Microscopy 62(1): 81–93 (2013)
doi: 10.1093/jmicro/dfs093........................................................................................................................................................................................................................................................

Review

High-speed atomic force microscopy
Toshio Ando*
Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan*To whom correspondence should be addressed. E-mail: tando@staff.kanazawa-u.ac.jp..............................................................................................................................................................................................

Abstract..............................................................................................................................................................................................

Keywords

atomic force microscopy, high-speed AFM, realtime imaging, dynamics ofproteins, biomolecular processes, bioimaging..............................................................................................................................................................................................

Received

27 September 2012, accepted 2 December 2012; online 4 January 2013..............................................................................................................................................................................................

Introduction
Directly observing individual protein molecules inaction at a high spatiotemporal resolution has long
been a holy grail of biological science. An ideal microscopy technique performing this type of observation has to meet all the following conditions (seeTable 1): (i) in-liquid specimen imaging, (ii) highspatial resolution, (iii) high temporal resolution,(iv) low invasiveness to the specimen and (v) direct imaging of the specimen without the use of
markers (in other words, the morphology of the specimen itself is resolved). For example, ﬂuorescencemicroscopy that is reinforced by super-resolution
techniques surpassing the diffraction limit [1] meetsthe ﬁrst through the fourth conditions, but is neverable to satisfy the ﬁfth condition in principle.

Although many efforts have been made to develop
liquid electron microscopy techniques capable of observing non-stained biological specimens in solutions[2], the electron dose required to achieve high contrast and spatial resolution instantaneously kills the
sample. Solving this problem is a highly difﬁcult,if not impossible, task. Atomic force microscopy(AFM) meets the ﬁrst, second and the ﬁfth conditions and moderately meets the fourth condition, but
its temporal resolution (30 s at its highest level) is
too low to trace the dynamics of biological molecules
in real time. However, when I started to develop
high-speed AFM (HS-AFM) two decades ago, the low
temporal resolution of AFM had intuitively seemed
to me to be an issue that could be circumvented,
although I had no rational strategies at that time.

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High-speed atomic force microscopy (HS-AFM) has been developed as anano-dynamics visualization technique. This microscopy permits direct
observation of structure dynamics and dynamic processes of biologicalmolecules in physiological solutions, at a subsecond to sub-100 ms temporal resolution and an 2 nm lateral and a 0.1 nm vertical resolution.
Importantly, tip–sample interactions do not disturb the biomolecules’
functions. Various functioning proteins including myosin V walking on an
actin ﬁlament and bacteriorhodopsin responding to light have been successfully visualized with HS-AFM. In the quest for understanding thefunctional mechanisms of proteins, inferences no longer have to be madefrom static snapshots of molecular structures and dynamic behavior of
optical markers attached to proteins. High-resolution molecular movies
obtained from HS-AFM observations reveal the details of molecules’
dynamic behavior in action, without the need for intricate analyses andinterpretations. In this review, I ﬁrst describe the fundamentals behind
the achieved high imaging rate and low invasiveness to samples, andthen highlight recent imaging studies. Finally, future studies are brieﬂy
described.

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Table 1. Feasibility comparison of three types of microscopy
Conditions
Microscopy

In-liquid specimen High spatial resolution High temporal resolution Low invasiveness Direct imaging

Fluorescence microscopy W
Electron microscopy;
Atomic force microscopy W

W

W

W

×

W

;
×

×
;

W

W

W

W , Fully met; ;, Partially met or conditional; ×, impossible or extensive efforts required to meet.
‘Direct imaging’ means that the morphology of an object itself is resolved.

is no doubt that this new microscopy will be widelyused in biological research in the near future. This
review does not intend to describe speciﬁc HS-AFM
techniques, but instead provides the fundamentals
behind the high imaging rate and low invasivenessto the sample, highlights recent imaging studies ofproteins and outlines ongoing and future studies toexpand the scope of HS-AFM’s application to biological studies. For technical details of the instrumentation, more application studies and experimental
procedures for imaging, see reviews in [17,31,32],
respectively.

Fundamentals behind high imaging rate
Principles of tapping mode AFM

Among a number of operation modes, HS-AFM
employs the tapping mode [33]. In this mode, thecantilever is oscillated at its ﬁrst resonant frequency
fc. A stylus probe attached to the free end of thecantilever is intermittently brought into contact
with the sample, which signiﬁcantly reduces thefriction between the tip and the sample duringlateral scanning of the sample stage. The mechanical response of the cantilever upon contact, i.e.
change in the cantilever’s oscillation amplitude, ismeasured, and then the sample stage is moved in
the z-direction to recover the amplitude back to its
initial set point value through feedback control (seeFig. 1). This series of operations is repeated manytimes for different sample surface points duringlateral scanning of the sample stage. Consequently,
the movement of the sample stage traces the
sample surface, and therefore, the feedback controller output at each x–y position represents the
sample height at the position. Thus, the sample
surface topography is constructed by the recorded
feedback controller output.

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AFM was originally invented to visualize atoms
on solid surfaces [3] but is now routinely used inbiological sciences [4,5] for high-resolution imagingof biological samples under physiological conditions [6,7], as well as force measurements to estimate the strength of intra- and intermolecularbonds at single-molecule level [8,9] and the elasticity of biological surfaces [10–12]. Soon after theadvent of AFM, several groups attempted toobserve dynamic biological events despite the very
low imaging rate of AFM: Hansma and colleagues
observed the clotting process by ﬁbrin at 1 min
intervals [13], and a group including one of theinventors of AFM (Binnig) also tried to image thedynamic processes of live cells being infected by avirus [14] and the binding of antibodies to the
S-layer of bacterial cells [15]. In 1992, Binnig predicted that ‘in biology, use of the force microscope
will probably become quite common because of its
ability to deliver ﬁlms of processes’ [16]. However,
we had to wait 16 years until this prediction cametrue.
In 2008, the instrumentation development for
HS-AFM was completed [17]. Toward this completion, various techniques and devices were developed: small cantilevers [18–20], optical beam
deﬂection detectors for detecting deﬂection of asmall cantilever [20,21], fast amplitude detectors[17,20], fast scanners [17,22,23], a counterbalancemethod to avoid the generation of unwanted vibrations of the scanner [20,23], an active damping
method for the z-scanner [24], a compensator for
drift in the cantilever excitation efﬁciency [25] anda new feedback controller [25]. Since the completion of HS-AFM instrumentation development, thetechnique’s ability to reveal the structure dynamics
and dynamic processes of proteins has been
demonstrated continuously [26–30]. As such, there

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83

ﬂat but varies with time as

Feedback bandwidth

There is always a time delay (τ0) in feedback
control. This time delay and other factors, which
will be speciﬁed later, limit the maximum possible
imaging rate Rmax. A quantitative relationshipbetween Rmax and these limiting factors is given
below. For simplicity, it is assumed that the sample
surface proﬁle in the x–z plane has a sinusoidal
shape with a periodicity λ and height h0. When thesample-stage scanner is moved in the x-direction at
velocity Vs, the sample height h under the cantilevertip changes with time as h0hðtÞ ¼
 ½sinð2pf tÞ þ 1;2

ð1Þ

where f = Vs/λ. During feedback scanning, the zscanner moves in the direction opposite to thesample height so that the sample surface looks ﬂatwhen viewed from the cantilever. However,
because of the time delay τ0, the z-scanner moves
at feedback frequency f as
 h0
 ½sinð2pf t  uÞ þ 1;zðtÞ ¼ 
2

ð2Þ

where θ = 2πfτ0. Therefore, the sample surface whenviewed from the cantilever does not look perfectly

  

u
u
SðtÞ ¼ hðtÞ þ zðtÞ ¼ h0 sin
cos 2pf t  : ð3Þ
2
2
The deviation signal S(t) is feedback error. When the
cantilever-free oscillation amplitude and the amplitude set point are set at A0 and As (=rA0; 0 < r < 1),
respectively, the cantilever tip in contact with thesample surface is pushed upward by the sample toan extent of D(t) = S(t) + A0(1 − r) provided that the
sample is more rigid than the cantilever.
The feedback bandwidth fB, which is a value
characterizing the speed performance of feedback
operation in the AFM system, is usually deﬁned by
the feedback frequency at which a 45° phase delay
occurs in the feedback scan. Thus, fB = 1/(8τ0).
Because the feedback frequency f should notexceed fB, the condition f < fB holds, which limits
the scan speed Vs as Vs < λfB. By all our efforts toincrease the response speed of all devices contained in the feedback loop, particularly of themechanical devices (i.e. cantilever and scanner), fB
is now enhanced up to 110 kHz. For a scan size inthe x-direction W and number of scan lines N, themaximum possible imaging rate Rmax is given by
Rmax = (λfB)/(2WN) when no sample fragility is considered. For example, for fB = 110 kHz, W = 200 nm,

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Fig. 1. Schematic of tapping-mode AFM setup. The cantilever is vertically oscillated at its ﬁrst resonant frequency. The cantilever deﬂection
is detected by the optical beam deﬂection detector comprised of the laser diode and the split photodetector. The differential signal from thesplit photodetector is converted into an amplitude signal by the amplitude detector. The detected amplitude signal is compared with thepredetermined set point, and their difference (error signal) is processed (mainly integrated) by the PID feedback controller to produce adriving signal for displacing the z-scanner so that the error signal approaches zero. By this control, the oscillation amplitude of the cantilever(and hence, the vertical tip–sample interaction force) is mostly maintained constant during the lateral scanning of the sample stage. The D/Aconverters implemented in the PC output drive signals to laterally move the sample stage, while the A/D converter records the feedback
controller output representing the sample height.

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N = 100 and λ = 10 nm, we obtain Rmax = 27.5
frames s−1 (fps).

Sample fragility as a limiting factor
of imaging rate

where kc and Qc are the spring constant and quality
factor in water of the cantilever, respectively. The
degree of sample fragility can be expressed inseveral ways. For example, sample fragility can be
expressed as the maximum possible tapping force
F0 under which the sample still retains its full biological function. The condition F0 > Fmax and Eq.
(4) determine the maximum possible phase delay
θmax for given values of kc, Qc, A0, h0 and r. Thisθmax modiﬁes Rmax as
Rmax ¼

ð2lfB umax Þ:ðpW NÞ

ð5Þ

Another factor that determines θmax arises from thecondition that the cantilever deﬂection D(t) should
always be positive. When D(t) < 0, and when no adhesive force is exerted between the tip and thesample, the tip loses contact with the sample
surface at its bottom swing of oscillation. When the
tip completely detaches from the sample surface,the error signal becomes saturated at A0(1 – r), regardless of the tip’s distance from the sample
surface. In order for the tip to lightly tap thesample surface, the magnitude of A0(1 – r) has to be
set at a small value. When the error signal is saturated under this light-tapping conditions, it takes
time for the tip to land on the surface again, i.e.
‘parachuting’ occurs. During parachuting, no information on sample topography is obtained. The condition that D(t) should always be positive results in u
A0 ð1  rÞ  h0 sin
. 0:
ð6Þ
2

Effect of tip tapping on sample

HS-AFM was developed with the goal of observing
dynamic events of functioning biomolecules.
Therefore, we must quantitatively consider any
adverse effects of the tip tapping on the sample.
Note that the mechanical quantity that affects the
sample (i.e. causes a momentum change) is not theforce itself but the impulsive force (the product of
the force acting on the sample and the time forwhich the force acts). The small cantilevers, which
we have been using routinely, have kc = 0.2 N m−1,
fc = 3.5 MHz in air and 1.2 MHz in water and Qc = 2
in water. As described later, non-invasive imaging
can be carried out for various proteins under conditions of A0 = 1 nm, r = 0.8–0.9 and θ = 20°. Under
these conditions, the magnitude of the maximum
tapping force estimated using Eq. (4) is 100 pN
for a typical protein height of h0  5 nm. However,the force acting time is short (100 ns or less).
Thus, the magnitude of the impulsive force is small.
The effect of tip tapping on the sample can also
be estimated from the energy loss of an oscillating
cantilever. When a cantilever with kc = 0.2 N m−1 isoscillated at A0 = 1 nm, its mechanical energy is
only 24 kBT, where kB is Boltzmann’s constantand T is 300 K. This value is comparable with theenergy of ATP hydrolysis, 20 kBT. Even when thisamount of energy is completely lost and used toexcite the sample, the energy would be distributed
into many degrees of freedom within the tapped
portion of the sample and quickly dissipate into
more degrees of freedom, including those of surrounding water molecules (as discussed below). In
actual imaging experiments, such events of complete energy loss would happen only very rarely, if

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Because of the feedback error, the cantilever tip
taps more strongly as it scans a sample surface thatis increasing in height (i.e. as the tip is heading‘uphill’ on the surface), and more weakly as it scansa sample surface that is decreasing in height (i.e.
heading ‘downhill’). The maximum tapping force
Fmax is approximately expressed as
  
 
kc
u
Fmax ¼
 A0 ð1  rÞ þ h0 sin
;
ð4Þ
Qc
2

For example, for A0 = 1 nm, h0 = 5 nm and r = 0.8,
Eq. (6) gives θmax < 4.6°. Therefore, the maximum
possible imaging rate is reduced to 0.1 × (λfB)/(2WN). Thus, this parachuting problem is the largest
obstacle in achieving a high imaging rate under lighttapping conditions. However, this difﬁcult problem
has been solved by a new feedback controller (adynamic proportional-integral-derivative (PID) controller) we developed [25], in which the feedback
gain is automatically increased when A exceeds As(i.e. when the tip is scanning at a downhill region on
the sample).

T. Ando High-speed AFM

Recent progress in bioimaging by HS-AFM

In 2008, HS-AFM was started to be utilized tosolve biological issues that had to date been difﬁcult or impossible to address by other methods.
Since then, the number of publications on bioimaging by HS-AFM has gradually increased, increasing rapidly in the last three years (see review [31]).
The bioimaging studies conducted thus far havecovered a wide range of dynamic molecular processes and structure dynamics, which can be classiﬁed into (i) structure dynamics of proteins in action[26–28,34–37], (ii) selfassembly processes [31,38,39],(iii) dynamic protein–protein and protein–DNA
interactions [40–44], (iv) diffusion processes [30,45–47], (v) molecular processes associated with enzymatic reactions [31,32,48–50] and (vi) dynamics occurring with DNA origamis [51–55]. Moreover,non-biological dynamic phenomena, such as thephotodegradation of single chains of a chiral helical
π-conjugated polymer [56], the phase transition ofdetergent aggregates [57] and dissolution of photoresists [58] have been captured on video. Below we
describe two of our own recently published imaging
studies of proteins in action.

Rotary catalysis of axle-less F1-ATPase

The minimum unit of F1-ATPase, α3β3γ subcomplex,
is a rotary motor that retains full ATPase activity(Fig. 2a) [59–61]. The coiled-coil part of the rotary
shaft γ is inserted in the central cavity formed by astator ring α3β3, where three α and three β subunits
are arranged alternately [62]. Three ATP-bindingsites are located at the α–β interfaces, mainly in theβ subunits. The rotation of γ occurs in the counterclockwise direction as viewed from the exposed
globular portion of γ (or from the C-terminal side ofα3β3). In the ATPase cycle, three β subunits take different chemical states: ATP-bound, ADP-bound andnucleotide-free (empty) states [59,62]. Each chemical state cyclically propagates among the threeβ subunits (i.e. rotary catalysis occurs). Therefore,
there is strong cooperativity among the β subunits.
How is this cooperativity created? There is no
direct contact between any pairs of β subunits andat any instance the β–γ interaction is always different for the three β subunits because γ has no symmetry. Considering these facts, Wang and Oster [63]proposed that interactions with γ control and determine the conformational and catalytic states of individual β subunits. This idea was reinforced bystudies showing that backward mechanical rotationof γ with external force reverses the chemical reaction toward ATP synthesis [64,65], whereas forced
forward rotation accelerates ATP binding [66]. Thisview was challenged by the ﬁnding that even when
γ is shortened so that most γ–β interaction sites are
eliminated, the short γ still rotates unidirectionally[67]. However, in single-molecule optical microscopy, there is no way to observe the rotary catalysis
after the complete removal of γ. Thus, we exploitedthe power of HS-AFM to solve this issue [28].
α3β3 with Lys7-tags at the N-termini of its β subunits was covalently immobilized on an aminoreactive mica surface. An image of the C-terminalside of α3β3 without nucleotide shows a pseudosixfold symmetric ring (Fig. 2b). Each subunit has an
upwardly protruding portion at the inner top side of
the ring, but these portions are higher at three alternately arranged subunits than at the other three.
Comparison of this image with a simulated AFM
image (Fig. 2d) constructed from a crystal structureof nucleotide-free α3β3 (PDB code, 1SKY) (Fig. 2c)

Downloaded from http://jmicro.oxfordjournals.org/ at Kanazawa University on February 24, 2013at all, and overall, the cantilever oscillation amplitude is maintained at r × A0 = 0.8–0.9 nm. On
average, the amount of energy loss after each tap isvery small: ½kcA20(1 – r 2)/Qc = 2.3–4.4 kBT.
The tip oscillating at high fc in water iteratively
taps the sample. When one image is captured for an
area of Wx × Wy nm2 at an imaging rate R, a molecule with a top surface area (Atop nm2) perpendicular to the tip is tapped fc/R × Atop/(WxWy) times perframe. For example, under the conditions (fc  1
MHz, R = 12.5 fps, and Wx × Wy = 45 × 22 nm2) used
for non-invasive imaging of axle-less F1-ATPase (i.e.
the γ subunit is removed from F1-ATPase) [28], one
molecule of axle-less F1-ATPase (Atop  100 nm2) istapped 8000 times per frame and 4 million times
in total during the successive 40 s imaging.
Nevertheless, the axle-less F1-ATPase is fully functional throughout the imaging, as shown later. This
result clearly indicates that the energy transferredfrom the oscillating tip to the sample never accumulates in the sample but instead dissipates quickly(much faster than the oscillation period of the cantilever, 1 μs).

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M I C R O S C O P Y , Vol. 62, No. 1, 2013[68] revealed that the three subunits showing higher
protrusions are the three β subunits. In 1 mM
AMP-PNP, the shape of α3β3 became triangular andthe central hole became obscure (Fig. 2e). Only one
subunit had a higher protrusion together with an outwardly extended distal portion, which was identiﬁed
to be an empty β subunits by comparison of the experimental AFM image with a simulated one (Fig. 2g)
constructed from a crystal structure of nucleotidebound α3β3γ subcomplex (PDB code: 1BMF) [62]from which γ was removed (Fig. 2f). In addition, thetwo subunits with retracted distal portions were identiﬁed to be nucleotide-bound β subunits.
When imaged in 2–4 μM ATP at 12.5 fps, distinct
conformational dynamics appeared at the β subunits(Fig. 3a shows images obtained in 2 μM ATP; see thesupplementary data online, Movie S1 was acquired in
3 μM ATP). Each β subunit exhibited a conformational transition between the outwardly extended highstate (open (O) state) and the retracted low state(closed (C) state). The following prominent features
were observed: (i) only one β subunit assumes the O
state and (ii) when the O-to-C transition occurs at
one β subunit, the opposite C-to-O transition occurs
simultaneously at its counterclockwise neighbor
β subunit in most cases. Thus, the O conformation
propagates counterclockwise (as noted by the changing locations of the highest pixels, each marked
with a red circle, in Fig. 3a). The ATP hydrolysis
rates at 2, 3 and 4 μM ATP were approximately
one-third the rates of conformational propagation at
2, 3 and 4 μM ATP, respectively. Thus, the conformational propagation is tightly coupled with the ATPasereaction. As shown in Fig. 3b, the cumulated angle ofthe highest pixel with discrete 120° steps revolves linearly with time until the end of imaging (40 s total).
Thus, the function of α3β3 is completely retained
throughout the imaging. As mentioned above, the
α3β3 molecule is tapped with the oscillating tip 4
million times in total during this successive imaging.

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Fig. 2. AFM images of α3β3 subcomplex at C-terminal surface. (a) Schematic for F1-ATPase. (b) Averaged AFM image obtained innucleotide-free condition. (c) C-terminal surface of crystal structure of nucleotide-free α3β3 subcomplex. (d) Simulated AFM image of (c).
(e) Averaged AFM image obtained in 1 mM AMP-PNP. (f ) C-terminal surface of crystal structure of α3β3 subcomplex obtained in ATP.
(g) Simulated AFM image of (f ).

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87

These dynamic transitions indicate that theO-to-C transition occurs when ATP is bound to an
empty β subunit while the C-to-O transition occurs
when an ADP-bound β subunit releases ADP. So theempty, ADP-bound and ATP-bound β subunits arearranged counterclockwise in this order, and therefore, the observed conformational propagation
demonstrates rotary catalysis by the α3β3 subcomplex. Thus, we reach the important conclusion thatthe intrinsic cooperativity responsible for torquegeneration to rotate γ is elicited through β–β interplay alone, and that γ passively undergoes torque torotate [28].

Bacteriorhodopsin in response to light

Bacteriorhodopsin (bR) is a light-driven proton
pump, transferring protons across the membranefrom the cytoplasmic side to the extracellular side
[69,70]. bR comprises seven transmembrane αhelices (named A–G) surrounding the retinal
chromophore covalently bound to Lys216 via a protonated Schiff base [71,72]. In the native membrane(referred to as the purple membrane), bR

assembles into trimers, which are arranged in ahexagonal lattice. bR has been extensively studied
by optical spectroscopy techniques. Upon absorption of light, photoisomerization from the all-transconformation to the 13-cis conformation of theretinal takes place. This isomerization induces protontransfer from the Schiff base to Asp85, which triggers
a cascade of changes in the bR structure. A series ofintermediates designated J–O are deﬁned, among
which M (M410) having an absorption peak at 410
nm and the longest lifetime is the only intermediatecontaining a deprotonated Schiff base [70]. In contrast to this well-deﬁned spectroscopy of bR during
its photocycle, the overall dynamic structural change
in bR upon photoactivation has long been elusive,
despite the establishment of detailed static structurein the unphotolyzed state by X-ray crystallography
and electron microscopy [73–75]. Thus, bR is one of
the best targets for which the power of HS-AFM may
be demonstrated.
The photocycle of wild-type (WT) bR at neutral
pH proceeds very quickly (10 ms). To slow down
the photocycle, we used the D96N bR mutant
which has a longer photocycle (10 s at pH 7) but

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Fig. 3. Rotary propagation of conformational change of α3β3 subcomplex at C-terminal surface. (a) Successive images of C-terminal surfaceof α3β3 subcomplex in 2 μM ATP, captured by HS-AFM at 12.5 fps. The highest pixel position in each image is marked with the red circle.
(b) Time evolution of the 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 β subunits in the open state.

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still acts as a proton pump [76]. Native purple membranes containing D96N bR were directly placed ona bare mica surface. Figure 4a shows successive
images of the D96N mutant at the cytoplasmic
surface captured at 1 fps [26]. Upon illumination
with green light (532 nm, 0.5 μW), bR markedly
changed its structure (as observed by comparisonof images at 1 and 2 s) and returned to the groundstate within a few seconds after illumination was
discontinued. This structural change was speciﬁcally induced by green light, not by blue or red light,
which is consistent with the fact that bR in the
ground state effectively absorbs green light. The
centers of mass of all activated bR monomers
moved outward by 0.7–0.8 nm from the center ofthe respective trimers and rotated counterclockwiseby 7–8° around the center of the respective trimers.
The images shown in Fig. 4b and c are averaged
images under dark and illumination conditions, respectively (also see the supplementary data online,
Movie S2); the averaging was performed using
several trimers appearing in each frame, considering the hexagonal lattice arrangement of thetrimers. From these images, it is evident that each
protrusion is split into larger and smaller sections(arrows 1 and 2) upon light illumination. Comparedwith the atomic model of the α-helical cytoplasmic

ends in the unphotolyzed state [72], the protruding
areas around helices E and F shift outward fromthe trimer center under illumination. Therefore, theprotrusion movement visualized here is ascribed tothe displacement of the E–F loop. The minor protrusion (arrow 2) likely corresponds to the position
around helices A and B. Note that owing to indiscernible alterations at the extracellular surface, theoverall position of each bR molecule does notchange. The photoactivated state observed by
HS-AFM was attributed to the spectroscopically
identiﬁed MN intermediate [35] (it exhibits outward
tilt of helix F similar to that of the N state of WT
while the Schiff base is still deprotonated like theprecedent M state because of the absence of the
proton donor residue (Asp96) for the Schiff base).
Thus, the photo-induced structural change in bR,
which has long been studied but elusive, is now
clearly revealed by direct visualization with HS-AFM.
As a result of the outward displacement, nearestneighbor bR monomers (one set is circled in lightblue circle in the image at 1 s in Fig. 4a), eachbelonging to a different adjacent trimer (shown by
the three white circles), appear to transiently assemble (as indicated by the light-blue circle in theimage at 2 s). We deﬁne the triad of nearestneighbor monomers belonging to different trimers

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Fig. 4. Structural change in bR induced by light absorption. (a) Successive high-speed AFM images of the cytoplasmic surface of D96N bR indark and illuminated conditions, captured at 1 fps. The green bars at 2 and 3 s indicate illumination of 532 nm green light with an intensity of
0.5 μW. A bR trimer is highlighted by the white triangle in the image at 0 s. In the image at 1 s, three adjacent trimers are encircled with whitecircles, while a trefoil comprised of three nearest-neighbor monomers, each belonging to one of the encircled trimers, is indicated with thelight-blue circle. The same trefoil is encircled in light blue in the images at 2 and 3 s. (b) Averaged AFM image of trimer in dark conditions.
(c) Averaged AFM image of trimer under illumination. Under illumination, the protrusion is split into larger (arrow 1) and smaller (arrow 2)
sections.

T. Ando High-speed AFM

Unique features of HS-AFM imaging

As demonstrated above, the entire structure of aprotein molecule and its minute changes with time
can be captured in an AFM movie. Unlike ﬂuorescence microscopy, wherein the subset of molecular
events observed is dependent on the selected sitewhere the probe is placed (and hence, dynamic
events to be observed are preselected), various
dynamic events of a molecule appear in a molecular movie without preselection. Therefore, we canhave a great chance of learning multiple molecularactions in one imaging experiment. Moreover, molecular actions that appear in a video can often beinterpreted in a straightforward manner, without intricate analyses and interpretations, leading to convincing, if not irrefutable, conclusions. These
excellent features of HS-AFM imaging signiﬁcantly

facilitate and accelerate our understanding of thefunctional mechanisms of proteins.
However, HS-AFM often achieves this superiority
at the cost of intensive sample preparation. For successful visualization of dynamic events in biomolecular systems, certain conditions are often requiredof the sample preparation, particularly for the substrate surface on which sample molecules areplaced. The substrate surface must have an appropriate afﬁnity for the molecules. To observe
dynamic interactions between different species ofmolecules, selective attachment of one of thespecies of molecules to a surface is required.
Otherwise, there is no chance of observing theirinteraction. The surface must be ﬂat in HS-AFM,
unlike ﬂuorescence microscopy. Moreover, in AFM(and HS-AFM), the sample can be viewed from only
one direction, that is, perpendicular to the surface.
Therefore, to observe speciﬁc molecular features,
molecules sometimes have to be oriented selectively on the surface, as in the case of the observationof myosin V molecules walking on an actin ﬁlament[27]. Methods that are used to fulﬁll these conditions are described in articles [32,79,80].

Future prospects of HS-AFM studies
Dynamic imaging of isolated biomolecularsystems

Various bioimaging studies using HS-AFM havebeen undertaken in recent years. Nonetheless, there
are countless biomolecular systems remaining to be
imaged. Even for biomolecular systems that havealready been studied in detail by other approaches,
HS-AFM allows us to inquire into more details orquestions that have been difﬁcult or impossible toaddress by other methods, giving us a great chanceof making unexpected new ﬁndings. This is indeed
exempliﬁed in our studies of F1-ATPase and bR, as
described above, as well as myosin V [27] and cellulase [29]. Importantly, the HS-AFM instrument wehave developed is now commercially available fromthe Research Institute of Biomolecule Metrology,
Inc. (Tsukuba, Japan). Small cantilevers made ofsilicon nitride (9–10 μm long, 2 μm wide and 130 nm
thick; BL-AC10DS, Olympus, Tokyo) with fc = 1.5
MHz in air and 0.6 MHz in water, kc = 0.1 N m−1
and a quality factor Qc  2 in water are alreadyDownloaded from http://jmicro.oxfordjournals.org/ at Kanazawa University on February 24, 2013as a ‘trefoil’, to distinguish such a structure fromthe original trimers. To investigate the possible
effect of bR–bR interaction within a trefoil on thephotocycle, bR was illuminated with green light ofvarying intensity (0.007–0.5 μW). Under weak illumination, only one monomer in each trefoil was
activated in most cases. When only one monomer
was activated in a trefoil, it decayed with a time
constant of 7.3 ± 0.58 s. In contrast, under stronger
illumination, two or three monomers within atrefoil tended to be activated together. Interestingly,
the decay of each monomer markedly depended on
the order of its activation: the monomer that was
activated last among the activated monomers in a
trefoil decayed with a shorter time constant of
2.0 ± 0.16 s, whereas the decay kinetics of themonomers activated earlier did not follow a single
exponential, and the average decay time lengthenedto 13 s. Thus, bR–bR interaction within an illuminated trefoil creates both positive and negative cooperative effects in the decay kinetics as the initialbR recovers.
In contrast to previous expectations [77,78], thesecooperativities do not occur within a trimer, but
within a trefoil. This ﬁnding was made possible byclosely looking at individual bR molecules at a highspatial resolution, in real space and real time. It isdifﬁcult to imagine how such cooperativity could beobserved using techniques other than HS-AFM.

89

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M I C R O S C O P Y , Vol. 62, No. 1, 2013

commercially available. Smaller cantilevers are also
being commercialized by NanoWorld (Neuchâtel,Switzerland). Thus, there is no doubt that continued
and expanded exploration of dynamic HS-AFM
imaging will be carried out for a wide range of biomolecular systems in the near future, which will accelerate and cultivate a better understanding of
how biomolecules function. Thereby, HS-AFM willtransform traditional structural biology and singlemolecule biophysics.

The fast scanners that we have developed are optimized for high-speed imaging of isolated biomolecules; their resonant frequencies are very high, but
the scan range is limited to 1 μm, 4 μm and 0.8–1 μm
in the x-, y-, and z-directions, respectively (when
100 V is applied at maximum). Therefore, large
samples such as live bacterial and mammalian cells
have not been able to be imaged by HS-AFM until
recently. To overcome the scan range limitation,
we have recently developed a wide-area scannerthat can be displaced up to 43 μm in the x- andy-directions and 2.5–4 μm in the z-direction (when
100 V is applied at maximum). The resonant frequency of these x- and y-scanners is low (1.6 KHz),
but can be extended to 10 kHz by an inverse
compensation-based active damping technique. The
resonant frequency of the z-scanner (50–100 kHz) isdetermined by the piezoactuator used (and hence
approximately by the maximum possible displacement). This resonance is removed by the previously
developed Q-control technique [24], but there is no
way to extend the bandwidth at present. Using thisscanner and another medium-range scanner, imagesof whole live bacterial cells were recently acquired.
After taking an image of an entire magnetic bacterial
cell, a localized portion of its outer surface wasimaged fast, visualizing porin molecules that weredensely packed but rapidly diffusing [45]. This ability
to acquire wide-range images will create a new opportunity to observe the dynamics of molecules onhigher order structures such as live mitochodoria,
nuclei and neuronal spines. Given that the functional
dynamic processes of molecules and molecular assemblies occurring on these surfaces are largely

Imaging of dynamics of mammalian cells

In our currently developed HS-AFM method, thefeedback operation is continuously performed atevery point on the sample surface. Therefore, ittakes a longer time to acquire an image of a larger
object. However, single-nanometer resolution isoften not needed in the imaging of cell morphology.
If the feedback operation is performed only at
surface points corresponding to pixels of an image
to acquire, the imaging time can be signiﬁcantly
reduced. This type of feedback operation is already
employed in peak-force mode AFM [81] and in
hopping mode [82] (also called AC-mode [83] orpulse mode [84,85]) scanning ion conductance microscopy. The cantilever base or the sample stage
is moved up and down at a hopping frequency fH ina relatively large hopping amplitude AH so that theprobe is intermittently brought into contact with, orclose to, the sample surface. The feedback scan is
performed only when the probe tip is retracted
from the sample surface. This technique can be
implemented in HS-AFM, allowing much faster acquisition of the entire image of a mammalian cellwith a spatial resolution much higher than that of
optical microscopy.
The imaging rate R for this type of HS-AFM isdetermined by R = fH/(Nx × Ny), where Nx and Ny
are the number of pixels in the x- and y-directions,
respectively. For example, in the case of a typicalz-scanner having a resonant frequency fz = 50 kHz
and a maximum displacement 4.3 μm at 100 V, we
can use fH  30 kHz. Therefore, a 10 × 10 μm2 area
can be imaged at 0.75 fps with Nx × Ny = 200 × 200( pixel size, 50 nm), provided that the z-scanner retraction can be performed quickly after detection oftip–sample contact. However, the z-scanner cannot
respond quickly even when its quality factor Qz is
reduced to 1 (the response time Qz/(πfz)  6.4 μs).
When AH = 100 nm is used, the z-scanner moves
down further during this time delay, and therefore,
the sample is indented 10 nm or more by the tip.
This problem can be solved by the use of az-scanner to which an additional piezoactuator witha high resonant frequency (1 MHz) is attached;

Downloaded from http://jmicro.oxfordjournals.org/ at Kanazawa University on February 24, 2013

Dynamic imaging of molecules on higher order
structures

unknown, HS-AFM will open new avenues for cellbiology as well.

T. Ando High-speed AFM

91.......................................................................................................................

13 Drake B, Prater C B, Weisenhorn A L, Gould S A C, Albrecht T R,
Quate C F, Cannell D S, Hansma H G, and Hansma P K (1989)
Imaging crystals, polymers, and processes in water with the atomic
force microscope. Science 243: 1586–1589.

with this additional piezoactuator, the z-scanner
can be retracted immediately after detection of tip–
sample contact.

.......................................................................................................................

Supplementary data

14 Häberle W, Höber J K H, Ohnesorge F, Smith D P E, and Binnig G(1992) In situ investigations of single living cells infected by viruses.
Ultramicroscopy 42–44: 1161–1167.

Supplementary data are available at http://jmicro.
oxfordjournals.org/.

Acknowledgments

15 Ohnesorge F, Heckl W M, Häberle W, Pum D, Sara M, Schindler H,
Schilcher K, Kiener A, Smith D P E, Sleytr U B, and Binnig G (1992)
Scanning force microscopy studies of the S-layers from Bacillus coagulans E38-66, Bacillus sphaericus CCM2177 and of an antibody
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16 Binnig G (1992) Force microscopy. Ultramicroscopy 42–44: 7–15.
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17 Ando T, Uchihashi T, and Fukuma T (2008) High-speed atomic force
microscopy for nano-visualization of dynamic biomolecular processes. Progr. Surf. Sci. 83: 337–437.
.......................................................................................................................

18 Walters D A, Cleveland J P, Thomson N H, Hansma P K, Wendman
M A, Gurley G, and Elings V (1996) Short cantilevers for atomic
force microscopy. Rev. Sci. Instrum. 67: 3583–3590.

Funding.......................................................................................................................

This work was supported by a grant-in-aid for Basic
Research S (20221006 and 24227005) from the
MEXT, Japan, by a Knowledge Cluster Initiative
Project and by a CREST project of the Japanese
Science and Technology Agency.

.......................................................................................................................

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I thank Takayuki Uchihashi, Noriyuki Kodera, Daisuke Yamamoto,
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