Reprinted from

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High-Speed Atomic Force Microscopy
Toshio Ando, Takayuki Uchihashi, and Noriyuki Kodera

Jpn. J. Appl. Phys. 51 (2012) 08KA02# 2012 The Japan Society of Applied Physics

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http://dx.doi.org/10.1143/JJAP.51.08KA02

High-Speed Atomic Force Microscopy
Toshio Ando1;2 , Takayuki Uchihashi1;2 , and Noriyuki Kodera2
1
2

Department of Physics, Kanazawa University, Kanazawa 920-1192, Japan
Bio-AFM Frontier Research Center, Kanazawa University, Kanazawa 920-1192, Japan

Received January 11, 2012; accepted May 7, 2012; published online August 20, 2012
The technology of high-speed atomic force microscopy (HS-AFM) has reached maturity. HS-AFM enables us to directly visualize the structure anddynamics of biological molecules in physiological solutions at subsecond to sub-100 ms temporal resolution. By this microscopy, dynamicallyacting molecules such as myosin V walking on an actin filament and bacteriorhodopsin in response to light are successfully visualized. Highresolution molecular movies reveal the dynamic behavior of molecules in action in great detail. Inferences no longer have to be made from static
snapshots of molecular structures and from the dynamic behavior of optical markers attached to biomolecules. In this review, we first describe
theoretical considerations for the highest possible imaging rate, then summarize techniques involved in HS-AFM and highlight recent imaging
studies. Finally, we briefly discuss future challenges to explore. # 2012 The Japan Society of Applied Physics

1. Introduction

The function of proteins is tightly related to their structures,and thus, structural biologists have been determining theirdetailed structure. However, the obtained structures are
essentially static, whereas proteins are dynamic in nature.
The dynamic behavior of proteins in action has been mainly
studied by single-molecule ﬂuorescence microscopy.1,2) A
ﬂuorescent spot emitted from a ﬂuorophore attached to a
speciﬁc locus of a protein is recorded on video and the time
sequence of its position (or polarization angle) is analyzed inhigh precision. Nevertheless, protein molecules themselves
are invisible in such observation. Hence, the simultaneous
observation of the structure and dynamics of proteins haslong been infeasible. We have to infer how proteins operate
to function from gleaned data with signiﬁcant resolution
gaps. The inference is often aﬀected by assumptions andinterpretations of the data, and therefore diﬀerent conclusions are sometimes derived from the same experimental
data. We often need many years to reach a convincing
conclusion on the functional mechanism of a protein. Thus,
it is evident that we should develop more direct methods thatcan provide great detail of molecular action without intricate
analyses and interpretations.
Atomic force microscopy (AFM) is unique in its
capability of directly capturing high-resolution images ofvital biological samples in aqueous solutions.3,4) AFM wasthus expected to play a role in bridging the gap between
structural biology and single-molecule biophysics. However,
AFM was not able to meet this expectation because of its
low imaging rate. The visualization of a sample by AFM
requires the acquisition of sample height information overmany points on the sample surface (see Fig. 1). During theacquisition at each point, the following operations arecarried out. A stylus probe attached to the free end of
a cantilever is brought into contact with the sample. The
mechanical response of the cantilever upon this contact(change in, e.g., deﬂection, oscillation amplitude, oscillation
phase, or resonant frequency, depending on the operation
mode of AFM used) is measured and then the sample stage
is ﬁnally moved in the z-direction to recover the mechanicalstate of the cantilever back to a given condition (i.e., set
point) through feedback control. For this recovery, theclosed feedback loop spends a certain amount of time mainly
because of the slow response of the mechanical devices (i.e.,cantilever and z-scanner). This series of operations is

Fig. 1. (Color) Schematic for general conﬁguration of tapping-mode
AFM system.

repeated many times for diﬀerent sample surface points
during the lateral scanning of the sample stage. Thus, it takes
a long time (at least 30 s) to capture an image of the sample.
Molecules moving on the substrate surface are, therefore,
blurredly imaged or cannot be imaged at all.
Hansma’s group and our group independently started
around 1993 to develop high-speed AFM (HS-AFM) in thetapping mode. Various devices and techniques, such as smallcantilevers,5–10) high-speed scanners,8,9,11–16) active damping16,17) and feedback control16,18) techniques, and electronics8,9,12,13,16–21) have been developed. After a proof-ofprinciple stage, the technology of HS-AFM has reachedmaturity.22–24) Although the speed performance depends on
imaging conditions, it can generally capture an image of aprotein molecule within 100 ms or less. Importantly, thestructure and function of fragile proteins are not disturbed
by the interaction with an oscillating cantilever tip.22–24) This
high-speed and low-invasiveness performance provides a
new opportunity to visualize dynamically functioning
protein molecules in great detail. As has been demonstrated
in recent imaging studies,22–32,34) the visualized dynamic
images of proteins can provide information inaccessible byother approaches, giving much insight into how proteinsfunction. Remarkably, dynamic images can be interpreted
straightforward without intricate analyses and interpretations, making it possible to attain irrefutable conclusions.
There seems no doubt that continued exploration ofcountless biological phenomena will be carried out using
HS-AFM in the near future. This review is written with the

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Fig. 3. (Color online) Scanner movement tracing sample surface withsinusoidal shape with periodicity of , without parachuting. Gray (red in the
web version) broken line, trajectory of scanner movement; black (blue in the
web version) broken line, inverted trajectory of scanner movement. The sumof the black broken line and the sample surface is the tracing error.
Fig. 2. (Color online) Schematic for feedback loop and devices containedin the loop. The input to this feedback loop is the sample height variations
right under the cantilever tip, and the output is the z-scanner movement in
the z-direction. The output from the PID controller is used as an image
signal. The main time delays are indicated with  c for the response of thecantilever oscillation amplitude to tip–sample contact, p for tip
parachuting, m for detection of the cantilever oscillation amplitude, and z
for z-scanner movement.

intention of providing potential users of HS-AFM with an
overview of techniques implemented in the HS-AFM
instrument as well as precautions and devices for successful
imaging. In this review, we also brieﬂy discuss futurechallenges to overcome the limitations of current HS-AFM.

is deﬁned by the feedback frequency at which =4 phasedelay occurs in tracing the sample surface. Thus, we canderive the following relation; fB ¼ 1=ð16 open Þ. The main
operation of the PID control is performed by the
I-operation, i.e., the integration of error signals (diﬀerence
between the cantilever oscillation amplitude A and its set
point As ). The P- and D-operations compensate for the delay
in I-operation by a factor of  > 1, so that the feedback
bandwidth is modiﬁed as

fB ¼
:
ð1Þ
16open

2. Highest Possible Imaging Rate

2.2 Excessive force

2.1 Feedback bandwidth

Under the condition of the cantilever free oscillation
amplitude A0 and its set point As , a peak force Fp is exerted
from the oscillating cantilever tip to the sample by Fp ¼
kc ðA20  A2s Þ1=2 =Qc ,34) where kc is the spring constant andQc is the quality factor of the cantilever in water. Thisestimation is made under a provisional condition that no
feedback delay occurs. How large is the excessive force
exerted onto the sample due to feedback delay? For
simplicity, we assume that the sample has a sinusoidal
surface corrugation characterized with the wavelength  inthe x-direction and the maximum height h0 (see Fig. 3).
When the sample stage is moved at velocity Vs in thex-direction, the sample height h right under the cantilevertip changes as hðtÞ ¼ h0 =2  sinð2Vs t=Þ. As long as thecantilever tip always touches the sample surface at thebottom of swing, the z-scanner moves as zðtÞ ¼ h0 =2 sinð2Vs t=  close Þ. The diﬀerence between hðtÞ and zðtÞis the feedback error (‘‘residual topography’’, S) expressedas

We chose the tapping mode among various operation modesof AFM such as contact, tapping, and frequency-modulation(FM) modes. In the tapping mode, the cantilever is oscillated
in the z-direction at its ﬁrst resonant frequency. The resulting
intermittent contact between the tip and sample caneliminate friction force during lateral scanning and thus
minimize damage to and deformation of fragile biologicalsamples. The FM mode, in which the cantilever is oscillated
in the z-direction with a small amplitude less than 1 nm,
enables highly sensitive force detection. Nonetheless,frequency detection is rather complicated and slower thanamplitude detection. In the tapping mode, the vertical tip
force acting on the sample is controlled by a proportionalintegral-derivative (PID) controller so that the cantilever
oscillation amplitude (and thus the tip–sample interaction
force) is kept constant. Precise and fast feedback control ismandatory for fast and low-invasiveness imaging.
Figure 2 shows the devices contained in the closed
feedback loop as well as the time delays with each of which
the corresponding device responds when used independently(i.e., time delay intrinsic to each device). The input to thisfeedback loop is the variation in the sample height, while theoutput is the displacement of the z-scanner. The output fromthe PID controller is used as an image signal (i.e., a sample
height signal). The total sum of the intrinsic time delays isreferred as the open-loop time delay (open ). The total delayin the closed feedback loop (close ) is diﬀerent from open .
The relationship between them is approximately given as
close ¼ 2 open , as far as the feedback gain is maintained at
1.16) The phase delay in the feedback loop close is givenby close ¼ 2f close , where f is the feedback frequency atwhich the z-scanner is displaced. The feedback bandwidth fB

SðtÞ ¼ hðtÞ þ zðtÞcloseclosecos 2Vs t= ¼ h0 sin:ð2Þ22
The cantilever tip experiences this residual topography inaddition to a constant height of A0 ð1  rÞ, where r is thedimensionless set point (i.e., r ¼ As =A0 ). Therefore, when
feedback delay occurs, the maximum excessive force isexerted by kc h0 sinðclose =2Þ=Qc in an uphill region of thesample. On the other hand, when SðtÞ þ A0 ð1  rÞ < 0 in a
downhill region of the sample, the tip completely detaches
from the sample surface and time elapses until the tip landson the surface again (i.e., parachuting). During parachuting,the sample height information is completely lost. Therefore,

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the time over which parachuting continues also contributes
to the closed-loop time delay.
2.3 Highest possible imaging rate

The highest possible imaging rate (i.e, highest possible
frame rate) Rmax depends on the feedback bandwidth fB as
well as on imaging conditions, the spatial frequency of thesample surface corrugation (1=) to be imaged, and sample
fragility. The imaging conditions are given by the scan range
in the x-direction (W) and the number of scan lines (N).
When the x-scanner is displaced at velocity Vs , the z-scanneris scanned at a frequency f ¼ Vs = and one image iscaptured with a time T ¼ 2NW=Vs . Thus, we obtain T ¼
2NW=f . When the sample is strong enough so that it is not
aﬀected by the excessive tip force due to the =4 phasedelay, the feedback frequency f can go up to fB . In this case,
we obtain Rmax ¼ fB =ð2NWÞ. In reality, fragile biologicalmolecules would not withstand a large excessive force. By
denoting the maximum phase delay allowable for a fragile
molecule as max , the highest possible imaging rate is given
byRmax ¼

2max fB
:
NW

ð3Þ

Here, note that the mechanical quantity aﬀecting the sample(i.e., altering the momentum) is not the force itself but theimpulse of force, i.e., force  a time period over which theforce is applied. For our small cantilevers with a resonant
frequency of 1.2 MHz in water, the force applying period is100 ns or less.
3. Time Delays
3.1 Time delay with cantileverAs seen below, the cantilever is the most inﬂuential delay
component among devices in the feedback loop (Fig. 2).
First, the tip, which is oscillating at the ﬁrst resonant
frequency of the cantilever, fc , should tap the sample surface
at least once per pixel; thus, we obtain fc > 2Np-x Np-y =T ,where Np-x and Np-y are the numbers of pixels in the x- andy-directions, respectively. For example, to capture an image
with 100  100 pixels at 25 fps, fc should be at least 500 kHz
in an aqueous solution, when this tapping issue is only
considered. Upon tapping, a brief stepwise force is exerted
to the cantilever. The cantilever responds to this force with a
response time given by c ¼ Qc =ðfc Þ. In addition, it takes
time to measure the cantilever oscillation amplitude, theminimum time m of which is given by m ¼ 1=ð2fc Þ. Thus,
for the cantilever alone, we have a delay time of Qc =ðfc Þ þ
1=ð2fc Þ, when the cantilever oscillation amplitude can bemeasured at every half cycle of the oscillation. The timedelay is about 0.95 s for our small cantilevers with fc 
1:2 MHz and Qc  2 in water (vide x4.1).
3.2 Time delay with scanner

The response of the z-scanner with the ﬁrst resonant
frequency fz and the quality factor Qz is characterized by asecond-order transfer function. The phase delay ’z at the
feedback frequency f can be estimated from the phasefrequency relationship (i.e., the Bode phase plot) of thetransfer function. The time delay z is given by z ¼
’z =ð2f Þ. For our z-scanner with fz ¼ 370 kHz and the

maximum displacement 1 m (vide x4.1), the phase delay
is approximately 8.3 when driven at 100 kHz through anactive damping circuit with a mock z-scanner constructedusing an LRC circuit17) (vide x4.4.2), and thus,  z  0:23 s.
By active damping, Qz is moderately suppressed to 2 to
avoid a large phase delay. The resonance at 370 kHz is not
excited by driving at 100 kHz.
3.3 Time delay with parachuting

Other delay factors arise from electronic circuits containedin the feedback loop and from parachuting. Our analogue
electronics circuits [a preampliﬁer and a signal conditioner
used in the optical beam deﬂection (OBD) detector formeasuring cantilever deﬂection, a piezodriver for thez-scanner, and a PID controller] are minor delay factors.
Parachuting is apt to occur frequently and prolonged when
the cantilever amplitude set point As is set close to the freeoscillation amplitude A0 so that the oscillating tip lightly
taps the sample. This issue is problematic and a major
obstacle to making high-speed AFM imaging compatible
with low-invasiveness imaging. The maximum set point rmax
with which parachuting does not occur can be obtained using
the relation SðtÞ þ A0 ð1  rmax Þ ¼ 0 and is expressed asrmax ¼ 1 h0
close
:
sin
A0
2

ð4Þ

For example, when h0 ¼ 2A0 and rmax ¼ 0:9, we obtainclose ¼ 5:7 . This means that to avoid parachuting, we haveto decrease the imaging rate so that the feedback frequency fbecomes much lower than the feedback bandwidth, i.e., f ¼½close =ð=4Þ fB  0:13fB . In other words, when scanned at
f ¼ fB , signiﬁcant parachuting occurs and thus most of thedownhill regions of the sample cannot be imaged at all. We
have solved this diﬃcult problem by inventing a new
feedback controller termed ‘‘dynamic PID controller’’.18)
This controller makes low-invasiveness imaging possible
without deterioration of the scan speed because it cancompletely suppress parachuting as far as the condition As 
0:9  A0 is used (vide x4.4.3). The analytical expression ofparachuting time (delay time) is given elsewhere.16)
4. Components and Control Techniques for HS-AFM

As comprehensive descriptions of devices and techniquesdeveloped for HS-AFM have already been given elsewhere,16) only their outlines are described here. Our HSAFM instrument is now commercialized by the ResearchInstitute of Biomolecule Metrology (Tsukuba, Japan).
4.1 Small cantilevers

Several types of small rectangular cantilevers made ofsilicon nitride have been developed by Olympus incollaboration with our group. The high-end small cantilevers(Olympus BL-AC7DS-KU2, custom-made) that we havebeen routinely using for imaging studies are 6–7 m long,
2 m wide, and 90 nm thick [Fig. 4(a)]. They have thefollowing mechanical properties: fc  1:2 MHz in water(3:5 MHz in air), Qc  2 in water, and a spring constantkc  0:2 N/m. The commercially available small cantilevers(Olympus BL-AC10DS-A2) are 9–10 m long, 2 m wide
and 130 nm thick, and have fc  0:6 MHz in water (1.5 MHz
in air), Qc  2 in water, and kc  0:1 N/m.

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apex radius is usually reduced to 5 nm [Fig. 4(d)] andsometimes to 0:5 nm (yield: 10%), judging from the
AFM images obtained with the sharpened tips.23) Since thetip radius directly inﬂuences the resolution of the captured
images, we need to improve our method to enhance the yieldof sharp tips with an apex radius of 0:5 nm. The low yieldis due likely to the fact that the apex with a 0:5 nm radius
produced during plasma etching tends to collapse withfurther etching.
Materials harder than carbon will probably produce sharp
tips with a higher yield. Although we have not tried yet, we
can prepare EBD tips of tungsten or a mixture of tungsten
and carbon. A gas injection system, which is adaptable to
SEMs and can mix diﬀerent gases under computer control, iscommercially available (Omniprobe OmniGISÔ).
4.3 Fast amplitude detector

Fig. 4. Scanning electron micrographs of small cantilevers. (a) Lever andbird-beak-shaped tip. (b) EBD tip grown on an original bird-beak-shaped
tip. (c) Distal end of EBD tip before plasma etching. (d) Distal end of EBD
tip after plasma etching. (e) Wing portion of previous version of smallcantilever. (f ) Wing portion of most recent version of small cantilever.

The change in the angle ’ of the free end of a cantileveris given by ’ ¼ 3z=ð2LÞ, where z is the verticaldisplacement of the free end and L is the cantilever length.
For the short cantilevers, the angle-based OBD detection of
cantilever deﬂection7,8,35) is, therefore, advantageous overdistance-based detection, for example, by optical interferometric methods.36,37) The short cantilevers have 10–20 times
higher sensitivity for OBD detection than conventional
cantilevers. We have been using an OBD detector optimized
for small cantilevers.8)
4.2 Cantilever tip

The tip of small cantilevers manufactured by Olympus has abird-beak-like shape with an apex radius range of 15–24 nm[Fig. 4(a)].10) Therefore, we have to attach a sharper tip tothe bird-beak-shaped tip in order to obtain high-resolution
images. Although attaching38,39) or directly growing40) a thincarbon nanotube onto an original cantilever tip is reported,
their procedures are too demanding for routine use. Usinga scanning electron microscope (SEM of the low-vacuumtype), we routinely prepare electron-beam-deposited (EBD)
tips under an atmosphere of phenol gas sublimated fromphenol crystals [Figs. 4(b) and 4(c)].16) They grow at
1 m/min under a low-vacuum condition. The tip can be
sharpened by nitrogen or oxygen plasma etching.16) The

We developed two types of fast amplitude detectors: peakhold and Fourier types (Fig. 5). The peak-hold type[Fig. 5(a)] holds the sinusoidal output from the splitphotodiode sensor (quadrant PIN diode sensor) at its peak
and bottom, and outputs their diﬀerence as the peak-to-peakamplitude at every half cycle of cantilever oscillation.8) The
sample/hold timing signals are generated using either the
split-photodiode sensor output itself (internal mode) or asignal synchronized with the cantilever excitation signal(external mode). In addition to a decrease in the cantilever
oscillation amplitude, the resonant frequency of the cantilever shifts towards a higher frequency when repulsive
force acts on the tip. This frequency shift results in a phaseshift of the cantilever oscillation relative to the excitation
signal. Therefore, in the external mode where the sample/
hold timings are independent of the phase shift, the detected
amplitude signal is aﬀected by the phase shift. Therefore, theexternal mode is more sensitive to the tip–sample interaction
than the internal mode. Here, note that the phase shift of an
oscillating small cantilever is 1;000 times larger than thatof conventional cantilevers because of the large resonant
frequency-to-spring constant ratio of the small cantilevers.
Although this amplitude detection is fastest, it is apt to beaﬀected by the thermal motion of the cantilever because
only two-point voltages of the split-photodiode sensor output
are used. In the Fourier type [Fig. 5(b)], the Fourier sine andcosine coeﬃcients (a and b, respectively) are calculated forthe cantilever’s ﬁrst resonant frequency component of the
split-photodiode sensor signal, and ða2 þ b2 Þ1=2 is output at
every cycle of cantilever oscillation.16) This detector is lesssusceptible to the thermal eﬀect (1=4) than the peak-holdtype.
4.4 High-speed scanner

Some conditions are required for the high-speed scanner: (a)a high resonant frequency, (b) a small number of resonantpeaks within a narrow frequency range, (c) suﬃcientmaximum displacement, (d) a small crosstalk between thethree scan axes, and (e) a small quality factor. Here, several
techniques that have met these conditions are described.
4.4.1 Mechanical design and counterbalance

The quick displacement of a piezoactuator exerts a large
impulsive force to the supporting base, causing vibrations of

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(a)

(b)

Fig. 5. Circuit diagrams for two types of fast amplitude detectors: (a) peak-hold type and (b) Fourier type.

during its displacement.16) The y-scanner displaces a blockcontaining the x- and z-scanners, the x-scanner displaces a
block containing the z-scanner, and the z-scanner displacesa sample stage. Each block is connected to a surrounding
mechanism through a pair of ﬂexures. The ﬂexures are
ﬂexible in the displacement direction, but are stiﬀ in theperpendicular directions. This connection between the x-, y-,and z-scanners through the ﬂexures minimizes crosstalk
between the three scan axes. The scanner’s surrounding
mechanism and ﬂexures are monolithically fabricated from astainless steel block or a duralumin block. The gaps in thescanner are ﬁlled with an elastomer to passively damp thevibrations.
Fig. 6. (Color) High-speed scanner. A sample stage (glass rod) is glued
on the top surface of the z-piezoactuator using nail enamel. 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 scanner block are ﬁlled with an elastomer for passive
damping.

the base and the surrounding mechanisms and in turn of theactuator itself. To alleviate the vibrations, counterbalancemethods have been introduced into the x- and z-scanners(Fig. 6). For the z-scanner, two identical piezoactuators
are attached to a supporting base at the counterplanes.16)
Impulsive forces are countered by the simultaneous displacement of the actuators of the same length in the counter
directions. In another method used for the z-scanner, apiezoactuator is held at the four rims parallel to thedisplacement direction so that its center of mass remains
unchanged during displacement.15) In this method, themaximum displacement becomes one half of that in theﬁrst method, but the resonant frequency remains almost
unchanged from that in free oscillation. For the x-scanner, apiezoactuator is held at the two opposite planes (perpendicular to the displacement direction) with a pair of ﬂexures sothat the center of mass of the actuator remains unchanged

4.4.2 Active damping

The fast scanners designed as above have a small number ofresonant peaks. The resonant peaks generally consist of thestructural resonance (arising from the scanner mechanism)and the resonance of the actuators themselves. The resonantpeaks of the x-scanner comprise both types of resonances,and the frequency of its structural resonance is lower thanthat of the x-piezoactuator itself. Both resonances can be
simultaneously eliminated by a feedforward compensation
method because it is scanned in a periodic pattern (isosceles
triangle waveform).16) Let us express the Fourier transformof a series of isosceles triangles as Fð!Þ and the transfer
function of the x-scanner as Gð!Þ. In the compensation
method, the inverse Fourier transform of Fð!Þ=Gð!Þ iscalculated and output to the x-scanner driver through adigital-to-analog (D/A) converter. The ﬁrst 10 terms in theinﬁnite series of the inverse Fourier transform are practicallysuﬃcient for driving the x-scanner in a well-approximatedisosceles triangle waveform. This method can extend thex-scanner bandwidth as long as the piezodriver to be used
has suﬃcient bandwidth for higher frequencies.
This feedforward method cannot be applied to thez-scanner because its displacement is not periodic and

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actuator.18) This drift compensator enables stable lowinvasiveness and high-speed imaging when operating
together with the dynamic PID controller, even under theconditions of A0 ¼ 1{2 nm and As  0:9  A0 .
5. Sample Stage and Hydrodynamic Pressure

Fig. 7. Active damping of z-scanner vibrations. The LRC circuit (mockz-scanner) is constructed so that its transfer function is similar to that of thereal z-scanner.

unspeciﬁed beforehand. When an alternative feedback
method for damping is used, the displacement or speed of
the z-scanner has to be measured, but it is diﬃcult to do so.
We developed a feedback Q-control method in which theoutput from a ‘‘mock z-scanner’’ (LRC circuit) is used.17)
The mock z-scanner is characterized with a transfer function
similar to that of the real z-scanner. An input signal to thez-piezodriver is also input to the mock z-scanner, and theminus derivative of the output from the mock z-scanner isadded to the output signal from a PID feedback controller(Fig. 7). As a result, the quality factor Qz is reduced.
4.4.3 Dynamic PID control and drift compensation

As shown in Fig. 3, the plus–minus signs of the error signal
are diﬀerent between the uphill and downhill regions of thesample. Therefore, the sign of the error signal indicates
which region is being scanned. We used this indicator toeliminate the tip parachuting eﬀect. Simply, the feedback
gain is increased in the downhill region while the gain is
kept constant in the uphill region.18) There are several ways
to automatically change the feedback gain depending on theerror signal (or on the cantilever oscillation amplitude). We
employed the following method. A threshold level Aupper is
set at (or slightly above) the amplitude set point As . Whenthe cantilever oscillation amplitude A exceeds this thresholdlevel, the diﬀerential signal (A  Aupper ) is ampliﬁed andadded to the error signal. The error signal that contains an
extra signal is fed to the conventional PID controller. The‘‘false error signal’’ which is larger than the ‘‘true error
signal’’ produces a quicker feedback response; therefore,
A quickly becomes smaller than Aupper and the feedback
operation automatically returns to the normal mode. Thus,
even with As very close to A0 , the parachuting period isshortened markedly or can be completely eliminated.
To minimize the tapping force, a cantilever free oscillation amplitude A0 ¼ 1{2 nm is used. Therefore, the diﬀerence between A0 and As is very small (0:1{0:2 nm) under
the condition As  0:9  A0 . However, the oscillation
power of an excitation piezoactuator gradually decreases
with time because the excitation eﬃciency becomes low
probably owing to the temperature rise of the piezoactuator.
Thus, A0 easily decreases by >0:2 nm owing to this drift,
and therefore, stable imaging is diﬃcult under this condition.
This problem was solved by maintaining a constant average
amplitude of a second harmonic oscillation of the cantileverby controlling the power driving the excitation piezo-

In high-speed imaging, the sample-stage z-scanner moves athigh frequencies (50–100 kHz); hence, a substrate, which is
attached to the sample stage and in contact with a buﬀer
solution, also moves at high frequencies in the z-direction.
This fast movement of the substrate produces hydrodynamic
pressure. An oscillating cantilever and its supporting baseare positioned very close to the substrate surface (here, thesupporting base, which we call the ‘‘wing’’ [Figs. 4(e) and4(f )], is the portion that bridges the cantilever beam and thelarge base directly in contact with the cantilever holder).
Therefore, they are apt to be moved by the hydrodynamic
pressure, particularly when the sample stage is large. The
movement results in a slow response of the cantilever
oscillation amplitude to z-scanner displacement. Therefore,
in a HS-AFM system with a sample-stage scanner rather
than a tip scanner, the size of the sample stage should be
minimized.9) As a sample stage, we routinely use a glass rodof 1.5–2 mm diameter and 2 mm height to the upper end ofwhich a substrate (usually a mica disk of 1 mm diameter and<0:1 mm thickness) is glued with epoxy adhesive. The
bottom end of the glass rod is glued onto the top of thez-piezoactuator with nail enamel.
Further precaution is required to minimize the hydrodynamic pressure eﬀect. Even with a small sample stage, thehydrodynamic pressure is large at its central area. In fact,
when a cantilever tip is placed at the central area andbrought into contact with the substrate surface, the responseof the cantilever oscillation amplitude to quick displacementof the sample stage is delayed.41) In the worst case, this delay
extends to a few s which is signiﬁcantly longer than theminimized total time delay in the feedback loop. When the
cantilever tip is placed in an edge region of the sample stage
so that the whole cantilever chip minimally overlaps with thesample stage, no time delay due to the hydrodynamic
pressure is observed. The angle between the top surface ofthe sample stage and the cantilever (the entire cantileverchip) is also an important factor. When the angle is small(5 ), the wing of the cantilever is subjected to thehydrodynamic pressure, and thus, the cantilever response
is slowed. We usually set the angle at 10 .
The design of the wing is also an important factor forminimizing the hydrodynamic pressure eﬀect. In a previous
design, the wing is large [Fig. 4(e)], but it is much smaller inthe current one [Fig. 4(f )]. The cantilevers with the latter
design are much less aﬀected by the hydrodynamic pressure.
Unfortunately, this excellent improvement is not yet
employed for commercial small cantilevers produced by
Olympus.
The shape and length of cantilever tip are also importantfactors to consider. When the tip is shorter, the cantilevergets closer to the sample surface at the bottom of the swing
and thus the solution conﬁned between them becomes moresqueezed, resulting in oscillation damping. This squeeze
eﬀect also occurs in the space conﬁned by the tip and thesample surface when the tip is wider. Not only damping but

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Fig. 8. Response of cantilever oscillation amplitude while approaching
substrate surface in water and upon contact with the surface. The gray line
indicates the response of a small cantilever having an EBD tip of 1 mlength grown on the original bird-beak-shaped tip, while the black line
indicates that of a cantilever with only a bird-beak-shaped tip.

also the deterioration of sensitivity to the tip–sample
interaction is caused by the squeeze eﬀect.41) Figure 8
shows the force curves obtained using small cantilevers with
diﬀerent tip shapes and lengths. When a 1.5 m long tip
with a bird-beak shape is used, the cantilever oscillation
amplitude becomes smaller with approaching the substratesurface, whereas when the tip has an additional 1-m-longneedle-shaped tip, the oscillation amplitude is kept constantuntil the tip comes into contact with the substrate surface.
Moreover, when brought into contact with the surface, theoscillation amplitude more sharply decreases with the longer
tip than with the shorter one.
6. Substrate Surfaces

The substrate surface on which a sample is placed plays
an important role in dynamic AFM imaging. To observe
dynamically acting protein molecules under physiologicalconditions, the substrate surface must have an appropriate
aﬃnity for the molecules. When the aﬃnity is too low,
protein molecules diﬀuse rapidly on the surface and cannot
be clearly imaged even by HS-AFM. When the aﬃnity is too
high, the molecules often lose their biological function. To
observe dynamic interactions between diﬀerent proteins,
selective protein attachment to a surface is required.
Otherwise, there is no chance to observe their interaction.
Thus, we have to prepare an appropriate substrate surface
depending on the dynamic biomolecular event to bevisualized. As substrate surfaces, bare24–27,31,32,42) or chemically treated23) mica surfaces, planar lipid bilayer surfaces,22)
and the surfaces of two-dimensional crystals of streptavidin
formed on a biotin-containing lipid bilayer have been used
for dynamic AFM imaging.33,43)
Mica (natural muscovite or synthetic ﬂuorophlogopite)
has been frequently used as the substrate source because of
its surface ﬂatness at the atomic level over a large area. It has
a net negative charge and is therefore quite hydrophilic. A
bare mica surface adsorbs various proteins by electrostaticinteraction. We can control the aﬃnity for a speciﬁc protein
by varying the ionic strength or pH, or by adding divalentcations such as Mg2þ . For the immobilization of fractionatedmembranes containing membrane proteins, the bare mica

surface is most useful. A water layer of 1 nm thickness
separates the membranes from the mica surface, enabling the
motion of membrane proteins within the membranes.44,45)
In fact, bacteriorhodopsin (bR) in a non-crystalline region
rapidly diﬀuses in the purple membrane attached to a baremica surface.26) By chemically treating a mica surface,proteins can be covalently immobilized onto the surface. For
instance, F1 -ATPase was attached onto a mica surface in a
selective orientation; the mica surface was ﬁrst coated withamino silane and then treated with an amino crosslinker,
glutaraldehyde. F1 -ATPase with (Lys)7 introduced to the
N-terminus of the - and -subunits was covalently
immobilized onto the surface, in an end-up orientation with
the C-terminal surface at the top.23)
The surface of two-dimensional crystals of streptavidin
is particularly useful for the selective immobilization ofhomo-oligomeric protein complexes, such as GroEL andactin ﬁlaments because a stable pinning of the complexes atmultiple biotinylated sites is possible.33,46) Importantly, thesurface is resistant to the nonspeciﬁc binding of proteins.
However, for nonoligomeric proteins, their tethering to thesurface through a single biotinylated site sometimes results
in a rapid motion around the link. The high mobility can be
reduced using reactive dibiotin compounds.
Planar lipid bilayer surfaces can be used for both speciﬁc
and nonspeciﬁc (electrostatic) immobilizations of proteins.22,46) For the speciﬁc immobilization of biotinylated
proteins and His-tag-conjugated proteins, lipids with biotin
and Ni–NTA (nickel–nitrilotriacetic acid) at the polar headgroups can be used, respectively. Multiple-point pinningis sometimes required for stopping the rapid motion ofmolecules tethered to the surfaces. Planar lipid bilayer
surfaces prepared using electrically neutral phospholipids
are very resistant to the nonspeciﬁc binding of proteins.
For electrostatic immobilization, lipids with charged headgroups can be used. Unlike bare mica surfaces, the surface
charge density can be controlled using diﬀerent fractions ofa charged lipid. The charge polarity can also be altered using
positively or negatively charged lipids. For the dynamic
AFM imaging of myosin V on an actin ﬁlament, a planar
lipid bilayer surface comprising a biotinylated lipid andpositively and neutrally charged lipids was used.22)
7. Imaging Studies

Around 2008, the instrumentation of HS-AFM was nearlymature, and thus, the use of HS-AFM was started to solve
biological issues that have been diﬃcult or impossible toaddress by other methods. Since then, the number ofpublications on bioimaging by HS-AFM has gradually
increased, rapidly increasing in the last two years. The
imaging studies conducted thus far have covered a wide
range of dynamic molecular processes and structure
dynamics: e.g., myosin V walking on an actin ﬁlament,22)
rotary catalysis of rotorless F1 -ATPase,23) bacteriorhodopsin
in response to light,24,25) dynamics of bacteriorhodopsin
2D crystals,26,27) ATP-induced structural changes in P2X4
receptors,28) cellulase hydrolyzing cellulose ﬁbers,29,30)amyloid-like ﬁbril formation from cleaved lithostathine,31)
formation of supported planar lipid bilayers,47) reaction of
DNA restriction-modiﬁcation enzyme,48,49) unwrapping of
nucleosomes,50,51) GroEL-GroES interaction regulated by

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(a)

(c)

(b)

Fig. 9. (Color) Structural change in bR induced by light absorption. (a) Successive high-speed AFM images of the cytoplasmic surface of D96N bR in thedark or illuminated conditions. Frame rate, 1 fps; pixel size, 200  200 pixels. A bR trimer is highlighted by the white triangle. The green bars at 2 and 3 s
indicate illumination of 532 nm green light of 0.5 W. (b) Schematic of arrangement of bR molecules and bR trimers in the dark. The arrows indicate thedirections in which respective bR molecules are displaced upon light illumination. (c) Schematic of arrangement of bR molecules and bR trimers underillumination. 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.

ATP hydrolysis by GroEL,14,33,52) undulation motion of an
intrinsically disordered protein,42) membrane-mediated interaction between c-rings of ATP synthase,32) collapse of
three-dimensional DNA origamis,53,54) defect diﬀusion inprotein 2D crystals,43) enzymatic reactions in DNA origamis,55,56) and translocation of a DNA nanorobot on a track
formed on a DNA origami tile.57) Moreover, nonbiologicaldynamic phenomena, such as the photodegradation of single
chains of a chiral helical -conjugated polymer,58) the phasetransition of detergent aggregates,59) and photoresists
dissolution,60) have also been studied. Fantner et al. studied
the kinetics of antimicrobial-peptide-induced morphologicalchanges in individual bacterial cells using a HS-AFM
instrument and the small cantilevers they developed.61) In
this section, some imaging studies performed by our groupare highlighted. Several molecular movies can be seen at our
homepage(http://www.s.kanazawa-u.ac.jp/phys/biophys/index.htm).
7.1 Bacteriorhodopsin in response to lightbR functions as a light-driven proton pump, transferring
protons across the membrane from the cytoplasmic side to
the extracellular side.62,63) bR comprises seven transmembrane -helices (named A–G) surrounding the retinal
chromophore covalently bound to Lys216 via a protonatedSchiﬀ base.64,65) In the purple membrane, bR assembles into
trimers, which are arranged in a hexagonal lattice. Uponabsorption of light, photoisomerization from the all-transconformation to the 13-cis conformation of the retinal takes
place. This isomerization induces proton transfer from theSchiﬀ base to Asp85, which triggers a cascade of changes inthe bR structure. A series of intermediates designated J–O
are deﬁned by spectroscopy, and M (M410 ), which has anabsorption peak at 410 nm and the longest lifetime, is the
only intermediate containing a deprotonated Schiﬀ base.63)
The light-induced conformational changes in bR in thefrozen activated state of the wild type (WT) and bR mutants
have been studied by X-ray crystallography and electron

microscopy.66–68) The extent of the changes reported varies
depending on the methods used, but is generally small (0.1–
0.3 nm). Until our recent study, the dynamic nature of theconformational changes has remained elusive owing to thelack of techniques feasible for the high-resolution visualization of the dynamic structural changes of proteins. Our groupsucceeded for the ﬁrst time in capturing high-resolution
successive images showing the dynamic changes of bR in
response to light using HS-AFM.24,25)
The photocycle of the wild type at neutral pH proceeds
very fast (10 ms). To slow down the photocycle, we thenused the D96N bR mutant that has a longer photocycle(10 s at pH 7) but still retains an ability of proton
pumping.69) Native purple membranes containing D96N
bR were directly placed on a bare mica surface. Figure 9(a)shows successive images of the D96N mutant at thecytoplasmic surface captured at 1 fps. Upon illumination
with green light (532 nm, 0.5 W), bR markedly changed its
structure (compare images at 1 and 2 s) and returned to theunphotolysed state within a few seconds after light-oﬀ. Notethat this structural change is speciﬁcally induced by green
light not by blue or red light, and that bR in the groundstate eﬀectively absorbs green light. All the activated bR
monomers moved outwards by 0.7–0.8 nm from the center ofthe respective trimers and rotated counterclockwise by 7–8around the center of the respective trimers, as illustrated byarrows in Fig. 9(b). However, the overall position of eachbR 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 M are very short. However, it remained uncertain
whether the photoactivated state detected by AFM also
contained the N intermediate that follows M. This issue wassolved in a further study.25) In this study, the D96N mutants
were alternatively illuminated with green light and blue
light. It is well known that the absorption of blue light by

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retinal in M410 induces the back isomerisation reaction ofretinal. The HS-AFM observation showed that the bR
molecules activated by green light illumination quickly
return back to the ground state upon subsequent blue lightillumination. This means that the excited state detected by
HS-AFM only contains the M intermediate. Moreover, it
was also revealed by this observation that the conformationof bR is tightly governed by the conformation of retinal.
As a result of the outward displacement, nearest-neighborbR monomers, each belonging to a diﬀerent adjacent trimer,transiently assemble, as illustrated in Fig. 9(c). Here, we use
a new designation, ‘‘trefoil’’, for the triad of nearest-neighbormonomers belonging to diﬀerent trimers to distinguish itfrom the original trimer. To investigate a possible eﬀect ofbR-bR interaction within a trefoil on the photocycle, bR
was illuminated with lights of diﬀerent intensities (0.007–
0.5 W). Under weak illumination, only one monomer ineach trefoil was activated in most of the cases. When only
one monomer was activated in a trefoil, it decayed with atime constant of 7:3  0:58 s. In contrast, under stronger
illumination, two or three monomers within a trefoil tended
to be activated together. Interestingly, the decay of eachmonomer markedly depended on the order of its activation.
The monomer that was activated latest among the activatedmonomers in a trefoil decayed with a shorter time constantof 2:0  0:16 s. On the other hand, the decay kinetics of theearly activated monomers did not follow a single exponential, and the average decay time lengthened to 13 s. Thisobservation indicates that 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 a trefoil under illumination engenders
both positive and negative cooperative eﬀects in the decay
kinetics as the initial bR recovers. It is interesting to notethat the cooperativity does not occur within a trimer but
within a trefoil. As a consequence of both positive andnegative cooperative eﬀects, the turnover rate of thephotocycle is maintained constant, on average, irrespectiveof the light intensity. This seems to indicate why bR trimers
assemble into a hexagonal lattice in the native state.
7.2 Cellulase hydrolyzing cellulose fibers

Cellulose is the most abundant biomass and component offeed crops used in the livestock industry. Therefore, theeﬃcient conversion of cellulose to fuels and the improvement of cellulose digestibility particularly for ruminants areimportant issues in agricultural industries. One of the
problems in bioreﬁning applications to cellulose is the low
enzymatic hydrolysis of cellulosic raw materials. To
maximize the hydrolysis eﬃciency, various studies havebeen performed on individual cellulases, cellulase cocktails,70) and the pretreatment of cellulose ﬁbers. One of themost serious problems is the rate of hydrolysis being quickly
slowed down with time via an unknown mechanism.71,72)
Biochemical analyses are ineﬀective for resolving this issue
because the reaction occurs not in a bulk solution but at asolid–liquid interface. We attempted to determine themechanism by observing cellulase molecules hydrolyzing
cellulose ﬁbers using HS-AFM.30) This approach wasdemonstrated to be eﬀective for such a unique enzymatic
reaction.

7.2.1 Traffic jam of cellulase on cellulose fibers

The cellulolytic ascomycete fungus Trichoderma reesei (Tr)secretes two extracellular cellulases TrCel7A and TrCel6A
that hydrolyze the glycosidic linkages of crystalline
cellulose to produce cellobiose (-1,4-glucose dimmer).73)
TrCel7A hydrolyzes cellulose from its reducing end, while
TrCel6A does it from the nonreducing end.74) These
enzymes have similar two-domain structures [Fig. 10(a)]composed of a smaller cellulose-binding domain (CBD)
and a larger catalytic domain (CD). The active site is locatedin a relatively long tunnel formed in CD: 5 nm long in thecase of TrCel7A and 2 nm long in the case of TrCel6A.75,76)
Therefore, these enzymes have been thought to consecutively hydrolyze a crystalline cellulose chain without
releasing the chain, and thus, have been expected toprocessively and unidirectionally move along a cellulose
ﬁber.73–77)
Crystalline cellulose I was placed on a bare surface ofhighly ordered pyrolytic graphite (HOPG) and TrCel7A wasadded to the bulk solution. As shown in Fig. 10(b),TrCel7A, in fact, moved processively and unidirectionallyon the surface of crystalline cellulose.29,30) The unidirectional movement of TrCel7A was not smooth but anintermittently repeated stop-and-go behavior [Fig. 10(c)].
When TrCel7A continuously moved within a certain range,
its velocity was 7.1 nm/s on average. The overall average
speed was 5.3 nm/s, without considering molecules thatcompletely halted during imaging. One hydrolytic cycle
produces one cellobiose unit of approximately 1 nm inlength. Therefore, the hydrolysis rate is estimated to be5/s. This value is about 300 times higher than thebiochemically measured hydrolysis rate. This large discrepancy seems mainly due to the limited number of available
reduced ends in the crystalline cellulose.
Crystalline cellulose IIII , which has a crystalline polymorphic form diﬀerent from that of cellulose I , is formed
through a super-critical ammonium treatment of cellulose I
and can be more eﬀectively degraded by TrCel7A.78)
Biochemical analysis indicated that TrCel7A has a higher
aﬃnity for cellulose IIII than for cellulose I , but its
maximum hydrolysis rate is only slightly higher than thehydrolysis rate of cellulose I .78) The HS-AFM observationof the TrCel7A movement on the surface of cellulose IIII
revealed a ‘‘traﬃc jam’’ of bound enzyme molecules[Fig. 10(d)]. When one molecule was halted, many following molecules stacked behind it, causing a traﬃc jam.
However, after additional molecules had also become
blocked, the enzyme molecules restarted to move again on
the surface, and a cellulose bundle was peeled oﬀ thecrystalline cellulose [Fig. 10(e)]. These phenomena of traﬃc
jam and ﬁber peeling may be able to account for the rapid
retardation of cellulose hydrolysis rate with time. The
distance between neighboring traﬃc lanes is smaller than thesize of TrCel7A. Therefore, the enzyme molecules moving
along neighboring lanes have a large chance of colliding
with each other. Before adding TrCel7A, the surface of
crystalline cellulose ﬁbers is smooth. When a TrCel7A
molecule is hydrolyzing one lane of cellulose, it is likely thatit also partially peels lanes adjacent to the lane beinghydrolyzed by the molecule. This peeling is likely to resultin a go-and-stop behavior of the molecule as well as in a

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(a)

(c)

(a)

(b)

(d)

(b)

(e)

(c)

Fig. 10. (Color) Dynamics of cellulase TrCel7A hydrolyzing cellulose
ﬁbers captured by HS-AFM. (a) Schematic of cellulases with two-domain
structure. (b) Successive AFM images showing processive andunidirectional movement of TrCel7A on crystalline cellulose I . Frame rate,
1.1 fps. Three molecules are marked with arrows 1–3. (c) Intermittent ‘‘goand-stop’’ movement of TrCel7A molecules on cellulose I . The movementsof seven molecules are shown with diﬀerent marks. The broken line
represents the average velocity of TrCel7A (3.5 nm/s) estimated in previous
HS-AFM observation.29) (d) Successive AFM images showing traﬃc jam of
TrCel7A bound to crystalline cellulose III. Frame rate, 3.3 fps. The
congestion is marked with arrows. (e) Successive AFM images showing
ﬁber peeling (marked with arrows) of crystalline cellulose III by TrCel7A
after congestion. Frame rate, 3.3 fps.

Fig. 11. (Color) Synergy between TrCel6A and TrCel7A in cellolose
hydrolysis. (a) Biochemically detected synergy in cellobiose production
from cellulose III. The green and red plots are the rates of cellobiose
production by TrCel6A and TrCel7A, respectively, and the blue plot shows
the synergy between the two enzymes. The dotted line indicates the simple
sum of cellobiose production calculated from the green and red plots.
(b) Successive AFM images showing crystalline cellulose III ﬁbers beinghydrolyzed by TrCel6A alone. No appreciable degradation of the ﬁbers isobserved. The time shown in each image indicates a lapse after the additionof TrCel6A. Frame rate, 3.3 fps. (c) Successive AFM images showing rapid
degradation of crystalline cellulose III ﬁbers in the presence of both
TrCel6A and TrCel7A. TrCel7A was added 8.3 min after the initial additionof TrCel6A. The time shown in each image indicates a lapse after the
addition of TrCel7A. Frame rate, 3.3 fps.

rough surface of crystalline cellulose; thus, TrCel7A
molecules are apt to be halted. Once one molecule isstacked, many molecules around it are also stacked. Most
likely, this is the mechanism underlying the rapid retardationof cellulose hydrolysis rate.

TrCel6A [although an image at 3.6 min is shown in
Fig. 11(b), the appearance was almost unchanged even at
8 min]. However, the addition of TrCel7A at 8.3 min
abruptly altered the microﬁbril appearance; the cellulose
ﬁbrils were rapidly degraded and almost completely
disappeared at 3.6 min after the addition of TrCel7A[Fig. 11(c)]. This observed synergistic eﬀect was much
more drastic than the biochemically observed one and was
independent of the order of addition of the two enzymes. We
noticed that bound TrCel6A molecules often detached from
cellulose IIII without showing appreciable processive run.
This fact and the observed synergy suggest that TrCel6A canalso work as an endo-glucanase. The short active-site tunnelof TrCel6A may be able to open and allow the generation ofnicks in the middle of crystalline cellulose; these nicks,
therefore, become starting and ending points for TrCel7A
activity. Thus, the synergy observed is not an exo–exo
synergy but an endo–exo one.

7.2.2 Endo-exo synergy

As reported previously,74,79) a cocktail of TrCel7A and
TrCel6A synergistically increases the rate of cellobiose
production [Fig. 11(a)]. This eﬀect seems peculiar because
cellulose hydrolysis from both reducing and nonreducing
ends cannot explain the synergy. To understand themechanism underlying this synergistic eﬀect, the cellulose
degradation process was imaged ﬁrst in the presence of
TrCel6A alone. After a while, TrCel7A was added while
imaging was ongoing. When TrCel6A was alone, nosigniﬁcant change was observed in the appearance of
cellulose IIII microﬁbrils even 8 min after the addition of

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(a)

(b)

(c)

(d)

Fig. 12. (Color) Walking M5-HMM captured by HS-AFM. (a) Successive AFM images showing processive movement of M5-HMM in 1 M ATP. Frame
rate, 7 fps. (b) Schematic of two-headed bound M5-HMM. (c) Successive AFM images showing hand-over-hand movement in 1 M ATP. Frame rate, 7 fps.
The swinging lever is highlighted with a thin white line. (d) Schematic explaining the images in C. The mark ‘‘+’’ indicates the plus ends of actin ﬁlaments.

7.3 Walking myosin V on actin filament

Double-headed myosin V (M5), which is a member of the
myosin superfamily and functions as a cargo transporter incells, has been extensively studied by single-molecule
ﬂuorescence microscopy and optical-trap namometry.80) It
moves processively along an actin ﬁlament towards the plus
end of the ﬁlament81,82) in a hand-over-hand manner witha 36 nm advance83–85) for every ATP hydrolysis cycle.86)‘‘Hand-over-hand’’ means that the two heads step alternately,exchanging leading and trailing roles at each step, verymuch like ‘‘walking’’. Nevertheless, comprehensive descriptions of the walking behavior and how force generation iscoupled with the ATPase reaction have not been attained
until our recent HS-AFM observation of the motor.22)
7.3.1 Step motion of myosin V

To observe moving M5 molecules on actin ﬁlaments, theﬁlaments have to be immobilized on a surface, while M5should be free from the surface. Therefore, selective
immobilization is required. As a substrate surface, we ﬁrst
used an electrically neutral phospholipid bilayer surface
containing biotin-lipid, formed on mica. Partially biotinylated actin ﬁlaments were immobilized onto this surface
through streptavidin with a low surface density. Tailtruncated M5 (M5-HMM) was never bound to this surface.
Although these conditions were ideal, we included apositively charged lipid 1,2-dipalmitoyl-3-trimethylammonium-propane (DPTAP, 5%) in the bilayer. The appropriately charged surface facilitated observing the sideways
ﬁgure of actin-bound M5-HMM. M5-HMM moving processively with 36 nm steps was clearly visualized at 7 fps[Fig. 12(a)]. The two-headed bound M5-HMM showed
unique structural features [illustrated in Fig. 12(b)]; theneck–motor domain junction appears smooth in the leadinghead but is V-shaped in the trailing head without exception,
because the neck regions emerge from diﬀerent parts of the
motor domain [Figs. 12(a) and 12(b)]. The short coiled-coiltail was mostly tilted towards the minus end of actin. Thesefeatures can be used to determine the actin polarity whenbound M5-HMM is stationary. When DPTAP was absent in
the lipid bilayer, the average translocation velocity was
similar to that measured by ﬂuorescence microscopy under
the same solution condition, indicating no eﬀect of the tip–
sample interaction on the motor activity. However, we could
not see detailed molecular behavior during a step as it wascompleted within a frame.
To slow down the step, streptavidin molecules were
further placed on the substrate surface as moderate obstacles
to the advance. This method allowed the visualization of
stepping processes [Fig. 12(c); illustrated in Fig. 12(d)].
After trailing head detachment, the leading head appeared tospontaneously rotate from the reverse arrowhead orientation
towards the arrowhead orientation (the term ‘‘arrowhead’’originates from the conﬁguration of single-headed myosinbound to an actin ﬁlament in the rigor state). Before
completing this rotation, the leading head brieﬂy halted bycolliding with a streptavidin molecule placed on the way of
its natural path. In this halted state, the detached trailinghead was most distant from the actin ﬁlament (thus, the twoheads were aligned nearly straight, pointing in opposite
directions) and slightly rotated around the neck-neckjunction [the second frame in Fig. 12(c)]. Then, the leadinghead overcame the streptavidin blockade and completely
rotated to the arrowhead orientation. Accompanied by thisfurther rotation, the trailing head bound to a forward site of
the actin ﬁlament to become a new leading head, completing
one step. Here, it was clearly revealed that during and before
the completion of a step, the trailing head never interacts

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with actin and hence passively moves forwards, driven by
the rotating leading head. Thus, the ‘‘inchworm’’ mechanism(a molecule moves forward like an inchworm, in which one
head always leads while the other head follows) considered
for kinesin87) as well as the ‘‘biased diﬀusion’’ mechanism(a molecule moves forward by directional diﬀusion according to a potential slope created along the actin helix)
proposed for M588) can be ruled out. The rotation of theleading head is the swinging leverarm motion itself initially
proposed by Hugh Huxley for the powerstroke of muscle
myosin.89)

(a)

(b)

7.3.2 Tension generation

The seemingly spontaneous rotation of the leading headfollowing trailing head detachment suggests that intramolecular tension for the advance already existed in the twoheaded bound molecule. In this bound state, the trailing headis in the arrowhead orientation. This orientation is natural atleast for the ADP-bound or nucleotide-free head. Hence, theintramolecular tension is not produced by the bound trailinghead. On the other hand, the leading head is not in thenatural orientation (i.e., in the reverse arrowhead orientation)and hence pays an energy cost to generate the intramoleculartension. In fact, the bound leading head is slightly curved
outwards, while the trailing head is straight [Figs. 12(a) and12(b)]. Upon trailing head detachment, the constraint
keeping the bound leading head in the unnatural orientationis removed and hence the leading head spontaneously rotates
forwards, meaning that the bound leading head is in astrained pre-stroke state and that the leverarm swing is notaccompanied by chemical transitions.
This inferred view was reinforced by the observation thatthe short coiled-coil of two-headed bound M5-HMM wassometimes unfolded and then the leading head rotated fromthe reverse arrowhead orientation to the arrowhead orientation, very much similarly to the swinging leverarm motion
observed during a step [Fig. 13(a)]. The uncoiling is caused
by intramolecular tension, not by the interaction with theoscillating cantilever tip. This is suggested by the fact that afraction of single-headed species is contained in the sample
of M5-HMM puriﬁed through actin binding and dissociation.90)
In general, during the actomyosin ATPase cycle, thestrained pre-stroke state has been thought to be always
formed through Pi release from an ADP-Pi bound head
weakly interacting with actin.91,92) Thus, it may beconsidered that the strained pre-stroke state, which is formed
directly by the binding of the ADP-bound leading head toactin, does not participate in the forward step in ATP.
However, we observed molecular behavior in ATP thatindicates that it does participate in the step. In two-headed
bound M5-HMM, both of the motor domains frequently
exhibited brief dissociation and reassociation on the sameactin ﬁlament (or a brief translocation by around 5 nm
along the actin ﬁlament), whereas M5-HMM remained
at approximately the same position on the ﬁlament[Fig. 13(b)]. We have termed this behavior ‘‘foot stomp’’.
The foot stomp was more frequently observed at the leadinghead than at the trailing head. The brieﬂy detached leadinghead does not carry bound Pi because Pi must have beenreleased from an ADP-Pi-bound M5 head when it is initially(c)Fig. 13. (Color) Other dynamic events of M5 HMM observed by
HS-AFM. (a) Unfolding of coiled-coil tail of two-headed bound M5-HMM.
Top image, before unfolding; bottom image, after unfolding. (b) Successive
AFM images showing foot stomp at leading head observed in 1 M ATP.
The events of leading head detachment from actin are indicated witharrowheads. Imaging rate, 7 fps. (c) AFM image showing sharply bent
leading head of two-headed bound M5-HMM in the nucleotide-free
condition. The mark ‘‘+’’ indicates the plus ends of actin ﬁlaments.

bound to actin.93) This rapid Pi release is a general featureof actomyosin ATPase reaction. Nevertheless, the detachedleading head with only ADP bound rebinds to actin, still in
the reverse arrowhead orientation, and then swings forward
following trailing head detachment.
The detachment of the leading head during foot stomp is
not caused by ATP binding to the leading head. As describedin the next section, a nucleotide-free leading head is often
sharply bent. We never observed a sharply bent leading head
before foot stomp.
7.3.3 Mechanism of hand-over-hand movement

The leading head of two-headed bound M5-HMM wasstraight (slightly curved outwards) in ADP or ATP, while it
was often sharply bent in the nucleotide-free condition[Fig. 13(c)]. Therefore, just by looking at the conformationof the leading head, we can judge whether or not the leadinghead contains nucleotides. To estimate the ADP dissociation
rate constant at the leading head, M5-HMM bound to actin
at various concentrations of ADP was observed, and then the
lifetime of straight leading head was analyzed as a functionof ADP concentration. The ADP dissociation rate constant at
the leading head was estimated to be 0.1/s. This means thatADP is released from the leading head every 10 s, on
average. However, M5-HMM walks many steps for 10 s,
meaning that ADP does not dissociate from the leading
head. That is, before ADP dissociation from the leading
head, the leading head is switched to the trailing head during
walking. ADP dissociation, the subsequent ATP binding,and the resulting detachment from actin solely occur at thetrailing head. This is the basis underlying the processive
hand-over-hand movement. This mechanism was inferred

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previously from various indirect experiments86,94–98) but isnow clearly and directly demonstrated by the high-speed
AFM observation.
7.4 Summary of imaging studies

As demonstrated in the three imaging studies highlighted
above, the HS-AFM imaging of proteins in action is a
powerful approach to revealing how proteins operate tofunction. Although only from one direction, the wholeﬁgure of a protein molecule and its minute change withtime appear in an AFM movie. Unlike ﬂuorescencemicroscopy, wherein the subset of molecular events
observed is dependent on the site where the probe isplaced, various dynamic events of a molecule appear in amolecular movie without planned selection, meaning thatboth expected and unexpected dynamic molecular behaviors
can appear simultaneously. Therefore, we can have a greatchance of learning several facts of molecular action in
one imaging experiment. Moreover, molecular actions thatappear in a video can often be interpreted straightforward
without sophisticated analysis, leading to the drawing ofconvincing and irrefutable conclusions. These general
excellent features of HS-AFM imaging signiﬁcantly facilitate and accelerate our understanding of the functional
mechanism of proteins.
8. Outlook

Continued studies on a wide range of interesting dynamics
of puriﬁed biological molecules will be actively carried out
using HS-AFM. HS-AFM will be used routinely in the near
future, similarly to the historical path taken by conventional
AFM. However, we need to explore dynamic eventsoccurring not only with puriﬁed molecules but also withthose on the surfaces of live cell membranes and organelles.
The plasma membranes of live eukaryotic cells are
extremely soft, and hence, largely deformed by contact
with a cantilever tip before appreciable deﬂection of thecantilever is detected. Therefore, at present, it is impossible
to image molecules on the membranes using AFM.
However, the outer surfaces of live bacterial cells andorganelles such as mitochondria and nucleus are rather
rigid. Nonetheless, molecular architectures on these surfaces
have not been studied much using AFM. We can only ﬁnd a
few examples of such imaging studies, e.g., nuclear porecomplexes on a nuclear envelop,99,100) the hexagonal
S-layer on the surface of Corynebacterium glutamicum,101)
and the peptidoglycan organization on the surface ofLactococcus lactis.102) Since dynamic molecular eventsoccurring on these surfaces are largely unknown, it isworthwhile to attempt to image them using HS-AFM. This
direction of studies will be more actively performed.
As mentioned above, the plasma membranes of live
eukaryotic cells are extremely soft. This is due to the wide
area of the membranes. For small structures, such as
dendritic spines (0:5 m in diameter) where most excitatory synapses reside, we may be able to visualize
individual proteins on their surfaces. Even without molecular resolution, observing dynamic morphological changes insuch cellular structures is intriguing and meaningful becauseof their inaccessibility from other approaches. For example,
spine enlargement associated with long-term synaptic

potentiation,103,104) a cellular model of learning and memory,takes place together with the exocytosis and endocytosis ofvesicles.105–107) However, the detailed dynamics of membrane traﬃcking during spine growth are unknown.
To visualize molecules on large eukaryotic cell surfaces,
we need a noncontact condition. The noncontact imaging
capability in liquids has already been achieved by scanningion-conductance microscopy (SICM).108) Owing to theprogress of fabrication techniques for producing very sharp
glass capillaries with a small pore at the apex, the spatial
resolution of SICM has reached a few nm.109) Immobile
protein molecules with a size of 14 nm on living cell
membranes have been successfully imaged.110) Moreover,
hopping-mode SICM, which is similar to the previously
developed AC-mode SICM,111) has recently been designed
to prevent the collision of the capillary electrode probe withtall samples.112) However, the bandwidth of ion-conductancedetection cannot easily be increased, because of the highimpedance to ionic current through the small pore of thecapillary electrode as well as of slow ionic currents inaqueous solutions. Nevertheless, we think that increasing the
bandwidth is not impossible. Scanning tunneling microscopy
has a high impedance (hence a low bandwidth) problem but
it has been overcome.113) By adapting our high-speed
scanning setup based on fast scanner and control techniquesachieved in HS-AFM, various types of high-speed scanningprobe microscopy techniques including high-speed SICM
will be developed in the future as innovative tools forbiological research.
Acknowledgements

We would like to thank Daisuke Yamamoto, Hayato
Yamashita, Mikihiro Shibata, Ryota Iino, Hiroyuki Noji,Kiyohiko Igarashi, Hideki Kandori, and the many present
and past students in our lab for collaborations or discussions
on topics described in this review. Long-term ﬁnancial
support by NEDO, JST (CREST Project), JSPS (Grant-inAid for Basic Research S), MEXT (Knowledge Cluster
Initiative Project), and the Mitsubishi Foundation is also
gratefully acknowledged.

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