Review
pubs.acs.org/CR

Terms of Use

Filming Biomolecular Processes by High-Speed Atomic ForceMicroscopy
Toshio Ando,*,†,‡,§ Takayuki Uchihashi,†,‡,§ and Simon Scheuring∥†

Department of Physics, and ‡Bio-AFM Frontier Research Center, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan§
CREST, Japan Science and Technology Agency, 4-1-8 Honcho, Kawaguchi 332-0012, Japan∥U1006 INSERM/Aix-Marseille Université, Parc Scientiﬁque et Technologique de Luminy Bâtiment Inserm TPR2 bloc 5, 163 avenue
de Luminy, 13288 Marseille Cedex 9, France
8.6. Diﬀusion and Interaction of Membrane
Proteins
8.6.1. Interaction between bR Trimers in 2D
Crystal Lattice
8.6.2. Interaction of C-Rings in Purple Membrane
8.6.3. Interaction of AQP0 in Native Eye Lens
Membrane
8.6.4. Diﬀusion and Interaction of OmpF
8.7. Self-Assembly Processes
8.7.1. Amyloid Fibril Formation
8.7.2. 2D Crystallization of Annexin A5
8.7.3. Supported Lipid Bilayer Formation
8.8. Other Topics on Protein Dynamics
8.8.1. Cellulase
8.8.2. Intrinsically Disordered Proteins
8.8.3. Dynamics of Vacancy Defects in 2D
Crystals of Proteins
9. Dynamic Imaging of Live Cells
9.1. Bacterial Cell Surfaces
9.1.1. Bacteriolysis of Bacillus subtilis Subjectedto Lysozyme
9.1.2. Surface Change of E. coli Subjected to
Antimicrobial Peptide
9.1.3. Imaging of Single Molecules on Live
Bacteria Surfaces
9.2. Dynamic Phenomena Occurring on Eukaryotic Cell Surfaces
9.3. Diﬀusion and Interactions of Membrane
Proteins on Eukaryotic Live Cells
10. Perspectives
11. Concluding Remarks
Appendix
A.1. Resolution
A.2. Software
A.3. Simulation of AFM Image
A.4. Correlation Coeﬃcient Analysis
A.5. Tracking Analysis of Proteins Moving along a
Filament
Author Information
Corresponding Author
Notes

CONTENTS
1. Introduction
2. Principle of AFM Imaging
3. Speed Limits of AFM Imaging
4. Minimizing Invasiveness: The Tip−Sample Interaction
5. Small Cantilevers
6. Hydrodynamic Pressure
7. Substrate Surfaces
7.1. Mica Surfaces
7.2. Lipid Bilayer Surfaces
7.3. Surface of 2D Crystals of Streptavidin
8. Dynamic Imaging of Proteins
8.1. General Remarks on Dynamic Structural
Study of Motor Proteins
8.2. Myosin V Walking on Actin Filament
8.2.1. Visualization of Lever-Arm Swing
8.2.2. Directional Rule8.2.3. Foot Stomp and Tension Generation
8.2.4. Asymmetric ADP Dissociation Kinetics
8.2.5. Chemo-mechanical Coupling in Walking
Myosin V
8.2.6. Remaining Issues in the Walking Mechanism of Myosin V
8.3. Rotary Catalysis of α3β3 Subcomplex of F1ATPase
8.4. General Remarks on Membrane Proteins
8.5. Structural Changes of Membrane Proteins
8.5.1. Bacteriorhodopsin Responding to Light
8.5.2. Role of Trimer−Trimer Interaction in bR
Function
8.5.3. Up−Down Motion of Ca2+-ATPase
8.5.4. P2X4 Receptor in Response to ATP© 2014 American Chemical Society

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Special Issue: 2014 Single Molecule Mechanisms and Imaging
Received: July 16, 2013
Published: January 30, 2014
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Biographies
Acknowledgments
Abbreviations
References

Review

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Table 1. Feasibility Comparison of Three Types ofMicroscopy

1. INTRODUCTION
In the history of science and technology, researchers have always
undertaken endeavors to enhance the degree of “directness ofmeasurement”. With “directness of measurement”, we conceptualize that the essence of the object under investigation, thestructure, dynamics, and function of biological samples in thescope of this Review, is entirely and straightforwardly assessed by
the measurement, bypassing hypotheses and intricate data
analysis. When the degree of measurement directness is low,
conclusions derived from the gleaned data often diﬀer dependingon the formulation of hypotheses, analysis models, and the datainterpretation. That is why conceptual consensus about a speciﬁc
issue of the studied object is rarely reached based on indirect
data. This is often the case for studies of structure−function
relationship of proteins. Previously, scientists had to understanddistinct qualities of proteins by measuring a sample containing ahuge number of molecules. The results are ensemble-averaged
quantities (often at equilibrium) that provide only limited
information on the proteins of study. Proteins are dynamic innature and work at the single-molecule level. Protein molecules
ﬂuctuate, undergo structural changes, bind to and dissociate frominteraction partners, and traverse a range of energy and chemical
states during molecular action. Most, if not all, of these dynamics
and their statistical distributions are convoluted and hidden in
ensemble averaging measurements.
To overcome the limitations of ensemble measurements,
single-molecule biophysics was developed more than two
decades ago, with the use of ﬂuorescence microscopy,1,2 optical
spectroscopy,3,4 and optical and magnetic tweezers,5,6 whose
performances were further improved by the advancements ofimproved optical microscopes, lasers, electronics, computers,
and high-sensitivity video cameras and sensors. Using thesetechniques, our understanding of the functional mechanism ofproteins has made signiﬁcant steps forward. Moreover, superresolution optical microscopy techniques bypassing thediﬀraction limit for ﬂuorophore localization have recently been
added to ﬂuorescence microscopy.7−9 However, the degree of
directness of measurement is still limited, because the protein
molecules themselves are invisible in these single-molecule
measurements. Protein structure is typically studied by X-ray
crystallography, electron microscopy (EM), and nuclear
magnetic resonance (NMR) spectroscopy. To date, thesetechniques have revealed detailed three-dimensional structuresof over 94 000 proteins (Protein Data Bank (PDB), http://www.
rcsb.org/pdb/home/home.do), with a growth rate of about 8000
novel structures per year (2010−2013). Yet, these techniques
make use of ensemble averaging, and, more seriously, theobtained structures are merely limited to static snapshots of ﬁxed
conformations. Thus, the simultaneous and direct observation of
structure, dynamics, and function of single protein molecules haslong been infeasible, and hence the materialization of a techniqueallowing such an observation has long been awaited in biologicalsciences.
An ideal microscopy technique that allows simultaneous
observation of structure, dynamics, and function of single protein
molecules has to meet all of the following conditions (see Table
1): (i) in-liquid specimen imaging, (ii) high spatial resolution,(iii) high temporal resolution, (iv) low invasiveness to thespecimen, and (v) direct imaging of the specimen without the useof markers (in other words, resolving the structure of thespecimen itself). Although eﬀorts have been made to develop
environmental electron microscopy techniques capable ofobserving unstained biological specimens in solutions,10 thestrong electron dose that is required to achieve high contrast andspatial resolution instantaneously denatures the sample.
Achieving the above-described goals by EM is a highly diﬃcult,if not impossible, task. Conventional atomic force microscopy11(AFM) meets most of the above-mentioned conditions, except
for the third condition, that is, high temporal resolution, and thefourth condition, that is, low invasiveness, is only moderately
satisﬁed.
AFM was originally invented to visualize atoms on solid
surfaces.11 The basic principle of AFM imaging is that a sharp tip
attached to the free end of a microcantilever probes the sample
surface, while this tip is being scanned over the sample surface(more details are given in the next section). Today, AFM can beused for a wide range of specimen made of any material in any
environment (vacuum, air, and liquids). When used formeasuring biological samples in liquid, the tip is brought into
contact with the sample, and the mechanical response of thecantilever upon this contact is measured. The sample stage (orthe cantilever base) is moved in the Z-direction to keep constantthe cantilever’s mechanical state (and hence, the mechanical tip−
sample interaction force) at a given condition, that is, a set point,
through feedback control. This series of operation is repeatedmany times, so that the tip traces the whole sample surface toprovide the sample surface topography. AFM is now routinely
used in biological sciences (see reviews12,13) not only for highresolution imaging of biological samples under physiologicalconditions,14,15 but also for force measurements to estimate thestrength of intra- and intermolecular bonds at the singlemolecule level16,17 and the elasticity of biological objects.18−20
However, as mentioned above, the AFM’s low imaging rate(about 1 min is required to capture an image) has precluded theanalysis of the vast majority of dynamic biological processes andtherefore limited its usefulness for biological studies.
This shortcoming of AFM imaging can in principle besigniﬁcantly improved. Encouraged by this expectation and3121

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Table 2. HS-AFM Imaging Studies on Proteins and Cells
year

targets and subjects

2006

actin filament gliding on myosin and GroEL−
GroES interaction24
conformational change of GroEL and GroEL−
GroES interaction30
DNA translocation and looping by type III
restriction enzyme31
identification of intrinsically disordered regions of
FACT protein32
DNA−BAL 31 nuclease interaction33
anisotropic diffusion of point defects in streptavidin
2D crystals34
dynamic equilibrium at the edge of bR 2D
crystals35
unidirectional translocation of cellulase along
cellulose fibers36
ATP-induced conformational change in P2X4
receptor37
structural change of CaM and actin polymerization
on streptavidin 2D crystals38
purple membrane in contact-mode HS-AFM42

2007
2008

2009

2010

translocation of EcoRII restriction enzyme along
DNA40
structural change of bR trimer in a stretchedmembrane41
membrane-mediated protein−protein interaction39
height change of acid-sensing ion channel 1a upon
acidification43
photoinduced structural change in bR44

2011

process of forming supported planar lipid bilayers45
effect of ClpX on FtsZ polymerization46
self-assembly of amyloid-like fibrils47
myosin V walking on an actin filament48
time course of cell death by antimicrobial peptide68
bR in response to alternate two-color
illumination49
height change in sarcoplasmic reticulum Ca2+ATPase50

substratesurfacesyearbare mica
bare mica
bare mica
bare mica
bare mica
streptavidin
2D crystals
nativemembrane
HOPG

2012

bare mica andSLB
streptavidin
2D crystals
nativemembranebare mica
nativemembraneb
nativemembraneSLB

2013

nativemembranebare mica
bare mica
bare mica
SLB
live cellanativemembranenativemembranetargets and subjects
structural fluctuations of CENP-W complexed with
CENP-T51
rotary catalysis of rotorless F1-ATPase52
pathways of ATP-independent unwrapping of
nucleosomes53
concerted DNA cleavage by Type IIF Restriction
Enzyme SfiI54
slide and hopping motion of Rad54 on DNA55
dynamics of synthetic proteo-nucleic building
blocks56
unidirectional movement and traffic jams ofcellulases hydrolyzing cellulose57
formation of synthetic DNA−protein complex
formation58
diffusion and interaction of OmpF molecules incrowded condition59
cooperative interaction dynamics of aquaporin-0 in
eye lens cell membranes60
diffusion of crowded porin molecules on live
bacterial cell69
formation of complexes between DNA and singlestranded DNA-binding protein65
dynamics of DNA cytosine deaminase bound tosingle-stranded DNA67
structural changes of NMDA receptors61
photoactivated structural changes in isolated
bacteriorhodopsin trimers62
phosphorylation-coupled intramolecular dynamics
of unstructured FACT protein63
dynamic processes in bacteriolysis and endocytosis
in live cells70
diffusion and interaction of AQP0 on live cells71
oligomerization of DNA cytosine deaminase on
single-stranded DNA66
self-assembly of sphingomyelin-specific toxin64

substratesurfacesbare mica
modified mica
modified mica
modified mica
bare mica
bare mica
HOPG
modified mica
SLB
nativemembrane
live cellamodified mica
modified mica
SLB
nativemembraneabare mica
live cellsa
live cellsa
modified mica
SLBaNative membranes and live cells are placed on chemically modiﬁed
mica surfaces. bNative membranes are suspended over nanowells. In
the other cases, native membranes are placed and SLBs are formed onbare mica surfaces. Streptavidin 2D crystals are formed on biotincontaining SLBs.

possible.68−71 Most of the biological HS-AFM studies onproteins and cells performed since 2006 are listed in Table 2.
This Review focuses on application studies of HS-AFM, inwhich dynamics of proteins and live cells are visualized. For
application studies of HS-AFM on the dynamics of DNA andDNA-binding proteins, see ref 72. Here, we also describe the HSAFM setup, substrate surfaces, and precautions to be consideredin HS-AFM imaging experiments.73 Before these topics aredescribed, the physical backgrounds of the high imaging rate andlow invasiveness are explained. In the last part of this Review,
future prospects of HS-AFM studies are brieﬂy outlined.

motivated by the desire to visualize single protein molecules inaction, attempts to increase the AFM imaging rate were initiatedaround 1993 by the Hansma and Ando groups independently,
and later by the Miles group. Through the development of newdevices and techniques (see reviews21−24) and the improvementof the ﬁrst version of prototypic instruments,25,26 HS-AFM wasrealized around 200823 (for the history of development of HS-AF
techniques, see review22). Now in 2013, state of the art HS-AFM
can capture dynamic images of biomolecules at sub-100 ms
temporal and submolecular lateral resolution. Importantly, thefunction of fragile molecules is not disturbed by the interaction
with the cantilever tip (detailed in section 4, Minimizing
Invasiveness: The Tip−Sample Interaction). This performance
made it possible to directly observe the structural dynamics anddynamic processes of biomolecules in action, providing astraightforward understanding of how the biomolecules operate
to function.27−29 In the past few years, various biomolecularsystems and their dynamic events have been studied using HSAFM, continuously demonstrating the innovative power of thisnovel microscopy to reveal details of molecular action
inaccessible to other approaches.30−67 Very recently, the analysis
of dynamic events occurring in live cells has also become

2. PRINCIPLE OF AFM IMAGING
AFM can be operated in several imaging modes.74,75 These are
generally classiﬁed into contact mode, intermittent contact
mode, noncontact mode, and other modes. Figure 1 shows atypical experimental setup for intermittent contact mode AFM.
For HS-AFM imaging of biological samples, the intermittent
contact mode is predominantly used, which will be detailed
below. However, ﬁrst, brief descriptions are given of the contact
and noncontact modes; other modes are out of the scope of thisReview.
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Figure 1. Schematic of experimental setup for intermittent contact mode AFM.

surface) of the cantilever, respectively. This is because theapparent spring constant of the cantilever is changed from kc to kc− k by the tip−sample interaction. In the attractive regime, k is
positive, and hence, Δfc < 0, whereas in the repulsive regime, k is
negative, and hence, Δfc > 0. Before imaging, a set point
frequency is deﬁned, which is shifted with respect to the original
resonance frequency. During scanning, the feedback control tries
to get the actual frequency back to the set point frequency bydriving the Z-scanner, which restores the desired tip−sample
interaction corresponding to the set point frequency. The true
noncontact condition is achieved only in vacuum or in air,because a high quality factor of the cantilever oscillation Qc is
required for highly sensitive detection of Δfc at any instant during
the scanning. In solution, Qc is small because of damping of thecantilever oscillation due to the viscous drag eﬀect of thesolution. Moreover, to obtain a relatively large Δfc, the oscillating
cantilever tip must always be close to the sample surface where
the force gradient is steep, meaning that the cantilever oscillation
amplitude must be kept very small (<0.2 nm). Nevertheless, FMmode AFM has recently been used also in aqueous solutions to
obtain high-resolution images of biological samples78−80 underconditions where the oscillating tip makes contact with thesample. Liquid FM-AFM is, however, limited to samples withsmall surface roughness (<1 nm). Because of this limitation, theFM-mode is generally inappropriate for biological HS-AFM thatis desired to be applicable to a wide range of samples.
In intermittent contact mode, also called tapping mode or
amplitude modulation (AM) mode,81 the cantilever is excited tooscillate at or near its ﬁrst resonance frequency. The excitation
frequency is ﬁxed at a constant value. The oscillating tip interacts
with the sample surface at its bottom swing (about 10% of theoscillation cycle), by which the oscillation amplitude is reduced.

In contact mode, the cantilever tip is always in contact with thesample while scanned across the sample surface during
topography acquisition. The cantilever is deﬂected by the tip−
sample interaction. The extent of cantilever deﬂection, and hence
the tip−sample interaction force, is kept constant during
scanning by feedback control. The force applied on the sample
can be minimized by the use of contact mode cantilevers withsmall spring constants with respect to bending by the normal
force. In contrast, the spring constant is high for cantileverbending and twisting caused by lateral friction forces. Therefore,
when the sample surface roughness is relatively large, high lateral
friction forces are exerted by the dragging tip onto the sample,
eventually resulting in signiﬁcant deformation and damage of
biological samples or displacement of samples that are weakly
attached to the substrate surface. Therefore, this mode has been
most successfully used for ﬁrmly immobilized samples with small
surface roughness such as bacteriorhodopsin (bR) embedded in
the crystalline purple membrane.76
In noncontact mode, the so-called frequency-modulation(FM) mode is typically used.77 In the FM mode, the cantilever isalways oscillated at its ﬁrst resonant frequency, using a selfoscillation circuit. The self-oscillation means that the cantileverdeﬂection signal is used to generate the cantilever excitation
signal, so that the frequency of the excitation signal is the same as
the cantilever resonance frequency that varies upon tip−sample
interaction. The gradient of the tip−sample interaction force inZ-direction, k, induces a shift (Δfc) of the cantilever resonancefrequency, approximately by
Δfc = −(fc /2kc)k(1)

where kc and fc are the spring constant and the original resonancefrequency (under free oscillation while the tip is very far from the3123

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This amplitude reduction is not only due to energy dissipation.
As will be described later, the cantilever resonance frequency
shift (and hence a phase shift relative to the excitation signal) also
results in amplitude reduction, depending on the excitation
frequency used. During scanning, the oscillation amplitude A is
kept constant at the amplitude set point As by feedback control.
The cantilever deﬂection is measured typically by the opticalbeam deﬂection (OBD) system (see Figure 1). The laser beam
reﬂected back by the cantilever is guided to the segmented (bicell
or quadrant) photodiode sensor. The output signals from thephotodiode sensor are conditioned to produce a diﬀerential
signal (S1 − S2) or a normalized diﬀerential signal (S1 − S2)/(S1 +S2) corresponding to the cantilever deﬂection. Either one ofthese two signals is fed into the deﬂection-to-amplitude
convertor, followed by subtraction of its output amplitude signal
from a set point voltage corresponding to the set point
amplitude. The resulting signal (i.e., feedback error) is fed into
the proportional-integral-derivative (PID) feedback controller.
The PID controller’s output signal is input to the Z-piezodriverfor signal ampliﬁcation to displace the Z-scanner so that the
feedback error approaches zero. The output from the PID
controller represents the sample height, because the Z-scanner
movement driven by the ampliﬁed PID output signal traces the
sample surface, although its movement is the inverse of thesample height. Therefore, the PID output signal is used toconstruct the sample topography in a computer. The signals fordisplacing the X- and Y-scanners, deﬁning the scan area underinvestigation, are generated by the computer through a twochannel digital/analog (D/A) converter and fed into therespective piezodrivers.

Figure 2. Eﬀects of feedback delay on the tip force exerted onto thesample. (a) Movement of Z-scanner (red line) tracing sinusoidally
shaped sample surface (black line) with periodicity λ and amplitude h0/2, when no parachuting occurs. Because of the delay of the Z-scanner
movement, tracing error (blue line) is produced. (b) Tip force (blueline) exerted onto the sample when the set point force (black line)
corresponding to the set point amplitude is relatively large. (c) Tip force(blue line) exerted onto the sample when the set point force (black line)
is set at a small value.

determined by A0, As, kc, and Qc, and approximately expressed
as82

3. SPEED LIMITS OF AFM IMAGING
The input to the feedback loop is the change in sample heightunder the cantilever tip, which is caused by lateral scanning of thesample stage relative to the cantilever. The ﬁnal output from thefeedback loop is the displacement of the Z-scanner. If it waspossible to keep the cantilever oscillation amplitude precisely
constant at its set point value by feedback control, the sample
surface would look perfectly ﬂat when viewed from the cantilever.
In reality, however, this is not the case because of an inevitable
time delay in feedback control. Several electronic and physicaldevices are contained in the feedback loop, each of which
responds to the corresponding input with a certain time delay,
and, therefore, the Z-scanner cannot move “at the same time” asthe change of sample height.
Here, we brieﬂy discuss how the imaging speed limit of AFM isdetermined by the time delay τ0 of the feedback loop.28 For
simplicity, the sample surface is assumed to have a sinusoidal
shape with a periodicity λ in the X−Z plane (Figure 2a). This
assumption is a justiﬁed generalization of the real situation
because any sample surface can be expressed as a Fourier integralof sine and cosine functions. When the sample stage is scanned in
the X-direction at velocity Vs, the sample height under the tipchanges with time at frequency f = Vs/λ. Therefore, the Z-scanner
is moved at the same frequency f in the direction opposite to thesample height, with a phase delay θ = 2πfτ0 with respect to thechange of the sample height under the tip during scanning(Figure 2a). Because of this feedback error, in the uphill region of
the sample the cantilever oscillation amplitude A is smaller thanAs, whereas in the downhill region A is larger than As. Therefore,
in the uphill region the tip is pushed more strongly into thesample than the set point force Fs, while in the downhill region
the interaction force is smaller than Fs (Figure 2b). Fs is

Fs = kc[(1 − α) × (A 0 2 − A s 2 )]1/2 /Q c(2)where α (0 < α < 1) represents the ratio of amplitude reduction
caused by the cantilever resonance frequency shift over the total
amplitude reduction. As described in the next section, α isapproximately 0.5 for short cantilevers optimized for HS-AFM.
When the excessive force in the uphill region is too excessive, thebiological sample is deformed and its biological activity iscompromised or completely lost. When As is set closer to A0 sothat Fs becomes smaller, the tip−sample interaction force
becomes zero in the downhill region (Figure 2c), meaning thatthe tip loses contact with the sample surface at its bottom swing,
resulting in inability to acquire the sample height information.
Once the contact is lost, the error signal (A − As) is saturated at asmall value of (A0 − As), irrespective of how far the tip is apartfrom the sample surface at its bottom swing. Therefore, once thecontact is lost, it takes long to bring the tip back to sample
interaction (i.e., “parachuting”). The maximum possible phasedelay not causing parachuting is approximately 10 times smaller
than the phase delay limit acceptable for not producing excessive
force of disturbing protein functions in the uphill region.28,73
Thus, maybe counterintuitively, tip parachuting is the major
obstacle in simultaneously achieving both high-speed and lowinvasive performances of AFM.
This problem has been solved by a new feedback control
technique called “dynamic PID control”, in which a false error
signal proportional to the real error signal is automatically added
to the real error signal when A exceeds As (i.e., when the downhill
region of the sample is being scanned).83 This has proveneﬃcient to bring the tip back into sample contact even at very
high scan velocities. Therefore, to estimate the limits of speed
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impulsive force (i.e., the product of the force acting on the sample
and the time during which this force is applied). The time during
which the force acts is short (∼100 ns or less, i.e., ∼10% of theoscillation cycle of ∼1 μs) for small cantilevers with 1.2 MHz
resonance frequency in water. Thus, the magnitude of theimpulsive force is small, in the atto-Newton second range (∼5 ×
10−18 N s).
The low-invasiveness can also be considered in terms ofenergy. An oscillating cantilever loses part of its oscillation energy
during every tap by (1 − α) × kc(A02 − As2)/(2Qc) on average.
For the same conditions as those detailed in the aforementioned
imaging studies, this energy loss only amounts to 1.2−2.2 kBT(when α = 0.5), where kB is the Boltzmann constant and T is thetemperature of 300 K in the experiments. Even when theoscillation energy is completely lost at once by the tip−sample
interaction, it amounts only to 24 kBT (when α = 0), comparable
to the energy of ATP hydrolysis ∼20 kBT. In actual imaging
experiments, such events of complete oscillation energy loss
happen only very rarely, if at all.

and invasiveness, we focus on the eﬀect of feedback phase delayon the sample damage. Denoting θmax as the maximum allowable
phase delay at which the sample is not aﬀected by excessive force,the highest possible feedback frequency f max is given by f max =θmax/(2πτ0), and hence the highest possible scan velocity in the
X-direction Vsmax is given by Vsmax = λθmax/(2πτ0). When the scansize in the X-direction is W and the number of scan lines is N, thehighest possible imaging rate Rmax frames/s (fps) is given by Rmax
= λθmax/(4πNWτ0). Instead of using τ0, we can use a morepractical parameter, the feedback bandwidth f B at which π/4phase delay occurs, that is, f B = 1/(8τ0). Thus, Rmax is ﬁnallyexpressed as
R max = 2λθmaxfB /(πNW )(3)

f B varies depending on the ratio between sample height h0 andthe cantilever free oscillation amplitude A0 but is generally ∼100
kHz when proteins with height 1−5 nm are imaged, in our HSAFM instrument.23 According to our experiences of HS-AFM
imaging of proteins, θmax is approximately π/9.73 Therefore,
under the conditions of N = 100, W = 150 nm, and λ = 10 nm,
Rmax is about 14.8 fps.

5. SMALL CANTILEVERS
For the above-mentioned imaging studies, custom-made smallcantilevers (Olympus BL-AC7DS-KU5) have been used that are
6−7 μm long, 2 μm wide, and 90 nm thick (Figure 3a). They have

4. MINIMIZING INVASIVENESS: THE TIP−SAMPLE
INTERACTION
For HS-AFM imaging, small cantilevers with high resonant
frequencies in water fc of 400 kHz to 1.2 MHz are used.
Therefore, the sample is tapped many times by the oscillating
cantilever tip during successive image acquisition. Here, weestimate the actual numbers of tapping for two cases. When an
image of a sample area Wx × Wy nm2 is captured at an imaging
rate R fps, a molecule with a top surface area (Atop nm2)
perpendicular to the tip is tapped fc/R × Atop/(WxWy) times perframe. Under the conditions (fc ≈ 1.2 MHz, R = 12.5 fps, and Wx
× Wy = 45 × 22 nm2) used for the noninvasive imaging of the
α3β3 subcomplex of the F1-ATPase,52 one α3β3 subcomplex (Atop∼100 nm2) is tapped approximately 9600 times per frame and ∼5
million times in total during the 40 s of successive imaging.
Under the conditions (fc ≈ 1.2 MHz, R = 7 fps, and Wx × Wy =150 × 75 nm2) used for the noninvasive imaging of tail truncatedmyosin V moving on an actin ﬁlament,48 one of the myosin’s two
motor domains (Atop ∼25 nm2) is tapped with the tip ∼380 times
per frame and ∼60 000 times in total during the 23 s movie
acquisition time. The α3β3 subcomplex and the myosin V did notshow any activity loss throughout the image acquisition periods.
Thus, the energy transferred from the oscillating cantilever to thesample does not accumulate in the biological sample but
dissipates quickly, within a time range much shorter than theoscillation period of the cantilever of ∼1 μs, into many degrees of
freedom of many atoms including those of surrounding watermolecules.
The other mechanical parameters of small cantilevers used inthese imaging studies are kc ≈ 0.2 N/m and Qc ≈ 2 in water. The
free oscillation amplitude A0 and set point amplitude As were set
approximately at 1 nm and 0.8−0.9 nm, respectively. Therefore,
according to eq 2, the set point force Fs is approximately 30−42
pN. However, due to the phase delay in feedback control theactual applied force is higher; a maximum force as large as ∼80
pN must sometimes have been exerted during the scanning overthe uphill regions of the samples. Why does this magnitude offorce not disturb the function of these biomolecules? We shouldﬁrst note that the mechanical quantity that aﬀects the sample(i.e., causes a change in momentum) is not the force but the

Figure 3. Small cantilever. (a) Scanning electron microscopy (SEM)
image of small cantilever. The inset shows an electron-beam-deposited(EBD) tip grown on an original bird-beak-shaped tip. (b) Magniﬁed
SEM image of EBD tip.

a resonance frequency fc ≈ 1.2 MHz in water (3.5 MHz in air), a
quality factor Qc ≈ 2 in water, and typical spring constant kc ≈ 0.2
N/m. The commercially available small cantilevers are 9−10 μm
long, 2 μm wide, and 130 nm thick, and have fc ≈ 0.6 MHz inwater (1.5 MHz in air), Qc ≈ 2 in water, and kc ≈ 0.1 N/m(Olympus BL-AC10DS-A2), or 6 μm long, 2 μm wide, and 200
nm thick, and have fc ≈ 0.4 MHz in water (1.2 MHz in air), Qc ≈2 in water, and kc ≈ 0.15 N/m (NanoWorld UCS-1.2, Neuchâtel,
Switzerland). Small cantilevers have advantages in addition totheir fast response to tip−sample interaction. As indicated by eq
1, the cantilever resonance frequency is increased by repulsive
tip−sample interaction, and this increase becomes greater withincreasing ratio fc/kc. Small cantilevers designed for HS-AFM
have large fc/kc ratios, approximately 1000 times larger ascompared to conventional cantilevers. Therefore, with smallcantilevers an increase of the resonance frequency results in adecrease (or increase) of the oscillation amplitude depending on
the excitation frequency used fex, as shown in Figure 4. The
resonance frequency increase also results in a phase advance
relative to the excitation signal, as shown in the bottom of Figure
4. Roughly speaking, the amplitude decreases when the excitation
frequency is lower than the free oscillation resonance frequency
fc, and vice versa. The most eﬀective excitation frequency fexm forgaining the largest amplitude decrease is approximately ex3125

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The last advantage of small cantilevers is that they enhance thesensitivity of cantilever deﬂection detection by the OBD
detection method. The deﬂection of a cantilever alters theangle of its free end (Δφ), which is detected by the OBD detector(see Figure 1). For a given displacement Δz of the cantilever free
end in Z-direction, a shorter cantilever results in a larger angle
change following Δφ = 3Δz/2L, and thus gives higher
displacement detection sensitivity. Because of this highsensitivity and the strongly diminished thermal noise eﬀect
with short (6−10 μm) cantilevers, the displacement as small as∼0.1 nm can be detected even with high bandwidth detection (afew MHz). This high detection sensitivity allows better force
control and hence greatly contributes to the low-invasive
performance of HS-AFM.
In AFM, the spatial resolution in the lateral direction depends
largely on the cantilever tip apex radius. The tip apex radius ofcommercially available small cantilevers is not necessarily smallenough to acquire high-resolution images. Now short cantilevers
with sharp electron beam deposited (EBD) or carbon nanoﬁber
tips are commercially available. They are however expensive andnot of perfect reproducible quality. When a scanning electron
microscope (SEM) and a plasma etcher are available, sharp EBD
tips can be fabricated in the laboratory, and, moreover, used shortcantilevers featuring EBD tips can be recycled many times,
followed by cleaning in the plasma etcher. As a sublimate forgrowing EBD tips, phenol powder is used to control thechemistry of the deposit.73
A small amount of the phenol powder is placed in a container
with holes of ∼0.1 mm in diameter in the lid located inside theSEM chamber. Short cantilevers are placed on the lid, near theholes to favor deposition of decomposed phenol vapors onto the
HS-AFM tip. A spot-mode electron beam is focused onto eachoriginal HS-AFM cantilever tip, which produces the growth of astylus composed of amorphous carbon at a growth rate of ∼17
nm/s. The focused electron beam is irradiated onto the original
tip for ∼1 min typically, which results in ∼1 μm length of an EBD
tip with an apex radius of 15−25 nm, hence providing an
excellent aspect ratio. The EBD tip can then be sharpened byargon or oxygen plasma etching, which reduces the apex radius to
4−5 nm (Figure 3b). In the best case, it can be reduced to <1 nm,
although this occurs rarely, ∼10% of the etching procedures.73
The mechanical durability of this sharp tip is not superb but stillgood enough to perform several experiments. It is notable that
the EBD tip can be completely removed by oxygen plasma
etching over one day. Therefore, a small cantilever chip can bereused many times as long as the cantilever is intact and new EBD
tips can be grown onto it and sharpened.

Figure 4. Amplitude and phase response curves of a cantilever as afunction of excitation frequency when the cantilever is freely oscillating(solid line) and when the cantilever tip is inﬂuenced by negative force
gradient (repulsive regime of tip−sample interaction) without energy
dissipation (broken line). This interaction increases the cantilever’s
resonant frequency. When the cantilever is excited at frequency fex, theresonant frequency shift results in amplitude reduction by ΔA and inphase advance by Δθ.

pressed as fexm = fc × (1 − 0.35/Qc).84 Thus, there are two typesof amplitude decreases: one due to energy dissipation andanother due to the resonance frequency change. These decreases
are most eﬀectively added when the excitation frequency fexm isused, providing high sensitivity for tip−sample interaction
detection. According to measurements with small cantilevers,
the tip−sample contact induces phase advances by ∼11° under
the practical conditions used for noninvasive imaging of proteins.
From a numerical analysis, this amount of phase advance
contributes to the measured amplitude decrease by approximately 50%, when fex is optimized at fexm.
Another advantage of small cantilevers is the small thermal
noise eﬀect on amplitude detection. According to theequipartition theorem, the average cantilever deﬂection by
thermal excitation is a function of only temperature T and thespring constant kc:
Δz 2 1/2 = (kBT /kc)1/2(4)where Δz is the cantilever deﬂection at the free end. The
averaged thermal deﬂection amounts to 0.14−0.2 nm at 300 K
for a cantilever with a spring constant kc = 0.1−0.2 N/m. The
average deﬂection values of 0.14−0.2 nm are comparable withthe amplitude damping (A0 − As) = 0.1−0.2 nm used fornoninvasive imaging of proteins. However, the thermal noise
density is distributed over a large frequency range from 0 to
approximately 2fc, that is, 0−2 MHz. Therefore, the thermal
noise density is small for small cantilevers because fc is high, ascompared to conventional cantilevers where the total thermal
noise density is accumulated within a narrow frequency range ofabout 0−30 kHz. In oscillating AM mode, the frequency regionused for imaging is approximately the feedback frequency fcentered on the resonance frequency fc. Thus, a band-pass ﬁlter
cleans the diﬀerential output signal of the segmented photodiodesensor by eliminating the thermal noise substantially. Thus, for acantilever with a higher fc, the thermal noise has less eﬀect on the
cantilever amplitude measurement.

6. HYDRODYNAMIC PRESSURE
In our HS-AFM instrument,23 which is now commerciallyavailable (HS-AFM 1.0, RIBM, Tsukuba, Japan), the sample
stage (and not the cantilever tip) is scanned in all three
dimensions (lateral dimensions X, Y and vertical dimension Z),
which simpliﬁes the instrument structure and gives additional
stability to the OBD detection. In HS-AFM imaging of proteins,the Z-scanner is displaced at 20−100 kHz. This high-frequency
displacement produces and exerts high hydrodynamic pressure,particularly to the cantilever and its supporting base positioned
near the sample stage surface.85 Under such hydrodynamic
pressure, the cantilever and the chip itself tend to move, often
deteriorating the dynamic performance of the cantileverresponse to relevant tip−sample interactions.73 In the worst
case, the cantilever response delay becomes longer than the3126

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Reviewsection 8.8.1) and (v) (described in detail in a complementary
review article72). More details about imaging studies on surface
conditions (i−iii) are also described elsewhere.38,73,87,88

minimized delay of the feedback loop, meaning that thehydrodynamic perturbation becomes a major limiting factor for
HS-AFM operation. To minimize the hydrodynamic pressure,the dimensions of the sample stage must be designed as small as
possible, 1−2 mm in diameter.73,85,86 To further circumvent
disturbance by hydrodynamic pressure, the lateral overlap
between the sample stage and the cantilever chip must be
minimized, meaning that only objects placed at the peripheral
regions of the sample stage should be imaged.73,86 Moreover, thecantilever tip (tip together with EBD tip) should be longer than
2.5 μm. When the tip is shorter than this value, the cantilever getsclose to the sample surface in a distance range where the solution
conﬁned between them is signiﬁcantly “squeezed” at the bottomof cantilever swing, resulting in oscillation damping as well as in ablunt change of the oscillation amplitude upon weak tip−sample
interaction. This leads to a drastic deterioration of the cantileverdeﬂection detection sensitivity during imaging.73,86 Commercially available short cantilevers do not suﬃciently consider the
problem raised by hydrodynamic perturbation and the tip lengthis only ∼1 μm, highlighting the necessity for EBD tip growth inlaboratories to achieve successful HS-AFM operation.

7.1. Mica Surfaces

The surface of freshly cleaved natural muscovite or synthetic
ﬂuorophlogopite mica has frequently been used for AFM
imaging because it provides atomically ﬂat surfaces over large
areas. The mica surface is negatively charged in aqueous solution
because K+ ions located between the cleaved layers are desorbed
from the surface upon cleavage and contact with water.
Therefore, the cleaved surface facilitates electrostatic adsorptionof positively charged molecules. Monovalent cations (K+, Na+,and Li+) weaken the adsorption aﬃnity for many proteins in theorder K+ > Na+ > Li+. In analogy, strongly negatively charged
molecules such as DNA can be adsorbed on bare mica surface
through the addition of divalent cations including Mg2+, Ca2+,
Ni2+, or Zn2+ in the buﬀer. Because electrostatic adsorption isnonspeciﬁc, proteins adsorbed on the bare mica surface cannot
be expected to have speciﬁc orientations on the surface but some
proteins do so. For example, GroEL, a barrel shaped moleculewith a double-ring structure, tends to bind to the surface in theend-up orientation.24,30 Probably such preferential adsorption
orientation is due to speciﬁc uneven charge distribution on thesurface of the biological molecule. The bare mica surface isparticularly useful for fractionated native membranes or SLBs
containing membrane proteins because a thin water layer (∼1
nm thick) separates the lower membrane surface from the mica
surface,89,90 sometimes resulting in no direct interaction of theproteins with the mica surface. In fact, as observed on bR in the
purple membrane and outer membrane F (OmpF) inreconstituted membranes, molecules in the noncrystallized
regions rapidly diﬀuse in such supported membranes.35,59,62
The mica surface can be functionalized in several ways. The
surface contains silanol Si−OH, which can react with organosilanes to form siloxane bonds Si−O−Si, and hence various
functional groups can be introduced to the surface.91 3Aminopropyltriethoxysilane (APTES) has frequently beenused to graft NH2 groups to the mica surface, aﬀording a
positively charged surface. However, caution has to be taken in
the preparation because APTES contains three ethoxy groups
and can therefore polymerize laterally and vertically in aqueous
solution, often resulting in increased surface roughness. 1-(3Aminopropyl)silatrane (APS) also can be used for the same
purpose.66,92 APS is less reactive than APTES and resistant tohydrolysis and polymerization at neutral pH, which guarantees
smooth surfaces.88 The NH2-functionalized surfaces can bereacted with various chemicals for further functionalizations. For
example, the reaction with the homobifunctional reagent
glutaraldehyde can provide an amino-reactive surface, onto
which proteins can be covalently immobilized, often in speciﬁcorientations. This strategy was used for HS-AFM observation of
F1-ATPase52 and isolated bR trimers,62 as described in sections
8.3 and 8.5.2, respectively.

7. SUBSTRATE SURFACES
The choice of substrate surfaces on which samples are placed is a
key to successful dynamic imaging by HS-AFM. Unlike singlemolecule ﬂuorescence microscopy, the surface roughness should
be small enough to identify molecules of interest that havetypically single nanometer dimensions, and in the mostdemanding cases the surface should be atomically ﬂat. Moreover,because AFM can visualize the sample only from one direction
perpendicular to the substrate surface, sample molecules
sometimes have to be attached to the surface in a speciﬁcorientation so that their characteristic structural features andportions of interest of the molecules can be visualized. These
requirements are common in both static and dynamic AFM
imaging. However, in dynamic AFM imaging, sample molecules
should not strongly interact with a surface so as to be kept
functionally intact. Nevertheless, when the interaction is too
weak, the molecules detach from the surface or move too fastalong the surface to be clearly imaged with HS-AFM. When
dynamic events of the interaction between two diﬀerent species
of molecules are to be imaged, selective immobilization is often
required, meaning that only one species of molecules areimmobilized onto the surface without loss of biological activity,
while the other species of molecules should be free from thesurface and interact only with the surface-immobilized partner.
Thus, establishment of various substrate surface (as well as the
sample) conditions is required depending not only on theproperties of molecules to be imaged but also on the nature of the
dynamic events to be studied.
Thus far, ﬁve types of surfaces have been used for HS-AFM
imaging of biological molecules: (i) bare or chemically modiﬁed
mica, (ii) supported lipid bilayers (SLBs) formed on mica, (iii)
two-dimensional (2D) crystals of streptavidin grown on planar
lipid bilayers containing biotin lipid,34,38,87 (iv) highly oriented
pyrolytic graphite (HOPG),36,57 and (v) DNA origami tiles
placed on mica surface.72 Native membranes are usually placedon bare mica surface, very similar to condition (ii). In a rare case,membrane samples are placed on a substrate surface withnanowells, so that narrow portions of the membranes are
suspended.41 Live cells are often attached to chemically modiﬁed
mica surfaces.68−71 Characteristics and use of these surfaces are
summarized in this section except for (iv) (brieﬂy described in

7.2. Lipid Bilayer Surfaces

The surfaces of self-assembled monolayers (SAMs) and bilayersof lipids have been used as substrate surfaces not only for AFM
imaging but also for various other purposes.93 The lipid SAMs
have been prepared using the Langmuir−Blodgett method; amonomolecular layer of lipids is ﬁrst formed on the water−air
interface and then transferred to a solid substrate surface bypulling the surface out of the liquid phase. This method requires a
special apparatus and skills. In comparison with this method, SLB
3127

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artiﬁcial membranes on a solid surface can be prepared moreeasily and are more amenable for biological studies than lipid
SAMs.94,95 Deposition of small unilamellar vesicles (SUVs, or
small liposomes) on the bare mica surface leads to the rupture ofliposomes by electrostatic interactions with the surface, followed
by fusion of the small SLB patches and resulting in the formationof a SLB with negligible surface roughness covering essentiallythe entire mica support. The surfaces of SLBs formed with lipids
having electrically neutral zwitterionic polar head groups, such as
phosphatidyl choline (PC) and phosphatidyl ethanolamine(PE), are known to be unfavorable for nonspeciﬁc binding ofproteins.96,97 Starting from a PC or PE SLB, various types of
surfaces can be prepared by including diﬀerent types of lipids inthe electrically neutral SLB.98,99 Therefore, the surface of SLBs
can be used for electrostatic adsorption or speciﬁc immobilization of proteins.87 Unlike the bare mica surface, the surface
charge density and polarity can easily be controlled using thelipid bilayer strategy, facilitating the adjustment of its aﬃnity forproteins. For example, lipids with the following charged headgroups are available: negatively charged head groups phosphatidylserine (PS), phosphoric acid (PA), and phosphatidylglycerol(PG); positively charged head groups trimethylammoniumpropane (TAP) and ethylphosphatidylcholine (EPC). Importantly,
various lipids having functional groups, such as biotin and Ninitrilotriacetic acid (NTA), at their polar heads are commerciallyavailable. They enable speciﬁc attachment of streptavidin- or Histag-conjugated proteins to the surface of SLBs.87 When
streptavidin is immobilized on the biotinylated lipid bilayer,
biotinylated proteins can further be immobilized on top of thestreptavidin.
SLBs are highly ﬂuid when lipids with unsaturated alkyl chainssuch as DOPC (1,2-dioleoyl-sn-glycero-3-phosphocholine) are
their main constituents. When protein molecules are bound withlow surface density to the surface of such SLBs, they diﬀuse too
fast to be imaged even with HS-AFM.87 However, when thesurface density of proteins is high enough for neighboring
protein molecules to interact with each other, the molecular
diﬀusion is signiﬁcantly decreased. In some cases, proteins form
2D crystals.100 When dense protein packing or 2D crystallizationis achieved, only a speciﬁc protein surface is exposed, and hence
high-resolution imaging is possible, as demonstrated with 2D
crystals of streptavidin101 and annexin V102 formed on thesurfaces of ﬂuid SLBs that contain biotinylated lipids ornegatively charged lipids, respectively. When proteins bound toa SLB surface at low density are to be observed, lipids withsaturated alkyl chains such as DPPC (1,2-dipalmitoyl-sn-glycero3-phosphocholine) should be used as a main component of the
SLB to attain low ﬂuidity. For example, low ﬂuidic SLBs withbiotin lipid were used to immobilize partially biotinylated actin
ﬁlaments in the HS-AFM imaging study of myosin V walking on
the ﬁlaments.48

Figure 5. Streptavidin and its 2D crystal. (a) Schematic of streptavidin
molecule placed on biotin lipid-containing lipid bilayer surface. ○,biotin-binding sites unoccupied with biotin; ●, biotin-binding sites
occupied with biotin. (b) AFM image of streptavidin 2D crystal formedon biotin-containing SLB (top) and height proﬁle along the white line
shown in the AFM image (bottom).

exposed to the aqueous environment and accessible (Figure 5a).
Therefore, biotinylated samples can be speciﬁcally immobilizedon the surface of streptavidin 2D crystals.38,87 This surface isparticularly useful for selective immobilization of homooligomeric proteins such as actin ﬁlaments, GroEL, and ATPases
associated with diverse activities (AAA) proteins because a stablepinning of the complexes at multiple biotinylated sites is possible.
In fact, the 2D crystal surface has been used for HS-AFM imagingof actin polymerization processes at the plus and minus ends ofan actin ﬁlament,38 dynamic association and dissociation
between GroEL and GroES,38 Ca2+-induced conformationalchanges of calmodulin,38 and ATP-invoked structural changes ofan AAA ATPase, p97.103 For monomeric proteins, the tetheringto the surface through a single biotin binding site is locally ﬁxed
but can result in a rotational motion around the link. This
mobility can however be reduced using reactive dibiotin
compounds. Moreover, His-tag-conjugated proteins can be
speciﬁcally immobilized on the surface of streptavidin 2D
crystals by the attachment of biotin-NTA to the surface.

8. DYNAMIC IMAGING OF PROTEINSIn this main section of this Review, we show various types ofmolecular imaging on proteins (and one case of lipids) by HSAFM, which have been conducted in the past few years. The
variety of studies could be classiﬁed in several ways: according tophenomena such as structural changes and diﬀusional processesor according to the nature of the proteins, soluble or membraneassociated. Here, we have decided to discuss the HS-AFM
achievements in topical groups of (i) motor proteins (myosin V
and F1-ATPase), (ii) structural changes of membrane proteins(bR, Ca2+-pump, P2X4 receptor), (iii) diﬀusion and dynamic
interaction of membrane proteins (bR, c-rings of ATP-synthase,aquaporin-0 (AQP0), and OmpF), (iv) self-assembly processes(amyloid-like ﬁbril formation, 2D crystallization of annexin A5,
SLB formation), and (v) other topics (cellulase enzymatic action,
intrinsically disordered proteins, and diﬀusion of point vacancy
defects in 2D protein crystals). These are described below in this7.3. Surface of 2D Crystals of Streptavidin

Streptavidin is a homo tetramer with dihedral D2 symmetry(Figure 5a). Importantly, it is not favorable to nonspeciﬁc
binding of many proteins, while each subunit has a high aﬃnity
biotin binding site. Streptavidin 2D crystals are easily formed on
the surface of a ﬂuid SLB containing biotin-lipid. These 2D
crystals are monomolecular layers with surface roughness <0.2
nm (Figure 5b), given appropriate 2D crystallization conditions(pH and ionic strength) have been chosen.38,87 On the SLBs, twoof the four biotin binding sites of streptavidin face the lipid
bilayer and are occupied by biotin, whereas the other two are3128

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Figure 6. Structure of myosin V, actomyosin ATPase reaction scheme, and prevailing view on conformational states of myosin. (a) Schematic ofmolecular structure of myosin V. (b) AFM image of myosin V. (c) Reaction scheme of actomyosin ATPase. (d) Schematic for backward leaning
prestroke and forward leaning poststroke conformations. The mark “+” indicates the plus end of an actin ﬁlament (red). The motor domain (yellow)
binds to an actin ﬁlament in the same orientation, in both conformational states, while the neck (blue) is leaned in diﬀerent directions. The forward leverarm swing is supposed to occur on the actin-bound head when Pi is released from myosin. The backward lever-arm swing is supposed to occur on the
detached head when ATP is hydrolyzed to ADP−Pi.

superfamily were also found to mechanically act on membranes
to perform remodeling of their morphology.110 Thus, thedeﬁnition of motor proteins has now been remarkably extended.
All proteins that perform mechanical work by hydrolyzing
nucleotides or using electrochemical energy are considered to be
motor proteins. During the chemical reaction cycle, they globallychange their structure in a large extent as well as the mode ofinteraction with partner or substrate proteins, DNA, or subunits
that they are constituted of. These dynamic changes produce
mechanical actions, such as pushing, pulling, unwinding,winding, rotating, squeezing, transporting, and sliding. Thesemechanical actions in themselves are their biological function.
Therefore, the direct visualization of these dynamic changes atthe single molecular level is considered to show their functions
themselves on screen. As such, these diverse motor proteins are
excellent targets for HS-AFM analysis.
There are many ring-shaped motor proteins comprisingmultiple ATPase subunits, represented by F1-ATPase, GroEL,
and AAA proteins. The ATP-binding sites of these proteins are
typically positioned at the interface between adjacent subunits,suggesting coupling of ATP binding and hydrolysis events withchanges of intersubunit interactions followed by a synchronizedglobal change in the ring structure; the chemical state ofindividual subunits would be inﬂuenced by the intersubunit
interactions and vice versa. Therefore, the interplay between
subunits is expected to elicit cooperative ATP binding andhydrolysis and possibly rotational propagation of the chemical
and conformational states over the ring, which drive themechanical actions tightly linked to the function of these
proteins.109,111 Detailed analysis of individual subunits and theirstructural changes by HS-AFM hence represents a powerfulnovel approach to unraveling expected but so far unstudied
dynamic events. If an ATP analog visualizable even with AFM isavailable, HS-AFM should also be able to visualize the expected
cooperative binding and hydrolysis of the ATP analog andconcomitant structural changes at the subunit level. Once
simultaneous detection by HS-AFM of the chemical andconformational states of an ATPase becomes possible, ourunderstanding of the fundamental problem of biological energy
transduction to mechanical work in ATPase motors will begreatly enhanced: How is the chemical energy liberated by ATPhydrolysis used to execute mechanical work? However, without
this simultaneous detection, HS-AFM imaging of myosin Vorder. Application studies on live cells, which have become
possible only very recently thanks to the development of fastwide-area scanners and the integration of optical microscopy
with HS-AFM, are shown in the following section. DNA andDNA−protein complexes are also important targets of HS-AFM.
In fact, several HS-AFM studies have already been performed onthese systems mainly by the Takeyasu, Lyubchenko, andSugiyama groups.31,33,40,54,55,58,65−67,104 However, we do notdescribe these studies here as they are reviewed elsewhere.72
Movies of molecular actions captured by HS-AFM havecorroborated several previous reports or reinforced speculated
phenomena concerning the structure and dynamics of proteins.
However, the dynamic molecular images have provided not only
corroborative “direct visual evidence” for inferences but also
solved long-standing questions that have previously been diﬃcult
or impossible to address by other approaches. Therefore, HSAFM has successfully provided more and deeper insights into
how the proteins operate to function. We emphasize this aspectin each application study described below, willing to attract broadinterest of chemical biologists and biophysicists for the use of
HS-AFM to unravel biological questions.
8.1. General Remarks on Dynamic Structural Study of Motor
Proteins

Quite a while ago, myosin and dynein were the only proteins
called motor proteins. These motors were shown to hydrolyze
ATP to convert the chemical energy of ATP into mechanicalwork. During the ATPase cycle, myosin interacts with actin
ﬁlaments to cause muscle contraction, while dynein interacts
with microtubules to beat cilia and ﬂagella. Along with theprogress of cell biology and molecular biology, actin105 and many
classes of myosins were found in all eukaryotes and discovered tobe responsible for several types of cellular motilities.106 Kinesin
was ﬁrst described in the squid giant axon107 and then in all
eukaryotes as a motor responsible for cargo transport along
microtubules and other mechanical functions. F1-ATPase wasfound to be a rotary motor that links ATP generation orhydrolysis to the directional transmembrane passage of protons
or ions.108 Functionally diverse AAA (or AAA+) proteins werealso found, which function by exerting force on substrate proteinsor protein aggregates to degrade, translocate, or disaggregate
them.109 DNA-dependent ATPases such as condensin, cohesion,and helicases were found to mechanically act on DNA totransform its structure or assemblies. GTPases of the dynamin3129

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Review(M5), which will be described in detail below, has elucidated theessence of this fundamental problem with nearly ultimate
clearness.

The processivity and nonprocessivity are determined, respectively, by whether or not the lifetime of the strongly bound A−
M−ADP is predominant over the other states (except for the A−
M state waiting for ATP binding).127 That is, in nonprocessive
myosins, Pi-release is the rate-limiting step, while in processive
myosins, ADP-release is rate-limiting.127
The key idea in the prevailing view on the chemo-mechanicalcoupling in myosin motility, which has been mainly derived frommuscle myosin studies, is that the myosin head is supposed totake two diﬀerent conformations, prestroke and poststroke
conformations corresponding to diﬀerent angles between the
motor domain and the neck domain (often called “lever-arm”),
depending on the nucleotide states (Figure 6d); the headassumes the prestroke conformation in the ADP−Pi bound state,while it assumes the poststroke conformation in the ADP-boundstate as well as in the ATP-bound state. As such, the detachedATP-bound head swings back the lever-arm upon ATPhydrolysis (i.e., ATP→ADP−Pi), while upon the chemicaltransition from ADP−Pi to ADP, the actin-bound head is
supposed to swing the lever-arm (around the neck−motor
domain junction) from the backward leaning orientation to theforward leaning orientation, resulting in exertion of sliding force
to the bound actin ﬁlament in the backward direction (Figure6d).123−126 The angle between the motor and neck domains thatis taken by the prestroke conformation facilitates the binding of
the ADP−Pi-bound head to actin in the backward leaningorientation and at the same time impedes its binding to actin in
the forward leaning orientation. The A−M−ADP−Pi state withthe prestroke conformation is considered to be a higher-energy
state; that is, the energy liberated by ATP hydrolysis is stored in
the myosin head. Upon Pi release, the myosin head is supposed
to use the stored energy by swinging the lever-arm forward to
execute a powerstroke, after which a lower-energy state of A−
M−ADP is supposed to be formed. For processive myosins, thisscheme has to be modiﬁed because there is a large chance thatboth heads are attached to actin. After Pi release, the leading headcannot swing the lever-arm that is pulled back by the actin-boundtrailing head. In this two-headed bound state where both heads
are bound to ADP, intramolecular tension is generated. As such,the energy liberated by ATP hydrolysis is considered to be stored
in the tension-generating state and eventually used for lever-arm
swing at the leading head that takes place following trailing headdetachment from actin. Direct HS-AFM imaging of processive
M5 has considerably shaken this concept, as detailed below.
8.2.1. Visualization of Lever-Arm Swing. The lever-arm
swing as powerstroke in muscle myosin was proposed by Huxley
in 1969.128 Since then, numerous studies were performed toobtain evidence for this hypothesis, not only for muscle myosin
but also for M5. Nevertheless, even with the use of singlemolecule techniques, no unambiguous experimental evidence forthis mode of action was attained. Instead, other models were
proposed, which brought long lasting debate and confusion129,130 (although at a higher level) in the ﬁelds of muscle
contraction and myosin motor action in general. The confusion
was gradually cleared up with time, but the swinging lever-arm
hypothesis has remained a proposal lacking solid data as it was
before, until its visualization by HS-AFM.48 This example of thelong lasting divergence of myosin functional hypotheses clearly
indicates a general diﬃculty of obtaining clinching evidence fordynamic actions of proteins even with the use of welldisseminated single-molecule techniques. In other words, thedegree of directness of measurement was not yet high enough.
Therefore, M5 lever arm motion was naturally chosen as a

8.2. Myosin V Walking on Actin Filament

The double-headed motor protein, M5 (Figure 6a,b), is amember of the myosin superfamily and functions as a cargotransporter in cells (see review112). Each “head” contains an Nterminal motor domain and a long neck domain consisting of six
IQ motifs, to each of which calmodulin is attached (in some casesa calmodulin-like light chain). After the neck region, M5 isdimerized by a coiled-coil tail, which is followed by a C-terminal
globular tail domain able to bind to various cargos via adapter
proteins. Unlike muscle myosin (i.e., myosin II), M5 does not
self-assemble into oligomers but operates as a single molecule.
Remarkably, M5 moves highly processively on actin ﬁlaments
over long distances,113,114 facilitating the observation ofindividual M5 molecules at work. M5 movement and motor
mechanism have extensively been studied by single-molecule
ﬂuorescence microscopy and optical-trap nanometry. Using
these technologies, it has been shown that M5 moves along actin
ﬁlaments toward the plus end113 in a “hand-over-hand”
manner,115,116 advancing 36 nm113,117 per ATP hydrolysis
cycle.118 The 36 nm stride corresponds to a half pitch of theright-handed, double-helical structure of an actin ﬁlament, and,
therefore, M5 moves approximately on a plane (only slightly
biased for a left-handed spiral).119 “Hand-over-hand” means thatthe two heads step alternately, exchanging leading and trailingroles with each step, very much like “bi-pedal walking”. In
addition, the mechanism underlying the alternate steps was
suggested to arise from asymmetric kinetics of ADP dissociationfrom the two heads; ADP dissociation at the trailing head is more
accelerated than at the leading head120 and/or ADP dissociation
at the leading head is decelerated.118,120−122 However, despite
numerous and extensive studies, the heart of the motor
mechanism, that is, how the tension for the forward step isgenerated in the molecule, coupled with the ATPase reaction,and how the energy liberated by ATP hydrolysis is used, hasremained elusive. These central questions were previously
studied to a great extent on muscle myosin mostly byensemble-averaging techniques (see reviews123,124) and considered to have reached a consensus to some extent (although direct
and irrefutable evidence has been lacking). The chemomechanical coupling and nature of the chemical energy usage
in M5 has been considered to be the same as (or very similar to)that in muscle myosin (see reviews125,126). Yet, actual attempts to
examine these central issues have not been carried out until the
recent HS-AFM study on M5.48 Before describing this HS-AFM
study in detail, the prevailing view of the chemo-mechanicalcoupling in an ATP hydrolysis cycle in myosin is summarized
below.
The simplest scheme of the actomyosin ATPase cycle is shownin Figure 6c. The nucleotide-free head tightly bound to actin
detaches from the actin immediately after binding to ATP,quickly followed by hydrolysis of the bound ATP to ADP−Pi.
When the ADP−Pi bound head is attached to actin, the bound Pi
dissociates from the head, which is followed by the formation of astrongly bound tertiary complex A−M−ADP (A and M denoteactin and myosin, respectively) and then by ADP dissociation,completing one ATPase cycle. The main role of actin in the
ATPase reaction is to accelerate the otherwise very slow Pi and
ADP dissociation from a myosin head. The degree of acceleration
for Pi and ADP dissociation varies among the classes of myosin.
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Figure 7. Tail-truncated myosin V (M5-HMM) movement on actin ﬁlament captured by HS-AFM. All of the images were taken at a frame rate of 7 fps.
(a) Successive AFM images showing processive movement of M5-HMM in 1 μM ATP when positively charged lipid is absent in the planar lipid bilayer(PLB) surface. The arrowhead indicates one of the streptavidin molecules attached to the PLB surface. (b) Successive AFM images showing processive
movement of M5-HMM in 1 μM ATP when positively charged lipid is present in the PLB. The arrows indicate the coiled-coil tail pointing to the minus
end of actin. The arrowhead indicates one of the streptavidin molecules attached to the PLB surface. (c) Clips of successive images showing longprocessive run of M5-HMM in 1 μM ATP (14 steps are recorded). (d) Schematic explaining structural features of two-headed bound M5-HMM
observed in the presence of nucleotides. (e) Successive AFM images showing stepping process in 1 μM ATP. The swinging lever is highlighted with athin white line.

streptavidin that were attached beforehand to the SLB with alow surface density. The tail-truncated M5 (M5-HMM) was notfound to adhere to this surface but only interacted with the actin
ﬁlaments, on which it moved unidirectionally. In various
concentrations of ATP, M5-HMM moved with velocities V
that followed a modiﬁed Michaelis−Menten relationship as afunction of ATP concentration:

paradigm target to illustrate the power of HS-AFM in providing
unambiguous direct data of complex molecular processes.48
To visualize moving M5 molecules on actin ﬁlaments, actin
ﬁlaments had to be immobilized on a surface, whereas M5
molecules had to be free enough from the surface to interact with
the immobilized actin ﬁlaments. To this end, the surface of an
SLB containing DPPC and biotin-cap-DPPE in a weight ratio of
9:1 was ﬁrst used as a substrate surface. Partially biotinylated
actin ﬁlaments were immobilized on the surface throughV = ds × [1/(k1[ATP]) + 1/k 2]−1
3131

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Reviewwhere ds is the step size, k1 is the second-order ATP binding rate
constant, and k2 is the ﬁrst-order ADP dissociation rate constant.
This relationship holds because ADP release is the major ratelimiting step,127 ATP binding is also a rate-limiting step at low
ATP concentrations (other chemical steps occur much faster),and each discrete translocation step is tightly coupled to one ATPhydrolysis.118 The observed step size was ds = 37.2 ± 12.5 nm,
irrespective of the ATP concentrations. The estimated values ofthe maximum velocity Vmax (≡k2ds) = 403 ± 7 nm s−1, Km for
ATP (≡k2/k1) = 5.4 ± 0.5 μM, k1 = 2.0 ± 0.9 μM−1 s−1, and k2 =10.8 ± 3.8 s−1 were identical to those measured by total internal
reﬂection ﬂuorescence microscopy (TIRFM) under the same
buﬀer and temperature conditions,116,131 indicating no eﬀects ofthe tip−sample and surface−sample interactions on motor
activity. However, as shown in Figure 7a, most of the molecules
moved perpendicularly to the substrate surface, so that theircharacteristic structure was not well resolved.
To facilitate weak sideways attraction of M5-HMM onto the
SLB surface, 5% of a positively charged lipid DPTAP (1,2dipalmitoyl-3-trimethylammonium-propan) was added to the
SLB. Although the maximum translocation velocity was reducedby the addition of DPTAP to 73% of the full velocity observed
without DPTAP, characteristic moving M5-HMM could clearlybe visualized in a sideways view, as shown in Figure 7b. This
reduction of velocity was mainly caused by the apparent
reduction of the second-order ATP binding rate constant. The
scan size was set to a small range (typically to about 130 × 65nm2) to achieve a relatively high imaging rate (7 fps) and aminimum mechanical load to M5-HMM molecules. However,processive runs (4−16 steps corresponding to 144−576 nm
translocation) in 0.1−2 μM ATP were observed, during which
the moving molecules were tracked by manual shift of the narrowscan area (Figure 7c). The occasional failure of continual tracking
occurred when the target molecule collided with another M5ahead on the actin ﬁlament, when a portion of discontinuous
substrate surface was encountered, or when the manual tracking
was not appropriately performed. The sideways HS-AFM
imaging of two-headed bound M5-HMM consistently exhibitedthe following structural features. The neck−motor domainjunction appears smooth in the leading head but is V-shaped in
the trailing head without exception (Figure 7d). The shortcoiled-coil tail is mostly tilted toward the minus end of the actin
ﬁlament (Figure 7b, arrows). These features are consistent withelectron microscopy observations132 and could be used todetermine the actin polarity when bound M5-HMM wasstationary.
Despite the successful visualization of moving M5-HMM
molecules, no detailed molecular behavior during a step could be
resolved because single steps were completed faster than theacquisition time of an individual frame (1/7 s). To slow the step
movement of M5, additional streptavidin molecules were placedon the substrate surface (the biotinylated SLB) as moderateobstacles to the walking of M5-HMM. This method allowed thevisualization of stepping processes as shown in Figure 7e. After
trailing head detachment, the leading head appeared tospontaneously rotate from the backward leaning orientation
toward the forward leaning orientation. Before completing thisrotation, the leading head brieﬂy halted by colliding with astreptavidin molecule placed in the way of its natural path, andthe detached trailing head could be visualized in a conformation
far away from the actin ﬁlament and slightly rotated around theneck−neck junction. The leading head then overcame thestreptavidin blockade and completely rotated to the forward

leaning orientation. Accompanied by this further rotation, thetrailing head attached to a forward site of the actin ﬁlament tobecome the new leading head, completing one step. Thus,
dynamic processes in the forward step were directly visualized athigh spatial resolution. The observed rotation of the leading headcorresponds to the swinging lever-arm motion to propel themolecule forward.128 Hence, by direct imaging, HS-AFM could
provide irrefutable evidence for the Huxley’s hypothesis. It isnoteworthy that before completing a step the detached trailinghead never interacted with actin and hence passively moved
forward, driven by the rotating leading head, ruling out somemodels that had suggested myosin motility to occur in the“inchworm” like model considered for kinesin133 or in the“biased diﬀusion” model proposed for single-headed myosin II
and for single-headed myosin V,134 in which the trailing head wasconsidered to slide on the actin ﬁlament to get close to and passbeyond the leading head.
8.2.2. Directional Rule. After the imaging of many
unidirectionally moving M5-HMM molecules, a “directionalrule” was noticed regarding the relationship between the moving
direction and the direction from which M5-HMM was bound toan actin ﬁlament. M5-HMM molecules that were moving fromleft to right were bound to actin ﬁlaments from top, while thosemoving from right to left were bound to actin from bottom asshown in Figure 8a. This rule was unrelated to the previously

Figure 8. Directional rule and its mechanism. (a) Typical HS-AFM
images of two-headed M5-HMM showing the directional rule; there is aﬁxed relationship between the moving direction (arrows) and the
direction (arrowheads) from which M5-HMM bind to an actin ﬁlament.
(b) Schematic showing M5-HMM bound to actin at the left ﬂanks of the
motor domains. (c) Schematic showing the reason for the directional
rule. In the M5-HMM shown in black, which does not follow thedirectional rule, the motor domains have to get into the narrow space
between the actin ﬁlament and the substrate surface, which hardly
occurs.

observed tendency of moderate left-handed spiral motion ofwalking M5 around a freely suspended actin ﬁlament,119 because
this directional rule held even in the absence of ATP, where theactin polarity could be judged from the above-mentioned
structural features of two-head bound M5-HMM. The directional rule suggested a geometry-speciﬁc positioning of the
motor domain on the actin ﬁlaments. As seen in AFM images of
two-headed bound M5-HMM (e.g., Figure 8a), the motor
domain looked brighter than actin, meaning that the boundmotor domain is taller than the top surface of the actin ﬁlament.
Hence, as shown in Figure 8b, the directional rule indicated that3132

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single-molecule polarized ﬂuorescence microscopy.137 Thus, thefoot stomp is an inherent behavior of M5.
The foot stomp at the leading head raised an importantquestion as to the chemo-mechanical coupling in this motor. The
brieﬂy detached leading head does not carry bound Pi because Pi
has already been released from the ADP−Pi-bound head sinceinitial attachment of the head to actin. It is well-known that Pi
release occurs immediately after the binding of an ADP−Pibound head to actin.127 Nevertheless, the detached only ADPbound leading head rebound to actin still in the backward leaningorientation, and then swung the lever-arm forward following
trailing head detachment, like in a normal step process. This
indicated that tension generation for forward movement canoccur without transitioning through the ADP−Pi bound state,
but directly in the ADP-bound state. Thus, as a major surprise,
the tension generation for forward movement does not seem torequire chemical energy be supplied by ATP hydrolysis.
This external energy-less tension generation was further
conﬁrmed by the HS-AFM observation of two-headed bound
M5-HMM in the presence of ADP; sometimes the short coiledcoil tail unwound, after which the monomerized leading head
immediately rotated toward the forward leaning orientation,
similar to the swinging lever-arm motion (Figure 9b). The
coiled-coil region unwinding suggested again that the distortionof the actin-bound leading head, which is caused just by binding
to actin without any chemical transition, is the source of theintramolecular tension generation for forward movement.
From these HS-AFM observations, the following scenario wasdrawn as to the tension generation and the lever-arm swing topropel the molecule forward. In the two-headed bound state, thetrailing head is in the forward leaning orientation, which is naturalat least for the ADP-bound or nucleotide-free head. On the other
hand, the leading head is not in the natural orientation (i.e., it is in
the backward leaning orientation) and therefore pays an energy
cost that generates the intramolecular tension. The energy cost is
not supplied by the ATPase reaction. Upon trailing headdetachment, the constraint keeping the bound leading head in
the unnatural orientation is released, and hence the leading headspontaneously rotates forward. This scenario is not consistent
with the commonly accepted view of chemo-mechanicalcoupling in myosin motors detailed above. According to thisformerly prevailing view, the leading head of M5-HMM, in theﬁrst place, would not bind to actin without going through the
M−ADP−Pi state, and in the second place, tension would not be
generated without transitioning from A−M−ADP−Pi to A−M−
ADP; the transition has been supposed to be an important step inwhich the chemical-to-mechanical energy transduction occurs.
Before discussion of how the prevailing view has to be changed
based on HS-AFM observations (section 8.2.5), more details of
the foot stomp are described below.
The foot stomp was observed not only in the presence of ATP
but also in the presence of ADP only and under nucleotide-free
condition, always with higher frequencies at the leading head. In
ADP and under nucleotide-free condition, detachment of thetrailing head, which was followed by a forward step, was only
rarely observed; that is, the foot stomp at the trailing head mostly
occurred as brief translocation along the actin ﬁlament. The
higher frequency of foot stomp at the leading head in any
nucleotide condition implied that the leading head’s aﬃnity for
actin is lower than the trailing head’s aﬃnity. This is consistent
with the fact that when binding to actin, the leading head has topay an energetic cost by generating intramolecular tension.
Interestingly, the foot stomp was frequently accompanied by a

the motor domains are bound to actin at their left ﬂank, provided
that the M5-HMM molecule faces the advancing direction. If an
actin ﬁlament was rotated about its long axis on the substratesurface, the outer surface of the motor domains would hit thesubstrate surface at some angle, as shown in Figure 8c. Thus, thedirectional rule originates from the steric hindrance eﬀect of thesurface, which is reinforced by a weak attractive surface−sample
interaction. An important conclusion derived here is as follows:when M5 stands on an actin ﬁlament and faces the advancing
direction, the motor domains do not grasp the top surface of theactin ﬁlament but grasp the right ﬂank of the ﬁlament, as shownin Figure 8b. Although not explicitly described before in electron
microscopy 3D reconstruction studies of the actin-bound M5
head, this conclusion is consistent with the 3D structure of thebinary complex.135
8.2.3. Foot Stomp and Tension Generation. Interestingly, while M5-HMM was in the two-headed bound state in
ATP containing buﬀer, the motor domain of the leading headfrequently exhibited brief dissociation and reassociation on thesame actin ﬁlament, whereas the molecule remained at
approximately the same position on the ﬁlament, as shown in
Figure 9a. Similarly, it was observed that the motor domain of the

Figure 9. M5-HMM’s foot stomping at the leading head, unfolding of
coiled-coil tail, and conformational transition of the leading headcaptured by HS-AFM. (a) Successive AFM images showing foot stomp
at the leading head in the presence of 1 μM ATP. As pointed by thearrowheads, the leading head are brieﬂy detached from actin. Because
the detached head is moving fast, it does not appear in the images. Frame
rate, 7 fps. (b) Unfolding of the coiled-coil tail of two-headed bound M5HMM. Left images, before unfolding; right images, after unfolding. (c)HS-AFM images showing conformational transition between straightand bent conformations of the leading head in the presence of 0.1 μM
ADP. The symbol “+” indicates the plus ends of actin ﬁlaments (b, c).

trailing head exhibited a brief translocation approximately by ±5nm along the actin ﬁlament. These behaviors were termed “footstomp”. The foot stomp of the leading head was observed about 3
times more often than stomping of the trailing head. Although
not well documented, a foot-stomp-like behavior was previously
suggested by ﬂuorescence microscopy observations of walking
M5 molecules.136 More recently, the foot stomp was further
conﬁrmed by the observation of walking M5 by high-speed
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Reviewdomains. The sharp kink is likely to occur to release the large
strain accumulated in the neck, suggesting that the neck−motor
domain junction is stiﬀer in the absence of nucleotides than thatof the nucleotide-bound head.
To examine the nucleotide-dependent ﬂexibility of the neck−motor junction, the distribution of forward leaning orientation of
single-headed M5 (Figure 11a) was compared between the
nucleotide-free and ADP-bound conditions. Under nucleotidefree conditions (Figure 11b,c), the orientation angle relative tothe long axis of the actin ﬁlament was 34 ± 10°. In ADP (Figure
11d,e), however, the angle distribution was wider with two peaks
at 29 ± 11° and 51 ± 15°, which may be relevant to two diﬀerent
ADP-bound states in equilibrium.123 Therefore, the hingearound the neck−motor domain junction of the ADP-boundhead is more ﬂexible than that of the nucleotide-free head. This
result may be related to the angle distributions measured byelectron spin resonance for a spin probe introduced to theregulatory light chain of myosin in rigor and contracting muscle
ﬁbers.139
The sharply kinked conformation that was often adopted by
the leading head under the nucleotide-free condition could
provide a useful indicator of whether or not the leading headcontained a nucleotide. Even in the presence of 0.1 μM ATP, theleading head neck was mostly straight (>98%), suggesting thatthe leading head almost always retains ADP until the trailing headbinds to ATP and then detaches from actin. In the presence oflow concentrations of ADP, the leading head exhibited alternate
switching between the straight and sharply kinked conformations. From the proportion (Figure 12a) and the lifetime (Figure12b) of the straight leading head as a function of ADP
concentration, the kinetic parameters for ADP binding/dissociation on the leading head were estimated: the ADP
dissociation constant, Kd = k2/k1 = 0.075 ± 0.013 μM; the ADP
dissociation rate constant, k2 = 0.100 ± 0.004 s−1; the secondorder ADP binding rate constant, k1 = 1.3 ± 0.3 μM−1 s−1. The
value of k2 is 70 times smaller than the corresponding rate for the
trailing head estimated above from the dependence of thetranslocation velocity on the ATP concentration (k2 = 7.2 ± 1.5s−1). This degree of asymmetry between the two heads is similar
to that previously reported.121 The value of k1 is 10 times smaller
than the corresponding value, 12.6 μM−1 s−1, previously
measured in a solution study on single-headed M5.127 The
ADP dissociation rate constant of 0.1 s−1 means that, on average,
one ADP is released from the leading head every 10 s. However,
M5-HMM walks many steps during 10 s. Thus, the sequential
events of ADP release, the subsequent ATP binding, and theresulting head dissociation take place solely at the trailing head,
which is the basis underlying the processive hand-overhand
stroke generation. This mechanism had been inferred fromvarious indirect experiments120−122 but was clearly and directly
demonstrated by the HS-AFM observation.
Because ADP never dissociates from the leading head while
the M5-HMM molecule is walking in the presence of ATP, thesharp kinking of the leading head in the nucleotide-free condition
has no relevance to powerstroke generation. Moreover, the brief
detachment of the leading head observed as a foot stomp in thepresence of ATP is not caused by ATP binding to the headbecause the sharply kinked conformation was never observed
before the brief detachment of the leading head.
8.2.5. Chemo-mechanical Coupling in Walking Myosin
V. How can the prevailing view of the chemo-mechanicalcoupling in myosin motors be changed or modiﬁed to bereconciled with the novel results that emerged mostchange in the head separation (the distance between the attachedmotor domains). The direction of the change was well correlatedwith the head separation before the foot stomp, as shown in
Figure 10a (for foot stomp at the leading head in 1 μM ATP) and

Figure 10. Displacement of motor domain of leading (a) and trailing (b)heads after foot stomping as a function of head separation before thefoot stomping. Each right-hand ﬁgure shows the histogram ofdisplacement.

Figure 10b (for foot stomp at the trailing head in 1 μM ATP).
That is, a shorter head separation before foot stomp tended toresult in a longer one after foot stomp, and vice versa. Histogramsof the displacement accompanied by a foot stomp had threepeaks separated by ∼5 nm, which was approximately the distance
between adjacent actin monomers in one strand of an actin
ﬁlament (righthand panels in Figure 10a and b). Very similar
relationships were also observed in the presence of ADP andunder nucleotide-free condition. These results indicated thatboth heads have a propensity of searching for the energeticallyfavorable binding sites on an actin ﬁlament, but the leading head
does that more often because its binding aﬃnity is diminished
due to the molecular stress (enabling the next forward step) thatit features.
8.2.4. Asymmetric ADP Dissociation Kinetics. Even
under the nucleotide-free condition, both heads of M5-HMM
were bound to the same actin ﬁlament. However, unlike in thepresence of nucleotides, where the leading head almost always(>95%) took a nearly straight conformation (slightly curved
outward), the leading head often exhibited a sharply kinked
conformation and alternated back and forth between thisconformation and the nearly straight conformation (Figure9c), indicating that these conformations are in equilibrium. The
hinge portion on the neck where the kink occurs was very close to
the motor domain. Although the structure was not well resolvedin the AFM image, the kink appeared to occur at the neck−motor
domain junction, as suggested by electron micrographs.138 In the
sharply kinked conformation, both heads had the forward leaningorientation around the motor domain, and thus both necks
appeared to emerge from the front of the respective motor
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Figure 11. Angle distributions of actin-bound single-headed M5. The single-headed M5 is formed by natural uncoiling of the short tail of two-headbound M5-HMM. (a) Schematic showing the angle measured. (b,d) HS-AFM images of bound single-headed M5 under the nucleotide-free condition(b) and in 1 mM ADP (d). The plus ends of actin ﬁlaments, which are judged from the directional rule shown in Figure 8, are indicated with “+”. The
measured angles are shown in the images. Imaging rate, 3 fps. Scale bar, 30 nm. (c,e) Histograms of the angle measured for bound single-headed M5
under nucleotide-free condition (c) and in 1 mM ADP (e). The red curves represent the results of the best ﬁt to the sum of two Gaussian distributions.

tional states.123−126 This view has to be changed as follows: amyosin head adopts two (or more) conformations even under agiven nucleotide (chemical) condition, and goes back and forth
between diﬀerent conformations, but its dynamic equilibrium isbiased to one side depending on the nucleotide condition. As
described in the supplementary data in a cryoelectron
microscopy study by Volkmann et al.,135 both bent and straightconformations appeared in the actin-bound single-headed M5 inthe presence of AMPPNP or ADP·AlF4, without a large
population diﬀerence. This means that the energy diﬀerence
and the energy barrier between the two conformational states is< ∼2 kBT (T = 300 K), much smaller than the chemical energy to
be liberated by ATP hydrolysis (∼20 kBT). Thus, the following
conclusion can be formed: although the conformational changeof the myosin head caused by the back swing of the lever-arm is
an eﬀective strategy for facilitating its binding to actin in the
backward leaning orientation, the energy required to produce theprestroke conformation is much less than that provided by ATPhydrolysis.
According to the prevailing view, the chemical energy liberated
by ATP hydrolysis is supposed to be converted to mechanicalenergy and stored in the actin-bound leading head of M5 with
ADP. This state of the leading head is supposed to be formed
only through Pi release at the head. The Pi release is supposed toinduce a pre- to poststroke conformational change in the leading
head, which generates intramolecular tension for forward
movement. In this view, the chemical energy liberated by ATPhydrolysis is supposed to be transduced to mechanical energy instraightforwardly from the HS-AFM observations describedabove?
According to the prevailing view, the leading head of M5HMM would not be able to bind to actin when the leading head isin the ADP-bound or nucleotide-free states because both heads
are supposed to assume a poststroke conformation (i.e., straightneck−motor domain junction). Moreover, the trailing head of
M5-HMM would not be able to bind to actin when AMPPNP(mimicking the posthydrolysis ADP−Pi) is bound to the trailinghead because the head is supposed to assume a prestroke
conformation (i.e., bent neck−motor domain junction). In
reality, both heads were bound to an actin ﬁlament in the
nucleotide-free and ADP-bound states as well as even in the
AMPPNP-bound state. The distinct prestroke and poststroke
conformations have been derived from electron microscopy and
X-ray crystallography studies of nucleotide-free and nucleotidebound heads of myosins from various sources.135,140−144 Most ofthese numerous studies agree that the neck−motor domainjunction is bent in the ADP−Pi state, so that the lever-arm swings
back toward the motor domain (i.e., direction opposite to theforward lever-arm swing) upon ATP hydrolysis. Although thisback swing to form a prestroke conformation facilitates theleading head of M5 to bind to actin in the backward leaningorientation, this conformation also occurs in the ﬂexible ADPbound head (otherwise, two-headed binding to actin could notoccur in the presence of ADP). The previous static structural
studies have often been interpreted as if a tight one-to-one
relationship would exist between the chemical and conforma3135

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supposed to spontaneously move forward once the trailing headdetaches from actin upon ATP binding. Energy is required tobreak the rigor bond formed by the actin-trailing head complex.
Therefore, ATP binding itself should supply the energy forbreaking the rigor bond of the trailing head.
How much energy is required for this bond to break? The
unbinding (rupture) force was previously measured to be 15 pN
using AFM.16 The rupture distance was also estimated to be 1.7−
2.5 nm in this study. This rupture distance is unusually long as
compared to, for example, the 0.23 nm for an antigen−antibody,145 the 0.15−0.3 nm for streptavidin−biotin,146 andthe 0.05−0.3 nm for α-actinin−actin dissociation.147 This longrupture distance is consistent with the fact that actin−myosininterfaces contain bonds formed between ﬂexible loops.148,149
From the values of rupture force and rupture distance, the energy
required to rupture a single actin−myosin rigor bond is 6−9 kBT(T = 300 K), which is 30−45% of the chemical energy of ATPhydrolysis (∼20 kBT). This mechanically estimated value is ingood agreement with the value of 9 kBT estimated biochemicallyfor the free-energy change in the dissociation A−M → A + M.150
For the ATP (T)-induced dissociation A−M + T → A + M−T,
the free-energy change is biochemically estimated to be ∼10
kBT,150 which is again similar to that for the mechanicaldissociation A−M → A + M. However, one may argue that ATP
is not yet hydrolyzed in the step where a rigor actin−myosin
complex dissociates, and, therefore, the energy of ATP hydrolysis
is not yet used during this phase. However, the account balanceof energy should be considered for the whole cycle of the ATPasereaction, and hence the ATP-binding energy can be used as amain part of the energy of ATP hydrolysis.
8.2.6. Remaining Issues in the Walking Mechanism of
Myosin V. The above argument on the chemo-mechanical cyclein walking M5 puts forward a new principle for the energy
transduction and motor function mechanism of M5, which issummarized below and can probably be extended to all myosins
with slight modiﬁcations.
(i) Only a very small fraction of the chemical energy liberated
by ATP hydrolysis is used for the back swing of the lever-arm toform the prestroke conformation. Even without any chemicalenergy input, the back swing can occur thermally, but its
probability is somewhat smaller than that of the chemicallyinduced case.
(ii) No energy is used for the generation of intramoleculartension for forward movement (i.e., no chemical-to-mechanicalenergy transduction takes place in tension generation).
(iii) The forward lever-arm swing of the leading headspontaneously occurs upon trailing head detachment, without
any chemical transitions at the leading head.
(iv) The chemical energy of ATP hydrolysis is mainly used todissociate the rigor complex of actin and the trailing head.
(v) The main role of ATP hydrolysis does not lie in supplying
energy but in ensuring the unidirectional mechanical cycle byshifting the conformational equilibrium to either of the pre- andpoststroke conformations.
The most striking feature in this novel view is that the strained
two-headed bound molecule can be formed without a chemicalenergy input and that M5 moves spontaneously forward once thetrailing head detaches from actin. If this was really the case, wewould, however, appear to encounter a perpetuum mobile
problem. Even in the presence of ADP, the foot stomp occurs at
both leading and trailing heads of a two-headed bound molecule,with more frequent occurrence at the leading head than at thetrailing head. When the trailing head detaches from actin, theFigure 12. Leading head conformations and their dependence on
nucleotides. (a) Population of straight and sharply bent conformations
under diﬀerent nucleotide conditions. The numbers in (frames) and[molecules] represent those of examined images and molecules,
respectively. (b) Average lifetime ⟨τST⟩ during which the leading headassumes the straight conformation in various ADP concentrations (thenumber of events used to obtain the average is shown in ⟨events⟩). The
error bar indicates standard error. The line curve indicates the best ﬁt
result obtained by analyzing the data using the following equation: ⟨τst⟩= nNF/kS−B + nADP/k−, where kS−B represents the spontaneous transition
rate of the nucleotide-free leading head from the straight conformation
to the sharply bent conformation, k− represents the ADP dissociation
rate constant for the leading head, and nNF and nADP are, respectively, thefractions of nucleotide-free and ADP-bound leading head under a given
ADP concentration, which can be calculated using the ADP dissociation
constant Kd of 0.075 μM estimated from the data shown in (a).

the form of intramolecular tension. Nevertheless, HS-AFM
observations indicated that the intramolecular tension can bedirectly generated by the binding of the ADP-bound leading headto actin without passing through the Pi-releasing step at theleading head. Thus, the energy liberated by ATP hydrolysis is not
used for the tension generation. Therefore, the way in which the
ATP hydrolysis energy is actually used for myosin V walking(perhaps also for mechanical work performed by all members ofthe myosin superfamily) must be reconsidered. Because thestrained (tension-generating) two-head bound state can beformed without the use of ATP hydrolysis energy, the molecule is3136

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Figure 13. HS-AFM imaging of α3β3 subcomplex of F1-ATPase under diﬀerent nucleotide states. (a) Crystal structure of α3β3γ subcomplex. The α-, β-,and γ-subunits are shown in cyan, pink, and yellow, respectively. (b) Typical HS-AFM image of α3β3 subcomplex immobilized on a chemically treatedmica surface without nucleotide (150 × 150 pixels; imaging rate 1 fps). (c) Filtered HS-AFM image of C-terminal side of the α3β3 subcomplex without
nucleotide. (d) C-terminal side of crystal structure of the nucleotide-free α3β3 subcomplex. The α and β subunits are colored in cyan and pink,respectively. The C-terminal DELSEED motif of the β subunits corresponding to the high protruding portions is highlighted in red. (e) Simulated AFM
image constructed from the structure in (d). (f) Filtered AFM image of the N-terminal side of the α3β3 subcomplex without nucleotides. (g) N-terminalside of the crystal structure of the α3β3 subcomplex. (h) Simulated AFM image constructed from the structure in (g). (i) Filtered AFM image of the Cterminal side of the α3β3 subcomplex at 1 mM AMP-PNP. (j) Crystal structure of the α3β3 subcomplex with bound nucleotides. This structure is
obtained by removing γ from the crystal structure of α3β3γ subcomplex. (k) Simulated AFM image constructed from the structure in (j).

molecule would take a forward step by the spontaneous swing
performed by the leading head. On the other hand, when theleading head detaches from actin, the molecule would not take abackward step because the trailing head is bound to actin in astable orientation (i.e., the forward leaning orientation). Thus,
without any energy input, the M5 molecule appears to step
forward many times (albeit slowly), even with the less frequent
occurrence of foot stomps at the trailing head. This is forbidden
perpetuum motion. However, in this gedanken experiment, two
factors are not well taken into consideration: (i) a single-headed
bound myosin V undergoes rotational Brownian motion aroundthe average position biased to the forward direction, which is dueto the ﬂexible neck−motor domain junction, and (ii) the
population of two conformations of the ADP-bound detachedhead is biased toward the poststroke conformation. Considering
these factors, the detached trailing head can have two possibilitiesof binding to the original or frontward actins, while the detachedleading head can have two possibilities of binding to the originalor backward actins. Thus, the principle of detailed balance would
hold between the forward step after trailing head detachment andthe backward step after leading head detachment, resulting in nonet movement of the molecule. The ratio of foot stomp
occurrence between the leading and trailing heads was observed
to be 4:1 in the presence of 1 mM ADP. This ratio of 4:1 appears
to be insuﬃcient for the detailed balance. However, as describedin section 8.2.3, the foot stomp at the trailing head mostly occurs
as a brief translocation approximately by ±5 nm along the actin
ﬁlament, which does not allow the forward swing of the leading
head. Therefore, when the brief translocation is omitted and only
the events of brief detachment from actin are counted as footstomp, the ratio of foot stomp occurrence between the two heads
becomes much larger, guaranteeing the detailed balance.
The novel principle mentioned above predicts that even in thepresence of saturated ADP, M5 would processively move
forward if the trailing head is repeatedly detached by somemeans instead of ATP. If each head of M5 was labeled with a
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conformational and catalytic states of individual β subunits tosequentially generate torque.162 This proposal appeared
reasonable because the γ subunit is asymmetric around its long
axis, and hence at any instance the β−γ interaction is always
distinct among the three β subunits. In fact, some biochemical
studies suggested that the α3β3 ring would not possess intrinsic
cooperativity on its own,155,163,164 supporting the contention
that the γ subunit would mediate the interplay among β subunits.
The view that the γ subunit is a “dictator” was reinforced bystudies showing that backward mechanical rotation of the γsubunit using external force reverses the chemical reaction
toward ATP synthesis,165,166 whereas forced forward rotation
accelerates ATP binding.167
However, a few biochemical studies suggested the intrinsic
cooperativity of α3β3 ring168,169 even though they were notwidely accepted. Furthermore, the “γ-dictator” model wasrecently challenged by the ﬁnding that even when the mostinteraction sites between β and γ are abolished, F1 retains its
active catalytic power and capability to rotate the shortened γ
subunit unidirectionally.170,171 To make it clear whether intrinsic
cooperativity exists within the α3β3 ring itself, the three β
subunits in the α3β3 ring alone (i.e., γ is absent) undergoing
conformational changes should be examined. However, theobservation of the rotational conformational changes by optical
microscopy requires the attachment of an optical probe to the γ
subunit.172 In one experiment, the polarization imaging of asingle ﬂuorophore ﬁrmly attached to a single β subunit in F1ATPase has successfully revealed the conformational change ofthe β subunit coupled with rotation of the γ subunit.173 However,
it is rather diﬃcult or impossible to simultaneously observe theconformations of all three β subunits in a single α3β3 ring using
optical techniques. Thus, HS-AFM imaging of the α 3β 3subcomplex in ATP was carried out to verify this issue that hadremained unanswered by other approaches.52
On bare mica surface, all molecules of the α3β3 subcomplex
rapidly diﬀused on the surface, which made it diﬃcult to image
them clearly. Therefore, the subcomplex had to be immobilized
somehow. To observe the C-terminal face of the α 3 β 3subcomplex, the subcomplex had to be placed on a substratesurface with the C-terminal side up. To force the molecules into
this orientation, a Lys7-tag was introduced at the N-terminus ofeach β subunit, and the mica surface was made amino-reactive
using APTES and glutaraldehyde.
Figure 13b shows a typical HS-AFM image of the α3β3
subcomplex without nucleotides. A ﬁltered image (3 × 3 pixel
median average ﬁlter) of the α3β3 showed a pseudo-6-foldsymmetric ring. Each subunit had an externally protruding
portion at the distal region of the ring as well as an upwardly
protruding portion at the inner top side of the ring. Yet three ofthese upward protrusions were higher at alternately arranged
three subunits than the other three (Figure 13c). The simulated
AFM image (Figure 13e, section A.3 in Appendix) of the Cterminal side of the subcomplex constructed from the crystal
structure of nucleotide-free α3β3 subcomplex shown in Figure
13d corresponded well to the observed image, indicating that the
N-terminal side was selectively attached to the mica support. By
comparison between the observed and simulated images, thethree subunits with higher protrusions were identiﬁed as the βsubunits. However, the selective attachment to the mica was not
perfect; about 30% of the molecules showed a smaller ring
without externally protruding portions, which corresponded to
the N-terminus face (Figure 13f−h).

chromophore and the photoexcited chromophore temporarily
altered the conformation of the head to dissociate it from actin,light pulses illuminating only the trailing head should make themolecule step forward, because the strain stored in the leadinghead would induce a forward step. Repetition of this directed
illumination only at the trailing head would have to be expected
to result in a long-distance forward movement of the molecule.
Instead of this method, the AFM cantilever tip, replacing theaction of ATP binding, could be used to detach the trailing headby injecting mechanical energy into the head. Such guided
manipulation of a single molecule by HS-AFM is a futurechallenge, which certainly will provide novel insights into thechemo-mechanical coupling in M5 as well as motor proteins ingeneral.
8.3. Rotary Catalysis of α3β3 Subcomplex of F1-ATPase

F1-ATPase is the soluble part of the FoF1-ATP synthase andworks as an ATP-driven rotary motor.151,152 The F1-ATPaseconsists of ﬁve kinds of subunits with a stoichiometry of
α3β3γ1δ1ε1. The α3β3γ subcomplex of F1-ATPase is a minimum
stable complex with the full ATPase and motor activities; the γ
subunit rotates in the stator α3β3 ring while ATP is hydrolyzed by
the ring.153−155 The concept of the “rotary catalysis” of F1 was
ﬁrst proposed by Boyer as a natural consequence of elaborate
biochemical studies on the “binding change mechanism” for the
ATP synthase.156 In the binding change mechanism, all three
nucleotide-binding subunits in FoF1-ATP synthase are always indiﬀerent states and alternately exchange the states during ATP
synthesis and hydrolysis. The physical rotation of the γ subunit
within the α3β3 ring was postulated for explaining the binding
change mechanism.156 Later, the ﬁrst crystal structure of bovine
mitochondrial F1-ATPase revealed by Walker and colleagues
showed that the α and β subunits are arranged alternately,
forming a ring, while the central γ subunit forms the central shaft
deeply penetrating into the ring’s central hole, and that thecatalytic sites of ATP synthesis and hydrolysis are located at the
α−β interfaces, mainly on the β subunits.157 Thus, thepostulation of a rotating γ subunit seemed to be a structurally
supported realistic concept. Biochemical158,159 and optical160
analyses also implied the rotation of the γ subunit, and, ﬁnally, theunidirectional and counterclockwise rotation was directly
observed by single-molecule optical imaging, in which a
ﬂuorophore-labeled actin ﬁlament attached to the γ subunit
was visualized to rotate around the center of the stator α3β3ring.108,161 To achieve this unidirectional, successive rotation, thethree catalytic β subunits must undergo cooperative andsequential conformational changes, coupled to ATP hydrolysis.
However, before the advent of HS-AFM, no experiments wereable to be conducted to examine how cooperative catalysis in the
F1-ATPase took place.
In Walker’s crystal structure mentioned above, three catalytic
sites are in diﬀerent nucleotide-bound states (Figure 13a): one
ATP-bound (αTP−βTP interface), one ADP-bound (αDP−βDP
interface), and one nucleotide-free (αE−βE interface). The
structure taken by both βTP and βDP, in which their C-terminal
domains swing toward γ, is designated as the closed
conformation, while the structure taken by βE, in which the Cterminal domain swings away from γ, is designated as the open
conformation. The rotation of the γ subunit has been considered
to be driven by the open/close bending motion of the β subunits
because the two conformational states appear to push or bepushed by the γ subunit in the crystal structure. Wang and Oster
proposed that interactions with the γ subunit would control the3138

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Figure 14. HS-AFM imaging of α3β3 subcomplex in ATP. (a) Clips of successively captured HS-AFM images of the C-terminal side of α3β3 subcomplex
in 2 μM ATP (imaging rate, 12.5 fps). The highest pixel in each image is indicated by the red circle. (b) Tight correlation between two types ofconformational changes in β subunits observed in ATP; diﬀerent heights of protrusion (“H”, high; “L”, low) and extended/retracted (“E”/”R”)
conformations of the distal region. (c) Time course of cumulated angle of the highest pixel. The inset shows a trajectory, superimposed on an AFM
image, of the highest pixels corresponding to the high protrusions of extended β subunits (412 frames). The center of rotation is deﬁned by the averaged
X- and Y-positions of the highest pixels, and the cumulated angles are calculated relative to the ﬁrst frame.

the opposite R-to-E transition usually occurred simultaneouslywith its counterclockwise neighbor β subunit. Consequently, theE state conformation propagated in the counterclockwise
direction, with a discrete angle of 120° (Figure 14c).
In the above statement, the diﬀerent conformations of the βsubunits were described only on the basis of their appearances,
without quantitative measure. For quantitative descriptions, a
two-dimensional correlation coeﬃcient can be used to evaluate
the image similarity or dissimilarity between a given image and areference image (section A.4 in Appendix). Figure 15a shows thehistograms of the correlation coeﬃcients calculated for the
successive images of each β subunit observed in 2 μM ATP using
an image of a β subunit in the E state as a reference. The
correlation coeﬃcients distributed around two distinct peaks.
The peaks at larger and smaller coeﬃcients correspond to the E
and R states, respectively. Typical time courses of theconformational states taken by individual β-subunits within an
α3β3 subcomplex observed in the presence of 2 μM ATP are
shown in Figure 15b. In all images observed in the presence of 2−
4 μM ATP (8746 images in total), the RRE state was dominant(82% of total). The REE state was also observed (14.5%). Other
states, RRR (3%) and EEE (0.5%), were rare. When “+1” wasgiven for a counterclockwise shift of the RRE state, while “−1”was given for a clockwise shift of the RRE state, the cumulated
number exhibited the rotary propagation of the RRE state

In the presence of a nonhydrolyzable ATP analog, AMPPNP,
the ring became triangular because two of the alternately
arranged three subunits showed inwardly retracted distal
portions. These two retracted subunits also showed lower
protrusions, and consequently the ring showed a single highprotrusion (Figure 13i). A simulated AFM image constructedfrom the crystal structure of the F1-ATPase obtained in ATP157
from which the γ subunit was removed showed an asymmetricring very similar to the observed image (Figure 13j and k). By
comparison between the observed and simulated images again,
the subunit with the high protrusion was identiﬁed to be an
empty β subunit, and the two retracted subunits with low
protrusions were identiﬁed to be AMPPNP-bound β subunits.
Under the nucleotide-free condition and in the presence of
AMPPNP, the observed structures were always stationary,
whereas in the presence of ATP, distinct dynamic changes in
the conformation of the three β subunits occurred (Figure 14a).
Each β subunit underwent a conformational transition between
the outwardly extended high state (E state) and the retracted low
state (R state). The outwardly extended and inwardly retracted
conformations were well correlated with the high and low
protrusion states, respectively (Figure 14b). The mostprominent features were that only a single β-subunit adapted
the E state, as statically observed in the presence of AMPPNP,
and that when the E-to-R transition occurred at one β subunit,
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Figure 15. Analysis of HS-AFM images of α3β3 subcomplex of F1-ATPase using correlation coeﬃcient. (a) Correlation coeﬃcient histograms calculated
for each β subunit (n = 220) observed in 2 μM ATP. An AFM image of an extended β subunit was used as a reference for the analysis of each β subunit.
(b) Time courses of correlation coeﬃcients. The white and gray backgrounds represent periods of extended high and retracted low states, respectively.
Solid lines show the mean correlation coeﬃcient for each period. (c) Time evolution of cumulated number of counterclockwise shifts of the RRE state.
“○” and “×” correspond to RRE and other states (REE, EEE and RRR), respectively. The increase in the cumulated number indicates that the extended
β subunit in RRE shifts counterclockwise. (d) Distributions of the number of successive shifts of the “RRE” state in clockwise (top) andcounterclockwise (bottom) directions at 2 μM ATP. The values for “mean” indicate the average number of successive shifts, and the values for “s.d.”indicate the standard deviations.

generation to rotate the γ subunit is elicited through the β−βinterplay alone. Because direct interactions between β subunits inthe α3β3 ring are considered to be infeasible from the crystal
structure, the β−β interplay is very likely to take place via theintervening α subunits. This inference could be corroborated by
the following observations, in addition to the observation
showing conformational changes of the α subunits that were lessvisible than those of the β subunits. During HS-AFM imaging, asingle subunit rarely dissociated from the α3β3 ring (Figure 17),
presumably due to imperfect covalent binding of the ring to themica surface and transient, overly strong tip−sample interaction.
When either a single β subunit or a single α subunit was lost fromthe α3β3 ring, the rotary propagation completely halted, althoughsome cooperative conformational transitions still occurred
between the remaining β subunits.
The results of HS-AFM observations on the α3β3 ring in ATP
deﬁnitely excluded the “γ-dictator” model170 that only theinteraction with the γ subunit would determine the conformational and catalytic states of β subunits.174,175 However, the rate
constant of ATP binding for the α3β3 subcomplex estimated fromthe ATP concentration-dependent dwell time of the E state was∼1 × 106 M−1 s−1, which is more than 10 times lower than thatfor a subcomplex containing the γ subunit, α3β3γ.176 Moreover,without γ, propagation of the conformational states occasionallyoccurred in the opposite direction. Thus, the γ subunit plays a(Figure 15c). The propagation was unidirectional, rotating
counterclockwise with an eﬃciency of 83% (in 2 μM ATP) over
the total shift events examined (n = 371) (Figure 15d).
The dwell times in the E and R states of each β subunit
decreased with increasing ATP concentration from 2 to 4 μM
(Figure 16a−c). This fact indicated that ATP binding is ratelimiting in this range of ATP concentrations, consistent with theMichaelis−Menten constant of 12 μM for ATP hydrolysis
determined biochemically. The histograms of dwell times of theE state followed single exponential functions (Figure 16a−c,
upper panels), whereas those of the R state were well ﬁtted withthe model of two consecutive ATP binding events at the othertwo β subunits triggering the transition of the β subunit from theR state to the E state (Figure 16a−c, lower panels). The rate
constants of the counterclockwise shift of the RRE state werecomparable with the initial rates of ATP hydrolysis measured
biochemically (Figure 16d). Therefore, the E-to-R transition
occurs when ATP is bound to an empty β, while the R-to-E
transition occurs when an ADP-bound β releases ADP. As such,the empty, ADP-bound, and ATP-bound β subunits are arranged
counterclockwise in this order, and hence the observed
conformational propagation demonstrated rotary catalysis by
the α3β3 subcomplex alone without the γ shaft.
The HS-AFM observations of the α3β3 subcomplex directly
indicated that the intrinsic cooperativity responsible for torque3140

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Figure 16. Kinetics analysis of conformational change of α3β3 subcomplex. Histograms of dwell times for the E (top) and R (bottom) states in various
concentrations of ATP (a, 2 μM, n = 266; b, 3 μM, n = 426; c, 4 μM, n = 374). Dwell time distributions for the E state ﬁt well with single exponential
decay functions (solid lines on top). The distributions for the R state ﬁt with functions representing consecutive reactions with two identical time
constants: A × t × exp(−t/τ) (solid lines on bottom). (d) ATP concentration dependence of the rate of conformational change of β subunit determinedby ﬁtting the dwell-time histograms of the E and R states, shown in (a)−(c), and the initial rate of ATP hydrolysis determined by biochemical assay atthree diﬀerent ATP concentrations.

8.4. General Remarks on Membrane Proteinsrole in establishing an oligomeric structure with the full ATPase
activity and reinforcing the cooperativity elicited by the β−βinterplay.
Because the γ subunit passively rotates, an object (notnecessarily made of polypeptides) that can be inserted into thecentral hole of the α3β3 would be able to rotate driven by therotational conformational changes of the β subunits. This
experiment is a future challenge. Moreover, it is also an intriguingchallenge to examine the chemo-mechanical coupling in F1ATPase using another approach. Because there is a strong
chemo-mechanical coupling in the β subunits (i.e., the chemical
states determine the respective conformational states), an
opposite direction of coupling (i.e., determination of thechemical states by the mechanical states) is likely to occur,
which is reminiscent of the fact that the forced clockwise rotationof the γ subunit by external force can reverse the chemical
reaction to ATP synthesis165,166 but diﬀerent from it. The β
subunits take the E (outwardly extended high) and R (inwardly
retracted low) conformational states corresponding to the
nucleotide-free and nucleotide-bound states, respectively. In a
buﬀer solution containing ADP (without ATP), the protrusion of
one β subunit in the E conformational state (i.e., in the
nucleotide-free state) can be pushed using a cantilever tip, which
may convert the conformation to that of the R state, triggering
ADP binding to the subunit as well as ADP dissociation from oneof the neighboring β subunits. As a result, ATP-less rotation ofthe γ subunit would occur, similar to the ATP-less walking of M5
on actin ﬁlaments, as predicted above.

Membrane proteins have been, and are still to some extent, theorphans of structural and molecular biology. Membrane proteinsmust, to adopt their native fold within the hydrophobic lipid
membrane core, expose hydrophobic amino acids, in stark
contrast to soluble proteins that bury hydrophobic amino acids
within the protein interior. On both membrane-protruding faces,
typically termed intracellular and extracellular surfaces, membrane proteins are hydrophilic, exposing polar or charged aminoacids. This general architecture is the basis of their amphiphilic
nature. It is this feature that rendered biochemical, functional,
and structural characterization of membrane proteins diﬃcult fora long time. It took 25 years from the ﬁrst soluble protein(myoglobin) structure solved177 to the ﬁrst membrane protein(reaction center) atomic structure determination.178 At the date
this Review is written, scientists dispose of 1264 membraneprotein structures of 406 unique proteins,179 while a total of 91
022 structures are disposed of the PDB.180 Hence, the structural
information on membrane proteins of ∼1% of all structures isstill in unfavorable relation to the ∼25% of all genes coding formembrane proteins.181 The majority of the membrane protein
structures have been solved by X-ray crystallography, a minor
fraction by EM crystallography (e.g., ref 182) and NMR
spectroscopy (e.g., ref 183). Anyway, regarding the diﬃculties
in analyzing membrane protein structures, alternative techniquesthat provide structural information, even at medium resolution,
were strongly welcomed. Such data can be provided by single
particle EM (e.g., ref 184) and AFM (e.g., ref 185). Beyond
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Figure 17. Cessation of rotary conformational change propagation by loss of a single subunit. (a,d) AFM images before and after loss of (a) β1 or (d) α1
within α3β3 subcomplex in 2 μM ATP. Imaging rate, 12.5 fps. (b,e) Time courses of correlation coeﬃcients for the three β subunits designated in (a) and(d), respectively. Solid horizontal lines show the mean correlation coeﬃcients for the periods of E and R states. The states are judged by examining
whether the correlation coeﬃcients are above or below a threshold (0.978). β1 was lost at 16.2 s (b) and α1 was lost at 9.4 s (e) as indicated by the verticalbroken lines. (c,f) Cumulated angles of open β subunit positions measured using the highest pixel position in each frame. (c) and (f) correspond to (b)and (e), respectively.

that hold pigments in place.15,186 The majority of membraneproteins occur in at least two relatively stable conformations,activated and inactivated for receptors,187 inside-open andoutside-open for transporters,188 and of course the short-lived
intermediates between these conformations. Thinking theconcept of protein conformational changes to the end, onemust conclude that many proteins are actually in a permanentchange of conformations, such as the mitochondrial ATP
synthase. Indeed, the ATP synthase is subject to the proton
gradient across the inner mitochondrial membrane during its
entire “life span” and is therefore probably in continuous rotation(if it is not complexed with its inhibitory subunit), highlighting
the importance of assessing protein conformational dynamics
beyond static structure analysis. It is very challenging to study
these conformational changes. Techniques that do not allow insitu structure assessment typically need to block the proteins in

structural analysis, the analysis of membrane protein dynamics isof key importance for the understanding of membranecomplexes. Several levels of dynamics of membrane proteinscan be distinguished, such as conformational changes, diﬀusion,and interaction. We will treat these aspects individually below.
However, it must readily be highlighted that the concomitant
analysis of structure and dynamics necessitates the use of atechnique with high spatial and temporal resolution. This has notbeen possible so far, and the HS-AFM allowing imaging of single
molecules at high spatiotemporal resolution is opening novel
ways to assess superstructure/structure/function relationships ofmembrane proteins.
In contrast to what a single crystallographic protein structure
might suggest, only very few proteins accommodate a single ﬁxed
conformation, such as proteins that serve rather as scaﬀolds than
as functional entities, for example, light-harvesting complexes
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Figure 18. HS-AFM imaging of D96N bR mutant. (a,b) HS-AFM images of the cytoplasmic side (a) and the extracellular side (b) of D96N bR mutant(imaging rate, 1 fps) under dark (left panels) and green light illumination (right panels) (λ = 532 nm, 0.5 μW). bR trimers are highlighted by the white
triangles. The insets in (a) display respective averaged images of trimers. The light-blue circles in (a) indicate a trefoil, and the green bars in (a) and (b)indicate green light illumination. (c) Decay after ﬂash-illumination of the activated bR molecules observed at diﬀerent pH values (7, 8, and 9). The
exponential decay constants (τ) at pH 7, 8, and 9 are 6.7, 25, and 48 s, respectively. The inset shows the absorbance change at 410 nm (that is, decay of
the M-intermediate) after ﬂash illumination of D96N measured at various values of pH (7, 8, or 9). (d) Traces (red, blue, and purple marks) of thecenters of mass of bR molecules under dark−illumination cycles are superimposed on the image of D96N in the dark. A bR trimer is encircled with thewhite triangle, and the positions of the α-helical ends facing the cytoplasmic surface (labeled with A−G) are derived from an atomic model of theunphotolyzed state.

bundles.192 Over the following about 25 years, the entire
photocycle of bR was characterized functionally in detail,190 andhigh-resolution X-ray structures were determined.193
A bR monomer comprises seven transmembrane α-helices(named A−G) enclosing the chromophore retinal.194,195 bR
assembles into trimers, which are packed into 2D hexagonallattices.191 In the photocycle, photoisomerization from all-transto 13-cis conformation of the retinal takes place with subsequent
primary proton transfer from the Schiﬀ base to Asp85, which
triggers a cascade of changes in the bR structure. A series ofintermediates designated J, K, L, M, N, and O are deﬁned byspectroscopy, and the M (M410 with the light-absorption peak at
410 nm) is the only intermediate containing a deprotonatedSchiﬀ base.190 The M and N states have long lifetimes, whereas
the other states have much shorter lifetimes. The light-induced
conformational changes in bR have been investigated by various
methods.189,196 In particular, crystallographic structures in thefrozen activated state of the wild type (WT) and bR mutants havebeen solved at the atomic resolution.196−199 The common viewthat has been reached through these studies on the bR structure

both conformations. HS-AFM, although limited to surface
contouring and to a ∼30 ms dynamic range, opens a novel
avenue for the direct visualization of membrane protein
conformational dynamics. In this feat, it combines two additional
advantages; ﬁrst single molecules can be analyzed and hence theprocess action of individual proteins can be studied, and second
studying proteins in a close-to-native environment brushes aside
any interpretational doubt that could emerge, for example, forproteins subject to intermolecular contacts in crystals.
8.5. Structural Changes of Membrane Proteins

8.5.1. Bacteriorhodopsin Responding to Light. A
paradigm membrane protein system is the bacterial light-driven
proton pump bR in the purple membrane of Halobacterium (H.)
salinarum.189,190 It was identiﬁed and functionally characterized
in the early 1970s,191 and is nowadays perhaps the best
characterized membrane protein. Its natural tendency to form
highly ordered 2D arrays has been advantageous for structural
studies, kick-starting EM crystallography.192 It was in thesestudies that the ﬁrst experimental proof was acquired that
membrane proteins actually cross the membrane as α-helix
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Figure 19. Cooperative eﬀects on the decay kinetics of D96N. (a) The top ﬁgure shows the decay of activated bR monomers (as depicted by the redcircle), while the other monomers (the gray circles) within the respective trefoils are not in the activated state. The exponential decay constant is 7.3 ±0.58 s. On the other hand, when one or two nearest neighbor molecules (as depicted by green circles in the middle ﬁgure) in a trefoil are alreadyactivated, the bR monomer activated latest (as depicted by the blue circles in the middle ﬁgure) decays faster. The exponential time constant is shortened
to 2.0 ± 0.16 s. In contrast, the early activated molecules within a trefoil (as depicted by green circles in the bottom ﬁgure) show a nonexponential broad
distribution of decay time with an average of 13.3 s. (b) Decay rate of activated D96N under diﬀerent light intensities. The upper graph shows the decayof activated molecules under weak illumination. The decay constant is 5.4 ± 0.34 s. The lower graph shows the decay of activated molecules underrelatively strong illumination. The decay constant is 6.1 ± 0.37 s.

resolution. It was natural that this intriguing topic was studied
soon after the advent of HS-AFM, although it appeared as achallenging task to study the bR photocycle by HS-AFM. The
ﬁndings obtained thereby surprisingly provided novel unexpected insights into the bR photocycle, so far unappreciatedby other technical and structural approaches.44 The observation
successfully revealed the conformational changes much larger
than 0.3 nm as well as the cooperative interplay between excitedbR molecules in the 2D crystal.44,49 Moreover, the trimer−trimer
interaction energy was quantitatively estimated by theobservation of dynamic interaction between diﬀusing trimers
and the crystalline lattice edge in the membrane.35 Finally, HSAFM imaging of various bR mutations was performed to identify
the amino acids responsible for the interaction between trimers
leading to lattice assembly, revealing the purposes for bR
assembly into regular lattices.62 All of these HS-AFM studies aredescribed as topics of this section (structural changes ofmembrane proteins), except for the study of the trimer−trimer
interaction energy (described in section 8.6.1).
The photocycle of wild type bR takes place within about 10 ms
at neutral pH; hence bR in the photoexcited state is too shortlived to be observed even using HS-AFM. Therefore, the purple

during the photocycle is that the proton channel on thecytoplasmic surface is opened by the tilting of helix F away fromthe protein center.197 This alteration is followed by rearrangement of the interhelix E−F loop, thus resulting in relatively global
conformational changes in the M and N intermediates.200−202
Despite the consensus on the qualitative description ofconformational changes, no agreement was reached for thequantitative aspect (reported changes were small, ranging from
0.1 to 0.3 nm) because of the lack of techniques enabling direct
visualization of the conformational change of bR underphysiological condition.
The 2D character of purple membrane promoted the
beginnings of high-resolution AFM imaging and the assessmentof image resolution and quality.203 Following the pioneering
AFM study of bR in the purple membrane by Engel’s group,203
bR has been repeatedly imaged to demonstrate the increasing
high-resolution power of AFM. In fact, unprecedented highresolution images capable of distinguishing the individualinterhelix loops of bR were obtained again by Engel’s
group.76,204 Nevertheless, until recently the physiologically
relevant structural changes in bR (i.e., response to light) havenever been studied using AFM because of its lack in temporal
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visualized here is considered to represent the displacement of the
E−F interhelical loop. This displacement was clearly seen in the
averaged image of the excited state bR (compare the averaged
images under dark and illumination, which are inset in Figure
18a).
As a consequence of the outward displacement of the E−F
loop in each bR monomer, the three nearest-neighbor bR
monomers, each belonging to a diﬀerent adjacent trimer, always
transiently assembled. The triad of the nearest-neighbormonomers was designated as “trefoil” to distinguish it from theoriginal trimer (trefoils under dark and illumination conditions
are indicated by light-blue circles in the left and right panels in
Figure 18a). Interestingly, this transient assembly in trefoils
altered the decay kinetics of the activated state. The decay
kinetics of the activated bR monomers in three diﬀerent
situations are shown in Figure 19a. Under weak illumination,
only one monomer in each trefoil was activated in most cases.
When only one monomer was activated in a trefoil, it decayedwith a time constant of 7.3 ± 0.58 s (see top panel in Figure 19a).
In contrast, under stronger illumination, two or three monomers
within a trefoil tended to be activated together. Interestingly, thedecay of each activated monomer within a trefoil was altereddepending on the order of its activation. The monomer that was
activated latest among the activated monomers in the trefoil
decayed with a shorter time constant of 2.0 ± 0.16 s (see middle
panel in Figure 19a). On the other hand, the decay kinetics of theearly activated monomers did not follow a single exponential, andthe averaged decay time lengthened to about 13 s (see bottom
panel in Figure 19a). This observation indicated that the early
activated monomers do not return to the ground state as long as
the adjacent monomers within the trefoil are in the active state.
Thus, the transient assembly of excited bR molecules within atrefoil elicits both positive and negative cooperative eﬀects in the
decay kinetics as the initial bR recovers. Some cooperativity
regarding the photoexcitation of bR was previously proposed andconsidered to occur within a trimer.207,208 However, the HSAFM observation revealed that the cooperative eﬀects are caused
by bR−bR interactions within a trefoil, not within a trimer,
shading a completely novel view onto the assembly of bR trimers
in the hexagonal lattice.
When all of the activated molecules were counted regardless of
their activation order and of the number of activated monomers
in a trefoil, the lifetimes of the activated state were well ﬁt tosingle exponential functions under both weak and strong lightillumination conditions, and their decay constants were nearly
identical, irrespective of the intensity of illuminated light (Figure19b). This was explained by the coexistence of the positive andnegative cooperativity eﬀects under strong light intensity. The
two cooperative eﬀects are nearly canceled out with each otheron averaging, resulting in an approximately constant decay rate
independently of the light intensity. Under a conceivableassumption that the conformational change turnover of anactivated bR monomer is tightly coupled with proton transfer by
the bR monomer, the average rate of proton-pumping byactivated bR would be nearly conserved regardless of lightintensity. As a result of the cancelation eﬀect, the cooperativeeﬀects are hardly detected by ensemble averaging approaches. As
such, this discovery of cooperative eﬀects in a trefoil best
exempliﬁes the importance of closely looking at individual
molecules by HS-AFM. Even for extensively studied biomolecular systems such as bR, undisclosed important mechanisms
still exist to be discovered, some of which can be newly disclosed

membrane containing D96N bR mutant was used. This mutant is
known to have a longer photocycle (∼10 s) but retains an abilityof proton pumping.205 D96N bR lacks the proton donor residue(Asp96) for the Schiﬀ base, rendering the M-intermediate very
long and hence the photocycle so slow; the proton forreprotonation of the Schiﬀ base during the decay of the M
state has to be supplied from the aqueous milieu. A state exactly
the same as the N state of the wild-type bR, which follows the M
state and contains a protonated Schiﬀ base, has been considered
not to exist in the excited D96N bR. To avoid photoactivation ofbR with the laser light used in the OBD detection, an infraredlaser (980 nm) was used for the OBD detector. The HS-AFM
observation was performed in a solution of pH 7, 8, 9, or 10 tohave diﬀerent photocycle times.
Figure 18a shows two images of D96N bR clipped fromsuccessive images at the cytoplasmic surface captured at an
imaging rate of 1 fps, under repeated dark−illumination cycles.
Individual bR molecules within individual trimers were clearly
resolved (left panel in Figure 18a; captured under dark), as
observed on the cytoplasmic surface with conventional slow
contact-mode AFM.206 During illumination with green light(532 nm, 0.5 μW; green bars in Figure 18a), the bR molecules
drastically changed their structure (right panel in Figure 18a).
After the light was switched oﬀ, the excited bR molecules
returned back to the unphotolyzed-state conformation in a few
seconds. The conformational changes were highly reproducible
in repeated dark−illumination cycles. On the other hand,discernible conformational changes were never observed at theextracellular surface (Figure 18b). To ensure that the conformational changes observed here were not artifacts such as a tipforce-induced conformational change,204 the decays of theexcited state after ﬂash illumination at various pH values wereanalyzed (Figure 18c). It is well-known that the lifetime of the Mintermediate of D96N is prolonged at alkaline pH205 as observedwith UV−visible spectroscopy (the inset in Figure 18c). The
decay detected by HS-AFM also showed strong pH dependence,
similar to that measured by UV−visible spectroscopy. Thisagreement indicated that the conformational change observed by
HS-AFM is surely an intrinsic nature related to the bR
photocycle. Moreover, the fact that the conformational change
observed with green light illumination was not observed when
red or blue light was used for illumination reinforces thisconclusion. Note that the conformational change rarely occurredunder blue light illumination because the ground state having anabsorption peak at 570 nm absorbs blue light slightly.
Figure 18d shows the trajectories that the centers of mass of bR
molecules took during the dark−illumination cycles (red, blue,
and purple marks), which are superimposed on an AFM image
captured under the unphotolyzed state. Here, the “center ofmass” for each bR monomer was calculated from thecorresponding surface area and height distribution in the HSAFM topograph. The trajectories were highly centrosymmetricand oriented outward from the trimer centers under the lightillumination condition. All activated bR monomers exhibited adisplacement of 0.7−0.8 nm on average, which is much larger
than those estimated before from electron microscopy and X-ray
crystallography.198,199 In comparison to the atomic model of the
α-helical cytoplasmic ends in the unphotolyzed state (A−G in
Figure 18d), the protruding areas around helices E and F shifted
outward from the trimer center under illumination. In addition, aprevious AFM imaging study of bR has assigned the prominent
protrusion on the cytoplasmic face in AFM images of bR to thecytoplasmic E−F loop.204,206 Therefore, the bR movement3145

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Figure 20. Eﬀect of alternate two-color illumination on the conformation of D96N bR observed by HS-AFM. (a) Displacement of centers of mass forthree D96N bR molecules within a trimer measured as a function of time at pH 8 on the cytoplasmic side. The green bars show the periods of green lightillumination. (b) Displacement of centers of mass for three D96N bR molecules within a trimer measured as a function of time at pH 8 on thecytoplasmic side. The green and blue bars show the periods of illumination of green and blue light, respectively. The average displacement induced bygreen light is 0.80 ± 0.13 (mean ± s.d.) nm. (c) Decay of the activated state of D96N bR at pH 8 detected by HS-AFM. Green and blue regions indicate
application of green and blue light, respectively. Illumination periods of both green and blue light are 3 s.

Figure 21. HS-AFM images of cytoplasmic surfaces of purple membranes containing bR mutants (a, W10I; b, Y131I; c, W12I; d, Y135I; e, W12F).
Imaging rates: (a, b, and e) 1 fps, (c and d) 33 fps.

by HS-AFM imaging, thanks to the directness of the acquired
data of single-molecule imaging.
It is known that the all-trans to 13-cis isomerization of retinal
can be reversed by a second illumination with a diﬀerent
wavelength.209 After the retinal is isomerized to the 13-cis form
by green illumination, blue light (λ = 408 nm) illuminationinduces a back reaction of the retinal to the ground state (all-transform). However, it has not yet been evident whether the protein
structure would tightly be coupled to the isomerization of theretinal molecule because the spectroscopically detected reversal
does not necessarily indicate the reversal of the entire protein
conformation. This issue was addressed by the direct visualization of D96N bR under alternate illumination of green andblue light.49 Figure 20a shows the displacement of the centers of
mass, as a function of time, for three diﬀerent bR monomers at
pH 8. Once bR molecules were activated by green lightillumination, the activated state was mostly maintained evenafter the light was turned oﬀ (Figure 20a). This was because thelifetime of the activated state of D96N bR at pH 8 is much longer
than the duration used (10 s) for green light illumination. In thecase of alternative illumination, bR molecules underwent
alternate conformational changes between the excited andground states synchronized with the switching between green
and blue light illumination (Figure 20b). The blue light-induced
conformational change back to the ground state is caused by thephotoback-reaction, not by the normal turnover of thephotocycle. Because bR in the ground state also slightly absorbs
blue light, deactivated bR was sometimes reactivated under theblue light (blue arrows in Figure 20b). The number of activatedmolecules rapidly decreased nearly to zero after the brief blue
light illumination (Figure 20c). These observations clearly
showed that the bR protein structure is completely governed by
the conformational state of the retinal molecule in its core.
8.5.2. Role of Trimer−Trimer Interaction in bR
Function. It has long been a question: For what purpose is bR
assembled into a crystal lattice in the native state? As described in
section 8.5.1, the conformational change of bR during thephotocycle elicits cooperative eﬀects due to interactions between
excited bR molecules within trefoils. This suggests that the lattice
formation would have a role in the physiological function of bR.
Thus far, only a few studies have been presented for the
elucidation of the origin of the trimer−trimer interaction leading
to the crystal lattice and for the estimation of the energy involvedin the interaction.210−213
Amino acid residues in bR responsible for trimer−trimer
association leading to lattice assembly were recently identiﬁed by
the combination of HS-AFM imaging and directed mutagenesis.
A previous neutron diﬀraction study has suggested that some of
the aromatic residues (W10, W12, Y131, and F135) locatedvertically no farther than 5 Å into the extracellular side of themembrane and laterally at the outer rim of the trimers are
involved in lattice formation.211 To more speciﬁcally identify
essential amino-acid residues for the lattice assembly, ﬁve bRmutants (W10I, Y131I, W12I, F135I, and W12F) were preparedand imaged by HS-AFM.62 Figure 21 shows images of these bR
mutants. As can be seen in these AFM images, the W10I and
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Figure 22. HS-AFM imaging of isolated D96N/W12I bR trimers. (a) HS-AFM image under dark of D96N/W12I bR trimers anchored on chemically
modiﬁed mica substrate (imaging rate: 1 fps). (b,c) Clips of successively captured HS-AFM images of a D96N/W12I bR trimer (b) under green lightillumination (indicated by green rectangles) and (c) after the light was turned oﬀ (imaging rate: 2 fps). The insets are schematic illustrations of theobserved bR conformations. The gray and green circles indicate the conformations of bR monomers in the ground and excited states, respectively.

above strongly suggested that the cooperativity measured on the
single-molecule level should be lost when bR was unable to forma lattice. Figure 23a shows the population of activated D96N and
D96N/W12I bR molecules as a function of time after the start ofcontinuous illumination. The rise time constant of D96N/W12I
bR was estimated to be τ = 1.9 ± 0.06 s, which was 4-fold longer
than that of D96N bR (τ = 0.49 ± 0.01 s) under the same lightintensity. On the other hand, the decay kinetics after light-oﬀ wasalmost identical for both mutants (τ ≈ 10 s), on average overmany bR molecules (Figure 23b). As expected, the decay kineticsof three bR molecules within an isolated D96N/W12I bR trimer
was conﬁrmed to be independent of their activation order, likethe case of three bR molecules within a D96N bR trimer, but
unlike the case of three bR molecules within a D96N bR trefoil. A
possible eﬀect of anchoring the molecules onto the chemically
modiﬁed mica on the activation and decay kinetics was excluded
because D96N bR immobilized onto the chemically modiﬁed
mica showed kinetics with time constants similar to those for
D96N bR simply physisorbed on a bare mica surface (insets in
Figure 23a and b). Therefore, the chemical anchoring has no
signiﬁcant eﬀect on the photocycle, and, consequently, the risedelay observed with D96N/W12I bR is intrinsically caused bylattice disruption. Because the rise rate linearly increased withincreasing illumination intensity (Figure 23c), the retardation
was likely to be attributable to a lower eﬃciency of theconformational change or to a smaller optical absorption crosssection of the D96N/W12I mutant. The latter was, however,ruled out because the diﬀerence spectrum between the groundand 100% excited states of D96N/W12I bR measured at low
temperature (210 K) was identical to that of D96N bR.

Y131I mutants formed a lattice (Figure 21a and b). On the other
hand, the lattice was not formed in the cases of W12I and F135I
mutants (Figure 21c and d), but as indicated by the dotted
triangles in the images trimers were visualized as isolated entities
diﬀusing fast within the membrane. These results indicated thatboth W12 and F135 residues play an essential role in assemblingbR trimers. The W12F mutant also formed a crystal lattice, as
shown in Figure 21e, indicating that aromatic residues at
positions 12 and 135 associate with each other (homotopically or
heterotopically) possibly through a hydrophobic stacking
interaction mediated by the sugar moieties of glycolipids
contained in the membrane.211,214
Because the bR trimer isolation method was established by theabove study, the question, “for what purpose bR is assembled
into a crystal lattice in the native state?”, was addressed by HSAFM imaging.62 Double mutations, D96N and W12I, wereintroduced to bR to obtain isolated trimers with a slow
photocycle. Because D96N/W12I bR trimers diﬀused fast in
the membrane, they were immobilized on a mica surface treatedwith APTES and then activated with glutaraldehyde (Figure22a). Upon green light illumination, the D96N/W12I bR mutant
exhibited conformational changes similar to that observed with
the D96N bR mutant (i.e., outward movement of the E−F loops;
Figure 22b; imaging rate, 2 fps). After light was turned oﬀ, theexcited-state conformation returned back to the ground-state
conformation within a few seconds (Figure 22c).
The kinetics of photoactivation during continuous green
illumination, as well as the decay kinetics after light-oﬀ, wasanalyzed to see a possible eﬀect of lattice disruption on thephotoreaction cycle, especially because the results presented
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Figure 23. Eﬀects of lattice disassembly on the photoreaction kinetics of bR. (a) Time courses of accumulation of activated bR molecules detected by
HS-AFM after the start of continuous light illumination (0.2 μW). Red plots, unanchored D96N bR; blue plots, anchored D96N/W12I bR. The
exponential time constants (τ) of D96N and D96NW12I are 0.49 ± 0.01 and 1.9 ± 0.06 s, respectively. The inset shows the time course of theaccumulation of activated bR in an anchored D96N bR sample. The exponential time constant is 0.58 ± 0.01 s. (b) HS-AFM-detected decay of activatedstates of unanchored D96N bR (red plots) and anchored D96N/W12I bR in full active trimers (blue plots) after light (0.2 μW) was switched oﬀ. The
exponential time constants of D96N and D96NW12I are 10.4 ± 0.10 s and τ = 9.6 ± 0.04 s, respectively. The inset shows the decay of activatedmolecules for anchored D96N after light was turned oﬀ. The exponential time constant was 9.3 ± 0.04 s. (c) Rate constants of rise phase as a function oflight intensity for D96N (red plots) and D96NW12I (blue plots) measured by HS-AFM. The inset shows the rate constants of the rise phase as afunction of light intensity for D96N measured by UV−visible spectroscopy. (d) Models of photocycles in D96N bR and D96N/W12I bR. Upper
illustrations indicate the lattice-assembled D96N bR (left) and disassembled D96N/W12I bR (right). The bottom schematics show the modeled
photocycles in D96N bR (left) and D96N/W12I bR (right). In the photocycle of D96N bR, the presence of a direct recovery pathway to the groundstate from MX with no E−F loop displacement is minor as indicated by the dashed arrow. The rate constants, k1, k2, k3, and k4, for transitions between
diﬀerent states are assigned as shown in the right bottom schematic. Numerical analysis of HS-AFM detected conformational change kinetics based on
this model gives k1 = 2.04 s−1, k2 = 100.0 s−1, k3 = 0.1 s−1, and k4 = 0 s−1 (for D96N bR) or 370.4 s−1 (for D96N/W12I bR).

nonproductive pathway, no signiﬁcant structural change occurs
in bR and no proton is pumped out; however, the kinetics of thispathway is much faster than that of the original pathway. All of
the results obtained in this study were quantitatively well
explained by mathematical simulations based on this model.62
Thus, trimer−trimer association plays an essential role inproviding bound retinal with an appropriate environment tomaintain its full photoreactivity and in maintaining the natural
photoreaction pathway. The bR lattice in purple membrane is
not formed nonspeciﬁcally for the improvement of packing
density. It is formed because the photocycle is more eﬃcient inthis assembly. These ﬁndings illustrate the fantastic power of HSAFM’s direct acquisition of dynamic data at the single-molecule
level and how this power could contribute to the discovery ofimportant novel aspects in one of the best-studied membraneprotein system.
8.5.3. Up−Down Motion of Ca2+-ATPase. Ca2+ is aregulator of muscle contraction. Skeletal muscle contracts whenCa2+ is released from the sarcoplasmic reticulum (SR) into thecytoplasm. During this activation, Ca2+ binds to troponinassociated with actin ﬁlaments and tropomyosin, and thereby
the inhibition of actomyosin ATPase activity by troponintropomyosin is removed.215 Ca2+ is pumped back into the lumenof the SR by the action of Ca2+-ATPase (an energy driven Ca2+pump) existing in the SR membrane, resulting in muscle
relaxation.216−218 Two Ca2+ ions are transported per ATPase
cycle.218 The Ca2+-ATPase, sarcoplasmic reticulum Ca2+-ATPase
1a (SERCA), is a member of the P-type ATPase superfamily,
which forms a phoshoprotein intermediate and undergoes large

Therefore, the reduction of the rate at which the excited state
conformation is accumulated during continuous illumination
should be attributed to a reduced eﬃciency of adopting theactivated conformation in D96N/W12I bR. This suggested that a
nonproductive photoreaction pathway would additionally occur
in disassembled D96N/W12I bR, through which the initial
photoactivated state (this unspeciﬁed state is designated MX)
would be de-excited to the ground state G without passing
through the M state (see Figure 23d). In addition to HS-AFM
observations, spectroscopic and proton pumping assay investigations were carried out for both D96N and D96N/W12I
samples to ﬁnd the origin of the reduced rate of M state
accumulation, but details of these studies are omitted here.
The results obtained from HS-AFM imaging, optical spectroscopy, and proton pumping assay are summarized as follows:(i) the M state with an outwardly displaced E−F loop exists in thephotocycle of both D96N bR and D96N/W12I bR; (ii) after thestart of continuous illumination, the M state accumulation ofdisassembled D96N/W12I bR occurs with a rate ∼4-times lower
than that of assembled D96N bR; (iii) the accumulation rate ofboth samples linearly increases with increasing light intensity;(iv) the optical absorption cross section is very similar between
D96N bR and D96N/W12I bR; and (v) the introduction of
W12I mutation into WT bR does not appear to aﬀect the proton
pumping activity of bR within experimental accuracy. Taking
these results into consideration, a novel model of the photo
reaction was proposed (Figure 23d); the lattice disruption causes
the photoreaction pathway to branch into two (the original
productive pathway and a new nonproductive pathway). In the3148

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while the T → S transition is expected to take place depending on
the Ca2+ concentration. To clear these issues experimentally,
Yokokawa and Takeyasu observed the dynamic height changes of
SERCA in real time, using HS-AFM.50
Puriﬁed SR vesicles containing SERCA were brieﬂy treatedwith a detergent, deocycholate (DOC), to break the vesicles. The
DOC-treated SR sample was adsorbed on a bare mica surface.
The resulting membrane fragment had a thickness of 5.1 ± 0.6
nm on mica surface. The SR membrane contained well-isolated
SERCA molecules with a density less than a few molecules per
μm2. The molecules in the membrane were essentially immobile(very slowly diﬀusing) with an averaged 2D diﬀusion coeﬃcientof 0.4 ± 0.2 nm2/s, although the reason for this immobility was
unspeciﬁed. The immobility allowed the repeated heightmeasurements of a single molecule for relatively long time
periods with single-line scanning26,30 and imaging by HS-AFM.
Figure 25a shows successive AFM images of a SERCA
molecule in the membrane captured at an imaging rate of 1 fps inthe presence of both 10 nM ATP and 100 μM free Ca2+. The
cross sections along the X- and Y-axes vaguely showed height andshape changes between the T and S states of the SERCA
molecule, likely corresponding to compact and Y-shaped
conformations, respectively. The single line scanning, in whichan AFM tip was repeatedly scanned along the X-axis only, now
clearly displayed rapid up-and-down motion of SERCA as a spikewave with an average lifetime of 0.15 ms (Figure 25b, arrowheadin the lower panel). As expected, when the ATP concentration
was increased to 1 μM, the frequency of spike wave appearance
was signiﬁcantly increased (Figure 25c), whereas the lifetime ofindividual spike waves was independent of the ATP concentration. The short-lived SERCA in the T state was 2.3 ± 0.4 nm
taller than that in the S state (n = 65). This value of heightincrease is consistent with the height diﬀerence between the E1·
2Ca2+ form and the other more compacted forms of SERCA(E1ATP, E1P, E2P, and E2) in the X-ray crystallographic
structures.224 To conﬁrm this correspondence, the height of
SERCA from the membrane surface was analyzed to produce
histograms (Figure 26). The average height was 5.4 ± 0.8 nm in
the presence of 100 μM free Ca2+ without ATP (Figure 26a),which is in good agreement with the height of the cytoplasmic
domain estimated from the X-ray crystallography data of the E1·
2Ca2+ form (S state). At a lower free Ca2+ concentration (10nM), the height was similar to that measured in 100 μM free Ca2+
without ATP (Figure 26b). In contrast, in the presence of 10 μM
thapsigargin (TG), which is known to retain SERCA in a form
analogous to E2,226 the histogram showed two peaks around 5.4± 0.7 and 7.2 ± 1.0 nm (Figure 26c). These results strongly
supported the inference that the height change observed in thepresence of ATP and Ca2+ is caused by the conformationalchange between E1·2Ca2+ and the other compact forms of
SERCA during the ATPase reaction cycle.
The height change of SERCA molecules was measured in thepresence of various ATP concentrations and two diﬀerent free
Ca2+ concentrations (Figure 27a). In lower Ca2+ concentration(10 nM), spike waves (quick up-and-down changes) were only
rarely observed; instead, the lifetime of the elevated conformation became remarkably longer (>200 ms) provided that aconcentration of ATP > 10 nM was present. The long lifetime ofthe elevated conformation at low Ca2+ concentration could haveresulted from lowered ATPase activity in low Ca2+ concentrations.227 On the other hand, in higher Ca2+ concentration(100 μM), the number of spike waves increased with increasing
ATP concentration. Therefore, as expected, the T → S transition

conformational changes during the course of ATP hydrolysis.217
SERCA has been well characterized structurally and functionally.219,220 The reaction mechanism of P-type ATPase iscommonly interpreted according to the Albers-Post
scheme.221,222 In this scheme, the ATPase has two distinctgroups of states during the catalytic cycle, which are traditionallydesignated as E1 and E2 (Figure 24a).223 In the E1 state, theFigure 24. ATPase reaction and structure of sarcoplasmic reticulum
Ca2+-ATPase 1a (SERCA). (a) Simpliﬁed Albers−Post scheme forCa2+-ATPase reaction. (b) 3D structures of E1Ca2+ and E2(TG) formsof SERCA in lipid bilayer: Reprinted with permission from ref 225.
Copyright 2003 John Wiley & Sons, Inc.

ATPase has a high aﬃnity for Ca2+, whereas in the E2 state, it has
a low aﬃnity for Ca2+. The ion-binding sites, which are locatedwithin the transmembrane domain, are open toward thecytoplasm in the E1 state and toward the lumen of the SR (orextracellular side) in the E2 state. Crystal structures of SERCA
have been determined for various physiological states.224,225 The
basic structure of SERCA comprises a large cytoplasmic
headpiece, a transmembrane domain made of 10 (M1-M10) αhelices, and short lumenal loops (inside the sarcoplasmic
reticulum). Four of the transmembrane helices, M2-M5, havelong cytoplasmic extensions. The cytoplasmic headpiece consistsof three domains, which are referred to as actuator (A),nucleotide binding (N), and phosphorylation (P) domains. A
large global conformational change takes place during thereaction cycle: the three cytoplasmic domains (A, N, and P
domains) are separated in the E1·2Ca2+ state into a Y-shaped
conformation (left schematic in Figure 24b) but tightly packed in
a compact conformation in the other states (right schematic in
Figure 24b).
From this global structural change, the height of the E1·2Ca2+intermediate is smaller (short (S) state) than those of the otherintermediates (tall (T) states). According to the simpliﬁed
Albers−Post scheme (Figure 24a), the S → T transition is
expected to take place depending on the ATP concentration,
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Figure 25. HS-AFM imaging of SERCA dynamics in the presence of nucleotide and Ca2+. (a) Successive HS-AFM images in the presence of 10 nM ATP
and 100 μM free Ca2+ captured at a frame rate of 1 fps. The cross sections along the X- and Y-axes are shown in the corresponding lower panels. The
broken line indicates a height of 5.5 nm from the membrane surface. (b,c) Single line scan (2D observation) HS-AFM images of SERCA in the presenceof 100 μm free Ca2+ and in the presence of (b) 10 nM and (c) 1 μM ATP, respectively. The lower panels show the X-axis cross sections positioned at theline indicated by the arrow beside the 2D HS-AFM images. Reprinted with permission from ref 50. Copyright 2011 John Wiley & Sons, Inc.

occurred depending on Ca2+ concentration, meaning that thistransition takes place at the Ca2+-binding step (E1 → E1Ca2+transition), and that the E1 state, whose conformation has not yet
been resolved, also has an elevated conformation.
Under near physiological conditions (1.0 mM ATP and 100
μM free Ca2+), SERCA molecules appeared to maintain theelevated conformation for a long time without T → S transition(Figure 27b). Spikes were not observed even with a much faster
X-line scanning rate up to 1 kHz, which corresponded to a time
resolution of 50 μs (i.e., the cantilever tip stayed on the moleculefor 50 μs). This fact suggested two possible explanations: one isthat the lifetime of E1Ca2 is <50 μs; the other is that SERCA has
no shorter conformation under high ATP concentrations (> ∼1mM). Considering the activation eﬀect of high concentrations(> ∼1 mM) of ATP on the ATPase activity and the Ca2+
uptake228 as well as the ATP concentration-independent lifetimeof the elevated conformation, it was proposed that SERCA would
not transit through the E1Ca2+ state at ATP concentrations
higher than ∼1 mM, and hence would follow a catalytic pathway
diﬀerent from the ordinary Albers−Post scheme. However, thepossibility that the lifetime of E1Ca2 is much shorter than 50 μs,
or the possibility that the chemical state of E1Ca2+ does exist but
there is not suﬃcient time for E1Ca2+ to take a shorter
conformation, cannot be ruled out.
8.5.4. P2X4 Receptor in Response to ATP. P2X receptors
are ATP-gated membrane cation channels transferring cations
across the membrane from the extracellular side to thecytoplasmic side. Seven subtypes of subunits (P2X1−P2X7)
have been identiﬁed.229 The channels are formed as multimers(mainly trimers) of subunits, homomeric or heteromericchannels.230 Each subunit has a large extracellular domain
capable of binding ATP, two transmembrane domains
responsible for the formation of the channel gate, and small N
and C terminal intracellular domains.229 All P2X receptors arepermeable to small monovalent cations, and some havesigniﬁcant Ca2+ or anion permeability.231 P2X receptors areabundantly distributed and involved in diverse physiologicalprocesses from synaptic transmission to inﬂammation andsensation of taste and pain.231 The crystal structure of thezebraﬁsh P2X4 receptor (P2X4R) in the resting closed-channel
state was recently solved (Figure 28a,b).232 The three ATPbinding sites are located at the intersubunit interfaces within thetrimer, suggesting that ATP binding promotes subunit rearrangement and ion channel opening. The ion channel opening of
P2X4R is regulated by the extracellular Ca2+ level.233 Electrophysiology studies have shown that P2X4R opens a small cationpermeable channel pore at a high concentration of Ca2, whereas
in the absence of Ca2+, it forms a large pore allowing permeationof large cationic molecules.234 However, structural studies on thechannel opening and Ca2+ eﬀects on it have been absent until thefollowing HS-AFM study by Shinozaki and collaborators.37
First, isolated (membrane-dissociated) P2X4R attached topolylysine-coated mica was imaged in the absence of ATP and
Ca2+. The still images showed circular, ellipsoid, or triangularparticles, although their sizes were approximately similar to eachother (12.6 ± 0.2 nm in diameter and 2.3 ± 0.1 nm high). The
height value of 2.3 nm is much smaller than the values of either
the height or the width of the molecule estimated from the3150

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ATP adsorption on the polylysine-coated surface. Yet this isunlikely, considering the ATP binding capacity of the surface. In
1 mM ATP, most of the molecules appeared to be trimers with adiameter of 14.2 ± 0.2 nm, a height of 3.0 ± 0.1 nm, and a
distance between lobes of 8.7 nm ± 0.1 nm. The trimer
appearance implied that the molecules would mostly stand on
the substrate with the extracellular top face up, which would be
however hardly understood because the observed height (3.0nm) was much smaller than the distance between theextracellular and intracellular ends (9.8 nm) and because thesmaller surface (∼3 nm in diameter) of the N- and C-terminal
intracellular domains would hardly bind to mica surface
preferentially. The imaged trimers did not exhibit a three-foldaxis of symmetry but heterogeneous asymmetric shapes. Thisappearance suggested that the molecules do not stand uprightbut lean on the substrate surface.
For time-lapse imaging of P2X4R at 2 fps, the sample wasplaced on a bare (i.e., no polylysine-coated) mica surface andimaged. Under this condition, many P2X4R molecules shifted
position during imaging, but some were successfully imaged. The
images again often showed asymmetric shapes, indicating thatthe molecules leaned sideward on the mica surface. To observe
the time course of ATP-induced structural changes of P2X4R,
ﬂash photolysis of caged ATP was used (200 μM). Upon ﬂash
photolysis, individual subunits became visible, and the subunit−
subunit distance gradually increased, but the images were blurry.
Because symmetric molecules were not observed and the images
were blurry, a single-particle correlation averaging method wasapplied to reconstruct 3-fold-symmetric images. The symmetrized images showed that the trimeric structure became moreevident with time and that the central pore was being dilated withtime (0−5 s) after uncaging of caged-ATP without Ca2+ (Figure
28c).
Next, P2X4R reconstructed in a lipid bilayer on a mica surface
was imaged. In the absence of ATP, the appearance of P2X4R was
circular with 11.4 ± 0.3 nm diameter and protruding 1.8 ± 0.1
nm high from the lipid bilayer surface. This height value wasagain too small as compared to the height of the extracellular partin the atomic structure (7 nm). In the presence of ATP, P2X4R
showed a trimeric structure with 13.3 ± 0.3 nm diameter and 2.1± 0.1 nm high from the lipid bilayer surface. The trimeric
structure was more apparent as compared to that observed withisolated P2X4R but was asymmetric again. The reconstructed 3fold-symmetric images of P2X4R in ATP again showed pore
dilation. On the other hand, in the presence of 2 mM Ca2+, nopore dilation occurred in the 3-fold-symmetrized images thatwere reconstructed from raw images obtained several seconds
after uncaging of caged-ATP. To conﬁrm the Ca2+-dependenceof large molecule permeation through P2X4R reconstructed in alipid bilayer, an ethidium bromide (EtBr)-uptake assay was
performed as follows. The lipid membrane containing P2X4R
was suspended over a 500 μm hole ﬁlled only with a buﬀer
solution. In the external side, an EtBr solution with or without
Ca2+ was placed. In the absence of Ca2+, the red ﬂuorescence of
EtBr started increasing in the hole within a few seconds after the
addition of ATP and then increased gradually. On the other
hand, in the presence of 2 mM Ca2+ in the external solution, no
increase of ﬂuorescence was observed. Thus, the P2X4R sample
reconstructed in the lipid membrane was conﬁrmed to be active.
Therefore, the ATP induced pore dilation observed in theabsence of Ca2+ was veriﬁed to be relevant to the conformationalchange of P2X4R responsible for the permeation activity.

Figure 26. Histograms of the height diﬀerences between the top of
SERCA and the surface of the membrane. Statistical section analyses of
SERCA were performed with the data obtained in the presence of (a)100 μM free Ca2+ (n = 78), (b) 10 nM free Ca2+ (n = 54), and (c) 10 nM
free Ca2+ and 10 μM TG, after 30 min incubation (n = 82). The lines are
Gaussian ﬁts for the histograms. Reprinted with permission from ref 50.
Copyright 2011 John Wiley & Sons, Inc.

atomic structure of P2X4R:232 the distance between the ends ofthe extracellular and intracellular parts (9.8 nm); the width of theextracellular top face (7.5 nm). The reason for this large
discrepancy is unclear. Moreover, the orientation of observed
P2X4R molecules on the polylysine-coated mica surface could
not be judged from their images and size values. When 100 μM
ATP was added, no signiﬁcant changes appeared in the P2X4R
images, but addition of 1 mM ATP induced large structuralchanges of P2X4R after several to 30 min. The reason such a longincubation time and high concentrations of ATP were required
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Figure 27. Typical single line scan height data of SERCA obtained with diﬀerent solution conditions. (a) Representative single line scan data obtained atincreasing ATP concentration in the range 0−100 μM and in the presence of 10 nM and 100 μM free Ca2+. (b) Single line scan data for ∼2 s in thepresence of both 1 mM ATP and 100 μm free Ca2+. The lower and higher broken lines indicate heights of 5.5 and 8.0 nm measured from the membranesurface, respectively (a, b). Reprinted with permission from ref 50. Copyright 2011 John Wiley & Sons, Inc.

Figure 28. Structure of P2X4 receptor. (a) Atomic structure of N and C termini deletion mutant of homotrimeric P2X4 receptor viewed parallel to themembrane. Each subunit is depicted in a diﬀerent color. The gray bars suggest the boundaries of the outer (out) and inner (in) leaﬂets of the membrane
bilayer. (b) Atomic structure of N and C termini deletion mutant of homotrimeric P2X4 receptor viewed from the molecular 3-fold axis (from theextracellular side). Reprinted with permission from ref 232. Copyright 2009 Nature Publishing Group. (c) Successive AFM images showing ATPinduced structural changes of P2X4 receptor. Before activation, P2X4 receptor was in circular shape and exhibits some ﬂuctuation (−2.5 to ∼0.0 s).
Caged ATP (200 μM) was uncaged at 0 s, after which the structure of P2X4 receptor changed to a trimer structure within 0.5 s. P2X4 receptor thenexhibited a further structural change and adopted a pore dilation-like conformation. Note that the images are not directly captured ones but are thoseobtained by averaging images of 10 P2X4 receptor particles to reconstruct 3-fold symmetric images at given elapses of time.

8.6. Diﬀusion and Interaction of Membrane Proteins

Nevertheless, two issues remain elusive in this HS-AFM imaging
study. First, the observed heights of the isolated P2X4R and theexposed part of the membrane embedded P2X4R do notcompare favorably with those revealed by the atomic structure of
P2X4R published after this HS-AFM study. It has often beenrecognized that the height of protein molecules appears smaller
than the real height when they are strongly adsorbed on substratesurfaces. However, it seldom occurs when protein molecules areplaced on or embedded in inert lipid bilayers. Second, it is unclear
whether the observed asymmetric structures are inherent to thisreceptor protein.

The diﬀusion and interaction dynamics of membrane proteinsare emerging topics in membrane biology. Their importance has
been recognized with the paradigm shift of the scientists’ view ofthe membrane superstructure (e.g., ref 235): In the early 1970s
biologists conceptualized the cell membrane as a 2D ﬂuid of lipid
molecules in which the relatively few membrane proteins freely
diﬀuse.236 This concept has been amended over the last 30 years(e.g., ref 237), because indirect experimental evidence
accumulated that the membrane features a highly heterogeneous,complex, and intricate superstructure, laterally compartmentalized, densely packed, locally even rigid, and organized. It goes far
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Figure 29. Association and dissociation of bR molecules at crystal edges captured by HS-AFM. (a) Clips of successive HS-AFM images on the borders
between the bR crystal and noncrystalline areas (imaging rate, 3.3 fps). The bR trimers in the crystal are encircled by thin lined triangles. Newly bound(0.6 s) and dissociated (2.1 s) trimers are indicated by red and thick lined white triangles, respectively. (b) Schematic representing the binding manner ofa trimer at the crystal edge (I, II, and III) and in the crystal interior (VI). Roman numerals indicate the number of interaction bonds (dotted lines)
containing W12 residue. (c) Histograms showing lifetimes for the type II and III binding events. The histograms were ﬁtted with single-exponentialfunctions (dotted lines).

facilitate the cell to reorganize proteins in space and in time, toform temporal functional units to perform a particularbiochemical function,, or to modulate the function of amembrane protein.241 HS-AFM appears therefore as aparticularly promising tool for the study of membrane protein
structure, dynamics and interactions: (i) its high signal-to-noise
ratio allows direct visualization of individual membrane proteins
without labeling, (ii) the label-free submolecular resolution
imaging quality warrants additionally the analysis of themolecular environment in the ﬁeld of exploration, and (iii) thehigh spatiotemporal resolution of about 1 nm and 30 ms allows
the assessment of characteristic physical parameters of diﬀusion
and interaction like diﬀusion constants and properties, andinteraction strength and geometry in great detail. Indeed, while
optical microscopy works at comparable frame rate, only thehigh-resolution imaging capacity of HS-AFM allows the direct
visualization of the molecule itself and the partner molecules,
opening a novel avenue of structural and dynamical studies ofprotein−protein interaction in lipid membranes, which will be
described below.
8.6.1. Interaction between bR Trimers in 2D Crystal
Lattice. By observing bR trimers diﬀusing in the noncrystalline
area in the purple membrane and their temporal interaction withthe crystal edges using HS-AFM, the energy for association
between bR trimers through hydrophobic stacking interaction
between aromatic residues was estimated.35 Successive HS-AFM
images of the purple membrane frequently showed repeatedassociation and dissociation of bR trimers at the crystal edges(Figure 29a), indicating that the bR crystal is in dynamic
equilibrium with bR molecules located in the noncrystalline area.
In Figure 29a, bR trimers in the crystal are highlighted by thethin-lined triangles. At 0.6 s, two bR trimers (red triangles) werebeyond the objective of this Review to overview the data that led
to this conceptual change, but the major contributions camefrom optical microscopy showing anomalous diﬀusion behaviorof membrane constituents trapped in compartments and/or
assembling into clusters, biochemical data showing the existenceof supercomplexes of membrane proteins of very high molecularweight and nonsolubilizable membrane domains, and fromfreeze-fracture and tomography EM analyses revealing densely
grained membrane areas. It has, however, so far remained
impossible to visualize a native membrane at molecularresolution and describe the structure, dynamics, and interactionsof membrane proteins within it at high spatial and temporal
resolution. Conventional AFM has been successful in some casesand provided high-resolution views, which was however limited
to highly specialized membranes with exceptional protein
arrangement and density such as photosynthetic membranes orlens membranes.15,238 Light microscopy has undergone dramatic
progress in the recent years, with a signiﬁcant improvement of
the localization resolution of membrane constituents undernative conditions.239 However, the resolution is molecular at bestand is based on the high spatial localization of a single labeledprotein, implying that one cannot see a second protein in close
proximity to the ﬁrst. Electron microscopy has equallyprogressed for superstructural analysis notably with the development of tomography of vitriﬁed samples.240 This technique
however analyzes samples at low temperatures and in ultrahighvacuum and is therefore unable to observe dynamics.
Furthermore, it is a somewhat less powerful technique for theobservation of membrane integral constituents, because thecontrast between the atoms of the transmembrane proteins andthe lipids is limited. Biologically, the diﬀusion and interactions ofmembrane proteins are of crucial importance. These features3153

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Figure 30. H. salinarum ATP-synthase c-ring motion monitored by HS-AFM and energy proﬁle of interaction between c-rings. (a) A series of HS-AFM
images over a time span of 1309 ms (clipped from a longer movie) for two ATP synthase c-rings in dynamic interaction (imaging rate, 5.35 fps). Two crings assembled in a dimer that dynamically changed the center-to-center distance. Rarely, the dimer dissociated and a monomeric c-ring remained (t =561) until a diﬀusing c-ring approached the attractive regime of the monomeric c-ring (t = 748 and 935 ms) and associated (t = 1122 ms). Arrows
indicate diﬀusing c-ring monomers. (b) Three examples of ATP-synthase c-ring assemblies. (Left to right) Close dimer (center-to-center distance 80 Å);
elongated dimer (center-to-center distance 120 Å); and monomer. (c) Interaction between ATP synthase c-rings as a function of center-to-center
distance: (left) histogram (n = 91) of the center-to-center distance of c-rings; (right) energy landscape of membrane-mediated interaction between crings.

as illustrated by the dotted lines in Figure 29b (see the casemarked with “VI”). As exempliﬁed in Figure 29a, successive AFM
images showed many association and dissociation events, inwhich bR trimers were bound to diﬀerent sites at the border
between the crystal and noncrystalline areas. These events wereclassiﬁed into types “I”, “II”, and “III” depending on the numberof interaction sites involved. Type II binding events werepredominant (74%), whereas type I (6%) and type III (20%)
events were less frequent. The type I binding events occurredwith too short bound lifetimes to appear in clear images,
preventing reliable statistics. Figure 29c shows histograms of the
lifetimes of type II and type III bindings that were measured
using HS-AFM images taken at an imaging rate of 10 fps. Eachhistogram was well ﬁtted by a single exponential function. The
average lifetimes τ2 of type II binding and τ3 of type III binding
were estimated to be 0.19 ± 0.01 and 0.85 ± 0.08 s, respectively.

newly bound to the crystal edge. At 2.1 s, two bR trimers (white
triangles) dissociated, and another trimer (red triangle in the topof image) was bound to the crystal edge. One of the two
dissociated trimers remained in the crystal edge for ∼0.9 s. Not
only bR trimers but also bR dimers and monomers wereobserved to bind to and dissociate from the crystal edge, thesemuch less frequently than trimers. The bound lifetimes of bR
monomers and dimers were shorter than that of trimers. Among
many observed binding events, the binding of trimeric bR
occurred predominantly (82%), indicating that trimers aremostly formed and stable in the noncrystalline region.
The intertrimer interaction energy was estimated from thebound lifetime of newly associated bR trimers at the crystal edge
and its dependence on the number of interaction sites. From thepacking arrangement of bR trimers in the 2D crystal, each trimer
appears to interact with the surrounding trimers through six sites,3154

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Center-to-center distances shorter than 8.0 nm were neverobserved. From the favorable center-to-center distance of 10.3
nm, the interaction energy landscape described an attractive
regime that extended by a radius of about 3.5 nm in themembrane plane. Membrane-mediated protein−protein interaction has been intensively studied by theory and computation,
depicting an interaction proﬁle with an attractive well at shortdistance (about 5 nm around a protein) that can be extended by afaint attractive regime at larger distance.241 The general idea has
emerged that the membrane protein perturbs the lipid bilayer
structure and that the proteins associate to each other tominimize the energetic cost of this perturbation. Typically, themembrane proteins force the bilayer to bend so as for it to adapt
to the rigid hydrophobic core of their transmembrane segments;
this bending is predicted to relax over a distance comparable tothe bilayer thickness. In agreement, HS-AFM measurements
showed that the lipid bilayer between the two c-rings, even if
these were in an elongated dimer conformation with up to 3 nm
lipid bilayer space between them, adapted to the hydrophobic
thickness of the protein and was 0.5−1.0 nm thicker, as
compared to the bilayer environment far from the c-rings.
Furthermore, the interaction energy landscape showed that at a
center-to-center separation of 12.5 nm, hence at an elongation
distance of about 4 nm from protein−protein contact (in good
agreement with the bilayer thickness), the attraction was only
about −1 kBT, and hence thermal motion could separate the two
proteins, in good agreement with theoretical considerations.
8.6.3. Interaction of AQP0 in Native Eye Lens
Membrane. As seen in the above analysis of bR and A-ATP
synthase c-rings in native purple membranes, HS-AFM has
proven powerful in revealing the interaction dynamics andenergy and conformational changes of bacterial membraneproteins.35,39 However, a step toward a deeper understanding of
membrane structure and membrane protein interaction can begained from eukaryotic systems. In eukaryotic cell membranes, avariety of phospholipids are found, and, importantly, nonphospholipid membrane constituents such as cholesterol areevidenced to drive membrane superstructure.246 The variety of
constituents with diﬀerent physicochemical properties leads to anatural segregation of the membrane constituents, given that not
all interactions are in the range of kBT. Indeed, this led to theconcept of mosaicity of the membrane, supported by indirect
biochemical and biophysical data.235,237 This vision of themembrane comprises both structural and dynamic aspects.
Therefore, HS-AFM appeared to be a most promising tool toinvestigate membrane constituent segregation in eukaryotic
systems.237 This is, however, not so easy, because the membranes
must be puriﬁed from a mammalian tissue, and suﬃcient purity
must be achieved for analysis by HS-AFM while it must be
assured that the native assembly of the membranes remains
unaltered. In mammalian eye lens cell membranes, membranechannels form supramolecular domains termed junctional
microdomains responsible for metabolite ﬂux, ion and watertransport, and cell−cell adhesion.247,248 Junctional microdomains are mainly constituted of lens-speciﬁc aquaporin-0(AQP0) and connexins (Cx), the ﬁrst forming stable tetramers
and the second hexamers termed connexons. AQP0 and Cx
constitute about 60% and 15%, respectively, of the entire
membrane protein amount. Conventional AFM allowed thecharacterization of the supramolecular arrangement of these
proteins in junctional microdomains at high resolution, showing
the AQP0 assembly in regular square arrays edged and separated
by densely but nonregularly packed connexons.238 Both of these

The longer lifetime of type III binding than that of type II
obviously arises from a relationship of E3 < E2 < 0, where E2 andE3 are the association energies responsible for type II and type III
interactions, respectively. The average lifetime ratio, τ2/τ3, isgiven by τ2/τ3 = exp[(E3 − E2)/kBT] (T is room temperature in
Kelvin). Because the type II and III bonds involve two and threeelementary bonds, respectively, the energy diﬀerence E3 − E2should correspond to the association energy of the single
elementary bond formed by hydrophobic stacking interaction
between W12 and F135.62 This elementary association energy
was estimated to be about −1.5 kBT from the ratio of τ2/τ3 = 0.22.
This value approximately coincides with that measured by
diﬀerential scanning calorimetry.210
8.6.2. Interaction of C-Rings in Purple Membrane.
Although less abundant than bR, the purple membrane of H.
salinarum also contains an ATP synthase that uses thetransmembrane proton gradient generated by bR to produce
ATP. Little is known about this ATP synthase: It is an A-ATP
synthase (A for archaea), but sequence analysis indicatessimilarities to both F- and V-type ATP synthases.242 Despiteextensive studies of purple membranes using conventional
AFM,206 these molecules have long escaped being observed,
probably due to their mobility outside the crystalline bR arrays.
Taking advantage of the fast frame acquisition rate of HS-AFM,
A-ATP synthase c-rings were observed in the lipid bilayer regionsat the edges of bR 2D lattices (Figure 30).39 Mass spectroscopyof the investigated sample corroborated the presence of proteins
with a mass of 7517 and 7618 Da corresponding to the native cring sequence.243 C-rings occurred as rings with 6.5 ± 0.5 nm
diameter and 1.8 nm protrusion height, which formed dimers in
the membrane. Dimer formation has been described for
mitochondrial ATP synthases, but the prokaryotic homologues
have been thought to be monomers. The data hence suggested
that the A-ATP synthase forms superstructures reminiscent of
those found in mitochondria.244,245 The dynamic HS-AFM
imaging allowed one to monitor the interaction of the c-rings in
the membrane plane at 5.35 fps.39 The c-ring interaction was
highly dynamic; the center-to-center distance changed as afunction of time (Figure 30a), and sometimes one c-ring evendissociated from the assembly (Figure 30a, arrows). Hence, three
diﬀerent c-ring association states occurred (Figure 30b): themonomer, when one c-ring was found without a partner
molecule, the close dimer, when two c-rings were in directcontact and the center-to-center distance was between 8.0 and
10.0 nm, and the elongated dimer, when two c-rings wereseparated with center-to-center distances of about 12.0 nm. The
mean square displacement (MSD) as a function of the lag-timeΔt revealed diﬀerent diﬀusion regimes for the c-ring monomer
and dimer states. Fitting the generalized diﬀusion equation MSD∝ Δtα to the experimental data, the exponent α reported free
diﬀusive and subdiﬀusive behavior for the monomer and dimer,
respectively. The corresponding diﬀusion constants D were 125
and 1.5 nm2/s, following MSD = 4DΔt. From the dynamic
description of the c-ring interaction, a distance-dependent
probability density function could be derived (Figure 30c, left)that could be translated into an interaction energy proﬁle, p(d) =p∞ exp[−E(d)/kBT] (Figure 30c, right), where p(d) is the
distance dependent probability, E(d) is the related interactionenergy, and T is 300 K. This energy landscape described the cring interaction with a maximum strength of −3.5 kBT at a centerto-center distance of about 10.3 nm. At shorter distances, soft
hard-body repulsion occurred, probably originating from thecontact and intercalation of exposed amino acid side chains.
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Figure 31. HS-AFM analysis of vertical and lateral association of lens-speciﬁc AQP0 in native eye lens membranes. (a) Contact mode HS-AFM imagingof eye lens junctional microdomains during nanodissection captured at 1.02 fps. Initially two stacked membrane layers were images (frame t = 0.0 s)
from which the top layer is sequentially removed in distinct membrane fractions of area of about 10 000 nm2 (from t = 4.9 s to t = 38.2 s). Finally, theentire top membrane of the junctional microdomain was removed (t = 70.5 s), revealing that the bottom layer comprised protein domains of the same
shape and localization as the top layer (labeled with 1 and 2, in frames t = 0.0 s and t = 70.5 s). (b) High-resolution HS-AFM visualization of AQP0
tetramers in dynamic association to and dissociation from square AQP0 array (imaging rate, 4 fps). The assembly and disassembly of AQP0 occurring at
the edge of the junctional microdomain array were observed (see arrows in frames t = 0.0 s and t = 2.5 s).

unit cell dimension and about 1 nm protrusion height, edged byconnexon rings that were nonordered and protruded further by
about 2 nm. HS-AFM medium resolution observations readily
revealed protein motion at the edges of the junctional
microdomains. Following, high spatial and temporal resolution
HS-AFM imaging (single-molecule resolution at 4 fps) of thejunctional microdomains depicted the assembly of AQP0 andconnexons to and their disassembly from junctional microdomains (Figure 31b). Following the same statistical analysis of
the image series as has been performed on bR,35 the molecules
associating with and dissociating from the microdomain edge
were classiﬁed according to the number of their intermolecular
contacts (Figure 31b, arrows), although only relevant statistics
for molecules making one and two neighbor contacts could beassessed, and their residence lifetimes at these positions were
evaluated. Lifetimes of τ2 = 9.8 ± 0.5 s and τ1 = 0.7 ± 1.0 s were
found for AQP0 with two and one neighbors, respectively. Usingτ1/τ2 = exp[(E(two bonds) − E(one bond))/kBT], an interactionenergy of −2.7 kBT was determined for a single AQP0−AQP0
interaction. To test these experimental results, a single parameter
Monte Carlo simulation was run following p(diss) = exp[(n ×E(bond)/kBT)], where p(diss) was the probability of an AQP0 to
dissociate, n is the number of neighbors, and E(bond) is the energy
per single bond. Such simulations (over 300 000 simulation
steps) with the use of the experimentally determined energetic
term of −2.7 kBT (independent of the molecular concentration at
the beginning of the simulation) showed AQP0 assembly in silico
forming junction microdomains. In contrast, simulations at lower−1.7 kBT or higher −3.7 kBT interaction energies either
prohibited formation of domains or formed domains withmembrane proteins are not only channels, but act as celladhesion molecules by means of their extracellular loops thatinteract homotypically with the proteins from the neighboring
cell; two head-to-head assembled AQP0 tetramers form a socalled thin-junction, and two head-to-head assembled Cx
hexamers form a so-called gap-junction. 249,250 HS-AFM
provided novel insights into both the vertical (cell−celladhesion) and the lateral (domain formation) associations ofthese junctional microdomain proteins.60 The lens cell
membrane preparations maintained the native double membranearchitecture of the junctional microdomains (Figure 31a, frame t= 0.0 s). Taking advantage of the nanomanipulation capacity of
the cantilever tip, HS-AFM in contact mode under application of
additional forces allowed the dissection of junctional microdomains (Figure 31a). Analysis of the junctional microdomain
areas and circumference as a function of time during microdomain dissection revealed that the adhesion function of theproteins was cooperative; entire membrane patches dissociatedat once (Figure 31a, compare frames t = 14.7 and 19.6 s). From
this study, the size of adhesive units could be determined as about
100 × 100 nm2 assembly of about 250 AQP0 tetramers, inagreement with imaging of the junctional microdomainarchitecture. Additionally, by imaging of the double layer(prior to dissection) and the single layer (following dissection),junctional microdomains proved to be the homotypic head-tohead association of AQP0 and connexons for cell−cell-adhesion(numbered (1) and (2) in Figure 31a). After junctional
microdomain dissection, the extracellular face of the proteins
could be studied on the single layers at high resolution. AQP0
was arranged in a characteristic square lattice with 6.3 ± 0.1 nm
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Figure 32. HS-AFM analysis of the diﬀusion and interaction dynamics of trimeric OmpF. (a) High-resolution HS-AFM visualization of the dynamic
movement of about 70 OmpF trimers in the membrane (imaging rate, at 2.1 fps). The resolution is high enough to allow determining the lateral position
and the rotational angle of every single channel. (b) The eﬀect of local membrane crowding on the diﬀusion behavior of individual OmpF trimers. The
average position (black star) and positional standard deviation (black error bars) were determined for each molecule by the lateral and angular trackingof the proteins in subsequent frames (colored lines and arrows). From the average positions, the Voronoi tessellation of the membrane surface wascalculated (black meshwork) depicting the free membrane area that each molecules experiences. Molecules housed in large Voronoi cells (i.e., large freemembrane space) revealed high mobility, while molecules in locally and densely packed regions were essentially immobile. Four trimers in a rather solid
association are outlined (1) in correspondence to the molecules outlined (1) in frame t = 1908 ms in (a). A highly mobile single molecule is outlined (2)corresponding to the molecule outlined (2) in frame t = 2385 ms in (a).

of boxes within the protein regions then was plotted versus thelogarithm of the size of the boxes to quantify the fractal
dimension D deﬁned by the slope of the plot. The OmpF
distribution in the membrane had a high value of D ≈ 1.7, while asimulation of OmpF arranged in a 2D-crystal gave D ≈ 1.0. Sucha high fractal dimension is characteristic of diﬀusion-limited
aggregation;257 hence the molecular movements result in a
dynamic equilibrium and arrange at any instant for the proteins
to be exposed to a wide variety of molecular surfaces in themembrane plane. Beyond this case example, the calculation of
the fractal dimension appears to be an appealing quantity toassess membrane protein assembly, as it reports in a single
number the complexity of the molecular distribution mosaicity.
Moving from static to dynamic analysis, a wide distribution of
dynamic behaviors of individual proteins was observed; somemolecules participated in stable associations with others (Figure
32a, outline 1), while some diﬀused in less dense membraneregions between the aggregates (Figure 32a, outline 2). The
eﬀect of the local membrane crowding on the dynamics of themolecules, which was determined by Voronoi tessellation,
showed that diﬀusion scaled roughly in inverse relation to the
local protein density (Figure 32b). The Voronoi cell area
histogram described roughly a Gaussian distribution withsmallest Voronoi cells of about 7000 Å2 and largest cells of
about 20 000 Å2, peaking at about 12 000 Å2 (Figure 32b). In
comparison, OmpF packed in 2D crystals within a membrane
area of 5500 Å2 per trimer.258 In agreement with the membrane
area dispersion, some molecules displayed corralled subdiﬀusive
behavior (Figure 32b, outline 1), and others diﬀused freely(Figure 32b, outline 2) with a slope of 1.1 in a log(MSD) versuslog(Δt) plot and with a maximum displacement velocity of about
25 nm/s. Because the diﬀusion behavior and the local membrane
density were coupled, averaging of the local molecule velocities
showed that certain membrane areas were favorable formolecular motion while others were in a static phase. Close

noncharacteristic morphology. Finally, connexons were revealed
to have a diﬀerent association behavior; their ﬁnal arrangement
was nonordered. Additionally, connexons associated anddissociated so rarely that the dynamics could only sporadicallybe observed by HS-AFM, precluding determination of an energy
term that should be higher than that of AQP0. However, thisstability might well be related to their function as moleculessustaining relevant forces in cell−cell adhesion.251
8.6.4. Diﬀusion and Interaction of OmpF. HS-AFM
imaging of the diﬀusion and interaction of OmpF provided themost detailed information and maybe the most general insights
to date about membrane protein interactions in the lipid plane.59
OmpF and its homologues are widely distributed on the cellsurface of many bacteria, where they act as an ion and solutechannel and for phage recognition, and form a transienttranslocon complex with vitamin B12 receptor BtuB for colicin
import.252−254 In the outer membrane of E. coli, there are about
105 OmpF copies inserted, covering at least 50% of the cellsurface.255,256
OmpF were reconstituted at a realistic protein density of about
50% membrane coverage into a bilayer containing all of the lipid
constituents of the native E. coli system. In particular, thepresence of lipopolysaccharides associated in complex with
OmpF was assured. HS-AFM allowed the analysis of thestructure and dynamic aspects of about 70 OmpF trimers in eachframe at high spatial and temporal resolution; individual subunits
were resolved, and movie acquisition rates were 2−5 fps. From astatic point of view, the proteins embedded at low 50% density inthe ﬂuidic lipid bilayer were found to be loosely distributed overthe membrane at any instant during the observation (Figure32a). To assess the protein distribution quantitatively, fractal
analysis, using the box method, was performed. In this in silico
procedure, the individual protein outlines were compared tooverlaid square boxes of various sizes. Boxes that fall into the ﬁeldof the protein outlines were scored. The logarithm of the number
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formations possibly occur in the intermediates, which control
the self-assembly pathways and kinetics.263 As in the cases ofpolymerization of actin to actin ﬁlaments and tubulin to
microtubules, small ligand molecules (e.g., ATP in the case ofactin and GTP in the case of tubulin) and their chemical
reactions are sometimes involved in protein self-assembly.264
Moreover, polarity or anisotropy exists in the assembled
structures of proteins, and hence the assembly kinetics diﬀers
from one place to others in the growing structures. Thus, selfassembly of proteins is highly complex, and hence it is diﬃcult todecipher the entire molecular process using static, nonvisualization, or ensemble-averaging approaches. HS-AFM can simultaneously visualize several assembly processes that are ongoing
sequentially and in parallel at diﬀerent portions of growing
structures, and therefore is the most powerful approach to
studying self-assembly of proteins as well as other molecules.
Here, three examples of HS-AFM observations on self-assembly
processes are described.45,47,86
8.7.1. Amyloid Fibril Formation. Amyloids are ﬁbrous
protein aggregates, and their deposition is often found in tissues
from patients with diﬀerent types of diseases and disorders.
Therefore, amyloids are associated with various diseases,
particularly neurodegenerative diseases such as Alzheimer’s,
Huntington’s, Parkinson’s, Creutzfeldt−Jacob, and prion diseases,265,266 although in some cases amyloids are functional inliving organisms.267 Rather than precipitated insoluble aggregates, intermediate aggregated species have recently been
implicated in cellular toxicity and are likely to be the source ofaberrant pathology in amyloid diseases.268 About 30 diﬀerent
proteins including peptides have been identiﬁed as the major
components of the deposits associated with diseases. The
propensity of proteins to aggregate into amyloid-like structures
has been experimentally shown to be well correlated with adecreased conformational stability in the native state.269
Therefore, misfolding (or conformational changes) is believed
to be prerequisite to amyloidogenesis in most cases. The
structures of proteins in amyloid, intermediate species occurringin the growth process, and the outline of the growth kinetics havebeen well studied. Nonetheless, the dynamic processes andmultiple pathways through which the ﬁbers are formed havelargely been elusive. Polarity/anisotropy-dependent growth
kinetics has also been elusive because of a lack of usefultechniques.
Lithosthatine (also named Reg-1) of 144 amino acids is amember of proteins that generate amyloid-like ﬁbers. It is
produced by pancreas acinar cells and secreted into pancreatic
juice. It tightly binds to calcium carbonate and may act as an
inhibitor of calcium carbonate precipitation to prevent clogging
of pancreatic duct.270 In chronic calcifying pancreatitis, theprotein forms deposits in the pancreatic duct. Lithosthatine isalso expressed in other organs including stomach and brain.
Lithosthatine is susceptible to self-proteolysis under speciﬁc pH
conditions, producing a soluble N-terminal undecapeptide and a
C-terminal fragment of 133 amino acids. The cleaved protein(called S1) forms protease-K-resistant ﬁbrils271 that deposit in
the brain of patients with Creutzfeldt−Jacob disease andGertsmann−Sträussler−Scheinker syndrome, especially during
the very early stages.
HS-AFM was used to observe the ﬁbrillization process carriedout by lithosthatine.47 Upon 15 min incubation of the S1 form oflithosthatine generated by tryptic digestion of the full-length
protein, globular structures with 15−28 nm in diameter
appeared. They seemed to be small assemblies of lithosthatine

inspection of the molecules that remained stable over 2 min of
HS-AFM movie recording showed that these proteins wereassociated with other channels in speciﬁc aggregation conformations using speciﬁc surface interactions. Interestingly, theseinteractions often implied a symmetry relation with the neighbormolecule, such as dimers arranged in a p2 symmetry with 72 Å
center-to-center distance or trimers arranged in a lattice of p3
symmetry reminiscent of that adopted by bR, with 86 Å centerto-center distance. Statistical analysis of the interaction between
the proteins that were in dynamic encounter and dissociationfrom each other showed that OmpF channels followed a speciﬁc
interaction pathway. It appeared that the interaction contact sites
at the periphery of the molecule were the last stable contact
points before dissociation and could mediate processes into
stable assembly. Indeed, the interaction pathway implied sliding
and rotational motion of OmpF on the surface of the neighbormolecule; the molecules were going from the closest association
with a center-to-center distance of ∼7.2 nm (where the two ﬂat
surfaces of the proteins were intercalated into each other with p2
symmetry over an asymmetric association) to anotherconﬁguration with p2 symmetry, where the partners were onlyin contact at the periphery of the molecules with a large centerto-center distance of ∼10 nm. This ﬁnding was similar to the HSAFM observation of speciﬁc interactions of bR molecules via
exposed aromatic residues,62 an aspect that should be tested
further in future experiments. Coarse-grained molecular
dynamics simulations259 have been used to analyze theinteraction pathway computationally, which further underlines
the importance of angular lipids in the membrane protein−protein interactions. HS-AFM imaging and such simulations
agreed with each other in that several stable association patterns
occurred and were dynamically adopted. The patterns ranged
from an energetically favored association minimizing interaction
distance and maximizing protein−protein interfaces to makeprotein−protein contacts via peripheral portions strongly
mediated by the membrane lipids. This series of events andvariety of conformations are in a dynamic equilibrium
maintained by thermal energy, probably assuring the versatilefunctionality of OmpF in recognition and transport.
8.7. Self-Assembly Processes

Self-assembly is universal in biological systems and underlies theformation of a wide variety of ordered biological architectures.
This strategy that nature has adopted is largely diﬀerent from that
we do to produce man-made machines, in which we needblueprints as well as process charts. In biological self-assembly,
the blueprints and process charts are mostly encoded in theelements themselves to be assembled. Although molecular selfassembly is an excellent strategy for automated and energy-saved
construction of large ordered structures in most cases, it
sometimes works in a wrong way to produce substances
detrimental to health, as typically exempliﬁed in prion-relateddiseases. Thus, self-assembly of proteins in particular has been an
important topic not only in biophysical studies and engineering
applications but also in the studies of the pathogenesis of diseases
and their prevention and treatment.260,261
As is generally seen in self-assembly, protein self-assembly
starts from the formation of nucleation clusters, which provide alow-energy barrier pathway to further growth. Multiple
intermediates often appear during growth, and hence the growth
process toward the ﬁnal ordered structure usually proceeds
through multiple sequential and parallel (i.e., branched)pathways with diﬀerent kinetics.262 Conformational trans3158

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Figure 33. High-speed AFM images showing the time lapse of lithosthatine protoﬁbril elongation and association. Elongation of single protoﬁbrils isshown by white arrows in (a), (b), and (d). Some protoﬁbrils did not grow in length, as indicated by white arrowheads in (a) and (b). Interestingly, theend of an elongating protoﬁbril was fused with one end of a pre-existing protoﬁbril, forming a continuous structure, as marked with a light-blue arrow in(b). Protoﬁbrils were often laterally associated with another, at least for one part of their length. The complete sideways association between two
protoﬁbrils occurred in a zipper-like mechanism (pink arrows). Elongation of a protoﬁbril also occurred, being stacked on a ﬁbril, as indicated with thelight-blue arrowheads in (c). The growth of a protoﬁbril or ﬁbril was stopped when the growing end encounters a side edge of a second ﬁbril, forming a
false branching point with a certain range of contact angle, as indicated with red arrowheads in (d). Winding of two protoﬁbrils formed helical ﬁbrils (e)(the inset is a ﬁltered image).

top of laterally associated protoﬁbrils (light-blue arrowheads in
Figure 33c). The lateral association of protoﬁbrils to ﬁbrils took
place in a zipper-like mechanism (pink arrows in Figure 33b). In
the initial stage, two protoﬁbrils made contact with each other ina small region, showing a Y shape. The separated protoﬁbrils
were getting closer to each other by a gradual increase of thecontact region, and eventually the two protoﬁbrils werecompletely associated to form a helical ﬁbril similar to otheramyloid ß ﬁbrils (Figure 33e). It was also clear that a branchedstructure previously observed with lithosthatine ﬁbrils272 is a false
structure. The growth of a protoﬁbril or ﬁbril was stopped when
the growing end encountered a side edge of a second ﬁbril,forming a false branching point with a certain range of contact
angles (red arrowheads in Figure 33d).
Thus, the dynamic visualization of lithosthatine ﬁbrillization
revealed various elementary processes contained in theﬁbrillization, including the existence of polarity in a protoﬁbril(and hence also in a matured ﬁber) with markedly diﬀerent
elongation rates at the head and tail ends of a protoﬁbril.
8.7.2. 2D Crystallization of Annexin A5. Annexins are aclass of soluble proteins comprising more than 150 members,
which bind in a Ca2+-dependent manner to anionic phospholipids, particularly those containing serine at the polar heads.

tetramers with 9 nm in diameter and 2.5 nm in thickness. After 30
min incubation, protoﬁbril formation and its growth to longer
protoﬁbrils and thicker ﬁbrils were observed (Figure 33).
Because no entities (monomers or small oligomers) to be
added to the growing ends were visible, they must be ﬂoating anddiﬀusing in the bulk solution. Lengthening of protoﬁbrils took
place at 27−52 nm/s, in two ways: (i) addition of lithosthatine
oligomers to the ends of protoﬁbrils (white arrows in Figure 33a,b, and d) and (ii) fusion of a growing protoﬁbril to a pre-existing
protoﬁbril (light-blue arrow in Figure 33b). Interestingly, one
end of each of some pre-existing protoﬁbrils did not change its
position (white arrowheads in Figure 33a and b), suggesting amarked diﬀerence in the oligomer addition rate at the two ends ofa protoﬁbril and therefore the existence of polarity in the
protoﬁbril. Thus, the fusion of a growing protoﬁbril to a preexisting nongrowing protoﬁbril (light-blue arrow in Figure 33b)
is very likely to take place by head-to-tail connection. This issue
may be able to be veriﬁed directly by cutting across a protoﬁbril
using a cantilever tip and by the following observation ofelongation at the exposed ends, as has been done for an actin
ﬁlament.38
Elongation of ﬁbrils appeared to occur separately at eachconstituent protoﬁbril. In fact, a protoﬁbril was elongated on the3159

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Figure 34. Dynamic process of 2D crystal formation of annexin A5 captured by HS-AFM. The image at t = 0 s shows the lipid bilayer surface on whichannexin A5 molecules are diﬀusing. The diﬀusion appeared as large noise. After a CaCl2 solution was injected at t = 7 s, 2D crystals were formed throughinitial nucleation step, gradual assembling of nuclei, and rapid growth of crystal patches by the addition of molecules to the crystal edges. Scan area, 400 ×
400 nm2; imaging rate, 1 fps.

process seemed still high because the growth rate was low. As
seen at the top left of the images at t = 3 min and 39 s, 4 min and
24 s, and 5 min and 6 s, the growth rate was much higher for alarger cluster, which already had shown a well-ordered crystal
structure. During this rapid growth of the crystal with a much
lower energy barrier for monomer addition, the precursor
clusters were engulfed into the growing crystal, as seen in theimages at t = 5 min and 6 s and 5 min and 25 s. As seen in theimage at t = 6 min and 13 s, the lipid bilayer surface within theobservation area of 400 × 400 nm2 was ﬁnally covered by the
crystal.
After completion of crystallization, a highly ordered structurewith p6 symmetry appeared as previously reported279 (Figure
35a; scan range, 150 × 150 nm2; imaging rate, 2 fps). In the p6
crystal, the A5 trimers are arranged in a hexagonal honeycomb
pattern as shown in Figure 35b. Each “hole” in the honeycomb
structure tended to be occupied with a relatively mobile trimer(colored by pink in Figure 35b). The caged central trimer
underwent a more relaxed interaction with its surrounding cage
than trimers forming a part of the honeycomb lattice. Because ofthis mobility, the caged trimer has shown a less sharply deﬁned
density in the EM images of the crystal.279,280 The hole indicatedby the arrow at t = 0 s in Figure 35a was occupied by a trimer in
the next frame. This trimer was weakly bound and then soon
dissociated at t = 1 s. The resulting hole was ﬁlled again with atrimer at t = 1.5 s, but this trimer was now bound more stably.
The hole formation and ﬁling occurred at several places in the
crystal (images between 18 and 23.4 s). Because surface unboundmolecules had been washed away, this observation indicated thattrimers not incorporated in the crystal exist on the crystal surface
and are rapidly diﬀusing on it. HS-AFM imaging also revealed
rotational diﬀusion of a caged trimer weakly bound to thesurrounding cage (Figure 36; scan range, 50 × 50 nm2; imaging
rate, 5 fps). The caged trimer encircled in the image at t = 0 srotated counterclockwise by a 60° step. In the cage surroundedwith six trimers, the central trimer can assume six stable positions
with identical association energy. This rotational motion
indicated the association energy to be approximately −1 kBT.

Because of this property, annexins are involved in many
membrane-related functions: certain exocytic and endocytictransport, anchoring other proteins including actin cytoskeleton
to the cell membrane, organization of membrane domains
including rafts, and the regulation of ion ﬂux across the cell
membrane.273 Annexins not only bind to the cytosolic surface ofcell membranes as single molecules but also form lateral
assemblies at least on model membranes.274 In the early stage
in apoptosis, the lipid compositions in the inner and outermembrane leaﬂets of the bilayer are altered. In particular, afraction of phosphatidylserine increases at the outer leaﬂet, which
recruits extracellular annexins to form 2D crystals of annexins on
the surface.275 The crystals are supposed to induce membrane
indentation and subsequent inward vesicle budding and change
membrane curvature and therefore cell shape.276 Among many
types of annexins, 2D crystallization of annexin A5 on model
membranes has been well characterized by cryo-electron
microscopy and AFM.102,277,278 In the 2D crystals, trimers
represent the fundamental oligomeric state, which is stabilized by
the contact between trimers.
HS-AFM observations provided novel insights into several
dynamic events concerning 2D crystallization of annexin A5.86
As a model membrane, a lipid bilayer consisting of DOPC,
DOPS, and DOPE at ratio 5:2:3 w/w formed on bare mica wasused. The observed crystallization process is shown in Figure 34(scan range, 400 × 400 nm2; imaging rate, 1 fps). In the initialstage of the observation, CaCl2 was omitted from the observation
buﬀer. Under this condition, images with many spike noises
appeared (see the image at t = 0 s), which is due to annexin A5
molecules rapidly diﬀusing on the lipid bilayer surface. The image
gradually changed with time after the injection of CaCl2 (ﬁnalconcentration, 3 mM) at t = 7 s. As shown in the image at t = 31 s,
much less mobile small particles appeared, indicating the ﬁrst
stage of self-assembly (likely to be trimer formation). The size ofthe particle then increased with time as shown by the image at t =3 min and 39 s, indicating the formation of precursor clusters bymonomer addition to the initial clusters from surrounding
diﬀusing monomers. However, the energy barrier in this addition
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Figure 35. Dynamics occurring in annexin A5 crystal captured by HS-AFM. (a) Binding and dissociation dynamics of annexin A5 trimer on the 2D
crystal (scan area, 150 × 150 nm2; imaging rate, 2 fps). The hole in the honeycomb lattice indicated by the arrow in the image of t = 0 s was ﬁlled at 0.5 s
by a trimer that had been diﬀusing on the crystal surface. In the next 0.5 s, the trimer was dissociated. Similar events occurred at several places in the
crystal. (b) Model of the 2D crystal of annexin A5. The caged central trimer in the honeycomb structure is colored in pink.

Figure 36. Successive HS-AFM images showing rotational diﬀusion of a trimer trapped in a lattice cage of 2D annexin A5 crystal. Scan area, 50 × 50 nm2.
Imaging rate, 5 fps.

strong cooperative interactions between molecules in the crystal.
Finally, the entire crystal disappeared within a few minutes.
8.7.3. Supported Lipid Bilayer Formation. The supported lipid bilayer (SLB) formation occurs by self-assembly of
lipids, which is considered to proceed in three steps:281 (i) small
lipid vesicles (small unilamellar vesicles (SUVs)) are attached toa mica surface and then rupture because the vesicles are largely
deformed by the electrostatic interaction with the negatively
charged mica, (ii) upon rupture, small patches of SLB are formed,

The disassembly process of the 2D crystal was also monitored
by HS-AFM following chelation of Ca2+ ions. Figure 37 shows
HS-AFM images clipped from the successive images (scan range,
500 × 500 nm2; imaging rate, 1 fps). At t = 8 s after the injectionof an EGTA solution into the observation buﬀer at t = 0 s (during
imaging), the 2D crystal form was still maintained. After a while,some trimers, especially weakly bound trimers caged in the holes,
were removed from the lipid membrane. Once a disrupted region
appeared, the 2D crystal rapidly collapsed from there, indicating
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Figure 37. Disassembly process of 2D annexin A5 crystal captured by HS-AFM. An EGTA solution was injected into the sample chamber at 0 s. Scanarea, 500 × 500 nm2. Imaging rate, 1 fps.

the LNT followed by fusion with the interacting SLB patch.
When an LNT interacted with the edge of a SLB patch at one of
the free ends, the size of the patch was often getting smaller withtime, and, more surprisingly, the length of the LNT became
shorter, as shown in Figure 38d.
With further lapse of time, the length and density of LNTs
increased, and the size of SLB patches also increased. As a result,long LNTs were jammed into narrow spaces between large SLB
patches, thus pushing each other (images at 20 and 20.9 min in
Figure 38a). Some LNTs were collapsed in this congested
condition to give larger SLB patches, whereas other LNTs
quickly disappeared, very likely to be detached from the surface
because exposed mica surface became unavailable for their
attachment.
The high stability of LNTs can be accounted for by the smaller
surface curvature (and hence smaller surface tension), as
compared to that of small globular liposomes. In the sideways
surfaces of the edge regions of SLB patches and the ends of
LNTs, the polar head groups rather than alkyl chains are exposed.
The arrangements of lipid molecules at these surfaces areenergetically unfavorable as compared to those in the ﬂat surface
area of SLB patches and the longitudinal surfaces of LNTs.
Therefore, to reduce the fraction of these unfavorable surfaces,
SLB patches and LNTs grow by incorporating free lipids and byfusing with the respective kind of partners. As mentioned above,
when one of the ends of an LNT was brought into contact with asideways edge of an SLB patch, the area of the patch became
smaller with time, and the length of the LNT also became shorter
with time. The SLB patches are thin (a few nanometers) while
the LNTs are ∼20 nm in diameter, so that the arrangements oflipids facing at the sideways edges of SLB patches areenergetically unfavorable, as compared to those of lipids at theends of LNTs. This energetic diﬀerence would result in lipid
transfer from the SLB to the LNT, explaining the size reductionof the SLB. Nonetheless, it is hard to understand the shorteningof the LNT.

and then (iii) the patches are fused into larger SLBs, andeventually the entire mica surface is completely covered by thebilayers. However, when this entire process was imaged for theﬁrst time using HS-AFM, unexpected phenomena appeared,45
which are described below.
The lipid composition of small vesicles used was DOPC,
DOPS, and biotin-cap-DPPE in ratio 3:1:1 (w/w). The vesicles
were suspended in a buﬀer solution of pH 7.4 containing 2 mM
CaCl2. Images clipped from successive HS-AFM images captured
over 25 min after the deposition of the vesicles onto a mica
surface are shown in Figure 38a (imaging rate, 1.03 fps; scan size,
800 × 800 nm2). One of the unexpected phenomena observed
was the formation of lipid nanotubes (LNTs) with a uniform
diameter of ∼20 nm. Note that lipid nanotubes were recently
observed (by ﬂuorescence microscopy) to be formed from giant
vesicles containing biotin-lipid when the vesicles were exposed to
streptavidin.282
Short LNTs had already been formed in the vesicle sample
before the deposition onto mica surface as they appeared
together with small globular vesicles in the AFM images
immediately after the deposition. Unlike the globular vesicles,
the short LNTs did not rupture on the mica surface and grew
longer with time. However, the growth rate was markedly
diﬀerent from one LTN to another. The reason for this diﬀerence
was unclear. The fastest one was the LNT that appeared at the
top right and was growing toward left at ∼10 nm/s (marked withthe arrow in the image at 5.3 min). When a growing end
encountered the side of a pre-existing LNT, the growth was
stopped. The diﬀerence in the growth rate between the two ends
of a LNT was uncertain because the two ends were in somewhat
diﬀerent environments, and because the growth rate wasmarkedly diﬀerent among LNTs. The growth mainly occurredby the addition of free lipids to the free ends but sometimes
occurred by the end-to-end fusion of LNTs, as shown in Figure
38b.
The LNTs also interacted with the sideways edge regions of
SLB patches either at their longitudinal sides or at one of the freeends. When interacting at the longitudinal side, the LNTsometimes bent along the sideways edge of the SLB, as shown in
Figure 38c. The longitudinal contact rarely resulted in collapse of

8.8. Other Topics on Protein Dynamics

8.8.1. Cellulase. Cellulose is a predominant component ofplant cell walls,283 and therefore the most abundant biomass andcomponent of feed crops used in the livestock industry.
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Figure 38. HS-AFM imaging of dynamic events occurring in the process of SLB formation on mica. (a) Clips of successive HS-AFM images captured at
1.03 fps for a scan size of 800 × 800 nm2 over 25 min (Z color scale, 25 nm). The white arrows indicate rapidly growing lipid nanotubes with ∼20 nm
height. The light-blue arrow indicates the interaction between a patch of SLB and one end of a lipid nanotube. The arrowheads indicate liposomes with
7.5−30 nm. (b) Successive HS-AFM images showing fusion of two lipid nanotubes at the ends. (c) A HS-AFM image showing bending of a nanotube
interacting with the sideways edge of a SLB patch at the longitudinal side. (d) Clips of successive HS-AFM images showing the interaction of a lipid
nanotube with the edge of SLB patch at one of the ends. The size of the SLB patch is getting smaller, while the length of the nanotube is becoming
shorter.

polypeptide linker rich in serine and threonine residues that are
O-glycosylated.290−292 In general, the CBD is known to enhancecellulose hydrolysis by increasing the eﬀective enzyme
concentration on the surface of the insoluble substrate293
because loss of the CBD causes a signiﬁcant decrease in the
crystalline cellulose degradation rate, but has a less eﬀect on thehydrolysis of soluble or amorphous cellulose. As such, in theinitial step of cellulose hydrolysis, the cellulases adsorb onto thesurface of crystalline cellulose through the CBD. The CD has arelatively long tunnel formed by surface loops with catalyticamino acids extending from the central fold of the CD. A
cellulose chain threads through the tunnel and is hydrolyzed.291,292 Therefore, it has been considered that CBHs
hydrolyze crystalline cellulose chains in a processive manner,
making consecutive cuts without releasing the chain.294,295 The
CBHs are generally categorized into two main types: CBH I

Therefore, the eﬃcient conversion of cellulose to fuels and theimprovement of cellulose digestibility particularly for ruminants
are important issues in agricultural industries. Cellulose is a linear
polymer of β-1,4-linked glucose units.283 Cellulose chains arepacked into ordered arrays to form insoluble microﬁbrils, which
are stabilized by cross-links involving intermolecular hydrogenbonds.284 To degrade cellulose, many organisms produce
cellulases. Cellulase hydrolyzes β-1,4-glucosidic linkages of thepolymer. While all types of cellulases can hydrolyze amorphous
cellulose, only a limited number of cellulases can hydrolyze
crystalline cellulose.285 The enzymes that hydrolyze crystalline
cellulose are called cellobiohydrolases (CBHs) because themajor product of the reaction is cellobiose, a soluble β-1,4glucosidic dimer.286 CBHs consist of two-domain structures: acatalytic domain (CD) and a cellulose-binding domain (CBD)(Figure 39a).287−289 The two domains are connected by a ﬂexible
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Figure 39. Unidirectional processive run of cellulase on crystalline cellulose ﬁber captured by HS-AFM. (a) Schematic of cellulase (cellobiohydrolase:
CBH) with two-domain structure attached onto the surface of crystalline cellulose. (b) Clips of successive AFM images showing processive andunidirectional movement of TrCel7A on crystalline cellulose Iα. Imaging rate, 3.3 fps. Two molecules are marked with the arrows with diﬀerent colors.
(c) Intermittent “go and stop” movement of TrCel7A molecules on cellulose Iα. The movements of seven molecules are shown with the marks with
diﬀerent colors. (d) Velocity distribution of the linear movement of TrCel7A on crystalline cellulose Iα (n = 188). The histogram was approximated by
the combination of two Gaussian distribution curves (blue line) with mean ± SD values of −0.32 ± 3.4 (red) and 7.1 ± 3.9 nm/s (green). (e) Successive
AFM images showing traﬃc jam of TrCel7A bound to crystalline cellulose Iα. Imaging rate, 3.3 fps. The congestion is marked with the arrows.

deciphering these persistent unsolved issues, HS-AFM imaging
was recently carried out on CBHs hydrolyzing crystalline
cellulose ﬁbers.36,57
For HS-AFM observations, crystalline cellulose ﬁbrils were
immobilized on the hydrophobic surface of HOPG. The ﬁbrils
are expected to orient with their hydrophobic surfaces in contact
with the HOPG surface. By the symmetry of crystalline cellulose,
the opposite hydrophobic surfaces appear on the top, being
exposed to the solution. Cellulases bind to these exposedhydrophobic surfaces through CBD’s hydrophobic binding
capability. As such, the use of HOPG has an advantage against
the use of mica. During HS-AFM imaging of cellulose ﬁbrils on
the HOPG surface, a solution containing cellulase was injected
into the observation chamber.
The CBH I (TrCel7A) belonging to GH family 7 (Cel7) issecreted by the industrially important cellulolytic ascomycete

works processively from the reducing end of cellulose, while
CBH II works processively from the nonreducing end ofcellulose.296 The CD of CBH I belongs to the glycosidehydrolase (GH) family 7, and the CD of CBH II belongs to the
GH family 6.
Despite numerous studies, the mechanism of the enzymatic
digestion of crystalline cellulose has remained a major unsolved
problem in agricultural science and technology. For example, it
has been known that the hydrolysis rate decreases rapidly withtime,297 but its mechanism has not yet been understood.
Furthermore, several studies have indicated that CBH I and CBH
II types of enzymes exhibit strong synergy for the hydrolysis ofcellulose,298,299 but the reason for the synergy eﬀect has been
elusive. The main reason for the diﬃculty in understanding theseissues is the lack of experimental methods to monitor the
enzymatic reaction at the single molecular level. With the hope of3164

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Figure 40. HS-AFM imaging of TrCel7A moving on crystalline cellulose IIII. (a) Clips of successive AFM images of TrCel7A on crystalline cellulose IIII.
Imaging rate, 3.3 fps. (b) Schematics of sections of crystalline cellulose Iα and cellulose IIII. (c) Clips of successive HS-AFM images showing ﬁbrillationof crystalline cellulose IIII after congestion of TrCel7A molecules. Imaging rate, 3.3 fps.

fungus Trichoderma reesei. TrCel7A is one of the best-studied
CBHs. Figure 39b shows clips of successive HS-AFM images of
TrCel7A molecules on the crystalline cellulose Iα prepared fromthe cell wall of green algae. It was evident that TrCel7A was
moving unidirectionally and processively on the cellulose surface,making a line. The movement of TrCel7A was observed only on
the hydrophobic top surface of the cellulose crystal.
Figure 39c shows displacements of seven TrCel7A molecules
as a function of time. In this analysis, the center of mass of eachmolecule was tracked over consecutive images (section A.5 inAppendix). Interestingly, the movement of molecules sometimes
halted or was slowed (the open green and closed red plots in
Figure 39c) probably due to the presence of obstructions on the
crystalline cellulose surface, whereas some molecules moved
continuously without halt. A histogram of the measured velocity
for 188 molecules comprised two Gaussian distributions withaverage velocities of −0.32 ± 3.4 and 7.1 ± 3.9 nm/s (Figure39d). This may be consistent with the view that TrCel7A has twomodes of adsorption on a cellulose surface: the productive
adsorption mode in which both CD and CBD contribute to thebinding and the nonproductive adsorption mode in which the
enzyme binds only via the CBD.293 The overall average velocity
for the seven molecules was 5.3 ± 4.9 nm/s. From this velocity,
the hydrolysis rate was estimated to be 5 s−1, taking into accountthat one hydrolysis drives the movement by 1 nm, which is thelength of cellobiose. Surprisingly, this hydrolysis rate is 500 times
larger than that (0.01 s−1) estimated biochemically.300 This large
discrepancy is probably because in the biochemical estimation, it
has been assumed that all molecules productively work on thecellulose surface. However, in reality the number of reduced ends
available for TrCel7A binding is limited, and moreover the
movement of molecules is often hampered by traﬃc jams as seenin Figure 39e.

The fabric form of cellulose Iα can be converted to cellulose IIII
polymorph by supercritical ammonia treatment. This conversion
dramatically enhances the cellulose hydrolysis by TrCel7A.301 A
biochemical analysis has indicated that TrCel7A has a higher
aﬃnity for cellulose IIII than for cellulose Iα, but its maximum
hydrolysis rate is only slightly higher than cellulose Iα.301 Figure
40a shows clips of successive HS-AFM images of TrCel7A
molecules processively moving on a cellulose IIII surface. The
number of molecules on the cellulose IIII surface appeared larger
than that on the cellulose Iα surface, as shown in Figure 40a, but
the velocity of the molecules was similar for both surfaces. In thecase of cellulose Iα, the molecules moved only along limited lanes,
whereas, on cellulose IIII, TrCel7A molecules moved over almost
the whole surface. This explains the biochemically obtained
aﬃnity diﬀerence between cellulose Iα and IIII. It also agrees withprevious studies284,302 showing that cellulose Iα has a hydrophobic surface at (110) surface (Figure 40b, the left schematic),
whereas cellulose IIII has a hydrophobic surface not only at (110)
surface but also at (100) surface (Figure 40b, the right
schematic). Therefore, cellulose IIII has more lanes available
for cellulase binding and movement than cellulose Iα.
When a molecule halted, many following molecules stacked
behind it on the cellulose IIII surface, causing a traﬃc jam.
However, after several additional molecules were blocked at thisplace, the molecules restarted to move. In this occasion, acellulose bundle was peeled oﬀ from the crystalline cellulose IIII,
as shown in Figure 40c. These observations suggested thefollowing sequential events to occur: (i) an obstruction on thesurface of crystalline cellulose (and hence a rough surface) is
formed during the digestion, (ii) a single TrCel7A molecule isunable to overcome the obstacle by its own generating force andis therefore halted, and (iii) the accumulated molecules at the
obstacle work together with increased force to get the obstacleout of the way (i.e., pealing of a cellulose bundle), resulting in
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Figure 41. Synergistic hydrolysis by TrCel6A and TrCel7A visualized by HS-AFM. (a) Synergy, measured biochemically, between TrCel6A and
TrCel7A in cellobiose production from cellulose IIII. The green and red plots show the rates of cellobiose production by TrCel6A alone and TrCel7A
alone, 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) Clips of successive AFM images of cellulose IIII after injection of TrCel6A at 0 s. The degradation of cellulose
was not discernible even at 8 min. (c) Clips of successive AFM images after the additional addition of TrCel7A at 0 s (at 8.4 min after the initial additionof TrCel6A). The cellulose was rapidly degraded and almost disappeared at 8 min. Imaging rate, 3.3 fps for (b) and (c).

TrCel6A alone (Figure 41c). Eventually, the cellulose crystals
almost disappeared at ∼4 min after the addition of TrCel7A. The
observed synergy was independent of the order of the addition of
TrCel6A and TrCel7A. A very similar synergy was also observed
when both TrCel6A and TrCel7A were added simultaneously.
The HS-AFM observation showed that TrCel6A was randomly
bound to the cellulose ﬁbril but not the end of the ﬁbril, and the
movement was less processive. Therefore, the synergy observed
could be rather called the endo−exo synergy,306 although
TrCel6A has generally been deﬁned as an exoglucanase. One
possible mechanism of the synergetic degradation is that the twoloops forming the active site tunnel of TrCel6A can open,
allowing generation of nicks in the middle of crystalline cellulose,
and that these nicks become starting and ending points for the
TrCel7A activity.
8.8.2. Intrinsically Disordered Proteins. It has recently
been recognized that a new class of functional proteins exist,
which lack a well-deﬁned ordered structure in the entire length orin large domains under native conditions.307−310 These proteinsare generically called intrinsically disordered proteins (IDPs).
According to computational predictions based on genomes,∼30% of eukaryotic proteins are mostly disordered, and morethan 50% eukaryotic proteins have long disordered regions
consisting of more than 40 consecutive amino acid residues.311
IDPs contrast strikingly with a classical structure−function
paradigm of proteins. In a classical view, the structure of all
functional proteins must be well ordered, and forming deﬁned
tertiary structures is considered to be a unique strategy forestablishing speciﬁc interactions with ligands and partner
proteins (lock-and-key model); as such, proteins with largely
disordered structures have been considered to be denatured anddysfunctional, until the discovery of a large group of IDPs. The
thin and ﬂexible intrinsically disordered regions (IDRs)
contained in IDPs take a structural ensemble consisting of alarge number of random conformers. Upon binding to a target
molecule, the disordered structure is converted to a deﬁnedordered structure or remains partially or even fully disordered.
When folded into an ordered structure, this folding is termed

restarting linear movement. The traﬃc congestion is very likely
to be the mechanism underlying the rapid reduction with time ofcellulose hydrolysis rate. The distance between neighboring
traﬃc lanes is smaller than the size of a TrCel7A molecule.
Therefore, the enzyme molecules moving along neighboringlanes have a large chance to collide with each other. When a
TrCel7A molecule is hydrolyzing one lane of cellulose, it is likelythat the molecule partially peels lanes adjacent to the lane that themolecule is hydrolyzing. This peeling results in the production ofrough surfaces and obstacles. Note that before the addition of
TrCel7A, the surface of crystalline cellulose ﬁbers was smooth.
It is known that the mixture of two major CBHs, TrCel7A and
TrCel6A, synergistically enhances the hydrolysis of cellulose IIII
(Figure 41a). This eﬀect has been called the exo−exo synergy.300
Either of these two enzymes has an active site located in a tunnel
that passes through the whole CD. The active site tunnel of
TrCel6A is formed by two surface loops and is about 20 Å long,whereas that of TrCel7A is approximately 50 Å long, comprising
six loops.290−292,303 Previous transmission electron microscopic
observations of crystal cellulose partially hydrolyzed by both
TrCel6A and TrCel7A suggested that the enzyme cocktail
degraded the cellulose from both ends of the crystal. TrCel7A
degrades the crystal from the reducing end, leading to ﬁbrillation,
thinning of the crystal, or narrowing of the crystal end, whereas
TrCel6A hydrolyzes the crystal from the nonreducing end, lessprocessively than TrCel7A, thereby sharpening the crystal
tip.304,305 It is however diﬃcult to explain this type of synergistic
action based on their exoenzymatic activities. To clarify this issue,
the hydrolysis process of crystalline cellulose IIII in the presenceof both TrCel6A and TrCel7A was imaged.
Figure 41b shows clips of successive HS-AFM images taken forcellulose IIII in the presence of TrCel6A alone. The appearanceof the cellulose crystals did not signiﬁcantly change during
incubation over 8 min, even though many enzyme molecules
were observed on the cellulose surfaces. Upon addition of
TrCel7A at 8.3 min, the enzyme molecules started moving frommany points on the cellulose surfaces, and the degradation of
crystalline cellulose was dramatically faster than the case with
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Figure 42. Intrinsically disordered dFACT protein and eﬀect of phosphorylation on the structure visualized by HS-AFM. (a,b) Clips of successive HSAFM images of phosphorylated dFACT-WT (a) and dephosphorylated dFACT-10SA (b). Imaging rate, ∼14 fps; scan area, 100 × 100 nm2 with 80 × 80
pixels; Z-scale, 4.0 nm. The observed molecular features of dFACT are schematized according to the deﬁnitions in (c). These schematics were drawn
freehand by tracing the AFM images by visual estimation. (c) Schematics representing the characteristics of dFACT observed by HS-AFM (upper
schematic) and its side-view (lower schematic). Gray-colored ellipses and black-colored thick solid lines represent the globular domains (GD) and the
intrinsically disordered regions, respectively. (d) Histograms of distance D1−2 indicated in (c) measured for dFACT-WT (upper) and dFACT-10SA(bottom). (e) Histograms of height H2 indicated in (c) measured for dFACT-WT (upper) and dFACT-10SA (bottom). The histograms appear to have
two peaks at 1.6−1.7 and ∼1.1 nm, which correspond to HMG associated with an IDR and HMG alone, respectively.

“coupled folding”.312 Certain IDPs can promiscuously interact
with several target proteins, and the corresponding coupledfolding results in diﬀerent ordered structures adapted to thetarget proteins.313,314 This adaptation furnishes such IDPs withmultifunctionality. IDPs are often involved in transcription,translation, and cellular signal transduction. The adaptability,
multifunctionality, and promiscuity are IDPs’ attributes essential
for functioning in diverse biological processes as regulators.
Despite increasing recognition of the functional importance of
IDPs, the analysis of their disordered structures has been diﬃcult.
The local structure of IDPs with molecular weights less than ∼50
kDa (usually less than ∼20 kDa) can be determined by NMR, but
the overall random structure cannot be revealed by this method.

Electron microscopy cannot visualize the thin and random
structures. X-ray crystallography is entirely useless for IDPs
because IDPs cannot be crystallized. Except for computational
predictions, there have been no eﬀective means to identify theoverall random structure of IDRs until the advent of HS-AFM. It
was recently demonstrated that HS-AFM could visualize the thin
and random structures of IDRs, using Drosophila melanogaster
FACT (facilitates chromatin transcription) protein (dFACT)
overexpressed in E. coli, as a test sample.32 A chromatin
remodeler, FACT, is a heterodimer complex consisting of SSRP1and SPT16 subunits.315−317 HS-AFM images of dFACT showed
two wiggling tail-like segments with diﬀerent lengths that
protruded from the main body of dFACT; their contour lengths
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dFACT inhibits the formation of a stable complex of dFACT andnucleosomal DNA.326 The phosphorylated sites reside on an
acidic ID segment adjacent to an HMG domain-ﬂanking basic ID
segment of the SSRP1 subunit. The phosphorylation increases
the number of negative charges in the acidic segment. Therefore,
the DNA binding inhibition is conceived to be caused either by
an increased repulsive interaction between the acidic ID segment
and DNA or by the association between the acidic and basic ID
segments. To elucidate the inhibitory mechanism, HS-AFM
imaging was carried out for dFACT (from which the shorter IDR(888−1044) and the N-terminal amino peptidase like domain(1−401) in the SPT16 subunit were deleted) expressed in Sf9
insect cells, in fully phosphorylated (WT) and nonphosphorylated forms.63 The nonphosphorylated form was obtained bysite-directed mutagenesis to Ala at 10 phosphorylation sites(Ser/Thr) contained in the acidic ID segment of the SSRP1
subunit.
HS-AFM images captured at 15 fps showed a similar structureon the whole for phosphorylated WT dFACT (dFACT-WT)
and dephosphorylated mutant (dFACT-10SA) (Figure 42a,b); a
large globular domain (GD1) was connected to a tail-likestructure (i.e., IDR of SSRP1), as previously observed.32
However, in both constructs, two small globular domains,termed GD2 and GD3, were found on the IDR in the middle andat the distal end of the IDR, respectively (Figure 42a,b).
Therefore, the entire IDR was divided into two (IDR1−2 andIDR2−3); IDR1−2 connects GD1 and GD2, while IDR2−3 connects
GD2 and GD3 (Figure 42c). GD3 located at the distal end of the
IDR was consistently observed, without any appearance
diﬀerences between the two constructs, whereas GD2 locatedin the middle of the IDR appeared temporally and vanished, with
diﬀerent lifetimes between the two constructs; the GD2 ofdFACT-WT exhibited a longer lifetime than that of dFACT10SA. Moreover, the length of the entire IDR appeared to be
shorter in dFACT-WT than in dFACT-10SA. This length
diﬀerence was reﬂected from the much shorter length of IDR1−2
in dFACT-WT. The length of IDR2−3 was similar between thetwo constructs. To express these structural features quantitatively, the parameters of end-to-end distance and height shown in
Figure 42c were obtained for both constructs by analyzing manysuccessive HS-AFM images captured for three molecules of eachconstruct. The most distinct diﬀerence between the twoconstructs occurred in the end-to-end distance D1−2 between
GD1 and GD2 (Figure 42d); the distance was shorter in dFACTWT by ∼2 nm. The next distinct diﬀerence occurred in theheight H2 of GD2 (Figure 42e); dFACT-WT predominantly tooka high globular state (1.6−1.7 nm), while dFACT-10SA took alow state (∼1.1 nm). However, these structural states were notstatic but dynamic. The shorter GD1−GD2 distance state and thelower GD2 states were taken even by dephosphorylated dFACT10SA. Moreover, the position of GD2 was consistent with theposition of the HMG domain, the height ∼1.1 nm corresponded
to the size of HMG (∼1 nm), and the height of the IDRs was
0.4−0.6 nm. Thus, the following conclusions were derived:63 (i)the acidic segment of SSRP1’s IDR electrostatically interacts withboth the region containing the basic segment of SSRP1’s IDR
and the HMG domain to evoke partial folding of the IDR, (ii) the
folded and unfolded states are in dynamic equilibrium, (iii)
extensive phosphorylation of the acidic segment of the IDR shifts
the equilibrium toward the folded state, and (iv) this structural
bias to the folded state blocks the binding of nucleosomal DNA
to the region containing the basic IDR segment and the HMG
domain.

were 17.5 and 27.5 nm on average. HS-AFM images of mutants,
M1 and M2, in which either predicted IDRs were deleted,
conﬁrmed that these wiggling segments are IDRs, and that thelonger ID region belongs to the SSRP1 subunit while the shorterone belongs to the SPT16 subunit. Note that it is hard to visualize
IDRs using conventional AFM even when the sample is dried in
air and immobilized onto a substrate surface because thin IDRs(∼1 nm thick) become thinner under this condition.
Interestingly, both IDRs showed a similar macroscopic
ﬂexibility (or stiﬀness). The stiﬀness of a polymer chain can bequantiﬁed by the persistence length p of the chain, that is, thelength over which the average correlation in the tangential
direction decays by 1/e. In 2D, the mean-square point-to-point
distance ⟨r2⟩ of a uniform chain is given byr 2(l) 2D = 4pl × [1 − (2p /l) × (1 − exp( −l /2p))](6)

where l is the contour length between two points on the chain.318
In the AFM images obtained for the tail-like structures ofdFACT, their polypeptide chains were not resolved. Therefore,
the persistence lengths that can be obtained from the images arethose for overall (macroscopic) string structures. The analysis of
AFM images of the ID regions with eq 6 gave a persistence lengthof 11−12 nm for both ID regions. Microscopic persistence length
Lp can be approximately determined on the basis of anassumption that the polypeptide chain length L is a multiple ofthe number of amino acids and the likely average distance
between adjacent amino acid residues (∼0.34 nm) and on therelationship:

R2 = 4LpL(7)

where R is the end-to-end distance. This equation holds,
provided that L is much longer than Lp. Note that unlike themacroscopic persistence length, the microscopic persistence
length does not correspond to the stiﬀness of a polypeptide chain
but represents the degree of its folding; a shorter microscopic
persistence length reﬂects a higher degree of folding. The
microscopic persistence length was ∼1.1 nm for both ID regions(slightly shorter with the longer ID region, likely due to thepresence of the ordered high-mobility group (HMG) domain in
the longer ID region within the SSRP1 subunit). Microscopic
persistence lengths of structured and randomly coiled proteins
have previously been estimated by the measurement of force−
extension curves using AFM.319−321 The microscopic persistence
length of the PEVK region (186 amino acid residues), which is
responsible for the elastic property of titin and thought to form arandom coil, is reported to show a wide range of Lp (0.4−2.5 nm;
0.93 nm on average).320 On the other hand, for proteins withordered structures, the reported values for the microscopic
persistence length are within a range of 0.36−0.5 nm.319,321
Thus, the long microscopic persistence length (∼1.1 nm) of the
IDRs of dFACT protein is a ﬁngerprint of their very loose
structures. A recent HS-AFM study on other proteins’ IDRs has
further conﬁrmed long persistence lengths.322 The nature ofhigher looseness of IDRs than entropically folded random coils
reﬂects the electrostatic repulsion between charged amino acids
rich in IDRs, which reduces the entropy-driven folding.
The interaction of IDRs with target proteins and DNA can be
modulated by post-translational modiﬁcations, including phosphorylation, acetylation, methylation, sumoylation, and ubiquitination.323−325 A recent study has shown that extensive
phosphorylation of the IDR belonging to the SSRP1 subunit of
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Figure 43. Migration of monovacancy defects in streptavidin 2D crystal captured by HS-AFM. (a) Schematic of streptavidin arrays in a C222 crystal.
Unit lattice vectors are indicated: the a-axis includes rows of contiguous biotin-bound subunits, while the b-axis includes rows of contiguous biotinunbound subunits. (b) High-speed AFM images of streptavidin 2D crystal and monovacancy defects therein. The monovacancy defects are enclosed bydashed squares and circles. The directions of the lattice vectors of the crystal are also indicated. Imaging rate, 2 fps; scan area, 150 × 150 nm2; the numberof pixels, 200 × 200. (c) Schematic showing a streptavidin molecule being transferred to the neighboring monovacancy defect along the a-axis. (d)Schematic showing a streptavidin molecule being transferred to the neighboring monovacancy defect along the b-axis. The “u−u bonds” are shown inred, while the “b−b bonds” are shown in green (c and d). (e) Clips of successive HS-AFM images showing fusion of two monovacancy defects into a
divacancy defect. Imaging rate, 2 fps; scan area, 140 × 140 nm2; pixels, 200 × 200.

As demonstrated above, high-speed AFM is powerful in
identifying and characterizing IDRs of proteins. In conventional
methods, even the identiﬁcation of IDRs is diﬃcult and timeconsuming. Moreover, high-speed AFM can detect order−
disorder transitions of IDRs that occur naturally with free IDRsor are biased to the ordered state either by post-translational
modiﬁcation or by interaction with target molecules. In the near
future, high-speed AFM will visualize the dynamic processes inwhich IDRs are folded into ordered structures upon interaction
with their target molecules and drugs.
8.8.3. Dynamics of Vacancy Defects in 2D Crystals ofProteins. In 2D crowded conditions, protein molecules
frequently interact with each other even under a very weak
attractive force between the molecules, which sometimes resultsin the formation of 2D crystals. The 2D crystallization of proteins
occurs naturally as seen in bR in the purple membrane and AQP0
in the intercellular junctional microdomains, as well as artiﬁciallyas seen in electron crystallographic studies.100,327,328 Whenvacancy defects are present in the 2D crystal of a protein, thedefects are very likely to move in the crystal because proteins
molecules facing a vacancy defect lose some of intermolecularbonds, and therefore tend to move into the defect, leaving a newvacancy defect behind. In fact, this phenomenon was observed by
HS-AFM in the orthorhombic C222 crystals of streptavidin
formed on biotin-containing SLBs.34 Note that the 2D crystal has
P2 symmetry,38 but here the notation C222 for the symmetrygroup follows a previous report.328
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Reviewmultivacancy defects increased for both crystal axes, except forthe case of divacancy defects. For example, Da and Db of tetravacancy defects increased ∼3.5 times and ∼2.5 times,
respectively, as compared to those of monovacancy defects.
The higher mobility of larger point defects increases theirprobability to encounter other point defects to form larger point
defects with further higher mobilities. During the crystal growthin the presence of free streptavidin in the bulk solution, thisacceleration eﬀect should promote the removal of point defects
from the crystalline regions. This would be one of themechanisms for defect-free crystal formation not only in 2D
crystals but also in 3D crystals. This issue should be examined in
the future by imaging the crystal growth in the presence of freestreptavidin in the bulk solution. The mechanism underlying thelarger mobility of larger multivacancy defects is at a glance
considered to be contrary to the fact that many intermolecularbonds have to be broken for a large multivacancy defect to move.
However, this seemingly peculiar tendency was rationallyinterpreted by the consideration of two factors.34 One factor isthe lower stability of the crystal regions adjacent to a larger point
defect in comparison with the crystal regions adjacent to asmaller defect. Another is that there are multiple paths along
which a multivacancy defect migrates to one of their nearestneighbor positions.

The lipid composition used was DOPC, DOPS, and biotincap-DOPE (7:2:1 w/w). The C222 crystal of streptavidin withlattice constants a = 5.9 nm, b = 5.9 nm, and γ = 90° (Figure 43a)
was prepared using a relatively high ionic buﬀer solution
containing 2 mM CaCl2 (pH 7.4). Monovacancy defects in thestreptavidin 2D crystals were produced by increasing the tapping
force exerting from the oscillating tip to the sample. The imaging
was carried out at 2 fps over 150 × 150 nm2 scan areas.
Monovacancy defects were mobile, with an anisotropic feature
with respect to the two axes of the crystalline lattice: larger
mobility along the b axis than the a axis (Figure 43b). The a axis
includes the rows of contiguous biotin-bound subunits, whereas
the b axis includes the rows of contiguous biotin-unboundsubunits (Figure 43a). The linear relationship between the
MSDs and time intervals gave the diﬀusion rate constants of Da =20.5 nm2/s for the a-axis component of diﬀusion and Db = 48.8
nm2/s for the b-axis component of diﬀusion. This anisotropy inthe mobility (Db > Da) arises from a free energy diﬀerence
between the biotin-bound subunit−subunit interaction andbiotin-unbound subunit−subunit interaction. When a streptavidin molecule adjacent to a monovacancy defect moves to thedefect site along the a axis, two intermolecular bonds between
biotin-unbound subunits (“u−u bond”) and one intermolecularbond between biotin-bound subunits (“b−b bond”) are broken(see Figure 43c). On the other hand, when it moves to the defectsite along the b axis, one u−u bond and two b−b bonds arebroken (see Figure 43d). Therefore, the diﬀerence in theactivation energies Ea and Eb for the step movement of amonovacancy defect along the respective a and b axes simply
corresponds to the diﬀerence between the free energy changes
Gu−u and Gb−b produced by the formation of the respective u−ubond and b−b bond: Eb − Ea = Gu−u − Gb−b. Therefore, theobserved relationship Db > Da indicates Gu−u < Gb−b; that is, theu−u bond aﬃnity is higher than that of the b−b bond. The ratioof the two diﬀusion rate constants (Db/Da) can be expressed by
Db/Da = exp[−(Eb − Ea)/(kBT)]. Thus, from the observed valueof Db/Da = 2.4, the free energy diﬀerence Gu−u − Gb−b was
estimated to be approximately −0.88 kBT (T ≈ 300 K), whichcorresponds to −0.52 kcal/mol.
This free energy diﬀerence must aﬀect the 2D crystal growth,
resulting in asymmetric crystal growth rate along perpendicularaxes, hence in an anisotropic crystal morphology.328 In fact, thestreptavidin C222 crystals formed under biotinylated lipid layers
are known to have X-, H-, and rectangle-shaped morphologies.329,330 The anisotropy estimated from the aspect ratios of theC222 crystals has been reported to be 2−3 (about 2 at neutralpH),331 coinciding with the diﬀusion constant ratio (Db/Da =2.4). Supposing that the aspect ratio of a crystal is proportional to
the ratio of the free energies of attractive interactions that occur
along the crystal axes (Wulﬀ’s rule),332 Gu−u/Gb−b is approximately 2. This relationship and Gu−u − Gb−b ≈ −0.88 kBT resultin Gb−b ≈ −0.88 kBT and Gu−u ≈ −1.76 kBT. The small freeenergy Gb−b ≈ −0.88 kBT suggests that the C222 crystal could be
disrupted when all of the biotin binding sites are bound to biotin
because the number of b−b bonds per molecule in the C222
crystal lattice is two; hence the association free energy permolecule amounts to only −1.8 kBT. This was conﬁrmed by theobservation that the ordered lattices were immediately disrupted
when the crystals were exposed to free biotin in the bulk
solution.34
The monovacancy defects sometimes fused into a divacancy
defect (Figure 43e). Through several fusions, larger multivacancy
defects were formed.34 As the size increased, the diﬀusivity of

9. DYNAMIC IMAGING OF LIVE CELLSOne of the prospective high-impact applications of HS-AFM isreal-time monitoring of dynamic phenomena occurring in live
cells and isolated intracellular organelles, including morphological changes of cells, ﬁlopodium formation and retraction,
phagocytic and endocytic processes, fusion and ﬁssion of
mitochondria, synaptic formation in neurons, and enlargementof spines associated with memory and learning. Furthermore,
HS-AFM would oﬀer a unique opportunity to visualize dynamic
events of membrane proteins occurring on the surfaces of live
cells and isolated intracellular organelles. However, imaging ofthese phenomena has been one of the largest challenges for HSAFM. The larger size and extremely soft surfaces of these objects
require a large scan range and high sensitivity for tip−sample
interaction detection, respectively. Recently, fast scanners
covering an XY scan range of up to 10 × 10 μm2 and even up
to 46 × 46 μm2 have been developed.68,70,333 Despite theirnoncompact structures due to the use of large piezo actuators aswell as displacement magniﬁcation mechanisms, the high-speed
scan performance is only slightly inferior to the fast scanners witha much smaller maximum scan range. Using these wide-areascanners, HS-AFM imaging of live cells has just recently been
embarked on. Thus far, the number of successful studies is still
very limited, and even these successful studies do not yet fully
solve biological questions of importance. However, it isworthwhile to look at the current state of live cell imaging by
HS-AFM to know its potential as well as techniques that should
be further developed or combined with the current HS-AFM tofacilitate this endeavor.
9.1. Bacterial Cell Surfaces

9.1.1. Bacteriolysis of Bacillus subtilis Subjected to
Lysozyme. Because of the nonpathogenicity and amenability togenetic manipulation, Bacillus (B.) subtilis has been widely used
as a model system for investigating the roles of autolysins (i.e.,bacteriolysis caused by digestion of the cell wall peptidoglycan ofbacteria by enzymes that are produced from the bacteria
themselves) in the cell wall metabolism of endospore-forming
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Figure 44. Process of lysozyme-induced bacteriolysis captured by HS-AFM. (a) Time-lapse images of bacteriolytic process of B. subtilis subjected tolysozyme. Lysozyme was injected at t = 240 s. Scan range, 5 × 5 μm2; imaging rate, 0.05 fps (20 s/frame); pixels, 256 × 256. (b) Time course of RMS
roughness of the outer surface of B. subtilis subjected to lysozyme. Lysozyme was injected at t = 32 s. Solid line shows the result of best ﬁtting (timeconstant, τ = 36.8 s) by a single exponential function for the time course between t = 112 s and t = 300 s. (c) Corresponding clips of successive AFM
images of the outer surface of B. subtilis at diﬀerent time lapses. Values of RMS roughness were calculated for the area encircled by the white broken line
shown in the image at t = 86 s.

After t = 320 s, the surface morphology gradually became varied,
and wrinkle structures perpendicular to the long cell axis
appeared. At the same time, the cell gradually swelled (t = 1060,
1460, and 1580 s), indicating lowered resistance to osmoticpressure due to the degradation of the hard cell wall. Finally, at t =1720 s, the cell exploded by osmolysis, resulting in partial
exposure of the intracellular contents. The detailed morphological change induced by lysozyme at higher ﬁnal concentrationof 200 μM was also observed at a faster imaging rate 0.5 fps for a 2
× 1.5 μm2 area with 120 × 90 pixels (Figure 44c). The cell surface
roughness increased rapidly with a time lag of ∼90 s after the
injection of lysozyme at t = 32 s (Figure 44b). Here, the surface
roughness was expressed by the root-mean-square (RMS)
roughness of the area indicated by the white rectangle in Figure
44c. The roughness was almost constant (∼93 nm) before theinjection of lysozyme, whereas it increased to ∼125 nm after theGram-positive bacteria, and the structure of peptidoglycan andspores.334,335 It is well-known that B. subtilis is also susceptible tothe antibacterial action of lysozyme, which hydrolyzes 1,4-beta
linkages between N-acetyl glucosamine and N-acetyl muramic
acid, leading to degradation of peptidoglycan in the cell wall.334
This degradation process of peptidoglycan induced by lysozyme
was captured by HS-AFM.70
The B. subtilis bacteria were immobilized on a glass stage
coated with poly-L-lysine. HS-AFM imaging was conducted in a
culture medium to maintain bacteria alive. Figure 44a shows
time-lapse images of the bacterium before and after the additionof lysozyme (imaging rate, 0.05 fps; scan size 5 × 5 μm2 with 256
× 256 pixels). Before the injection of lysozyme (t = 0 s), the cellsurface appeared smooth. At t = 240 s, lysozyme (ﬁnalconcentration, 80 μM) was injected into the observation
solution, and therefore the imaging was somewhat disturbed.
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injection. The increase in the surface roughness was well ﬁtted bya single exponential function with a time constant of τ = 36.8 s, as
indicated by the solid line in Figure 44b. High-resolution imaging
on the bacterial cell surface was also carried out before and after
the addition of lysozyme, at an imaging rate 0.5 fps for a 500 ×
500 nm2 area with 200 × 200 pixels (Figure 45). Fine structures

increase of the cell induced by CM15 is consistent with anelectron microscopy study.337 Interestingly, the cell “1” started
changing within 13 s after the addition of CM15 and the change
was completed in ∼60 s, whereas the cell “2” did not startchanging until ∼80 s, and the surface alterations were notcompleted in ∼120 s, suggesting that the two cells are not in thesame stage in the cell cycle, or revealing individuality concerning
their sensitivity to CM15. Correlation between the degree of
surface roughness increase and the cell death was conﬁrmed by
HS-AFM-combined ﬂuorescence microscopy using a ﬂuorescentindicator of cell viability.
Time evolution of the surface roughness along the long axis ofseveral bacteria (assessed by the phase signal) was analyzed toquantify the kinetics of the change. Figure 46b shows
representative cross sections of a single bacterium surface at
diﬀerent incubation stages. The marked increase of the surface
roughness appeared at ∼80 s after the addition of CM15. Figure
46c shows the normalized RMS roughness for nine cells as afunction of incubation time. The onset time of the roughnesschange depended largely on individual cells, ranging from 40 s(bacterium 7) to more than 4 min (bacterium 2), with an averageof 155 ± 89 s. The time for each bacterium to complete thechange (smooth to corrugated) was more consistent; 50% of the
maximum roughness increase occurred within 52 ± 16 s after the
onset of roughness change.
On the basis of these results, it was proposed that sterilizationof bacteria by CM15 is a two-stage process, that is, incubationand execution phases, and that the bulk sterilization rate isdominated by the incubation phase, rather than the execution
phase.
9.1.3. Imaging of Single Molecules on Live Bacteria
Surfaces. Dynamics of membrane proteins in isolated native
membrane fragments and those reconstituted in synthetic
membranes can be visualized at high resolution with HSAFM.35,37,41−44,49,50,59,61 However, it would be more ideal tovisualize membrane proteins in live cell membranes at the singlemolecule level, using HS-AFM. Little is known about dynamic
actions of membrane proteins in real environments, where
membrane proteins interact with other proteins and areinﬂuenced by membrane potential and ionic species partitioned
across the membrane. Therefore, such in situ dynamic imaging
will certainly expand the usefulness of HS-AFM in biologicalstudies. Previously, visualization of the molecular organization onlive cell surfaces was performed for the hexagonal S-layer ofCorynebacterium glutamicum,338 the peptidoglycan organizationof Lactococcus lactis,339 and the germinating spores of B.
atrophaeus.340 However, only a few attempts of HS-AFM
visualization have thus far been made on molecules on live cellsurfaces.69,71 The ﬁrst attempt was carried out for a bacterium,69
which will be described below.
The magnetotactic spirillum Magnetospirillum (M.) magneticum AMB-1 was used as a test sample.69 The cells werechemically immobilized on mica, which had been covered with
poly-L-lysine and then treated with glutaraldehyde. The viabilityof the immobilized cells was assessed using a ﬂuorescentindicator of bacterial viability. Most of the cells were alive for atleast 1 h after the immobilization. Figure 47a and b shows the cellsurface captured for scan ranges of 150 × 150 nm2 (imaging rate,
0.5 fps) and 100 × 100 nm2 (imaging rate, 1.43 fps), respectively.
The cell surface was coarse and covered all over with regularly
arranged particles with similar size (Figure 47a). The magniﬁed
image (Figure 47b) showed reticulate structures with a dent ineach central region, which were irregularly moving, indicating

Figure 45. Bacterial surface change by lysozyme captured by HS-AFM.
(a,b) Magniﬁed views of outer surface of B. subtilis (a) before and (b)after the injection of lysozyme. Scan range, 500 × 500 nm2; imaging rate,
0.5 fps; pixels, 256 × 256.

on the cell wall were clearly visible, as shown in Figure 45a (t = 0s). Soon after the injection of lysozyme, the ﬁne structure on thecell surface became blurry due to the degradation of the cell wall,
and a wrinkled structure with a periodicity of about 300 nm
appeared (Figure 45b).
9.1.2. Surface Change of E. coli Subjected to
Antimicrobial Peptide. Antimicrobial peptides (AMPs)
produced by a vast array of organisms comprise a diverse classof natural antibiotics, which are considered to be a major part ofthe innate immune system of a host (a ﬁrst line of defense against
pathogenic invasion by microbes). The peptides display a wide
range of antimicrobial activities against bacteria, viruses, andfungi. The binding of these peptides to a bacterium is followed by
the disruption of cytoplasmic membranes, leading to bacterialcell death.336 Because antimicrobial peptides have a potential astherapeutic alternatives to current antibiotics, numerous studies
have been conducted to understand how AMPs interact with andact on cell membranes. However, there have been no direct
studies to elucidate the early stage kinetics of the membrane
disruption in live cells at high spatiotemporal resolution. Fantnerand colleagues studied this issue by imaging surface morphological changes of bacterial cells subjected to an AMP using a HSAFM instrument and small cantilevers developed by them.68
E. coli cells were immobilized on a cover slide coated with polyL-lysine and then imaged in water. CM15, which is a linear,synthetic hybrid AMP composed of the ﬁrst seven residues of thececropin A and residues 2−9 of the bee venom peptide mellitin,
was added to the HS-AFM ﬂuid cell to a ﬁnal concentration of 50
μg/mL using a ﬂow-through ﬂuid exchange system with a dual
syringe pump. HS-AFM images were captured at 0.077 fps for 3
× 3 μm2 areas. For the HS-AFM observation, the phase of thecantilever oscillation relative to the excitation signal was used as
the imaging signal (rather than the topography) because the ﬁne
structures of the bacterial surface were more conspicuous thanthe height image. This is because the topography contains a largeheight background against small surface corrugations of thebacteria surface. Figure 46a shows time-lapse phase images of thesurfaces of E. coli cells after the addition of CM15 at t = −6 s. The
surface of the cell marked with “1” changed from smooth torough with time due to the CM15 action. The surface roughness
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Figure 46. Process of antimicrobial peptide-induced bacteriolysis captured by HS-AFM. (a) Clips of successive HS-AFM images of E. coli subjected to
CM15 antimicrobial peptide. CM15 was injected at t = −6 s. Imaging rate, 0.08 fps (12.5 s/frame); pixels, 1024 × 256. (b) Cross sections along the longaxis of a bacterium showing the time progression of the surface variation. (c) Averaged surface variations of individual bacterial cells as a function of timeafter injection of CM15. Reprinted with permission from ref 68. Copyright 2010 Nature Publishing Group.

imaged with the same tip at the same or lower loading force. Thechange of the reticulate structure during the dissection is shownin Figure 48b. By the increased loading force during imaging(frames 3−7), the reticulate structure was disrupted, resulting inappearance of diﬀusing triangular-shaped molecules in themembrane patch (frame 13 in Figure 48b). Most of themolecules showed a trimeric form (Figure 48c). To identify thetrimeric protein, proteins contained in the outer membrane
fraction were analyzed by sodium dodecyl sulfate-polyacrylamide
gel electrophoresis and then by matrix-assisted laser desorption/
ionization time-of-ﬂight tandem mass spectrometry of thetrypsin-digested gel slices. The most abundant protein in theouter membrane fraction was a 40-kDa protein (∼80% of thetotal protein content), which was identiﬁed as a porin
homologue (amb0025: Msp1). Therefore, it was concluded
that the major fraction of the molecules densely covering theouter membrane of M. magneticum AMB-1 is porin trimers.

that the overall structure is loosely packed. Many particles wereseen on the embankments surrounding dents (Figure 47b,
arrows). The central positions of 23 dents in the reticulate
structure were tracked. The trajectories of four of the 23 dents areshown in Figure 47c. Despite the highly crowded condition, their
MSDs were approximately linear as a function time (Figure 47d,
inset), from which diﬀusion constants were estimated. Figure
47d shows a histogram of the diﬀusion constants, which could beﬁtted with a log-normal distribution, resulting in a mean diﬀusion
constant of 3.2 ± 0.4 nm2/s.
M. magneticum AMB-1 belongs to α-proteobacteria, and is
enveloped by the outer and cytoplasmic membranes. To identify
the composition of the reticulate structure, the outer membraneof M. magneticum AMB-1 was isolated, placed on a bare mica
surface, and observed by HS-AFM. The HS-AFM images showeda reticulate structure very similar to that observed on live cellsurfaces (Figure 48a). As previously demonstrated,38,71,341,342
the cantilever tip can be used to manipulate and dissect samples
by increasing the loading force, and the treated samples can be
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Figure 47. Molecular imaging of bacterial outer surface by HS-AFM. (a,b) HS-AFM images of the outer surface of live M. magneticum AMB-1 for (a) 150
× 150 nm2 area and (b) 100 × 100 nm2 area, captured at 0.5 and 1.43 fps, respectively. Small indents and particles surrounding the indents were seen
over the entire cell surface. Compactly arranged particles forming a reticulate structure were seen at the rims of these indent (arrows in (b)). (c)Trajectories of four reticulate structures superimposed on HS-AFM image (200 × 200 pixels, 2 fps). These trajectories were drawn by tracing the
movement of each indent. (d) Distribution of diﬀusion constants for individual trajectories. The data followed a log-normal distribution characterized bya mean diﬀusion constant D = 3.2 ± 0.4 nm2/s (red solid line). Inset shows MSDs of individual trajectories as a function of time.

Figure 48. HS-AFM images of isolated outer membrane of M. magneticum AMB-1 adsorbed on mica. (a) Reticulate structure similar to those observedon the bacterial outer surface. (b) HS-AFM observation of tip-induced dissection process of the reticulate structure in an outer membrane patch.
Numbers indicate the frame number. Triangle-shaped molecules were seen after the dissection of the reticulate structure. (c) Highly magniﬁed HS-AFM
image of trimeric molecules observed after dissection. Imaging rates, 1.0 fps; pixels, 100 × 100 (a and b) and 160 × 160 (c).

Figure 49. Dynamics occurring on the apical surface of HeLa cell captured by HS-AFM. (a) Fluorescence microscopic image of GFP-transfected HeLa
cells. The cantilever is indicated by the white arrow. (b) Clips of successive HS-AFM images of the apical surface of a live HeLa cell. Growth of aﬁlopodium is indicated by arrows. Scan range, 5 × 5 μm2; imaging rate, 0.2 fps; pixels, 200 × 200.

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Figure 50. Dynamics observed by HS-AFM on the peripheral surface of HeLa cell. (a,b) Topographic image of HeLa cell at an peripheral region (a)before and (b) after the addition of cytochalasin D. Thin black lines in (a) indicate ﬂow of small objects. Black dots in (b) show that no ﬂow is occurring.
Scan range, 5 × 5 μm2; imaging rate, 0.2 fps; pixels, 200 × 200. (c) Clips of successive images showing the process of endocytosis. Initially formed pits arehighlighted by the dotted circles. Scan range, 5 × 5 μm2; imaging rate, 0.2 fps; pixels, 200 × 200.

9.2. Dynamic Phenomena Occurring on Eukaryotic Cell
Surfaces

morphological changes at the apical surface of the cell cultured
on the glass rod were captured by HS-AFM at 0.2 fps for 5 × 5
μm2 with 200 × 200 pixels (Figure 49b). HS-AFM cannot image
ﬁlopodia perpendicularly extending from the cell surface due tothe tip force. Indeed, even during imaging at low interaction
forces, the surface morphology was deformed. Nonetheless, thedynamic growth of some ﬁlopodia was visualized as shown witharrows in Figure 49b.
Next, the tip was positioned at the peripheral region of the cell
where the cell is much thinner and hence the eﬀective stiﬀness ishigher due to the underlying hard substrate. At a peripheral
region of the HeLa cell, sheet-like structures with ∼100 nm
height were captured at 0.2 fps for 5 × 5 μm2 with 200 × 200
pixels (Figure 50a). Interestingly, the HS-AFM movie clearly
showed a continuous ﬂow of small objects directed from the cell
periphery toward the center of the cell body (Figure 50a). On theother hand, after bath application of cytochalasin D (an inhibitorof actin polymerization), the continuous ﬂow ceased completely(Figure 50b). This indicated that the ﬂow observed in the HSAFM movies is the actin retrograde ﬂow. As a matter of course,
the ﬂow is occurring in the cell interior. HS-AFM visualizes the

Imaging of plasma membranes of live eukaryotic cells with AFM
is more challenging than that of outer membranes of live bacteria
cells because the plasma membranes are extremely soft andhence easily deformed by the contact of the AFM tip. An attempt
to observe by HS-AFM dynamic events taking place on theplasma membranes of live eukaryote cells was recently made.70
As a test sample, HeLa cells, one of the most commonly usedimmortal cell lines derived from human cancer cells, wereimaged. The HeLa cells were cultured on a poly-L-lysine-coatedglass rod at 37 °C in 5% CO2. The glass rod was the sample stage
routinely used for HS-AFM imaging. For positioning of the AFM
tip on a deﬁned place of a large HeLa cell, a simple ﬂuorescent
microscope system was incorporated into the HS-AFM system.
Figure 49a shows a ﬂuorescent image of a HeLa cell in which
GFP was overexpressed. The cantilever could be seen as a dark
shade (arrow in Figure 49a), and hence the cantilever tip position
could be easily adjusted on a target region of the cell surface.
Numerous actin-based ﬁlopodia dynamically repeat extension
and retraction at the apical surface of a live HeLa cell.343 Dynamic
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dynamics of cytoskeleton in the cell interior as convolution
between the surface morphology and the local mechanicaldiﬀerences.344
Endocytosis is a cellular process, in which a cell brings
molecules or substances into the cell interior from the exterior by
engulﬁng them with the cell membrane. The engulfment
typically begins with the formation of nanoscale pits on the cellsurface. Recently, this pit formation and disclosure have been
directly visualized on Cos-7 cells by using scanning ion
conductance microscopy (SICM; see section 10), which canoperate in noncontact condition.345 Even with tapping mode
HS-AFM, this process was successfully captured using HeLa cells
at an imaging rate 0.1 fps for 5 × 5 μm2 with 200 × 200 pixels,70 asshown in Figure 50c. In the HeLa cell, mEGFP-Rab5(Rab5Q79L)-CAAX was overexpressed. The small GTPase
Rab5 is known to be crucial in regulation of endocytosis, and its
constitutive active mutant Rab5(Rab5Q79L) stimulates endocytosis.346−348 Replacement of the C-terminus of Rab5 with a
CAAX box derived from k-Ras enhances the plasma membrane
localization of Rab5. As seen in Figure 50c, each of the two pits
encircled by broken lines was capped by a membrane protrusionthat grew with time from the membrane surrounding each pit.

eukaryotic cells. Numerous questions concerning membranesuperstructure and domain formation, and their related dynamics
and responses to environmental stimuli, await to be investigated.351

10. PERSPECTIVES
HS-AFM just recently opened a new avenue to directly andclosely look at biological molecules in action and even dynamic
events occurring in live cells. What biological questions ofimportance will be further addressed and solved by HS-AFM
imaging? How extensively will this new microscopy be used inbiological science? Are higher imaging rates achievable? Is it
possible to combine other techniques with HS-AFM to make it
more useful? Perspectives on these issues are brieﬂy described
below.
Nils Walter predicted as follows: “In a decade or two fromnow, each static structure from X-ray crystallography in ourbiochemistry textbooks may be accompanied by a dynamic videoof the same biopolymer in actual biological action.”352 Thus far,
we have videos of only four proteins that can be placed in suchtextbooks: bR,44,49 myosin V,48 F1-ATPase,52 and P2X4R.37 In
other HS-AFM imaging studies conducted so far, protein
structures are not well resolved (for example, cellulase,57 Ca2+
pump,50 NMDA receptor,61 restriction enzymes31,40,54), or
diﬀusion and interaction processes of proteins are imaged
instead of biological actions while their structures are resolved(for example, c-rings,39 OmpF,59 and aquaporin-060,71). Thereare countless proteins that are interesting to be investigated using
HS-AFM: condensin assembling chromatins into a chromosome,helicase unwinding a double-stranded DNA, RecBCD unwinding a double-stranded DNA and digesting the resulting single
strands, proteasomes digesting polyubiquitinated proteins, E3
ubiquitin ligase self-ubiquitinating or ubiquitinating substrate
proteins, dynein moving on microtubules, AAA proteinsdisaggregating aggregated proteins, ABC transporters transporting substrates, dynamin squeezing membrane tubes, proteintransport apparatuses transporting proteins, MreB and ParM inpolymerization, many channel proteins opening and closing theirchannels during gating, DNA polymerase synthesizing acomplementary DNA strand, and ribosome synthesizing apolypeptide. These are limited examples of important biomolecular systems that have been untouched by HS-AFM. In
principle, any biomolecular action that occurs in the milliseconds
to seconds time range is a potential target for HS-AFM structuraland dynamics analysis. The limited number of systems studied so
far is solely because the number of HS-AFM users is limited. Thissituation will quickly change because HS-AFM instruments withhigh-speed (>10 fps) and low-invasiveness performances are
now commercially available, and small cantilevers optimized for
HS-AFM are also commercially available. By a synergy eﬀect ofincreasing number of HS-AFM users and concomitant
accumulation of the know-how to conduct successful HS-AFM
imaging (preparations of substrate surfaces and samples), thenumber of dynamic videos to be placed in future biochemistry
textbooks will increase. Furthermore, the equipment will become
handier by future enhanced development and less costly. The
latter is also true for consumables like the small cantilevers. As
such, Nils Walter’s prediction will surely come true.
The speed performance of the current HS-AFM instrument
including small cantilevers will not be signiﬁcantly improved.
Maybe 100 fps could be achieved, but even to reach this rate theimaging area will have to be narrowed to ∼20 × 20 nm2. This isbecause the small cantilevers represent the current speed limit.

9.3. Diﬀusion and Interactions of Membrane Proteins on
Eukaryotic Live Cells

Indirect evidence has been accumulated that membrane proteins
and lipids in eukaryotic cells are organized in functional rafts,
work together in multiprotein assemblies, modulate theirfunction by interaction, temporally associate in signal ortransport complexes, etc.349 Direct visualization of a native cellsurface under physiological conditions allowing the stimulationand the observation of a dynamic membrane rearrangement
would provide tremendous advances for the understanding of
membrane related processes. Most recently, ﬁrst breakthroughexperiments have been reported, indicating the feasibility of highspeed and high resolution imaging on eukaryotic cells using HSAFM techniques.70,71 First, the development of a wide-range
scanner that can overview an entire cell while maintaining highresolution capacity has already proven powerful to visualize
dynamic cellular processes on live eukaryotic cells at highresolution, as described above.70 In another approach, theintegration of optical microscopy devices into a HS-AFM setup
allowed the placement of the HS-AFM tip onto areas of interest
on a cell and subsequent high-resolution high-speed imaging
using a fast narrow-area scanner. With this latter technology, HSAFM has been used to visualize individual water channels AQP0
binding to and dissociating from junctional microdomains
directly on lens ﬁber cells.71 As has been demonstrated fordissected AQP0 junctional microdomains by conventional and
HS-AFM,60,238,350 imaging AQP0 on a live cell corroborated
their assembly in square arrays with approximate 6.5 nm unit celldimensions, but importantly revealed novel insights into their
physiological function: although only very slowly (D = 0.5 nm2/s), the junctional microdomains diﬀused freely in the lens cell
membrane. This motional freedom of the junctional microdomains is the key to an understanding of the maintenance ofadhesion between lens cells while the lens tissue undergoes
elastic deformation when focusing to diﬀerent distances. Thispilot study of HS-AFM on live cells addressed a highly specialized
membrane system of a highly specialized cell. However, thetremendous potential of HS-AFM for cell biology applications
was shown by the proof of concept that individual membraneproteins could be imaged even in the membranes of live
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removed the limitation imposed to the sample-stage scan HSAFM that only a small sample stage can be used to achieve highspeed scanning of the sample stage. One can now use large
samples. When the ﬂuorescence microscope combined with the
tip-scan HS-AFM instrument is operated in the total internal
reﬂection mode, simultaneous single-molecule HS-AFM/ﬂuorescence microscopy observations can be performed, as
demonstrated.358 Therefore, in addition to imaging of live
eukaryotic cells and isolated intracellular organelles, thiscombined system allows simultaneously observing, for example,
the dynamic binding and dissociation events of ﬂuorescentligands on single protein molecules and the structural changes ofthe molecules concomitantly occurring with the ligand binding
and dissociation.
Tip-scan mode HS-AFM combined with optical microscopy
allows the full use of the light microscope; not only TIRFM
observations but also confocal and super-resolution ﬂuorescencemicroscopy observations can be performed simultaneously with
HS-AFM imaging. Moreover, optical tweezers can be installed in
the combined system. Therefore, HS-AFM can visualize
biological molecules under external force. In the current singlemolecule AFM-FS, only the magnitude of force applied to aprotein molecule and the extension of the molecule under force
can be measured,359 meaning that the molecular structure
deformed or unfolded by the applied force cannot be visualized.
This is also the case in the current optical trap FS.360 The HSAFM visualization of molecules under external force will provide
a new opportunity to study the eﬀect of applied force on theaction of the molecules as well as detailed unfolding and refolding
processes of the molecules.
Tip-enhanced ﬂuorescence microscopy has resolution <10nm.361 This technique is based on the highly enhanced electricﬁeld close to a laser-illuminated metal or silicon tip,362,363 andcan be installed in a tip-scan mode HS-AFM setup to materialize
super-resolution/high-speed ﬂuorescence microscopy combinedwith HS-AFM. Unlike some of current super-resolution
ﬂuorescence microscopy systems, no signiﬁcant background
ﬂuorescence appears even when relatively high concentrations of
ﬂuorophores exist in the bulk solution because only ﬂuorophores
very close to the cantilever tip can be detected.

Generally, to increase the resonant frequency, one must
compromise with the cantilever stiﬀness and vice versa. The
currently most advanced small cantilevers (resonant frequency
1.2 MHz in water, spring constant 0.2 N/m) deem to have almost
achieved the ultimate goal of balancing these two mechanicalquantities. Much smaller cantilevers might be fabricated, butbuilding a reliable and sensitive OBD detector for such very tiny
cantilevers seems very diﬃcult. Contact mode HS-AFM42 could
potentially circumvent the limitations imposed by the cantileverresonant frequency. It would, however, only be applicable tosolid, well adhering, and ﬂat samples because high friction forces
are unavoidable.
Short cantilever HS-AFM can also be used as a force
measurement tool for high-speed force spectroscopy (HS-FS).
The fast response time of the cantilever, and the fast piezo
elements and electronics will allow force measurements on
biomolecules with orders of magnitude faster rates thanconventional AFM.353 In particular, the small cantilevers featureseveral advantages: high force sensitivity (as discussed above inthe context of imaging), fast response, and low hydrodynamic
drag. We anticipate that HS-FS will have a wide range of
biological applications, notably allowing force application onmolecules with speeds comparable to molecular dynamics
simulations.
SICM, originally developed by Hansma,354 could be analternative to HS-AFM for live cell imaging if the scan speed wasincreased to that of the current HS-AFM. Instead of a cantilever,
SICM uses an electrolyte-ﬁlled glass capillary with an Ag/AgClelectrode inserted from the back, and measures an ionic current
going through a small pore at the apex.355 When the tip apex isplaced close to the sample surface, the impedance for the ioniccurrent is increased. Therefore, the sample surface can bedetected under noncontact condition by solely measuring theionic current. The achieved highest spatial resolution of SICM islaterally 3−6 nm.356 This resolution was obtained by theadvanced fabrication technique for producing very sharp
capillaries with a small pore at the apex, notably by the use of acomputer controlled puller equipped with a CO2 laser to melt thecapillary and the use of quartz capillaries with a high melting
point.357 High-speed SICM with a noncontact imaging capability
would be useful for observing the surface of extremely soft
surfaces of live eukaryotic cells.
The information that can be acquired with HS-AFM is limited,particularly in imaging of live eukaryotic cells and isolated
intracellular organelles. Only dynamic events occurring at theirouter surfaces are observable. Dynamic events occurring there
are often linked with dynamic events occurring in their interiors,
which can be observed with ﬂuorescence microscopy. Therefore,
HS-AFM should be combined with powerful ﬂuorescencemicroscopy. The current HS-AFM instrument employs the
sample-stage scan mode. This design is inadequate for making
simultaneous HS-AFM and ﬂorescence-microscopy observations
possible, because the sample stage moves relative to the objective
lens for ﬂuorescence microscopy and a crucial area to be imaged
simultaneously by ﬂuorescence microscopy is blocked by thecantilever, although suﬃcient ﬂuorescence microscopy opticalsignal can be acquired so as to guide the HS-AFM observation at
speciﬁc loci of cells.70,71 To have high performance optical
microscopy together with HS-AFM, a HS-AFM instrument isneeded that employs the tip-scan mode (the cantilever chip is
scanned at least in the X- and Y-directions). Very recently, a tipscan HS-AFM instrument was developed and combined with aninverted ﬂuorescence microscope.358 This new instrument

11. CONCLUDING REMARKS
The advent of AFM was hailed with acclamations by biologists.
This is because AFM is the ﬁrst and unique microscopy thatallows high-resolution visualization of active biological samples
in aqueous solutions. However, in contrast to the initial
excitement, the enthusiasm for bio-AFM imaging graduallysubsided. This is because biologists could not be satisﬁed onlywith still images. In 1992, one of the inventors of AFM, Gerd
Binnig, predicted that “in biology, use of the force microscope
will probably become quite common because of its ability todeliver ﬁlms of processes.”364 However, AFM was too slow to
ﬁlm biological processes. The eﬀorts to increase the imaging rateof bio-AFM started around 1993 and continued (see review365),
which were, however, not so visible for biologists for a while.
However, tremendous strides were recently accomplished indeveloping biocompatible HS-AFM providing high spatial andtemporal resolution and low-invasiveness. Consequently,
proteins in native biological action were readily captured onvideo. Gerd Binnig’s prediction has now partially come true;
unlike his prediction, HS-AFM has not yet become “quite
common”. Yet it is most certainly just a matter of time.
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Table 3. Resolutions Achieved by Diﬀerent Structural Biology Techniques
technique
X-ray diﬀraction
NMR
EM
crystallography
EM single particle
ﬂuorescencemicroscopy
AFM

sample3D crystal
protein in
solution
2D crystal
protein in
solution
proteins in
cells
adsorbed
proteins

resolution range“high-resolution”

3D electron density
3D distanceconstraints
2D or 3D nuclear
density
3D nuclear density

0.5−8.0 Å in x, y, z0.1−1.0 Å distance constraints

0.5−1.0 Å in x, y, z0.1−0.5 Å distance constraints

366,367
183,368

1.8−25 Å in x, y dimensions, lower resolution
in z dimension
5.0−25 Å

1.8−4.0 Å in x, y dimensions, lower
resolution in z dimension
3−5 Å

250,369
184,370

ﬂuorescent label
localization
surface topography200−1000 nm

10 nm spatial localization239

1−50 nm in x, y dimensions, 0.1−1.0 nm in z
dimension

1−2 nm in x, y dimensions, 0.1−0.2 nm in z
dimension

informationrefs

using new techniques, which often leads to the creation of new
ideas. In this sense, new techniques most contribute to theprogress in science. We hope that HS-AFM will be one of suchnew techniques and that this Review will provide biologicalresearchers with a chance of considering the use of HS-AFM in
their studies to directly observe their molecular and cellular
phenomena under study at unprecedented high spatiotemporal
resolution.

After describing the basic principle of AFM imaging, wequantitatively discussed how fast successive imaging could be
performed by HS-AFM. The highest possible imaging rate wasshown to be a function of several parameters. We found that themagnitude of maximum applied tip-force endurable by the
sample most severely restricts the imaging rate. The tip-force thatexceeds the set point force is caused by the delay in the feedback
control. This delay is minimized in HS-AFM but never vanishes
completely because of the “chasing-after” nature of the feedback
control. The spatial frequency of the sample surface corrugation
to be imaged was also shown to be a key parameter for thedetermination of the maximum possible imaging rate. Even forthe same sample surface corrugation, the detectable highest
spatial frequency depends on the sharpness of the cantilever tip.
The higher spatial frequency can be detected by the sharper tip,which however results in the necessity of taking the images atlower rates.
We did not describe the HS-AFM instrumentation in detail,but rather focused on small cantilevers, sample stages, substratesurfaces, and their associated issues, which HS-AFM users
themselves must deal with. The substrate surfaces are a key
element for successful HS-AFM imaging. Because proteins havemultifarious faces, we should be able to prepare many diﬀerent
substrate surfaces depending on the target proteins andmolecular processes to be visualized. Despite the necessity of
having a variety of substrate surfaces, its menu is still limited
because of the limited number of proteins attempted to bevisualized by HS-AFM. This issue will be improved along withincreasing number of HS-AFM studies.
We started the main part of this Review (i.e., application
studies on proteins) with walking myosin V and described thisHS-AFM study quite in detail. This is because we feel that it is themost impressive and representative demonstration of the abilityof HS-AFM to provide impregnable evidence for the nature of
the dynamic molecular structural changes involved in a crucial
biological mechanism. This study best exempliﬁed the mostimpactful feature of HS-AFM that direct observation of single
protein molecules in biological action immediately andstraightforwardly recounts in detail and provides great insights
into how the protein functions. In addition to several HS-AFM
studies of isolated proteins, we also reviewed two HS-AFM
studies of in situ imaging of protein molecules in live cells.
Because molecular processes on the membrane surfaces of live
cells and the surfaces of intracellular organelles are largely
unknown, HS-AFM will certainly also be intensely applied toprovide novel insight into these topics in the near future.
A Nobel laureate, Sydney Brenner, mentioned “progress inscience depends on new techniques, new discoveries, and new
ideas, probably in that order.” New discoveries are often made

APPENDIX
A.1. Resolution

Very diﬀerent resolution ranges are attained by various
techniques used in structural biology, as summarized in Table
3. The assessment of resolution of AFM topographs is a more
complex problem than one could initially anticipate, and often
AFM users describe their molecular imaging assessment withqualitative terms such as “high-resolution” or “submolecularresolution”, although sometimes the same qualitative terms havebeen used for completely diﬀerent structural details analyzed byother techniques.
The assessment of the resolution in AFM is a diﬃcult task. On
the one hand, the scanning process, with the fast and slow scanaxes, results in an inhomogeneous resolution in X and Y
dimensions. On the other hand, the tip radius crucial for surface
contouring is often unmeasured. For this reason, and for others
like the unknown contribution of electrostatic, chemical, and vander Waals forces, the image contrast transfer function is not
deﬁnable. Today, AFM scientists still debate about image
resolution. Initially, the imaging of membrane protein 2D
crystals has suggested by the visual inspection of diﬀraction spots
in calculated power spectra that the AFM could acquire
topography details with down to 6 Å resolution.371 It has
however been shown that the surface convolution by the AFM
tip of radius comparable to the features or unit cell of the proteins(nanometer scale) imperatively results in higher order diﬀraction
spots not corresponding to realistic image resolution, and hencein overestimation of the imaging resolution.372 An approachcloser to the genuine resolution assessment like the one that has
been used in single particle electron microscopy appeared more
appropriate.372,373 Sometimes it was proposed that like for
optical microscopy, the capability of separating two features fromeach other could be used as resolution assessment in AFM.
However, this leads to an underestimation of the imaging
resolution. This is because AFM contours topography and does
not transform a point signal into a point spread function like an
optical microscope does. For example, AFM can contour a most
protruding surface feature and at the same time document
information about the asymmetric nature of the surrounding
lower topography protein structure; see the HS-AFM top3178

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Table 4. Software Packages That Allow Image Processing
softwarenamewebpageaccessimageJ

http://rsb.info.nih.gov/ij/

freephotoshop

http://www.adobe.com/products/
photoshop.html
http://gwyddion.net
http://www.liv.ac.uk/∼sdb/
ImageSXM/
http://2dx.org/
http://blake.bcm.edu/emanwiki/
EMAN2
http://imagescience.de/imagic/

commercial

gwyddion
image SXM
2dx
EMAN2
IMAGICMRC
SIMPLE

http://www2.mrc-lmb.cam.ac.uk/
image2000.html
http://simple.stanford.edu

free
free
free/open source
free/open source

applications
single pixel manipulation, various operations (Java-based plugins can be downloaded for awide range of image processing tasks)
single pixel manipulation
various scanning probe microscopy speciﬁc image visualization and analysis operations
single pixel manipulation, various scanning probe microscopy speciﬁc image visualization
and analysis operations
2D-crystal processing
2D-crystal processing, single particle averaging

commercial
nonproﬁt prices
free for academia

2D-crystal processing, single particle averaging

free/open source

single particle averaging

2D-crystal processing

ography of the α3β3 subcomplex of F1-ATPase (Figure 13).
When one follows the logics of distinguishing intensity peaks, a
low resolution (of about 3 nm) would be estimated on thesemolecules from the distance between the peaks of the subunits.
This is certainly not the case, because the resolution was highenough to unambiguously distinguish between the conformational states of the molecules by cross-correlation comparison
with a surface rendering of the atomic structure.52 Indeed theAFM contouring is structurally dependent on objects that arelarger than the radius of the tip, where a structure oriented
resolution assessment similar to single particle electron
microscopy is most appropriate. However, when contouring
objects of size similar to the tip radius or smaller, then obviously
the tip convolutes the features in a point spread function manner.
Recently, FM-AFM has reported the resolution of substructure
on individual lipid headgroups in supported lipid bilayers.374
This ﬁnding remains controversial because of the documented
diﬀusion of lipids even in gel phase bilayers.375 Such a highresolution could so far only be acquired on crystalline lipids in a
regular arrangement, and certainly not on a protein. Taking
together, the highest AFM imaging resolution on protein
molecules is of about 1 nm in lateral dimensions and 0.1 nm invertical dimension. This is in agreement with both (i)comparison with atomic structure surfaces rendered to diﬀerent
resolutions and (ii) theoretical considerations concerning the tip
radius. The AFM imaging resolution has not been improved
since pioneering works of high-resolution imaging by contact
mode AFM on bR and aquaporins.203,371,376 However, HS-AFM
in AM mode, which has now achieved the same resolution anddocumented protein dynamics,44 is much less invasive52 and canachieve such high resolution on highly corrugated and relatively
loosely attached molecules.48

meaning of the image.372 In any case, ﬁltering results in altered(diminished) image resolution. One may choose a soft mean ormedian ﬁlter to decrease imaging noise and/or scan related noise(scan lines), which enhances the visibility of the object to beanalyzed. However, only features visible in the original raw data
set should be scientiﬁcally interpreted. For molecular imaging,several image processing packages “inherited” from EM can beused to average and analyze images of molecules; when themolecules are ordered in 2D crystals, lattice oriented algorithms
can be used, while when the molecules are randomly arranged,
single particle averaging approaches can be used.377 Thesemethods function typically well, but care must be taken not toover-interpret the resolution. This is mainly due to the fact that
EM-speciﬁc program algorithms have been designed to workwith a large amount of well-deﬁned (white) noise overlaid to themolecule images. In AFM, the signal-to-noise ratio is muchhigher, and the noise is of diﬀerent nature. Hence, AFM images
produce high signal diﬀraction spots at high resolution whenanalyzed using EM software packages as shown378 anddiscussed.372 Importantly, dynamic processes must be tracked
in the analysis of successive HS-AFM images. For this purpose,
single particle tracking software packages or plugins developed
for ﬂuorescence optical microscopy can be “inherited”.379 Again,
these programs function relatively well for HS-AFM particle
tracking data analysis. Yet there are some limitations; forexample, ﬂuorescence particles can “blink” (disappear andreappear in subsequent frames), and the scan line noise ispresent in AFM images, which might inﬂuence the tracking
algorithm.380 Clearly, there is still a lot of work to be done by anda major interest for the HS-AFM community (and the AFM
community in general) to gather eﬀorts to standardize image
processing and analysis tools adapted to the speciﬁcities of the
microscopy technique. Typically, standardization of processes,
analysis, and statistical assessment provide a major increase in thecredibility of a technical approach and will therefore boost theesteem and use of AFM and HS-AFM in biological sciences ingeneral.

A.2. Software

HS-AFM data visualized must be analyzed. A hopefully helpful
but certainly not complete list of software that can be used toanalyze AFM data in general and HS-AFM data in particular isgiven in Table 4.
Typically, images are brieﬂy pretreated (mostly ﬂattened forscan line non-horizontality) by the AFM-speciﬁc data acquisition
software. After transformation to a more general ﬁle format (tiﬀ,stack of tiﬀs), images can be contrast adjusted and ﬁltered using
any pixel-oriented image treatment software. Of course, caremust be taken to consider that contrast adjustment meanschanging the height scale in AFM topographs and suchalterations must be reliably tracked to maintain the scientiﬁc

A.3. Simulation of AFM Image

Construction of simulated AFM images from protein crystal
structures can be performed using software, for example, theSPM simulator (Advanced Algorithm Systems Co., Tokyo,
Japan). The cantilever tip is modeled as a circular cone of various
angles with a small sphere of various radiuses at the apex. Eachatom in a protein is modeled as a hard sphere with acorresponding van der Wars radius. The simulated images are
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Figure 51. Simulated AFM images of the C-terminal side of α3β3 subcomplex obtained using diﬀerent tip apex radiuses indicated at the top of thecorresponding images.

Figure 52. Correlation coeﬃcient analysis of AFM images. (a) Experimentally acquired AFM image of α3β3 subcomplex in nucleotide-free condition
that was chosen as a reference image and ROIs therein used in the two-dimensional correlation coeﬃcient analysis for the determination ofconformational states of each β subunit. The transparent boxes represent ROIs for three β subunits. (b) Simulated AFM image of α3β3 subcomplex in thepresence of ATP and the ROI (encircled with the white rectangle) containing a β subunit in the E conformational state (i.e., nucleotide-free state). Scalebar, 5 nm. (c) A clip from successively acquired HS-AFM images of α3β3 subcomplex in the presence of ATP and the ROI (encircled with the white
rectangle). Scale bar, 5 nm. (d) Time course of correlation coeﬃcients calculated to determine the conformational states of the β subunit shown in the
ROI of (c). For this calculation, the simulated AFM image and the ROI as a reference shown in (b) were used. Bimodal change of the correlationcoeﬃcient is clearly seen: the larger r (0.98) and smaller r (around 0.95) represent the E and R conformational states, respectively.

processed by a low-pass ﬁlter with an appropriate cut-oﬀ
wavelength that is adjusted to be similar to the spatial resolutionof experimentally obtained AFM images. By the comparison ofexperimentally obtained images to images simulated with various
cone angles and tip apex radiuses, one can approximately
determine the cone angle and the tip apex radius used in theexperiment. The procedure is exempliﬁed here on the α3β3
subcomplex of F1-ATPase, as illustrated in Figure 51 (the coneangle was ﬁxed at 10°).

HS-AFM images of the protein, a reference frame and ROIs arechosen; the ROI in each image fully contains the protein
molecule or the subunit to be analyzed. The 2D correlationcoeﬃcient r deﬁned below is calculated frame-by-frame for each
ROI. The 2D correlation coeﬃcient r is deﬁned as:r=ΣmΣn(Imn − I ̅ )(R mn − R̅ )(ΣmΣn(Imn − I ̅ )2 )(ΣmΣn(R mn − R̅ )2 )(8)Here, Imn and Rmn are intensities at pixel point (m, n) in a ROI to
be analyzed and those in the reference ROI in the reference
frame, respectively. I ̅ and R̅ are mean values of the intensity
matrices I and R, respectively. When the analyzed image and thereference one are the same, the correlation coeﬃcient r becomes

A.4. Correlation Coeﬃcient Analysis

A 2D correlation coeﬃcient calculation can be performed toassign HS-AFM images of a protein to speciﬁc conformational
states of the protein or a subunit in the protein. For a series of3180

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Figure 53. Tracking procedures for a cellulase molecule linearly moving along a cellulose ﬁbril (see text).

1. If they are completely diﬀerent, the correlation coeﬃcient r
becomes zero. Therefore, one can evaluate the similarity of two
images from the value of r. Of course, to assign an AFM image to
one of the speciﬁc conformational states, the conformational
state of the protein (or the subunit) contained in the reference
ROI should be determined anyhow. This determination can bemade again by this correlation coeﬃcient analysis, when crystal
structures of the protein are available for its speciﬁc conformational states. In this case, using the correlation coeﬃcient
analysis, the ROI of the reference image experimentally obtainedis compared to ROIs of the simulated images constructed as
mentioned above. The procedure is exempliﬁed for the βsubunits within the α3β3 subcomplex of F1-ATPase, as shown in
Figure 52, and its results are shown in Figure 15.

(5) In the second frame, the ROI made in the ﬁrst frame istranslated along the linear trajectory obtained above by the
distance expected from the average translocation velocity
determined above. The 2D correlation coeﬃcient r for the
ROI in the second frame is calculated, in which the ROI of theﬁrst frame is used as the reference image. To ﬁnd the ROI
position that gives the maximum value of r, the ROI of the second
frame is translated within tolerance of 5 × 5 pixels. At the ROI
position where the maximum value of r is obtained, the position
of ROI in the second frame is ﬁxed. Next, the center of mass ofthe molecule is calculated within the ROI, which represents the
position of molecule in the second frame.
(6) The same procedures described above are repeated for theremaining frames (Figure 53f).

A.5. Tracking Analysis of Proteins Moving along a Filament

AUTHOR INFORMATION

For statistical analysis of velocities of individual molecules
linearly moving on a ﬁlamentous structure, the center of mass ofeach molecule can be tracked semi-automatically for its
successive AFM images. Because AFM images sometimes
contain spike noises due to instant adsorption of other molecules
to the ﬁlaments, fully automatic tracking is diﬃcult. Here, thesemi-automatic tracking procedure is shown below for the case ofcellulase molecules moving on crystalline cellulose ﬁbers, but
applicable to other systems where objects to be analyzed are
moving approximately in a linear path.
(1) To reduce spike noises in the AFM images, all raw images
are processed by a 2 × 2 pixel averaging ﬁlter before the analysis.
An example is given for the ﬁrst frame as shown in Figure 53a(raw image) and Figure 53b (ﬁltered image).
(2) A single cellulase molecule in the ﬁrst frame is selected, anda ROI is drawn by encircling the molecule with a rectangle(Figure 53c).
(3) Within the ROI, the image is compensated for the heightslope by ﬁrst-order plane ﬁtting, and then the center of mass in
the ROI is calculated (red dot, Figure 53c). This height slope
compensation is sometimes required especially when the heightof the ﬁlamentous structure is not uniform.
(4) In the last frame of the successive images, the same
molecule is encircled with a rectangle, followed by the heightslope compensation and then calculation of its center of mass ofthe molecule (Figure 53d). Next, the determined centers of massof the target molecule in the ﬁrst and last frames are connected bya linear line, which gives the ﬁrst approximation for the trajectory(Figure 53e). An average translocation velocity of the molecule isdetermined using the distance between the centers of mass in theﬁrst and last frames and the time interval between them.

Corresponding Author*E-mail: tando@staﬀ.kanazawa-u.ac.jp.
Notes

The authors declare no competing ﬁnancial interest.
Biographies

Toshio Ando studied physics, applied physics, and biophysics at Waseda
University and received his Dc. Sci. degree in 1980 from Waseda
University. After working on the mechanism of muscle contraction andﬂuorescence techniques for six and half years as a postdoctoral fellow
and then an assistant research biophysicist at UC San Francisco (ManuelMorales’s lab), he was appointed to the faculty at Kanazawa University,where he is currently a Full Professor of Physics and the Director of the
Bio-AFM Frontier Research Center. His research interests include theelucidation of functional mechanism of proteins, particularly of motor
proteins, and the instrument development of high-speed scanning probe
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provide novel insights into the processes taking place in biologicalmembranes. Simon Scheuring has been awarded a European ResearchCouncil (ERC) grant (2012).

microscopes. He has received many awards for his development of highspeed AFM and its biological applications.

ACKNOWLEDGMENTS
We thank colleagues of our laboratories and collaborators fortheir enthusiasm to pioneer biological research by the development and use of HS-AFM techniques. HS-AFM research in the
Ando lab is supported by a HFSP Grant (no. RGP0069 to T.A.),
Grant-in-Aid for Basic Research from the MEXT, Japan (nos.
20221006 and 24227005 to T.A., nos. 23115008 and 30326300
to T.U.), Grants-in-Aid for Scientiﬁc Research on Innovative
Areas from the MEXT, Japan (no. 21113002 to T.A.), a JST’s
program on Development of Systems and Technology for
Advanced Measurement and Analysis (to T.A.), the Knowledge
Cluster Initiative Project from the MEXT, Japan (to T.A.), and
JST/ALCA (to T.U.). HS-AFM research in the Scheuring lab is
supported by a European Research Council (ERC) Grant (no.
310080), and Agence National de la Recherche (ANR) grants(nos. ANR-12-BS10-009-01 and ANR-12-BSV8-0006-01).

Takayuki Uchihashi received his B.Sc., M.Sc. in Physics from Hiroshima
University, Japan in 1993 and 1995, respectively. In 1998, he obtained
his Dc.Eng. in Electronics from Osaka University, Japan. From 1998 to2000, he worked at the Joint Research Center for Atom Technology(JRCAT) in Tsukuba as a research associate. In 2000, he joined theDepartment of Electronic Engineering, Himeji Institute of Technology.
In 2002, he moved to Trinity College in Dublin and worked as a seniorresearcher. In 2004, he joined the Department of Physics, Kanazawa
University, and since 2006 he has been an associate professor. His
research interests include the instrumentation of scanning probe
microscopy and its application to biological science.

ABBREVIATIONSAAAATPases associated with diverse cellular activities
ADP
adenosine diphosphateAMamplitude modulation
AMP
antimicrobial peptide
AMPPNP adenylyl-imidodiphosphate
APS
1-(3-aminopropyl)silatrane
APTES
3-aminopropyltriethoxysilane
AQP0
aquaporin-0
ATPadenosine triphosphateA5annexin 5bR
bacteriorhodopsin
CBHcellobiohydrolases
CBD
cellulose-binding domainCDcatalytic domain
D/A
digital to analog
DNA
deoxyribonucleic acid
DOC
deocycholate
DOPC
1,2-dioleoyl-sn-glycero-3-phosphocholineDPPC
1,2-dipalmitoyl-sn-glycero-3-phosphocholine
DPPE
1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine
EBDelectron beam deposition
EGTA
ethylene glycol tetraacetic acidEM
electron microscopy
EPC
ethylphosphatidylcholine
FACT
facilitates chromatin transcription
fps
frames per secondFMfrequency modulation
GFP
green ﬂuorescent proteinGHglycoside hydrolase
HMG
high-mobility group
HMM
heavy mero myosin
HOPG
highly ordered pyrolytic graphite
IDP
intrinsically disordered protein
IDR
intrinsically disordered region
mEGFP monomeric enhanced GFP
M5
myosin V
MSD
mean square displacement
NMR
nuclear magnetic resonanceNTA
nitrilotriacetic acid
OBD
optical beam deﬂection

Simon Scheuring is the founder and head of the “Bio-AFM-Lab”
laboratory U1006 INSERM/Université Aix-Marseille. He is a trained
biologist from the Biozentrum at the University of Basel, Switzerland(1992−1996). During his Ph.D. in the laboratory of Andreas Engel(1997−2001), he learned electron microscopy and atomic force
microscopy, and got interested in membrane proteins. During thisperiod he worked on the structure determination of aquaporins incollaboration with Peter Agre. During his postdoc at the Institut Curie in
Paris, France, in the laboratory of Jean-Louis Rigaud (2001−2004), helearned membrane physical chemistry and developed atomic force
microscopy for the study of native membranes. As a permanent
researcher (2004−2007) and junior research director (2007−2012), he
set up his lab at the Institut Curie in Paris, France. He has been awarded
an INSERM Avenir grant (2005) and the “Médaille de la Ville de Paris”(2007). He then moved to Marseille (2012) where he built a larger
laboratory named “Bio-AFM-Lab” administrated by INSERM and
Université Aix-Marseille, where he was promoted to senior research
director (2012). His objective is to head a dynamic young research teamwith members from diﬀerent scientiﬁc ﬁelds ranging from biology,
physics, and engineering to develop and use atomic force microscopy to
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OmpF
PC
PDB
PE
PG
PID
PS
RMS
SAM
SERCA
SICM
SLB
SR
SSRP
TAP
WT

Review(29) Ando, T. FEBS Lett. 2013, 587, 997.
(30) Yokokawa, M.; Wada, C.; Ando, T.; Sakai, N.; Yagi, A.;
Yoshimura, S. H.; Takeyasu, K. EMBO J. 2006, 25, 4567.
(31) Crampton, N.; Yokokawa, M.; Dryden, D. T. F.; Edwardson, J.
M.; Rao, D. N.; Takeyasu, K.; Yoshimura, S. H.; Henderson, R. M. Proc.
Natl. Acad. Sci. U.S.A. 2007, 104, 12755.
(32) Miyagi, A.; Tsunaka, Y.; Uchihashi, T.; Mayanagi, K.; Hirose, S.;
Morikawa, K.; Ando, T. ChemPhysChem 2008, 9, 1859.
(33) Sugasawa, H.; Sugiyama, Y.; Morii, T.; Okada, T. Jpn. J. Appl. Phys.
2008, 47, 6168.
(34) Yamamoto, D.; Uchihashi, T.; Kodera, N.; Ando, T. Nanotechnology 2008, 19, 384009.
(35) Yamashita, H.; Voïtchovsky, K.; Uchihashi, T.; Contera, S. A.;
Ryan, J. F.; Ando, T. J. Struct. Biol. 2009, 167, 153.
(36) Igarashi, K.; Koivula, A.; Wada, M.; Kimura, S.; Penttilä, M.;
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outer membrane protein F
phosphatidyl choline
protein data bankphosphatidyl ethanolamine
phosphatidylglycerol
proportional integral derivative
phosphatidylserine
root-mean-squareself-assembled monolayersarcoplasmic reticulum Ca2+-ATPase
scanning ion conductance microscopy
supported lipid bilayersarcoplasmic reticulum
structure speciﬁc recognition protein
trimethylammoniumpropane
wild type

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