BB42CH17-Ando

ARI

3 April 2013

15:19

ANNUALREVIEWS

Further

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Click here for quick links to
Annual Reviews content online,
including:• Other articles in this volume• Top cited articles• Top downloaded articles• Our comprehensive searchHigh-Speed AFM andApplications to Biomolecular
Systems
Toshio Ando,1,2 Takayuki Uchihashi,1,2and Noriyuki Kodera2
1

Department of Physics and 2 Bio-AFM Frontier Research Center, Kanazawa University,
Kanazawa 920-1192, Japan; email: tando@staff.kanazawa-u.ac.jp

Annu. Rev. Biophys. 2013. 42:393–414

Keywords

The Annual Review of Biophysics is online atbiophys.annualreviews.org

atomic force microscopy, proteins in action, high-resolution imaging,
dynamics of biomolecules, nanostructural dynamics

This article’s doi:
10.1146/annurev-biophys-083012-130324
c 2013 by Annual Reviews.
Copyright All rights reserved

Abstract
Directly observing individual protein molecules in action at high spatiotemporal resolution has long been a holy grail for biological science. This isbecause we long have had to infer how proteins function from the static
snapshots of their structures and dynamic behavior of optical makers attached to the molecules. This limitation has recently been removed to alarge extent by the materialization of high-speed atomic force microscopy(HS-AFM). HS-AFM allows us to directly visualize the structure dynamics
and dynamic processes of biological molecules in physiological solutions,at subsecond to sub-100-ms temporal resolution, without disturbing theirfunction. In fact, dynamically acting molecules such as myosin V walking onan actin ﬁlament and bacteriorhodopsin in response to light are successfully
visualized. In this review, we ﬁrst describe theoretical considerations for thehighest possible imaging rate of this new microscope, and then highlightrecent imaging studies. Finally, the current limitation and future challenges
to explore are described.

393

BB42CH17-Ando

ARI

3 April 2013

15:19

Contents

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 394
FEEDBACK OPERATION AND MAXIMUM POSSIBLE IMAGING RATE . . . . . 395
Feedback Operation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396
Other Factors Limiting Feedback Frequency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397
Maximum Possible Imaging Rate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398
CANTILEVER, SAMPLE STAGE, AND SUBSTRATE SURFACE . . . . . . . . . . . . . . . 398
Small Cantilevers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 398
Sample Stage and Hydrodynamic Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400
Substrate Surfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400
IMAGING OF PROTEINS IN ACTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401
Myosin V Walking on Actin Filament . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402
Rotary Catalysis of Rotorless F1 -ATPase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405
FUTURE CHALLENGES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 407
Wide-Area Observation and In Situ Imaging of Dynamic Processes . . . . . . . . . . . . . . . 407
Faster Wide-Area Observation and Dynamic Imaging of Cell Morphology . . . . . . . . 408
High-Speed Noncontact Imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 408

INTRODUCTION

AFM: atomic force
microscopy

394

Atomic force microscopy (AFM) was originally invented to visualize atoms on solid surfaces (10).
In biological sciences, this microscopy is now routinely used to directly acquire high-resolution
images of biological samples under physiological conditions, without sample staining (66, 81, 82).
AFM is also used for the recognition and localization of speciﬁc molecules (37, 91) as well as force
measurements to estimate the strength of intra- and intermolecular bonds at the single-molecule
level (26, 67, 108), the elasticity of biological surfaces (19, 98), and the osmotic pressure of live
cells (90). Therefore, AFM is a nano-toolbox for biology (reviewed in References 64 and 65).
However, the limited scan speed of AFM limits its usefulness. It takes time (at least 30 s) to acquire an image, and therefore, molecules moving on the substrate surface are imaged as a bluror cannot be imaged at all. To overcome this limitation, the development of high-speed AFM(HS-AFM) was started around 1993. Through the initial prototypes (5, 103, 104) and their extensive improvements (8, 9, 24, 25, 29, 47, 50, 51, 53, 101), HS-AFM is now materialized (3, 7).
Although the speed performance depends on imaging conditions, current HS-AFM can generally
capture an image of biological molecules within 100 ms or less. Importantly, the structure and function of fragile molecules are not disturbed by the interaction with a cantilever tip. This high-speed
and low-invasive performance opens up a new opportunity to visualize dynamically functioning
biological molecules in great detail. As has been demonstrated in recent imaging studies (12, 40,
52, 86, 99), the visualized dynamic images of proteins can provide information inaccessible withother approaches, giving great insight into how the proteins function. Remarkably, the dynamic
images can be interpreted straightforwardly without intricate analyses and interpretations, making
it possible to attain ﬁrm conclusions. This review summarizes the fundamentals of HS-AFM, highlights recent imaging studies of proteins, and outlines ongoing and future challenges to expand
the scope of its application to biological studies.
Ando·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 2013

15:19

Laser diodeSplit photodiode

Piezoactuator
for excitation

MirrorCantilever base

Focusing
lens

Differential
amplifierSine wave
generator

Amplitude
detector

Cantilever

Freq.

Amp.

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Sample stageSample-stage
scannerSet pointz

P- Gain

I- GainyMonitorx

D- Gain

z

y

x

DA-y

DA-x

PID-feedback
controller

AD-zComputer
Piezo driver

Figure 1
Schematic for general conﬁguration of a tapping-mode AFM system (see text for details).

FEEDBACK OPERATION AND MAXIMUM POSSIBLE IMAGING RATEVisualization by AFM of the topography of a sample placed on a substrate surface requires
acquisition of sample height information over many points on the sample surface (Figure 1;
Supplemental Movie 1, follow the Supplemental Material link from the Annual Reviews
home page at http://www.annualreviews.org). A stylus probe attached to the free end of a
cantilever is brought into contact with the sample. Mechanical response of the cantilever uponthis contact is measured and then the sample stage is ﬁnally moved in the z-direction to recover
the mechanical state of the cantilever back to a given condition (i.e., set point) through feedback
control. For this recovery, the closed 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 repeated many times for different sample surface points during lateralscanning of the sample stage. Among several imaging modes of AFM (reviewed in References
31 and 63), the tapping mode (120) (also called amplitude modulation mode or intermittent
contact mode) is often used for biological samples. In this mode, the cantilever is oscillated in thez-direction at its ﬁrst resonant frequency (Supplemental Movie 1) so that the tip intermittently
taps the sample surface. This tapping results in decreased amplitude as well as a phase shift
relative to the excitation signal (101). The intermittent contact can eliminate friction force duringlateral scanning, which minimizes deformation of fragile biological samples. Described below
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

Set point: the target
value of a controlled
variable395

BB42CH17-Ando

ARI

3 April 2013

15:19

Parachuting

λSample surface

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

h0
Tracing error
VS

Scanner trajectory
Figure 2
Scanner movement tracing sinusoidally shaped sample surface with periodicity λ and amplitude h0 /2, when no parachuting occurs. The
inset shows a trajectory of the bottom swing position of the tip (broken line) relative to the sample surface when parachuting occurs.

is the feedback operation to maintain the cantilever oscillation amplitude (and thus tip-sample
interaction force) constant. Next, the highest possible imaging rate of tapping-mode HS-AFM
is derived as a function of the feedback bandwidth and various parameters.

Feedback Operation
In the closed feedback loop (see Figure 1), the input to the loop is the variation of sample heightright under the cantilever tip and the output is the displacement of the z-scanner. The output fromthe feedback controller is used as a signal representing the sample height. There are several devices
in the feedback loop: a cantilever, an optical beam deﬂection (OBD) detector for the detectionof cantilever deﬂection, an amplitude detector (deﬂection-to-amplitude converter), a feedback
controller, a piezoelectric driver, and a piezoactuator-based z-scanner. Each of these devices hasa certain time delay in the response to the corresponding input signal. Therefore, the feedback
control cannot perform without time delay. This delay deteriorates the speed performance of the
microscope, which is described below.
For simplicity, it is assumed that the sample surface proﬁle in the x-z plane has a sinusoidal
shape with a periodicity λ and an amplitude h0 /2 (Figure 2, black line). When the sample-stage
scanner is moved in the x-direction at velocity Vs , the sample height h right under the cantilevertip changes with time as
OBD: optical beam
deﬂection

396

h(t) = (h 0 /2) × sin(2π f t),

1.

where f = Vs /λ. During feedback scan, the z-scanner moves in the direction opposite to the sample
height so that the sample surface looks ﬂat when viewed from the cantilever. However, because
Ando·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 201315:19

of the time delay (τ 0 ) in the closed feedback loop, the z-scanner moves at feedback frequency f asz(t) = −(h 0 /2) × sin(2π f t − θ),2.

where θ = 2π fτ 0 (Figure 2, red line). Therefore, the sample surface when viewed from thecantilever does not look perfectly ﬂat but varies with time as

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

S(t) = h(t) + z(t) = h 0 sin(θ/2) cos(2π f t − θ/2).

3.

Quality factor: ameasure representing
small damping (energy
loss per cycle) in aresonant system

The deviation from a ﬂat surface (i.e., feedback error) due to the feedback delay is indicated withthe blue line in Figure 2. When cantilever free oscillation amplitude and the amplitude set point
are set at A0 and As (= rA0 ; 0 < r < 1), respectively, the cantilever tip in contact with the sample
surface is pushed upward by the sample to an extent of D(t) ≡ S(t) + A0 (1 − r) so long as the
sample is rigid enough compared with the cantilever.
The feedback bandwidth fB , which is a value characterizing the speed performance of feedback
operation in the AFM system, is usually deﬁned by a feedback frequency at which 45◦ phase delay
occurs in the feedback scan. Thus, fB = 1/(8τ 0 ). Because the feedback frequency f should notexceed fB , a condition f < fB holds, which limits the scan speed Vs as Vs < λfB .

Other Factors Limiting Feedback Frequency
However, two other factors limit f more severely than fB . One comes from the condition that D(t)should always be positive. Otherwise, at downhill regions of the sample, the cantilever tip cannot
make contact with the sample surface at the bottom swing of oscillation; i.e., parachuting occurs(Figure 2, inset). Once detached, the error signal is saturated at A0 (1 − r). When r is set close to
1 so that the tapping force exerted from the oscillating cantilever tip to the sample is minimized,
the saturated error signal is small, and hence the parachuting time is prolonged (51, 92). During
parachuting, the sample topography cannot be recorded at all. The condition under which noparachuting occurs is expressed as
A0 (1 − r) − h 0 sin(θ/2) > 0.

4.

To minimize the tapping force, small A0 has to be used in addition to r close to 1. For example,
for A0 = h0 /5 and r = 0.9, Equation 4 limits the phase delay as θ < 2.3◦ , which severely
limits the feedback frequency f as f < 0.05 fB . The other condition that limits f comes from the
maximum possible tapping force under which the structure and function of the biological sample
are retained. The maximum tapping force F pmax is exerted at an uphill region of the sample, which
can be approximately expressed as
Fpmax = (kc /Q c ) × [A0 (1 − r) + h 0 sin(θ/2)],

5.

where kc and Qc are the spring constant and the quality factor in water of the cantilever, respectively.
For example, under the condition of F pmax = 100 pN, kc = 200 pN/nm, Qc = 2, A0 = 1 nm,
h0 = 5 nm, and r = 0.9 [these values are realistic ones as values close to these values are actuallyused in successful imaging of functioning proteins (40, 52, 86, 99)], Equation 5 limits the phasedelay as θ < 20.7◦ , which moderately limits f as f < 0.46fB .
Thus, the parachuting problem is the severest limiting factor for the maximum possible feedback frequency and hence for the scan speed Vs . However, we have successfully eliminated thisproblem by developing a feedback controller that can automatically change its gain parameters
depending on the cantilever oscillation amplitude during imaging; feedback gain is increased at
the downhill region of the sample so that parachuting only rarely occurs (51). Even when it occurs,
the parachuting time is reduced signiﬁcantly.
www.annualreviews.org • High-Speed AFM for Filming Biomolecules397

BB42CH17-Ando

ARI

3 April 2013

15:19

Here, note that the mechanical quantity affecting the sample (i.e., causing momentum change)
is not the tip force itself acting on the sample but the impulsive force (the product of the force acting
on the sample and the time for which the force acts). When the cantilever resonant frequency ishigh (∼1 MHz in water), as is the case for small cantilevers optimized for HS-AFM, the force
acting time is approximately 100 ns or less, which guarantees no signiﬁcant effect of F pmax =100 pN on fragile protein molecules.

Maximum Possible Imaging Rate

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

The maximum possible imaging rate Rmax is a function of fB and the imaging condition (the scansize in the x-direction W, the number of scan lines N, and the spatial frequency of sample height
corrugation to be imaged 1/λ) as well as of the maximum possible phase delay θ max in the feedback
operation that depends largely on the sample fragility. For given W, N, and Vs , one frame ofimage is captured with time T = (2WN)/Vs . The maximum possible scan speed Vs max is given byθ max /(π /4) × λfB . Thus, Rmax is given by
Rmax = 2θmax λ fB /(π W N ).

6.

For example, under a realistic condition for imaging protein molecules by our HS-AFM ( fB =110 kHz, θ max = π /9 (i.e., 20◦ ), λ = 10 nm, W = 150 nm, and N = 100), Equation 6 gives
Rmax = 16.3 frames per second (fps). The high fB achieved (110 kHz) is approximately 1,000 times
higher than that of conventional AFM systems.

CANTILEVER, SAMPLE STAGE, AND SUBSTRATE SURFACE
HS-AFM is materialized by the achievement of three conditions to meet high-speed and lowinvasive performance: (a) extensive reduction of time delays in the response of all devices containedin the feedback loop (reviewed in Reference 7), (b) damping of mechanical vibrations caused by
fast displacement of the scanner (7, 53), and (c) feedback control technique that can eliminate
tip parachuting even when As is set close to A0 (51). Details of various techniques and devices
that have achieved these conditions are described elsewhere (7). The latest HS-AFM instrument
developed by us is commercially available from the Research Institute of Biomolecule Metrology
Co., Ltd. (Tsukuba, Japan). Here, we therefore focus on devices (small cantilevers, sample stage,
and substrate surfaces) and associated issues that HS-AFM users themselves must deal with.

Small Cantilevers
When a cantilever tip taps the sample surface, a brief stepwise force is exerted on the cantilever.
The cantilever responds to this force with a response time τ c = Qc /π fc , where fc is the ﬁrst
resonant frequency of the cantilever in water. Measuring the cantilever oscillation amplitude
requires time, at least τ m = 1/2fc . Thus, the resonant frequency fc should be as high as possible,
whereas kc should be small for fragile biological samples. Qc can be naturally small in water. The
size of the cantilever has to be small to achieve both high fc and small kc (50, 104). Several typesof small rectangular cantilevers made of Si3 N4 have been developed by Olympus (Tokyo, Japan)
in collaboration with our group. The high-end small cantilevers (Olympus BL-AC7DS-KU5,
custom-made) that we have been routinely using for imaging studies are 6–7 μm long, 2 μm wide,and 90 nm thick (Figure 3a). They have fc = 1.2 MHz in water (3.5 MHz in air), Qc ≈ 2 in water,
and kc = 0.2 N/m. The commercially available small cantilevers (Olympus BL-AC10DS-A2) are
9–10 μm long, 2 μm wide, and 130 nm thick, and have fc = 0.6 MHz in water (1.5 MHz in
398

Ando

·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 2013

15:19

a

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

1 μm

b

1 μm

c
Cantilever baseCantileverMica disk

Figure 3
Cantilever, tip, and relative arrangement of cantilever and sample stage. (a) Scanning electron microscope(SEM) image of a small cantilever. (b) SEM image of an electron-beam-deposited (EBD) tip grown on an
original bird-beak-shaped tip. (c) Relative arrangement between sample stage and cantilever forcircumventing the disturbance by hydrodynamic pressure produced by quick displacement of the z-scanner.

air), Qc ≈ 2 in water, and kc = 0.1 N/m. Small cantilevers are also available from NanoWorld(Neuchâtel, Switzerland).

Small cantilevers have other advantages. The total thermal noise (kB T/kc )1/2 (where kB is
Boltzmann’s constant and T is the temperature in Kelvin) (2) is distributed over frequencies up toslightly above fc . Therefore, a cantilever with a higher fc has a lower noise density. In the tapping
www.annualreviews.org • High-Speed AFM for Filming Biomolecules399

BB42CH17-Ando

ARI

3 April 2013

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

EBD: electron beam
deposition (deposited)

15:19

mode, the frequency region used for imaging is approximately the feedback frequency centered on
the resonant frequency. Thus, for a cantilever with a higher fc , thermal noise has less effect on theamplitude measurement. The deﬂection of a cantilever alters the angle of its free end (Δϕ), which
is detected by the OBD detector (see Figure 1). Therefore, for a given displacement Δz of thecantilever free end in the z-direction, a shorter cantilever results in a larger angle change (Δϕ =3Δz/2L) and thus gives higher displacement detection sensitivity. Because of a small thermal noise
effect and this high sensitivity with small cantilevers 6–10 μm long, the displacement as small as<0.1 nm can be detected even with high bandwidth detection (a few megahertz). The deﬂectionto-amplitude conversion can be performed every half or one cycle of cantilever oscillation using
fast amplitude detectors developed (5, 7).
The bird-beak-shaped tip of the small cantilevers produced by Olympus is approximately
1.5 μm long and its apex radius is 15–24 nm. Small cantilevers that have an additional carbon
nanoﬁber tip with an apex radius <10 nm grown on the bird-beak-shaped tip have become commercially available (Olympus, BL-AC10FS-A2) (49, 97). When a scanning electron microscope(SEM) is available, electron-beam-deposited (EBD) tips can be grown under an atmosphere of gas
sublimated from materials. Various materials can be used for the deposition of carbon, platinum,
tungsten, and silicon dioxide. A gas injection system, which is adaptable to SEMs and can mix
different gases under computer control, is commercially available (Omniprobe OmniGISTM ). The
grown tips can be sharpened by nitrogen or oxygen plasma etching. The tip apex radium is usually
reduced to ∼5 nm and sometimes to ∼0.5 nm (100).

Sample Stage and Hydrodynamic Pressure
In high-speed imaging, the sample-stage z-scanner is displaced at high frequencies (20–100 kHz).
This fast scan exerts a hydrodynamic pressure on the cantilever and its supporting base placed inclose vicinity to the sample surface, which tends to move them, particularly when the sample stage
is large (4). Their movement results in a slow response of the cantilever oscillation amplitude to zscanner displacement. In the worst case, this delay extends to a few microseconds (6, 100), which ismuch longer than the minimized time delay in the closed feedback loop. As a sample stage, a small
glass rod 1.5–2 mm in diameter and 2 mm high is routinely used for HS-AFM imaging (Figure 3c).
To further circumvent the disturbance by hydrodynamic pressure, the cantilever is positioned
in a way that the whole cantilever chip minimally overlaps with the sample stage when viewed fromthe top (Figure 3c). Moreover, the cantilever tip longer than ∼2.5 μm is prepared by the growthof a >1-μm-long EBD tip on the original tip (Figure 3b). When the tip is shorter, the cantilevergets closer to the sample surface at the bottom of swing and thus the solution conﬁned between
them becomes more squeezed, resulting in oscillation damping and the deteriorated detection
sensitivity of tip-sample interaction (6).

Substrate Surfaces
A substrate surface, on which a sample is placed, holds the key to successful HS-AFM imaging(reviewed in References 109 and 111). The surface should be ﬂat enough so that the molecules ofinterest deposited on it can be easily identiﬁed. Observing dynamically acting protein molecules
requires the substrate surface to loosely bind the molecules to allow them to retain their physiological function. However, the sample-surface interaction is essential to avoid too rapid Brownian
motion of the sample, particularly for single-protein molecules isolated completely from othermolecules. Observing dynamic interactions between different proteins often requires selective
protein attachment to a surface. As substrate surfaces, bare or chemically treated mica surfaces,
400

Ando

·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 2013

15:19

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

planar lipid bilayer (PLB) surfaces, and the surfaces of two-dimensional crystals of streptavidin
formed on a biotin-containing PLB have been used for dynamic AFM imaging. The propertiesof these surfaces are summarized below.
Mica surfaces. Mica (natural muscovite or synthetic ﬂuorophlogopite) has been frequently usedas 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 DNA and various
proteins by electrostatic interaction. We can control the adsorption strength by varying the ionicstrength or pH or by adding divalent cations such as Mg2+ and Ni2+ (especially for attaching negatively charged samples) (17, 72). Monovalent cations markedly change the afﬁnity; Li+ > Na+ > K+
for every protein (18). For fractionated membranes containing membrane proteins, the bare mica
surface is useful. A water layer of ∼1 nm thickness separates the membranes from the mica surface,enabling the motion of membrane proteins within the membranes (46, 76, 113). Proteins, including
proteins in membranes, can be covalently immobilized onto chemically treated mica surfaces (43).

Ni–NTA:nickel–nitrilotriaceticacid
DOPC: dioleoylphosphatidylcholine
P2X4 receptor: oneof the ATP-gatedmembrane cation
channels
OmpF: outermembrane protein F

Planar lipid bilayer surfaces. PLB can easily be formed on a bare mica surface by the depositionof liposomes (58). The PLB surfaces can be used for both speciﬁc and electrostatic immobilizations of proteins. For the speciﬁc immobilization of biotinylated proteins and His-tag-conjugatedproteins, lipids with biotin and nickel–nitrilotriacetic acid (Ni–NTA) at the polar head groups canbe used, respectively. Multiple-point pinning is sometimes required for stopping the rapid motionof molecules tethered to the surfaces. PLB surfaces prepared with electrically neutral phospholipids are resistant to the nonspeciﬁc binding of proteins. For electrostatic immobilization, lipids
with charged head groups can be used. Unlike bare mica surfaces, the surface charge density andpolarity can be varied by using different fractions of a charged lipid and by using positively or
negatively charged lipids, respectively.
Streptavidin two-dimensional crystal surfaces. The two-dimensional crystals of streptavidin
can easily be formed on highly ﬂuidic PLBs containing lipids with unsaturated alkyl chains (such as
DOPC, dioleoylphosphatidylcholine) and a biotin-containing lipid (11, 20). Their surface is particularly useful for the selective and stable immobilization of homo-oligomeric protein complexes
because pinning the complexes at multiple biotinylated sites is possible (109). Not only biotinylated samples but also His-tag-conjugated samples can be immobilized using biotin-Ni-NTA
compounds as a linker. Importantly, the surface is resistant to the nonspeciﬁc binding of proteins.

IMAGING OF PROTEINS IN ACTION
The imaging studies conducted thus far have covered a wide range of dynamic molecular events(reviewed in References 3, 14, and 44). These events are classiﬁed into (a) structure dynamics
of proteins including myosin V on actin (52), rotorless F1 -ATPase (99), P2X4 receptors (87),bacteriorhodopsin (bR) (85, 86), Ca2+ pump (118), and intrinsically disordered FACT protein(61); (b) self-assembly processes including amyloid-like ﬁbril formation from cleaved lithostathine(57) and PLB formation (3, 33); (c) dynamic protein-protein interactions including GroELGroES (8, 109, 119), membrane-mediated association between c-rings of ATP synthase (15),bR trimer-trimer (13, 113), and DNA-histone (60, 94); (d ) diffusion processes including OmpF
reconstructed in lipids at high density (12), porin trimers on a live bacterial cell surface (112),
defects in protein two-dimensional crystals (110), and DNA-bound Rad54 (79); (e) molecular
processes associated with enzymatic reactions including cellulase hydrolyzing cellulose ﬁbers (39,40) and DNA restriction-modiﬁcation enzymes (32, 93); and ( f ) dynamics occurring with DNA
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

401

BB42CH17-Ando

ARI

3 April 201315:19

origamis (22, 23, 80, 96, 107). Below we give two examples of imaging studies of proteins inaction to demonstrate the power of HS-AFM.
HMM: heavy
meromyosin(tail-truncated myosin)

Myosin V Walking on Actin FilamentAnnu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Double-headed myosin V (M5) functions as a cargo transporter in cells (reviewed in 83) and moves
processively along an actin ﬁlament toward the plus end of the ﬁlament (56, 77) in a hand-overhand
manner with a 36-nm advance (28, 106, 116) for every ATP hydrolysis cycle (78). Hand-overhandmeans that the two heads step alternately, exchanging leading and trailing roles at each step, verymuch like walking. We visualized the walking molecules by HS-AFM (52). Below, we describe
an experimental setup for this visualization and what we can learn by closely looking at walking
molecules.
Selection of substrate surface. Partially biotinylated actin ﬁlaments were immobilized on thesurface of PLB containing an electrically neutral phospholipid, a biotin lipid, and a positively
charged lipid, through streptavidin with a low surface density. When the positively charged lipid
was absent, tail-truncated M5 (M5-HMM) was never bound to the surface and only interactedwith the immobilized actin ﬁlaments to move unidirectionally. The velocity of the movement was
identical to that measured by ﬂuorescence microscopy under the same buffer solution. However,most of the molecules were moving, orienting perpendicularly to the surface, so that their structure
was not well resolved (Figure 4a). When a positively charged lipid was included in the bilayer at
an appropriate density, we could observe the characteristic sideways topography of the molecules
processively moving with ∼36-nm steps at slightly lower velocity (Figure 4b; Supplemental
Movie 2). However, we could not see detailed molecular behavior during a step because it wascompleted within a frame time (1/7 s). To slow down the step, streptavidin molecules were further
placed on the substrate surface as moderate obstacles to the advance. This method allowed thevisualization of stepping processes as shown in Figure 4c,d (see also Supplemental Movie 3).
No effect of tip-sample interaction on motor activity. When streptavidin molecules as moderate obstacles were absent, we could continuously track a moving molecule (up to ∼20 steps) by
shifting the scan area manually, while observing the scanned images on a computer display (Supplemental Movie 4). The undegraded velocity of the observed movement for a long distanceindicates no effect of the tip-sample interaction on the motor activity. Let us discuss this issue ina way different from that mentioned in Other Factors Limiting Feedback Frequency, above. In
this imaging with a scan size of 150 × 75 nm2 , at 7 fps, we used a cantilever having fc ≈ 1 MHz inwater, kc ≈ 0.2 N/m, and Qc ≈ 2 in water. The oscillating cantilever tip therefore taps the sample
surface 12.7 times per 1 × 1 nm2 . As the two-dimensional size of the motor domain is roughly
5 × 5 nm2 , the motor domain is tapped with the tip ∼320 times during one frame time (1/7 s) andmuch more (>50,000 times) during the successive imaging, clearly indicating no accumulation
but quick dissipation of the energy given to the molecule by the tapping.
In this imaging, the cantilever free oscillation amplitude A0 and amplitude set point As were set
at ∼1 nm and 0.8–0.9 As , respectively. On average, the energy of the oscillating cantilever dissipates
after every tapping by 1/2kc (A0 2 − As 2 )/Qc = 2.3–4.4 kB T, where T is 300 K. Even when theoscillation energy is completely lost, its energy loss is only 24 kB T (similar to the energy of ATPhydrolysis, ∼20 kB T ). Even if the motor domain is mechanically excited by this amount of energy,
the energy would quickly dissipate into many degrees of freedom including those of surrounding
water molecules. Thus, so long as small free oscillation amplitude is used for a cantilever with
402

Ando

·

Uchihashi·

KoderaBB42CH17-AndoARIa

3 April 2013

15:19

b
147 msd
147 ms

147 ms++1,174 msc

1,027 ms

Streptavidin

1+

293 ms2

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

1,320 ms

1,320 ms

440 ms3

1,467 ms

2,201 ms

587 ms
– end

+ end

e
1,614 ms

2,347 ms

734 ms1,760 ms

2,494 ms

50 nm

14.99 s+15.33 s3,374 ms

3,814 ms

50 nm

30 nm

30 nm

Figure 4
Walking tail-truncated myosin V (M5-HMM) and unfolding of coiled-coil tail captured by high-speed atomic force microscopy(HS-AFM). (a) Successive AFM images showing processive movement of M5-HMM in 1 μM ATP when positively charged lipid isabsent on the planar lipid bilayer (PLB) surface. Frame rate, 7 fps. (b) Successive AFM images showing processive movement of
M5-HMM in 1 μM ATP when positively charged lipid is present on the PLB surface. Arrows indicate the coiled-coil tail pointing to
the minus end of actin. (c) Successive AFM images showing hand-overhand movement in 1 μM ATP. The swinging lever is highlighted
with a thin white line. (d ) Schematic explaining the images in panel c. (e) Unfolding of the coiled-coil tail of two-headed bound
M5-HMM. Top image, before unfolding; bottom image, after unfolding. The symbol “+” indicates the plus ends of actin ﬁlaments.
The arrowheads show some of streptavidin molecules. Vertical dashed lines show the centers of mass of the motor domains. The frame
rates used are 7 fps for panels a–c and 3 fps for panel e. The z-scales are 18 nm for panel a and 10.5 nm for panels b, c, and e. Adaptedwith permission from Reference 52.

www.annualreviews.org • High-Speed AFM for Filming Biomolecules403

BB42CH17-Ando

ARI

3 April 201315:19

kc ≈ 0.2 N/m, the function of M5-HMM (and other proteins) is not disturbed by the oscillating
tip even when tapped many times.

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Spontaneous swing of leading head in hand-overhand movement. As shown in Figure 4c
and Supplemental Movie 3, after trailing head detachment, the leading head appeared to spontaneously rotate from the reverse arrowhead orientation toward the arrowhead orientation (theterm arrowhead originates from the conﬁguration of single-headed myosin bound to an actin
ﬁlament in the rigor state). Before completing this rotation, the leading head brieﬂy halted bycolliding with a streptavidin molecule placed in the way of its natural path, and the detachedtrailing head was most distant from the actin ﬁlament and slightly rotated around the neck-neckjunction. Then, the leading head overcame the streptavidin blockade and completely rotated tothe arrowhead orientation. Accompanied by this further rotation, the trailing head was bound to a
forward site of the actin ﬁlament to become a new leading head, completing one step. Here, it was
clearly revealed that before the completion of a step, the trailing head never interacted with actin
but passively moved forward, driven by the rotating leading head. The rotation of the leadinghead is exactly the swinging lever arm motion proposed by Huxley (38) for the powerstroke ofmuscle myosin, a hypothesis that existed for a long time without clinching evidence.
Foot stomp and unwinding of the coiled-coil tail. The seemingly spontaneous rotation of
the leading head following trailing head detachment suggests that intramolecular tension for theadvance has already existed in the two-headed bound molecule. In this bound state, the trailinghead is in the arrowhead orientation, which is natural at least for the ADP-bound or nucleotide-freehead. Nevertheless, the leading head is not in the natural orientation (i.e., in the reverse arrowheadorientation) and therefore pays an energy cost to generate the intramolecular tension, which is
implied in the slightly curved appearance of the leading head. Upon trailing head detachment, theconstraint keeping the bound leading head in the unnatural orientation is removed and hence theleading head spontaneously rotates forward, meaning that the bound leading head is in a strained
prestroke state and that the lever arm swing is not accompanied by chemical transitions.
Interestingly, during the two-headed bound state in ATP, the motor domain of the leading headfrequently exhibited brief dissociation and reassociation on the same actin ﬁlament, whereas themolecule remained at approximately the same position on the ﬁlament (Supplemental Movies 3
and 4). Similarly, the motor domain of the trailing head exhibited a brief translocation by ∼ ± 5 nm
along the actin ﬁlament. We termed these behaviors foot stomp. The foot stomp was observed
more frequently at the leading head than at the trailing head (approximately 3:1). Although not
well documented, a foot-stomp-like behavior was previously suggested in ﬂuorescence microscopy
observations of walking myosin V molecules (89, 95). Thus, the foot stomp is an inherent behaviorof myosin V.
The foot stomp at the leading head seems to raise an important issue of the chemomechanicalcoupling in this motor. The brieﬂy detached leading head does not carry bound Pi because Pi
release occurs immediately after the initial binding of the ADP–Pi-bound head to actin (21).
Nevertheless, the detached leading head with only ADP bound rebinds to actin still in the reversearrowhead orientation, and then swings forward following trailing head detachment, indicatingthat tension generation for forward movement can occur without transitioning through an ADP–
Pi-bound state. It can occur in the ADP-bound state. Thus, the tension generation for forward
movement does not seem to require that chemical energy be supplied by ATP hydrolysis.
During the two-headed bound state in ADP, the short coiled-coil tail was sometimes unwound,
immediately after which the monomerized leading head rotated toward the arrowhead orientation,
similar to the swinging lever arm (Figure 4e). Again, this unwinding suggests that the distortion
404

Ando

·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 2013

15:19

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

of the actin-bound leading head, which does not require chemical energy, is the source of theintramolecular tension generation for forward movement.
Mechanism of hand-overhand movement. The leading head of two-headed bound M5-HMM
was straight (slightly curved outward) in ADP or ATP, whereas it was often sharply bent in the
nucleotide-free condition. Therefore, the conformation of the leading head indicates whether
the leading head contains nucleotides. From AFM movies of actin-bound M5-HMM in various
concentrations of ADP, the ADP dissociation rate constant at the leading head was estimated to be
0.1 s−1 . Thus, ADP 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. ADP
dissociation, the subsequent ATP binding, and the resulting detachment from actin solely occur
at the trailing head. This is the basis underlying the processive hand-overhand movement. Thismechanism was inferred previously from various indirect experiments (27, 70, 73, 75, 78, 102) but
is now clearly and directly demonstrated by the HS-AFM observation.

Rotary Catalysis of Rotorless F1 -ATPase
The α3 β3 γ subcomplex of F1 -ATPase (a part of ATP synthase) is the minimum complex forthe full ATPase activity. About half the length of the long γ subunit is inserted into the central
cavity formed by a ring-shaped α3 β3 where three α subunits and three β subunits are arranged
alternately (1). Three ATP binding sites locate at the α-β interfaces, mainly in the β subunits.
The α3 β3 γ subcomplex is a rotary motor (34, 48, 68, 115) (Figure 5a). The γ subunit rotatesin the stator α3 β3 ring driven by rotary hydrolysis of ATP at the three β subunits. The rotation
occurs in the counterclockwise direction as viewed from the exposed side of the γ subunit (or fromthe C-terminal side of α3 β3 ). In the ATPase cycle, three β subunits take different chemical states:
ATP-bound, ADP-bound, and nucleotide-free (empty) states (1, 34). Each chemical state cyclically
propagates over the three β subunits. Thus, there is strong cooperativity between β subunits.
How is the cooperativity essential for torque generation to rotate the γ subunit engendered
without direct contact between the β subunits? In every instance, the β-γ interaction is different
among the three β subunits because the γ subunit has no symmetry. In consideration of thisfact, it was proposed that interactions with the γ subunit control the conformational and catalytic
states of individual β subunits (105). This idea was reinforced by studies showing that backward
mechanical rotation of the γ subunit with external force reverses the chemical reaction toward
ATP synthesis (42, 74), whereas forced forward rotation results in accelerated ATP binding (41).
This view was challenged by the ﬁnding that even when the γ subunit is shortened so that most γ-βinteraction sites are abolished, the short γ subunit still rotates unidirectionally (30, 62). However,
because single-molecule optical microscopy requires attachment of a probe to the γ subunit forthe observation of rotary catalysis, it cannot examine whether the rotary catalysis (hence, thecooperativity) occurs with α3 β3 alone. This issue was solved by HS-AFM imaging of α3 β3 in
ATP (99), as described below.
HS-AFM imaging of α3 β3 subcomplex. α3 β3 with Lys7 -tags at the N termini of the β subunits was covalently immobilized to a mica surface that was ﬁrst coated with 3-aminopropyltriethoxysilane and then treated with glutaraldehyde. An image of the C-terminal side of α3 β3without nucleotide shows a pseudo-sixfold symmetric ring (Figure 5b). Each subunit has an upwardly protruding portion at the inner top side of the ring, but these portions are higher at threealternately arranged subunits than at the other three. When this image was compared with a simulated AFM image (Figure 5d) constructed from a crystal structure of nucleotide-free α3 β3 (PDB
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

405

BB42CH17-Ando

ARI

3 April 2013

15:19

a

γ

β

z-scaleAnnu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

b

9.3 nm

5 nm

c

αE

αE

βE

βE

5 nm

0 nm

d

αE

0 nm

8.5 nmββEz-scale

e

α

f

βE

g

αE

αTP

βDP

βTP
αDP

h
0s

β2

0.56 sβ11.44 s1.68 s2.24 s2.32 sβ3

Figure 5
Atomic force microscopy (AFM) images of the α3 β3 subcomplex at the C-terminal surface. (a) Schematic for F1 -ATPase. (b) Averaged
AFM image obtained in nucleotide-free condition. (c) The C-terminal surface of crystal structure of a nucleotide-free α3 β3subcomplex. (d ) Simulated AFM image of panel c. (e) Averaged AFM image obtained in 1-mM AMP-PNP. ( f ) The C-terminal surfaceof crystal structure of an α3 β3 subcomplex obtained in ATP. ( g) Simulated AFM image of panel f. (h) Successive AFM images showing
counterclockwise rotary propagation of conformational change of an α3 β3 subcomplex at the C-terminal surface captured byhigh-speed AFM in the presence of 2 μM ATP, at 12.5 fps. The red circle marks the highest pixel position in each image. The color
scale placed at the right-hand side of panels b and e indicate the z-scale for the respective images. Adapted with permission from
Reference 99.

406

Ando

·

Uchihashi·

KoderaAnnu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

ARI

3 April 2013

15:19

ID: 1SKY) (88) (Figure 5c), the three subunits showing higher protrusions were identiﬁed as βsubunits.
In 1 mM AMP-PNP, the shape of α3 β3 became triangular and the central hole became obscure(Figure 5e). Only one subunit had a higher protrusion and an outwardly extended distal portion.
This is very likely to be a β subunit. If so, two subunits whose distal parts are retracted toward
the center are β subunits, in consideration of the alternate subunit arrangement. This assignment
was conﬁrmed by the comparison of the AFM image and a simulated AFM image (Figure 5g)
constructed using a crystal structure of a nucleotide-bound α3 β3 γ subcomplex (PDB ID: 1BMF)(1) from which the γ subunit was removed (Figure 5f ). In addition, it was clariﬁed that the βsubunit with the highest protrusion and an extended distal portion is empty and that the two β
subunits with retracted distal portions are nucleotide bound.
When imaged in 2–4 μM ATP at 12.5 fps, distinct conformational dynamics appeared at theβ subunits (Figure 5h; Supplemental Movie 5). Each β subunit exhibited a conformationaltransition between the outwardly extended high state (open, O, state) and the retracted low state(closed, C, state). The following prominent features were observed: (a) Only one β subunit assumes
the O state as in the presence of AMP-PNP, and (b) when the O-to-C transition occurs at one β
subunit, the opposite C-to-O transition occurs simultaneously at its counterclockwise neighbor
β subunit in most cases. Thus, the O conformation propagates counterclockwise (Figure 5h, redcircles). The ATP hydrolysis rates at 2, 3, and 4 μM ATP were in approximate agreement withthe rates of conformational revolution at 2, 3, and 4 μM ATP, respectively.
These dynamic transitions indicate that the O-to-C transition occurs when ATP is bound toan empty β subunit and that the C-to-O transition occurs when an ADP-bound β subunit releases
ADP. So, the empty, ADP-bound, and ATP-bound β subunits are arranged counterclockwise inthis order, and therefore, the observed conformational propagation demonstrates rotary catalysis
by the α3 β3 subcomplex. Thus, we reach the most important conclusion: The intrinsic cooperativity responsible for torque generation to rotate the γ subunit is elicited through β-β interplay
alone and the γ subunit passively undergoes torque to rotate (99).

FUTURE CHALLENGES
As exempliﬁed by these imaging studies, HS-AFM can provide high-resolution movies of biologicalmolecules in action from which we can learn a great detail about how the molecules operate tofunction. However, current HS-AFM has the following limitations: (a) The scan range of thehigh-speed scanner is limited to 1 μm, 4 μm, and 1 μm in the x-, y-, and z-directions, respectively,to achieve high resonant frequencies for fast scan, and (b) because the cantilever tip makes contact
with the sample, very soft surfaces such as the membranes of live eukaryotic cells are largely
deformed by the contact, which disables the visualization of molecules on such soft surfaces.
Below, we describe ongoing and future endeavors to overcome these limitations.

Wide-Area Observation and In Situ Imaging of Dynamic Processes
The size of a wider-area scanner becomes larger, resulting in lower resonant frequencies and hence
a lower highest-possible scan speed. This adverse effect has been recently removed by a simple
inversion-based feedforward control technique (7) and by an enhanced iterative inverse control
technique (54, 114). Therefore, even with a scanner capable of scanning over ∼40 × 40 μm2 , aline scan of ∼100 Hz is now possible without production of vibrations. However, because these
inverse control techniques capable of extending the bandwidth cannot be applied to the z-scanner,
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

407

BB42CH17-Ando

ARI

3 April 2013

15:19

its bandwidth is still limited. Nonetheless, a large sample such as a live eukaryotic cell can now be
imaged within ∼60 s when a relatively small number of scan lines (∼200) are used.
After taking a whole topography image of a large biological sample, we can image molecules
in a local area of interest on the sample surface when the surface is relatively rigid. For example,
it is possible to observe dynamics of molecules on the surfaces of live bacterial cells, intracellularorganelles (such as mitochondria and nuclei), and small structures such as neuronal spines. In fact,
in situ visualization of porin trimers moving on the outer surface of a live magnetic bacterium wasrecently accomplished (112). Such in situ dynamic molecular imaging will have a great impact oncell biology because dynamic molecular processes occurring on these surfaces are largely unknown.

Faster Wide-Area Observation and Dynamic Imaging of Cell Morphology
Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

For the imaging of a whole cell, many-pixel images with single-nanometer resolution are unnecessary. Nonetheless, in the tapping mode, the feedback operation has to be continuously performed
for a huge number of points on the surface; thus, it takes a long time to image a whole cell. Thissituation can be improved by the use of a different operation mode employed by scanning ion conductance microscopy (SICM). This mode, called AC mode (71), pulse mode (36, 55), or hopping
mode (69), has been devised to avoid probe-sample collision; a probe (glass capillary) is moved
up and down during lateral scanning of the sample stage. Therefore, the feedback operation is
discretely carried out only at lateral positions where the sample height information is acquired.
Its imaging acquisition time is given by the number of pixels contained in an image divided by
the up-and-down frequency fud . With fud = 20 kHz, a 50 × 50 μm2 image with 250 × 250 pixels
can be acquired within approximately 3 s. The resonant frequency of commercially available stack
piezoactuators displaceable up to 10 μm is approximately 30 kHz when their ends on one side are
ﬁxed. Therefore, this image acquisition time is realistic as far as the sample stage can be quickly
retracted upon tip-sample contact.

High-Speed Noncontact Imaging
A noncontact condition is mandatory for visualizing molecules on the extremely soft surfaces of
eukaryotic cells. SICM has already achieved this condition (16, 35). With the use of very sharp
glass capillaries with a small pore at the apex, the spatial resolution of SICM has reached a few
nanometers (117). Immobile protein molecules with a size of ∼14 nm in live cell membranes havebeen successfully imaged (84). Moreover, not only imaging but also smart electrophysiologicalmeasurements are possible with SICM (59). However, the bandwidth of ion conductivity detection
is low, because of the high impedance to ionic current through the small pore of a capillary electrodeand because of slow ionic currents. But, there is a possibility of increasing the bandwidth. Scanningtunneling microscopy also has a high impedance (hence a low bandwidth) problem but it has alreadybeen overcome by a circuitry technique (45). The slow ionic current problem can be solved by
the positioning of the counter-electrode close to the capillary end.
Noncontact AFM called solution-vibration AFM is currently being developed by the Andogroup. The sample stage is vibrated at a high frequency (1–3 MHz) with small amplitude, which
in turn vibrates the sample solution placed on the top of the sample stage. Although the vibrations
are transmitted to a place far from the sample surface, only solution vibrations at places in close
proximity to the sample surface are affected by the presence of the sample. In fact, when the tip
end was close to the surface of a polystyrene bead with a diameter of 150 nm (at a distance of<50 nm), the bead was visualized clearly by the phase detection of the cantilever vibrations (H.
Watanabe, T. Saito & T. Ando, unpublished result).
408

Ando

·

Uchihashi·

KoderaBB42CH17-Ando

ARI

3 April 2013

15:19

SUMMARY POINTS
1. Using HS-AFM, we can directly observe the structure dynamics and dynamic processes of
biomolecules, at subsecond to sub-100-ms temporal and submolecular spatial resolution,
without disturbing their function.
2. Visualized dynamic images of biomolecules can provide information inaccessible withother approaches, giving great insight into how the molecules function.
3. Dynamics of molecules that appear in AFM movies can be interpreted straightforwardly
without intricate analyses and interpretations, making it possible to attain ﬁrm conclusions.

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

4. In situ dynamic imaging of biomolecules is now becoming possible.
5. HS-AFM will transform structural biology and single-molecule biophysics.

DISCLOSURE STATEMENT
The authors are not aware of any afﬁliations, memberships, funding, or ﬁnancial holdings thatmight be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS
This work was supported by Japan Science and Technology Agency (Project of Core Research for
Evolutional Science and Technology), Grant-in-Aid for Basic Research (S) from Japan Society for
the Promotion of Science (20221006 and 24227005), and Ministry of Education, Culture, Sports,
Science and Technology Japan (Knowledge Cluster Initiative Project).
LITERATURE CITED
1. Abrahams JP, Leslie AG, Lutter R, Walker JE. 1994. Structure at 2.8 Å resolution of F1 -ATPase frombovine heart mitochondria. Nature 370:621–28
2. Albrecht TR, Grütter P, Horne D, Rugar D. 1991. Frequency modulation detection using high-Q
cantilevers for enhanced force microscope sensitivity. J. Appl. Phys. 69:668–73
3. Ando T. 2012. High-speed atomic force microscopy coming of age. Nanotechnology 23:062001
4. Ando T, Kodera N, Maruyama D, Takai E, Saito K, et al. 2002. A high-speed atomic force microscopefor studying biological macromolecules in action. Jpn. J. Appl. Phys. 41:4851–56
5. Ando T, Kodera N, Takai E, Maruyama D, Saito K, et al. 2001. A high-speed atomic force
microscope for studying biological macromolecules. Proc. Natl. Acad. Sci. USA 98:12468–72
6. Ando T, Uchihashi T. 2012. High-speed AFM and imaging of biomolecular processes. In NanoscaleLiquid Interfaces: Wetting, Patterning and Force Microscopy at Molecular Scale, ed. T Ondarçuhu, JP Aimé,pp. 711–40. Singapore: Pan Stanford Publ.
7. Ando T, Uchihashi T, Fukuma T. 2008. High-speed atomic force microscopy for nanovisualization of dynamic biomolecular processes. Prog. Surf. Sci. 83:337–437
8. Ando T, Uchihashi T, Kodera N, Miyagi A, Nakakita R, et al. 2005. High-speed AFM for studying the
dynamic behavior of protein molecules at work. e-J. Surf. Sci. Nanotechnol. 3:384–92
9. Ando T, Uchihashi T, Kodera N, Miyagi A, Nakakita R, et al. 2006. High-speed atomic force microscopy
for studying the dynamic behavior of protein molecules at work. Jpn. J. Appl. Phys. 45:1897–903
10. Binnig G, Quate CF, Gerber C. 1986. Atomic force microscope. Phys. Rev. Lett. 56:930–33
11. Blankenburg R, Meller P, Ringsdorf H, Salesse C. 1989. Interaction between biotin lipids and streptavidin in monolayers: formation of oriented two-dimensional protein domains induced by surface recognition. Biochemistry 28:214–21
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

3. Reviews various
bioimaging studies byhigh-speed AFM.

5. Presents firsthigh-speed AFM,
capturing moving
protein molecules at
12.5 fps.

7. Extensively reviews
various techniquesleading to theestablishment ofhigh-speed AFM forbiological studies.

409

ARI

3 April 2013

15:19

12. Casuso I, Khao J, Chami M, Paul-Gilloteaux P, Husain M, et al. 2012. Characterization of the motionof membrane proteins using high-speed atomic force microscopy. Nat. Nanotechnol. 7:525–29
13. Casuso I, Kodera N, Le Grimellec C, Ando T, Scheuring S. 2009. High-resolution high-speed contact
mode atomic force microscopy movies of purple membrane. Biophys. J. 97:1354–61
14. Casuso I, Rico F, Scheuring S. 2011. High-speed atomic force microscopy: structure and dynamics of
single proteins. Curr. Opin. Chem. Biol. 15:704–9
15. Casuso I, Sens P, Rico F, Scheuring S. 2010. Experimental evidence for membrane-mediated proteinprotein interaction. Biophys. J. 99:L47–49
16. Chen C-C, Zhou Y, Baker LA. 2012. Scanning ion conductance microscopy. Annu. Rev. Anal. Chem.
5:207–28
17. Costa LT, Pinto JR, Moraes MB, de Souza GGB, Sorenson MM, et al. 2004. Chemical treatment of
mica for atomic force microscopy can affect biological sample conformation. Biophys. Chem. 109:63–71
18. Czajkowsky DM, Shao Z. 2003. Inhibition of protein adsorption to muscovite mica by monovalentcations. J. Microsc. 211:1–7
19. Dague E, Alsteens D, Latgé J-P, Verbelen C, Raze D, et al. 2007. Chemical force microscopy of single
live cells. Nano Lett. 7:3026–30
20. Darst SA, Ahlers M, Meller PH, Kubalek EW, Blankenburg R, et al. 1991. Two-dimensional crystals of
streptavidin on biotinylated lipid layers and their interactions with biotinylated macromolecules. Biophys.
J. 59:387–96
21. De La Cruz EM, Wells AL, Rosenfeld SS, Ostap EM, Sweeney HL. 1999. The kinetic mechanism of
myosin V. Proc. Natl. Acad. Sci. USA 96:13726–31
22. Endo M, Hidaka K, Sugiyama H. 2011. Direct AFM observation of an opening event of a DNA cuboid
constructed via a prism structure. Org. Biomol. Chem. 9:2075–77
23. Endo M, Katsuda Y, Hidaka K, Sugiyama H. 2010. Regulation of DNA methylation using different
tensions of double strands constructed in a deﬁned DNA nanostructure. J. Am. Chem. Soc. 132:1592–97
24. Fantner GE, Hegarty P, Kindt JK, Schitter G, Cidade GAG, et al. 2005. Data acquisition system forhigh speed atomic force microscopy. Rev. Sci. Instrum. 76:026118
25. Fantner GE, Schitter G, Kindt JH, Ivanov T, Ivanova K, et al. 2006. Components for high speed atomic
force microscopy. Ultramicroscopy 106:881–87
26. Fernandez JM, Li H. 2004. Force-clamp spectroscopy monitors the folding trajectory of a single protein.
Science 303:1674–78
27. Forgacs E, Cartwright S, Sakamoto T, Sellers JR, Corrie JE, et al. 2008. Kinetics of ADP dissociationfrom the trail and lead heads of actomyosin V following the power stroke. J. Biol. Chem. 283:766–73
28. Forkey JN, Quinlan ME, Shaw MA, Corrier JET, Goldman YE. 2003. Three-dimensional structuraldynamics of myosin V by single-molecule ﬂuorescence polarization. Nature 422:399–404
29. Fukuma T, Okazaki Y, Kodera N, Uchihashi T, Ando T. 2008. High resonance frequency force microscope scanner using inertia balance support. Appl. Phys. Lett. 92:243119
30. Furuike S, Hossain MD, Maki Y, Adachi K, Suzuki T, et al. 2008. Axle-less F1 -ATPase rotates in thecorrect direction. Science 319:955–58
31. Gan Y. 2009. Atomic and subnanometer resolution in ambient conditions by atomic force microscopy.
Surf. Sci. Rep. 64:99–121
32. Gilmore JL, Suzuki Y, Tamulaitis G, Siksnys V, Takeyasu K, et al. 2009. Single-molecule dynamics
of the DNA−EcoRII protein complexes revealed with high-speed atomic force microscopy. Biochemistry48:10492–98
33. Giocondi M-C, Yamamoto D, Lesniewska E, Milhiet P-E, Ando T, et al. 2010. Surface topography ofmembrane domains. Biochim. Biophys. Acta 1798:703–18
34. Gresser MJ, Myers JA, Boyer PD. 1982. Catalytic site cooperativity of beef heart mitochondrial F1
adenosine triphosphatase. Correlations of initial velocity, bound intermediate, and oxygen exchange
measurements with an alternating three-site model. J. Biol. Chem. 257:12030–38
35. Hansma PK, Drake B, Marti O, Gould SA, Prater CB. 1989. The scanning ion-conductance microscope.
Science 243:641–43
36. Happel P, Hoffmann G, Mann SA, Dietzel ID. 2003. Monitoring cell movements and volume changes
with pulse-mode scanning ion conductance microscopy. J. Microsc. 212:144–51

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

410

Ando

·

Uchihashi·

KoderaAnnu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

ARI

3 April 2013

15:19

37. Hinterdorfer P, Dufrêne YF. 2006. Detection and localization of single molecular recognition eventsusing atomic force microscopy. Nat. Methods 3:347–55
38. Huxley HE. 1969. The mechanism of muscular contraction. Science 164:1356–66
39. Igarashi K, Koivula A, Wada M, Kimura S, Penttilä M, Samejima M. 2009. High speed atomic force microscopy visualizes processive movement of Trichoderma reesei cellobiohydrolase I on crystalline cellulose.
J. Biol. Chem. 284:36186–90
40. Igarashi K, Uchihashi T, Koivula A, Wada M, Kimura S, et al. 2011. Trafﬁc jams reduce hydrolytic
efﬁciency of cellulase on cellulose surface. Science 333:1279–82
41. Iko Y, Tabata KV, Sakakihara S, Nakashima T, Noji H. 2009. Acceleration of the ATP-binding rate of
F1 -ATPase by forcible forward rotation. FEBS Lett. 583:3187–91
42. Itoh H, Takahashi A, Adachi K, Noji H, Yasuda R, et al. 2004. Mechanically driven ATP synthesis by
F1 -ATPase. Nature 427:465–68
43. Jonkheijm P, Weinrich D, Schröder H, Niemeyer CM, Waldmann H. 2008. Chemical strategies forgenerating protein biochips. Angew. Chem. Int. Ed. 47:9618–47
44. Katan AJ, Dekker C. 2011. Leading edge, minireview: high-speed AFM reveals the dynamics of single
biomolecules at the nanometer scale. Cell 147:979–82
45. Kemiktarak U, Ndukum T, Schwab KC, Ekinci KL. 2007. Radio-frequency scanning tunneling microscopy. Nature 450:85–88
46. Kim J, Kim G, Cremer PS. 2001. Investigations of water structure at the solid/liquid interface in thepresence of supported lipid bilayers by vibrational sum frequency spectroscopy. Langmuir 17:7255–60
47. Kindt JH, Fantner GE, Cutroni JA, Hansma PK. 2004. Rigid design of fast scanning probe microscopes
using ﬁnite element analysis. Ultramicroscopy 100:259–65
48. Kinosita K Jr, Adachi K, Itoh H. 2004. Rotation of F1 -ATPase: how an ATP-driven molecular machine
may work. Annu. Rev. Biophys. Biomol. Struct. 33:245–68
49. Kitazawa M, Ohta R, Okita T, Tanaka J, Tanemura M. 2007. Mechanical properties of single carbon
nanoﬁbers grown on tips of scanning probe microscopy cantilevers by ion irradiation. Jpn. J. Appl. Phys.
46:6324–28
50. Kitazawa M, Shiotani K, Toda A. 2003. Batch fabrication of sharpened silicon nitride tips. Jpn. J. Appl.
Phys. 42:4844–47
51. Kodera N, Sakashita M, Ando T. 2006. Dynamic proportional-integral-differential controller
for high-speed atomic force microscopy. Rev. Sci. Instrum. 77:083704
52. Kodera N, Yamamoto D, Ishikawa R, Ando T. 2010. Video imaging of walking myosin V by
high-speed atomic force microscopy. Nature 468:72–76
53. Kodera N, Yamashita H, Ando T. 2005. Active damping of the scanner for high-speed atomic force
microscopy. Rev. Sci. Instrum. 76:053708
54. Li Y, Bechhoefer J. 2009. Model-free iterative control of repetitive dynamics for high-speed scanning inatomic force microscopy. Rev. Sci. Instrum. 80:013702
55. Mann SA, Hoffmann G, Hengstenberg A, Schuhmann W, Dietzel ID. 2002. Pulse-mode scanningion conductance microscopy—a method to investigate cultured hippocampal cells. J. Neurosci. Methods
116:113–17
56. Mehta AD, Rock RS, Rief M, Spudich JA, Mooseker MS, et al. 1999. Myosin-V is a processive actin-based
motor. Nature 400:590–93
57. Milhiet PE, Yamamoto D, Berthoumieu O, Dosset P, Le Grimellec C, et al. 2010. Deciphering thestructure, growth and assembly of amyloid-like ﬁbrils using high-speed atomic force microscopy. PLoS
ONE 5:e13240
58. Mingeot-Leclercq M-P, Deleu M, Brasseur R, Dufrêne YF. 2008. Atomic force microscopy of supported
lipid bilayers. Nat. Protoc. 3:1654–59
59. Miragoli M, Moshkov A, Novak P, Shevchuk A, Nikolaev VO, et al. 2011. Scanning ion conductance microscopy: a convergent high-resolution technology for multi-parametric analysis of living cardiovascular
cells. J. R. Soc. Interface 8:913–25
60. Miyagi A, Ando T, Lyubchenko YL. 2011. Dynamics of nucleosomes assessed with time-lapse high-speed
atomic force microscopy. Biochemistry 50:7901–8
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

51. Presents a new
feedback controller thatsolves the difficult
parachuting problem.

52. Presents astonishing
AFM movies of myosin
V walking on actin
filament.

411

ARI

3 April 2013

15:19

61. Miyagi A, Tsunaka Y, Uchihashi T, Mayanagi K, Hirose S, et al. 2008. Visualization of intrinsicallydisordered regions of proteins by high-speed atomic force microscopy. Chem. Phys. Chem. 9:1859–66
62. Mnatsakanyan N, Krishnakumar AM, Suzuki T, Weber J. 2009. The role of the βDELSEED-loop of
ATP synthase. J. Biol. Chem. 284:11336–45
63. Moreno-Herrero F, Colchero F, Gómez-Herrero J, Baró AM. 2004. Atomic force microscopy contact,
tapping, and jumping modes for imaging biological samples in liquids. Phys. Rev. E 69:031915
64. Müller DJ, Dufrêne YF. 2008. Atomic force microscopy as a multifunctional molecular toolbox in
nanobiotechnology. Nat. Nanotechnol. 3:261–69
65. Müller DJ, Helenius J, Alsteens D, Dufrêne YF. 2009. Force probing surfaces of living cells to molecularresolution. Nat. Chem. Biol. 5:383–90
66. Müller DJ, Janovjak H, Lehto T, Kuerschner L, Anderson K. 2002. Observing structure, function andassembly of single proteins by AFM. Prog. Biophys. Mol. Biol. 79:1–43
67. Nakajima H, Kunioka Y, Nakano K, Shimizu K, Seto M, et al. 1997. Scanning force microscopy of
the interaction events between a single molecule of heavy meromyosin and actin. Biochem. Biophys. Res.
Commun. 234:178–82
68. Noji H, Yasuda R, Yoshida M, Kinosita K Jr. 1997. Direct observation of the rotation of F1 -ATPase.
Nature 386:299–302
69. Novak P, Li C, Shevchuk AI, Stepanyan R, Caldwell M, et al. 2009. Nanoscale live-cell imaging using
hopping probe ion conductance microscopy. Nat. Methods 6:279–81
70. Oguchi Y, Mikhailenko SV, Ohki T, Olivares AO, De La Cruz EM, Ishiwata S. 2008. Load-dependent
ADP binding to myosins V and VI: implications for subunit coordination and function. Proc. Natl. Acad.
Sci. USA 105:7714–19
71. Pastré D, Iwamoto H, Liu J, Szabo G, Shao Z. 2001. Characterization of AC mode scanning ionconductance microscopy. Ultramicroscopy 90:13–19
72. Pastré D, Piétrement O, Fusil S, Landousy F, Jeusset J, et al. 2003. Adsorption of DNA to mica mediatedby divalent counterions: a theoretical and experimental study. Biophys. J. 85:2507–18
73. Purcell TJ, Sweeney HL, Spudich JA. 2005. A force-dependent state controls the coordination of processive myosin V. Proc. Natl. Acad. Sci. USA 102:13873–78
74. Rondelez Y, Tresset G, Nakashima T, Kato-Yamada Y, Fujita H, et al. 2005. Highly coupled ATP
synthesis by F1 -ATPase single molecules. Nature 433:773–77
75. Rosenfeld SS, Sweeney HL. 2004. A model of myosin V processivity. J. Biol. Chem. 279:40100–11
76. Sackmann E. 1996. Supported membranes: scientiﬁc and practical applications. Science 271:43–48
77. Sakamoto T, Amitani I, Yokota E, Ando T. 2000. Direct observation of processive movement by individual myosin V molecules. Biochem. Biophys. Res. Commun. 272:586–90
78. Sakamoto T, Webb MR, Forgacs E, White HD, Sellers JR. 2008. Direct observation of themechanochemical coupling in myosin Va during processive movement. Nature 455:128–32
79. Sanchez H, Suzuki Y, Yokokawa M, Takeyasu K, Wyman C. 2011. Protein-DNA interactions in highspeed AFM: single molecule diffusion analysis of human RAD54. Integr. Biol. 3:1127–34
80. Sannohe Y, Endo M, Katsuda Y, Hidaka K, Sugiyama H. 2010. Visualization of dynamic conformationalswitching of the G-quadruplex in a DNA nanostructure. J. Am. Chem. Soc. 132:16311–13
81. Scheuring S, Fotiadis D, Möller C, Müller SA, Engel A, et al. 2001. Single proteins observed by atomic
force microscopy. Single Mol. 2:59–67
82. Scheuring S, Sturgis JN. 2005. Chromatic adaptation of photosynthetic membranes. Science 309:484–87
83. Sellers JR, Weisman LS. 2008. Myosin V. In Myosins: A Superfamily of Molecular Motors, Vol. 7: Proteins
and Cell Regulation, ed. LM Coluccio, pp. 289–323. Berlin: Springer. 482 pp.
84. Shevchuk AI, Frolenkov GI, Sanchez D, James PS, Freedman N, et al. 2006. Imaging proteins in membranes of living cells by high-resolution scanning ion conductance microscopy. Angew. Chem. Int. Ed.
45:2212–16
85. Shibata M, Uchihashi T, Yamashita H, Kandori H, Ando T. 2011. Structural changes in bacteriorhodopsin in response to alternate illumination observed by high-speed atomic force microscopy. Angew.
Chem. Int. Ed. 50:4410–13

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

412

Ando

·

Uchihashi·

KoderaAnnu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

ARI

3 April 2013

15:19

86. Shibata M, Yamashita H, Uchihashi T, Kandori H, Ando T. 2010. High-speed atomic force
microscopy shows dynamic molecular processes in photo-activated bacteriorhodopsin. Nat.
Nanotechnol. 5:208–12
87. Shinozaki Y, Sumitomo K, Tsuda M, Koizumi S, Inoue K, et al. 2009. Direct observation of ATP-induced
conformational changes in single P2X4 receptors. PLoS Biol. 7:e1000103
88. Shirakihara Y, Leslie AG, Abrahams JP, Walker JE, Ueda T, et al. 1997. The crystal structure of thenucleotide-free α3β3 subcomplex of F1 -ATPase from the thermophilic Bacillus PS3 is a symmetrictrimer. Structure 5:825–36
89. Shiroguchi K, Kinosita K Jr. 2007. Myosin V walks by lever action and Brownian motion. Science
316:1208–12
90. Stewart MP, Helenius J, Toyoda Y, Ramanathan SP, Müller DJ, et al. 2011. Hydrostatic pressure andthe actomyosin cortex drive mitotic cell rounding. Nature 469:226–30
91. Stroh C, Wang H, Bash R, Ashcroft B, Nelson J, et al. 2004. Single-molecule recognition imaging
microscopy. Proc. Natl. Acad. Sci. USA 101:12503–7
92. Sulchek T, Yaralioglu GG, Quate CF, Minne SC. 2002. Characterization and optimization of scan speed
for tapping-mode atomic force microscopy. Rev. Sci. Instrum.73:2928–36
93. Suzuki Y, Gilmore JL, Yoshimura SH, Henderson RM, Lyubchenko YL, et al. 2011. Visual analysis ofconcerted cleavage by type IIF restriction enzyme SﬁI in subsecond time region. Biophys. J. 101:2992–98
94. Suzuki Y, Higuchi Y, Hizume K, Yokokawa M, Yoshimura SH, et al. 2010. Molecular dynamics of
DNA and nucleosomes in solution studied by fast-scanning atomic force microscopy. Ultramicroscopy
110:682–88
95. Syed S, Snyder GE, Franzini-Armstrong C, Selvin PR, Goldman YE. 2006. Adaptability of myosin V
studied by simultaneous detection of position and orientation. EMBO J. 25:1795–803
96. Tanaka F, Mochizuki T, Liang X, Asanuma H, Tanaka S, et al. 2010. Robust and photocontrollable
DNA capsules using azobenzenes. Nano Lett. 10:3560–65
97. Tanemura M, Okita T, Yamauchi H, Tanemura S, Morishima R. 2004. Room-temperature growth ofa carbon nanoﬁber on the tip of conical carbon protrusions. Appl. Phys. Lett. 84:3831–33
98. Touhami A, Nysten B, Dufrêne YF. 2003. Nanoscale mapping of the elasticity of microbial cells byatomic force microscopy. Langmuir 19:4539–43
99. Uchihashi T, Iino R, Ando T, Noji H. 2011. High-speed atomic force microscopy reveals rotary
catalysis of rotorless F1 -ATPase. Science 333:755–58
100. Uchihashi T, Kodera N, Ando T. 2012. Guide to video recording of structure dynamics and dynamic
processes of proteins by high-speed atomic force microscopy. Nat. Protoc. 7:1193–206
101. Uchihashi T, Yamashita H, Ando T. 2006. Fast phase imaging in liquids using a rapid scan atomic force
microscope. Appl. Phys. Lett. 89:213112
102. Veigel C, Schmitz S, Wang F, Sellers JR. 2005. Load-dependent kinetics of myosin-V can explain its
high processivity. Nat. Cell. Biol. 7:861–69
103. Viani MB, Pietrasanta LI, Thompson JB, Chand A, Gebeshuber IC, et al. 2000. Probing protein–protein
interactions in real time. Nat. Struct. Biol. 7:644–47
104. Viani MB, Schäffer TE, Paloczi GT, Pietrasanta LI, Smith BL, et al. 1999. Fast imaging and fast force
spectroscopy of single biopolymers with a new atomic force microscope designed for small cantilevers.
Rev. Sci. Instrum. 70:4300
105. Wang H, Oster G. 1998. Energy transduction in the F1 motor of ATP synthase. Nature 396:279–82
106. Warshaw DM, Kennedy GG, Work SS, Krementsova EB, Beck S, et al. 2005. Differential labeling of
myosin V heads with quantum dots allows direct visualization of hand-over-hand processivity. Biophys.
J. 88:L30–32
107. Wickham FJS, Endo M, Katsuda Y, Hidaka K, Bath J, et al. 2011. Direct observation of stepwise
movement of a synthetic molecular transporter. Nat. Nanotechnol. 6:166–69
108. Williams M, Fowler SB, Robert B, Best RB, Toca-Herrera JL, et al. 2003. Hidden complexity in themechanical properties of titin. Nature 422:446–49
109. Yamamoto D, Nagura N, Omote S, Taniguchi M, Ando T. 2009. Streptavidin 2D crystal substrates forvisualizing biomolecular processes by atomic force microscopy. Biophys. J. 97:2358–67
www.annualreviews.org • High-Speed AFM for Filming Biomolecules

86. Presents AFM
movies showing bR in
response to light andcooperativity by bR-bR
interactions.

99. Presents AFM
movies showing rotary
propagation ofconformational changesof rotorless F1 -ATPase.

413

ARI

3 April 2013

15:19

110. Yamamoto D, Uchihashi T, Kodera N, Ando T. 2008. Anisotropic diffusion of point defects in a twodimensional crystal of streptavidin observed by high-speed atomic force microscopy. Nanotechnology
19:384009
111. Yamamoto D, Uchihashi T, Kodera N, Yamashita H, Nishikori S, et al. 2010. High-speed atomic force
microscopy techniques for observing dynamic biomolecular processes. Methods Enzymol. 457:B541–44
112. Yamashita H, Taoka A, Uchihashi T, Asano T, Ando T, et al. 2012. Single molecule imaging on living
bacterial cell surface by high-speed AFM. J. Mol. Biol. 422:300–9113. Yamashita H, Voı̈tchovsky K, Uchihashi T, Antoranz Contera S, Ryan JF, et al. 2009. Dynamics ofbacteriorhodopsin 2D crystal observed by high-speed atomic force microscopy. J. Struct. Biol. 167:153–
58
114. Yan Y, Wu Y, Zou Q, Su C. 2008. An integrated approach to piezoactuator positioning in high-speed
atomic force microscope imaging. Rev. Sci. Instrum. 79:073704
115. Yasuda R, Noji H, Kinosita K Jr, Yoshida M. 1998. F1 -ATPase is a highly efﬁcient molecular motor thatrotates with discrete 120◦ steps. Cell 93:1117–24
116. Yildiz A, Forkey JN, McKinney SA, Ha T, Goldman YE, et al. 2003. Myosin V walks hand-over-hand:single ﬂuorophore imaging with 1.5-nm localization. Science 300:2061–65
117. Ying L, Bruckbauer A, Zhou D, Gorelik J, Shevchuk A, et al. 2005. The scanned nanopipette: a newtool for high resolution bioimaging and controlled deposition of biomolecules. Phys. Chem. Chem. Phys.
7:2859–66
118. Yokokawa M, Takeyasu K. 2011. Motion of the Ca2+ -pump captured. FEBS J. 278:3025–31
119. Yokokawa M, Wada C, Ando T, Sakai N, Yagi A, et al. 2006. Fast-scanning atomic force microscopy
reveals the ATP/ADP-dependent conformational changes of GroEL. EMBO J. 25:4567–76
120. Zhong Q, Inniss D, Kjoller K, Elings VB. 1993. Fractured polymer/silica ﬁber surface studied by tapping
mode atomic force microscopy. Surf. Sci. 290:L688–92

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

BB42CH17-Ando

414

Ando

·

Uchihashi·

KoderaBB42-Frontmatter

ARI

13 April 2013

7:12

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Contents

Annual Review ofBiophysics
Volume 42, 2013

Doing Molecular Biophysics: Finding, Naming, and Picturing Signal
Within Complexity
Jane S. Richardson and David C. Richardson p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 1
Structural Biology of the Proteasome
Erik Kish-Trier and Christopher P. Hill p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p29
Common Folds and Transport Mechanisms of Secondary
Active Transporters
Yigong Shi p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p51
Coarse-Graining Methods for Computational Biology
Marissa G. Saunders and Gregory A. Voth p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p73
Electrophysiological Characterization of Membrane
Transport Proteins
Christof Grewer, Armanda Gameiro, Thomas Mager, and Klaus Fendler p p p p p p p p p p p p p p p95
Entropy-Enthalpy Compensation: Role and Ramiﬁcations in
Biomolecular Ligand Recognition and Design
John D. Chodera and David L. Mobley p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 121
Molecular Mechanisms of Drug Action: An Emerging View
James M. Sonner and Robert S. Cantor p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 143
The Underappreciated Role of Allostery in the Cellular Network
Ruth Nussinov, Chung-Jung Tsai, and Buyong Ma p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 169
Structural Insights into the Evolution of the Adaptive Immune System
Lu Deng, Ming Luo, Alejandro Velikovsky, and Roy A. Mariuzza p p p p p p p p p p p p p p p p p p p p p 191
Molecular Mechanisms of RNA Interference
Ross C. Wilson and Jennifer A. Doudna p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 217
Molecular Trafﬁc Jams on DNA
Ilya J. Finkelstein and Eric C. Greene p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 241

vii

BB42-Frontmatter

ARI

13 April 2013

7:12

Advances, Interactions, and Future Developments in the CNS, Phenix,
and Rosetta Structural Biology Software Systems
Paul D. Adams, David Baker, Axel T. Brunger, Rhiju Das, Frank DiMaio,
Randy J. Read, David C. Richardson, Jane S. Richardson,
and Thomas C. Terwilliger p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 265
Considering Protonation as a Posttranslational Modiﬁcation
Regulating Protein Structure and Function
André Schönichen, Bradley A. Webb, Matthew P. Jacobson, and Diane L. Barber p p p p p 289

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Energy Functions in De Novo Protein Design: Current Challenges
and Future Prospects
Zhixiu Li, Yuedong Yang, Jian Zhan, Liang Dai, and Yaoqi Zhou p p p p p p p p p p p p p p p p p p p p p 315
Quantitative Modeling of Bacterial Chemotaxis: Signal Ampliﬁcation
and Accurate Adaptation
Yuhai Tu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 337
Inﬂuences of Membrane Mimetic Environments on Membrane
Protein Structures
Huan-Xiang Zhou and Timothy A. Cross p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 361
High-Speed AFM and Applications to Biomolecular Systems
Toshio Ando, Takayuki Uchihashi, and Noriyuki Kodera p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 393
Super-Resolution in Solution X-Ray Scattering and Its Applications to
Structural Systems Biology
Robert P. Rambo and John A. Tainer p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 415
Molecular Basis of NF-κB Signaling
Johanna Napetschnig and Hao Wu p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 443
Regulation of Noise in Gene Expression
Alvaro Sanchez, Sandeep Choubey, and Jane Kondev p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 469
Evolution in Microbes
Edo Kussell p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 493
Protein Structure Determination by Magic-Angle Spinning Solid-State
NMR, and Insights into the Formation, Structure, and Stability of
Amyloid Fibrils
Gemma Comellas and Chad M. Rienstra p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 515
Structural Studies of RNase P
Alfonso Mondragón p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 537
On the Universe of Protein Folds
Rachel Kolodny, Leonid Pereyaslavets, Abraham O. Samson, and Michael Levitt p p p p p p 559

viii

Contents

BB42-Frontmatter

ARI

13 April 2013

7:12

Torque Measurement at the Single-Molecule Level
Scott Forth, Maxim Y. Sheinin, James Inman, and Michelle D. Wang p p p p p p p p p p p p p p p p 583
Modeling Gene Expression in Time and Space
Pau Rué and Jordi Garcia-Ojalvo p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 605
Mechanics of Dynamin-Mediated Membrane Fission
Sandrine Morlot and Aurélien Roux p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 629

Annu. Rev. Biophys. 2013.42:393-414. Downloaded from www.annualreviews.org
by Kanazawa University Medical Library Branch on 05/13/13. For personal use only.

Nanoconﬁnement and the Strength of Biopolymers
Tristan Giesa and Markus J. Buehler p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p p 651
Solid-State NMR of Nanomachines Involved in Photosynthetic
Energy Conversion
A. Alia, Francesco Buda, Huub J.M. de Groot, and Jörg Matysik p p p p p p p p p p p p p p p p p p p p p p 675
Index
Cumulative Index of Contributing Authors, Volumes 38–42 p p p p p p p p p p p p p p p p p p p p p p p p p p p 701
ErrataAn online log of corrections to Annual Review of Biophysics articles may be found at
http://biophys.annualreviews.org/errata.shtml

Contentsix