Pflugers Arch - Eur J Physiol (2008) 456:211–225
DOI 10.1007/s00424-007-0406-0

INVITED REVIEW

High-speed AFM and nano-visualizationof biomolecular processes
Toshio Ando & Takayuki Uchihashi & Noriyuki Kodera &Daisuke Yamamoto & Atsushi Miyagi &Masaaki Taniguchi & Hayato Yamashita

Received: 24 September 2007 / Accepted: 15 November 2007 / Published online: 20 December 2007
# Springer-Verlag 2007

Abstract Conventional atomic force microscopes (AFMs)
take at least 30–60 s to capture an image, while dynamic
biomolecular processes occur on a millisecond timescale or
less. To narrow this large difference in timescale, various
studies have been carried out in the past decade. These
efforts have led to a maximum imaging rate of 30–60 ms/
frame for a scan range of ~250 nm, with a weak tip–sample
interaction force being maintained. Recent imaging studies
using high-speed AFM with this capacity have shown that
this new microscope can provide straightforward andprompt answers to how and what structural changes
progress while individual biomolecules are at work. This
article first compares high-speed AFM with its competitor(single-molecule fluorescence microscopy) on various
aspects and then describes high-speed AFM instrumentation and imaging studies on biomolecular processes. The
article concludes by discussing the future prospects of thiscutting-edge microscopy.
Keywords Microscopy . Imaging . ATP hydrolysis .
Motion . Kinetics

T. Ando (*) : T. Uchihashi : N. Kodera : D. Yamamoto :A. Miyagi : M. Taniguchi : H. Yamashita
Department of Physics, Kanazawa University,
Kakuma-machi,
Kanazawa 920-1192, Japane-mail: tando@kenroku.kanazawa-u.ac.jp
T. Ando : T. Uchihashi : N. Kodera : D. Yamamoto
CREST/JST,
Sanban-cho, Chiyoda-ku,
Tokyo 102-0075, Japan

Introduction
Protein molecules dynamically change their structureduring functioning. Therefore, their functional mechanisms
cannot be understood only from the ensemble average
information such as crystal structures at different staticstates and biochemical properties. We need single-molecule
techniques that will allow us to track and measureindividual biomolecules at work. The recent developments
in fluorescence video microscopy allow us to observe thetranslational or rotational motion of the individual fluorescent spots that are emitted from fluorophores attached tobiomolecules. Such observations are valuable for dissecting
the dynamic behavior of the labeled biomolecules in action[1–3]. However, we still need to infer from the observed
behavior of the fluorescent spots how the labeled molecules
actually behave. For this, if possible, we should directly
observe single molecules themselves at nanometer spatial
resolution and millisecond temporal resolution. This would
be a more straightforward approach to understanding
biomolecular behavior than observing fluorescent spots.
Unlike optical microscopy, AFM can directly visualize
unstained biomolecules in physiological solutions at nanometer resolution. This unique capability inspired researchers to apply it to the observation of biomolecular processes.
For example, in 1989, Paul Hansma and his colleagues
observed at ~1-min intervals the fibrin clotting process
which was initiated by the digestion of fibrinogen withthrombin [4]. Using tapping mode AFM [5], Hansma,
Bustamante, and their colleagues obtained time-lapse
images (~30-s intervals) of the RNA replication reaction
by DNA and RNA polymerase [6] and of the onedimensional diffusion of RNA polymerase along a DNA
strand [7]. However, as biomolecular processes generally
occur on a millisecond timescale or less, the image

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acquisition time (30 s to a few minutes) was too long tocapture images in real time.
Attempts to increase the AFM imaging rate were
initiated in 1991 by Quate and his colleagues. Theirattempts are made with the aim of fast inspection orlithography of a wide range of surfaces of hard materials[8], rather than of observing biomolecular processes.
Images (a few micrometer scan range) of a metalsemiconductor field-effect transistor were taken at a frame
rate of ~0.3 s/frame in constant-height mode (i.e., no
feedback scan in the z-direction). This study proceeded
towards the development of cantilevers with integrated
sensors and/or actuators [9, 10] and cantilever arrays with
self-sensing and self-actuation capabilities [11–13]. Unfortunately, fabrication of these cantilevers could be made only
in relatively large dimensions, resulting in low resonant
frequencies (although higher than those of conventional
tube scanners) and large spring constants. Insulation
coating of the integrated cantilevers for the use in liquids
further lowered the resonant frequency [14]. Thus, thisapproach has not come to fruition of realizing high-speed
AFM for biological research.
Studies aiming at realizing high-speed AFM forbiological research were pioneered around 1994. Some
results from these studies gradually began making anappearance during 1996–2002. In 1998, a fast-imaging
study was reported [15], in which an image of DNA wastaken in 1.7 s using small cantilevers and an opticaldeflection detector [16] designed for them. A more
complete high-speed AFM system was reported by us in2001 [17] and 2002 [18], wherein a high-speed scannerand fast electronics were introduced in addition to smallcantilevers (resonant frequencies ~600 kHz in water) andan optical deflection detector for them. An imaging rate of
80 ms/frame was achieved, and therefore, moving myosin
V molecules were filmed in successive images. This stage
is considered as the first stage of high-speed bioAFM
development.
In the second stage (2003–2007), the capability of highspeed AFM was further enhanced by the introduction of
new devices such as active damping techniques forminimizing vibrations of the z-scanner [19] and a new
feedback controller that could maintain a low tip–sample
interaction force without sacrificing the scan rate [20].
Various imaging studies were attempted to capture biologicalprocesses on video. Some of these studies demonstrated theusefulness of high-speed AFM for elucidating the molecularmechanisms of proteins [21, 22]. Now, the most advanced
tapping-mode high-speed AFM can capture images on video
at ~30–60 ms/frame for a scan range of ~250 nm under
feedback operation capable of preventing weak protein–protein interactions from being disturbed by the scanning tip[23, 24].

Pflugers Arch - Eur J Physiol (2008) 456:211–225

High-speed AFM is becoming a truly useful tool forbiological sciences. We expect that in the near future, it will
be fully established and commercially available, leading to a
rapid increase in the user population. This article aims toprovide potential users and developers with a concise introduction to high-speed AFM with regard to its features relative
to single-molecule optical microscopy, instrumentation, application to biological studies, current capabilities and limitations,and future prospects. AFM movies have been placed at http://
www.s.kanazawa-u.ac.jp/phys/biophys/roadmap.htm.

Comparison with TIRFM
Total internal reflection fluorescence microscopy (TIRFM;[1–3]) is a single-molecule technique for tracking dynamic
biomolecular processes that competes with high-speed
AFM. It therefore seems worthwhile to compare these two
microscopic techniques, as they are complementary innature and have advantages and disadvantages over eachother.
Spatial resolution
The greatest advantage of AFM is, of course, its highspatial resolution. The spatial resolution of optical microscopy is given by Abbe’s law: d=0.5λ/NA, where λ=wavelength of monochromic light and NA=limiting numerical aperture of an objective. The
spatial resolutionofﬃ
pﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
AFM is approximately given by 2rt ðΔzÞmin, where rt =radius of curvature of the tip apex and (Δz)min =resolution
limit of the sample height detection. For typical values of rt =5 nm and (Δz)min =0.3 nm with bioAFM, the spatial
resolution becomes ~2 nm, which is ~100 times higher thanthat with conventional fluorescence microscopy. This resolution is still approximately ten times higher than a more
recently developed fluorescence microscopy, namely, stimulated-emission-depletion (STED) fluorescence microscopy[25, 26]. However, this high resolution of AFM is not
sufficient to detect the binding of very tiny ligands (such as
ATP) to a protein. In TIRFM, we can monitor the binding
events using fluorescent ligands [27].
Time resolution
Fluorescence video microscopy is usually performed at an
imaging rate of 33 ms/frame. This is comparable to themost advanced high-speed AFM, although the imaging areaof optical microscopy is much wider than that of AFM.
Fluorescence video microscopy can be performed at faster
imaging rates, using bright fluorescent labels and fastervideo cameras. The imaging rate of AFM can be enhanced
to 1 ms/frame [28]. This is accomplished by discarding thePflugers Arch - Eur J Physiol (2008) 456:211–225

tapping-mode operation and the feedback control for
maintaining a constant tip–sample interaction force. Without feedback control, it would be impossible to observe
biomolecular processes.

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recognition imaging is compatible with high-speed imaging.
This issue will be discussed later. Due to the nonselective
visibility with AFM, the substratum surface should have verysmall roughness. This condition narrows the range of our
choice for a substratum.

NoninvasivenessCombined optical microscope and high-speed AFM
With regard to noninvasiveness to samples, optical microscopy has the strongest advantage, as long as illumination
wavelengths longer than ~500 nm are used. In high-speed
tapping mode AFM, the tip–sample interaction force cannot
be reduced to a negligible level for samples that containdelicate biomolecular interactions. Presently, a truly noncontact condition is realized only with in-vacuum AFM[29]. High-speed noncontact AFM (nc-AFM) for in-liquidsamples will therefore become an important target in thenear future.
Floating molecules
The advantage of AFM is that floating molecules in thesolution are completely invisible, and therefore, theirconcentrations can be increased. When we study the
dynamic interaction of a floating protein with the counterpart protein attached to the substratum, invisibility offloating molecules is quite advantageous. In TIRFM, theconcentration of floating molecules with fluorophores
cannot be increased beyond 50~100 nM, as an elevationin the concentration results in too much background
fluorescence [27].
Visibility
With AFM, every object attached to a substratum andthe substratum surface itself are visible (i.e., nonselective visibility). This feature is in marked contrast to
TIRFM. With TIRFM, only fluorescently labeled molecules are visible, a feature that appears advantageous, as
we can extract only what we would like to see.
However, we always have to bear in mind that key
information may be missed in the selective visualization.
In addition, with TIRFM, photobleaching imposes
serious limitations not only on the length of successive
imaging but also on the measurement of slow kinetic
association and dissociation events (i.e., difficulty in
distinguishing the dissociation events from photobleaching; [30]). With AFM, it is sometimes hard to identify
observed molecules contained in a multicomponent
system; with TIRFM, however, this can be easily
achieved by specific labeling with fluorophores or by
GFP techniques. A recently developed technique, “recognition AFM” [31, 32], appears to solve this problem.
However, it has not been experimentally tested whether

As described above, the two microscopic techniques havecomplementary features. For single-molecule observationsof biomolecular processes, combining these techniques intoa single instrument would therefore be useful. For example,
we can visualize dynamic protein structural changes byhigh-speed AFM while recording simultaneously opticalsignals of chemical reactions such as ATP hydrolysis at
individual protein molecules. Macromolecules such asprotein, DNA, or complexes such as chromosomes are
stretched by optical tweezers while their structural changes
are being imaged by AFM. Chemical specification of an
object imaged by AFM can be made from its Raman or
fluorescence signals. There have been a number of reports
on conventional slow AFM combined with confocal
fluorescence [33–36] or conventional Raman scattering[37, 38] microscopes, coherent anti-Stokes Raman scattering [39, 40] microscopes, or TIRFM [41–43]. When still
images are taken, simultaneous recording of optical and
AFM images is not necessarily required. When simultaneous recording is required, various precautions arenecessary. Cantilever tips of amorphous silicon andamorphous silicon nitride are known to be strongly photoluminescent [44–46], while crystalline silicon tips are muchless luminescent [33]. EBD carbon tips are also lessluminescent. As silicon nitride tips are usually amorphous,and therefore, its luminescence has to be filtered out, whichnarrows our choice of wavelengths used in optical
microscopy. We need to use crystalline silicon tips or
EBD carbon tips particularly in multicolor confocal
imaging. UV, near UV and visible lights are absorbed inthis order by gold often used for cantilever coating. The
thermal expansion coefficient of gold is larger than thoseof silicon and silicon nitride, and therefore, light illumination causes cantilever deflection [24]. In tapping mode
AFM, DC deflection of a cantilever does not affect AFM
imaging, so that the photothermal bending of a cantilevershould not be a problem as far as its bending is not toolarge and the light power is stable. In addition, by
switching off the illumination light during the trace scan,and by switching it on during the retrace scan [33], we cantake optical images without disturbing AFM imaging.
However, heating a cantilever causes a problem of
increasing the temperature of a solution around thecantilever. Therefore, we have to avoid using UV andnear UV lights.

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electron beam deposition in naphthalene gas and sharpen it(~4 nm in radius) by plasma etching in argon gas.

Maximum imaging rate
While feedback control to maintain the constant tip-sample
interaction force is indispensable for high-speed bioAFM, it
limits the imaging rate considerably. The reachable feedback frequency (feedback bandwidth) is determined by thesum of time delays that occur with various devices
contained in the feedback loop, such as the cantilever,
amplitude detector, feedback controller, and z-scanner. The
maximum image acquisition rate Vimage is estimated fromthe feedback bandwidth fB, scan range (Ws), and thenumber of scan lines (Ns). Vimage is approximately given by
Vimage ¼

wfB
aw
P
¼
2WS NS 16WS NS t n

ð1Þ

where w is the apparent width of features on the surface,Στn is the sum of time delays, α (~1.4) is a factor by which
the differential operation contained in feedback control
compensates for a phase delay in the feedback loop [20].
w is approximately given by the convolution of the tip
pﬃﬃﬃﬃﬃﬃﬃ
radius rt and the sample radius rs (i.e., w ¼ rt rs ). Here,the feedback bandwidth is defined as a feedback frequency
that results in a phase delay of 45° in tracing the sample
topography. This phase delay produces a feedback error, andtherefore, the tip–sample interaction force is not perfectly
maintained at a constant level. When the sample is delicate
enough to be damaged or disturbed by the extra force causedby the feedback error, Vimage should be smaller than thatgiven by Eq. 1. By introducing various devices mentioned
below, the feedback bandwidth now stands at ~130 kHz,
which approximately corresponds to the maximum imaging
rate of ~30 frames/s for a scan range of ~240 nm and 100
scan lines.

Instrumentation of high-speed AFM
The basic structure of high-speed AFM is similar to that ofconventional AFMs. However, various devices have been
optimized for high-speed scanning. The optimized devices
are given below.

Optical beam deflection detector
To focus an incident laser beam onto a small cantilever,a ×20 objective lens with a long working distance of 8 mm
(Nikon, CFI Plan Fluor ELWD 20×C; NA, 0.45) is used.
The beam reflected back from the cantilever is collectedwith the same lens. The incident and reflected laser beams
are separated using a quarter wavelength plate and apolarization splitter [15–17]. The laser driver is equippedwith a radio frequency power modulator for reducing thelaser beam coherence [48]. The photosensor consists of afour-segmented Si PIN photodiode (3 pF, 40 MHz) and acustom-made fast amplifier/signal conditioner (~20 MHz).
Fast amplitude detector
Conventional rms-to-dc converters use a rectifier circuit anda low-pass filter and require at least several oscillation
cycles to output an accurate rms value. To detect thecantilever oscillation amplitude at periodicity of a half
oscillation cycle, we developed a method to capture thepeak and bottom voltages and then produce their difference
as the amplitude [17]. The amplitude detection timeτ d ð¼ 1=2fc Þ is 0.42 μs for our small cantilevers with aresonant frequency of ~1.2 MHz in water.
High-speed scanner
We employed flexure stages made of blade springs for the x
and y scanners (Fig. 1). The flexure stages are made bymonolithic processing to minimize the number of resonantpeaks [22]. The y-scanner displaces the x-scanner, and thex-scanner displaces the z-scanner on the top of which asample stage is placed. The maximum displacements of thex and y scanners at 100 V are 1 and 3 μm, respectively. The
x-piezoactuator is held at both its ends with flexures, so that

Cantilevers
Most advanced cantilevers for high-speed AFM are manufactured by Olympus using silicon nitride [47]. They are ~6 μm
long, ~2 μm wide, and ~90 nm thick, and are coated withgold of thickness ~20 nm, which results in a resonant
frequency fc of ~3.5 MHz in air and ~1.2 MHz in water,
spring constant kc ~0.2 N/m, and quality factor Qc ~2.5 inwater. Therefore, their response time τ c ð¼ Qc =πfc Þ is 0.66 μs
in water. The cantilever tip is not sharp enough (~17 nm inradius at the apex). We usually attach an additional tip using

Fig. 1 Scanner structure. a Top view. Note that the x-piezoactuator(X) is sandwiched between two sets of flexures. b 3D view. A dummy
z-piezoactuator is placed to take a weight balance with the zpiezoactuatorPflugers Arch - Eur J Physiol (2008) 456:211–225

its center of mass is hardly displaced and, consequently, nolarge mechanical excitation is produced. The x-scannershows resonant peaks at 45, 65 kHz, and higher frequencies; however, these peaks are not large. The z-piezoactuator (maximum displacement, 2 μm at 100 V; self-resonant
frequency, 360 kHz) is held only at the four corners of its
surface that is perpendicular to the displacement direction.
The z-piezoactuator can be displaced almost freely in boththe counter directions, and consequently, impulsive forces
are barely exerted on the holder; thus, only a very low
mechanical excitation is produced to the scanner mechanics.
This holding method has an additional advantage in that theresonant frequency is not lowered by holding, although the
maximum displacement decreases by half. The x-scanner isactively damped either by the previously developed Qcontrol technique [19] or by feedforward control using
inverse compensation [49, 50]. The z-scanner is also actively
damped either by the Q-control technique or by inverse
compensation. A circuit that automatically produces aninverse transfer function for a given transfer function wasdeveloped [24, 51]. By this compensation, the z-scanner
bandwidth fs is extended to ~500 kHz, and the quality factor
Qs is reduced to ~0.5. Therefore, its response timeτ s ð¼ Qs =πfs Þ is ~0.32 μs.

215

avoided completely, resulting in a significant increase in the
feedback bandwidth and a decrease in the tapping force.
Drift compensator
The drift in the cantilever excitation efficiency poses a
problem, especially when As is set close to 2A0. For
example, as the efficiency is lowered, 2A0 decreases andconcomitantly, App decreases. The feedback system interprets this decrease in App as the tip interacting with thesample too strongly and, therefore, withdraws the sample
stage from the cantilever; this, of course, is an incorrect
operation. This withdrawal tends to dissociate the cantilevertip completely from the sample surface, especially when As
is close to 2A0. Thus, the dynamic PID controller does notoperate properly if the cantilever excitation efficiency is notstabilized. Drifts in the excitation efficiency can be detectedby monitoring drifts in the second harmonic amplitude of
cantilever oscillation averaged over a time period longer
than the image acquisition time [52]. To maintain theconstant difference (2A0 −As), we controlled the output gainof a wave generator connected to the excitation piezoactuator [20]. With this drift compensation, very stable imaging
was achieved, even with a small difference (2A0–As)=
0.4 nm.

Dynamic PID control
Other devices (piezo driver, phase detector)
To observe biological processes that contain delicate
protein–protein interactions, the tip–sample interaction
force has to be minimized. The tapping force exerted fromthe oscillating cantilever tip to the sample can be reduced
by using an amplitude set point r close to 1. However, under
this condition, the oscillating cantilever tip tends to detachcompletely from the sample surface at steep down-hillregions of the sample. Once detached, like “parachuting,” ittakes time for the tip to touch the surface again. Parachuting
results in loss of sample topography information and in a low
feedback bandwidth. During parachuting, the error signal(difference between the cantilever’s peak-to-peak oscillation
amplitude App and the peak-to-peak set point amplitude As) is
saturated at (2A0 −As)=2A0(1−r), where 2A0 is the freeoscillation peak-to-peak amplitude. A shallower set point
results in a smaller saturated error signal and hence prolongs
the parachuting time τp. The feedback gain cannot be
increased to shorten τp, as a larger gain induces overshootat up-hill regions of the sample, resulting in the instability of
the feedback operation. To solve this problem, we devised a
new PID controller named a “dynamic PID controller” that
can automatically change the feedback gain depending on
the oscillation amplitude [20]. In other words, the feedback
gain is increased when App exceeds a threshold level that is
set between As and 2A0 (or at As). By this technique, theparachuting time τp is significantly reduced, or parachuting is

The piezodriver for the z-scanner is custom-made andcharacterized with a maximum output voltage of 50 V, rms
noise <2 mV, high slew rate of 1,000 V/μs, and bandwidthof 3 MHz for capacitive loads up to 2 nF. Our smallcantilevers have a ~1,000 times larger value of fc/kc, whencompared with conventional cantilevers. Therefore, a large
phase shift occurs by tip–sample interaction even with asmall Q, showing the potential for fast phase-contrast
imaging. We developed a fast-phase detector [53] basedon a design by Stark and Guckenberger [54]. This device
can detect phase shifts within a single oscillation cycle
and, more importantly, at any timing within a cycle.
Because of this feature, we can choose the detection
timing at the point where the largest phase shift occurs. In
addition, it can be inferred whether the tip–sample
interactions conserve or dissipate energy. This is because
the phase shift due to an energy-conservative interaction
decreases very quickly with time, while the phase shift
caused by an energy-dissipative interaction is maintained
over the oscillation cycle [53].
Photothermal actuation of cantilevers
The tip–sample distance can be controlled not only by thez-scanner but also by directly actuating the cantilever. An

216intensity modulated infrared laser beam was used tophotothermally deflect small cantilevers [24]. The slow
response of the photothermal expansion effect was eliminated by inverse compensation [24]. This method allowed
faster control of the tip–sample distance due to the muchhigher resonant frequency of the small cantilevers than thatof a z-piezoactuator.
Data acquisition system
High-speed data acquisition systems (DAQ) are commercially available. We have been using DA/AD boards(10 MHz, 12 bits) with memories of ~0.5 MB/channeland direct access memory (DMA) controller chips, together
with our own making software. DMA allows high-speed
data transfer without imposing a load to the computer CPU.
We can attain a minimum image acquisition time of ~3 ms
using the DA/AD boards. As DAQ with higher rates(100 MHz, 12–16 bits) are available, faster frame acquisition is possible. However, it takes time to display a bitmap
image even when data are transferred directly to a bitmap
memory area. For example, it takes about 6–10 ms todisplay a 256×256 pixel image. To avoid a time lag
between frames caused by this delay, we have to thin out
the frames to be displayed. Large amounts of data streamout of high-speed imaging. The maximum number ofacquirable successive frames depends on the memory size
equipped with the computer. For a typical memory size,
2 GB, of a personal computer, approximately 20,000
successive images with 12-bit resolution and 256×256
pixels can be recorded. For a frame rate of 30 ms/frame, we
can record continuously for 10 min. Quantitative descriptions of AFM image acquisition abilities of a given DAQ
are given by Fantner et al. [55, 56].

Comparisons of three high-speed AFM Systems
Three groups (Hansma, Ando, and Miles) have reported
instrumentation of high-speed AFM for biological samples.
Each AFM setup has unique features, advantages, anddrawbacks over the others. Unfortunately, confusion has
been occasionally seen in literatures where evaluation andcomparison of different setups are described. We cannot
simply discuss the comparative merits of high-speed AFM
setups just by the achieved imaging rates or scan velocities
in the x-direction because these quantities depend on
various factors that are not necessarily relevant to the
instrument performance.
Miles’ group has demonstrated ultrafast imaging (1,300
frames/s) of collagen fibers in an ambient environment[28]. This remarkable speed was achieved by AFM
operation in constant-height mode (without feedback scan

Pflugers Arch - Eur J Physiol (2008) 456:211–225

of the z-scanner), resonant scan of a tuning fork x-scanner(resonant frequency, 30–100 kHz; [57, 58]), and fast digital
handling of cantilever deflection data [28]. The cantilevers
used are not described well in their articles. The use of atuning fork as an x-scanner was pioneered by Seo et al.
[59, 60]. In an ambient environment, a strong attractive
capillary force is produced at thin water layers confined
between the tip and the sample. This force avoids the tip
being detached from the sample surface even at anextremely high tip velocity of >4 cm/s. In addition, thisattractive force prevents the dried collagen sample frombeing damaged by the rapidly scanning tip. However, theirhigh-speed AFM is limited to samples firmly attached to a
substratum in an ambient environment and, therefore, does
not seem applicable to the observation of biomolecular
processes.
Hansma’s group has developed various devices for highspeed AFM; fast scanners [56, 61], data acquisition systems[55, 56], an optical deflection detection system [15] forsmall cantilevers, and two types of small cantilevers [56].
Their designs seem to be made with intention of making
them compatible with the Veeco Nanoscope. Their smallcantilevers for contact mode imaging have a resonant
frequency of 150–350 kHz in air and a spring constant of
1–5 pN/nm, while those for tapping mode imaging have aresonant frequency of up to 2 MHz in air and a spring
constant below 2 N/m. Their scanners are designed withintention of observing relatively wide areas; a scan range of
6–15 μm in x- and y-directions and 6 μm in z-direction.
The scanners are equipped with strain gauge sensors forlinearizing the displacements by a closed loop controller.
Their lowest resonant frequency is 10–28 kHz. However, it
is not clear from their papers how high resonant frequency
is attained for the most important z-scanner. Judging fromthe piezoactuators used in the z-scanner, it must be around60 kHz (a few times lower than that of our z-scanner). It isalso not clear whether an active damping controller isimplemented in the z-scanner. As the digital feedback
system implemented in the Veeco Nanoscope III is used,the developed devices do not seem to be effectively utilized
to achieve high imaging rates. Thus, their images of
biological samples taken in a sub-second timescale wereobtained in the so-called error mode (error signals are used
to form images; [56, 62]).
As described in detail in the previous section, Ando’s
group has been developing various devices with strong
intention of realizing video rate imaging of biomolecular
processes. We have developed the whole high-speed AFM
setups without considering their compatibility to commercial AFMs. We have been concentrating on increasing the
feedback bandwidth as high as possible. We also have been
making efforts for low tapping forces exerted from an
oscillating tip to the sample. Sacrificing the scan range toPflugers Arch - Eur J Physiol (2008) 456:211–225

217

achieve a high feedback bandwidth is inevitable at present.
However, this issue may be resolved as discussed latter.

Imaging of biomolecular processes
High-speed AFM is not completely established yet as a tool
for studying biomolecular processes. Therefore, in thepresent stage, biological processes that have been expectedor known to occur have to be demonstrated by high-speed
AFM. In addition, techniques for preparing samples andsubstrates for their attachment have to be developed; thesetechniques are often different from those for still imaging.
Along with the gradual accumulation of successful imagingof known molecular processes, high-speed AFM will beconsidered a reliable tool, while newly filmed data on
unexplored biological processes will be widely accepted.
The imaging studies that have been performed by us andothers are described below.
Early imaging studies
Early imaging studies with high-speed AFM were performed mainly for evaluating the capacity of this new
microscope and for finding devices for improvement. Soon
after the first successful imaging of moving myosin
molecules at 80 ms/frame [17], it was realized that the
tip–sample interaction force was too strong to image fragile
samples such as actin filaments and microtubules, whichwere easily disrupted during imaging (Fig. 2). The
disruption of fragile samples was not only due to theinsufficient feedback speed but also due to the large tapping
force exerted from the oscillating tip to the sample. Using a
dynamic PID controller, the tapping force was greatly
reduced. For example, the unidirectional movement of
individual kinesin molecules along microtubules [21] wasobserved (Fig. 3) without microtubule disruption. However,in the observed kinesin movement, the kinesin molecules
were attached to both the mica surface and microtubules.
The movement of kinesin that was attached only to
microtubules was not observed, implying that the tip
removed kinesin that had attached only to microtubules.
In the observation of actin filament movement, weFig. 2 Actin filaments being disrupted during imaging. The number
attached to each image indicates the time (s) elapsed after imaging
began. Imaging rate, 80 ms/frame; scan size, 400 nm

Fig. 3 Movement of kinesin–gelsolin along a microtubule. The
number attached to each image indicates the time (s) elapsed after
imaging began. The arrowheads point at kinesin–gelsolin. Scale bar,
100 nm; imaging rate, 0.64 s/frame

experienced a similar problem. When the mica surface
was densely coated with myosin V, the gliding of actin
filaments was captured on video (Fig. 4; [22, 23]).
However, when the myosin V density was lowered, thegliding movement was hardly observed. These observations
indicated that with the assistance of a dynamic PID
controller, the tip–sample interaction force was successfully
reduced to a level at which protein denaturation wasprevented and self-assembled polymer structures were notdisrupted. However, the force was not reduced sufficiently
to prevent the delicate protein–protein association frombreaking.
It is important to observe how protein conformation
changes dynamically upon binding to its ligand. Protein
molecules always show Brownian motion due to thethermal agitation of surrounding water molecules. Therefore, changes in protein shape caused by this random
agitation have to be distinguished from ligand-induced
conformational changes. When the ligand associationdissociation kinetics is fast, the ligand-induced changes
occur repeatedly. A certain type of structural change that isrepeatedly observed can be judged as one induced by thereaction. On the other hand, when the kinetics is slow, this
distinction is difficult. For example, due to the low ATPase
activity of myosin V alone (~0.05/s), it was difficult tojudge whether the structural changes observed just once or
twice were, in fact, those induced by the binding of ATP (orits hydrolysis; Fig. 5).
To solve this problem, we combined a UV-flash
photolysis of caged compounds with high-speed AFM.
However, we first encountered a following technicalFig. 4 Actin filaments gliding over a surface that are densely coatedwith myosin V. The number attached to each image indicates the time(s) elapsed after imaging began. Scale bar, 200 nm; imaging rate, 1 s/
frame

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Pflugers Arch - Eur J Physiol (2008) 456:211–225

Fig. 5 Myosin V in ATP. The number attached to each image
indicates the time (s) elapsed after imaging began. The head portion of
myosin V molecule, which is circled, seems to change its conforma-tion from an extended form to a folded form. ATP concentration is
2 μM. Imaging rate, 0.5 s/frame; scan range, 300 nm

problem. An intense flash illumination by a 355-nm lasercaused large bending of the cantilever due to the photothermal expansion effect. This caused the tip to hit thesubstratum with a strong force, leading to tip damage. To
solve this problem, attenuated high-frequency laser pulses(~50 kHz) were applied, while the y-scanner was being
scanned towards the starting point after the completion of
one frame acquisition. In addition, during this period, thesample stage was withdrawn from the cantilever tip. This
combined method allowed us to observe the rotational
movement of the myosin V head around the head–neckjunction that occurred immediately after UV application(Fig. 6). We also applied this method to observe the heightchanges in GroEL upon binding with ATP and binding with
GroES.
Single-headed dynein C from chlamydomonous flagellahas a ring-shaped head from which a stem and a stalk
protrude. The stalk interacts with microtubules at its
globular distal end. Previously, electron micrographs ofdynein C revealed that the angle between the stalk and thestem differed between the no-nucleotide state and the ADPVi bound state [63]. We confirmed this result using AFM inaqueous solutions. As dynein C has a relatively highATPase rate even in the absence of microtubules, it mustbe easy to distinguish the ATPase-driven conformationalchange from its Brownian motion. In fact, high-speed AFM
imaging of dynein C alone in the presence of ATP revealed
that the stem moved back and forth between two positions
while the stalk and head were stationary [22, 23]. The two
positions approximately corresponded to the nucleotide-freeand ADP-Vi bound states. Besides, the frequency of thestem movement was similar to the ATPase rate. Therefore,

it seems certain that the observed periodic stem movement
is actually driven by the ATPase reaction.
Recent imaging studies
We learned from early imaging studies that reduction in thetapping force was essential to the establishment of highspeed AFM truly useful in biological research. To makedynamic PID control more effective, we developed a lownoise sensor amplifier. Due to the low noise, we could use
an amplitude set point of r>0.9, which made it possible toobserve actin filament gliding on a mica surface sparsely
coated with myosin V (Fig. 7). The bound heads of myosin
V were oriented along one direction [22, 23], like the wellknown arrowhead structure in muscles. From this structure,
the sliding actin filament polarity was identified, and thus,
it was shown that this filament moved toward the minusend (i.e., natural direction). However, we could not detectlarge conformational changes in the bound heads while theactin filament was moving unidirectionally.
In spite of this successful observation, we were unable toobserve the processive movement by single myosin V
molecules along actin filaments. In low ionic strength
solutions, myosin V tended to attach to the mica surface. Inhigh ionic strength solutions, it was nearly free from thesurface, but the affinity of myosin V for actin was lowered.
To circumvent this problem, we reduced the cantilever-free
oscillation amplitude from the usual ~5 to ~1 nm, thereby,
sacrificing the feedback bandwidth. With such a small
amplitude and in a high ionic strength solution containing a
low concentration of ATP, the processive movement was
finally captured on video at 0.1 s/frame. The two heads of a

Fig. 6 Myosin V head bending upon UV-flash photolysis of caged ATP. The number attached to each image indicates the time (s) elapsed after
imaging began. UV flash was applied between the first and second images. Scale bar, 30 nm; imaging rate, 80 ms/framePflugers Arch - Eur J Physiol (2008) 456:211–225

Fig. 7 Three-dimensional images of actin filament sliding movement
captured by high-speed AFM. The number attached to each image
indicates the time (s) elapsed after imaging began. Scale bar 30 nm,
imaging rate, 180 ms/frame

myosin V molecule were changing the leading and trailing
positions alternately with a walking stride of ~72 nm, i.e.,
in a hand-over-hand manner as demonstrated by singlemolecule fluorescence microscopy [64–67]. High-speed
AFM revealed the movement in greater detail than thatshown by fluorescence microscopy. The lead lever-arm
bent just before the rear head detached from the actin
filament, as previously shown by electron microscopy [68].
The detached rear head rotated around the junction between
the two lever arms and then attached to a frontward actin.
Just after the attachment, the new lead head moved farther
by ~5 nm along the actin filament. In the absence of
nucleotide or in the presence of ADP, only one head of amyosin V molecule attached to an actin filament. From theorientation of the bound head, it was clear that the boundhead was in the trailing position. On the other hand, in thepresence of a medium concentration of AMP-PNP, bothheads were associated with an actin filament for a longtime. This indicates that on binding to AMP-PNP, the leadhead conformation adapts itself so that it is able to bind toactin by rotating around the head–neck junction. Therefore,
with myosin V that is bound to an actin filament at both theheads in an ATP-containing solution, the trailing head must
contain ADP (or no nucleotide in a low ATP concentration),and the lead head must contain ATP or ADP-Pi. After thephosphate is released, the lead head rotates back to theprevious orientation, an orientation unfavorable for actin219

binding. However, the lead head cannot dissociate fromactin because the ADP-bound state has a high affinity for
actin. This energetically unfavorable conformation bends
the lead neck forward, thereby pulling the trailing head, sothat it detaches from actin and then being conveyed
frontward. Using these mechanical processes along thechemical reaction pathway, the two heads alternate theirpositions to walk along an actin filament.
Another example of biomolecular processes successfully
captured is GroEL–GroES association–dissociation cycles
controlled by the ATPase reaction. GroEL consists of 14
identical ATPase subunits that form two heptameric rings
stacked back to back [69, 70]. From a series of biochemical
studies [71, 72], it has been shown that there is positive
cooperativity for binding and hydrolysis of ATP in the samering and negative cooperativity between the two rings. Dueto this negative cooperativity, it has been considered that
GroEL binds with GroES at one ring while releasing
GroES from the other [73–75]. This alternate on–off
switching seems to exclude the concomitant binding of
GroES to both rings of GroEL. However, this issue hasremained controversial [76–80], because some electron
micrographs have shown a complex of GroES–GroEL–
GroES with a football shape. To place GroEL on asubstratum in a side-on orientation so that both the rings
are accessible to GroES, we prepared GroEL that were
biotinylated at the equatorial domains [81]. The GroEL wasattached to streptavidin 2D crystal sheets prepared onplanar lipid bilayers containing a biotin lipid. We could
capture the alternate on–off events. Surprisingly, before thealternate switching took place, a football structure appearedwith a high probability. Thus, the high-speed AFM
observation clearly solved the controversial issue.
Imaging with commercial high-speed AFM
Infinitesima (Oxford, U.K.) has recently produced VideoAFM™ equipped with a fast x-scanner made of a quartz
tuning fork, based on the studies by Miles’ group [57, 58].
A fast feedback system is not implemented to thiscommercial high-speed AFM. Although its marketing is
restricted only in Japan at present, Olympus (Tokyo, Japan)
has produced Nano Live Vision™ (Olympus, Tokyo,
Japan), which is a modified version of the microscope
developed by Ando’s group [17]. Users of the latter productrecently filmed dynamic processes of GroEL–GroES
interaction that were regulated by the ATPase reaction of
GroEL immobilized onto mica surface in an end-up
orientation [82]. Moreover, they observed formation anddissociation of a streptavidin-biotinylated DNA complex[83] and one-dimensional diffusion of a restriction enzyme
along a DNA strand which was followed by the cleavage
reaction [84]. A most recent study successfully captured

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translocation and looping of DNA by a type III restriction
enzyme, EcoP15I [85].

Substratum
The choice of the substratum on which a sample is placed isvery important to the AFM observation of biomolecular
processes. The preparation of substrata with very smallroughness have been explored extensively for still-image
observations [86]. However, these preparations have mainly
been devised to attach samples firmly onto them. Therefore,
they are often useless for visualizing biomolecular processes. The visualization of biomolecular processes requires thesubstratum to have various properties; for example, (a)appropriate binding affinity for the sample to ensure that its
physiological function is retained, (b) selective attachmentof a specific component in a multicomponent sample, and(c) attachment of molecules in the desired orientation. Mica(natural muscovite or synthetic fluorophlogopite) has beenfrequently used as a source substratum due to its surface
flatness at the atomic level over a large area. It has net
negative charges and therefore is quite hydrophilic. A baremica surface adsorbs various proteins by electrostaticinteraction. Except for some cases (such as GroEL
attachment in an end-up orientation), its binding orientation
is not unique, and selective attachment of a specific species
is not expected.
If possible, it is best to have, in the first step, a surface
on which the sample never attaches. Then, this surface ismodified so that only a specific species of molecules can
attach to it through a linker in a desired orientation. A
membrane surface with zwitterionic polar head groups suchas phosphatidyl choline (PC) and phosphatidyl ethanolamine (PE) is known to resist protein adsorption [87, 88].
Streptavidin is also useful for specific attachment of
biotinylated protein because it also resists nonspecificprotein adsorption. Mica-surface-supported planar lipid
bilayers [89] can be easily formed from liposomes as theirstrong hydrophilic interaction disrupts the liposomes [90].
Various lipids with functional groups that are attached topolar groups (e.g., biotin attached to PE, Ni-NTA attachedto GS) are available. They enable the specific attachment ofproteins labeled with biotin or his-tags onto planar lipid
bilayers. DOPC lipid is useful for the preparation of
streptavidin 2D crystals when it is used together withbiotinylated lipids [91, 92]. It contains an unsaturatedhydrocarbon on each of the two alkyl chains, which causes
the bending of the chains and, therefore, weakens theinteraction between neighboring DOPCs. This weak interaction lowers the phase-transition temperature of DOPC
lipid down to approximately −20°C, thereby affording large
fluidity to the planar bilayer at room temperature andPflugers Arch - Eur J Physiol (2008) 456:211–225

facilitating 2D-crystal formation of streptavidin (Fig. 8a).
The densely packed streptavidin does not diffuse easily. If
less diffusibility is necessary, the packed streptavidin can becross-linked using glutaraldehyde, which does not influence
its ability to bind to biotin. We used this surface for theselective attachment of biotinylated GroEL in a side-onorientation. DPPC contains no unsaturated hydrocarbons in
the alkyl chains; therefore, its phase-transition temperature
is high (~41°C), and it is appropriate for preparing planar
bilayers with low fluidity. For example, when planar
bilayers are formed with DPPC at a high temperature(~60°C) together with a certain fraction of DPPE-biotin,
streptavidin, which is sparsely attached to the surface,hardly diffuses at room temperature (Fig. 8b). When
DOPE-biotin is used together with DPPC, the sparsely
attached streptavidin diffuses at a moderate rate (Fig. 8c).

Fig. 8 Streptavidin on mica-supported planar lipid bilayers. (a)Streptavidin 2D crystals formed on bilayers of DOPC+DOPE-biotin.
Scale bar, 30 nm. (b) Streptavidin on bilayers of DPPC+DPPE-biotin.
Scale bar, 40 nm: imaging rate, 0.18 s/frame. (c) Streptavidin on
bilayers of DPPC+DOPE-biotin. Scale bar, 40 nm; imaging rate,
0.18 s/framePflugers Arch - Eur J Physiol (2008) 456:211–225

Future prospects
Speed and interaction force
As demonstrated in the successful imaging mentioned
above, high-speed AFM is becoming a useful tool inbiological sciences, particularly in the study of themolecular processes of proteins. To expand the scope of
biological samples that can be studied by high-speed AFM,
the imaging rate must be enhanced further and the tip–
sample interaction force must be significantly reduced. At
present, the cantilever is the main factor that limits boththese capabilities. Most advanced small cantilevers developed by Olympus seem to have almost achieved theultimate goal of balancing a high resonant frequency anda small spring constant. The question therefore is how thisseemingly impenetrable level can be surpassed.
Tip–sample interaction has been detected by measuring
the changes in the amplitude, phase, or resonant frequencyof an oscillating cantilever. With the small quality factor of
the cantilever that is necessary for obtaining a fast response,
these three quantities are not sensitive to the impulsive
interaction. In tapping mode AFM, where the excitation
frequency is fixed, a change in the resonant frequency isdetected as the phase shift relative to the excitation signal.
For a cantilever with a low quality factor, the phase does
not change sharply as a function of the resonancefrequency, resulting in low sensitivity of the phase to theinteraction. The influence of an impulse (~peak force×
interaction time) on the cantilever is distributed overharmonic frequencies (integral multiples of the fundamental
frequency). The cantilever oscillation gain is lower athigher harmonic frequencies, which gives rise to the low
sensitivity of the amplitude. As the impulsive force isexerted transiently in a short time, its peak force isrelatively large. This means that the peak force must be ahighly sensitive quantity. Although the force F(t) cannot be
detected directly and precisely, a rough estimate of F(t) canbe obtained from the cantilever’s oscillation waves z(t) by
putting z(t) into the cantilever motion equation and thensubtracting the excitation signal (i.e., inverse determination
problem; [93]). Here, we do not need to resort thedifferential of z(t), a process that yields noisy signals. An
operation where the phase of each Fourier decomposed
harmonic signal is shifted by π/2 and then multiplied by anappropriate gain is identical to differentiation. To make thismethod effective, the cantilever’s oscillation signal must be
detected with a wide bandwidth (at least up to 4×fc), andfast analog or digital calculation systems are necessary forconverting z(t) to F(t). In addition, a fast peak-hold system
is necessary to capture the peak force. Alternatively, thepeak force can be calculated using only Fourier coefficients, without obtaining the force signal as a function of

221

time. This sensitive method for detecting the tip–sample
interaction should lower the interaction force.
The ultimate way to minimize the interaction force isnoncontact imaging. Nc-AFM has been realized only invacuum by utilizing a significantly large quality factor of thecantilever in vacuum. If high-speed nc-AFM is realized inliquid conditions, we can use stiffer cantilevers with higher
resonant frequencies, which can promise higher imaging
rates. In addition, high-speed nc-AFM will make it possible
to image dynamic molecular processes that occur on living
cell membranes, a process that is, at present, absolutely
impossible, as living cell membranes are extremely soft. As
many important proteins exist on membranes, noncontact
imaging capability greatly expands the usefulness of highspeed AFM in life sciences. Extracellular stimuli (such as
drugs, endogenous ligands, or electric one) are first received
by membranes proteins (receptors or ionic channels) andthen transmitted to the interiors to produce various cellularactions. Various diseases are linked with membrane proteins,sugar chains in membrane glycoproteins, and several
intracellular molecules involved in signal transductions.
Therefore, high-speed nc-AFM will also make a great
contribution to disease detection, diagnosis, and drug
assessment and discovery.
An ultrasonic technique may have the potential of
realizing noncontact imaging; here, the cantilever tip andthe sample are ultrasonically excited at different frequenciesof f1 and f2 (f1, f2 >>fc) with |f1 −f2|~fc. Even when the tip is
not in contact with but close to the sample surface, the two
ultrasound waves must interfere with each other to produce
acoustic waves with a frequency of |f1 −f2| that excites thecantilever. A similar configuration has been used forscanning near-field ultrasound holography (SNFUH),
which has been developed for high-resolution subsurface
imaging [94]. Ion-conductance scanning probe microscopy(ICSPM) already satisfies the noncontact condition [95]. To
materialize high-speed ICSPM, we must find a method toincrease the bandwidth of ion-conductance detection, as
ionic currents through a small pore of the capillaryelectrode are very low.
High-speed AFM for intracellular imaging
SNFUH has already successfully been used in intracellularimaging [94]. In its application, a high-frequency acoustic
wave is launched from under the sample stage, and thiswave then propagates through the sample. Materials withdifferent elastic moduli embedded in the sample modulate
the phase and amplitude of the propagating acoustic waves.
These modulations are reflected on the acoustic interference
that occurs at the cantilever tip and, consequently, arereflected on the cantilever oscillation around its resonant
frequency. It does not seem difficult to combine SNFUH

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with the high-speed scanning techniques developed so far.
This combination will enable the filming of dynamic
processes of intracellular organelles. Although it ispresently unclear whether SNFUH has resolution in thez-direction, the images obtained with ultrasonic waves
launched at different angles should contain information
along the z-axis. Therefore, there may be a way toreconstitute a 3D image from multiple images obtained at
different launching angles. High-resolution 3D imaging of
intracellular structures in real time will revolutionize cellbiology and physiology. Various intracellular processes inliving cells and organisms that take place spontaneously or
responding to extracellular stimuli may be able to be
examined by high-speed SNFUH combined with optical
microscopy.
High-speed recognition AFM
Hinterdorfer and his colleagues recently developed a recognition imaging technique [31, 32]. In their technique, acantilever tip that is functionalized with a probe molecule
through a flexible short-linker is oscillated at a frequency
below resonance. Its oscillation is split into approaching andwithdrawing parts. The oscillation signals in the approaching part reflect the repulsive tip–sample interaction, providing topography images. The signals in the withdrawing partreflect the association events between the probe moleculeand its counterpart, providing recognition images. The
effective concentration of a probe molecule attached toa cantilever tip depends on the tether length. With atether length of 2 nm, the concentration is ~50 mM,
which is high enough for the association reaction to takeplace in 20 μs for a system with a typical association rate
constant of 1×106 M−1 s−1. This suggests that recognition
imaging at a moderate rate (~0.4 s/frame) is possible solong as the cantilever oscillation amplitude is small.
Another type of recognition imaging may be derived fromphase-contrast imaging [53] using a cantilever tip functionalized with a specific probe molecule. Its phasecontrast image will reveal the arrangement of the specificmolecules in the sample.
High-resolution and high-speed AFM
Atomic resolution had been attained with AFM only inultrahigh vacuum environments. However, recent studies
with frequency-modulation AFM (FM-AFM) haveachieved atomic resolution in a liquid environment byminimizing the sensor noise and using a small amplitude of
cantilever oscillation [48, 96]. In addition, the highresolution imaging capability of FM-AFM was also
demonstrated by successful imaging of individual waterlayers weakly interacting with lipid membrane surfaces

Pflugers Arch - Eur J Physiol (2008) 456:211–225[97]. In these studies, conventional cantilevers with arelatively small quality factor (~6) were used. As a muchlarger shift in the resonant frequency is expected to occur
with small cantilevers, it seems possible to achieve subnanometer resolution as well with high-speed tapping mode
AFM equipped with a fast frequency-shift (phase-shift)detector [53] and a sharp cantilever tip. Our recent
preliminary experiments filmed atomic resolution imagesof a mica surface in water at a frame rate of 37 frames/s,
although further noise reduction remained to be accomplished. AFM with both high-resolution and high-speed
capabilities has enormous potential; imaging of secondary
structures on protein surfaces and their dynamic changes,
mapping of rapid changes in the water layer structure at
protein surfaces, imaging of dynamic lipid–protein interaction on membranes, and detection of small-ligand interaction with its active site on a protein surface. It is impossible
to study these phenomena by using “conventional” highspeed AFM.
Direct seeing at in-liquid nanometer-scale worlds in realtime will markedly change our style of examining biologicalproblems. As discussed above, the usefulness of high-speed
imaging capability of AFM will effectively expand by othertechniques being combined. In the initial stage, high-speed
AFM will be actively used to observe a wide range of dynamic
processes with isolated proteins, protein assemblies, andprotein–DNA complexes. In the next stage, more complex
systems including living cells and organisms will becometargets of high-speed AFM combined with other techniques. In vivo and in vitro visualization of various
processes at the molecular level will become possible;
responses of membrane receptors to stimuli, nuclear
envelope formation and disassembly, chromosome replication and segregation processes, phagocytosis, proteins
synthesis in the endoplasmic reticulum, targeting processes of synthesized proteins through the Golgi apparatus,and etc. Thus, high-speed AFM-based visualization
techniques have a great potential of innovating not only
in biochemistry and biophysics but also in cell biology,
physiology, and pharmaceutical and medical sciences. To
open up such unprecedented fields, steady efforts have to
be carried out towards expanding the capability of highspeed AFM and the related techniques.

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