sites drives translocation of the DNA double strand to
extrude loops, thus efficiently leading to collision of themolecules. However, the ATPase activity is too low to
explain such a long translocation [123]. The high-speed
imaging showed that EcoP15I in fact translocated DNA
in an ATP-dependent manner and consequently caused
accumulation of supercoiling1 . The enzyme bound to its
recognition site also made non-specific contacts with otherDNA sites, thus forming loops (diffusive looping). The loops
formed were extruded by the ATPase-driven translocationof DNA. Thus, the two mechanisms (translocation anddiffusive looping) effectively reduce the distance between therecognition sites. However, recent single-molecule magnetic
tweezers experiments showed that a DNA double strand
stretched so as not to form loops could be cleaved, suggestingthat one-dimensional diffusion of an EcoP15I molecule
is sufficient to reach a second molecule attached to its
recognition site, and also that DNA cleavage occurs evenwhen an EcoP15I molecule within the dimeric complex isnot bound to its recognition site on the DNA [124]. A
recent review discussed these and other experimental results
to reconcile different models and provided suggestions forfurther experiments [125].

5. High-speed AFM imaging
As shown in table 1, from 2001–2008 the numberof publications on high-speed AFM imaging gradually
increased, and sharply increased in the most recent 2 years.
This leap reflects the recent maturity of the high-speed
AFM technique and the increase in the number of users.
A prototype of a high-speed AFM apparatus began to becommercially available from RIBM around 2006. In 2008 an
international consortium of high-speed bio-AFM was foundedby the cooperation of six laboratories throughout the world.
These laboratories are using the highest-end high-speed AFM
instruments designed by my group.
5.1. Early studies
Even in the period when the development of high-speed
AFM instruments was going on, a few groups carriedout high-speed bio-imaging experiments using the test
and prototypic apparatuses. Through imaging experiments,
my group examined the capability of test apparatuses
to find instrumental problems to improve. For example,
visualization of the following biomolecular processes
was attempted; calcium-induced dissociation of calmodulin(CaM) from the neck regions of myosin V (M5) [67],gliding movement of actin filaments over a myosin-coatedsurface (movie S1 available at stacks.iop.org/Nano/23/
062001/mmedia) [62], conformational change of singleheaded dynein C hydrolyzing ATP [62], kinesin movement
along microtubules [67] and association/dissociation between
GroES and GroEL under the control of the GroEL ATPase
cycle [62]. It is noteworthy that flash photolysis of caged
compounds was introduced for the first time to AFM
imaging [62, 67]. This technique is well matched withhigh-speed AFM when transient events triggered by therelease of ligands are visualized. It was used to observe CaM
dissociation from M5 upon Ca2+ release and GroES binding
to GroES upon ATP release (movie S2 available at stacks.
iop.org/Nano/23/062001/mmedia) [62, 67]. Even in theseearly trials, dynamic biomolecular events were successfully

1 See supplemental movie available at: www.pnas.org/content/suppl/2007/

07/13/0700483104.DC1/0843Movie3.avi.
7

Nanotechnology 23 (2012) 062001

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Table 1. Chronological progress of publications on high-speed AFM imaging.
Research
group

Year

na

Targets and subjects

2001
2002
2003

1
0
2

2004
2005

0
3

• Brownian motion of myosin V [6]None• Brownian motion of myosin V [50]• Kinesin movement along MT &CaM dissociation from myosin V [67]None
• Actin gliding, structural change of dynein C, GroES–GroEL
interaction [62]• Polyethylene oxide films during crystal growth [68]• Polymer films during crystallization from the melt [57]• MT destruction and conformational change of myosin V [69]
• Conformational change of GroEL and GroEL–GroES interaction [70]• DNA–nuclease interaction [71]• Crystallization of a CaCO3 thin film from supersaturated solution [72]
• Ultrafast imaging of collagen [58]• Brownian motion and photodegradation of π -conjugated polyrotaxane [73]• DNA translocation and looping by type III restriction enzyme [74]• Biotinylated DNA–streptavidin interaction [75]
• Anisotropic diffusion of point defects in streptavidin 2D crystals [76]• Identification of intrinsically disordered regions of proteins [77]• Human chromosomes in liquid [78]• DNA–nuclease interaction [79]• Effect of Ca2+ on vesicle fusion [80]• Dynamic equilibrium at the edge of bR 2D crystals [81]• Structural change of CaM and actin polymerization on streptavidin 2D
crystals [82]• Unidirectional translocation of cellulase along cellulose fibers [83]• Translocation of EcoRII restriction enzyme along DNA [84]• Fabrication and imaging of hard material surface [85]• Purple membrane in contact mode HS-AFM [86]• Thermal motion of π-conjugated polymer chain [87]• Opening of 3D hollow structure of DNA origami [88]• Opening of 3D hollow structure of DNA origami [89]• ATP-induced conformational change in P2X4 receptors [90]• Walking myosin V along an actin filament [91]• Photo-induced structural change in bR [92]• 2D crystal formation of annexin A–V and height change of p97 [93]• Analysis of components covering magnetosome surface [94]• Time course of cell death by antimicrobial peptide [95]• Dissolution of extreme UV exposed resist films under developing [96]• Dissolution of extreme UV exposed resist films under developing [97]• Process of forming supported planar lipid bilayer [98]• Self-assembly of amyloid-like fibrils [99]• Effect of ClpX on FtsZ polymerization [100]• Membrane-mediated protein–protein Interaction [101]• Photodegradation of chiral helical π-conjugated polymer chain [102]• Base-excision repair of damaged DNA in DNA origami scaffold [103]• Tension effect on DNA methylation [104]• Formation/disruption of G-quadruplex in DNA nanoscaffold [105]• Programmed self-assembly of DNA origami jigsaw pieces [106]• Programmed self-assembly of DNA origami jigsaw pieces [107]• Dissociation of histone from nucleosome [108]• Height change of acid-sensing ion channel 1a upon acidification [109]• Breaking of nanocapsules of DNA–azobenzene conjugate by UVlight [110]• Structural change of bR trimer in a stretched membrane [111]• Morphological transformation of detergent micells [112]• bR in response to alternating two-color illumination [113]• Rotary catalysis of rotorless F1 -ATPase [114]• Dislocation of cellulase hydrolyzing cellulose fibers [115]• CENP-W complexed with CENP-T [116]• Opening of DNA cuboid [117]• Synthetic molecular motor [118]• Ferritin adsorption onto a silicon substrate [119]

2006

3

2007

5

2008

5

2009

10

2010

21

2011 up to
September 2

8a n represents the number of publications in respective years.

8

Ando
Ando
Ando
Ando
Hobbs
Miles
Ando
Takeyasu
Takeyasu
Miles
Miles
Shinohara
Takeyasu
Torimitsu
Ando
Ando
Miles
RIBM
Torimitsu
Ando
Ando
Igarashi
Lyubchenko
Miles
Scheuring
Shinohara
Sugiyama
Sugiyama
Shinozaki
Ando
Ando
Ando
Ando
Fantner
Itani
Itani
Le Grimellec
MilhietOgura
Scheuring
Shinohara
Sugiyama
Sugiyama
Sugiyama
Sugiyama
Sugiyama
Takeyasu
Takeyasu
TanakaTorimitsu
Ando
Ando
Ando
Ando
Fukagawa
Sugiyama
TurberfieldYamamotoNanotechnology 23 (2012) 062001

Topical Review

Figure 6. High-speed AFM images showing the process of SLB formation. Lipid membranes were made of a ternary mixture made of
DOPC/DOPS/biotin-cap-DPPE in buffer (10 mM HEPES, 150 mM NaCl, 2 mM CaCl2 , pH 7.4). The liposome suspension (0.5 mg ml−1 )was then directly diluted into the buffer present in the liquid cell equipped in the high-speed AFM instrument and the bilayer formation was
imaged at 1.03 fps (scan size 800 × 800 nm2 ). SLB was formed from LNT (white arrows, approximately 20 nm in height) and liposomes(arrowheads, 7.5–30 nm in height). Z color scale, 25 nm. The light-blue arrow indicates the interaction between a patch of SLB and one endof an LNT.

group (movie S4 available at stacks.iop.org/Nano/23/062001/
mmedia) [98]. The lipid composition of small vesicles
used was DOPC:DOPS:biotin-cap-DPPE = 6:2:2 (w/w)
suspended in 10 mM HEPES–NaOH (pH 7.4), 150 mM
NaCl and 2 mM CaCl2 (DOPC: dioleoylphosphatidylcholine;
DOPS: dioleoylphosphatidylserine; biotin-cap-DPPE: 1,2dipalmitoyl-sn-glycero-3-phosphoethanolamine- N-(capbiotinyl)). The imaging was carried out over 25 min at 1.03 fps fora scan area of 800 nm × 800 nm. An unexpected phenomenon
observed was the formation of lipid nanotubes (LNTs) of∼20 nm diameter (figure 6). Short LNTs had already been
formed in the vesicle sample before the deposition onto amica surface as they appeared together with small globular
vesicles in the AFM images immediately after the deposition.
Unlike the globular vesicles, the short LNTs did not ruptureon the mica surface and grew longer by two mechanisms;(1) addition of free lipids to the free ends and (2) the
end-to-end fusion of LNTs. The LNTs also interacted withthe edge regions of SLBs either at their longitudinal sides
or at one of the free ends. In the latter case, the interacting
SLBs were getting smaller. With the lapse of time, the lengthand density of LNTs increased and the size of SLBs also
increased. As a result, long LNTs were jammed into narrow
spaces between large SLBs, thus pushing each other andeventually rupturing. After the rupture, the size of SLBs
immediately increased to cover large areas of the bare mica
surface.
As demonstrated by this observation, various dynamic
events appear simultaneously in a molecular movie (movie S4available at stacks.iop.org/Nano/23/062001/mmedia), which
is one of the powerful features of high-speed AFM. The high

5.2. Recent studies
As listed in table 1, several targets and subjects havebeen fully studied in the most recent 3 years. Here,these studies are classified as (1) 2D crystal systems,(2) isolated proteins, (3) protein self-assembly processes,(4) DNA-binding proteins and synthetic DNA devices,(6) manipulation and imaging, and (7) non-biological studies.
Below, at least one topic is highlighted for each class of study,
except for non-biological studies.
5.2.1. 2D crystal systems. 2D crystal systems studied thus
far are mica surface-supported planar lipid bilayers [98],detergent micelles (movie S3 available at stacks.iop.org/
Nano/23/062001/mmedia) [112], native membrane proteins(bacteriorhodopsin (bR) [81, 92, 113] and ATP-synthasec-rings in a native lipid membrane [101]), and 2D crystalsof streptavidin [76, 82] and annexin A–V [93] formed on
mica-supported planar lipid bilayers.
5.2.1.1. Planar lipid bilayers. Small lipid vesicles (singlelamellar vesicles), which are prepared by sonication of large
multi-lamellar vesicles, attach to a negatively charged mica
surface by electrostatic interaction and then rupture. Uponrupture, small patches of supported lipid bilayers (SLBs)
are formed. These patches are fused into larger SLBs andeventually the entire mica surface is completely covered by
the bilayers. This entire process was revealed for the first timeusing high-speed AFM.
The SLB formation process was recorded in depth in
the collaboration between my group and Le Grimellec’s
9Nanotechnology 23 (2012) 062001

Topical Review

Figure 7. Defect diffusion in streptavidin 2D crystal. (a) Schematic of a streptavidin molecule on biotinylated lipid bilayer. Two biotin
binding sites occupied by biotin are indicated by the closed circles. The open circles indicate biotin binding sites facing the aqueous solution
and are biotin-free. (b) High-speed AFM images of streptavidin 2D crystal with C222 lattice and monovacancy defects therein. The
monovacancy defects are enclosed by dashed squares and circles. The directions of the lattice vectors of the crystal are also indicated.
Successive images were obtained at an imaging rate of 2 fps with a scan area of 150 × 150 nm2 and the number of pixels was 200 × 200.
(c) Schematic of streptavidin arrays in C222 crystal. Unit lattice vectors are indicated. The a axis includes rows of contiguous biotin-boundsubunits, while the b-axis includes rows of contiguous biotin-unbound subunits.

stability of LNTs can be accounted for by the smaller surface
curvature (and hence smaller surface tension), compared withthat of small globular liposomes. In the sideways surfacesof the edge regions of SLBs and the ends of LNTs, thepolar head groups rather than alkyl chains are exposed.
The arrangements of lipid molecules at these surfaces areenergetically unfavorable compared with those in the flat
surface area of SLBs and the longitudinal surfaces of
LNTs. Therefore, to reduce the fraction of these unfavorable
surfaces, SLBs and LNTs grow by incorporating free lipids
and by fusing with the respective kind of partners. As
mentioned above, when one of the ends of an LNT was
brought into contact with a sideways edge of an SLB patch,
the size of the patch became smaller with time, indicatingthat lipids are transferred from the SLB to the LNT, although
the concomitant growth of the LNT was not apparent. The
SLBs are thin (a few nanometers) while the LNTs are ∼20 nm
in diameter, so that the arrangements of lipids facing at
the sideways edges of SLBs are energetically unfavorable,compared with those of lipids at the ends of LNTs. This
energetic difference would account for the lipid transfer. The
mechanism underlying LNT formation and the molecular
packing in an LNT are still elusive. It would be likely that the
LNTs have a double-layered or monolayer-stacked structurewith a hollow in the center, as has been revealed for different
LNTs [126, 127]. LNTs naturally occur in life and synthetic
LNTs are expected to have various applications including
encapsulation of drugs. A study similar to the above wasrecently carried out using a cationic surfactant, aiming at
an eventual goal of elucidating the process of cleaning bydetergents at the nanometer scale [112].
As will be described later, SLB surfaces are usefulfor anchoring a specific protein species without non-

specific adsorption of other species. This selective anchoring
facilitates AFM imaging of dynamic interaction events ofsamples containing multiple protein components [93].
5.2.1.2. 2D crystals of proteins on SLBs. 2D crystals ofproteins formed on SLBs have been widely studied for various
purposes; e.g. structural determination of proteins by electron
diffraction [128], understanding of crystal formation [129]and practical use of the surfaces as substrates. For the efficient
formation of 2D protein crystals on SLB surfaces, the SLB
should be highly fluidic. Lipids with unsaturated alkyl chainssuch as DOPC are often used for preparing highly fluidic
SLBs. To attach a protein to an SLB surface, lipids withfunctional groups or electrically charged lipids are included in
the SLB [93]. Here, atopic of dynamics involved in 2D protein
crystals is described.
The topic is diffusion of point vacancy defects inthe orthorhombic C222 crystal of streptavidin formed on
biotin-containing SLBs (movie S5 available at stacks.iop.
org/Nano/23/062001/mmedia) [76]. Note that the 2D crystal
has P2 symmetry [82] but here the notation C222 for thesymmetry group follows a previous report [130]. Streptavidin
is a homotetrameric protein with dihedral D2 symmetry andeach subunit specifically binds to one biotin. Streptavidin iseasily crystallized in 2D forms on biotinylated SLBs. On thebiotinylated SLBs, two of the four biotin binding sites face
the lipid layers and are occupied by biotin, while the othertwo are exposed to the aqueous environment and therefore
are biotin-free (figure 7(a)). The lipid composition used was
DOPC, DOPS and biotin-cap-DOPE (7:2:1 w/w). The C222
crystal of streptavidin with lattice constants a = 5.9 ± 0.1 nm,
b = 5.9 ± 0.3 nm and γ = 90◦ was prepared using a buffer
solution containing 10 mM HEPES–NaOH, 150 mM NaCland 2 mM CaCl2 (pH 7.4). Monovacancy defects in the10Nanotechnology 23 (2012) 062001

Topical Review

light-induced structural changes are described, while in
the latter, membrane-mediated protein–protein interaction is
described.

streptavidin 2D crystals were produced by increasing thetapping force exerted from the oscillating tip to the sample.
The imaging was carried out at 2 fps over 150×150 nm2 scan
areas.
As shown in figure 7(b), monovacancy defects were
mobile, with an anisotropic feature with respect to the twoaxes of the crystalline lattice; larger mobility along the baxis than the a axis. The a axis includes rows of contiguous
biotin-bound subunits, whereas the b axis includes rowsof contiguous biotin-bound subunits. The linear relationshipbetween the mean-square displacements and time intervals
gave the diffusion rate constants of Da = 20.5 nm2 s−1 for thea-axis component of diffusion and Db = 48.8 nm2 s−1 for theb-axis component of diffusion. This anisotropy in the mobility(Db > Da ) arises from a free energy difference between thebiotin-bound subunit–subunit interaction and biotin–unboundsubunit–subunit interaction. When a streptavidin molecule
adjacent to a monovacancy defect moves to the defectsite along the a axis, two intermolecular bonds between
biotin–unbound subunits (‘u–u bond’) and one intermolecularbond between biotin-bound subunits (‘b–b bond’) are broken.
On the other hand, when it moves to the defect site along
the b axis, one u–u bond and two b–b bonds are broken(see figure 7(c)). Therefore, the difference in the activationenergies Ea and Eb for the step movement of a monovacancy
defect along the respective a and b axes simply corresponds
to the difference between the free energy changes Gu−u andGb−b produced by the formation of the respective u–u bondand b–b bond; namely, Eb − Ea = Gu−u − Gb−b . Therefore,
the observed relationship Db > Da indicates Gu−u < Gb−b ;namely, the u–u bond affinity is higher than that of the b–bbond. The ratio of the two diffusion rate constants (Db /Da )can be expressed by Db /Da = exp[−(Eb − Ea )/(kB T)],
where kB is the Boltzmann constant and T is the absolutetemperature. Thus, from the observed value of Db /Da ∼ 2.4,
the free energy difference Gu−u − Gb−b is estimated to beapproximately −0.88kB T (T ∼ 300 K), which corresponds to−0.52 kcal mol−1 .
The monovacancy defects sometimes fused into a divacancy defect. Through several fusions, larger multivacancy
defects were formed. As the size increased, the diffusivityof multivacancy defects increased for both crystal axes,
although the diffusivity of divacancy defects was similar
to that of monovacancy defects. The higher mobility oflarger point defects increases their probability to encounter
other point defects to form larger point defects with further
higher mobilities. During the crystal growth in the presenceof free streptavidin in the bulk solution, this acceleration
effect should promote the removal of point defects from thecrystalline regions. This would be one of the mechanisms fordefect-free crystal formation not only in 2D crystals but also
in 3D crystals. This issue should be examined in the future byimaging the crystal growth in the presence of free streptavidinin the bulk solution.

bR in response to light. To explore appropriate conditions
for capturing AFM images of high spatial resolution andto demonstrate the high spatial resolution power of AFM,
the purple membrane, which contains highly packed 2D
arrays of photoreceptor bR molecules, has often beenused in preference as a sample to image [131]. In fact,
faithful high-resolution images capable of distinguishingthe individual interhelix loops of bR are obtained [132].
Nevertheless, the physiologically relevant structural changes
in bR (i.e. response to light) have never been captured by
AFM until recently. bR functions as a light-driven proton
pump, transferring protons across the membrane from thecytoplasmic side to the extracellular side [133, 134]. bR iscomprised of seven transmembrane α-helices (named A–G)
surrounding the retinal chromophore covalently bound to
Lys216 via a protonated Schiff base [135, 136]. In the purple
membrane, bR assembles into trimers and the trimers arearranged in a hexagonal lattice. Upon absorption of light,
photoisomerization from all-trans to 13-cis conformationof the retinal chromophore takes place. This isomerizationinduces proton transfer from the Schiff base to Asp85,
which triggers a cascade of changes in the bR structure.
A series of intermediates designated J, K, L, M, N and O
are defined by spectroscopy, and M (M410 ), which has anabsorption peak at 410 nm and the longest lifetime, is only
the intermediate containing a deprotonated Schiff base [134].
The light-induced conformational changes in bR in the frozen
activated state of the wild type (WT) and bR mutants
have been studied by x-ray crystallography and electron
microscopy [137–139]. The extent of the changes reported
varies, depending on the methods used, but it is generally
small (0.1–0.3 nm). The dynamic nature of the conformationalchanges, however, remained elusive due to the lack oftechniques amenable to the high-resolution visualization of
dynamic structural changes of proteins. My group, for the
first time, succeeded in capturing high-resolution successive
images showing dynamic changes of bR in response to lightusing high-speed AFM [92].
The photocycle of the wild type at neutral pH
proceeds very fast (∼10 ms). We attempted to visualize
the conformational changes at the highest possible imaging
rate of ∼80 fps by narrowing the imaging area but wereunable to detect discernible changes. To slow down thephotocycle, we then used the D96N bR mutant that has alonger photocycle (∼10 s at pH 7) but still retains a proton
pumping ability [140]. Figure 8(a) presents successive imagesof the D96N mutant at the cytoplasmic surface captured at
1 fps. Upon illumination with green light (532 nm, 0.5 µW),bR drastically changed its structure (compare images at 1 and2 s) and returned to the unphotolysed state in a few seconds
after light-off (movie S6 available at stacks.iop.org/Nano/23/
062001/mmedia). This structural change was reproducible inrepeated dark–illumination cycles. Note that this structuralchange is specifically induced by green light, not by blue

5.2.1.3. Native membrane proteins. Two topics on native
membranes proteins, bacteriorhodopsin (bR) and ATPsynthase c-rings, are highlighted here. In the former,
11

Nanotechnology 23 (2012) 062001

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Figure 8. Structural change in bR induced by light absorption. (a) Successive high-speed AFM images of the cytoplasmic surface of D96N
bR under dark or illuminated conditions. Frame rate, 1 fps; pixel size, 200 pixels × 200 pixels. A bR trimer is highlighted by the white
triangle. The green bars indicate illumination by 532 nm green light of 0.5 µW. (b) Schematic of arrangement of bR molecules and bR
trimers under dark. The arrows indicate the directions in which respective bR molecules displace upon light illumination. (c) Schematic ofarrangement of bR molecules and bR trimers under illumination. The three bR molecules in a trefoil form a transient assembly. The arrows
indicate the directions in which respective bR molecules displaced upon light illumination. (d), (e) Surface maps of the magnified images inthe dark (d) and under illumination (e). The position of each trimer center is denoted by the white dots. A monomer in the dark is indicatedby a single white arrow. Under illumination, the topography of the monomer splits into major and minor protrusions, as indicated by green
arrows 1 and 2, respectively. The positions of seven transmembrane α helices A–G are schematically indicated on the left bottom trimer
images.

or red light. This fact clearly indicates that the observed
structural change is not an artifact but intrinsic to the nature ofbR because bR in the ground state effectively absorbs green
light. All the activated bR monomers moved outwards fromthe centers of respective trimers and rotated counterclockwisearound the centers of respective trimers, as illustrated byarrows in figure 8(b). This movement was quantified by
calculating the center of mass of each bR monomer. The
average displacement was 0.7–0.8 nm and the averagerotation angle was 7◦ –8◦ . However, the overall position ofeach bR molecule does not change because of indiscernible
alterations at the extracellular surface. The photoactivatedstate observed by high-speed AFM could be attributed tothe spectroscopically identified M intermediate because thelifetimes of other spectroscopic intermediates occurring prior
to M are very short. However, it is uncertain whether thephotoactivated state detected by AFM also contains the N
intermediate that follows M. This issue was solved by anotherstudy as mentioned later [113].
Figures 8(d) and (e) respectively show magnified AFM
images obtained before and during illumination which
are superimposed on the atomic model of the α-helicalcytoplasmic ends in the unphotolysed state (A–G) [136]. The
protruding parts around helices E and F displace outwards
from the trimer center under illumination. Since a previous
static AFM imaging study of bR has assigned the prominent
protrusion in the AFM topographs of bR to the cytoplasmic

E–F loop [132], the protrusion movement visualized here is
ascribed to the displacement of the E–F loop. As a resultof E–F loop displacement, the protrusion is divided into two
sections (green arrows 1 and 2 in figure 8(e)), and the minor
protrusion (green arrow 2) likely corresponds to the position
around helices A and B. Previous x-ray crystallography andelectron microscopy studies could not detect the displacementof loop regions because of their thin and disordered structures.
This is the reason why these previous studies only detected asmall change (0.1–0.3 nm).
As a result of the outward displacement of the E–F
loop, nearest-neighbor bR monomers, each belonging to adifferent adjacent trimer, transiently assemble, as illustrated
in figure 8(c). Here, we use a new designation, ‘trefoil’,for the triad of nearest-neighbor monomers belonging todifferent trimers to distinguish it from the original trimer.
Notably, this transient assembly in a trefoil alters the decay
kinetics of the activated state. Figure 9(a) shows the timecourse of the displacement of the mass centers for six bRmonomers belonging to two different trefoils observed at
different light intensities (Mn 1–Mn 3; n indicates different
trefoils) (see movie S7 available at stacks.iop.org/Nano/23/
062001/mmedia). Under weak illumination, it is mainly thecase that only one monomer in each trefoil is activated (M1 3and M2 2 in the left panel of figure 9(a)). When only one
monomer is activated in a trefoil, it decays with a time
constant of 7.3 ± 0.58 s (the top panel of figure 9(b)). In
12

Nanotechnology 23 (2012) 062001

Topical Review

Figure 9. Cooperative effects on the decay kinetics in D96N bR. (a) Displacement of mass center positions as a function of time measured
for six bR monomers forming two different trefoils (Mn 1–Mn 3; n indicates different trefoils) under different light intensities. The green
regions correspond to the illumination, and their shade level indicates the relative light intensity (0.007 and 0.5 µW). The blue arrows
indicate the conformational changes of the monomers (M1 3, M2 1 and M2 3) that are activated subsequent to the activation of othermonomers in the respective trefoils. (b) The top figure shows the decay of activated bR monomers (as depicted by the red circle) while theother monomers (the gray circles) within the respective trefoils are not in the activated state. The exponential decay constant is 7.3 s. On theother hand, when one or two nearest-neighbor molecules (as depicted by green circles in the middle figure) in a trefoil are already activated,the bR monomer activated latest (as depicted by the blue circles in the middle figure) decays faster. The exponential time constant isshortened to 2.0 s. In contrast, the early activated molecules within a trefoil (as depicted by green circles in the bottom figure) show anon-exponential broad distribution of decay time with an average of 13.3 s.

(movies S8 and S9 available at stacks.iop.org/Nano/23/
062001/mmedia) [113]. M410 having an absorption peak at
410 nm effectively absorbs blue light. It is known that, after
M410 is formed upon green illumination, it can be driven backto the ground state (bR570 ) by blue light [141]. High-speed
AFM imaging under alternate illumination revealed that thephotoactivated conformation of bR is also driven back to
the unphotolyzed conformation without time delay. Thus,
conformation of bR is tightly coupled to the conformationof retinal. Next, it was examined whether the activated stateof bR observed by AFM contains not only M410 but also
other intermediates that are spectroscopically different from
M410 . To address this issue, the photocycle was initiated bybrief green light illumination and then blue light was briefly
applied with different delay times. Before the blue light isturned on, the fraction of activated molecules is gradually
decreasing in the normal course of the photocycle, but it isabruptly decreased nearly to zero after the brief blue lightillumination. This abrupt decrease was independent of thetime interval between the brief two-color illumination. Thus,

contrast, under stronger illumination, two or three monomers
within a trefoil tend to be activated together. Interestingly, thedecay of each monomer markedly depends on the order of
its activation. The monomer that is activated latest among theactivated monomers in the trefoil (blue arrows in the rightpanel of figure 3(a)) decays with a shorter time constant of
2.0 ± 0.16 s (the middle panel of figure 9(b)). On the other
hand, the decay kinetics of the early activated monomers (M1 2and M2 2 in the right panel of figure 9(a)) does not follow asingle exponential (the bottom panel of figure 9(b)) and theaveraged decay time lengthens to ∼13 s. This observation
indicates that the early activated monomers do not return to
the ground state as long as the adjacent monomers within
the trefoil are in the active state. Thus, bR–bR interaction
within the trefoil (not within the trimer) under illuminationengenders both positive and negative cooperative effects in the
decay kinetics as the initial bacteriorhodopsin recovers. As aconsequence, the turnover rate of the photocycle is maintained
constant, on average, irrespective of the light intensity.
This study was followed by experiments where bR
was alternately illuminated with green and blue light13

Nanotechnology 23 (2012) 062001

Topical Review

Figure 10. Membrane-mediated interaction between c-rings. (a) Successive high-speed AFM images of c-rings in a dimer. Two c-rings arein a dimer over a time span of 1309 ms. (b) Successive high-speed AFM images of c-ring dimer that dissociates and reforms. One c-ring isintermediately monomeric for ∼400 ms (white arrows in frames: 561, 748 and 935 ms) and subsequently reforms a dimer that resembles theinitial assembly. (c) Histogram of the center-to-center distance of c-rings. (d) Membrane-mediated two-protein interaction energy landscape.
Reprinted with permission from Elsevier.

it was concluded that the conformation of bR in the activatedstate observed by AFM solely corresponds to the M410 .
The above AFM observations of bR, particularly thebR–bR interaction in the trefoil, concretely exemplified thatminute and elaborate functional mechanisms of proteinsinaccessible by other techniques can be uncovered by direct
visualization of proteins in action at high spatiotemporal
resolution.

Scheuring’s group studied this membrane-mediatedprotein–protein interaction by high-speed AFM imaging of
the edge regions of the native purple membrane [101].
Many bR trimers diffuse in the fluidic edge regions
and a small number of ATP-synthase c-rings also diffuse
there [144]. The dynamic association and dissociation
between diffusing bR trimers and the crystalline edges
were previously imaged (movie S10 available at stacks.iop.
org/Nano/23/062001/mmedia) and analyzed by my groupand the association energy between two bR molecules
was estimated to be −1.5kB T (T = 300 K) per single
bond [81]. C-rings with 6.5 nm diameter were observed
to assemble into dimers with dynamic association distances
ranging from 8–12 nm (figure 10(a)). The dimers rarely
dissociated to monomers but were reformed after several
hundreds of milliseconds (figure 10(b)). The probability
distribution of the center-to-center distance of c-ring dimerp(d) was obtained from the AFM movie2 (figure 10(c)) andanalyzed by the Boltzmann law p(d) = p∞ exp[−U(d)/kB T],
where U(d) is the interaction energy and kB T is the

Membrane-mediated protein–protein interaction. Membraneproteins interact with the surrounding lipid molecules. This
interaction affects the structure, function and assembly of
the proteins. ‘Hydrophobic mismatch’ refers to the difference
between the hydrophobic thickness of the lipid bilayer andthe length of the hydrophobic region of the membraneembedded proteins [142]. This hydrophobic mismatch at theprotein–lipid interface alters the thickness and curvature of
the membrane to alleviate the mismatch. When two-protein
molecules are in close vicinity, they feel each other throughtheir perturbed membrane fields and then associate witheach other to further reduce the energy cost imposed by thehydrophobic mismatch [143].

2 Available

00900-8.
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thermal energy. The energy minimum of −3.5kB T occurred
at a center-to-center distance d = 10.3 nm (figure 10(d)).
This long-range attraction is likely to be caused by thehydrophobic mismatch. In fact, the hydrophobic thickness ofphosphatidyl glycerophosphate with a C17 acyl chain, a lipid
predominantly existing in the membrane, is ∼2.8 nm, whereas
the hydrophobic thickness of the c-rings is about 1 nm larger
than the hydrophobic thickness of the lipid bilayer (∼5.6 nm).
This presumption was further supported by the measurementof ∼1 nm elevated plateau of the lipid region between thec-rings.
5.2.2. Isolated proteins. Isolated proteins recently imagedby high-speed AFM are facilitates chromatin transcription (FACT) protein [77], cellulase hydrolyzing cellulose
fibers [83, 115], ATP-gated P2X4 receptor/channel [90],myosin V walking along actin filaments [91], acid-sensing
ion channel 1a under acidification [109] and α3 β3 subcomplexof F1 -ATPase (movie S11 available at stacks.iop.org/Nano/23/
062001/mmedia) [114]. Some of these studies are highlighted
here.
5.2.2.1. Intrinsically disordered FACT protein. There aremany proteins or large protein domains that lack a welldefined ordered structure under functional conditions [146,147]. These new class of proteins are called intrinsicallydisordered proteins (IDPs). IDPs take a structural ensemble
consisting of a large number of random conformers. Uponbinding to a target molecule, the disordered structure isoften converted to a defined ordered structure. In NMR, thelocal structure of IDPs can be determined but the overallrandom structure cannot be provided. Electron microscopy
cannot visualize the thin and random structures. Except forcomputational predictions, there was no way to identify theoverall random structure of IDPs.
High-speed AFM has recently demonstrated its ability
to visualize the thin and random structures of IDPs,
using FACT protein as a test sample [77]. Displacementof histone H2A/H2B dimers from nucleosomes by FACT
protein facilitates RNA polymerase II transcription [147]and chromatin remodeling [148]. FACT is a heterodimer
consisting of structure-specific recognition protein-1 (SSRP1)
and SPT16. Figure 11(a) shows AFM images of a FACT
oligomer with four ID regions that are extended from the main
body of the oligomer and undulating (movie S12 available
at stacks.iop.org/Nano/23/062001/mmedia). The monomer of
FACT protein showed two ID regions with different lengths(contour length: 17.5 and 27.5 nm on average). These ID
regions were identified by preparing mutants M1 and M2. In
M1, the predicted ID regions and the HMG-box domain of
SSRP1 were deleted, while in M2 the predicted ID region of
SPT16 was deleted. The longer ID region belongs to SSRP1
and the shorter one belongs to SPT16.
Interestingly, both ID regions showed a similar macroscopic flexibility (or stiffness). The stiffness of a polymer
chain is described by the persistence length p of the chain,that is, the length over which the average correlation in the

Figure 11. ID regions of FACT protein. (a) Successive high-speed
AFM images of oligomerized FACT protein. A lumpy shape andfour tail-like structures can be seen (see inset schematic). The
position of the tail-like structures fluctuates markedly over time.
The images were processed by using brightness-equalizing
software. Imaging rate, 6 fps; scan range, 150 × 150 nm2 .
(b) Persistence length analysis of ID region of M2 mutant (IDregion of SPT16 subunit is deleted). The square point-to-point
distance is plotted as a function of the macroscopic contour length
between the corresponding two points. The red line indicates thebest-fit curve of equation (1) with p = 12.1 nm. The inset image
indicates the relationship between the microscopic contour and themacroscopic structure. The microscopic contour is schematicallydrawn with a thin yellow line.

tangential direction decays by 1/e. In two dimensions, themean-square point-to-point distance of the chain is given by
hr2 (l)i2D = 4pl[1 − (2p/l)(1 − exp(−l/2p))],(1)

where l is the contour length between two points on the
chain [149]. In the AFM images obtained for the tail-likestructures of FACT, polypeptide chains are not resolved.
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Therefore, the persistence lengths that can be obtained
from the images are those for overall (macroscopic) string
structures. The analysis of AFM images of the ID regions
using equation (1) (figure 11(b)) gave a persistence lengthof 11–12 nm for both ID regions. Microscopic persistence
length Lp can be estimated based on an assumption that thepolypeptide chain length is a multiple of the number of aminoacids and the likely average distance between adjacent aminoacid residues (0.34 nm) and on the relationship hR2 i = 4Lp L(L, polypeptide chain length; R, 2D end-to-end distance). This
analysis gave a microscopic persistence length of ∼1.1 nm forboth ID regions (slightly shorter with the longer ID region,
likely due to the presence of the ordered HMG-box domain).
Microscopic measurements of persistence length havepreviously been performed by measuring force–extension
curves by AFM [150, 151], optical tweezers [152] andmagnetic tweezers [153]. The microscopic persistence lengthof the PEVK region (186 amino acid residues), which is
responsible for the elastic property of titin and is thought
to form a random coil, is reported to show a wide range ofLp (0.4–2.5 nm, on average approx. 0.93 nm) [151]. On theother hand, for proteins with ordered structures, the reported
values for the microscopic persistence length are within arange of 0.36–0.5 nm [150, 154]. Thus, the long microscopic
persistence lengths of the ID regions of FACT protein are
fingerprints of their very loose structures.
As demonstrated above, high-speed AFM is quite
powerful in identifying and characterizing ID regions ofproteins. In conventional methods, even only the identificationof ID regions is difficult and time-consuming. Moreover,high-speed AFM can detect order–disorder transitions of ID
regions (unpublished data) that occur naturally with some free
ID regions. In the future, high-speed AFM will visualize theprocess in which ID regions are folded into ordered structures
upon interacting with their target molecules and drugs.

proceeds as
A– M + ATP → A + M– ATP → A + M– ADP– Pi→ A– M– ADP– Pi → A– M– ADP → A– M,
where A and M represent actin and myosin head, respectively.
Immediately after binding to ATP, the actin-bound headdetaches from actin, and then the bound ATP is quicklyhydrolyzed to ADP–Pi. The ADP–Pi bound head has a low
affinity, yet a somewhat stronger affinity than the ATP-boundhead, for actin. When the ADP–Pi bound head is attached toactin, the bound Pi immediately dissociates from the head,
which is followed by the formation of a strongly boundcomplex of A–M–ADP. A nucleotide-free head also has astrong affinity for actin. These strong affinity complexes arecalled a ‘rigor complex’. Thus, a myosin head has two stateswith respect to actin binding, i.e. weak and strong boundstates. The main role of actin in the ATPase reaction is toaccelerate the otherwise very slow Pi and ADP dissociationfrom a myosin head. The degree of acceleration for ADP
dissociation varies depending on the class of myosin. The
processivity and non-processivity are determined by whether
or not the lifetime of A–M–ADP is predominant over the otherstates [161]. Double-headed myosins can be proteolyzed to
produce two single heads. A single-headed myosin binds toactin with a given angle relative to the actin filament. Except
for myosin VI, which uniquely moves towards the minus endof actin [162], the bound single head orients toward the plus
end of actin at least in the strong bound states. This orientation
towards the plus end of actin is called ‘arrowhead orientation’.
To observe M5 molecules walking along actin filaments,
the filaments have to be immobilized onto a substratesurface while M5 should be free from the surface. We
first used an electrically neutral phospholipid bilayer surface
containing biotin–lipid, formed on mica. To this surface,tail-truncated M5 (M5-HMM) were never bound, whereas
partially biotinylated actin filaments were immobilized
through streptavidin. However, in this condition, M5-HMM
mostly walked, standing perpendicular to the surface, andhence its molecular shape (sideways figure as illustrated
in figure 12(a)) could not be clearly imaged. To facilitate
the weak sideways adsorption of M5-HMM onto thebilayer surface, a positively charged lipid (1,2-dipalmitoyl3-trimethylammonium-propane (DPTAP), 5%) was includedin the bilayer. M5-HMM moving processively with ∼36 nm
steps was clearly visualized (figure 12(b), movie S13
available at stacks.iop.org/Nano/23/062001/mmedia). The
translocation velocity was similar to that measured byfluorescence microscopy when DPTAP was absent in the lipid
bilayer, indicating no effect of the tip–sample interaction on
the motor activity. The molecular process occurring during a
step, however, could not be resolved since a step completesvery fast.
Additional streptavidin molecules placed on the substratesurface as moderate obstacles to the advance allowed thevisualization of this process (movie S14 available at stacks.
iop.org/Nano/23/062001/mmedia). After the trailing headdetaches from actin, the leading head spontaneously rotatesfrom the reverse arrowhead orientation to the arrowhead

5.2.2.2. Walking myosin V. Motor proteins are highly
dynamic. They use chemical energy liberated by ATPhydrolysis to generate unidirectional force and motion. The
double-headed myosin V (M5), a member of the myosinsuper-family, moves processively along an actin filament,
towards the plus end of the filament (see the review [155]).
Facilitated by the processivity of the molecule [156, 157],
numerous single-molecule studies have demonstrated that it
moves ‘hand-over-hand’ with a 36 nm advance [158, 159]
for every ATP hydrolysis [160]. ‘Hand-over-hand’ means thatthe two heads step alternately, exchanging leading and trailingroles with each step, very much like ‘walking’. Nevertheless,the comprehensive descriptions of the walking behavior andthe chemomechanical coupling have not been attained until
our recent high-speed AFM observation of the molecule [91].
Before describing our observation results, let us
summarize established facts which are largely common to allclasses of myosins. A myosin head is comprised of a motor
domain and a neck domain. The motor domain contains twofunctional sites; the actin-binding and nucleotide-binding sites
which communicate with each other. The ATPase reaction
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Figure 12. High-speed AFM images of tail-truncated M5 (M5-HMM) interacting with actin filaments. (a) Schematic of two-headed bound
M5-HMM. (b) Successive high-speed AFM images showing processive movement of M5-HMM in 1 µM ATP. Frame rate, 7 fps; scan area,
130 × 65 nm2 . (c) Successive high-speed AFM images showing hand-over-hand movement in 1 µM ATP. Frame rate, 7 fps, scan area,
150 × 75 nm2 . The swinging lever is highlighted by a thin white line. (d) Schematic explaining the images in (c). (e) AFM images showing
unwinding of the short coiled-coil tail of two-headed bound M5-HMM observed in 50 mM ADP (left panel, before unwinding; right panel,
after unwinding). (f) An AFM image of two-headed bound M5-HMM captured under the nucleotide-free condition. The leading head is
sharply bent. (g) An AFM image of two-headed bound M5-HMM captured in the presence of 50 mM ADP. The leading head isapproximately straight. Vertical or horizontal dashed lines in (b), (c), (f) and (g) show the centers of mass of the motor domains, and a plus
sign is used to indicate the plus end of actin.

orientation (figures 12(c) and (d)). This rotation is theswinging lever arm motion itself initially proposed by
Huxley for muscle myosin [163], which had never beenclearly demonstrated before. The detached trailing headrotates around the advancing neck–neck junction. During therotation, the head becomes distant from and then gets closer
to the actin filament. Finally, it attaches to a forefront actin,
completing one step.
The spontaneous rotation of the leading head following
the trailing head detachment suggests that intramoleculartension for the advancement has already existed at theleading head of the two-headed bound molecule. This
supposition was directly demonstrated by the observationthat the short coiled-coil of two-headed bound M5-HMM
was sometimes unfolded and then the leading head rotatedfrom the reverse arrowhead orientation to the arrowheadorientation (figure 12(e), movie S15 available at stacks.
iop.org/Nano/23/062001/mmedia), very much similar to theswinging lever arm motion during stepping. Thus, we canderive the following conclusions. (1) By nature, both heads
most stably bind to actin in the arrowhead orientation but theleading head has to bind in the reverse arrowhead orientation
because it is linked to the trailing head at the neck–neckjunction and a favorable actin site is already occupied by
the trailing head (otherwise the molecule would be forced
to twist around the actin filament, paying more energy cost).
Thus, intramolecular tension for advancement is generated
at the leading head. (2) The detachment of the trailing headremoves this constraint imposed to the leading head, thus
resulting in the rotation of the leading head. (3) Therefore, thisrotation spontaneously occurs without chemical transitions in
the ATPase reaction.
It was further observed that the leading head of thetwo-headed bound molecule was briefly detached from actin
and rebound to actin without stepping (‘foot stomp’), and thenthe molecule stepped forwards (movie S16 available at stacks.
iop.org/Nano/23/062001/mmedia). As Pi is quickly released
when M5–ADP–Pi is initially bound to actin, the leading head
only contains bound ADP before and after foot stomping.
Nevertheless, the molecule can step forwards, indicating that
intramolecular tension for forward movement is generated
without transitioning through Pi release.
Moreover, it was observed that the leading head of thetwo-headed bound M5-HMM was often sharply bent in the
nucleotide-free condition (figure 12(f), movie S17 available atstacks.iop.org/Nano/23/062001/mmedia), while it was mostly
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straight in ADP and ATP (figure 12(g)). Therefore, just bylooking at the shape of the leading head, we can judge whether
or not the leading head contains nucleotides. We imaged
two-headed bound M5-HMM in the presence of various
concentrations of ADP to estimate the ADP dissociation
rate constant at the leading head. The rate came out to be
0.1 s−1 . This means that ADP is released from the leadinghead every 10 s, on average. However, M5-HMM walks many
steps for 10 s. Thus, we can conclude that during walking
ADP does not dissociate from the leading head (before ADP
dissociation, the leading head is switched to the trailing
head). ADP dissociation, and the subsequent ATP binding,and the resulting detachment from actin solely occurs at thetrailing head. This is the basis underlying the hand-over-handmovement. This mechanism had been suggested from indirect
experiments [160, 164–166] but is now clearly and directly
demonstrated by the high-speed AFM observation.
Just before foot stomping at the leading head, the head
never showed the sharply bent conformation which is uniqueto the nucleotide-free leading head. This fact reinforces ourconclusion that the leading head performing foot stomp
carries ADP and thus the brief detachment from actin (i.e. theinitial stage of foot stomp process) is not caused by binding of
new ATP to the leading head.
As seen above, many dynamic events of M5-HMM
appear in the molecular movies, facilitating a quick findingof several facts including those inaccessible with conventional
approaches.
5.2.2.3.

TrCel7A on a crystalline cellulose surface was only observedat >10 times higher concentrations, indicating a low affinityof the CD alone for cellulose. However, it moved at a velocity
similar to that of the intact wild-type molecule, indicating that
the CBD is not required for the movement. Since TrCel7A is asmall molecule, its molecular conformation was not resolvedby the AFM observation, and hence elucidating how themolecule changes its conformation during the processive run
remains a challenge.
P2X4 receptor in response to ATP. P2X receptors are
ATP-gated membrane cation channels and have seven
identified sub-types of subunits (P2X1 –P2X7 ) [171]. Eachsubunit has two transmembrane domains separated by alarge extracellular domain capable of binding to ATP andsmall intracellular domains at the N and C termini [171].
Channels form as multimers (thought to be mainly trimers)of subunits; homometric or heteromeric channels. All P2X
receptors are permeable to small monovalent cations andsome have significant Ca2+ or anion permeability [172].
P2X receptors are abundantly distributed and involved indiverse physiological processes from synaptic transmission
to inflammation and sensation of taste and pain [172].
Crystal structures of the zebrafish P2X4 receptor in theclosed-channel state are recently obtained (figures 13(a)and (b)) [173]. The three ATP binding sites are formed at
the intersubunits in the trimer, suggesting that ATP binding
promotes subunit rearrangement and ion channel opening.
The ion channel opening of P2X4 receptor is regulated by
the extracellular Ca2+ level [174]. In a high concentration of
Ca2+ , P2X4 receptor opens a small cation-permeable channel
pore, whereas in the absence of Ca2+ , it forms a large poreallowing permeation of large cationic molecules.
Shinozaki and collaborators imaged ATP-induced conformational changes of P2X4 receptor at 2 fps using two
assay systems [90]; isolated (membrane-dissociated) P2X4
attached to polylysine-coated mica and P2X4 reconstructedin a lipid bilayer. In both systems, asymmetric molecules
were observed. In the former system, the asymmetry might
be due to a leaned orientation of the receptor on the substratesurface. In the latter system it could not be specified. In theabsence of ATP, particles observed had a globular shape ora slightly triangular shape, whereas in the presence of ATP
under no Ca2+ , three subunits were resolved together with acentral pore. Interestingly, a pore dilation trimer structure wasrevealed, whereas in the presence of Ca2+ , no pore dilation
was observed. Since symmetric molecules were not observed,
they used a single-particle correlation averaging method toreconstruct threefold-symmetric images after a given elapseof time. The symmetrized images showed the central pore
being dilated with time after the addition of ATP underno Ca2+ (figure 13(c)). Since even the membrane-attachedreceptor shows non-symmetric structures, this non-symmetry
may arise from fluctuations of the subunit arrangement in thetrimer. In addition, it still remains questionable whether or
not the asymmetric trimer structures with pore dilation are an
intrinsic nature of this purinergic receptor.

Other dynamic processes of isolated proteins.

Cellulase hydrolyzing cellulose fibers. There are several
types of cellulases that hydrolyze cellulose fibers. Cellobiohydrolases (CBHs) are exocellulase that cleaves glycoside
linkages of crystalline cellulose I either from the reducing
or nonreducing end of cellulose to produce tetrasaccharides
or disaccharides [167, 168]. Trichoderma reesei (Tr) Cel7A
composed of a small cellulose-binding domain (CBD) and alarger catalytic domain (CD) is a member of CBHs. Cel7ahydrolyzes cellulose I from its reducing end to produce a
β-1,4-glucose dimmer [169]. The two-domain structure iscommon to glycosidases degrading insoluble carbohydrate
substrates. From biochemical kinetics analyses, TrCel7A isthought to consecutively hydrolyze the crystalline cellulose
chain without releasing the chain, and therefore is expected
to processively move in one direction during the hydrolysis
reaction [170]. Nevertheless, this long-standing issue had
never been directly revealed before the high-speed AFM
observation recently conducted by Igarashi et al [83] andby the collaboration between my group and Igarashi’sgroup [115].
From the biochemically measured hydrolysis rate,
TrCel7A was expected to move on a crystalline cellulose
fiber at 1 nm min−1 . In marked contrast with this value,
TrCel7A moved at 3.5 nm s−1 , ∼200 times faster than
expected. This large discrepancy suggests that a large numberof molecules cannot smoothly hydrolyze cellulose becauseof traffic problem [115]. The movement of CBD-truncated18

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Figure 13. Structure of P2X4 receptor. (a) Atomic structure of N and C termini deletion mutant of homotrimeric P2X4 receptor viewed
parallel to the membrane. Each subunit is depicted in a different color. The gray bars suggest the boundaries of the outer (out) and inner (in)leaflets of the membrane bilayer. (b) Atomic structure of N and C termini deletion mutant of homotrimeric P2X4 receptor viewed from themolecular threefold axis (from the extracellular side) (Reprinted with permission from the Nature Publishing Group [173]). (c) Successive
AFM images showing ATP-induced structural changes of P2X4 receptor. Before activation, P2X4 receptor was in circular shape andexhibits some fluctuation (−2.5 to ∼0.0 s). Caged ATP (200 µM) was uncaged at 0 s. After uncaging, the structure of P2X4 receptor
changed to a trimer structure within 0.5 s. Then, P2X4 receptor exhibited a further structural change and adopted a pore dilation-like
conformation. Note that the images are not directly captured ones but are those obtained by averaging images of ten P2X4 receptor particles
to reconstruct threefold symmetric images at a given elapse of time.

5.2.3. Protein self-assembly processes.
Thus far, this
topic other than 2D protein crystal formation has not beenstudied in depth using high-speed AFM, except for the studyof amyloid-like fibril formation by N-terminus-truncatedlithosthatine, conducted by the collaboration between theMilhiet group and my group [99]. The polymerization
processes of actin and tubulin, which are briefly studied, will
be described in section 5.2.5.
Amyloids are insoluble fibrous protein aggregates andtheir deposition in tissues causes diseases, particularly neurodegenerative diseases [175, 176]. However, the mechanismand dynamics of fibrillization have been elusive. Lithosthatine(also named Reg-1) of 144 amino acids is produced bypancreas acinar cells and secreted into pancreatic juice. Ittightly binds calcium carbonate and may act as an inhibitorof calcium carbonate precipitation to prevent clogging ofpancreatic duct [177, 178]. In chronic calcifying pancreatitis,the protein forms deposits in the pancreatic duct [179].
Lithosthatine is also expressed in other organs including
stomach and brain [180]. Lithosthatine is susceptible toself-proteolysis under specific pH conditions, producing asoluble N-terminal undecapeptide and a C-terminal fragmentof 133 amino acids. The cleaved protein (called S1)
forms protease-K-resistant fibrils [181] that deposit in thebrain of patients with Creutzfeld–Jakob disease [182] andGertsmann–Straüssler–Scheinker syndrome [183], especiallyduring the very early stages.
The S1 form of lithosthatine was generated by tryptic
digestion of the full-length protein. After 15 min incubation,
globular structures with 15–28 nm in diameter appeared. They
appear to be small assemblies of lithosthatine tetramers with
9 nm in diameter and 2.5 nm in thickness. After 30 min
incubation, protofibril formation and its growth to longer
protofibrils and thicker fibrils were observed (figure 14).
Lengthening of protofibrils takes place at 27–52 nm s−1 .
Elongation of protofibrils occurs in two ways; (1) additionof lithosthatine oligomers to the ends of protofibrils (whitearrows in figure 14) and (2) fusion of a growing protofibril
to a pre-existing protofibril (light-blue arrows in figure 14).
Interestingly, the end of the pre-existing protofibril to be
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Figure 14. High-speed AFM images showing the time lapse of lithosthatine protofibril elongation and association. Elongation of single
protofibrils is shown by white arrows in (a), (b) and (d). Some protofibrils do not grow in length, as indicated by white arrowheads in (a) and(b). Interestingly, the edge of an elongating protofibril can be fused with that of a pre-existing protofibril, forming a continuous structure, as
marked with a light-blue arrow in (b). Protofibrils are often laterally associated with another, at least for one part of their length. The
complete sideways association between two protofibrils occurs in a zipper-like mechanism (pink arrows). Elongation of a protofibril also
occurs, being stacked on a fibril, as indicated with the light-blue arrowheads in (c). The growth of a protofibril or fibril is stopped when thegrowing end encounters a side edge of a second fibril, forming a false branching point with a certain range of contact angle, as indicatedwith red arrowheads in (d). Winding of two protofibrils can form helical fibrils (e) (the inset is a filtered image). Scale bars, 30 nm.

encounters a side edge of a second fibril, forming a false
branching point with a certain range of contact angle (redarrowheads in figure 14(d)). Thus, the dynamic visualizationof lithosthatine fibrillization revealed various elementary
processes contained in the fibrillization.

fused with the growing protofibril does not change its
position (white arrowheads in figure 14(b)), which may
suggest a marked difference in the oligomer addition rate at
the two ends of a protofibril. Elongation of fibrils appears
to occur separately at each constituent protofibril. In fact,a protofibril is elongated on top of laterally associatedprotofibrils (light-blue arrowheads in figure 14(c)). The lateral
association of protofibrils to fibrils takes place in a zipper-likemechanism (pink arrows in figure 14(b)). In the initial stage,
two protofibrils make contact with each other in a small
region, showing a Y shape. Then, the distance between
the separated protofibrils is getting closer to each other by
gradual increase of the contact region, and eventually thetwo protofibrils are completely associated to form a helicalfibril similar to other amyloid ß fibrils (figure 14(e)). It
is also clear that a branched structure previously observedwith lithosthatine fibrils is a false structure. The growthof a protofibril or fibril is stopped when the growing end

5.2.4. DNA-binding proteins and synthetic DNA devices.
Regarding DNA–protein interactions, there are a wide rangeof subjects to be studied by high-speed AFM imaging but
only a few studies have thus far been performed; reaction by
methyltransferase restriction enzymes [84, 104], unwrappingof DAN–histone complex [108] and reaction by DNA baseexcision repair enzyme [103]. Synthetic DNA nanodevices
are also interesting targets of dynamic AFM imaging.
The specific interactions between complementary base pairs
make DNA a useful building material for constructing twoand three-dimensional nanostructures [184]. Various planned
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incomplete migration of the motor between a cut stator and an
adjacent intact stator. The irreversible migration to an adjacentstator appears as transitional shifts of the motor position.
These dynamic processes are more directly revealed thanprevious approaches. However, the spatiotemporal resolutionof the movie data was not high enough, so the expectedsequential events contained in the motor migration, suchas restriction enzyme binding, digestion of a motor-boundstator and intermediate states prior to stepping of the motor,
and branch migration, were not well resolved. More detailed
molecular events could be visualized if the arrangement of
the stators was altered from the vertical to the horizontal
orientation and the imaging rate is increased.

Figure 15. DNA motor movement along the track of DNA stators
formed on a DNA origami tile. (a) Motor mechanism: a nicking
enzyme (triangles) cuts the stator (green) when the stator is boundto the motor (magenta), revealing a toehold at the 30 end of the
motor that facilitates transfer of the motor to the adjacent intact
stator by branch migration. This chain of processes is repeated andthus the motor continuously moves along the track of stators until
reaching the track end. (b) Successive AFM images showing
discrete steps of a single DNA motor. The highest point on eachprofile along the yellow line is interpreted as the motor position andmarked by the red circle. The motor steps between four stators (bluelines). Imaging rate, 0.1 fps; vertical scale bar, 50 nm. Reprinted
with permission from the Nature Publishing Group.

5.2.5. Manipulation and imaging.
The cantilever tip canalso be used to manipulate the sample. Changes in the sample
after the manipulation can be traced by imaging using thesame tip. The use of high-speed AFM in this way has been
attempted in a few studies; formation of vacancy defects
in streptavidin 2D crystals and subsequent observation ofBrownian motion of the defects (as already described in
section 5.2.1.2) [76], breaking actin filaments in the presenceof G-actin and subsequent observation of repolymerization
processes [82], breaking microtubules in the presence of
tubulin and subsequent observation of repolymerization
processes (unpublished), and removal of magnetosomes froma substrate surface and subsequent observation of molecules
transferred from magnetosomes to the surface [94]. Amongthese studies, a study on actin polymerization is briefly
described below.
Actin filaments have a structural polarity, so that thepolymerization kinetics is different at the two ends of the filaments; a higher rate at the barbed end than at the pointed end.
Previous fluorescence microscopy studies on the polymerization dynamics with individual actin filaments have suggested
that a polymerization unit is not a monomer alone but anaverage of monomers, oligomers and fragments [187, 188].
However, analysis of precise molecular events occurring at
the filament edges was difficult due to the limited spatial
resolution.
In the high-speed AFM observation, partially biotinylated
actin filaments were attached to a surface of streptavidin
2D crystals formed on biotin–lipid bilayers in the presenceof non-biotinylated G-actin in the bulk solution. Instead offinding actin filament ends, they were produced by breaking
an actin filament by applying an increased tapping force.
This manipulation allows simultaneous visualization of bothbarbed and pointed ends in a relatively small imaging area.
After breaking, elongation was only observed at one of the
newly created ends in the observation period and its rate was
0.11 µm min−1 , which is comparable to the values obtained
previously by fluorescence microscopy. The elongation step
size is distributed around two values (3.1 and 5.2 nm),indicating monomer and dimer addition to the barbed end
in light of the axially staggered two-stranded structure.
Interestingly, a stepwise decrease (3.5 ± 1.4 nm) in thelength was occasionally observed, indicating that monomer
dissociation is predominant. Moreover, the stepwise decrease

structures can actually be constructed by rationally designingbase sequences of DNA. This technique has been used toprepare DNA origamis [89, 107, 117], capsules [110] andnanorobots [118]. Here, a study on dynamic imaging of ananorobot is taken up.
Dynamic imaging was conducted for a DNA nanorobot in
the collaboration between Sugiyama’s group and Turberfield’sgroup [118]. Before this study, several types of DNA
nanorobots with or without an autonomous motility andprocessivity have been synthesized including bipedal walkers
and rotary gears [185, 186]. However, the motion of thesenanorobots has been detected in indirect ways and therefore
mechanistic details of their operation have remained elusive.
The nanorobot they constructed consists of an extended
track formed on a DNA origami tile (∼70 nm × 100 nm) anda motor DNA. The track consists of a one-dimensional arrayof single-stranded attachment sites (stators) with the 30 end at
the bottom, separated by ∼6 nm. The motor is a single strandof DNA that is complementary to a stator. A motor–stator
duplex contains the recognition site for a restriction enzyme(Nt.BbvCl) that hydrolyzes a specific backbone linkage ofthe stator. The motor translocation is fueled by the DNA
hydrolysis. When the motor DNA hybridizes to a stator strand
to form a double strand, the enzyme cuts the stator andremoves a short segment from the stator. Therefore, at the 30
end of the motor a toehold for binding to the adjacent intact
stator is revealed. After the actual binding of the toehold,the motor completely dissociates from the nicked stator andcompletely associates with the neighboring intact stator bybranch migration (figure 15(a)). This process is repeated
and thus the motor moves processively in one direction
along the stator track. The film captured by high-speed AFM
at 0.1 fps showed that the motor DNA migrates forwards
with discrete steps of ∼7 nm and dwell times of ∼50 s(figure 15(b)) but often moves back and forth between
two adjacent stators. This back-and-forth movement suggests
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Topical Review

in the length often occurred after monomer addition. The
end structure is the same for both cases of monomer anddimer addition. Nevertheless, monomer dissociation more
frequently occurred after monomer addition. This suggeststhat the added monomer possesses ATP while the added dimer
possesses ADP.
The approach of manipulation followed by imaging will
have a wide variety of applications. As exemplified above,
self-assembly processes can be studied by tracing re-assembly
processes after breaking an assembled structure. One ofthe other useful applications is to examine a complicatedmolecular architecture of a large protein with several domains
or a molecular assembly with different subunits. Thisapproach is particularly useful when its structure projected
to a 2D plane is known from the electron micrographs
but its 3D structure is unknown. Another application is to
examine the cooperative interaction between subunits of aprotein. For example, ring-shaped AAA proteins, which areoften hexamers of identical ATPase subunits, are supposed
to show large structural changes during ATP hydrolysis
via cooperative interplay between subunits. The cooperative
structural changes will be abolished by removing one or two
subunits from the ring. This approach was recently utilized
to reveal the cooperative interplay between β subunits in theα3 β3 subcomplex of F1 -ATPase (a pseudo-sixfold-symmetricring) [114]. The cooperative structural change in β propagatedin the counterclockwise direction during ATP hydrolysis but
the occasional subunit dissociation completely stopped therotary propagation of the conformational state.

(3) Imaging of dynamic structural changes of molecules inaction. Typical examples in this class are studies on
walking myosin V along an actin filament [91], photoinduced structural change in bR [92, 113] and rotationallycatalyzing α3 β3 subcomplex of F1 -ATPase [114].
Thus, there are two types of dynamics studied byhigh-speed AFM; ‘interaction dynamics’ and ‘structuraldynamics’. In the former type of studies, structural dynamics
of individual molecules would have allowed more detailed
analysis of the observed interaction dynamics. The lack ofstructural information at the sub-molecular level arises fromthree factors. The first factor is the insufficient sharpness of a
cantilever tip. The tip apex radius of commercially available
small cantilevers is ∼15 nm, insufficient for sub-molecularresolution imaging. Small cantilevers with EBD tips are alsocommercially available but expensive. The apex radius ofan EBD tip is often 4–5 nm, still insufficient for resolving
sub-molecular structures of small protein molecules, althougha ∼0.5 nm radius can sometimes be obtained. So, smallcantilevers with carbon nanotube tips with an apex radiusof ∼1 nm are desired, but the procedure is laborious andtherefore cannot be used routinely. We need to explore new
methods and different materials for fabricating sharp andrigid EBD tips. The second factor is Brownian motion ofmolecules, particularly in isolated protein molecules. This is aproblem intrinsic to AFM imaging of dynamic events because
dynamic AFM imaging requires freedom of movement tomolecules to some extent. In some cases, the reduction of
the Brownian motion can be achieved by devising appropriate
substrate surfaces, as has been done for imaging walking
myosin V molecules [91]. The third factor is the orientationof the sample relative to the substrate surface. AFM imaging
can only be done from the direction perpendicular to thesubstrate surface. When a molecule stands up on the substratesurface, we cannot see the sideways figure of the molecule.
In some cases, a molecule can be placed on a substratesurface in a desired orientation by preparing appropriate
substrate surfaces, as has been done for imaging the sideways
figures of walking myosin V molecules [91]. It can also
be done by introducing a functional group to a specificlocus of a molecule and by preparing a substrate surface
able to bind to the functional group, as demonstrated in theobservation of the sideways figure of GroEL molecules [82]and the C-terminal top surface of the α3 β3 subcomplex of
F1 -ATPase [114]. Dynamic AFM imaging of biomolecular
processes imposes various requirements on ‘wet techniques’.
This issue is described elsewhere [82, 93].
Various conditions are required for successful dynamic
AFM imaging. Nevertheless, once successful, acquired
molecular movies are striking. Various dynamic events
appear unselectively and simultaneously in a molecularmovie, which facilitates quick finding of several facts
including those inaccessible with conventional approaches.
Thus, the high-speed AFM imaging of functioning biologicalmolecules has the potential of transforming the fields ofstructural biology and single-molecule biology. In addition,
this new microscopy is applicable to the observation of nonbiological dynamic phenomena occurring at the solid–liquid

5.3. Summary of high-speed AFM imaging studies
As described above, dynamic AFM imaging has already been
conducted for a variety of samples. About half of the studies
in the last 3 years seem to be aimed at exploring the capabilityof this new microscopy and groping for effective applications.
This situation is quite similar to that in the earlier bio-AFM
studies after the advent of AFM in 1986. The dynamic AFM
imaging studies carried out thus far can be classified according
to the information obtained.
(1) Imaging of dynamic events occurring in the interaction
between molecules, in which the observed structure ofindividual molecules has no importance in the derived
conclusion. Typical examples of this class are the studies
on membrane-mediated protein–protein interaction [101]and dynamic interaction of isolated bR trimers with theedge of bR 2D crystals [81].
(2) Imaging of dynamic events occurring in the interaction
between molecules, in which the structure of individual
molecules is not well resolved. Typical examples ofthis class are the studies on unidirectional translocationof cellulase along cellulose fibers [83, 115] andtransportation of synthetic DNA motor along a series of
DNA stators [118]. The study on translocation of EcoRII
restriction enzyme along DNA [74] also belongs to thisclass.
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Nanotechnology 23 (2012) 062001

Topical Review

interfaces [96, 97, 112], including electrochemical reactions,corrosions, catalytic reactions and cleaning with detergents.
This microscopy will, therefore, facilitate creation of new ‘wet
nanotechnology’.

coatings over nitride trench walls and malaria in red blood
cells. A high-frequency acoustic wave (frequency f1 ) is
launched from under the sample stage, and this wave thenpropagates through the sample. The propagated ultrasoundcan be detected by the tip as well as by the lever of a
cantilever with a resonant frequency of fc . To ensure thenear-field detection by the tip and hence high resolution,
another acoustic wave with a frequency f2 (|f1 – f2 | ∼ fc ) is
launched from the cantilever base so that nonlinear tip–sample
interaction generates an acoustic wave with a frequency|f1 – f2 | and therefore the cantilever is efficiently excited by
the generated acoustic wave. According to their interpretation
for the sub-surface imaging, materials embedded in thesample with different elastic modulus/density ratios modulate
the phase and amplitude of the propagating acoustic wave,
and hence the cantilever tip detecting the acoustic wave
senses the embedded materials. However, it is very likelythat embedded materials affect the elasticity of the sample
surface and this effect seems to be detected by the cantilevertip. In fact, when materials are embedded deeply from thesample surface (>1 µm; much shorter than the wavelength ofultrasound with a few MHz frequency), no sub-surface image
appears in the acoustic phase image (unpublished data). It has
been demonstrated that SNFUH has a capability of imaging
artificial solid particles introduced in cells [190]. However,the live cell membranes are extremely soft and therefore it islikely that the cantilever tip deeply pushes the cell membrane
to sense the solid particles.
Because of this likely indirect detection of shallowly
embedded materials in the contact mode, SNFUH will not
be widely used to image intracellular structures. Alternativetechniques need to be developed to materialize a true
imaging of the interior of live cells at high resolution. One
possibility is to increase the spatial resolution of scanning
acoustic microscopy (SAM) that can image the interior ofcells with low resolution (∼ a few micrometers), under the
non-contact condition [191]. Once an operational principle
for high-resolution and high-contrast detection of acoustic
waves in SAM is established, it will open up an avenue
to materialize high-speed diaphano-AFM not for observing
individual molecules but for observing intracellular organelles
and cytoskeletons in live cells. The non-contact condition
can remove feedback operation and thus will make ultrafast
imaging possible. We are currently attempting to find a way toenable high-resolution and high-contrast detection of acoustic
waves in SAM.

6. Future challenges
The current high-speed AFM is powerful in visualizing
dynamic events of isolated and purified biological molecules
but not applicable to the dynamic observation of the outersurface and the interior of live cells (i.e. in situ imaging).
Solutions for these challenges and the improvement of thecurrent high-speed AFM are now being developed.
6.1. Small cantilevers
The feedback bandwidth of the current high-speed AFM
apparatus exceeds 100 kHz, and consequently an imaging
rate of 25 fps is achieved under the condition of 200 nm
scan size and 100 scan lines. Can we go beyond? The main
device limiting the speed is the small cantilevers. Generally,
to improve the resonant frequency, one must compromise
the stiffness and vice versa. The most advanced smallcantilevers developed by Olympus are deemed to have almost
achieved the ultimate goal of balancing these two mechanicalquantities. Considering their practical use together with the
OBD detector, doubling the resonant frequency to ∼2.4 MHz
in water seems to be the upper resonant frequency limit,
provided the cantilever compliance is not sacrificed. Thus, theupper limit of the imaging rate is approximately 40 fps under
the same condition as above.
6.2. Scanner
The maximum scan range of a scanner employed in thecurrent high-speed AFM apparatus is limited to 1–4 µm inthe x and y directions and ∼1 µm in the z direction. The scan
range in the x and y directions has to be increased at least
ten-times and that in the z direction has to be increased >5times for imaging cells. As a matter of course, the highest
possible imaging rate is reduced. However, the reduction in
the speed of x-scanning may not be so significant because thebandwidth of the x-scanner can be increased by an inverse
compensation method [7]. After slowly imaging a wholecell surface, a selected small area of the cell surface can beimaged quickly using the same scanner. This imaging can bedone for bacterial cells but not for mammalian cells because
mammalian cells are extremely soft.

6.3.2. Non-contact high-speed AFM.
As mentioned
above, mammalian cells are extremely soft, so that they
are largely deformed by the contact with a cantilever tip
before detectable cantilever deflection appears. Therefore,
interesting dynamic molecular processes occurring on thelive cell surface cannot be imaged except for small cellular
structures such as dendritic spines. Thus far, a true non-contact
condition is only achieved under vacuum environments using
frequency-modulation (FM) AFM [192, 193]. In liquids
FM–AFM has been used but can only operate in the brief
contact condition. In addition, the oscillation amplitude of

6.3. Next generation of high-speed AFM
6.3.1. High-speed diaphano-AFM. An exciting report was
published in Science in 2005 by Shekhawat and Dravid [189].
They developed scanning near-field ultrasound holography(SNFUH) for high-resolution sub-surface imaging in air.
They presented sub-surface images such as gold nanoparticles
embedded in a polymer film, voids embedded in polymer
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Topical Review

a cantilever has to be .0.2 nm to achieve high detection
sensitivity of the tip–sample interaction, inapplicable to asurface with large corrugations.
When the sample stage is vibrated in the z direction at ahigh frequency, with small amplitude, this vibration transmits
to the solution placed on the stage. This transmission pattern
may depend on the sample and therefore the sample may
be able to be detected by a cantilever. However, even in thenear-field regime, both tip and lever of the cantilever sense
the water vibrations. The lever senses more efficiently than
the tip. The tip is aligned in parallel to the direction in which
the vibration transmits, leading to low sensitivity. Thus, thisapproach will not grant single-nanometer resolution in thelateral direction.
The non-contact imaging capability in liquids has
already been achieved by ion-conductance scanning probe
microscopy (ICSPM) [194]. Owing to the progress infabrication techniques for producing very sharp glass
capillaries with a small pore at the apex, the spatial resolutionof ICSPM has reached a few nanometers [195]. Immobile
protein molecules with a size of ∼14 nm on living cell
membranes have been successfully imaged [196]. However,it seems difficult to increase the imaging rate of ICSPM;
the bandwidth of ion-conductance detection cannot easily be
increased, because of the high impedance to ionic current
through the small pore of the capillary electrode.
Thus far, there are no potential methods to visualize
dynamic molecular events on extremely soft mammalian cellsurfaces. An alternative and compromising way may be toattach live cells with a thin film having small pores of∼100 nm diameter. Cell surfaces exposed to the pores are
rigid and therefore can be imaged.

of the high-speed bio-AFM consortium, P Hinterdorfer,
S Scheuring, P-E Milhiet, Ch Le Grimellec, E Lesniewska
and Y Lyubchenko, for shearing ideas and scientificinterests. Long-term financial support by NEDO, JST, MEXT,
Knowledge Cluster Initiative and the Mitsubishi Foundation is
gratefully acknowledged.

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7. Conclusion
To directly observe biological molecules at work was a holygrail in biology. Efforts over the last two decades at lastmaterialized this long-quested dream. In high-resolution AFM
movies, we can see how molecules are dynamically behaving,changing their structure and interacting with other molecules,
and hence we can quickly understand in stunning detail how
molecules operate to function. This new approach will spreadover the world and widely applied to a vast array of biologicalissues, leading to a number of new discoveries. The extensionof high-speed AFM to a tool for imaging live cells, whichallows direct in situ observation of dynamic processes ofmolecules and organelles, remains an exciting challenge but
will be made in the near future because it is a right and fruitfulgoal.

Acknowledgments
I would like to thank T Uchihashi, N Kodera, M Shibata,
D Yamamoto, H Yamashita, H Kandori and previous many
students in my lab for collaborations and discussions
on interesting topics I described in this review. Without
their dedication, high-speed bio-AFM would not have beenmaterialized. I would also like to thank the members
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