Available online at www.sciencedirect.com

Progress in Surface Science 83 (2008) 337–437
www.elsevier.com/locate/progsurf

Review

High-speed atomic force microscopy
for nano-visualization of dynamic
biomolecular processes
Toshio Ando a,b,c,*, Takayuki Uchihashi a,b, Takeshi Fukuma c,da

Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japanb
CREST, JST, Sanbon-cho, Chiyoda-ku, Tokyo 102-0075, Japanc
Frontier Science Organization, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japand
PRESTO, JST, Sanbon-cho, Chiyoda-ku, Tokyo 102-0075, Japan

Commissioning Editor: J. Yoshinobu

Abstract
The atomic force microscope (AFM) has a unique capability of allowing the high-resolution
imaging of biological samples on substratum surfaces in physiological solutions. Recent technological progress of AFM in biological research has resulted in remarkable improvements in both theimaging rate and the tip force acting on the sample. These improvements have enabled the direct
visualization of dynamic structural changes and dynamic interactions occurring in individual biological macromolecules, which is currently not possible with other techniques. Therefore, high-speed
AFM is expected to have a revolutionary impact on biological sciences. In addition, the recently
achieved atomic-resolution in liquids will further expand the usefulness of AFM in biologicalresearch. In this article, we ﬁrst describe the various capabilities required of AFM in biological sciences, which is followed by a detailed description of various devices and techniques developed forhigh-speed AFM and atomic-resolution in-liquid AFM. We then describe various imaging studies
performed using our cutting-edge microscopes and their current capabilities as well as their limitations, and conclude by discussing the future prospects of AFM as an imaging tool in biologicalresearch.
Ó 2008 Elsevier Ltd. All rights reserved.

*
Corresponding author. Address: Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa
920-1192, Japan. Tel.: +81 76 264 5663; fax: +81 76 264 5739.
E-mail address: tando@kenroku.kanazawa-u.ac.jp (T. Ando).

0079-6816/$ - see front matter Ó 2008 Elsevier Ltd. All rights reserved.
doi:10.1016/j.progsurf.2008.09.001

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T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

Keywords: AFM; High-speed AFM; Atomic force microscopy; Imaging; Visualization; Biomolecular processes;
Dynamic processes; Protein; Atomic-resolution; Cantilevers; Temporal resolution; BiomoleculesContents
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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Basic principle of AFM and various imaging modes . . . . . . . . . . . . . . . . . . . . . . . . .
History of AFM studies on biomolecular processes. . . . . . . . . . . . . . . . . . . . . . . . . .
Requirements for high-speed AFM in biological research . . . . . . . . . . . . . . . . . . . . .
Feedback bandwidth and imaging rate. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.1. Image acquisition time and feedback bandwidth . . . . . . . . . . . . . . . . . . . . . . .
5.2. Phase delays in open-loop and closed-loop . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.3. Feedback bandwidth as a function of various factors . . . . . . . . . . . . . . . . . . . .
5.4. Parachuting time . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.5. Integral time in the PID feedback control . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.6. Refinement of analytical expressions for sp and sI . . . . . . . . . . . . . . . . . . . . . .
5.7. Summary of guidelines for developing high-speed AFM . . . . . . . . . . . . . . . . . .
Optimization of devices for high-speed AFM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.1. Cantilevers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.2. Cantilever tip . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.3. Optical beam deflection detector for small cantilevers. . . . . . . . . . . . . . . . . . . .
6.4. Tip–sample interaction detection methods . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.4.1. Amplitude detectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.4.2. Force detectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.5. High-speed scanners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.5.1. Counterbalance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.5.2. Mechanical scanner design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.5.3. Caution regarding hydrodynamic effects. . . . . . . . . . . . . . . . . . . . . . . .
6.6. Active damping. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6.1. Notch filtering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6.2. Feedback control for active damping . . . . . . . . . . . . . . . . . . . . . . . . . .
6.6.3. Feed-forward control for active damping . . . . . . . . . . . . . . . . . . . . . . .
6.6.4. Practice of active damping of the scanner vibrations . . . . . . . . . . . . . . .
6.7. Dynamic PID control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.7.1. Dynamic PID controller . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.7.2. Performance of dynamic PID control . . . . . . . . . . . . . . . . . . . . . . . . . .
6.8. Drift compensator . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.9. Line-by-line feed-forward control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.9.1. Feed-forward controller . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.9.2. Performance of feed-forward compensation . . . . . . . . . . . . . . . . . . . . .
6.10. Photothermal actuation of cantilevers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.11. Data acquisition system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
High-speed phase detector . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Devices for atomic-resolution in liquids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.1. Operation mode . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.2. Cantilever stiffness . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.3. Oscillation amplitude. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
8.4. Deflection sensor noise . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Imaging of dynamic biomolecular processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.1. Early imaging studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.1.1. Initial attempts and lessons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.1.2. Flash photolysis combined with high-speed AFM . . . . . . . . . . . . . . . . .
9.1.3. Dynein C. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.2. Recent imaging studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.2.1. Myosin V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.2.2. Chaperonin GroEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
9.2.3. Lattice defects in 2D protein crystals . . . . . . . . . . . . . . . . . . . . . . . . . .
9.2.4. Intrinsically disordered regions of proteins . . . . . . . . . . . . . . . . . . . . . .
9.3. Imaging using commercially available high-speed AFM . . . . . . . . . . . . . . . . . .
Substrata for observing dynamic biomolecular processes . . . . . . . . . . . . . . . . . . . . .
High-speed phase-contrast imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
11.1. Phase mapping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
11.1.1. Compositional mapping on blended polymers . . . . . . . . . . . . . . . . . . .
11.1.2. Dependence of phase-contrast on detection timing . . . . . . . . . . . . . . . .
11.1.3. Phase imaging of myosin filament . . . . . . . . . . . . . . . . . . . . . . . . . . .
11.2. Phase-modulation imaging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
11.2.1. Distance dependence of phase-shift signal . . . . . . . . . . . . . . . . . . . . . .
11.2.2. PM imaging of GroEL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Atomic-resolution imaging in liquids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
12.1. Atomic-resolution imaging by FM–AFM in liquid . . . . . . . . . . . . . . . . . . . . . .
12.2. Biological applications of FM–AFM. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
12.2.1. Direct imaging of intrinsic hydration layers. . . . . . . . . . . . . . . . . . . . .
12.2.2. Direct imaging of lipid-ion networks . . . . . . . . . . . . . . . . . . . . . . . . .
Future prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
13.1. Imaging speed, interaction force, and noncontact imaging. . . . . . . . . . . . . . . . .
13.2. High-speed AFM for intracellular imaging . . . . . . . . . . . . . . . . . . . . . . . . . . .
13.3. High-speed-recognition AFM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
13.4. Ultra-high-resolution high-speed AFM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
13.5. High-speed AFM combined with optical microscope . . . . . . . . . . . . . . . . . . . .
Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Appendix A. Supplementary data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1. Introduction
The atomic force microscope (AFM) was invented in 1986 by Binnig et al. [1], four
years after the invention of the scanning tunneling microscope (STM) [2]. Unlike STM
or electron microscopy, AFM is unique in its ability to observe insulating objects, andhence, opened the door to the visualization of nanometer-scale objects in liquids. This
unique capability was received with excitement by researchers of biological sciences as biomolecules only show vital activities in aqueous solutions. Before the AFM era, the highresolution visualization of individual biopolymers (proteins, DNA) was only possible byelectron microscopy in a vacuum environment. Many AFM imaging studies have been
performed on various biological samples to explore the potential of this new microscope.

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Through these studies, techniques for obtaining high-resolution images have been developed. However, AFMs unique capability, i.e., the high-resolution visualization of ‘‘active
biomolecules” in solutions, does not seem to have contributed signiﬁcantly to answering
many biological questions.
One of the essential features of biological systems is ‘‘dynamics”. The functions of biological systems are produced through dynamic processes that occur in biopolymers, biosupramolecules, organelles, and cells. Therefore, what is required of AFM for biologicalsciences is the ability to rapidly acquire successive high-resolution images of individual
biomolecules at work. This is solely because this type of imaging is impossible using othertechniques. However, the imaging rate of conventional AFM is too slow to observe
dynamic behavior of active biomolecules. Thus, endowing AFM with high-speed imaging
capability is expected to have a revolutionary impact on biological sciences.
Over the past decade, various eﬀorts have been directed toward increasing the AFM
imaging rate. The most advanced high-speed AFM can now capture images at 30–
60 ms/frame over a scan range of 250 nm with 100 scan lines. Importantly, the tip–
sample interaction force has been greatly reduced without sacriﬁcing the imaging rate,
so that weak dynamic interactions between biological macromolecules are not disturbed
signiﬁcantly. Although the number of published reports is still limited, dynamic biomolecular processes have been successfully captured on video, some of which have revealed thefunctional mechanisms of proteins. As demonstrated by these studies, the newly acquired
high-speed imaging capability has greatly heightened the value of AFM in biological sciences. High-speed bio-AFM will be established soon and commercially available in a few
years, which is expected to increase the user population quickly. In this article, we attempt
to provide the potential users and developers with comprehensive descriptions of highspeed AFM including various techniques involved in the instrumentation, applications
to biological studies, current capabilities and limitations, and future prospects. In addition, we describe the recent progress in increasing the spatial resolution to the atomic levelfor frequency-modulation in-liquid AFM. Concise reviews on high-speed AFM were previously presented [3,4]. AFM movies placed at http://www.s.kanazawa-u.ac.jp/phys/biophys/roadmap.htm and at the publisher’s web site will give readers an indication of thepower of this state-of-the-art microscope.
2. Basic principle of AFM and various imaging modes
A typical setup of tapping-mode AFM is depicted in Fig. 1. The AFM is a sort of a‘‘palpation” microscope. It forms an image by touching the sample surface with a sharp
tip attached to the free end of a soft cantilever while the sample stage is scanned horizontally in 2D. Upon touching the sample, the cantilever deﬂects. Among several methods ofsensing this deﬂection, optical beam deﬂection (OBD) sensing is often used because of its
simplicity; a collimated laser beam is focused onto the cantilever and reﬂected back into
closely spaced photodiodes [a position-sensitive photodetector (PSPD)] whose photocurrents are fed into a diﬀerential ampliﬁer. The output of the diﬀerential ampliﬁer is proportional to the cantilever deﬂection. During the raster scan of the sample stage, the detected
deﬂection is compared with the target value (set point deﬂection), and then the stage is
moved in the z-direction to minimize the error signal (the diﬀerence between the detected
and set point deﬂections). This closed-loop feedback operation can maintain the cantileverdeﬂection (hence, the tip–sample interaction force) at the set point value. The resulting 3D

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Fig. 1. Schematic presentation of the tapping-mode AFM system. In the constant-force mode, the excitation
piezoelectric actuator and the RMS-to-DC converter are omitted [87].

movement of the sample stage approximately traces the sample surface, and hence, a topographic image can be constructed using a computer, usually from the electric signals thatare used to drive the sample stage scanner in the z-direction. Sometimes, the topographic
image is constructed using values obtained by summing the electric signals used for driving
the z-scanner and the error signals with an appropriate weight function. This method cangive a more accurate topographic image than the former method. In the operation mode(constant-force mode; one of DC modes or contact-modes) described above, the cantilevertip, which is always in contact with the sample, exerts relatively large lateral forces to thesample because the spring constant of the cantilever is large in the lateral direction.
To avoid this problem, tapping-mode AFM (one of dynamic-modes) was invented [5],in which the cantilever is oscillated in the z-direction at (or near) its resonant frequency.
The oscillation amplitude is reduced by the repulsive interaction between the tip andthe sample. Therefore, this mode is also called the amplitude-modulation (AM) mode.
The amplitude signal is usually generated by an RMS-to-DC converter and is maintained
at a constant level (set point amplitude) by feedback operation.
In AM–AFM, the cantilever oscillation amplitude decreases not only by the energy dissipation due to the tip–sample interaction but also by a shift in the cantilever resonant frequency caused by the interaction [6–8]. As the excitation frequency is ﬁxed at (or near) the
resonant frequency, this frequency shift produces a phase shift of the cantilever oscillation
relative to the excitation signal. When this phase shift is maintained by feedback operation

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and an image is constructed from the electric signals used for driving the z-scanner, thisimaging mode is called the phase-modulation (PM) mode. Alternatively, we can constructa phase-contrast image from the phase signal, while maintaining the amplitude at a constant level by feedback operation. More details of the PM mode and phase-contrast imaging are given in Chapters 7 and 11. Instead of using a ﬁxed frequency, it is possible to set
the excitation frequency automatically to the varying resonant frequency of the cantileverusing a self-oscillation circuit [9,10]. In this case, the phase of the cantilever oscillation relative to the excitation signal is always maintained at 90°, and the resonant frequency
shift is maintained at a constant level by feedback operation. This mode is called the frequency-modulation (FM) mode and is described in Chapters 8 and 12.
3. History of AFM studies on biomolecular processes
In this chapter, we brieﬂy describe the history of bio-AFM, focusing on studies on biological processes without covering a wide range of bio-AFM studies (more comprehensive
descriptions on the bio-AFM history are given in a recent article [11]). The history willshow that the observation of dynamic biomolecular processes started soon after the invention of AFM, whereas studies with the aim of realizing the fast imaging capability were left
until later.
In 1987, the in-liquid observation ability of AFM was demonstrated [12]. Interestingly,
the liquid used was not water but paraﬃn oil, as the surface of sodium chloride crystal wasobserved. Around 1988, cantilevers manufactured using microfabrication techniques
became available [13], and the OBD method for detecting cantilever deﬂection was introduced [14]; these devices promoted the AFM imaging of biological samples such as aminoacid crystals [15], lipid membranes [16], biominerals [17], and IgG [18]. Even at this very
early stage, Hansma and colleagues attempted to observe the dynamic behavior of biological samples in action. For example, they observed the ﬁbrin clotting process initiated by
the digestion of ﬁbrinogen with thrombin at 1 min intervals [19]. Some trial observationsof dynamic biological processes were also performed on the viral infection of isolated cells[20] and antibody binding to an S-layer protein [21]. We can imagine that it must havebeen diﬃcult at this early stage to observe these dynamic processes, as only the contactmode was available (tapping-mode was invented in 1993 [5]); In the constant-force mode,biomolecules weakly attached to a surface are easily dislodged by the scanning tip. At thistime, more eﬀort was directed toward attaining suitable conditions under which high-spatial-resolution images could be obtained [22–34]. Using the constant-force mode, Engeland colleagues continuously obtained very beautiful high-resolution images of membraneprotein systems such as gap junctions [22], E. coli OmpF porin [30,31], aquaporin-1 in red
blood cells [32], and bacteriorhodopsin [33,34].
Soon after the tapping-mode was invented, this mode was shown to be operational inliquid-environment [35,36]. The acoustic method, which is now often used to excite cantilevers, was introduced [35]. Later, it was shown that this mode also produces high-resolution images of membrane proteins [37]. The tapping-mode enabled the imaging of
biological samples weakly attached to a substratum, which led to a moderate revival ofresearch activity on the exploration of biological processes, although the imaging rate
was as low as before. For example, in 1994, Bustamante and colleagues imaged DNA diffusion on a mica surface [38] and DNA bending upon binding to k Cro protein [39], andHansma and colleagues imaged DNA digestion with DNase [40] and the DNA–RNA

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polymerase binding process [41]. These two groups continued these studies and obtained
time-lapse images (30 s intervals) of the RNA transcription reaction between DNA andRNA polymerase [42] and of the 1D diﬀusion of RNA polymerase along a DNA strand[43]. Other examples of dynamic processes that have been imaged are the proteolytic cleavage of collagen I by collagenase [44] and nuclear-pore closing by exposure to CO2 [45].
Attempts to increase the scan speed of AFM were initiated by Quate and colleagues(e.g., [46–48]). Their aim was to increase the speed of lithographic processing and the evaluation of a wide surface area of hard materials. For this purpose, they developed cantilevers with integrated sensors and/or actuators, and cantilever arrays with self-sensing andself-actuation capabilities. It was only possible to fabricate these sophisticated cantilevers
with relatively large dimensions, thus the resonant frequency was not enhanced markedly
and the spring constant was large. The insulation coating of the integrated cantilevers thatallows their use in liquids further lowered the resonant frequency. The approach they
employed was adequate for their purposes but unsuitable for the use of AFM in biologicalresearch, as the required conditions for high-speed AFM in the two diﬀerent ﬁelds areoften considerably diﬀerent. Therefore, their line of studies did not result in the realizationof high-speed AFM for biological research. However, note that their approach will also be
useful for high-speed bio-AFM if insulated and integrated small cantilevers with a small
spring constant can be fabricated in the future.
In 1993, the scan speed limit of contact-mode AFM was theoretically analyzed [49],focusing on the relationship between the cantilever’s mechanical properties and the scan
speed. Some eﬀorts aimed at increasing the bio-AFM scan speed were initiated shortly
before 1995. In fact, we started to develop high-speed scanners in 1994 and small cantilevers in 1997. Hansma’s group also started to develop devices for high-speed bio-AFM
around 1995. They presented the ﬁrst report on short cantilevers (23 lm by 12 lm) in
1996 [50], and subsequently a report on fast imaging in 1999, in which small cantilevers
and an optical deﬂection detector [51] designed for the small cantilevers were used toobtain an image of DNA in 1.7 s [52]. The following year, they imaged the formationand dissociation of GroES–GroEL complexes [53]. However, because of the limited feedback bandwidth, this molecular process was traced by scanning the sample stage only inthe x- and z-directions. We reported a more complete high-speed AFM system in 2001[54] and 2002 [55]. In this study, we developed a high-speed scanner, fast electronics, smallcantilevers (resonant frequency, 600 kHz in water; spring constant, 0.1 N/m) and an
OBD detector for the small cantilevers. An imaging rate of 12.5 frames/s was achieved,
and the swinging lever-arm-like motion of myosin V molecules was ﬁlmed as successive
images over a scan range of 240 nm. However, this was only the ﬁrst step in the development of truly useful high-speed AFM for biological sciences.
4. Requirements for high-speed AFM in biological research
Biological macromolecules are highly dynamic. Their functions results from dynamic
structural changes and dynamic interactions with other molecules. Motor proteins transport cargo to their destinations by ‘walking’ along their ﬁlamentous protein tracks [56].
Cytoskeletons undergo polymerization and depolymerization cycles under the action by
regulatory factors [57,58]. Tightly wound chromosomes are unraveled and the exposed
DNA double strands are separated by helicase proteins into single strands for replicationand transcription [59]. The winding and unwinding of DNA produces tension, which

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results in the formation of knots. The knots can be relaxed by the action of topoisomerases[60]; the tense helical strand is cut and thereby freely spins to relieve the tension, and thenthe broken strands are reconnected. A newly synthesized polypeptide is trapped in the cavity of a molecular chaperon, folds into a functional 3D entity, and then detaches into thesolution [61]. Outlined pictures of these dynamic biological processes have been depicted
through many indirect measurements from various angles. However, it is still diﬃcult toobtain detailed pictures of many systems. There are many biomolecular systems remaining
for which even outlines of their dynamic processes have never been obtained.
Dynamic biological processes generally occur on a millisecond timescale. Therefore,ﬁrstly, biological sciences require AFM to have the ability of ﬁlming the dynamic behaviorof a puriﬁed protein weakly attached to a substratum in a physiological solution. The
imaging rate required is at least a few frames/s, and ideally speaking, a few hundred
frames/s. Physiological functions are often produced by the interaction among a few species of molecules. If all the molecules are attached to a substratum, they have almost no
chance of interacting with each other. Therefore, the selective attachment of one species
of molecule to a surface is required. ‘‘Dynamic interaction” implies that the force involvedin the interaction is weak. The force acting in protein–protein interactions approximately
ranges from 1 pN to 100 pN. Even the single ‘‘rigor” complex of a muscle-myosin head
and an actin ﬁlament, which hardly dissociates in equilibrium, is ruptured quickly by apulling force of 15 pN [62]. The force produced by motor proteins during ATP hydrolysis is generally a few piconewtons (e.g., see [63]). Therefore, it is further required thatthe tip–sample interaction force can be maintained at a very small level during imaging.
However, we should note that the mechanical quantity which aﬀects the sample is not
the force itself but force impulse, i.e., the product of force and the time over which theforce acts. In tapping-mode high-speed AFM, the time of force action is short, and therefore, a relatively large peak force (<20 pN) would not aﬀect the sample signiﬁcantly.
When a multicomponent system contains diﬀerent species of proteins with a similar
shape and size, we need a means of distinguishing them. They may be distinguished byvery high-resolution imaging. However, in a sample whose dynamic processes are to beobserved, the movement of protein molecules is caused not only by physiological reactions
but also by thermal agitation. Thus, it is often diﬃcult to realize very high-resolution forsuch a moving sample. We need a high-speed recognition imaging technique to placemarks on a speciﬁc species of protein molecules while capturing the topographic images.
High-speed AFM will become more useful in biological sciences if it attains the capability of observing the ﬁne structures on living cell membranes. A large number of membrane proteins play important roles in the functions of cells. However, little is known
about their dynamic molecular processes. At present, AFM cannot be applied to theobservation of ﬁne structures on living cell membranes, as the membranes are extremely
soft compared with available cantilevers. Thus, it is necessary for high-speed AFM to havethe ability of noncontact imaging in liquids. Recent progress in the FM–AFM of liquids
has enabled the high-resolution imaging of individual hydration layers on lipid membranes[64] (see Section 12.2). This successful imaging of individual hydration layers suggests thatthe tip–sample interaction force must have been very weak. In addition, the high-resolution was attained using cantilevers with relatively small quality factors. Therefore, itappears to be possible to realize high-speed quasi-noncontact AFM using the FM modeor high-speed noncontact AFM (nc-AFM) using completely diﬀerent modes. We will discuss this issue in Section 13.1.

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The spatial resolution of optical microscopy is not suﬃcient for directly observing the
dynamic processes of intracellular organelles. A recently developed method of ﬂuorescencemicroscopy, stimulated-emission-depletion (STED) ﬂuorescence microscopy [65,66], has aspatial resolution of 20 nm. This high-spatial-resolution has to be compromised to attainhigh temporal resolution as the number of photons collected is limited [67]. The AFM hasgenerally been considered a microscope for observing surfaces. However, it was recently
demonstrated that an ultrasonic technique combined with AFM allows us to observe subsurface structures [68]. Hence, the realization of high-speed ‘‘diaphan-AFM”, which
would enable observing the dynamic processes of organelles in living cells, is requiredfor biological sciences.
5. Feedback bandwidth and imaging rate
In the development of high-speed AFM apparatuses, it is important to have practicalguidelines that can quantitatively indicate how each device performance aﬀects the scan
speed and the imaging rate. An early theoretical consideration of the scan speed limit incontact-mode AFM was given in [49]. Concerning tapping-mode AFM, the dependenceof feedback bandwidth on various factors has been qualitatively described [69]. Numericalsimulations were also performed for this purpose, including the eﬀect of the dynamics ofthe tip–sample interaction [70]. However, they are not suﬃcient as practical guidelines. In
this chapter, we derive the quantitative relationship between the feedback bandwidth andthe various factors involved in AFM devices and the scanning conditions, based on an idea
previously presented for the derivation [71].
5.1. Image acquisition time and feedback bandwidth
Supposing that an image is taken in time period T over the scan range W  W with N
scan lines, then the scan velocity Vs in the x-direction is given by Vs = 2WN/T. Assumingthat the sample has a sinusoidal shape with periodicity k, the scan velocity Vs requires a
feedback operation at frequency f = Vs/k to maintain the tip–sample distance. The feedback bandwidth fB should be greater than or equal to f and can therefore be expressed asfB P 2WN =kT :ð1Þ

Eq. (1) gives the relationship between the image acquisition time T and the feedback bandwidth fB. For example, for T = 30 ms with W = 240 nm and N = 100, the scan velocity is
1.6 mm/s. When k is 10 nm, fB P 160 kHz is required to obtain this scan velocity. Notethat the maximum scan velocity achievable under a given feedback bandwidth depends
on the spatial frequency contained in the sample topography.
5.2. Phase delays in open-loop and closed-loop
To determine how the open-loop phase delay is related to the closed-loop phase delay,
here we consider a simple feedback loop in constant-force-mode AFM (Fig. 2). The sample height variation under the cantilever tip, uin ðtÞ, introduced by the x-scan of the sample
stage is considered as the input signal to this system, and the z-scanner displacement,
uout ðtÞ, is considered as the output signal. The time dependence of the closed-loop
input–output relationship is represented by a transfer function K(s) expressed as

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Fig. 2. Block diagram for the feedback loop of constant-force mode AFM.

KðsÞ ¼

T ðsÞ
;1 þ T ðsÞð2Þ

where T(s) is the open-loop transfer function given by C(s)A(s)H(s)G(s) (see Fig. 2). The
frequency dependence of T(s) is given by T0(x)exp[iu(x)], where T0(x) and u(x) are thegain and phase delay, respectively. Therefore, the frequency dependence of K(s) is expressed asKðixÞ ¼

ðT 0 cos uðxÞ þ T 20 Þ þ iT 0 sin uðxÞ:1 þ 2T 0 cos uðxÞ þ T 20

ð3Þ

Thus, the closed-loop phase delay U(x) and the gain K0(x) are, respectively, given bytan UðxÞ ¼

 sin uðxÞ;T 0 ðxÞ þ cos uðxÞ

ð4Þ

and
qﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
K 0 ðxÞ ¼ T 0
1 þ 2T 0 cos uðxÞ þ T 20 :

ð5Þ

When feedback performance is satisfactory and the feedback gain K0(x) is maintained at
1, the open-loop gain is approximately T0(x) = 1/[2cosu(x)]. By substituting this relationship into Eq. (4), we obtain the relationship U(x) = p  2u(x). Here, the phase diﬀerence of ‘‘p” appears because the direction of the z-scanner’s displacement is opposite thatof the variations in the sample height. Thus, we can conclude that the closed-loop phasedelay is approximately twice the open-loop phase delay, provided the feedback gain is
maintained at 1. This is also true in tapping-mode AFM.
5.3. Feedback bandwidth as a function of various factors
From the conclusion obtained above, the time delay in the closed-loop feedback control
can be estimated by summing the time delays that are caused by the devices involved in the
feedback loop. The closed-loop phase delay h [2u(x)] is given by 2  2pfDs, where Ds
is the total time delay in the open-loop and f is the feedback frequency. In tapping-mode
AFM, the main delays are the reading time of the cantilever oscillation amplitude (sd), thecantilever response time (sc), the z-scanner response time (ss), the integral time (sI) of error
signals in the feedback controller, and the parachuting time (sp). Here, ‘‘parachuting”
means that the cantilever tip completely detaches from the sample surface at a steeply

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347

inclined region of the sample, and thereafter, time elapses until it lands on the surface
again. It takes at least a time of 1/(2fc) to measure the amplitude of a cantilever that is
oscillating at its resonant frequency fc. The response time of second-order resonant systems such as cantilevers and piezoactuators is expressed as Q/(pf0), where Q and f0 are
the quality factor and resonant frequency, respectively. The feedback bandwidth is usuallydeﬁned by the feedback frequency that results in a phase delay of p/4. On the basis of thisdeﬁnition, the feedback bandwidth fB is approximately expressed as




fc
2Qc 2Qs fc
1þþþ 2f c sp þ sI þ d ;ð6Þ
fB ¼ a8ppfs
where fs is the z-scanner’s resonant frequency; Qc and Qs are the quality factors of the cantilever and z-scanner, respectively. d represents the sum of other time delays and a represents a factor related to the phase compensation eﬀect given by the D component in the
proportional–integral-derivative (PID) feedback controller or in an additional phase compensator. From Eqs. (1) and (6), we can estimate the highest possible imaging rate in agiven tapping-mode AFM setup by examining the open-loop time delay Ds. However, thisestimation must be modiﬁed depending on the sample to be imaged, because the allowablemaximum phase delay depends on the strength or fragility of the sample.
5.4. Parachuting time
Here, we determine the conditions that cause parachuting, and obtain a rough estimateof the parachuting time and its eﬀect on the feedback bandwidth [71]. The theoreticalresults obtained here are compared with experimental data to reﬁne the analytical expression for the parachuting time.
When a sample having a sinusoidal shape with periodicity k and maximum height h0 is
scanned at velocity Vs in the x-direction, the sample height S(t) under the cantilever tip
varies asSðtÞ ¼h0
sinð2pftÞ;2

ð7Þ

where f = Vs/k. When no parachuting occurs, the z-scanner moves asZðtÞ ¼ 

h0
sinð2pft  hÞ:
2

The feedback error (‘‘residual topography”, DS) is thus expressed ashhDSðtÞ ¼ SðtÞ þ ZðtÞ ¼ h0 sin cos 2pft  :
2
2

ð8Þ

ð9Þ

The cantilever tip feels this residual topography (Fig. 3) in addition to a constant height of
2A0(1  r), where A0 is the free-oscillation amplitude of the cantilever and r is the dimensionless peak-to-peak amplitude set point. When the set point peak-to-peak amplitude isdenoted as As, r = As/(2A0). The maximum extra force exerted onto the sample due tofeedback error corresponds to a distance of h0 sinðh=2Þ. Therefore, an allowable maximum
phase delay ha , which depends on the sample strength, is determined by this distance. The
amplitude set point r is usually determined by compromising two factors: (1) increase intapping force with decreasing r, and (2) decrease in the feedback bandwidth with increasing

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Fig. 3. The residual topography to be sensed by a cantilever tip under feedback control. When the maximum
height of the residual topography is larger than the diﬀerence (2A0  As), the tip completely detaches from thesurface. The untouched areas are show
R t in gray. The average tip–surface separation hdi at the end of cantilever’s
bottom swing is given by hdi ¼ 2t10 t0 0 ½2A0 ð1  rÞ þ h0 sinðU=2Þ cosð2pftÞdt, where t0 ¼ b=2pf (see the text).
This integral results in hdi ¼ 2A0 ð1  rÞðtan b=b  1Þ [71].

r owing to parachuting. Therefore, the allowable maximum extra force approximately
corresponds to  2A0 ð1  rÞ, which gives the relationship of sinðha =2Þ  ð2A0 =h0 Þð1  rÞ.
When DSðtÞ þ 2A0 ð1  rÞ > 0; no parachuting occurs. Therefore, the maximum set
point rmax for which parachuting does not occur is given byrmax ¼ 1 

h0
h
sin :
2A0
2

ð10Þ

Eq. (10) indicates that rmax decreases linearly with h0/2A0 (Fig. 4a) and with phase delay inthe feedback operation (Fig. 4b).
The parachuting time is a function of various parameters such as the sample height h0,
the free-oscillation amplitude A0 of the cantilever, the set point r, the phase delay h, andthe cantilever resonant frequency fc. Its analytical expression cannot be obtained exactly.
As a ﬁrst approximation, we assume that during parachuting, Eq. (9) holds and the z-position of the sample stage does not move. During parachuting, the average separation
between the sample surface and the tip at the end of the bottom swing is given by
2A0(1  r)(tan b/b  1) (see Fig. 3), where b is given byb ¼ cos1 ½2A0 ð1  rÞ=h0 sinðh=2Þ:ð11Þ

The feedback gain is usually set to a level at which the separation distance of 2A0(1  r)
decreases to approximately zero in a single period of the cantilever oscillation. Therefore,
the parachuting time sp is expressed as
sp ¼ ðtan b=b  1Þ=fc :ð12Þ

However, the assumptions, under which the average separation during parachuting wasderived, are diﬀerent from the reality. As mentioned later (Section 5.6), the analyticalexpression for sp should be modiﬁed in light of the experimentally obtained feedback
bandwidth as a function of r and h0/A0.
5.5. Integral time in the PID feedback control
The main component of PID control is the integral operation. It is diﬃcult to theoretically estimate the integral time constant (sI) with which the optimum feedback control is
attained. Intuitively, sI should be longer when a larger phase delay exists in the feedback
loop. In other words, when a larger phase delay exists, the gain parameters of the PID controller cannot be increased. Therefore, sI must be proportional to the height of residual

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349

Fig. 4. The maximum set point rmax that allows the cantilever tip to trace the sample surface without complete
detachment from the surface. (a) Dependence of rmax on the ratio of the sample height h0 to the peak-to-peakamplitude 2A0 of the cantilever free-oscillation. The number attached to each line indicates the phase delay of the
feedback operation. (b) Dependence of rmax on the phase delay of feedback control. The number attached to eachline indicates a value of h0/2A0 [71].

topography relative to the free-oscillation amplitude of the cantilever. As the error signals
fed into the PID controller are renewed every half cycle of the cantilever oscillation, sI
must be inversely proportional to the resonant frequency of the cantilever. The feedback
gain should be independent of parachuting, because the gain is maximized so that optimum feedback control is performed for a nonparachuting regime. Thus, sI is approximately expressed as sI = jh0sin(h/2)/(A0fc), where j is a proportional coeﬃcient.
5.6. Reﬁnement of analytical expressions for sp and sI
We experimentally measured the feedback bandwidth as a function of 2A0/h0 and rusing a mock AFM system containing a mock cantilever and z-scanner [71] (Fig. 5).
The mock cantilever and z-scanner are second-order low-pass ﬁlters whose resonant frequencies and quality factors are adjusted to have the corresponding values of a real cantilever and z-scanner. This mock AFM system is useful for conducting a rapid inspection

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Fig. 5. Circuit diagram of a mock AFM system. The disturbance signal fed into the input 2 simulates sample
topography. The output simulates the oscillation of a cantilever tip interacting with a sample surface. The
amplitude change caused by the interaction is given by the diode [71].

of the feedback performance. The experimentally obtained feedback bandwidths areshown by the black lines in Fig. 6. Feedback bandwidths are theoretically calculated using
Eq. (6), j and b as variables (see Eqs. (11) and (12)) and known values of the other parameters. From this analysis, we obtained reﬁned expressions for b and sI as follows:b ¼ cos1 ½A0 ð1  rÞ=5h0 sinðh=2Þ;ð13Þ

sI ¼ 4h0 sinðh=2Þ=ðA0 fc Þ:

ð14Þ

Feedback bandwidths calculated using these reﬁned expressions are shown by the graylines in Fig. 6. They approximately coincide with the experimental data.
5.7. Summary of guidelines for developing high-speed AFM
The following are a summary of the guidelines for realizing AFM with a high-speed
imaging capability.

Fig. 6. Feedback bandwidth as a function of the set point (r) and the ratio (2A0/h0) of the free-oscillation peakto-peak amplitude to the sample height. The number attached to each curve indicates the ratio 2A0/h0. The
feedback bandwidths were obtained under following conditions: the cantilever’s resonant frequency, 1.2 MHz; Q
factor of the cantilever oscillation, 3; the resonant frequency of the z-scanner, 150 kHz; Q factor of the z-scanner,
0.5. Black lines, experimentally obtained feedback bandwidths using a mock AFM; gray lines, theoreticallyderived feedback bandwidths.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

351(1) All time delay components involved in the feedback bandwidth must be similar.
When even one component has a signiﬁcant time delay compared with the others,
the feedback bandwidth is governed by the slowest component.
(2) As the cantilever resonant frequency is involved in two time delay components, it is
the most important device for achieving a high-speed scan capability.
(3) The quality factors of the cantilever and z-scanner have to be lowered.
(4) The resonant frequency of the z-scanner should be high (ideally, at a level similar tothat of the cantilever).
(5) The derivative operation given by the D component of the PID controller or of anadditional phase compensator can compensate for the feedback delay. To make thisoperation eﬀective, the gain of the z-scanner resonant peaks at high frequencies haveto be lowered. Otherwise, the derivative operation produces signiﬁcant mechanicalvibrations and therefore cannot be used.
(6) The free-oscillation peak-to-peak amplitude of a cantilever should be a few times larger than the maximum sample height. However, this condition has to be compromised to reduce the tapping force exerted from the oscillating tip on the sample.
(7) As the tip parachuting signiﬁcantly lowers the feedback bandwidth, we have todevelop methods that can shorten the parachuting time or avoid parachuting. We
can avoid parachuting by using a small set point amplitude. However, as thisincreases the tip–sample interaction force, we have to ﬁnd an alternative to a small
set point amplitude.
(8) All electronics used should have bandwidths as high as possible.
(9) We have to bear in mind that techniques for control operations have a minor role in
the improvement of the scan speed. The highest priority has to be assigned to theimprovement of the scanner and cantilevers over the consideration of sophisticatedcontrol techniques. Then, we should resort to control techniques to alleviate, to someextent, the limitation imposed by the well-optimized hardware devices.
6. Optimization of devices for high-speed AFM
6.1. Cantilevers
The feedback delays related to the cantilever are the amplitude detection time and thecantilever’s response time, both of which decrease in inverse proportion to the resonant
frequency. The resonant frequency fc and the spring constant kc of a rectangular cantileverwith thickness d, width w, and length L are expressed assﬃﬃﬃﬃﬃﬃﬃﬃ
d
E
;
ð15Þ
fc ¼ 0:56 2
12q
L
and
kc ¼

wd 3
E;
4L3

ð16Þ

where E and q are Young’s modulus and the density of the material used, respectively.
Young’s modulus and the density of silicon nitride (Si3N4), which is often used as a material for soft cantilevers, are E = 1.46  1011 N/m2 and q = 3.087 kg/m3, respectively. To
2.2
3.087 x 103

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attain a high resonant frequency and a small spring constant simultaneously, cantilevers
with small dimensions must be fabricated.
In addition to the advantage in achieving a high imaging rate, small cantilevers haveother advantages. For a given spring constant, the resonant frequency increases withdecreasing mass of the cantilever. The totalpthermal
ﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ noise depends only on the spring constant and the temperature and is given by k B T =k c [9], where kB is Boltzmann’s constantand T is the temperature in Kelvin. Therefore, a cantilever with a higher resonant frequency has a lower noise density. In the tapping-mode, the frequency region used for
imaging is approximately the imaging bandwidth (its maximum is the feedback frequency)
centered on the resonant frequency. Thus, a cantilever with a higher resonant frequency isless aﬀected by thermal noise. In addition, shorter cantilevers have higher OBD detection
sensitivity, because the sensitivity follows D/=Dz ¼ 3=2L, where Dz is the displacement andD/ is the change in the angle of a cantilever free-end. A high resonant frequency and asmall spring constant result in a large ratio (fc/kc), which gives the cantilever high sensitivity to the gradient (k) of the force exerted between the tip and the sample. The gradient of
the force shifts the cantilever resonant frequency by approximately 0.5 kfc/kc. Therefore,
small cantilevers with large values of fc/kc are useful for phase-contrast imaging and FM–
AFM. The practice of phase-contrast imaging using small cantilevers is described in Chapter 7. The usefulness and limitation of small cantilevers having a large fc/kc in FM–AFM is
described in Section 8.2.
The small cantilevers recently developed by Olympus are made of silicon nitride and arecoated with gold of 20 nm thickness (Fig. 7). They have a length of 6–7 lm, a width of
2 lm and a thickness of 90 nm, which results in the resonant frequencies of 3.5 MHz in
air and 1.2 MHz in water, a spring constant of 0.2 N/m, and Q  2.5 in water. We are
currently using this type of cantilever, although it is not yet commercially available. It is
possible to manufacture smaller cantilevers by microfabrication techniques to attain ahigher resonant frequency as well as a small spring constant. Considering the balance
between their practical use and desirable mechanical properties, we cannot expect a resonant frequency in water of much higher than 1.2 MHz. It is not practical to use a cantilever with w < 2 lm, considering the diﬀraction limit of the optics in the OBD detector.
To keep w and the spring constant unchanged, d/L should be unchanged (see Eq. (16)).

Fig. 7. Electron micrograph of a small cantilever developed by Olympus. Scale bar, 1 lm.

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353

To double the resonant frequency under this condition, both d and L should be halved (seeEq. (15)). With such a short cantilever (3–4 lm long), the incident laser beam used in the
OBD detector tends to be eclipsed by the cantilever supporting base. In addition, theallowable tilt range of the supporting base relative to the sample substratum surface
becomes narrowed. Thus, the practical upper limit of the attainable resonant frequency
in water is at the very most 2 MHz.
6.2. Cantilever tip
The tip apex radius of the small cantilevers developed by Olympus is 17 nm [72],which is not suﬃciently small for the high-resolution imaging of biological samples. We
usually attach a sharp tip on the original tip by electron-beam deposition (EBD) in phenolgas. A piece of phenol crystal (sublimate) is placed in a small container with small holes(0.1 mm diameter) in the lid. The container is placed in a scanning electron microscope(SEM) chamber and cantilevers are placed immediately above the holes. A spot-mode electron beam is irradiated onto the cantilever tip, which produces a needle on the original tip
at a growth rate of 50 nm/s. The newly formed tip has an apex radius of 25 nm(Fig. 8a) and is sharpened by plasma etching in argon or oxygen gas, which decreases
an apex radius to 4 nm (Fig. 8b). The mechanical durability of this sharp tip is not highbut is still suﬃcient to be used to capture many images.
This piece-by-piece attachment of the tip is time-consuming. Batch procedures for
attaching a sharp tip to each cantilever have been attempted by the direct growth of eithera single carbon nanoﬁber (CNF) [73,74] or a carbon nanotube (CNT) [75] at the cantilevertip. Tanemura found that Ar ion beam-irradiation onto a carbon-coated cantilever produces a single CNF only at the apex of the original tip [73]. In this method, the growthorientation is easily controlled by adjusting the direction of ion-beam irradiation relative
to the cantilever plane. However, at present, this method requires a carbon coating on the
cantilever and cannot produce CNFs with a radius less than 10 nm. Very recently, we
attempted to grow a single CNT on an original cantilever tip by chemical vapor deposition(CVD) using ethanol as a carbon source and Co as a catalyst. Although the success rate is
10% at present, we have grown a CNT at the tip with the desired orientation (Fig. 9).

Fig. 8. Electron micrographs of an EBD tip grown on an original cantilever tip. (a) Before and (b) after
sharpening by plasma etching in argon gas.

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Fig. 9. Electron micrograph of a carbon nanotube tip directly grown on an original cantilever tip by a CVD
method.

Carbon tips probably absorb red-laser light used for OBD sensing, as the laser light istightly focused onto a free-end region of the cantilever and passes through it to someextent. This light absorption certainly elevates the temperature at the tip. In addition,
heating also occurs by red-laser light absorption at the gold coat, although the absorption
rate is very small. It remains to be examined how high the temperature increases using
samples which exhibit transition phenomena at temperature moderately higher than ambient temperature.
6.3. Optical beam deﬂection detector for small cantilevers
Schäﬀer et al. designed an OBD detector for small cantilevers [51]; a laser beam
reﬂected back from the rear side of a cantilever is collected and collimated using thesame lenses as those used for focusing the incident laser beam onto the cantilever. We
use the same method but instead of single lenses, an objective lens with a long working
distance of 8 mm (CFI Plan FluorELWD20C, NA, 0.45, Nikon) is used [54]. The
focused spot is 3–4 lm in diameter. The incident and reﬂected laser beams are separatedusing a quarter-wavelength plate and a polarization splitter (Fig. 10). Our recent highspeed AFM is integrated with a laboratory-made inverted optical microscope withrobust mechanics. The focusing objective lens is also used to view the cantilever andthe focused laser spot with the optical microscope. The laser driver is equipped with aradio-frequency (RF) power modulator to reduce noise originating in the optics [76].
Its details are described in Section 8.4. The photosensor consists of a 4-segment Si
PIN photodiode (3 pF, 40 MHz) and a custom-made fast ampliﬁer/signal conditioner(20 MHz).

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355

Fig. 10. Schematic drawing of the objective lens type of OBD detection system. The collimated laser beam is
reﬂected up by the dichroic mirror and incident on the objective lens. The beam reﬂected at the cantilever iscollimated by the objective lens, separated from the incident beam by the polarization beam splitter and k/4 wave
plate, and reﬂected onto the split photodiode [54].

6.4. Tip–sample interaction detection methods
Tip–sample interactions change the amplitude, phase, and resonant frequency of theoscillating cantilever. They also produce higher-harmonic oscillations. In this section,
we describe methods for detecting the amplitude and the interaction force. Methods for
detecting shifts in the phase and resonant frequency are described in Chapter 7 and Section
8.1, respectively.
6.4.1. Amplitude detectors
Conventional RMS-to-DC converters use a rectiﬁer circuit and a low-pass ﬁlter, andconsequently require at least several oscillation cycles to output an accurate RMS value.
To detect the cantilever oscillation amplitude at the periodicity of half the oscillation cycle,
we developed a peak-hold method; the peak and bottom voltages are captured and then
their diﬀerence is output as the amplitude (Fig. 11) [54]. The sample/hold timing signals
are usually made from the input signals (i) (i.e., sensor output signals) themselves. Alternatively, external signals (ii) that are synchronized with the cantilever excitation signals
can be used to produce the timing signals. This is sometimes useful for maximizing thedetection sensitivity of the tip–sample interaction because the detected signal is aﬀectedby both the amplitude change and the phase shift. This is the fastest amplitude detector
and the phase delay has a minimum value of p, resulting in a bandwidth of fc/4. A drawback of this amplitude detector seems to be the detection of noise as the sample/hold circuits capture the sensor signal only at two timing positions. However, the electric noise
picked up in this peak-hold method is less than that produced by the thermal ﬂuctuationsof the cantilever oscillation amplitude.
A diﬀerent type of amplitude (plus phase) detector can be simply constructed using ananalog multiplier and a low-pass ﬁlter. The sensor signal sðtÞ  Am ðtÞ sinðx0 t þ uðtÞÞ is356

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

Fig. 11. Circuit for fast amplitude measurement. The output sinusoidal signal from the split-photodiode ampliﬁer
is fed to this circuit. The output of this circuit provides the amplitude of the sinusoidal input signal at half
periodicity of the oscillation signal [54].

multiplied by a reference signal [2 sinðx0 t þ /Þ] that is synchronized with the excitation
signal. This multiplication produces a signal given by Am ðtÞ½cosðuðtÞ  /Þ  cosð2x0 þuðtÞ þ /Þ. By adjusting the phase of the reference signal and placing a low-pass ﬁlter after
the multiplier output, we can obtain a DC signal of  Am ðtÞ cosðDuðtÞÞ, where DuðtÞ is aphase shift produced by the tip–sample interaction. In this method, the delay in the amplitude detection is determined mostly by the low-pass ﬁlter. In addition, electric noise iseﬀectively removed by the low-pass ﬁlter.
A diﬀerent method (Fourier method) for generating the amplitude signal at the periodicity of a single oscillation cycle has been proposed [77]. In this method, the Fourier sineand cosine coeﬃcients (A and B) are calculated for the fundamental frequency from thepﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
deﬂection signal to produce A2 þ B2 . The maximum bandwidth of this detector is fc/8,
half that of the peak-hold method. The electric noise level in the Fourier method was similar to that in the peak-hold method (Fig. 12a). However, regarding the accuracy of amplitude detection, the performance of the Fourier method is better because the cantilever’s
thermal deﬂection ﬂuctuations can be averaged in this method. Thus, the Fourier method
is less susceptible to the thermal eﬀect than the peak-hold method, and consequently, thedetected amplitude variation of cantilever oscillation under a constant excitation power
were less than that detected by the peak-hold method (Fig. 12b).
6.4.2. Force detectors
The nonlinear impulsive tip–sample interaction induces higher-harmonic vibrations of
the cantilever. In the amplitude detection described in the previous section, these vibrations are nearly neglected. The eﬀect of the impulse (peak force  interaction time) on
the cantilever motion is distributed over harmonic frequencies (integral multiples of thefundamental frequency). When the amplitude of one of the higher-harmonic vibrations
is used for image formation, it can result in high-contrast images containing maps of material properties extracted by the mechanical tip–sample interaction [78–81]. The images
depend on the detected harmonic frequency.
Another attempt to use higher-harmonics has been made with the aim of increasing thedetection sensitivity of the tip–sample interaction. Since the impulsive force is exerted transiently in a short time, its peak force is relatively large. This means that the peak force
must be a highly sensitive quantity. The force F(t) cannot be detected directly because

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357

Fig. 12. Noise level comparison of two amplitude detection methods (peak-hold method and Fourier method).
The upper traces (red lines) represent the input signal. (a) Comparison of electric noise. A clean sinusoidal signal
mixed with white noise was input to the detectors. The RMS voltage of the white noise was adjusted to have thesame magnitude as that of the OBD photo sensor output. (b) Comparison of variations in the detected cantilever
oscillation amplitude. (For interpretation of the references to colours in this ﬁgure legend, the reader is referred to
the web version of this paper.)the cantilever’s ﬂexural oscillation gain is lower at higher-harmonic frequencies. F(t) can
be calculated from the cantilever’s oscillation wave z(t) by substituting z(t) into the equation of cantilever motion and then subtracting the excitation signal (i.e., inverse determination problem) [81–83]. Here, we do not need to resort to the diﬀerential of z(t), which isa process that yields noisy signals. An operation in which the phase of each Fourier
decomposed harmonic signal is shifted by p/2 and then multiplied by an appropriate gainis identical to using the diﬀerential. To ensure that this method is eﬀective, the cantilever
oscillation signal with a wide bandwidth (at least up to 4  fc) must be detected and fast
analog or digital calculation systems are necessary for converting z(t) to F(t). In addition, afast peak-hold system is necessary to capture the peak force.
By neglecting the friction force, F(t) can be calculated roughly by
F ðtÞ ¼

m
X
n¼2

F n ðtÞ ¼

m
X
ð1  n2 ÞðAn cos nxc t þ Bn sin nxc tÞ;

ð17Þ

n¼2

where Fn(t) represents the force component with a harmonic frequency of n  fc, An and Bn
are the Fourier cosine and sine coeﬃcients of the nth harmonic component, respectively,

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xc is the fundamental resonant angular frequency, and m indicates the upper limit of theseries terms to be included. Because the excitation force and the fundamental frequency
component of the interaction force cannot be separated, the term with n = 1 must be neglected. As the quality factor of small cantilevers is 2–3, the friction force does not contribute signiﬁcantly to the total force compared with the other forces. Fig. 13b and c showsforce signals F(t) that were obtained by oﬀ-line calculation using an oscillation signal ofa small cantilever weakly interacting with a mica surface.
Since the time width of the impulsive force is narrow, it appears to be diﬃcult to capture the peak force using a sample/hold circuit. Instead of capturing the peak force, we cancalculate it using the time t0 when the cantilever oscillation reaches the bottom. For theﬁrst
harmonic
component of cantilever oscillation, Z 1 ðtÞ ¼ A1 cos x0 t þ B1 sin x0 t ¼
ﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
ﬃq

A21 þ B21 cosðxc t  uÞ, the time t0 is given by t0 = (p + 2kp + u)/xc, where k is an integer
and u isP
the phase delay given by u = tan1(B1/A1). Thus, the peak force is calculated asm
F ðt0 Þ ¼ n¼2 F n ðt0 Þ. Although the timing of the peak-force may deviate from the timing
when the cantilever reaches the end of the bottom swing, we can adjust t0 so that the maximum force signal can be attained. By including more terms in the series given by Eq. (17),we can obtain a larger force signal, as shown in Fig. 13c. However, this probably increases
noise, because noise contained in the Fourier coeﬃcients is ampliﬁed by a factor of(1  n2). In addition, the Fourier coeﬃcients in the higher-harmonic components become
smaller with increasing n. Therefore, the maximum term to be included in the series isdetermined by considering the total noise contained in the calculated peak force. For
the real-time calculation of the force, a fast DSP system or a fast FPGA system is required.
We are now attempting to build a peak-force detection circuit with a real-time calculation
capability.
Recently, a method of directly detecting the impulsive force was presented [84,85]. The
torsional vibrations of a cantilever have a higher fundamental resonant frequency (ft) thanFig. 13. Force signals calculated from the oscillation signal of a cantilever interacting with a mica surface inwater. (a) Cantilever oscillation signal; (b) and (c) force signals calculated taking the higher-harmonics up to theﬁfth component (b) or up to the 8th component (c).

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359

that of ﬂexural oscillations (fc). Therefore, the gain of torsional vibrations excited by
impulsive tip–sample interaction is maintained at 1 over frequencies higher than fc. Here,
we assume that ﬂexural oscillations are excited at a frequency of fc. To excite torsional
vibrations eﬀectively, ‘‘torsional harmonic cantilevers” with an oﬀ-axis tip have been introduced [84,85]. Oscilloscope traces of torsional vibration signals indicated a time-resolved
tip–sample force. We recently observed similar force signals using our small cantilevers
with an EBD tip at an oﬀ-axis position near the free beam end. After ﬁltering out the fc
component from the sensor output, periodic force signals appeared clearly (Fig. 14). To
use the sensitive force signals for high-speed imaging, we again need a means of capturing
the peak force or a real-time calculation system to obtain the peak force. As we do notneed time-resolved force signals for imaging purposes, some simpliﬁcation for the calculation can be used to increase the calculation speed.
6.5. High-speed scanners
The high-speed driving of mechanical devices with macroscopic dimensions tends to
produce unwanted vibrations. Therefore, among the devices used in high-speed AFM,
the scanner is the most diﬃcult to optimize for high-speed scanning. Several conditions
are required to realize high-speed scanners: (a) high resonant frequencies, (b) a small number of resonant peaks in a narrow frequency region, (c) suﬃcient maximum displacements,(d) small crosstalk between the three axes, (e) low quality factors. The following sections(Sections 6.5.1–6.5.3) describe several techniques for satisfying these conditions simultaneously. Active damping techniques are described in Section 6.6. The performance ofour developed scanners is described in Section 6.6.4.
6.5.1. Counterbalance
The quick displacements of a piezoelectric actuator exert impulsive forces onto the supporting base, which cause vibrations of the base and the surrounding framework and in

Fig. 14. Force signal directly obtained from the torsional signal of a small cantilever with an oﬀ-axis tip. The
cantilever was excited at its ﬁrst ﬂexural resonant frequency (1 MHz) in water. The oﬀ-axis tip wasintermittently contacted with a mica surface in water. Upper trace, torsional vibrations of the cantilever; lower
trace, force signal obtained by ﬁltering the torsional signal using a low-pass ﬁlter to remove the carrier wave
(1 MHz). The torsional signal appears even under free-oscillation due to cross talk between ﬂexural and torsional
vibrations.

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turn, of the actuator itself. To alleviate the vibrations, a counterbalance method was introduced for the z-scanner [54]; impulsive forces are countered by the simultaneous displacements of two z-piezoelectric actuators of the same length in the counter direction(Fig. 15a). In this arrangement, the counterbalance works eﬀectively below the ﬁrst resonant frequency of the actuators but does not work satisfactory around the resonant frequencies. The vibration phase changes sharply around the resonant frequencies, andtherefore, a slight diﬀerence in the mechanical properties of the two actuators disturbs
the counterbalance.
It is possible to make diﬀerent z-scanner designs that can counterbalance the impulsive
forces more eﬃciently. An alternative design employs a piezoelectric actuator sandwiched
between two ﬂexures in the displacement directions (Fig. 15b). As a single actuator is used,
its center of mass as well as the entire mechanism barely moves when the mechanical properties of the two ﬂexures are similar to each other. This is true even at the resonant frequencies. This method can be used for both the x- and z-scanners. To ensure that thismethod is eﬀective when used for the z-scanner, the ﬂexure resonant frequency must be
increased, which requires a large spring constant of the ﬂexures, which reduces the maximum displacement of the scanner. A second alternative is that a piezoelectric actuatoris held only at the rims or corners of a plane perpendicular to the displacement direction(Fig. 15c). This allows the actuator to be displaced almost freely in both directions, andconsequently, the center of mass barely moves. The last alternative is that a piezoelectric
actuator is glued to (or pushed into) a circular hole of a solid base so that the side rims
parallel to the displacement direction are held (Fig. 15d). Even held in this way, the actuator can be displaced almost up to the maximum length attained under the load-free condition. We recently employed this method for the z-scanner (see Section 6.6.4).
A sample stage has to be attached to the z-piezoelectric actuator. To balance the weight,
a dummy sample stage is attached to the opposite side of the piezoelectric actuator (in the
conﬁguration shown in Fig. 15a, it is attached to the second z-actuator). The weight of thedummy sample stage is chosen by considering the hydrodynamic pressure produced
against the quick displacement of the sample stage. The sample-stage mass (ms) decreases
theﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
lowest resonant
frequency of the z-piezoelectric actuator (mass, mz) by a factor ofﬃp
1 þ 2ms =3mz when the conﬁgurationsp
depictedin Fig. 15b–d are used. For the conﬁguﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
ration shown in Fig. 15a, the factor is 1 þ ms =3mz .

Fig. 15. Various conﬁgurations of holding piezoelectric actuator for suppressing unwanted vibrations. The
piezoelectric actuators are shown in green, and the holders are shown in blue. (a) Two actuators are attached tothe base, (b) the two ends of an actuator are held with ﬂexures in the displacement direction, (c) an actuator isheld only at the rims or corners of a plane perpendicular to the displacement direction, (d) an actuator is glued tosolid bases at the rims parallel to the displacement direction. (i) Top view, (ii) side view. (For interpretation of thereferences to colour in this ﬁgure legend, the reader is referred to the web version of this article.)

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6.5.2. Mechanical scanner design
Tube scanners that have been often used for conventional AFM are inadequate forhigh-speed scanning, as their long and thin structure lowers the resonant frequencies inthe x-, y-, and z-directions. The structural resonant frequency can be enhanced by adopting a compact structure and a material that has a large Young’s modulus to density ratio.
However, a compact structure tends to produce interference (crosstalk) between the threescan axes. A ball-guide stage [54] is one choice for avoiding such interference. An alternative method is to use ﬂexures (blade springs) that are suﬃciently ﬂexible to be displaced
but suﬃciently rigid in the directions perpendicular to the displacement axis [86,87].
The ﬂexure-based stage is more stable and more easily built than the ball-guide stage. Notethat the scanner mechanism, except for the piezoelectric actuators, must be produced bymonolithic processing to minimize the number of resonant elements.
When we need a scan speed as high as possible, it is best to have asymmetric structuresin the x- and y-directions; the slowest y-scanner displaces the x-scanner, and the x-scannerdisplaces the z-scanner, as in our currently used scanner (Fig. 16). Using this scanner, themaximum displacements (at 100 V) of the x- and y-scanners are 1 lm and 3 lm, respectively. Two z-piezoelectric actuators (maximum displacement, 2 lm at 100 V; self-resonant
frequency, 360 kHz) are used in the conﬁguration shown in Fig. 15a. The gaps in the scanner are ﬁlled with an elastomer to passively damp the vibrations. This passive damping is
eﬀective in suppressing low-frequency vibrations.
As developed at Hansma’s lab, a symmetrical x–y conﬁguration has an advantage ofbeing capable of rotating the scan direction [86]. Aluminum or duralumin is often used
as a material for the scanner. Magnesium and magnesium alloys appear to be more suitable materials because of their larger mechanical damping coeﬃcients and larger ratios ofYoung’s modulus to density. However, from our experiences, only a slight improvement is
attained using these materials.

Fig. 16. Sketch of the high-speed scanner currently used for imaging studies. A sample stage is attached on the
top of the upper z-piezoelectric actuator (the lower z-piezoelectric actuator used for counterbalancing is hidden).
The dimensions (W  L  H) of the z-actuators are 3  3  2 mm3. The gaps are ﬁlled with an elastomer for
passive damping.

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An alternative design for the xy-scanner has been used in a high-speed scanning tunneling microscope (STM) [88]. Two shear-mode piezoelectric actuators are simply stacked to
produce a xy-scanner, on top of which a z-piezoelectric actuator (stack actuator) is
attached. This type of scanner is commercially available from PI (Physik Instrumente)GmbH (Germany). This conﬁguration results in a very compact structure. However, inshear-mode piezoelectric actuators, the polarization axis is perpendicular to the external
electric ﬁeld, and hence, a large electric ﬁeld cannot be applied. The displacement rate issmaller than one-tenth that of stack-mode piezoelectric actuators. To attain suﬃcient displacements, a high voltage has to be applied to relatively thick piezoceramic plates. As theattachment of a z-piezoelectric actuator onto the stacked xy-scanner results in the formation of a long resonator with a low resonant frequency, we need a specially designed
attachment to avoid this problem.
6.5.3. Caution regarding hydrodynamic eﬀects
When the z-scanner is displaced quickly, hydrodynamic pressure is generated from thereaction of the sample solution against the cantilever base (Fig. 17). We examined thiseﬀect by quickly moving the sample stage toward the cantilever, while keeping the tip incontact with the sample stage [55]. As shown in Fig. 17a, when the sample stage dimensions were 5  5 mm2, the cantilever vibrated at 20 kHz due to the hydrodynamic
force. When the size of the sample stage was reduced to 1 mm in diameter, almost no
vibrations appeared (Fig. 17b). Therefore, we have been using glass sample stages witha circular-trapezoid shape and a small top surface of 1 mm diameter. When the cantilever was oscillated at its resonant frequency and the tip was intermittently in contact withthe sample stage, no signiﬁcant vibrations appeared, even when using a larger sample
stage. However, we later recognized that even when no vibrations appeared, the hydrodynamic pressure decreased the feedback bandwidth in tapping-mode operation. After
examining this eﬀect by changing the cantilever position relative to the sample stage,
we found that this decrease in feedback bandwidth was due to the slight and slow displacement of the cantilever base. We were able to solve this problem by tightly ﬁxing thecantilever base using a stiﬀ blade spring or by ﬁxing the cantilever base at a position
near the cantilever beam.

Fig. 17. Disturbance on cantilever by hydrodynamic force and its dependence on the sample stage shape. The
sample stage is quickly moved toward the cantilever whose tip is in contact with a mica surface in water. The
lower signals represent the voltage applied to the z-piezoelectric actuator to move the sample stage. The uppersignals represent the cantilever deﬂection. (a) The sample stage is a slide glass cut to 5  5 mm2. (b) The sample
stage is a glass of circular-trapezoid shape with a small top surface of 1 mm diameter [55].

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363

As a solution to this problem, it seems best to alter the design of the cantilever base sothat it catches much less hydrodynamic pressure. In fact, after a cantilever base was partially cracked at the edge close to the cantilever beam, the base did not move by the exertedhydrodynamic pressure even when the base was not tightly ﬁxed.
6.6. Active damping
It is impossible to manufacture a scanner that exhibits no resonance around the harmonic frequencies contained in its driving signals. Therefore, the elimination of unwanted
vibrations from the scanner is a key to realizing high-speed AFM. Furthermore, the bandwidth of any mechanical system is limited by its dimensions and materials, and hence,some manipulation techniques are also required to overcome this limitation and expand
the scanner bandwidth. In this section, we describe several techniques related to theseissues. In high-speed AFM, z-scanner driving may be too fast to control even using
advanced digital mode controllers. Therefore, it is assumed below that active dampingof the z-scanner is performed using analog mode controllers. Vibration control is basicallyclassiﬁed into three types: notch ﬁltering, feedback control, and feed-forward control(Fig. 18). In the following sections, these three types of vibration control are explained
in detail.
6.6.1. Notch ﬁltering
Notch ﬁltering (Fig. 18a) is convenient for eliminating resonance with a single peak.
The transfer function N(s) (s = ix) of a standard two-zero, two-pole notch ﬁlter is given bys2 þ x0 s=q þ x20:
N ðsÞ ¼ 2s þ x0 s=Q þ x20

ð18Þ

At x = x0, N(s) becomes Q/q (hence, no phase change). This value gives the minimumtransmission rate. To eliminate the resonance of a scanner characterized by a simple transfer function GðsÞ ¼ x21 =ðs2 þ x1 s=Q1 þ x21 Þ, x0 and q must be tuned to x1 and Q1, respectively. This tuning results in the composite transfer function CðsÞ ¼ x21 =ðs2 þ x1 s=Q þ x21 Þ.
Therefore, the scanner vibrations are damped completely using Q = 0.5–0.7. However,some eﬀort is required to tune q as well as Q using analog notch ﬁlters. When the scanneris characterized by a more complicated transfer function with multiple peaks, it is important to ﬁnd whether the multiple peaks arise from multiple resonators connected in series
or in parallel. From the observed phase spectrum of the scanner, we can judge the type of
connection. When the resonators are connected in series, the phase of the transfer function

Fig. 18. Three types of active damping methods for suppressing the unwanted vibrations of the scanner. (a)Notch ﬁltering method, (b) feedback control method, (c) feed-forward control method. G(s) represents thetransfer function of a scanner to be controlled. H(s) represents the transfer function of a controller for active
damping.

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decreases monotonically and signiﬁcantly with increasing frequency, while for a parallel
connection, the phase shows an up-and-down feature. For a case of serial connection,
we can eliminate the multiple peaks by using notch ﬁlters connected in series. For the caseof a parallel connection, it is impossible to eliminate the peaks.
6.6.2. Feedback control for active damping
The feedback operator H(s) (see Fig. 18b) converts the resonant system G(s) to a target
system R(s) expressed asRðsÞ ¼GðsÞ
:
1  GðsÞH ðsÞ

ð19Þ

In the ﬁrst step, let us consider the simplest case where G(s) consists of a single resonant
element with resonant frequency x1 and quality factor Q1. The target system is expressed
as a single resonator with resonant frequency x0 and quality factor Q0. For these systems,
H(s) is expressed as
 2
  2

1
1
s
s
s
s

¼
þ
þ1 
þ
þ1 :
ð20Þ
H ðsÞ ¼GðsÞ RðsÞ
x21 Q1 x1x20 Q0 x0
To eliminate the second-order term from H(s), x0 should equal x1, which results in


1
1
s

:
ð21Þ
H ðsÞ ¼ 
Q0 Q1 x1
This H(s) is identical to a derivative operator with gain = 1 at the frequency^ ¼ Q0 Q1 x1 =ðQ1  Q0 Þ (Fig. 19a). By adjusting the gain parameter of the derivative operx
ator, we can arbitrarily change the target quality factor Q0. This method is simpler thannotch ﬁltering, as the tuning parameter is simply the gain. This method is known as‘‘Q-control” and is often used for controlling the quality factor of cantilevers [89–91].
For the Q-control of cantilevers with constant frequency excitation, we can change thephase of feedback signal by 90° using a phase shifter, instead of a derivative operator.
However, this method does not guarantee Q-control of a cantilever whose tip is transiently
contacting with the sample, as higher-harmonic components are produced by the contact.
If we do not eliminate the second-order term from H(s), then H(s) is equivalent to asystem consisting of a derivative operator and a second-derivative operator connectedin parallel (Fig. 19b). This second-derivative operator corresponds to ‘‘mass-control”.
Using this second type of Q-controller, we can extend the scanner bandwidth when a second-derivative circuit and a piezoelectric actuator driver, both having high bandwidths,
can be used. Here, note that when either Q-controller is applied to the z-scanner, we mustFig. 19. Diagram of feedback control for active vibration damping. (a) Simple Q-control system, (b) combined
control system able to extend the scanner bandwidth.

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365

Fig. 20. Feedback Q-control system with a mock scanner. M(s) represents the transfer function of a mockscanner. M(s) is similar to the transfer function G(s) of the real scanner. H(s) represents the transfer function of a
feedback Q-controller.

measure the displacement or velocity of the z-scanner. However, it is diﬃcult to do so.
This problem is solved by using a mock scanner M(s) (a second-order low-pass ﬁlter) characterized by the same transfer function as the z-scanner (Fig. 20) [92]. The practical use ofthis technique is described in Section 6.6.4.
Real scanners often exhibit multiple resonant peaks, and therefore, neither of themethods described above work well. When elemental resonators are connected in series,
we have to use mock scanners, M1(s), M2(s), . . ., each of which is characterized by atransfer function representing each elemental resonator. Each mock scanner is controlled
by a corresponding Q-controller. For example, when the scanner consists of two resonators connected in series, the composite transfer function C(s) of the total system isexpressed asCðsÞ ¼

1
1
 G1 ðsÞG2 ðsÞ:1  M 1 ðsÞH 1 ðsÞ 1  M 2 ðsÞH 2 ðsÞ

ð22Þ

Since M1(s) and M2(s) are the same as (or similar to) G1(s) and G2(s), respectively, Eq. (22)represents a target system consisting of damped resonators connected in series (see Eq.
(19)). Note that we can use M1(s) and M2(s) separately, unlike G1(s) and G2(s).
When the scanner consists of elemental resonators connected in parallel, active damping becomes more diﬃcult. Here, we only consider the case in which two elemental resonators are connected in parallel. The controller H(s) is expressed as
 2

1
1
1
s
s

¼ ax2

þ
þ
1
;
ð23Þ
H ðsÞ ¼bx22
x20 Q0 x0
G1 ðsÞ þ G2 ðsÞ RðsÞ
1
þ
x1
x2
2
2
2
2
s þ sþxs þ sþx
Q1

1

Q2

2

where a and b are the weight factors of the two resonators (hence, a + b = 1). To eliminate
the highest-order term from H(s), x20 should equal ax21 þ bx22 , which approximately resultsin (see Fig. 21)~
H ðsÞ  ½ðband-pass filterÞ  g  1  s=x:

ð24ÞFig. 21. Feedback Q-control system for the scanner characterized by a transfer function composed of two
resonators connected in parallel. M(s) is similar to the transfer function G(s) of the real scanner. BP(s) represents
a band-pass ﬁlter.

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T. Ando et al. / Progress in Surface Science 83 (2008) 337–437~ and the gain parameter g are, respectively, expressed as
The band-pass ﬁlter BP(s), x,BPðsÞ ¼ x2

ðQ ax2 þ Q bx1 Þs2

1;

s2 þ ðQ ax2 þ Q bx1 Þs þ 1
x21 x22
2
1


1
a
b
~ ¼1
;


x
Q0 x0 Q1 x1 Q2 x2


a
b
~
g ¼ cx
þ 1;
þQ2 x2 Q1 x1

ð25Þ

0

ð26Þ
ð27Þ

where c is given by



1
1
x20
1
x0
x1 x2
1 ax1 bx2
c¼ 2þ 2 2 2þ
:


þ
x0
Q1
Q0 Q2
x1 x2 x1 x2 x0 x1 x2 Q1 Q2

ð28Þ

The eﬀect of this controller H(s) is depicted in Fig. 22.
When the scanner’s transfer function is given by a combination of serial and parallel
connections of elemental resonators, it becomes diﬃcult to design a controller. It is best
to manufacture a scanner with a simple structure so that the number of resonators is
minimized.
6.6.3. Feed-forward control for active damping
The feed-forward control type of active damping (Fig. 18c) is based on inverse compensation (i.e., H(s)  1/G(s)). Generally, inverse compensation-based damping has an advantage in that we can extend the scanner bandwidth. This damping method is much easier toapply for the x-scanner than for the z-scanner, because for the former, the scan waves are
known beforehand and are periodic (hence, the frequencies used are discrete; integral
multiples of the fundamental frequency) [93–95]. Here, we ﬁrst consider the damping ofFig. 22. Eﬀect of feedback control for active damping on the resonant system that consists of two elemental
resonators connected in parallel. This eﬀect was theoretically obtained using the feedback Q-controller shown in
Fig. 21. (a) Bode plots of the resonant system before damping. (b) Bode plots of the resonant system after
damping.

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367

the x-scanner vibrations. The waveforms of the x-scan are isosceles triangles characterized
by amplitude X0 and fundamental angular frequency x0. Their Fourier transform is given
by
"
#
þ1
1
2 X
1
dðx  kx0 Þ
ðk : oddÞ:ð29Þ
F ðxÞ ¼ 2pX 0 dðxÞ  2
2
p k¼1 k 2
To move the x-scanner in the isosceles triangle waveforms, the input signal X(t) sent to thex-scanner characterized by the transfer function G(s) is given by the inverse Fourier transform of F(x)/G(ix), which is expressed as
þ1
X 0 4X 0 X
11
 2cosðkx0 tÞ ðk : oddÞ:ð30Þ
X ðtÞ ¼2
p k¼1 k 2 Gðikx0 ÞIn practice, the sum of the ﬁrst 10 terms in the series of Eq. (30) is suﬃcient. We cancalculate Eq. (30) in advance to obtain numerical values of X(t) and output them in succession from a computer through a D/A converter. This method is convenient and works
well. However, applying this method to the y-scanner is not practical because the period of‘‘y-return” (scanning towards the origin after acquiring a frame) is much shorter than the
total period of the y-scan (i.e., image acquisition time), and hence, we have to use too
many terms in the series expansion of the y-scan waveform. We can simply use a roundwaveform for the y-return. Rounding the vertexes of the triangle waveforms is also usefulfor the x-scanner, although the deviation of the scan trajectory from the triangle waveform
becomes large.
The transfer function G(s) usually changes after the repeated use of the x-scanner, andtherefore, more robust inverse-based control is required to avoid periodic examination ofG(s). To this end, feedback control can be applied, in addition to the feed-forward control
mentioned above, in order to compensate for the change in G(s) (and hence for the insufﬁcient feed-forward control) [96,97]. The feedback control reduces the deviation of theoutput trajectory from the desired trajectory, and consequently, can also reduce the hysteresis and creep eﬀects of the x-piezoelectric actuator.
Applying the inversion-based feed-forward active damping to the z-scanner is not easy
because its scan waves are not predictable, and it is diﬃcult to design an electronic circuitthat is characterized by the inverse transfer function of the z-scanner. It may be possible tofabricate such a circuit when the scanner exhibits a single resonant peak. There seem to be
two possible designs for the circuit. One is the parallel connection of a second-derivative
operator, a derivative operator, and a proportional gain (i.e., s2 =x20 þ s=ðQx0 Þ þ 1). The
other can be fabricated by using a serially connected LRC circuit driven by a current regulation ampliﬁer (Fig. 23) [98]. However, it is not easy to make this circuit.

Fig. 23. Current-regulated circuit for feed-forward active damping control (inverse compensation) in the single
resonator system.

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Fig. 24. Circuit diagram for APITF. G(s) represents a given transfer function for a real resonant system. M(s)represents a transfer function of a mock resonator for G(s). (a) Single loop circuit type. K(s) approximately gives
1/M(s). (b) Nested circuit for improving the approximation [111].

It is desirable to develop a technique that enables the automatic production of an
inverse transfer function (APITF) for a given transfer function. The transfer function
K(s) of the circuit shown in Fig. 24a is given by KðsÞ ¼ 1=½1 þ gðMðsÞ  1Þ; here, M(s)is a mock scanner. In the case of g = 1, a complete inverse transfer function, 1/M(s),can be realized. However, delays occur in the electronic components such as the operational ampliﬁers used in the circuit. Therefore, the gain factor g must be less than 1,
and consequently, the complete inverse transfer function cannot be realized. In fact,
K(s)M(s) has a large resonant peak at a frequency higher than the peak frequencies ofM(s). However, this peak can be eliminated using a notch ﬁlter. When the disagreement
between K(s) and 1/M(s) is signiﬁcant, we can reduce the diﬀerence by placing a phasecompensator (1 + derivative operation) immediately before or after the mock scannerand by using a multiple-loop circuit, as indicated in Fig. 24b. The eﬀectiveness of this technique is shown in Fig. 25, which was obtained by numerical calculations of K(s) for thecase that G(s) consists of two elemental resonators connected in parallel. The resonantpeaks are eliminated and the bandwidth is signiﬁcantly extended. Unfortunately, thismethod has a limitation; it does not work when G(s) has more than three poles. We have

Fig. 25. Eﬀect of feed-forward control for active damping on the resonant system that consists of two elemental
resonators connected in parallel. This eﬀect was theoretically calculated using the 7-fold-nested APITF system. (a)Bode plots of the resonant system before damping. (b) Bode plots of the resonant system after damping.

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been using this APITF technique for compensating the slow response of the photothermal
bending of a cantilever (see Section 6.10) and z-scanners (Section 6.6.4).
6.6.4. Practice of active damping of the scanner vibrations
In this section, we describe the practice of active damping applied to the scanners that
we are currently using (one type is shown in Fig. 16) and to other scanners underdevelopment.
Here, we ﬁrst show the eﬀect of active damping applied to the x-scanner. The active
damping was performed using an inverse compensation method. The x-scanner transfer
function (amplitude and phase spectra) is shown in Fig. 26. When the line scan wasFig. 26. Frequency spectra (bode plots) of mechanical response of the x-scanner.

Fig. 27. Eﬀect of feed-forward active-damping control on the x-scanner vibrations. (a) x-Scanner displacement
driven by triangle waves without damping. (b) x-Scanner displacement driven by triangle waves compensated by
the feed-forward control. (c,d) Mica surface images acquired at 32 frames/s (c) without and (d) with damping.

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performed at 3.3 kHz without damping, its displacement exhibited vibrations (Fig. 27a).
When it was driven in a waveform calculated using Eq. (30) with the maximum term
k = 17 in the series, the x-scanner moved approximately in a triangle waveform(Fig. 27b). This damping eﬀect was also veriﬁed by imaging mica surfaces at 33 frames/s over a scan range of 240  240 nm2 with 100 scan lines (this imaging speed corresponds
to an x-scan frequency of 3.3 kHz). When no active damping was applied, striation patterns parallel to the y-axis appeared (Fig. 27c). The periodicity of the patterns indicatesthat the vibrations had a frequency of 60 kHz, which coincided with the resonant peakfrequency shown in Fig. 26. When active damping was applied, the striation patterns completely disappeared (Fig. 27d). Here, note that the resonant vibrations in the x-direction
also produce vibrations in the z-direction.
In the z-scanner that we are currently using for imaging studies, the z-piezoelectric actuators have a resonant frequency of 410 kHz under free-oscillation. The counterbalancemethod employed is depicted in Fig. 15a. Using this method of counterbalancing, nostructural resonances appear at frequencies lower than the ﬁrst resonant frequency(171 kHz) of the two identical z-piezoelectric actuators (gray lines in Fig. 28). The resonant frequency of 171 kHz is derived from the fact that one end of each piezoelectric actuator is attached to the base (410/2 = 205 kHz) and the other end is attached to a sample
stage (or a dummy stage). Higher resonant frequencies also appear at around 350 kHz.

Fig. 28. Frequency spectra of mechanical response of the z-scanner. (a) Gain spectra, (b) phase spectra. Graylines represent the response without feedback Q-control, and black lines represents the response with feedback Qcontrol.

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Judging from the phase spectrum (gray line in Fig. 28b), the two main resonant elements
are connected in parallel. For the active damping of these resonant vibrations, we employ
the simple Q-control method (Fig. 19a) described in Section 6.6.2. In addition, to compensate for a phase delay caused by active damping, a (1 + derivative) circuit is inserted
between the Q-controller and the piezodriver. Although this method does not work perfectly, as mentioned in the Section 6.6.2, we use a mock z-scanner with two resonator circuits connected in parallel. This gives a better result compared with the use of a mockscanner with a single resonator. Using this method of Q-control, the ﬁrst resonance completely disappears (black line in Fig. 28a) and the phase delay is not signiﬁcantly deteriorated (black line in Fig. 28b). As the quality factor is reduced signiﬁcantly, the z-scanner
response time is markedly improved from 17.1 ls to 0.93 ls (Fig. 29). This improvement in
the response speed of the z-scanner is more clearly conﬁrmed by the cantilever deﬂection
responded to the stepwise variation of the set point level of the PID controller (Fig. 30).
The set point variation corresponds to a 100 nm height change. Without active damping,
the response is very slow (Fig. 30a). Using the active damping and phase compensation by(1 + derivative) operation, the input rectangular signals are well restored in the cantileverdeﬂection signal (Fig. 30b). The slight ringing behavior observed at the rising and fallingedges are due to the remaining resonances at around 350 kHz. These resonances could be
removed using a notch ﬁlter or a low-pass ﬁlter. We do not remove them, since such ﬁltering increases the phase delay. In practice, these remaining resonances negligibly disturb

Fig. 29. Eﬀect of active damping on the transient response of the z-scanner. In (a), the z-scanner has its natural
Q = 9, while in (b) the z-scanner is actively damped so that its Q is reduced to 0.5. The z-scanner began to be
oscillated at t = 100 ls by ac signals (lines (i)) of 150 kHz. The amplitude of the ac input signals was attenuatedwhen operated without active damping. The enveloping curves (gray lines) of the z-scanner’s displacements (lines(ii)) indicate the transient changes in the amplitude of the z-scanner’s oscillation.

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Fig. 30. Cantilever deﬂection responses to rectangular variation of the set point of the PID controller. The trace‘‘2” in (a) is obtained without active damping of the z-scanner vibrations, while the trace ‘‘2” in (b) is obtainedwith active damping. The traces ‘‘1” represent the rectangular set point variation.

the imaging of protein molecules, even at an imaging rate of 30 ms/frame over a scan
range of 240 nm with 100 scan lines. This is understandable from the relationshipf = Vs/k (see Section 5.1). Under these imaging conditions, the maximum feedback frequency is estimated to be about 200 kHz when the apparent feature size of the sample protein molecules is about 8 nm. Therefore, the z-scanner is not excited even at an imaging
rate of 30 ms/frame.
We recently attempted to develop other types of scanner in which a z-piezoelectric actuator is held either at the four side rims parallel to the displacement direction [99] (method
depicted in Fig. 15d) or at the rims and corners of a plane perpendicular to the displacement direction (method depicted in Fig. 15c). The gain and phase spectra of the mechanical response of the former z-scanner are shown with the blue lines in Fig. 31a and b. The
z-scanner exhibited large resonant peaks at 440 kHz and 550 kHz. The resonant frequencyof 440 kHz is very similar to that of the free-oscillation of the piezoelectric actuator. Judging from the phase spectrum, the two resonators are connected in series. We used two feedback active damping circuits connected in series (see Eq. (22)). The resulting gain andphase spectra are shown with the red lines in Fig. 31a and b, respectively. The peak at
440 kHz is almost completely removed and the frequency that gives a 90° phase delay
reaches 250 kHz.
The gain and phase spectra of the mechanical response of the latter z-scanner (a z-piezoelectric actuator was held as depicted in Fig. 15c) are shown with blue lines in Fig. 31c
and d. The z-scanner exhibited large resonant peaks at 370 kHz and 540 kHz. The reso-

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Fig. 31. Frequency spectra of mechanical response of two types of z-scanners. (a) and (b) are, respectively, thegain and phase spectra of a z-scanner whose piezoelectric actuator is held at the four rims parallel to thedisplacement direction (see Fig. 15d). (c) and (d) are, respectively, the gain and phase spectra of a z-scanner whose
piezoelectric actuator is held at the four corners of a plane perpendicular to the displacement direction (seeFig. 15c). The blue lines represent the responses without feed-forward active damping, and the red lines represent
the responses either with feedback Q-control ((a), (b)) or feed-forward active damping ((c), (d)). (Forinterpretation of the references to colour in this ﬁgure legend, the reader is referred to the web version of thisarticle.)nant frequency of 370 kHz is similar to that of the free-oscillation of the piezoelectric actuator (410 kHz). Judging from the phase spectrum, the two resonators are connected in parallel. Thus, we applied feed-forward active damping using a double-loop APITF circuit
containing a mock scanner composed of two LRC circuits connected in parallel. The
resulting gain and phase spectra are shown with the red lines in Fig. 31c and d, respectively. The peak at 370 kHz is completely removed and the frequency that gives a 90°
phase delay is increased from 370 kHz to 500 kHz. A problem that we have not been able
to resolve is the occurrence of small resonances around 50–80 kHz. This is likely to arise
from that holding the z-piezoelectric actuator is not tight enough.
Among various methods of holding a z-piezoelectric actuator (Fig. 15), the method
depicted in Fig. 15d [99] thus far resulted in the best performance regarding the high-frequency of the ﬁrst resonance, mechanical stability and small crosstalk between the x- andz-axes. The maximum displacement (1 lm) is suﬃcient for imaging biological macromolecules. Since the ﬁrst resonant frequency is suﬃciently high, it will not be excited even atvideo-rate scanning. Therefore, we do not have to use complicated active damping methods to completely eliminate the resonant bands. However, it seems the best to remove

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resonant peaks at high frequencies, so that the high gain margin of feedback control forthe constant tip–sample distance is attained.
6.7. Dynamic PID control
To observe biological processes containing delicate protein–protein interactions, thetip–sample interaction force has to be minimized. The tapping force exerted on the sample
by the oscillating cantilever tip can be reduced by using an amplitude set point r close to 1.
As described in Section 5.4, this set point condition induces parachuting, during which theerror signal saturates. The saturated error signal becomes smaller as r increases, and hence,the parachuting time sp is prolonged. The feedback gain cannot be increased to shorten sp,
since a larger gain induces overshoot at uphill regions of the sample, resulting in the instability of the feedback operation. In this section, we describe a new PID controller(‘‘dynamic PID controller”) developed to solve this problem [71].
6.7.1. Dynamic PID controller
With conventional PID controllers, the gain parameters are constant, irrespective ofwhether scanning is underway at uphill regions or downhill regions of the sample. To
avoid parachuting or to shorten the parachuting period, the gain parameters should be larger at downhill regions than at uphill regions. The amplitude signal is an indicator of thetype of region that is being scanned. Generally, at uphill regions the amplitude is smaller
than the set point amplitude, while it is larger at downhill regions. Thus, with the dynamic
PID controller that we developed, the gain parameters are automatically altered depending on the cantilever oscillation amplitude. A threshold level Aupper is set between the set
point peak-to-peak amplitude As and the free-oscillation peak-to-peak amplitude 2A0 (oris set to As). We usually use a value of Aupper that is much closer to As than 2A0, the reason
for which is mentioned later. When the cantilever peak-to-peak oscillation amplitude Ap–pexceeds Aupper, the gain parameters are increased. There are several methods of imple-

Fig. 32. Diagram of feedback loop with dynamic PID operator [71].

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menting this automatic alteration in the gain parameters. We insert a circuit called a‘‘dynamic operator” between the error signal output and the input of a conventional
PID circuit (Fig. 32). This dynamic operator functions as follows (see Fig. 33). When
(Ap–p  Aupper) > 0, the diﬀerential signal (Ap–p  Aupper) is ampliﬁed and added to theerror signal. The error signal that contains this extra signal is fed to the conventional
PID. This ‘‘false error signal”, whose value is larger than that of the ‘‘true error signal”,
produces a quicker feedback response, and therefore, Ap–p quickly becomes smaller thanAupper and the feedback operation automatically returns to the normal mode. Thus, evenwhen As is very close to 2A0, the parachuting period is shortened markedly, and hence,practically no feedback saturation occurs. When Aupper is far from As, the dynamic operator produces an abrupt jump in the error signal. This is the reason why we use a value ofAupper that is much closer to As than 2A0. Noted that to ensure that this dynamic PID controller is eﬀective even when the set point r is very close to 1, the amplitude signal should
not contain noise greater than 2A0 (1  r)/3.
Similar manipulation of the error signal can also be performed when Ap–p is smaller
than As. In this case, a new threshold level Alower is set with a value suﬃciently lower thanAs. When Ap–p becomes lower than Alower, the diﬀerential signal (Ap–p  Alower) is ampliﬁed and then added to the error signal (Ap–p  As). This can prevent the cantilever tip
from pushing into the sample too strongly, particularly at steep uphill regions of thesample.
These manipulative operations are implemented using the circuit shown in Fig. 34. The
circuit has three branches in the horizontal direction. The true error signal passes throughthe middle branch. A DC signal corresponding to (Aupper  As) is fed to the input terminalof the upper branch and passes through a subtracter to produce (Ap–p  Aupper). The output is ampliﬁed (with a gain of Gupper), passed through a precision diode circuit, and ﬁnallyadded to the true error signal. The precision diode circuit compensates for an oﬀset inherent in the diode chip. The ﬁnal output, L[Ap–p  Aupper]  Gupper + (Ap–p  As), is fed tothe conventional PID input terminal. Here, L is an operator such that L[x] = x (ifx > 0) or L[x] = 0 (if x < 0). The signal (Ap–p  Aupper) can be constructed directly from

Fig. 33. Schematic showing the principle of the dynamic PID control. Solid line: an amplitude-distance curve;gray line: an error signal used in the conventional PID control; and broken line: an error signal used in the
dynamic PID control [71].

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Fig. 34. Circuit diagram of dynamic operator [71]. For details, see the text.

Ap–p and Aupper. However, in this case, Aupper has to be tuned depending on the value ofAs, because Aupper has to be larger than As. On the other hand, using the method describedabove, a positive DC signal corresponding to (Aupper  As) is tuned independently of thevalue of As. The false error signal, L[Alower  Ap–p]  Glower + (Ap–p  As), is similarly
constructed in the lower branch.

Fig. 35. Feedback bandwidth as a function of the set point r, measured using the mock AFM system. Solid
curves: feedback bandwidths measured using the mock AFM system with a conventional PID controller; dottedcurves: feedback bandwidths measured using the mock AFM system with the dynamic PID controller. The solid
curves and the dotted curves are aligned from top to bottom according to the ratio 2A0/h0 = 5, 2, 1, and 0.5 [71].

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6.7.2. Performance of dynamic PID control
Dynamic PID control signiﬁcantly enhances the feedback bandwidth, particularly when
the set point r is close to 1 (see dotted curves in Fig. 35). The feedback bandwidth is independent of r, provided r is less than 0.9, indicating that parachuting does not occur. The
superiority of the dynamic PID control is also clear from captured images of a mock sample with steep slopes (Fig. 36a). The images were obtained using a mock AFM. Herein, amock cantilever with Q = 3 oscillating at its resonant frequency of 1.2 MHz is scannedover a mock sample surface (rectangular shapes with two diﬀerent heights) from left toright at a scan speed of 1 mm/s (frame rate of 100 ms/frame). Here, the height of the taller
rectangle is 2A0, and As is set at 0.9  2A0. When using the conventional PID controller,
the topographic image became blurred (Fig. 36b) due to signiﬁcant parachuting at steep
downhill regions. On the other hand, the use of the dynamic PID controller resulted ina clear image (Fig. 36c), indicating that almost no parachuting occurred. Dynamic PID
control improves both the feedback bandwidth and the tapping force simultaneously,
and is therefore indispensable in the high-speed AFM imaging of delicate samples. Thisis demonstrated by the high-speed imaging of fragile biological samples (Chapter 9).
6.8. Drift compensator
The drift in the cantilever excitation eﬃciency poses a problem, particularly when r is
close to 1 (i.e., As is set close to 2A0). For example, as the eﬃciency is lowered, 2A0
decreases and, concomitantly, Ap–p decreases. The feedback system interprets this decreasein Ap–p as an overly strong interaction of the tip with the sample and therefore withdraws
the sample stage from the cantilever; this is, of course, an incorrect operation. This withdrawal tends to dissociate the cantilever tip completely from the sample surface, particularly when As is close to 2A0. For example, when 2A0 = 5 nm and As = 4.5 nm (r = 0.9),
their diﬀerence is only 0.5 nm. The satisfactory operation of dynamic PID control undersuch a condition requires high stability of the excitation eﬃciency. The stabilization of theFig. 36. Pseudo-AFM images of a sample with rectangles with two diﬀerent heights. (a) A mock AFM sample.
The images were obtained using a conventional PID controller (b) or using the dynamic PID controller (c). Thesesimulations with the mock AFM system were made under the following conditions: the cantilever resonant
frequency, 1.2 MHz; quality factor of the cantilever oscillation, 3; the resonant frequency of the z-scanner,
150 kHz; and quality factor of the z-scanner, 0.5; the line scan speed, 1 mm/s; the line scan frequency, 1 kHz; theframe rate, 100 ms/frame, the ratio of 2A0 to the total sample height = 1; and r = 0.9 [71].

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excitation eﬃciency was previously attempted by Schiener et al. by using the second-harmonic amplitude of the cantilever oscillation to detect drifts [100]. The second-harmonic
amplitude is sensitive to the tip–sample interaction, and therefore, the drift in A0 is
reﬂected in the second-harmonic amplitude averaged over a period longer than the image
acquisition time. Instead of controlling A0, they controlled As to maintain a constant valueof the diﬀerence (2A0  As). However, this control method varies the tapping force andfeedback bandwidth because the sample height increases relative to the oscillation amplitude. To compensate for drift in the cantilever excitation eﬃciency, we also used the second-harmonic amplitude of cantilever oscillation, but instead of controlling As, we
controlled the output gain of a wave generator (WF-1946A, NF Corp., Osaka, Japan)
connected to the excitation piezoelectric actuator (Fig. 37) [71]. We only used an I-controller whose time constant was adjusted to 1–2 s (about 10 times longer than the image acquisition time). The performance of this method of drift compensation is shown in Fig. 38. A
sample of myosin V bound to actin ﬁlaments in solution was imaged successively for 3 min
at 100 ms/frame. Very stable imaging was achieved, even with the small diﬀerence

Fig. 37. Circuit diagram of compensator for drift in cantilever excitation eﬃciency. The peak frequency of the
band-pass ﬁlter is adjusted at the second harmonic frequency. Instead of using the band-pass ﬁlter and RMS-toDC converter, we usually use a lock-in ampliﬁer.

Fig. 38. Successive imaging of myosin V attached to actin ﬁlaments using a compensator for drift in the
cantilever-excitation eﬃciency. The imaging was successively made for 3 min at frame rate of 10 frames/s andwith A0 = 2.5 nm and r = 0.92. Only ﬁve images obtained at times indicated by arrows are shown. Black line: theoutput from the drift compensator; gray line: the second harmonic amplitude. At 3 min, the compensator was
switched oﬀ [71].

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379(2A0  As) = 0.4 nm. In addition, the fragile actin ﬁlaments did not disassemble during
imaging. The output signal from the slow controller (I-controller) increased with time,
indicating that the cantilever excitation eﬃciency decreased with time. On the other hand,the second-harmonic amplitude remained constant. When the slow controller output was
disconnected at 3 min, no image was obtained, because of the complete detachment of thetip from the sample.

6.9. Line-by-line feed-forward control
As described in Sections 5.4–5.6, one of the limiting factors in the feedback bandwidth of
AFM is the ratio of (2A0  As) [= 2A0(1  r)] to the maximum sample height h0. The difference (2A0  As) between the peak-to-peak free-oscillation amplitude and the set pointpeak-to-peak amplitude should be increased to increase the feedback bandwidth. On theother hand, it is necessary to bring the set point amplitude As close to the free-oscillation
amplitude 2A0 to reduce the loading force of the tip acting on the sample. Thus, the two
aspects of soft tapping and fast imaging are inherently in conﬂict. We have presented a solution to this problem by introducing the dynamic PID controller [71], as described in Section
6.7. An additional approach to solving this problem is to use a two-degrees-of-freedom(2-DOF) controller. This controller consists of two controllers; one is a conventional
PID feedback controller (or dynamic PID controller) and the other is a feed-forward

Fig. 39. Block diagram of two-degrees-of-freedom (2-DOF) control. Feedback and feed-forward controllers are
combined [101].

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open-loop controller that moves the sample stage according to the sample height variation
recorded in the previous line scan [93,101]. The feedback controller thus only compensates
for the height diﬀerence between the previous and current scan lines.
6.9.1. Feed-forward controller
Fig. 39 shows a block diagram of the 2-DOF control scheme that we developed. The
feed-forward controller consists of 12 bit D/A and A/D converters, two memory sets
and two high-speed analog switches. The feed-forward controller starts to record the input
signal into memory 1, which is triggered by the start signal from the scan controller. This
recording is synchronized with the pixel clock signals. After the clock number reaches thepreset number (i.e., the number of pixel  integer in one scan line), the recorded signal isthen output from memory 1, and the input signal is simultaneously recorded into memory
2. These processes are repeated until one frame is acquired. The signal fed to the feed-forward controller input is an appropriately weighted sum of the PID feedback output signal
and the error signal. Note that the error signal is also used to generate the feed-forward
signal. In aqueous environment, the Q-value of the cantilever is not large, and is typicallyless than 3. Therefore, the error signal reﬂects the deviation of the tip trajectory from theactual topography. The use of this error signal to generate the feed-forward signal is essential for compensating the feedback delay. The PID output to be fed to the feed-forward
controller should be attenuated (the gain is usually set at 0.5). This is because, when
the surface geometry is signiﬁcantly diﬀerent between two adjacent lines, the feed-forward
operation would deteriorate the feedback operation rather than alleviate the load. Regarding the error signal to be fed to the feed-forward controller, its input gain can only be
adjusted by considering the relationship between the OBD sensor sensitivity and the piezoelectric displacement eﬃciency.
6.9.2. Performance of feed-forward compensation
To quantitatively evaluate the performance of the feed-forward compensation, we used
the mock AFM circuit which is described in Section 5.6. For this evaluation, the oscillation frequency and amplitude of the mock cantilever were set at 800 kHz and 0.1 V,
respectively. A sinusoidal wave with a frequency of 7.3 kHz and an amplitude of 0.1 V
was used as the mock sample topography.
Fig. 40 shows a comparison of pseudo-AFM images produced by the mock AFM circuit with and without feed-forward compensation. Here, the amplitude set point r was set
at 0.98. The scan direction was from left to right. As shown in Fig. 40a and c, the image
obtained without feed-forward compensation exhibits a distorted sinusoidal shape due tothe parachuting of the tip. On the other hand, as shown in Fig. 40e and g, images obtainedusing feed-forward compensation have a sinusoidal shape without appreciable distortion.
The height of the sinusoidal shape in the image obtained using the feed-forward compensation is 1.8 times larger than that obtained without feed-forward compensation. In addition, as shown in Fig. 40d and h, the feed-forward compensation reduces the error signalby 25%.
To conﬁrm the applicability of the feed-forward compensation, we imaged soft biomolecules. We used microtubules as a test sample. Tubulins were puriﬁed from a pig brain andassembled into microtubules in a buﬀer solution by adding glycerin, guanosine-50 -triphosphate (GTP), and taxol at 37 °C. The microtubules were adsorbed on a mica surface andthen the AFM experiment was performed in the same buﬀer solution containing taxol.

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381

Fig. 40. Pseudo-AFM images obtained using mock AFM under only feedback ((a)–(d)) and 2-DOF control ((e)–(h)). Topography ((a) and (e)), topographic cross-sectional proﬁles ((c) and (g)), error images ((b) and (f)) anderror cross-sectional proﬁles ((d) and (h)). The feed-forward gain is 0.5 [101].

Fig. 41a and b shows topographic images taken with and without feed-forward compensation, respectively. These images were acquired at 1 s/frame, with an amplitude set pointof 0.9 and a free peak-to-peak amplitude of 6 nm. The x-scan direction was from left toright on the images. In the image acquired using the feed-forward compensation(Fig. 41a), a single microtubule with a height of 25 nm as well as isolated protoﬁlaments
can be clearly observed. At the scanning position marked by an arrow in Fig. 41b, thefeed-forward compensator was switched oﬀ and thereafter only the feedback control
was used. Without the feed-forward compensation, the right-hand side of the microtubule
shows tip parachuting and the image quality markedly deteriorates. Furthermore, the isolated protoﬁlaments on the mica surface are not observed. This is because the ratio of the
microtubule height of 25 nm to the peak-to-peak free-oscillation amplitude of 6 nm is toolarge compared with (1  0.9)  6 = 0.6 nm (see Eq. (10)), and therefore, the feedback
bandwidth is not suﬃcient to precisely trace the topography. When the set point amplitude
was reduced without feed-forward compensation to improve the image quality, the micro-

Fig. 41. AFM images of a microtubule on a mica surface obtained (a) with and (b) without the feed-forward
compensation. In (b), the feed-forward control was cut at the position marked by an arrow. In (a) and (b), theamplitude set point r for feedback control was 0.9 and the peak-to-peak free-oscillation amplitude was 6 nm. In(c), the set point was increased to 0.95 without the feed-forward compensation. All images were acquired at 1 s/
frame. The scan size was 200  200 nm2 with 128  128 pixels.

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tubule was completely disassembled. In Fig. 41c, the amplitude set point r was increased toabout 0.95 without the feed-forward compensation. It can be seen that the microtubule isdisassembled at the positions where the feed-forward compensator was switched oﬀ (asindicated in Fig. 41b). From these results, feed-forward compensation is eﬀective for
improving the feedback bandwidth and reducing the loading force on the sample.
6.10. Photothermal actuation of cantilevers
There is a limit to the enhancement of the bandwidth of a piezoelectric actuator-based
z-scanner due to the limited performance of available piezoelectric actuators. The upper
limit seems to be 500 kHz in practice, considering the required maximum displacement
and the eﬀect of the mass of the attached sample stage on the resonant frequency. The tip–
sample distance can be controlled not only by the z-scanner but also by direct cantileveractuation. As small cantilevers have a high resonant frequency in water, the direct actuation of a small cantilever is expected to aﬀord a high bandwidth for the control of tip–sample distance. At present, small self-actuation cantilevers cannot be manufactured. There
have been several reports describing the direct actuation of cantilevers coated with or
attached to ferromagnetic materials using a magnetic ﬁeld [102,103]. Unfortunately, themagnetic coating stiﬀens the cantilevers and the attachment of magnetic materials lowers
the cantilever resonant frequency. In addition, it is diﬃcult to produce a high-frequency
magnetic ﬁeld generator with high bandwidths. Acoustic radiation pressure (ARP) generated by a focused acoustic transducer with a frequency of 100–300 MHz has recently beenused for the direct actuation of a cantilever in liquids [104]. The force generated by the
ARP can be focused on an area of a few microns in diameter, and therefore, can be used
for the direct actuation of a small cantilever. However, the ARP also exerts forces on thesample, and therefore, a special arrangement is required for the implementation of thesample stage and the transducer [105,106]. Also, although it was not mentioned in thesearticles, it is likely that the sample solution temperature is increased by the ARP.
Another technique for direct cantilever actuation is to use a photothermal bending
eﬀect. The irradiation of a laser beam actuates a gold-coated cantilever either by the pressure induced by the radiation or by photothermal expansion [107–110]. In liquids, the latter eﬀect is dominant because of the low quality factor of cantilevers. We used a 980 nm IR
laser for the actuation and a 675 nm red laser for detecting the deﬂection [111]; these laser
beams were focused onto a small cantilever using an objective lens. The irradiation position of the IR laser onto the cantilever was adjusted so that the maximum cantilever deﬂection was attained. The DC deﬂection eﬃciency was about 1.1 nm/mw; the laser power was
not that actually irradiated onto the cantilever but that measured at the exit of the objective lens. The power of the IR laser can be modulated from 0 to 200 mW with a frequency
range of 0–10 MHz, and therefore, the maximum deﬂection of the cantilever should havebeen 220 nm. In practice, the deﬂection was 120 nm due to the partial eclipse of the incident laser light at the objective lens inlet. This deﬂection range is suﬃcient for imaging
protein samples. We attempted to use a 405 nm violet laser for the actuation and obtained
a larger deﬂection eﬃciency of 10 nm/mw. However, the biological samples were damagedby the irradiation.
As expected, the photothermal bending response of the cantilever to the laser power
modulation was slow. The cantilever response to the modulation with a rectangular wave
was well ﬁtted by a double-exponential function with time constants of 8.4 ls and 123 ls

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

383(inset in Fig. 42). This slow response is also apparent in the frequency response (curves
marked with (b) in Fig. 42). As shown with the curves marked with (a) in Fig. 42, this slow
response was successfully eliminated by inverse compensation, using an APITF circuit[111] (Fig. 24b). We examined how a high feedback bandwidth can be achieved by controlling the tip–sample distance using the present photothermal bending method. Under acondition of 2A0/h0 = 5, the feedback bandwidth of 100 kHz was obtained for a small cantilever having a resonant frequency of 1.2 MHz and a quality factor of 3 in water [111].
This feedback bandwidth is 43% higher than the case when the tip–sample distance is controlled by our piezoelectric actuator-based z-scanner with a resonant frequency of 170 kHz
and a quality factor of 0.5. Because of this high feedback bandwidth, myosin V molecules
were clearly imaged at a rate of 32 frames/s over a scan range of 240  240 nm2 with 100
scan lines [111].
Here, note that there is a drawback to direct cantilever actuation when it is used to control the tip–sample distance. The cantilever has to be deﬂected up to the maximum sample
height. Therefore, the dynamic range in the OBD sensor has to be large, resulting in lower
sensor sensitivity. This is independent of the means used for the direct cantilever actuation.
When a piezoactuator-based z-scanner is used to control the tip–sample distance, the DC
deﬂection of the cantilever is smaller than the maximum sample height.
6.11. Data acquisition system
High-speed data acquisition (DAQ) systems are commercially available. We have beenusing DA/AD boards (10 MHz, 12 bits) with a memory of 0.5 MB/channel and direct
memory access (DMA) controller chips, together with software that we have created.
DMA allows high-speed data transfer without imposing a heavy load on the computer
CPU. We can attain a minimum image acquisition time of 3 ms using the DA/AD

Fig. 42. Frequency spectra of the gain (solid lines) and phase (broken lines) for a small cantilever excited by theintensity-modulated IR laser (a) with and (b) without inverse compensation. Insert: The time–domain response of
the cantilever displacement driven by the laser modulated with a rectangular wave, without inverse compensation.
The response shown with dots is ﬁtted by a double-exponential-function curve (solid line), a[1  exp(t/s1)] + b[1  exp(t/s2)] with time constants of s1  1.84 ls and s2  123 ls [111].

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boards. As DAQ systems with higher rates (100 MHz, 12–16 bits) are available, fasterframe acquisition is possible. However, it takes time to display a bitmap image, even whendata are transferred directly to a bitmap memory area. For example, it takes about 6–
10 ms to display a 256  256 pixel image. To avoid a time lag between frames causedby this delay, we have to reduce the number of frames to be displayed. Large amountsof data stream out of high-speed imaging. The maximum number of acquirable successiveframes depends on the size of the computer memory. For a typical memory size of 2 GB
for a personal computer, approximately 20,000 successive images with 12-bit resolution
and 256  256 pixels can be recorded (when memory size required by the operational system is ignored). For a frame rate of 30 ms/frame, we can record continuously for 10 min.
Quantitative descriptions of the AFM image acquisition abilities of a given DAQ system
are given by Fantner et al. [112,113].
7. High-speed phase detector
One of the unique features of tapping-mode AFM, which is the operation mode ofour high-speed AFM, is the capability of simultaneously imaging the topography andcompositional variations of a sample surface [6–8,114–117]. The distribution of material
properties is measured by detecting the phase diﬀerence between the excitation signal
and the cantilever oscillation. The phase diﬀerence is related to several material properties such as viscoelasticity, elasticity, adhesion, hydrophobicity/hydrophilicity [6], andsurface charge [117]. The energy dissipation of an oscillating cantilever due to inelastic
tip–sample interactions is considered to be the main mechanism in the generation ofphase-contrast [7,118]. Combining the information on these properties with a topography with unprecedented temporal resolution would provide a deeper insight into functional mechanisms and dynamic processes of biomolecules. In this chapter, a novel
method of realizing high-speed phase detection and some of its applications aredescribed [119].
For high-speed phase-contrast imaging, the phase diﬀerence must be detected with asuﬃciently higher bandwidth than the feedback bandwidth used for topographic imaging.
A lock-in ampliﬁer, which is usually used in the phase-contrast imaging with a conventional AFM, cannot be applied to high-speed imaging as its bandwidth is too low; it is usually limited to less than kHz even without the use of ﬁlters. To achieve fast phase detection,a novel phase detector [119] was developed on the basis of a design by Stark and Guckenberger [120]. Fig. 43 shows the operational principle of the fast phase detection. A twochannel wave form generator produces sinusoidal and sawtooth waves with a given phase
diﬀerence. The sinusoidal wave is used to oscillate the cantilever. The cantilever oscillation
signal is fed to a phase detector composed of a high-pass ﬁlter, a variable phase shifter, azero-crossing comparator, a pulse generator, and a sample-and-hold (S/H) circuit. A pulse
signal is generated at either the rising or falling edge of the output signal of the zero-crossing comparator. The pulse signal acts as a trigger for the S/H circuit. When the trigger isgenerated, the amplitude voltage of the sawtooth signal is captured and retained by the
S/H circuit. Thus, the sawtooth signal acts as a phase-voltage converter. The variable
phase shifter enables us to vary the timing of the trigger signal generated within the cantilever oscillation cycle. This function is essential for obtaining the optimum phase-contrast because the phase-contrast markedly depends on the trigger-timing, as described in
Section 11.1.2.

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385

Fig. 43. Schematic diagram of the fast phase detection system for AFM [119].

Fig. 44 shows a marked diﬀerence in the phase detection speed between a conventional
RF lock-in ampliﬁer and the fast phase detector. The input signal is a sinusoidal wave with

Fig. 44. Input signals with a frequency of 900 kHz and phase responses measured using (a) a lock-in ampliﬁer
and (b) the fast phase detector. The phase of the input signal is shifted stepwise from 0° to 90° at time zero.
Rectangular waves depicted by broken lines in (b) indicate the trigger signals [119].

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a frequency of 900 kHz. The phase of this sinusoidal wave was shifted stepwise from 0° to
90° at time zero. Using the lock-in ampliﬁer, the input signal oscillates over 20 cycles until
the output phase reaches 90°, as shown in Fig. 44a. The time constant s measured fromthis transient response was evaluated to be 4.5 ls, which corresponds to a bandwidth ofabout 35 kHz. This bandwidth is only 1/2–1/3 of the feedback bandwidth of our highspeed AFM. On the other hand, the fast phase detector responded much rapidly as shownin Fig. 44b. The phase of the input signal was changed at the point indicated by ‘‘A”. The
trigger signal for the S/H circuit is shown by broken lines. The time constant s was evaluated to be 0.12 ls, which corresponds to a bandwidth of about 1.3 MHz. This is muchhigher than the feedback bandwidth of our high-speed AFM. The detected phase shift
did not change stepwise. This is due to the limited bandwidth of the operational ampliﬁers
used after the S/H circuit.
The main source of noise of the fast phase detector is jitter originating from voltage
noise contained in the PSPD output. The RMS value of jitter DtRMS for a sinusoidal wave
containing broadband white noise VRMS is given by
DtRMS ¼

V RMS:
2pAfoscð31Þ

Here, A and fosc are the amplitude and frequency of the sinusoidal wave, respectively. The
jitter gives rise to timing ﬂuctuations in the trigger pulse for the S/H circuit and therefore,
the phase error DPRMS (degrees) is given by
DP RMS ¼

V RMS
360
 360 ¼;
2pA
2pðSNRÞ

ð32Þ

where SNR is the signal-to-noise ratio of the sensor output. For the test signal shown in
Fig. 44b, SNR is about 17 (A = 0.45 V and V RMS ¼ 26 mV). This SNR gives a phase noiseof 3.3°, which is in good agreement with the magnitude of phase noise (3°) evaluated fromthe phase signal shown in Fig. 44b. In the actual experiment, SNR is less than 10. Therefore, the intrinsic phase noise is expected to be larger than 5.7° due to the thermal ﬂuctuations of the cantilever under a low-Q condition. However, since the bandwidth of
1.3 MHz is much higher than the imaging bandwidth (ca. 100 kHz), the phase noise canbe reduced using a low-pass ﬁlter.
8. Devices for atomic-resolution in liquids
The understanding of biological phenomena at the molecular level requires the direct
imaging of individual mobile molecules in real-time as well as in real space under physiological environment. Thus, for bio-AFM, improvement of the spatial resolution is also
important. Recently, there has been a great progress in improving the spatial resolution
for dynamic-mode in-liquid AFM [121,122]. In this chapter, we describe the key featuresof this novel technique [76,123]. Its applications to biological studies are described in
Chapter 12.
8.1. Operation mode
For AFM applications in liquid, contact-mode AFM [1] and AM–AFM [124] havemainly been used. Contact-mode AFM has been used for the subnanometer-scale imaging

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387

of proteins [34] and lipid molecules [125] in model biological membranes, while AM–AFM
has mainly been used for the nanometer-scale imaging of isolated biomolecules [126,127]and cells [35] attached to a substrate. However, the recent improvement in spatial resolution was brought by employing FM–AFM [9] that has mainly been used for atomic-scale
imaging in ultra high vacuum (UHV) environments. In this section, we describe the basic
principles and experimental setup for FM–AFM and its distinctive features compared withother operation modes.
Fig. 45a shows a typical experimental setup for FM–AFM. A cantilever is mechanicallyoscillated at its resonance frequency using a self-oscillation circuit. The cantilever vibration is induced by an actuator next to the cantilever. The cantilever deﬂection is typicallydetected with an OBD sensor. The deﬂection signal output from the pre-ampliﬁer is fed
into a phase shifter which adjusts the phase of the signal so that the phase diﬀerence
between the deﬂection and excitation signals (/) is kept at 90°. The output from thephase shifter is ampliﬁed by an automatic gain control (AGC) circuit to keep the cantilever
oscillation amplitude (A) constant. The output from the AGC circuit is routed back to theactuator to form a self-oscillation loop.
As the cantilever approaches the surface, the tip–sample interaction force (Fts) induces aresonance frequency shift (Dfc) given by [128]fc
Dfc ðzÞ ¼
2k c

pﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
oF ts
A2  Dz2ðz þ DzÞ
dðDzÞ;
pA2 =2
A oz

Z A

ð33Þ

where kc is the cantilever spring constant and z is the vertical tip position. This behavior isschematically shown in Fig. 45b. Eq. (33) shows that Dfc is proportional to the force gradient integrated over the cantilever oscillation range with a weight function. If A is smallenough for the force gradient to be assumed constant, Eq. (33) can be simpliﬁed to [9]Dfc ðzÞ ¼ 

fc oF ts:
2k c oz

ð34Þ

Fig. 45. (a) Typical experimental setup for FM–AFM. (b) Schematic model of amplitude and phase vs. frequency
curves of a cantilever. Dotted and solid lines show the curves with and without tip–sample interaction,
respectively.

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As the amplitude vs. frequency curve is shifted, the phase vs. frequency curve is also
shifted by Dfc. Since / is kept constant at 90° with the self-excitation circuit, the cantilever oscillation frequency (f) changes with the change in resonance frequency (fc), as
shown in Fig. 45b. Thus, Dfc is measured by directly measuring the frequency change ofthe cantilever vibration. In the FM–AFM setup, the deﬂection signal is fed into a frequency detector which outputs a voltage signal proportional to the frequency of the input.
A phase-locked loop (PLL) circuit is most commonly used as the frequency detector. The
Dfc signal is fed into a PI controller which controls the vertical position of the sample toregulate the tip–sample distance.
The advantages of using FM–AFM over contact-mode AFM include the precise control of the vertical tip position without instabilities especially in the short-range interaction
regime, the capability of imaging isolated molecules weakly bound to a substrate, andenhanced sensitivity to the short-range interaction force [129] and hence a higher spatial
resolution [130]. The advantages of FM–AFM over AM–AFM are still under discussion.
One of the clear advantages of the former is the capability of the quantitative and independent measurement of conservative and dissipative forces [131]. Practically, the frequency
signal in FM–AFM changes much more rapidly than the amplitude signal in AM–
AFM, which facilitates the precise control of the tip position near the surface [121]. This
does not necessarily mean that FM–AFM has a higher signal-to-noise ratio in the force
measurement but does explain why FM–AFM has been used for reproducible atomic-scale
imaging both in vacuum [10] and liquids [132].
8.2. Cantilever stiﬀness
In contact-mode AFM and AM–AFM for biological applications, relatively soft cantilevers (typically kc < 0.1 N/m) have been used while stiﬀ cantilevers (typically kc > 10 N/m) have been used for high-resolution imaging with FM–AFM. In this section, we
describe the necessity of using a stiﬀ cantilever and its merits and demerits.
The RMS amplitude spectral density (nzB) of cantilever Brownian vibration induced bythermal energy is given by [9]sﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
2k B T
1
nzB ¼
;
ð35Þ
pfc k c Qc ½1  ðf =fc Þ2 2 þ ½f =ðfc Qc Þ2where kB, T, and Qc are Boltzmann’s constant, the absolute temperature and the Q factorof the cantilever, respectively. The total RMS vibration amplitude zB induced by the thermal energy is obtained by integrating nzB over the whole frequency range or directly fromthe equipartition theorem and given by [9]sﬃﬃﬃﬃﬃﬃﬃﬃ
kBT
:
ð36Þ
zB ¼
kc
For cantilevers with kc of 0.1 N/m and 10 N/m, zB is about 0.2 nm and 0.02 nm, respectively. This thermal vibration amplitude ultimately determines the accuracy of control
over the vertical tip position. Note that the thermal vibration can be reduced once the cantilever is in contact with the surface, such as in contact-mode AFM. However, delicate true
atomic-resolution imaging requires precise control of the vertical tip position in the tip–

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389

sample distance range from near contact to atomic-scale contact. Therefore, the high stiﬀness of the cantilever is essential for true atomic-resolution imaging as well as for atomicscale manipulation.
The RMS frequency of noise spectral density (nf) arising from the thermal cantilevervibration is given by [76]sﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
k B Tfc:
ð37Þ
nf ¼
pk c Qc A2
This equation shows that nf decreases with increasing kc. From Eqs. (34) and (37), the minimum detectable force gradient j oFozts jmin for small amplitude approximation is given by [9]sﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
oF ts
4k c k B TB¼;
ð38Þ
oz minpfc Qc A2
where B is the measurement bandwidth. If the force gradient is approximately constant,the magnitude of the conservative force (Fc) is proportional to A. Thus, the minimum
detectable conservative force is given by [133]sﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
4k c k B TB:
ð39ÞðF c Þmin ¼
pfc Qcpﬃﬃﬃﬃ
This equation shows that (Fc)min increases proportionally to k c with increasing kc. Practically, a cantilever with a higher value of kc tends to have higher values of fc and Qc, which
partially compensate for the reduction of force sensitivity due to the high value of kc.
Table 1 shows the parameters of typical soft and stiﬀ cantilevers and those of a smallcantilever designed for high-speed AFM (Section 6.1). This table shows that the practical
diﬀerence in the force sensitivity between soft and stiﬀ cantilevers is not signiﬁcant. Furthermore, the high values of fc and Qc greatly facilitate ﬁnding a correct cantilever resonance and stabilize the self-oscillation in FM–AFM.
The small cantilever has a high value of fc in spite of the small value of kc. Thus, it
should provide better force sensitivity than the other cantilevers with parameters shownin Table 1, although this has not yet been experimentally conﬁrmed for FM–AFM. It
should also allow the tip to be scanned more than 10 times faster than that of the othercantilevers. Note that this cantilever still suﬀers from problems such as large thermal
vibration amplitude (Eq. (36)) and the instabilities described below. Therefore, it should
Table 1Comparison of various parameters in water between typical soft and stiﬀ cantilevers and a small cantilever
designed for high-speed AFM

Typical soft cantilever
Typical stiﬀ cantilever
Small cantileverkc (N/m)fc (kHz)Q(Fc)min (pN)
(B = 1 kHz)nzB (fpc)ﬃﬃﬃﬃﬃﬃﬃ(fm/ Hz)

0.1
30
0.2

5
130
1200

1.5
8
2.5

8
12
0.6

2800
74
165

kc, spring constant; fc, ﬁrst resonant frequency; Q, quality factor; (Fc)min, the minimum detectable conservative
force; nzB (fc), RMS amplitude spectral density of cantilever Brownian vibration at fc.

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be used for applications where ultimate spatial resolution is not necessary but high force
sensitivity and high scanning speed are essential.
As a tip approaches a surface, the attractive force between the tip and the sample
increases. Thus, the tip experiences an increase in the force gradient. In contact-mode
AFM, the tip suddenly jumps and comes in contact with the surface at a distance where
the force gradient exceeds kc. Therefore, the following condition must be satisﬁed forthe measurement of force without tip adhesion [134]:
kc >

oF ts
:
oz

ð40Þ

Such adhesion is referred to as ‘‘jump-to-contact” and hinders precise control of interaction between the tip front atom and the surface atom or molecules. Since the operation ofcontact-mode AFM requires soft cantilever, the only way to meet this condition is to adjust the solution conditions such as adding salts [135]. This practically limits the application range.
In dynamic-mode AFM, the tip adhesion can be prevented by increasing A to meet thefollowing condition [136]:
k c A > jF ts j:ð41Þ

Thus, a soft cantilever can be used without limitations in solutions, provided A is suﬃciently large. However, the use of large amplitude practically limits the spatial resolution
as explained below. Namely, high-resolution AFM imaging requires small amplitude operation without instabilities, which in turn requires high stiﬀness of the cantilever.
8.3. Oscillation amplitude
In AM–AFM, A has been set at 5–50 nm while the novel FM–AFM technique utilizes
an amplitude of 0.1–1 nm. In this section, we describe the necessity of using small oscillation amplitude and the practical requirements for realizing this condition.
Fts includes long-range forces (e.g., electrostatic force and van der Waals force) andshort-range forces (e.g., chemical interaction). To achieve true (i.e., non-averaged)
atomic-resolution in AFM, the short-range interaction between the tip front atom andthe surface atom or molecule must predominantly contribute to the distance feedback signal (i.e., z in contact-mode AFM, A in AM–AFM, Dfc in FM–AFM).
The reduction of A in dynamic-mode AFM allows the tip to stay in the short-range
interaction regime (typically z < 1 nm) for a longer time in an oscillation period. In the
short-range interaction regime, the eﬀect of the long-range interaction force on the force
gradient is negligible compared with that of the short-range interaction force. Therefore,
the use of small amplitude enhances sensitivity to the short-range interactions andreduces sensitivity to the long-range interactions, leading to a higher spatial resolution[129,130].
Noise in the distance feedback control (dz) in FM–AFM is determined by the ratio
between the frequency noise df and the slope of the Dfc vs. distance curve
dz ¼

df
:
oðDfc Þ=oz

ð42Þ

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From Eq. (37), df for bandwidth B is given bysﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
k B Tfc B
df ¼:
pk c Qc A2

391

ð43Þ

This equation shows that df is proportional to A1, whereas o(Dfc)/oz is known to be constant for small amplitudes and proportional to A3/2 for large amplitudes [129]. From Eq.
(42), dz is proportional to A1 for small amplitudes and A1/2 for large amplitudes. Therefore, there is an optimal amplitude with a minimum value of dz. This optimal amplitude
has been reported to be on the order of the decay length of the force components [129]; on
the order of 0.1 nm and 1 nm for short-range and long-range interaction forces, respectively. Therefore, small amplitude operation improves the sensitivity to the short-range
interaction force.
8.4. Deﬂection sensor noise
The noise densityofﬃ cantilever deﬂection sensors used for conventional AFMs is typipﬃﬃﬃﬃﬃﬃ
cally 100–1000 fm/
Hz[137] while that for high-resolution FM–AFM imaging in liquid ispﬃﬃﬃﬃﬃﬃﬃ
less than 20 fm/ Hz [76,123]. In this section, we describe the necessity of such a low-noisedeﬂection sensor and technical details for realizing such an excellent noise performance.
A deﬂection signal contains two major noise components. One is from the thermal
Brownian vibration of the cantilever and the other is from the deﬂection sensor itself.
The RMS value of the deﬂection noise density due to the thermal noise (nzB) is describedby Eq. (35), while that from the deﬂection sensor nzs can be regarded as a white noise at afrequency range relevant to dynamic-mode AFM. For dynamic-mode AFM, noise components in the frequency range from fc  B to fc + B inﬂuence its performance. Therefore, inorder to achieve the theoretically-limited noise performance given by Eqs. (38) and (39),nzs must be reduced to less than nzB in this frequency range.
From Eq. (35), nzB, at f = fc is given bysﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
2k B TQc:
ð44Þ
nzB ðfc Þ ¼
pfc k cFor theptypical
soft andpﬃﬃﬃﬃﬃﬃ
stiﬀ
ﬃﬃﬃﬃﬃﬃﬃ
ﬃ cantilevers with parameters shown in Table 1, nzB(fc) is
2.8 pm/ Hz and 74 fm/ Hz, respectively. Eq. (44) and these values show that a stiﬀer
cantilever has a lower thermal noise peak and hence requires a lower noise level of thedeﬂection sensor to achieve the theoretically-limited noise performance. In order to suppress the inﬂuence of nzs to less than 10% of that of nzB, nzs must be less than 1/3 oftheﬃ typical stiﬀ cantilever with parameters given in Table 1, nzs mustnzB. For
pﬃﬃﬃﬃﬃﬃbe
ﬃ less thanpﬃﬃﬃﬃﬃﬃ
25 fm/ Hz, which is much lower than a typical value (nzs = 100–1000 fm/ Hz) for a conventional deﬂection sensor.
The noise performance of a deﬂection sensor is also important for operating FM–AFM
with small oscillation amplitude. FM–AFM utilizes a deﬂection signal for producing acantilever excitation signal as well as for detecting Dfc. In a self-oscillation circuit, noise
components that aﬀects the cantilever oscillation spectrum are in the range from fc  fc/(2Qc) to fc + fc/(2Qc). Thus, in order to make the oscillation spectrum freepﬃﬃﬃ from the inﬂuence from the psensor
ﬃﬃﬃﬃﬃﬃﬃ noise, the above-mentioned criterion becomes 1= 2 times smaller,
namely 18 fm/ Hz

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As discussed above, true atomic-resolution imaging in liquid with dynamic-mode AFM
requires the use of a stiﬀ cantilever and a small oscillation amplitude, which in turnrequires a low-noise cantilever deﬂection sensor. Among various methods proposed for
detecting cantilever deﬂection, the OBD method is the most widely used technique due
to its simple setup and the ease of optical beam alignment. In the rest of this section,pﬃﬃﬃﬃﬃﬃﬃ
we describe how to improve the value of nzs for an OBD sensor to less than 18 fm/ Hz.
Fig. 46 shows a typical setup of an OBD sensor used for liquid-environment. A focusedlaser beam is irradiated onto the backside of the cantilever and the reﬂected beam is detectedusing a PSPD. The cantilever deﬂection results in an angular deviation of the reﬂected laser
beam, which in turn gives rise to a displacement of the laser spot on the PSPD. The PSPD
produces a diﬀerential current signal proportional to the laser spot displacement. The
current signal is converted to a voltage signal using an I–V converter and is fed into a
diﬀerential ampliﬁer, generating a voltage signal proportional to the cantilever deﬂection.
Major noise sources in the OBD sensors include noise arising from the laser beam and thephotodiode (PD) shot noise. Theoretically, the PD shot noise should determine the minimum
noise obtained using the OBD sensor. However, the noise performance of practical OBD sensors is often limited by the noise related to the laser beam. Here we presents a way to reduce
the laser-related noise to less than that of the PD shot noise, and a method of reducing theshot noise to achieve the theoretically-limited noise performance in FM–AFM.
A laser diode operated with a relatively small output power compared with its ratingshows intensity ﬂuctuations originating from spontaneous light emission. This noise isreferred to as ‘‘quantum noise”. As the laser power increases, the quantum noise becomes
negligible compared with the average laser power. Instead, the intensity ﬂuctuation due tothe hopping of the laser oscillation mode becomes evident in the high-output-power
regime. This noise is referred to as ‘‘mode-hop noise.” The intensity ﬂuctuation inducedby the quantum noise is eliminated using a diﬀerential ampliﬁer as a common-mode noise.
In contrast, that from the mode-hop noise is magniﬁed as a diﬀerential noise, which thus
signiﬁcantly increases the deﬂection noise.
The magnitude of the mode-hop noise does not show a monotonic dependence on the
output laser power but exhibits a steep increase at a threshold value [76]. Thus, the laser
power should not exceed the threshold for the onset of laser mode hopping. Since a high-Fig. 46. Typical experimental setup for an OBD sensor for liquid-environment AFM [123].

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output laser power is desirable for reducing the noise from the PD shot noise, the laser
power should be set at a value slightly below this threshold.
In addition to the intrinsic noises discussed above, there are two extrinsic noise sources
related to the laser beam. The reﬂection and scattering of the laser beam occasionallycause part of the laser beam to return to the optical resonator of the laser diode. This optical feedback generates additional laser oscillation modes, leading to an increase in mode
hopping. This noise is referred to as ‘‘optical feedback noise”. On the other hand, some of
the reﬂected or scattered laser beam is incident to the PSPD. This laser beam interfereswith other laser beams which travel through diﬀerent optical paths. Such unexpected optical paths are often unstable and hence the laser spot pattern on the PSPD shows a ﬂuctuation over a relatively long time scale. This noise is referred to as ‘‘optical interference
noise”. These extrinsic laser noises are particularly evident in liquid-environment AFMs.
In the setup for liquid-environment AFM, many interfaces exist in the optical path of thelaser beam. Such interfaces enhance the reﬂection and scattering of the laser beam, leading
to an increase in the optical feedback noise and optical interference noise [76].
The extrinsic laser noise can be suppressed by modulating the laser power with an RF
signal whose frequency is typically 300–500 MHz [138,139]. The RF laser power modulation changes the laser oscillation mode from single-mode to multimode. Since the mode
hopping takes place due to the competition among the possible laser oscillation modesin the optical resonator, the multimode laser oscillation is much less sensitive to the opticalfeedback than the single-mode laser oscillation. Consequently, RF modulation considerably reduces the mode hopping induced by the optical feedback. In addition, the multimode laser beam has lower coherence than the single-mode laser. Thus, RF modulationis also eﬀective for suppressing the optical interference noise.
Fig. 47 shows a waveform of a Df signal measured in water before and after turning oﬀ
the RF laser power modulation [76]. The magnitude of the frequency noise suddenly
increased after turning oﬀ the RF modulation, which shows that the RF modulation iseﬀective in reducing the extrinsic laser noises. Although RF modulation also reduces
the intrinsic mode-hop noise, it is often diﬃcult to completely suppress it. Thus, the average laser power should be set at a value below the threshold for the onset of the mode-hop
noise even with the RF laser power modulation.
Once the eﬀect of the laser beam noise is suﬃciently reduced, the PD shot noise starts topredominantly aﬀect the deﬂection noise. The RMS deﬂection noise density arising fromthe PD shot noise (nzp) is given by [123]Fig. 47. Waveform of the frequency shift signal measured in water before and after turning oﬀ the RF laser
power modulation. A = 5 nm, f0 = 140 kHz, k = 42 N/m [76].

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a0 ‘ cnzp ¼
6vnm ‘fsﬃﬃﬃﬃﬃﬃﬃﬃﬃﬃ
2e
;
gaP 0

ð45Þ

where g is the eﬃciency of photo-current conversion of the PSPD, a is the laser power
attenuation in the optical path from the laser source to the PSPD, e is an electron charge,
and nm is the refractive index of the medium of the operating environment. P0 is the laser
power measured at the output of the beam source. ‘f, ‘c, a0 are the focal length, cantilever
length, and the diameter of the collimated laser beam, respectively. v is the correction factor for a Gaussian laser beam. If we deﬁne a0 as the diameter of the beam cross-section at
1/e (or 1/e2) of the maximum laser intensity, v is 1.13 (or 1.60). The equation gives us adirect guideline for reducing the deﬂection noise. Namely, increasing ‘f, a and P0, anddecreasing a0 and ‘c will improve the noise performance of the deﬂection sensor using
the OBD method.
Fig. 48 shows a deﬂection noise density spectrum measured in water using a low-noise
OBD sensor developed as discussed above [123]. The experimentally measured spectrum(solid line) shows good agreement with the thermal noise spectrum (dashed line) calculatedwith Eq. (35)aroundthe resonance frequency. Such a low deﬂection noise density
pﬃﬃﬃﬃﬃﬃ
ﬃ(nzs = 16 fm/ Hz) ensures the theoretically-limited noise performance in FM–AFM evenwith a stiﬀ cantilever (kc = 21 N/m).
9. Imaging of dynamic biomolecular processes
Presently, the world wide number of high-speed AFM users is small. In addition, highspeed AFM has not yet been completely established as a tool for studying dynamic biomolecular processes, although the performance of high-speed AFM has been markedly
improved in the last 3–4 years. In exploring the potential of high-speed AFM, the ﬁrst step
must be to demonstrate biological processes that have been predicted or are known tooccur. As shown below, such demonstrations (including unpublished results) have beensuccessfully presented very recently at our laboratory and other places. Exceeding the level

Fig. 48. Frequency spectra of cantilever thermal Brownian motion measured in water. The black line shows
experimentally measured values while the dotted line shows theoretically calculated values for thermal Brownian
motion with Eq. (35). k = 21 N/m, f0 = 123 kHz, Q = 6.5 [123].

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of just demonstrations, some of these studies were able to give insights into the functional
mechanisms of proteins. In this chapter, we introduce these successful recent studies
together with early pioneering studies performed to develop this unprecedented ﬁeld ofhigh-speed bio-AFM studies. We hope that readers will be encouraged and convincedof the use of this new microscopic tool to ﬁnd the answers to numerous unsolved biological questions.
9.1. Early imaging studies
In this section, we describe early imaging studies using high-speed AFM performed at
our laboratory between 2001 and 2004.
9.1.1. Initial attempts and lessons
In 2001, the AFM imaging of moving protein molecules at an imaging rate of 80 ms/
frame was successfully achieved for the ﬁrst time [54]. This encouraged us to image various
biomolecular processes. However, we soon realized that the tip–sample interaction force
was too strong to image fragile samples such as actin ﬁlaments and microtubules, whichwere easily destroyed during imaging (Fig. 49). This was due not only to the insuﬃcient
feedback speed but also to the large tapping force exerted by the oscillating tip on the sample. Thus, imaging studies in this early stage were mainly performed to evaluate the capacity of this new microscope and to determine which devices had to be improved.
A prototype dynamic PID controller [140] and an active damping technique for the zscanner [92] were developed during this period. Using the dynamic PID controller, the tapping force was greatly reduced and the feedback bandwidth was enhanced. For example,
the unidirectional movement of individual kinesin molecules along microtubules wasFig. 49. Actin ﬁlaments and microtubules being disrupted during imaging. The number attached to each image
indicates the time (s) elapsed after imaging began. (Top panel) actin ﬁlaments with myosin V in ATP; imaging
rate, 80 ms/frame; scan size, 400 nm. (Lower panel) microtubules; imaging rate, 460 ms/frame; scan size, 200 nm.

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Fig. 50. Movement of kinesin–gelsolin along a microtubule. The kimera protein is a homodimer composed of
two heavy chains, each of which consists of the head–neck region of kinesin at the N-terminal and gelsolin at the
C-terminal. 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 [140,141].

observed (Fig. 50) without disassembling the microtubules [140]. However, in the observedkinesin movement, the kinesin molecules were attached to both the mica surface and themicrotubules. The movement of kinesin that was attached only to microtubules was notobserved. On the other hand, in the presence of AMP–PNP, wherein kinesin ﬁrmly
attaches to microtubules, kinesin molecules could be observed on the top surface of the
microtubules (Fig. 51). These observations imply that the tip removed the kinesin that
had been attached to only the microtubules in the presence of ATP. While observing

Fig. 51. Microtubule associated with kinesin–gelsolin in the presence of AMP–PNP. White arrows indicate
kinesin–gelsolin bound to the microtubule top surface while red arrows indicate kinesin–gelsolin bound to themicrotubule at the side surface. Scale bar, 20 nm.

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Fig. 52. Actin ﬁlament gliding over a surface that is densely coated with 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 [141].

the movement of actin ﬁlaments, we experienced a similar problem. When a mica surface
was densely coated with myosin V, the gliding movement of actin ﬁlaments was captured
on video (Fig. 52) [87,141]. However, when the myosin V density was lowered, actin ﬁlaments did not appear in the images, suggesting that they were removed by the scanningand oscillating tip. These trial observations indicated that, using the dynamic PID controller, the tip–sample interaction force could be successfully reduced to a level at which protein inactivation was prevented and self-assembled polymer were not disassembled.
However, the force was not reduced suﬃciently to prevent the delicate protein–protein
association from breaking.
9.1.2. Flash photolysis combined with high-speed AFM
It is important to observe how a protein conformation changes dynamically upon binding to its ligand. Protein molecules always exhibit Brownian motion due to the thermal
agitation from surrounding water molecules. Therefore, changes in protein shape causedby this random agitation have to be distinguished from ligand-induced conformationalchanges. When the ligand association–dissociation kinetics is fast, the ligand-inducedchanges occur repeatedly during a reasonable imaging period. A structural change thatis repeatedly observed can be judged to be one induced by the ligand reaction. On theother hand, when the kinetics is slow, this distinction is diﬃcult. For example, becauseof the low ATPase activity of myosin V alone (0.05/s), it was diﬃcult to judge whether
structural changes observed only once or twice were in fact those induced by the binding ofATP (or its hydrolysis) (Fig. 53).

Fig. 53. 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, appears to change its conformation from an extended
form to a folded form. ATP concentration is 2 lM. Imaging rate, 0.5 s/frame; scan range, 300 nm [3].

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To solve this problem, we combined the UV ﬂash photolysis of caged compounds withhigh-speed AFM. To generate UV ﬂashes, we ﬁrst used a mercury lamp equipped with a
UV ﬁlter and a mechanical shutter. This photolysis system was used together with caged
calcium to observe Ca2+-induced calmodulin (CaM) dissociation from the myosin V neckregions [140]. Myosin V possesses six IQ motifs at each extended neck domain, to which a
number of CaM molecules (5) and light chains (1) are attached in the absence of Ca2+
[142,143]. In the presence of Ca2+, some of these CaM molecules dissociate from myosin V[144,145]. The sample stage unexpectedly moved in the x-direction during the applicationof a UV ﬂash for 1 s and soon returned after shutting oﬀ the UV light. This movement wascaused by thermal expansion of materials exposed to the UV light. All the calmodulinmolecules (and also the light chains) dissociated instantaneously from the neck regions,which resulted in very thin necks. In addition, the coiled-coil part, which had connected
the neck–neck junction and the PEST (the amino acid motif that is thought to target cytoplasmic proteins for rapid proteolytic degradation) site, appeared to unwind. These
actions are in strong contrast to the recently established knowledge on Ca2+-induced
CaM dissociation (i.e., CaM dissociates only from the second IQ motif on each neck[146]). It appears that agitation caused by the UV-induced thermal expansion confused
the feedback operation, resulting in an excessive tip–sample interaction force.
We improved the combined AFM system using attenuated high-frequency laser pulses(50 kHz, 355 nm). Laser pulses were applied while the y-scan was performed towards thestarting point after the completion of one frame acquisition. In addition, during this period, the sample stage was withdrawn from the cantilever tip. This improved method
allowed us to observe the rotational movement of the myosin V head around the head–neck junction that occurred immediately after UV application to the caged ATP-containing solution [141] (Fig. 54). We also applied this method to observe the height changes inchaperonin GroEL upon binding to ATP and GroES [87,141] (Fig. 55). The ATP-induced
movement of myosin V on an actin ﬁlament was also imaged, although this rapid movement was too fast to be resolved in several frames.
9.1.3. Dynein C
Single-headed dynein C from chlamydomonous ﬂagella has a ring-shaped head fromwhich a stem and a stalk protrude. The stalk interacts with microtubules at its globular
distal end. Previous electron micrographs of dynein C revealed that the angle between
the stalk and the stem diﬀered between the nucleotide-free state and the ADP-Vi boundFig. 54. 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 [141].

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Fig. 55. GroES binding to GroEL immediately after ATP release by ﬂash-photolysis of caged-ATP. The number
attached to each image indicates the time (s) elapsed after imaging began. UV ﬂash light was applied shortly
before the frames marked with ‘‘F”. Scale bar, 100 nm; Imaging rate, 1 s/frame [141].

state [147]. We conﬁrmed this result using AFM in aqueous solutions. Since dynein C has
a relatively high ATPase rate even in the absence of microtubules, it is easy to distinguish
the ATPase-driven conformational change from Brownian motion. In fact, as shown in
Fig. 56, the 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 [87,141]. The two positions approximately corresponded to the nucleotide-freeand ADP-Vi bound states. Furthermore, the frequency of the stem movement was similar
to the ATPase rate, indicating that the observed periodic stem movement is driven by theATPase reaction.
9.2. Recent imaging studies
In this section, we introduce successful imaging studies performed on several dynamic
biomolecular processes. As the original manuscripts for most of these studies are now in
preparation, some of the imaging data are not displayed here.
9.2.1. Myosin V
From early imaging studies, we learned that reduction in the tapping force was essential
for the establishment of high-speed AFM for biological studies. To increase the eﬀectiveness of dynamic PID control, we developed a low-noise sensor ampliﬁer. Owing to theFig. 56. Periodic structural change in dynein C. The left panel is a schematic drawing of dynein C. The stem stays
mainly at the position shown in the frames 51 and 64 (corresponding to the nucleotide-free state), andoccasionally moves to the position shown in the frame 63 (corresponding to the ADP-Vi-bound state). Scale bar,
30 nm; imaging rate, 160 ms/frame [87]. The dynein C sample was gifted from K. Ooiwa and H. Sakakibara(Kansai Advanced Research Center).

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low-noise, we could use a set point r  0.9, which made it possible to observe actin ﬁlaments gliding on a mica surface sparsely coated with myosin V (Fig. 57) [87,141]. The
bound heads of myosin V were oriented at an angle relative to the actin ﬁlament, similar
to the well-known 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 minus end (i.e., the natural direction). However, it was not evident whether large conformational changes in the bound heads occurred while the actin ﬁlament was moving
unidirectionally.
Despite this successful observation, we were unable to observe the processive movementby single myosin V molecules along actin ﬁlaments. In low-ionic-strength solutions, myosin V tended to attach to the mica surface. In high-ionic-strength solutions, it mostly
remained free from the surface 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 nm to 1 nm, thereby reducing the tip–sample interaction force but 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 myosin V molecule alternated between leading
and trailing positions with a walking stride of 72 nm, i.e., in a hand-over-hand manner,

Fig. 57. Three-dimensional images of actin ﬁlament sliding movement. The number attached to each image
indicates the time (s) elapsed after initiating began. Scale bar 30 nm, imaging rate, 180 ms/frame [141].

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as also demonstrated by single-molecule ﬂuorescence microscopy [148–151]. High-speed
AFM revealed the movement in greater detail than ﬂuorescence microscopy. The leadlever-arm bent immediately before the rear head detached from the actin ﬁlament. Thisbent form was similar to that shown previously by electron microscopy [152]. The
detached rear head rotated around the junction between the two lever-arms and was thenattached to a frontward actin. Immediately after the attachment, the new lead head was
hopping and sometimes moved forward or backward by 5 nm along the actin ﬁlament.
In the absence of nucleotide or in the presence of ADP, only one head of a myosin V molecule was bound to an actin ﬁlament. From the orientation of the bound head, it was clearthat the bound head was in the trailing position. On the other hand, in the presence of a
medium concentration of AMP–PNP, both heads were associated with an actin ﬁlament
for a long time. This indicates that upon binding to AMP–PNP, the lead head conformation adapts itself so that it is able to bind to actin by rotating around the head–neck junction. Therefore, when myosin V is bound to an actin ﬁlament at both heads in an ATPcontaining 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 the phosphate
is released, the lead head rotates back to the previous orientation, which is unfavorable foractin binding. However, the lead head cannot dissociate from the actin because the ADPbound state has a high aﬃnity for actin. This energetically unfavorable conformation
causes the lead neck to bend forwards, thereby pulling the trailing head, so that it detaches
from the actin and is then conveyed forward. By these mechanical processes along thechemical reaction pathway, the two heads alternate their positions to walk along an actin
ﬁlament.
9.2.2. Chaperonin GroEL
Here, we show an example of high-speed AFM imaging by which a long-lasting controversial question regarding a biomolecular reaction was solved. Chaperonin GroEL consists of 14 identical ATPase subunits that form two heptameric rings stacked back toback [153,154]. A series of biochemical studies [155,156] have shown that there is positive
cooperativity in the binding and hydrolysis of ATP in the same ring, whereas there is negative cooperativity between the two rings. Owing to this negative cooperativity, it has beenpresumed that GroEL binds to GroES at one ring while releasing GroES from the other[157–159]. This alternate on–oﬀ switching appears to exclude the concomitant binding of
GroES to both rings of GroEL. However, this issue had remained controversial [160–164],
since some electron micrographs have shown a complex of GroES–GroEL–GroES with afootball shape. To place GroEL on a substratum in a side-on orientation so that bothrings are accessible to GroES, we prepared GroEL biotinylated at the equatorial domains[165]. GroEL was attached to streptavidin 2D crystal sheets prepared on planar lipid bilayers containing a biotin lipid. We could capture the alternate association and dissociation
events at the two GroEL rings. Surprisingly, before the alternate switching took place, afootball structure often appeared. Thus, the high-speed AFM observation clearly solved
the controversial issue.
9.2.3. Lattice defects in 2D protein crystals
As described in the next chapter, we have being preparing substratum surfaces by forming streptavidin 2D crystals on supported planar lipid bilayers [166]. The 2D crystal surfaces are useful for the selective attachment of biotinylated molecules. When we were402

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Fig. 58. Movement of point defects in 2D streptavidin crystal formed on supported lipid planar bilayer
containing biotinylated lipid. The point defects indicated by arrow heads are maintained as point defects during
the observation. Scan range, 150 nm; imaging rate, 0.5 s/frame [168].

observing the crystal surfaces with orthorhombic C222 symmetry, we noticed that latticedefects were sometimes formed at a few places and moved over the surfaces (Fig. 58). This
movement is caused by exchanges between the defect (empty) site and one of the surrounding ﬁlled sites, as no free streptavidin exists in the bulk solution. To study the vacancy
defect mobility and its participation in the crystal growth, monovacancy defects in thestreptavidin 2D crystals were systematically produced by increasing the tapping force onto
the sample from the oscillating tip, similarly to a method used previously [167]. Unexpectedly, this movement had a preference for one direction relative to the other [168]. The
movement projected onto each lattice axis showed a time–displacement relationship characteristic of Brownian motion but the diﬀusion coeﬃcients diﬀered depending on the axes.
Analysis of this dependence revealed that biotin binding aﬀects the contact aﬃnity
between two adjacent streptavidin molecules. Streptavidin is a homo-tetramer with subunits organized in dihedral D2 symmetry. Therefore, in the 2D crystals, biotin only binds
to the two subunits facing the planer lipid bilayers (Fig. 59a). There are two types of subunit–subunit interactions between adjacent streptavidin molecules: interactions between
biotin-bound subunits and interactions between biotin-unbound subunits (Fig. 59b).
One crystallographic axis (a-axis) is comprised of contiguous biotin-bound subunit pairs

Fig. 59. (a) Schematic of a streptavidin molecule on biotinylated lipid bilayer. Two biotin binding sites occupied
by biotin are indicated by the closed circles. The open circles indicate biotin binding sites facing the aqueous
solution and are biotin-free. (b) Schematic of streptavidin arrays in C222 crystal. Unit lattice vectors areindicated: the a-axis includes rows of contiguous biotin-bound subunits, while the b-axis includes rows ofcontiguous biotin-unbound subunits [168].

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whereas the other axis (b-axis) is comprised of contiguous biotin-unbound subunit pairs. A
previous study on the formation of streptavidin 2D crystal on supported planar lipid bilayers suggested that the interaction between biotin-unbound subunits is stronger than thatbetween biotin-bound subunits [169], while another study suggested the reverse relationship [170]. Our analysis of the lattice defect movement clearly revealed that the interaction
between biotin-bound subunits is weaker. In addition, from the diﬀusion constant ratio(Db/Da  2.4), the diﬀerence in the association free energy of the two types of subunit–subunit contacts were quantiﬁed to be 0.88 kBT (T  300 K), which corresponds to0.52 kcal/mol [168].
Fusion of two point defects into a larger point defect was often observed (Fig. 60). On
the other hand, ﬁssion of a multivacancy point defect into smaller point defects was rarely
observed [168]. We think that the ﬁssion often occurs but cannot easily be observed
because the two point defects formed immediately after the ﬁssion are quickly fused again.
Immediately after the ﬁssion, a streptavidin molecule becomes facing both the resulting
two smaller point defects. This streptavidin molecule contains only two intermolecularsubunit–subunit contacts, and therefore, quickly detaches from the lattice and moves intoeither of the two adjacent point defects. The imaging rate we used (0.5 s/frame) was not
suﬃciently high to observe this transiently occurred ﬁssion.
During the formation of streptavidin 2D crystals in the presence of free streptavidin in
the bulk solution, small point defects such as mono- and di-vacancy defects would not easily access to the free streptavidin molecules (Fig. 61a), and hence, have a tendency toremain in the crystals. However, the fusion of small point defects into a larger point defectfacilitates its access to the free streptavidin molecules (Fig. 61b) and thereby promotes theremoval of small point defects from the crystals. Interestingly, the defect mobility increases
with increasing defect size [168]. The higher mobility of larger point defects increases theirprobability to encounter other point defects to form yet larger point defects with further
higher mobilities. During the crystal growth in the presence of free streptavidin in the bulk
solution, this acceleration eﬀect also promotes the removal of point defects from the crystalline regions.
9.2.4. Intrinsically disordered regions of proteins
In addition to biological processes, high-speed AFM imaging is also useful for identifying structurally ﬂexible regions of proteins, particularly intrinsically disordered (ID)regions of proteins.

Fig. 60. Fusion of two monovacancy point defects into a divacancy point defect in streptavidin C222 crystal.
Imaging rate, 0.5 s/frame.

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Fig. 61. Filling vacancy defects in streptavidin 2D crystal with free streptavidin molecules in the bulk solution.
Small point defects (a) such as mono- and di-vacancy defects would not easily access to the free streptavidin
molecules, while larger point defects (b) easily access to them and hence are removed from the crystal.

Biomolecular binding speciﬁcity has been described for a quite while by the famous‘‘lock and key” mechanism, posing that a protein must be folded to give a high degreeof geometrical precision in molecular binding. This concept is recently being overturned
by a surprising yet credible ﬁnding that many proteins in the cell appear to be unfolded
most of the time. Unstructured segments mostly comprise ﬂexible linkers that play important roles in the assembly of macromolecular complexes and in the recognition of smaller
biomolecules. The functional importance of ID regions has recently been recognized, particularly in transcription, translation, and cellular signal transduction [171,172]. However,there are no useful techniques for analyzing unstructured segments of this sort at the single-molecule level. Indeed, X-ray crystallography and electron microscopy do not allow us
to directly observe the ID regions. Conventional slow AFM also cannot reveal such a thinand ﬂexible entity. Under ambient conditions, thin and ﬂexible unstructured polypeptides
tend to form lumps, or become ﬂattened due to their strong attachment to a surface. Suchﬂattened polypeptides are diﬃcult to visualize by AFM. In addition, they are not easily
adsorbed onto substrate surfaces in solution, even with the assistance of adhesive chemicals. Recently, we observed the facilitates chromatin transcription (FACT) protein in solution by high-speed AFM [173]. FACT is a heterodimer and displaces histone H2A/H2B
dimers from nucleosomes, thereby facilitating RNA polymerase II transcription[174,175] and chromatin remodeling [176]. The image of oligomerized FACT clearly
revealed undulating tail-like structures protruding from the main body of FACT
(Fig. 62) [173]. The image of a FACT monomer indicated that FACT contains two taillike structures of diﬀerent lengths [173]. This ﬁnding coincided with the ID regions predicted from the amino acid sequences. In fact, the AFM images of deletion mutants lacking either of the two predicted ID regions showed either of the shorter and longer tail-likestructures. Their macroscopic contour lengths of the shorter and longer ID regions showedwide distributions and were on average 17.8 nm and 26.2 nm, respectively.
We analyzed mechanical properties (stiﬀness) of the observed tail-like structures. The
stiﬀness of a polymer chain is described by the persistence length p of the chain, i.e., thelength over which the average correlation in the tangent direction decays by 1/e. In
two-dimensions, the mean square point-to-point distance of the chain is given by [177]T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

405

Fig. 62. Successive AFM images of oligomerized FACT captured at an imaging rate of 6 frames/s. The scan
range was 150 nm  150 nm. A lumpy shape and four tail-like structures can be seen (see cartoon inset on (a)).
The position of the tail-like structures ﬂuctuated markedly over time. The images are processed using brightnessequalizing software [173].


2phr ðlÞi2D ¼ 4pl 1  ð1  el=2p Þ ;
l
2

ð46Þ

where l is the contour length between two points on the chain. Here, note that the persistence length depends on the hierarchy of the concerned polymer structures. For example,
the persistence length of a double helical DNA strand is diﬀerent from those of thicker
strings formed by super-coiling of a DNA strand. In the AFM images obtained for the
tail-like structures of FACT, polypeptide chains are not resolved. Therefore, the persistence lengths that can be obtained from the images are those for overall (macroscopic)
string structures (see inset in Fig. 63). We analyzed the tail-like structures of the deletionmutants. The mean square point-to-point distance r2 as a function of l was well ﬁtted by
Eq. (46) for both cases, giving persistence lengths of 10.5 nm and 12.1 nm for the shorterand longer tail-likes structures, respectively (Fig. 63). These similar values indicate that thetwo ID regions have physically similar structures.
The Young modulus E is a material property that represents its stiﬀness and is related
to the macroscopic persistence length as E = pkBT/I [178], where I is the momentum ofinertia of the cross-section of the object. Assuming that the tail-like structures of FACT
are thin rods with a radius q, I is given by I = pq4/4. We estimated q (1–1.6 nm) fromthe width of the tail-like structures on several AFM images obtained using diﬀerent cantilever tips, taking into account the tip radiuses. The width is less aﬀected than the heightby the substrate surface and its estimation is reliable when the sample height is similar toor smaller than the tip radius. Thus, the Young modulus of the two tail-like structures wasestimated to be 9–58 MPa from their similar macroscopic persistence lengths. This value

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Fig. 63. Persistence length determination of the macroscopic structures of the tail segments of (a) the deletionmutant with a shorter tail and (b) the deletion mutant with a longer tail. The square point-to-point distance isplotted as a function of the macroscopic contour length between the corresponding two points. The red lines
indicate the best-ﬁt curves of Eq. (46) with (a) p = 10.5 nm and (b) p = 12.1 nm. The inset image indicates therelationship between the microscopic contour and the macroscopic structure. The microscopic contour isschematically drawn with a thin yellow line [173]. (For interpretation of the references to colour in this ﬁgurelegend, the reader is referred to the web version of this article.)estimated as a macroscopic stiﬀness of the ID regions is 1–2 orders of magnitude smaller
than those reported for globular proteins such as an actin monomer (2 GPa) [179] andlysozyme (0.5–2 GPa) [180,181]. The value for an actin monomer is estimated by theforce–extension relationship measured using single actin ﬁlaments. The value of 0.5 GPa
for lysozyme is estimated by AFM force-indentation experiments using a tip with a radiusof 20 nm [180] while the value of 2 GPa for lysozyme is estimated by measuring the speedof ultrasound propagation through a lysozyme solution [181]. Therefore, these values represent the macroscopic stiﬀness of these proteins. Interestingly, Young’s modulus esti-

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mated as macroscopic stiﬀness of bovine carbonate anhydrase II is reported to be in arange of 2–6 MPa when the sample is incubated in the presence of 2–6 M guanidine
HCl [182]. Although these values of Young’s modulus are somewhat smaller than thevalue for the ID regions of FACT, the structures of the ID regions are considered to beroughly similar to those of denatured proteins.
The unique high-speed AFM capability to detect unstructured regions of proteins is
also useful for successful protein crystallization, since the predicted ID regions can be conﬁrmed and deleted before crystallization to facilitate crystal growth.
9.3. Imaging using commercially available high-speed AFM
The full scale commercialization of high-speed AFM has not been achieved yet. To thebest our knowledge, two manufacturers have produced prototype instruments. Inﬁnitesima (Oxford, UK) has recently produced VideoAFMTM equipped with a fast x-scanner
made of a quartz tuning fork, based on the studies by Miles’ group [183,184]. A fast feedback system is not implemented in this prototype. Although its marketing is restricted toonly Japan at present, Olympus (Tokyo, Japan) has produced Nano Live VisionTM, which
is a modiﬁed version of the microscope previously developed by us [54]. One of the users of
the latter product has recently ﬁlmed the dynamic processes of GroEL–GroES interaction,
which was regulated by the ATPase reaction of GroEL immobilized onto a mica surface inan end-up orientation [185]. Moreover, the group observed the 1D diﬀusion of a restriction enzyme along a DNA strand, which was followed by the cleavage reaction [186]. A
more recent study successfully captured the translocation and looping of DNA by a
type-III restriction enzyme, EcoP15I [187]. Another group observed the formation anddissociation of a streptavidin–biotinylated DNA complex [188].
10. Substrata for observing dynamic biomolecular processes
To visualize dynamic biomolecular processes, the substratum surfaces on which a sample is placed should meet various requirements; for example, (a) the surface must have anappropriate binding aﬃnity for the sample so that the sample does not move too fast on it,(b) the functional activity of the sample should be retained on the surface, (c) the surface
should selectively attach a speciﬁc component in a multicomponent sample, and (d) the
sample should attach to the surface in a desired orientation. Mica (natural muscovite orsynthetic ﬂuorophlogopite) has frequently been used as a source substratum owing to
its surface ﬂatness at the atomic level over a large area. It has net negative charge andis therefore quite hydrophilic. A bare mica surface adsorbs various proteins by electrostatic interactions. Except in some cases (such as GroEL attachment in an end-up orientation), the orientation of adsorbed proteins is not unique, and the selective attachment ofa speciﬁc species is not expected. When the dynamic processes of a single species of protein
are to be observed without other proteins, a mica surface is useful. We can control theaﬃnity for a speciﬁc protein by varying the ionic strength or pH, or by adding divalentcations such as Mg2+. However, interesting biomolecular processes often occur in multicomponent systems.
It is desired to have, in the ﬁrst step, a surface that does not initially attach to the sample. Then, this surface is modiﬁed so that only a speciﬁc species of molecules can attach toit through a linker in a desired orientation. A membrane surface with zwitterionic polar

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head groups such as phosphatidyl choline (PC) and phosphatidyl ethanolamine (PE) is
known to resist protein adsorption [189,190]. Streptavidin is also useful for the speciﬁc
attachment of a biotinylated protein since it also resists nonspeciﬁc protein adsorption.
Mica-surface-supported planar lipid bilayers [191] can be easily formed from liposomes
as their structure is disrupted by the strong hydrophilic interaction with the mica surface[192]. Various lipids with functional groups that are attached to polar groups (e.g., biotin
attached to PE, Ni-NTA attached to phosphatidylserine (PS)) are commercially available.
They enable the speciﬁc attachment of proteins labeled with biotin or his-tags onto planar
lipid bilayers. Dioleoyl-phosphatidyl-choline (DOPC) is useful for the preparation of 2D
streptavidin crystals when it is used together with biotinylated lipids [193,194]. It contains
an unsaturated hydrocarbon in each of the two alkyl chains, which causes the bending of
the chains and therefore weakens the interaction between neighboring DOPCs. This weakinteraction lowers the phase-transition temperature of DOPC lipid to 20 °C, thereby
aﬀording considerable ﬂuidity to the planar bilayer at room temperature and thus facili-Fig. 64. Streptavidin on mica-supported planar lipid bilayers. (a) 2D streptavidin crystals formed on bilayers of
DOPC + DOPE-biotin. Scale bar, 30 nm. (b) Streptavidin on bilayers of DPPC + DPPE-biotin. Scan range,
200 nm: imaging rate, 0.18 s/frame. (c) Streptavidin on bilayers of DPPC + DOPE-biotin. Scan range, 200 nm;
imaging rate, 0.18 s/frame. The number attached to each image in (b) and (c) indicates the time (s) elapsed after
capturing the ﬁrst frame. The apparent diﬀerence between (b) and (c) in the size of streptavidin molecules are dueto the diﬀerence in the sharpness of the cantilever tips used [3].

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tating the 2D crystal formation of streptavidin (Fig. 64a). The densely packed streptavidin
does not diﬀuse easily. If less diﬀusibility is necessary, the packed streptavidin can be crosslinked using glutaraldehyde, which does not aﬀect its ability to bind to biotin. We used thissurface for the selective attachment of biotinylated GroEL in a side-on orientation.
Dipalmitoyl-phosphatidyl-choline (DPPC) contains no unsaturated hydrocarbons in thealkyl chains; therefore, its phase transition temperature is high (41 °C) and it is suitablefor preparing planar lipid bilayers with low ﬂuidity. For example, when planar bilayers areformed using DPPC at a high temperature (60 °C) together with a suitable fraction of
DPPE-biotin, streptavidin, which is sparsely attached to the surface, hardly diﬀuses atroom temperature (Fig. 64b). When DOPE-biotin is used together with DPPC, the sparsely attached streptavidin diﬀuses at a moderate rate (Fig. 64c) [3].
Some degree of protein adsorption onto a planar lipid bilayer surface is sometimes necessary to observe dynamic interactions between an anchored protein and a ﬂoating counterpart protein. When the counterpart protein contains a ﬂexible structure, it exhibits very
rapid Brownian motion even when the protein is in contact with an anchored protein. As
such a rapidly moving structure cannot be imaged even by high-speed AFM, it is necessary
to ﬁnd appropriate lipid polar groups to slow the motion. Although the relationships
between the properties of protein adsorption onto planar bilayer surfaces and the lipid
composition have been studied [195–197], we need more extensive studies on this issue.
11. High-speed phase-contrast imaging
11.1. Phase mapping
11.1.1. Compositional mapping on blended polymers
To demonstrate that high-speed phase-contrast imaging is possible using small cantilevers and the fast phase detector described in Chapter 7, we ﬁrst imaged poly(styrene–butadiene–styrene) (SBS) block copolymers as a test sample, as SBS ﬁlms are often used forphase imaging with tapping-mode AFM [114,198]. The SBS polymer was dissolved in toluene and then cast on a freshly cleaved mica surface, which was followed by drying underambient conditions. Prior to the observation, the sample was heated at 70 °C for 1 h. The
small cantilever used had a resonant frequency of 658 kHz and a quality factor of 2 inwater, and a spring constant of 0.2 N/m.
It is known that several types of phase separation patterns are formed using SBS ﬁlms
depending on the evaporation speed of the solvent [199]. Fig. 65 shows typical (a) topographic and (b) phase images obtained simultaneously at a rate of 84 ms/frame in distilledwater. Here, the phase detection timing was set at a regime in which the tip was
approached the surface. Clear phase-contrast images were obtained even at such a highimaging rate. In the phase-contrast images shown in Fig. 65b and d, darker regions correspond to an advanced phase with an average shift of approximately +6° relative tothe phase at brighter regions. Since the image contrast in the error signal is faint, thephase-contrast predominantly reﬂects the compositional heterogeneity of the SBS ﬁlm.
To identify the compositions of diﬀerent region in the topographic image, the peak-topeak set point amplitude was reduced from 11 nm to 7 nm while imaging. Fig. 65c andd shows topographic and phase-contrast images obtained with the reduced set point
amplitude, respectively. The higher topographic region shown in Fig. 65a was dented by
the increased loading force, as shown in Fig. 65c. The cross-section height proﬁles shown

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Fig. 65. (a, c) Topographic and (b, d) phase-contrast images of a SBS ﬁlm captured at 84 ms/frame in pure water.
The scan area is 200  200 nm2 with 100  100 pixels. The amplitude set point r is 0.85 (a, b) and the peak-topeak free amplitude is 13 nm. The amplitude set point is reduced to 0.54 (c, d). Solid and broken lines correspondto the cross-sectional proﬁles (e, f) obtained before and after reducing the amplitude set point, respectively.
Broken lines in (a)–(d) indicate the positions at which the cross-sections are obtained [119].

in Fig. 65e reveal that it was dented by more than 5 nm. Thus, the compositions of thehigher and lower regions in Fig. 65a are identiﬁed to be the less stiﬀ poly-butadiene(PB) and the stiﬀ poly-styrene (PS) domains, respectively. The darker (advanced phase)area in the phase-contrast image shown in Fig. 65d is wider than that in Fig. 65b. Thisis because the contact area between the tip and the surface was increased by the increased
loading force. However, the magnitude of the phase shift is independent of the loading
force, as can be seen in the cross-section phase proﬁles shown in Fig. 65f. Therefore,
the observed phase-contrast is caused by only compositional heterogeneities and is independent of the surface topography.
Fig. 66 shows (a) topographic and (b) phase-contrast images obtained at 896 ms/frameon a diﬀerent SBS ﬁlm. The conﬁguration of this sample is very diﬀerent from that shownin Fig. 65. This large diﬀerence was caused by a diﬀerence in the solvent evaporation condition. In this case, the topographic and the phase-contrast images are not consistent witheach other. The compositional heterogeneity revealed in the phase-contrast image is notclearly seen in the topographic image.
Thus, it was clearly demonstrated here that the combination of small cantilevers andthe fast phase detector enables highly sensitive and rapid phase detection of the cantileverFig. 66. (a) Topographic and (b) phase-contrast images of a SBS ﬁlm captured at 896 ms/frame in pure water.
The scan area is 350  350 nm2 with 200  200 pixels.

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oscillation. This capability allows us to simultaneously observe the surface topography
and the surface material heterogeneity map at speed greater than 10 frames/s.
11.1.2. Dependence of phase-contrast on detection timing
To determine conditions for the optimum phase-contrast as well as to obtain an insight
into the phase-shift mechanism, we investigated the dependence of the phase-contrast on
the phase detection timing within a cantilever oscillation period [119]. Fig. 67a shows an
oscillation signal of a cantilever tip that is tapping an SBS ﬁlm in water. The cantilever
oscillation ‘‘0 nm” corresponds to the situation that the tip is touching the SBS surface.
The oscillation signal is slightly distorted from the sinusoidal wave due to the inductionof nonharmonic components attributed to the cantilever’s low quality factor. Fig. 67b–d
shows typical phase images obtained at various detection timings. At the phase detection
trigger points in the region indicated by (i) in Fig. 67a, we observe only a faint phase-contrast as shown in Fig. 67b. Since the image contrast of the error signal is similar to thisphase-contrast, this faint phase-contrast is mainly due to the ﬂuctuations in amplitude.
On the other hand, the phase-contrast image obtained at the trigger points in region (ii)shows a relative phase advance of approximately +5° in the PB regions as shown in
Fig. 67c. Then, at the trigger points in region (iii), we can clearly observe the phase-contrast with a relative phase advance of approximately +10° in the PB region as shown in
Fig. 67d. The phase-contrast image obtained at the trigger points in region (iv) shows asimilar contrast to that shown in Fig. 67c. Thus, the magnitude of the phase shift strongly
depends on the timing of the trigger point; namely, it depends on the movement directionof the oscillating tip. The largest phase shift is observed when the tip approaches the surface. However, when the tip withdraws from the surface, a phase shift barely occurs. At
the positions where the tip velocity is almost zero, the phase shifts only slightly. We used

Fig. 67. Phase-contrast images of a PS–PB block co-polymer ﬁlm depending on the phase detection timing. (a)Cantilever oscillation signal and various trigger regions for phase detection. The amplitude set point r is 0.63 andthe peak-to-to-peak free-oscillation amplitude is 8.7 nm. The oscillation signal of ‘‘0 nm” corresponds to the
distance where the tip is in contact with the surface. The phase-contrast images shown in (b), (c), and (d) are
obtained at trigger points within regions (i), (ii), and (iii), respectively. The imaging rate is 1.2 s/frame, and thescan area is 350  350 nm2 with 200  200 pixels. (e) Phase image obtained in air. Broken circles indicate thepoly-butadiene regions. The imaging rate and scan area are 8 s/frame and 400  400 nm2 with 256  256 pixels,
respectively [119].

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three diﬀerent cantilevers with resonant frequencies of about 600 kHz, 800 kHz, and
1.2 MHz. These showed a similar trigger-timing dependence. In addition, the same tendency was observed regardless of the imaging rate, free oscillating amplitude, and set point
amplitude. Surprisingly, the phase-contrast images observed under the ambient conditions
revealed a diﬀerent contrast (Fig. 67e). The phase was slightly delayed in the PB regions
encircled by dotted lines; identiﬁcation of these PB regions was performed using the same
indentation method as that employed to obtain the image shown in Fig. 65c. Thus, themechanism of the phase shift signiﬁcantly diﬀers between air and liquid-environments.
In addition, the distortion of the sinusoidal waveform in air is smaller than that in water,
and the strong dependence of the phase-contrast on the trigger-timing is not observed.
This is because the quality factor is relatively high in air.
Fig. 68a and b shows the phase lag and oscillation amplitude, respectively, recorded inwater with diﬀerent trigger-timings as functions of the relative tip–surface distance. In the
PS region, the oscillation amplitude decreased more signiﬁcantly than in the PB region, as
the PB region is much softer than the PS region. The phase shows a delay for both the PS
and PB regions. The phase delay in the PS region measured at a trigger point in region (iii)(see Fig. 67a) is larger than that in the PB region, as shown in Fig. 68a. This large diﬀerence results in a clear contrast in the phase image, as exempliﬁed by Fig. 67d. On the other
hand, the phase delay measured at a trigger point in region (i) does not diﬀer signiﬁcantly
between the PS and PB regions, as shown in Fig. 68b. The vertical lines in Fig. 68a and b

Fig. 68. Phase lags and cantilever oscillation amplitudes as a function of the relative tip–surface distancemeasured on the trigger points in (a) region (iii) and in (b) region (i) shown in Fig. 65a. Solid and dotted curves
are obtained on PS and PB rich domains, respectively. (c) The phase lags measured on the PS (d) and PB (N)regions are plotted as a function of the trigger points. The phase values are measured at the tip–surface distanceindicated by broken lines in (a) and (b) [119].

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indicate the tip–sample distances where similar amplitude reductions occur in trigger
regions (i) and (iii). As shown in Fig. 68c, at these tip–sample distances (which are similar),the phase delays in the PS and PB regions change with time after the tip–sample contact.
The phase delay in the PS regions increases with time, while that in the PB region remained
almost constant. An interpretation of this diﬀerent behavior is given later.
Phase-contrast images cannot be interpreted straightforwardly because the phase isaﬀected by a number of instrumental and non-instrumental parameters [6,7,114–
117,199–202]. These studies on phase-contrast mechanisms have been performed only
under ambient or vacuum conditions. Instrumental parameters such as the free-oscillation
amplitude and the set point amplitude aﬀect the phase image contrast in a complex manner. It has been reported that a ﬂipped phase-contrast occurs when the tapping force levelis being changed [114,115,203]. Such contrast ﬂips generally occur because of a change inthe tip–sample interaction between the net attractive- and repulsive-force regimes[204,205]. However, we did not observe these phase-contrast ﬂips. In addition, the amplitude–distance and phase–distance curves (Fig. 68a and b) did not show the features of atransition from the ‘‘low-amplitude state” to the ‘‘high-amplitude state”, the transition
that takes place upon moving from the attractive regime to the repulsive regime [204].
Therefore, the tip–sample interaction is mostly repulsive (e.g., high-oscillation amplitudestate) in both the PS and PB regions under the present imaging conditions. Under the
repulsive-force conditions, the phase-contrast of a PS/PB blend ﬁlm observed under ambient conditions mainly reﬂects its mechanical properties [201,205]. In this case, the phasesignal is more delayed in the softer PB region than in the harder PS region. This can be
explained by the fact that the larger repulsive-force on the harder material induces a morepositive resonant frequency shift, but the damping constants on both materials are similar[201,205]. However, in water, the phase-contrast is reversed. Hence, the phase image contrast in water appears to be dominated by other properties of PS and PB than theirmechanical properties and is very likely to be aﬀected by the adhesion force between
the tip and the sample [202,206]. Both PB and PS are nonpolar materials but PB is relatively more polar than PS because the thermal treatment of PB results in the formation ofsome C@O groups or other polar groups such as OH [201]. Because the AFM tip used hereis made of hydrophobic amorphous carbon, the adhesion force derived from the hydrophobic interaction should be larger in the PS region than in the PB region. This diﬀerence
produces a larger phase delay in the PS region. In air, large capillary forces mask suchchemical interactions [202,206], but the mechanical properties are emphasized by the larger quality factor in air.
Here, we propose a possible interpretation of the trigger-timing dependence of the PB/
PS phase-contrast observed in water. A larger repulsive-force generated in the harder PS
region produces a greater phase advance after the tip comes in contact with the surface,because a larger positive frequency shift relative to the ﬁxed driving frequency occurs.
The water medium damps the cantilever oscillation, and hence, the phase advancedecreases within one oscillating cycle due to the low Q factor of the cantilever [207]. Therefore, the phase advance produced immediately after the tip–surface contact becomes smaller with time. On the other hand, the larger phase delay in the PS region caused by theadhesive force does not decay quickly within a single oscillation cycle, because of theenergy-dissipative interaction. As the adhesive interaction eﬀect is larger than the repulsive-force eﬀect, the net phase shift is a delay rather than an advance. Consequently, when
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Fig. 68c. For the softer and less adhesive PB regions, the phase advance due to the repulsive-force is smaller and the phase delay due to the adhesive interaction is also smaller.
However, the net phase delay in trigger region (i) is similar to the corresponding phasedelay observed in the PS regions. Since the contribution of the phase advance to the net
phase delay is small in the PB regions, the gross phase is almost maintained, as shownin Fig. 68c.
The insights given above lead to the following important conclusions. In phase-contrast
imaging in liquids, the optimization of the trigger point is essential for obtaining a goodsignal-to-noise ratio. Moreover, by selecting the trigger point or by simultaneously obtaining phase-contrast images at multiple trigger points, it is possible to separately image themechanical and chemical properties of the sample.
11.1.3. Phase imaging of myosin ﬁlament
To demonstrate the performance of the simultaneous topographic and phase-contrast
imaging of biological systems, here we present images obtained using myosin ﬁlaments
in a buﬀer solution. Skeletal muscle myosin ﬁlaments were obtained by diluting the monomer solution with a low-salt buﬀer [208]. The cantilever peak-to-peak free-oscillation
amplitude was about 12 nm and the amplitude set point was about 0.7. Fig. 69a and bshows typical topographic and phase-contrast images, respectively, obtained at an imaging
rate of 4 s/frame. The scan area was 2 lm  2 lm, which was relatively large for highspeed imaging and therefore the imaging was performed at relatively slow speed. The surfaces of the myosin ﬁlaments appear smooth in the topographic image. On the other hand,the phase-contrast image shows the structures of the ﬁlaments in more detail. Here, thetrigger-timing was tuned so that the maximum phase-contrast was obtained. Note thatthe phase on the myosin ﬁlaments is delayed relative to that on the mica surface. This
probably indicates diﬀerences in the mechanical properties between the biological sample
and the mica; the myosin ﬁlaments have higher elasticity and/or viscoelasticity than mica.
The phase-contrast on the myosin ﬁlaments indicates the non-uniformity of the mechanical properties.
11.2. Phase-modulation imaging
11.2.1. Distance dependence of phase-shift signal
Recently, it has been demonstrated that FM-mode AFM has atomic-resolution imaging capability even in liquids (the details are described in Chapter 12) [121–123,132]. In

Fig. 69. (a) Topographic and (b) phase-contrast images of myosin ﬁlaments captured at 4 s/frame in buﬀer
solution. The scan area is 2  2 lm2 with 256  256 pixels.

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FM mode, a cantilever is excited at its resonant frequency in a self-oscillation mechanism,
which increases its eﬀective Q-value. The increased Q-value slows the cantilever amplitude response. In addition, the bandwidth of the conventional phase-locked loop
(PLL) that is used to detect the frequency shift is usually limited to a few tens of kHz.
Thus, it appears diﬃcult to increase the imaging rate of FM–AFM. On the other hand,as described in Section 11.1, the phase of the cantilever oscillation excited at a ﬁxed frequency at near its resonant frequency is also sensitive to the tip–sample distance as wellas to the surface material properties of the sample [209]. When the material properties are
not signiﬁcantly inhomogeneous on the sample surface, the phase signal will vary withdependence almost solely on the tip–surface distance. As mentioned in Section 11.1, asmall Q-value can be used for the phase detection. Thus, phase-modulation mode
AFM (PM–AFM) appears promising for high-speed low-force imaging. In the simple
model of harmonic oscillation, the phase shift D/ induced by the elastic interaction isapproximately expressed by D/  Qk=k c , where k is the gradient of the interaction
force. Our small cantilever used in high-speed AFM has a Q-value of 2.5 in water
but the spring constant k c is much smaller than that used in FM–AFM. Therefore, phaseshifts that are detectable using our fast phase detector are produced by the tip–sample
interaction.
Fig. 70 shows the amplitude–distance and phase-shift–distance curves observed on amica surface in a buﬀer solution. The phase is advanced (i.e., the resonant frequency
is increased) due to the repulsive interaction and its sensitivity is 28 mV/°. The amplitude
signal starts to decrease slowly at the distance marked with ‘A’ in Fig. 70. From distance‘B’, it decreases more quickly. For imaging in the AM mode, the set point amplitude
should be set less than that at the distance B to obtain suﬃcient image quality. The slopeof the amplitude–distance curve at distance B is approximately 60 mV/nm. Meanwhile,
the phase-shift signal decreases almost linearly from the distance A with an average slopeof 280 mV/nm. Because of this higher sensitivity of the phase signal, the PM mode (thephase signal is used for the feedback control of the tip–sample distance) enables lowinvasive imaging on the sample as well as at higher spatial resolution than the AM
mode.

Fig. 70. Amplitude-distance (dotted line) and phase shift-distance (solid line) curves measured on a mica surface
in a buﬀer solution.

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Fig. 71. Successive AFM images of GroEL taken by (a)–(d) the AM mode and (e)–(h) by the PM mode. Scan
area is 400  400 nm2. Imaging rate is 800 ms/frame.

11.2.2. PM imaging of GroEL
Chaperonin GroEL was used as a test sample for the PM imaging, as its stacked ring
structure is fragile and is often bisected by strong tip forces [210]. Fig. 71a–d shows successive topographic images taken in the AM mode at 800 ms/frame. The set point amplitude was approximately 90% of the peak-to-peak free amplitude of 7.5 nm. In theseimages, a large fraction of GroEL molecules are bisected and are sometimes dislodged
by the scanning tip. On the other hand, in the PM mode, the bisection of GroEL rings
barely occurred as shown in Fig. 71e–h. In addition, the hole centered on the ring structure
is more clearly observed in the PM mode than in the AM mode, and the subunits areresolved in the PM mode (Fig. 72).

Fig. 72. High-resolution image of GroEL taken in the PM mode. Scale bar, 10 nm; imaging rate, 480 ms/frame.

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12. Atomic-resolution imaging in liquids
12.1. Atomic-resolution imaging by FM–AFM in liquid
Since the ﬁrst demonstration of true atomic-resolution by FM–AFM in 1995 [10], anumber of atomic-resolution FM–AFM images have been obtained on various surfaces
in UHV. However, the high-resolution imaging capability of FM–AFM had been limited
to UHV until recently. In 2005, Fukuma et al. presented a method of overcoming this limitation using an ultralow noise cantilever deﬂection sensor, stiﬀ cantilevers, and a small
oscillation amplitude [76]. In this section, we present the high-resolution FM–AFM
images that demonstrate the signiﬁcant progress in improving the spatial resolution in
dynamic-mode AFM for liquid-environment.
The ﬁrst true subnanometer-resolution imaging by FM–AFM in liquid was demonstrated by imaging a single crystal of polydiacetylene [poly-PTS; 2,4-hexadine-1,6-diolbis(p-toluene sulfonate)] in water, as shown in Fig. 73a and b [121]. Fig. 73a shows stripes
with a spacing of approximately 0.75 nm, which correspond to the spacing of the polyacetylene main chain. The dots along each stripe represent individual PTS side chains. The
image also shows molecular-scale defects. Fig. 73b reveals two protrusions corresponding
to the two adjacent carbon atoms in a PTS side chain. These images demonstrate the true
subnanometer-resolution (0.25 nm) of FM–AFM in liquid.

Fig. 73. FM–AFM images of (a), (b) the bc plane of a poly-PTS single crystal (fc = 136 kHz, k = 42 N/m) and(c), (d) muscovite mica surface taken in water (fc = 140 kHz, k = 42 N/m). (a) 30 nm  30 nm, A = 0.26 nm,
Dfc = +167 Hz, scanning speed: 839 nm/s. (b) 6 nm  4 nm, A = 0.20 nm, Dfc = +290 Hz, scanning speed:
448 nm/s. (c) 8 nm  8 nm, Dfc = +54 Hz, A = 0.24 nm, scanning speed: 671 nm/s; (c) 4 nm  2.5 nm,
Dfc = +157 Hz, A = 0.16 nm, scanning speed: 934 nm/s. The images were taken in constant height mode [121].

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Soon after the imaging of poly-PTS, the ﬁrst true atomic-resolution imaging by FM–
AFM in liquid was reported by imaging a muscovite mica surface in water, as shown in
Fig. 73c and d [132]. Fig. 73c shows a honeycomb-like pattern, which is characteristicof a cleaved mica surface. In the more magniﬁed image shown in Fig. 73d, bright spots
are resolved along the honeycomb pattern. These bright spots have been tentatively attributed to Al3+ ions in the aluminosilicate honeycomb mesh. The direct imaging of individual
ions irregularly aligned on the surface demonstrates that the true atomic-resolution can beobtained by FM–AFM in liquid.
12.2. Biological applications of FM–AFM
One of the most attractive applications of liquid-environment FM–AFM is the highresolution imaging of biological systems under physiological environments. Since the spatial resolution of FM–AFM in liquid is as small as the size of a single water molecule or asolvated ion, FM–AFM should make it possible to visualize the atomic-scale interaction
potential at the interface between biological systems and the physiological environment(i.e., water molecules and ions). In this section, we describe examples of such applications
reported in 2007 by directly probing the atomic-scale interactions between a model biological membrane and physiological solution [64,122].
12.2.1. Direct imaging of intrinsic hydration layers
Water molecules adjacent to biological membranes could potentially play a key role inimportant biological processes, particularly, when they form hydration layers. The hydration layers alter the local interaction potential at the water/lipid interface, which signiﬁcantly aﬀects the function of membrane proteins and molecular transport across themembranes. FM–AFM has a unique capability of measuring local interaction potential
with a nanometer-scale cross-section using a sharp tip as a probe. This capability is ideal

Fig. 74. (a) Example of Dfc vs. distance curve measured on a DPPC bilayer in phosphate buﬀer solution, showing
an oscillatory proﬁle with two peaks. The smoothed line (solid) is obtained by averaging the raw data (shaded)over the distance range of ±0.02 nm from each data point. (b) Force vs. distance curve converted from (a) using
the formula reported in Sader and Jarvis [211]. The tip velocity during the force curve measurements was 0.5 nm/
s. The curves shown here were measured during tip approach. The curves taken during tip retraction show
approximately the same proﬁle but were omitted from the ﬁgure for clarity [64].

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for directly probing the interaction that nanoscale objects (e.g., proteins and solvated ions)experience when they approach a membrane surface.
Fig. 74a shows an example of such applications: a Df vs. distance curve measured on amica supported DPPC lipid bilayer in phosphate buﬀer saline (PBS) solution [64]. The raw
data (gray line) was smoothed (black line) and was converted to a force vs. distance curve(Fig. 74b) using the equation reported by Sader and Jarvis [211]. The force curves obtainedon the DPPC bilayer typically show an oscillatory force proﬁle with one or two peaks. The
average separation of the two peaks is 0.28 ± 0.05 nm. This distance agrees with the size ofa water molecule, suggesting that the oscillatory force proﬁle corresponds to the sequential
removal of ordered water molecules.
The tip–sample distance is regulated on the basis of the assumption that Df increases
with decreasing tip–sample separation such as in the force branches indicated by arrows(i)–(iii) in Fig. 74a. Thus, the oscillatory proﬁle of the Df vs. distance curve implies existence of multiple feedback positions for a single set point value as indicated by the circles
in Fig. 74a. Namely, the tip can spontaneously jump between these positions even if the set
point is unchanged.
Fig. 75 shows examples of such spontaneous jumps during FM–AFM imaging of the
DPPC bilayer in PBS solution [64]. In the image shown in Fig. 75a, the tip is scanned fromthe lowest terrace (Terrace 1) and spontaneously jumps twice. The height of the spontaneous jumps is 0.2–0.3 nm, which agrees with the size of a single water molecule. The image
shows the primary and secondary hydration layers with molecular-scale corrugations corresponding to the headgroup positions as well as the headgroups themselves. Such stable
imaging of the hydration layers demonstrates the excellent force sensitivity and spatial resolution of FM–AFM in liquid. It also proves that the hydration layers next to the DPPC
membrane are suﬃciently stable to allow such imaging.
12.2.2. Direct imaging of lipid-ion networks
Under physiological conditions, biological membranes are surrounded by an electrolytic solution containing various metal cations. So far, spectroscopy experiments haverevealed that metal cations speciﬁcally interact with negatively charged moieties of thelipid headgroups [212–215]. These experiments, together with theoretical simulations[216], have led to an idea that individual ions may interact with multiple headgroups toform complex ‘‘lipid-ion networks”. This idea has been used to account for the observed
eﬀects of the ions such as the enhanced mechanical strength of membranes [217] and the
reduced mobility of the lipid molecules therein [218]. However, it has been diﬃcult toexperimentally access such lipid-ion networks due to the lack of a method able to investigate local lipid-ion interactions with Ångstrom resolution. Since FM–AFM allows us todirectly observe the water/lipid interface with Ångstrom resolution, it is an ideal tool
for such biological applications.
The DPPC membrane surface imaged by FM–AFM exhibits various structures depending on both time and location, reﬂecting the mobile nature of the ions interacting with thelipids [122]. FM–AFM images reveal the existence of at least two diﬀerent conﬁgurations
as shown in Fig. 76a and b, which are referred to as Structures 1 and 2, respectively. Structure 1 has pairs of two protrusions separated by 0.30 nm, corresponding to the phosphateand choline groups in the PC headgroups. Structure 2 has hexagonally arranged surface
groups consisting of two oval-shaped subunits with their longer axes parallel to eachother. Detailed analyses of the FM–AFM images and theoretical simulations [122] have420

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Fig. 75. Example of an FM–AFM image of the DPPC bilayer in phosphate buﬀer solution showing spontaneous
jumps during imaging. Image size: 8 nm  8 nm. Tip velocity, 120 nm/s. Imaging speed: 85 s/frame. (b) Line-byline ﬂattened image of (a). Scale bar, 1 nm. Height range: 0.1 nm (black to white). Fast and slow scan directions:left to right and top to bottom. The regions indicated by the numbers 1, 2, and 3 correspond to the Terraces 1, 2,
and 3 in (a). (c) Line-averaged height proﬁle of (b) plotted along the slow scan direction [64].

suggested that the Ångstrom-scale contrasts found in Structure 2 represent the averaged
position of mobile ions interacting with the lipid headgroups, namely, lipid-ion networks.
The two subunits in Structure 2 show enhanced ion occupancies around the two negatively

Fig. 76. FM–AFM images of the DPPC bilayer in PBS solution. Height scale: 0.12 nm. (a) Structure 1. Tip
velocity: 146 nm/s. (b) Structure 2. Tip velocity: 120 nm/s [122].

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Fig. 77. Sequential FM–AFM images of the same area of the DPPC bilayer in PBS solution. Height range:
0.1 nm. Tip velocity: 120 nm/s. Imaging speed: 85 s/image [122].

charged oxygen atoms in the phosphate group, which have been reported to be the primary binding sites for metal cations [213,218,219].
Sequential FM–AFM images obtained from the same area (Fig. 77) reveal that some ofthe surface groups change their conﬁguration upon the formation or disappearance oflipid-ion networks [122]. For example, Subunit S1 does not pair with another subunit inFig. 77a, whereas a pair of Subunits, S3 and S4, appears in the next image (Fig. 77b).
The observed structural changes due to the formation and disappearance of the lipidion networks indicate that the negatively charged phosphate groups share the positive
charge of cations, by which an attractive electrostatic force is exerted on all the headgroups involved in the network. The attractive interaction force mediated through suchcomplex lipid-ion networks should bind the headgroups together and increase the global
mechanical strength of the membrane. In fact, it is known that the addition of metal cations increases the mechanical strength of the DPPC bilayer [217]. The FM–AFM images
shown here reveal the submolecular-scale origin of this eﬀect of the ions on the mechanicalproperties of the biological membrane.
13. Future prospects
The imaging speed of high-speed AFM has reached 10–30 frames/s. Importantly, thescanning and oscillating cantilever tip hardly disturbs the physiological functions of delicate biological samples, provided the cantilever free-oscillation amplitude is suppressed tosome extent. As described in Section 9.2, the current high-speed AFM with this capacity
can be used to ﬁlm the dynamic biological processes of puriﬁed protein systems. The video
images can allow us to gain insights into their functional mechanisms in a much morestraightforward manner than other techniques. However, even the current performanceof high-speed AFM does not fully meet the requirements described in Chapter 4. In thischapter, we discuss whether and how the requirements can be fulﬁlled and what technologies must be developed further to achieve breakthroughs in this new microscopy ﬁeld.
13.1. Imaging speed, interaction force, and noncontact imaging
To expand the scope of biological samples that can be studied by high-speed AFM, theimaging rate must be enhanced further and the tip–sample interaction force must be422

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reduced. Presently, the cantilever is the factor that prevents the achievement of both ofthese improvements. Generally, to improve the resonant frequency, one must compromise
the stiﬀness and vice versa. The most advanced small cantilevers developed by Olympus
deem to have almost achieved the ultimate goal of balancing these two mechanical quantities. Considering their practical use, doubling the resonant frequency to 2.4 MHz inwater seems to be the upper resonant frequency limit, provided the cantilever complianceis not sacriﬁced. The question, therefore, is how can this seemingly unsurpassable level be
exceeded? If we can signiﬁcantly improve the detection sensitivity of the tip–sample interaction, we can use stiﬀer cantilevers and hence attain resonant frequencies that are a few
times higher. As described in Section 6.4.2, measuring the peak force instead of the amplitude will improve the sensitivity. Even when we realize this peak force detection as well asa piezoelectric actuator-based z-scanner with a realistic upper limit of the bandwidth of
1 MHz, the achievable imaging rate is at most 50–60 frames/s for a scan range of240  240 nm2, 100 scan lines and k  10 nm (see Eqs. (1) and (6)). Thus, even if we
continue the present line of development, we will not be amply remunerated. Nevertheless,at present we have to pursue this line while searching for alternatives.
The ultimate minimization of the tip–sample interaction force is attained by noncontact
imaging. Nc-AFM has only been realized in a vacuum environment by utilizing a cantilever with a signiﬁcantly large quality factor in vacuum. If high-speed nc-AFM is realized inliquid conditions, we can use stiﬀer cantilevers with much higher resonant frequencies,
which will promise markedly 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, impossible to study by AFM, since living cell membranes are extremely soft. As many important proteins exist on membranes and are
involved in various physiological functions, the noncontact imaging capability will greatly
increase the usefulness of high-speed AFM in life sciences. For example, extracellular stimuli (such as drugs, endogenous ligands, light or electricity) are ﬁrst received by membranes
proteins (receptors or ionic channels) and are then transmitted to the cell interior to produce various cellular actions. Membrane proteins also determine the eﬃciency of drugdelivery to the interior of cells. Various diseases are linked with membrane proteins, sugar
chains in membrane glycoproteins, and several intracellular molecules involved in signaltransductions. Therefore, high-speed nc-AFM will also make a great contribution to thedetection and diagnoses, and the discovery and evaluation of new drugs.
We recently realized that an ultrasonic technique combined with AFM (Fig. 78) has thepotential of realizing noncontact imaging. In this experimental setup, the sample and cantilever tip are ultrasonically excited at diﬀerent frequencies of f1 and f2, respectively (f1,f2 >> fc) with |f1  f2|  fc. When a sample is attached to a uniform substratum, the wavefront of the ultrasound produced by the sample stage oscillation traces the sample topography. This wavefront can probably be detected by a cantilever tip that is close to but notin contact with the sample surface. We recently examined this possibility and found thatthe tip can sense a surface that it is not in contact with, as shown in Fig. 79. The cantileverstarted to oscillate at its ﬁrst resonant frequency fc when the tip was 2 nm apart from thesurface, and its amplitude increased as it approached the surface. As the tip made contact
with and was pushed onto the surface, the amplitude decreased. This biphasic behavior of
the amplitude signal will not cause problems with the tip–sample distance control if the
DC deﬂection signal is used together with the amplitude signal. Thus, an ultrasonic technique combined with AFM can have the capability of noncontact imaging. However, we

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Fig. 78. Schematic of the proposed high-speed ncAFM setup. Ultrasonic waves are launched from the bottom of
the sample as well as from the cantilever base. The frequencies, f1 and f2, are much higher than the fundamental
resonant frequency fc of the cantilever, and their diﬀerence Df  |f1  f2| is similar to fc. The two ultrasonic waves
interfere with each other and produce acoustic waves with a frequency of Df, which excites the cantilever. The
wavefront of the ultrasonic wave with f1 is formed at the sample surface and sensed by the cantilever that is not in
contact with but is close to the surface.

Fig. 79. Amplitude- and DC deﬂection-distance curves in a buﬀer solution, obtained by the ultrasound technique
combined with AFM. The experimental setup is depicted in Fig. 76. The amplitude signal is for the cantilever
oscillation at its ﬁrst resonant frequency in water (1.2 MHz). The ultrasonic frequencies of f1 and f2 are 5.2 MHz
and 4.0 MHz, respectively. DC-deﬂection signal is obtained by ﬁltering the cantilever deﬂection signal using a
low-pass ﬁlter (cut-oﬀ frequency, 500 kHz).

also became aware of the diﬃculty of maintaining this noncontact detection conditioneven for a few minutes due to the instability of the ultrasonic wave propagation in liquids.
Removing this instability is a key to the realization of high-speed nc-AFM in liquids.

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The noncontact imaging capability in liquids has already been achieved by ion-conductance scanning probe microscopy (ICSPM) [220]. Owing to the progress in fabrication
techniques for producing very sharp glass capillaries with a small pore at the apex, the spatial resolution of ICSPM has reached a few nm [221]. Immobile protein molecules with asize of 14 nm on living cell membranes have been successfully imaged [222]. However, it
seems diﬃcult to increase the imaging rate of ICSPM; the bandwidth of ion-conductancedetection cannot easily be increased, because the ionic current through the small pore of
the capillary electrode is very low.
13.2. High-speed AFM for intracellular imaging
A setup similar to that shown in Fig. 78 has been used for subsurface imaging [68]. Thismethod of microscopy has been called scanning near-ﬁeld ultrasound holography (SNFUH)
and has successfully been used for intracellular imaging under ambient conditions [68]. In its
application, a high-frequency acoustic wave is launched from under the sample stage, andthis wave then propagates through the sample. Materials embedded in the sample with different elastic moduli modulate the phase and amplitude of the propagating acoustic wave.
These modulations aﬀect the nonlinear acoustic interference that occurs at the cantilever
tip and are consequently reﬂected in the cantilever oscillation at its resonant frequency.
SNFUH has no resolution in the z-direction. However, using multiple images obtained with
diﬀerent launching angles of the ultrasonic wave, it is probably possible to reconstitute a 3D
image. Combining SNFUH with high-speed scanning techniques will enable the high-resolution 3D imaging of various intracellular processes in living cells and organisms which takeplace spontaneously or as a result of their responses to extracellular stimuli.
As described in Section 13.1, the ultrasonic technique combined with AFM is capableof noncontact surface imaging. When this method is applied to living cells, the cantilever
oscillation will be aﬀected by the cell surface structures as well as by the intracellular structures. How can we distinguish these eﬀects? The acoustic wave that appears near the sample surface consists of two components. One component is produced by the oscillation of
the whole sample stage, which oscillates the sample surface together. This component is inphase with the ultrasound wave launched from under the sample stage and is used for noncontact surface imaging. We can control its phase relative to the ultrasonic wave propagated through the cantilever to minimize or maximize the acoustic-interferenceproduced cantilever oscillation at fc. The other component is that propagated throughthe sample stage and then the sample, and is used for the imaging of intracellular structures. Its phase relative to the ultrasonic wave propagated through the cantilever can also
be controlled to minimize or maximizing the interference-induced oscillation at fc. Thus,
we can select either cell surface imaging or intracellular imaging by tuning the phasebetween the two ultrasonic waves (one from under the sample stage and the other fromthe cantilever). In an experiment (Fig. 79), we observed that the variation of the phase difference altered the amplitude of the acoustic-interference-induced cantilever oscillation
when the two frequencies (f1 and f2) were appropriately chosen.
13.3. High-speed-recognition AFM
A recognition imaging technique (called ‘‘Trec”) has recently been developed by Hinterdorfer and colleagues [223,224]. A cantilever tip that is attached to a small number of

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probe molecules through a ﬂexible short-linker is oscillated at a frequency below the resonant frequency. Each oscillation is split into approaching and withdrawing parts. The
approaching parts of oscillation signals reﬂect the repulsive tip–sample interaction, providing topographic images. The withdrawing parts reﬂect the association reaction
between the probe molecule and its counterpart on a substratum, providing recognition
images. The eﬀective concentration of the probe molecule attached to the cantilever tip
depends on the tether length. Assuming that only one probe molecule can interact withthe counterpart molecule and a tether length is 2 nm, the concentration is approximately
50 mM, which is suﬃciently high for the association reaction to take place in 20 ls in asystem 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 provided the cantilever
oscillation amplitude is small. As described in Chapters 7 and 11, another method of recognition imaging can be derived from phase-contrast imaging [119] using a cantilever tip
functionalized with a speciﬁc probe molecule. Its phase-contrast image reveals thearrangement of speciﬁc molecules in the sample. As far as the motion freedom required
to the association reaction is provided to the probe molecule, this method can use ashorter tether length (<2 nm) for the probe attachment, and therefore, the eﬀective concentration of one probe molecule can be increased to much higher than 50 mM. Consequently, recognition imaging by phase-contrast detection may be performed much fasterthan by Trec.
13.4. Ultra-high-resolution high-speed AFM
As described in Chapter 12, recent studies using FM–AFM have achieved atomic-resolution imaging in a liquid-environment by minimizing the sensor noise and using smallamplitude cantilever oscillation [76,132,209]. In these studies, conventional cantilevers
with a relatively small quality factor (6) were used. Since a much larger shift in theresonant frequency occurs with small cantilevers, it appears possible to achieve subnanometer-resolution even using high-speed tapping-mode AFM (AM–AFM) equippedwith a fast phase-shift detector (Fig. 43) [119] and a sharp cantilever tip. In our recent
preliminary experiments using AM–AFM, we ﬁlmed atomic-resolution images of a mica
surface in water at a frame rate of 6 frames/s (Fig. 80). Even at an imaging rate of 37
frames/s, atomic-resolution imaging was achieved, although further noise reduction
remains to be accomplished. AFM with both ultra-high-resolution and high-speed capabilities has enormous potential for the imaging of secondary structures on protein surfaces and their dynamic changes, the mapping of rapid changes in the water layer
structures on protein surfaces, the imaging of dynamic lipid–protein interactions onmembranes, and the detection of small-ligand interactions with the active site on a protein surface. It is impossible to study these phenomena using ‘‘conventional” high-speed
AFM.
13.5. High-speed AFM combined with optical microscope
Total internal reﬂection ﬂuorescence microscopy (TIRFM) [225–227] has been used
to track the dynamic biomolecular processes of individual biological macromolecules.
TIRFM allows us to observe the translational or rotational motion of individual ﬂuorescent spots that are emitted from ﬂuorophores attached to biomolecules. Since optical

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Fig. 80. Atomic-resolution image of a mica surface in water captured at 175 ms/frame. One of the successively
captured 400 images is shown. Scan area, 4.4  4.4 nm2.

microscopy and high-speed AFM have advantages and disadvantages over each other,
combining these techniques into a single instrument would therefore be useful. For example, when high-speed AFM is applied to living cells, the use of this sort of combined system is essential. From the optical image covering a wide area of the sample, we can quickly
ﬁnd a much narrower area to be scanned by AFM without trial imaging at various places.
We can visualize the dynamic structural changes of proteins by high-speed AFM while
simultaneously recording the optical signals of chemical reactions such as ATP hydrolysis
at individual protein molecules. The chemical speciﬁcation of an object imaged by AFM
can be determined from its Raman or ﬂuorescence signals. There have been a number ofreports on conventional slow AFM combined with confocal ﬂuorescence [228–231], conventional Raman scattering [232,233], coherent anti-Stokes Raman scattering [234,235],
or TIRF [236–238] microscopes. When the simultaneous recording of optical and AFM
images is required, some precautions are necessary regarding the luminescence of thecantilever tip [239–241] and the photothermal bending of the cantilever [111]. In tapping-mode AFM, the DC deﬂection of a cantilever does not aﬀect AFM imaging;
thus, the photothermal bending of the cantilever should not be a problem provided
its bending is not too large and the light power is stable. If necessary, we can switch
oﬀ the illumination light during the trace scan and switch it on during the retrace scan
[228].
However, building a combined system may not be as easy as the case of conventional
AFM, because high-speed AFM imposes various restrictions on the OBD detector and thescanner. Micro-electro-mechanical fabrication techniques, which have been employed to
produce self-sensing and/or self-actuation cantilevers [46–48] and sensor-combined scanners [242,243], may be the key to the realization of a combined system as well as to thesigniﬁcant enhancement of high-speed AFM performance.

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14. Conclusion
We have described various studies carried out in the last decade with the aim of increasing
the imaging rate and spatial resolution of bio-AFM, as well as the basic concepts and principles behind the techniques. Moreover, we have summarized recently performed visualization
studies on dynamic biomolecular processes, which were currently infeasible with other techniques. The direct and real-time observation of dynamic biomolecular processes is straightforward and can give deep insights to the functional mechanisms. Therefore, this new
microscopy will markedly change our style of considering biological questions. Nevertheless,there are presently only a few setups of high-speed bio-AFM in the world which can capture
dynamic biomolecular processes at 10–30 frames/s, and consequently, the user population islimited. Besides, to our knowledge, only two manufacturers are producing small cantilevers
for high-speed bio-AFM and the most advanced small cantilevers are not commercially available yet. We hope that this current situation will be quickly improved by manufacturers.
In the near future, high-speed AFM will be actively used to observe a wide range ofdynamic processes that occur in isolated proteins, protein assemblies, and protein–DNA
complexes, which will certainly induce enormous acceleration of our understanding of
how protein functions are produced. In the next stage, more complex systems including
living cells and organisms will become targets of high-speed AFM after it is successfully
combined with other techniques such as ultrasonic techniques and optical microscopy.
The in vivo and in vitro visualization of various processes at the molecular level will
become possible including the responses of membrane receptors to stimuli, nuclear envelope formation and disassembly, chromosome replication and segregation processes,
phagocytosis, protein synthesis in the endoplasmic reticulum, and the targeting processesof synthesized proteins through the Golgi apparatus. Thus, high-speed AFM-based visualization techniques have great potential to bring about breakthroughs not only in biochemistry and biophysics but also in cell biology, physiology, and pharmaceutical andmedical sciences. To open up such unprecedented ﬁelds, steady eﬀorts have to be carriedout towards expanding the capability of high-speed AFM and the related techniques.
Acknowledgements
We thank N. Kodera, D. Yamamoto, M. Taniguchi, A. Miyagi, H. Yamashita and all
previous students for their dedicated studies for developing high-AFM. This work was
supported by the Japan Science and Technology Agency (JST; the CREST programand a Grant-in-Aid for Development of Systems and Technology for Advanced Measurement and Analysis) and the Japan Society for the Promotion of Science (JSPS; a Grant-inAid for Basic Research (S)).
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in the online version, at
doi:10.1016/j.progsurf.2008.09.001.
References[1] G. Binnig, C.F. Quate, Ch. Gerber, Atomic force microscope, Phys. Rev. Lett. 56 (1986) 930–933.

428

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437[2] G. Binnig, H. Rohrer, Ch. Gerber, E. Weibel, Surface studies by scanning tunneling microscopy, Phys. Rev.
Lett. 49 (1982) 57–61.
[3] T. Ando, T. Uchihashi, N. Kodera, D. Yamamoto, A. Miyagi, M. Taniguchi, H. Yamashita, Invitedreview: high-speed AFM and nano-visualization of biomolecular processe, Pﬂüg. Arch.- Eur. J. Physiol. 456(2008) 211–225.
[4] T. Ando, T. Uchihashi, N. Kodera, D. Yamamoto, M. Taniguchi, A. Miyagi, H. Yamashita, Review: highspeed atomic force microscopy for observing dynamic biomolecular processes, J. Mol. Recognit. 20 (2007)
448–458.
[5] Q. Zhong, D. Inniss, K. Kjoller, V.B. Elings, Fractured polymer/silica ﬁber surface studied by tapping
mode atomic force microscopy, Surf. Sci. 290 (1993) L688–L692.
[6] J. Tamayo, R. Garcı́a, Deformation, contact time, and phase contrast in tapping mode scanning force
microscopy, Langmuir 12 (1996) 4430–4435.
[7] J.P. Cleveland, B. Anczykowski, A.E. Schmid, V.B. Elings, Energy dissipation in tapping-mode atomic
force microscopy, Appl. Phys. Lett. 72 (1998) 2613–2615.
[8] G. Bar, Y. Thomann, R. Brandsch, H.-J. Cantow, M.-H. Whangbo, Factors aﬀecting the height and phase
images in tapping mode atomic force microscopy. Study of phase-separated polymer blends of poly(etheneco-styrene) and poly (2,6-dimethyl-1,4-phenylene oxide), Langmuir 13 (1997) 3807–3812.
[9] T.R. Albrecht, P. Grütter, D. Horne, D. Rugar, Frequency modulation detection using high-Q cantilevers
for enhanced force microscope sensitivity, J. Appl. Phys. 69 (1991) 668–673.
[10] F.J. Giessibl, Atomic resolution of the silicon (1 1 1)–(7  7) surface by atomic force microscopy, Science
267 (1995) 68–71.
[11] P. Parot, Y.F. Dufrêne, P. Hinterdorfer, C. Le Grimellec, D. Navajas, J.-L. Pellequer, S. Scheuring, Past,
present and future of atomic force microscopy in life sciences and medicine, J. Mol. Recognit. 20 (2007)
418–431.
[12] O. Marti, B. Drake, P.K. Hansma, Atomic force microscopy of liquid-covered surfaces: atomic resolution
images, Appl. Phys. Lett. 51 (1987) 484–486.
[13] G. Binnig, Ch. Gerber, E. Stoll, T.R. Albrecht, C.F. Quate, Atomic resolution with atomic force
microscope, Europhys. Lett. 3 (1987) 1281–1286.
[14] G. Meyer, N.M. Amer, Novel optical approach to atomic force microscopy, Appl. Phys. Lett. 53 (1988)
1045–1047.
[15] S. Gould, O. Marti, B. Drake, L. Hellemans, C.E. Bracker, P.K. Hansma, N.L. Keder, M.M. Eddy, G.D.
Stucky, Molecular resolution images of amino acid crystals with the atomic force microscope, Nature 332(1988) 332–334.
[16] O. Marti, V. Elings, M. Haugan, C.E. Bracker, J. Schneir, B. Drake, S.A. Gould, J. Gurley, L. Hellemans,
K. Shaw, A.L. Weisenhorn, J. Zasadzinski, P.K. Hansma, Scanning probe microscopy of biological samples
and other surfaces, J. Microsc. 152 (1988) 803–809.
[17] G. Friedbacher, P.K. Hansma, E. Ramli, G.D. Stucky, Imaging powders with the atomic force microscope:from biominerals to commercial materials, Science 253 (1991) 1261–1263.
[18] J.N. Lin, B. Drake, A.S. Lea, P.K. Hansma, J.D. Andrade, Direct observation of immunoglobulin
adsorption dynamics using the atomic force microscope, Langmuir 6 (1990) 509–511.
[19] B. Drake, C.B. Prater, A.L. Weisenhorn, S.A. Gould, T.R. Albrecht, C.F. Quate, D.S. Cannell, H.G.
Hansma, P.K. Hansma, Imaging crystals, polymers, and processes in water with the atomic force
microscope, Science 243 (1989) 1586–1589.
[20] W. Häberle, J.K.H. Höber, F. Ohnesorge, D.P.E. Smith, G. Binnig, In situ investigations of single living
cells infected by viruses, Ultramicroscopy 4244 (Part B) (1992) 1161–1167.
[21] F. Ohnesorge, W.M. Heckl, W. Häberle, D. Pum, M. Sara, H. Schindler, K. Schilcher, A. Kiener, D.P.E.
Smith, U.B. Sleytr, G. Binnin, Scanning force microscopy studies of the S-layers from Bacillus coagulans
E38-66, Bacillus sphaericus CCM2177 and of an antibody binding process, Ultramicroscopy 4244 (Part B)(1992) 1236–1242.
[22] J.H. Hoh, R. Lal, S.A. John, J.-P. Revel, M.F. Arnsdorf, Atomic force microscopy and dissection of gap
junctions, Science 253 (1991) 1405–1408.
[23] H.J. Appel, J. Colchero, A. Linder, O. Marti, J. Mlynek, Na,K-ATPase in crystalline form investigated byscanning force microscopy, Ultramicroscopy 4244 (Part B) (1992) 1133–1140.
[24] A.L. Weisenhorn, B. Drake, C.B. Prater, S.A.C. Gould, P.K. Hansma, F. Ohnesorge, M. Egger, S.P. Heyn,
H.E. Gaub, Immobilized proteins in buﬀer imaged at molecular resolution by atomic force microscopy,
Biophys. J. 58 (1990) 1251–1258.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

429[25] R.D. Edstrom, M.H. Meinke, X.R. Yang, R. Yang, V. Elings, D.F. Evans, Direct visualization ofphosphorylase–phosphorylase kinase complexes by scanning tunneling and atomic force microscopy,
Biophys. J. 58 (1990) 1437–1448.
[26] M. Egger, F. Ohnesorge, A.L. Weisenhorn, S.P. Heyn, B. Drake, C.B. Prater, S.A.C. Gould, P.K. Hansma,
H.E. Gaub, Wet lipid–protein membranes imaged at submolecular resolution by atomic force microscopy,
J. Struct. Biol. 103 (1990) 89–94.
[27] H.J. Butt, K.H. Downing, P.K. Hansma, Imaging the membrane protein bacteriorhodopsin with the atomic
force microscope, Biophys. J. 58 (1990) 1473–1480.
[28] S.A.C. Gould, B. Drake, C.B. Prater, A.L. Weisenhorn, S. Manne, H.G. Hansma, P.K. Hansma, J. Massie,
M. Longmire, V. Elings, B. Dixon Northern, B. Mukergee, C.M. Peterson, W. Stoeckenius, T.R. Albrecht,
C.F. Quate, From atoms to integrated circuit chips, blood cells, and bacteria with the atomic force
microscope, J. Vac. Sci. Technol. A 8 (1990) 369–373.
[29] S. Karrasch, M. Dolder, F. Schabert, J. Ramsden, A. Engel, Covalent binding of biological samples to solid
supports for scanning probe microscopy in buﬀer solution, Biophys. J. 65 (1993) 2437–2446.
[30] F.A. Schabert, A. Engel, Reproducible acquisition of Escherichia coli porin surface topographs by atomic
force microscopy, Biophys. J. 67 (1994) 2394–2403.
[31] D.J. Müller, A. Engel, Voltage and pH-induced channel closure of porin OmpF visualized by atomic force
microscopy, J. Mol. Biol. 285 (1999) 1347–1351.
[32] T. Walz, P. Tittmann, K.H. Fuchs, D.J. Müller, B.L. Smith, P. Agre, H. Gross, A. Engel, Surface
topographies at subnanometer-resolution reveal asymmetry and sidedness of aquaporin-1, J. Mol. Biol. 264(1996) 907–918.
[33] D.J. Müller, F.A. Schabert, G. Büldt, A. Engel, Imaging purple membranes in aqueous solutions at
subnanometer resolution by atomic force microscopy, Biophys. J. 68 (1995) 1681–1686.
[34] D.J. Müller, G. Büldt, A. Engel, Force-induced conformational change of bacteriorhodopsin, J. Mol. Biol.
249 (1995) 239–243.
[35] C.A. Putman, K.O. Van der Werf, B.G. de Grooth, N.F. van Hulst, J. Greve, Viscoelasticity of living
cells allows high resolution imaging by tapping mode atomic force microscopy, Biophys. J. 67 (1994)
1749–1753.
[36] M.A. Lantz, S.J. O’Shea, M.E. Welland, Force microscopy imaging in liquids using ac techniques, Appl.
Phys. Lett. 65 (1994) 409–411.
[37] C. Möller, M. Allen, V. Elings, A. Engel, D.J. Müller, Tapping-mode atomic force microscopy produces
faithful high-resolution images of protein surfaces, Biophys. J. 77 (1999) 1150–1158.
[38] C. Bustamante, D.A. Erie, D. Keller, Biochemical and structural applications of scanning force microscopy,
Curr. Opin. Struct. Biol. 4 (1994) 750–760.
[39] D.A. Erie, G. Yang, H.C. Schultz, C. Bustamante, DNA bending by Cro protein in speciﬁc andnonspeciﬁc complexes: implications for protein site recognition and speciﬁcity, Science 266 (1994) 1562–
1566.
[40] M. Bezanilla, B. Drake, E. Nudler, M. Kashlev, P.K. Hansma, H.G. Hansma, Motion and enzymatic
degradation of DNA in the atomic force microscope, Biophys. J. 67 (1994) 2454–2459.
[41] M. Guthold, M. Bezanilla, D.A. Erie, B. Jenkins, H.G. Hansma, C. Bustamante, Following the assembly of
RNA polymerase–DNA complexes in aqueous solutions with the scanning force microscope, Proc. Natl.
Acad. Sci. USA 91 (1994) 12927–12931.
[42] S. Kasas, N.H. Thomson, B.L. Smith, H.G. Hansma, X. Zhu, M. Guthold, C. Bustamante, E.T. Kool, M.
Kashlev, P.K. Hansma, Escherichia coli RNA polymerase activity observed using atomic force microscopy,
Biochemistry 36 (1997) 461–468.
[43] M. Guthold, X. Zhu, C. Rivetti, G. Yang, N.H. Thomson, S. Kasas, H.G. Hansma, B. Smith, P.K.
Hansma, C. Bustamante, Direct observation of one-dimensional diﬀusion and transcription by Escherichia
coli RNA polymerase, Biophys. J. 77 (1999) 2284–2294.
[44] H. Lin, D.O. Clegg, R. Lal, Imaging real-time proteolysis of single collagen I molecules with an atomic
force microscope, Biochemistry 38 (1999) 9956–9963.
[45] H. Oberleithner, H. Schillers, M. Wilhelmi, D. Butzke, T. Danker, Nuclear pores collapse in response to
CO2 imaged with atomic force microscopy, Pﬂüg. Arch.- Eur. J. Physiol. 439 (2000) 251–255.
[46] R.C. Barrett, C.F. Quate, High-speed, large-scale imaging with the atomic force microscope, J. Vac. Sci.
Technol. B 9 (1991) 302–306.
[47] S.C. Minne, S.R. Manalis, C.F. Quate, Parallel atomic force microscopy using cantilevers with integrated
piezoresistive sensors and integrated piezoelectric actuators, Appl. Phys. Lett. 67 (1995) 3918–3920.

430

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437[48] S.R. Manalis, S.C. Minne, C.F. Quate, Atomic force microscopy for high speed imaging using cantilevers
with an integrated actuator and sensor, Appl. Phys. Lett. 68 (1996) 871–873.
[49] H.-J. Butt, P. Siedle, K. Seifert, K. Fendler, T. Seeger, E. Bamberg, A.L. Weisenhorn, K. Goldie, A. Engel,
Scan speed limit in atomic force microscopy, J. Microsc. 169 (1993) 75–84.
[50] D.A. Walters, J.P. Cleveland, N.H. Thomson, P.K. Hansma, M.A. Wendman, G. Gurley, V. Elings, Short
cantilevers for atomic force microscopy, Rev. Sci. Instrum. 67 (1996) 3583–3590.
[51] T.E. Schäﬀer, J.P. Cleveland, F. Ohnesorge, D.A. Walters, P.K. Hansma, Studies of vibrating atomic force
microscope cantilevers in liquid, J. Appl. Phys. 80 (1996) 3622–3627.
[52] M.B. Viani, T.E. Schäﬀer, G.T. Paloczi, L.I. Pietrasanta, B.L. Smith, J.B. Thompson, M. Richter, M. Rief,
H.E. Gaub, K.W. Plaxco, A.N. Cleland, H.G. Hansma, P.K. Hansma, Fast imaging and fast force
spectroscopy of single biopolymers with a new atomic force microscope designed for small cantilevers, Rev.
Sci. Instrum. 70 (1999) 4300–4303.
[53] M.B. Viani, L.I. Pietrasanta, J.B. Thompson, A. Chand, I.C. Gebeshuber, J.H. Kindt, M. Richter, H.G.
Hansma, P.K. Hansma, Probing protein–protein interactions in real time, Nat. Struct. Biol. 7 (2000) 644–
647.
[54] T. Ando, N. Kodera, E. Takai, D. Maruyama, K. Saito, A. Toda, A high-speed atomic force microscopefor studying biological macromolecules, Proc. Natl. Acad. Sci. USA 98 (2001) 12468–12472.
[55] T. Ando, N. Kodera, D. Maruyama, E. Takai, K. Saito, A. Toda, A high-speed atomic force microscopefor studying biological macromolecules in action, Jpn. J. Appl. Phys. 41 (2002) 4851–4856.
[56] S.P. Gross, M. Vershinin, G.T. Shubeita, Cargo transport: two motors are sometimes better than one, Curr.
Biol. 17 (2007) R478–R486.
[57] K. Kalil, E.W. Dent, Touch and go: guidance cues signal to the growth cone cytoskeleton, Curr. Opin.
Neurobiol. 15 (2005) 521–526.
[58] C. Revenu, R. Athman, S. Robine, D. Louvard, The co-workers of actin ﬁlaments: from cell structures tosignals, Nat. Rev. Mol. Cell. Biol. 5 (2004) 635–646.
[59] T.M. Lohman, K.P. Bjornson, Mechanisms of helicase-catalyzed DNA unwinding, Ann. Rev. Biochem. 65(1996) 169–214.
[60] G. Charvin, T.R. Strick, D. Bensimon, V. Croquette, Tracking topoisomerase activity at the singlemolecule level, Ann. Rev. Biophys. Biomol. Struct. 34 (2005) 201–219.
[61] D. Thirumalai, G.H. Lorimer, Chaperonin-mediated protein folding, Annu. Rev. Biophys. Biomol. Struct.
30 (2001) 245–269.
[62] H. Nakajima, Y. Kunioka, K. Nakano, K. Shimizu, M. Seto, T. Ando, Scanning force microscopy of theinteraction events between a single molecule of heavy meromyosin and actin, Biochem. Biophys. Res.
Commun. 234 (1997) 178–182.
[63] J.J. Schmidt, C.D. Montemagno, Bionanomechanical systems, Annu. Rev. Mater. Res. 34 (2004) 315–337.
[64] T. Fukuma, M.J. Higgins, S.P. Jarvis, Direct imaging of individual intrinsic hydration layers on lipid
bilayers at Ångstrom resolution, Biophys. J. 92 (2007) 3603–3609.
[65] S.W. Hell, M. Dyba, S. Jakobs, Concepts for nanoscale resolution in ﬂuorescence microscopy, Curr. Opin.
Neurobiol. 14 (2004) 599–609.
[66] G. Donnert, J. Keller, R. Medda, M.A. Andrei, S.O. Rizzoli, R. Lührmann, R. Jahn, C. Eggeling, S.W.
Hell, Macromolecular-scale resolution in biological ﬂuorescence microscopy, Proc. Natl. Acad. Sci. USA
103 (2006) 11440–11445.
[67] V. Westphal, S.O. Rizzoli, M.A. Lauterbach, D. Kamin, R. Jahn, S.W. Hell, Video-rate far-ﬁeld opticalnanoscopy dissects synaptic vesicle movement, Science 320 (2008) 246–249.
[68] G.S. Shekhawat, V.P. Dravid, Nanoscale imaging of buried structures via scanning near-ﬁeld ultrasoundholography, Science 310 (2005) 89–92.
[69] T. Sulchek, G.G. Yaralioglu, C.F. Quate, S.C. Minne, Characterization and optimization of scan speed fortapping-mode atomic force microscopy, Rev. Sci. Instrum. 73 (2002) 2928–2936.
[70] J. Kokavecz, O. Marti, P. Heszler, Á. Mechler, Imaging bandwidth of the tapping mode atomic force
microscope probe, Phys. Rev. B 73 (2006) 155403.
[71] N. Kodera, M. Sakashita, T. Ando, Dynamic proportional–integral–diﬀerential controller for high-speed
atomic force microscopy, Rev. Sci. Instrum. 77 (2006) 083704.
[72] M. Kitazawa, K. Shiotani, A. Toda, Batch fabrication of sharpened silicon nitride tips, Jpn. J. Appl. Phys.
42 (Part 1) (2003) 4844–4847.
[73] M. Tanemura, T. Okita, H. Yamauchi, S. Tanemura, R. Morishima, Room-temperature growth of acarbon nanoﬁber on the tip of conical carbon protrusions, Appl. Phys. Lett. 84 (2004) 3831–3833.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

431[74] M. Kitazawa, R. Ohta, T. Okita, J. Tanaka, M. Tanemura, Mechanical properties of single carbon
nanoﬁbers grown on tips of scanning probe microscopy cantilevers by ion irradiation, Jpn. J. Appl. Phys. 46(2007) 6324–6328.
[75] C.L. Cheung, J.H. Hafner, C.M. Lieber, Carbon nanotube atomic force microscopy tips: direct growth bychemical vapor deposition and application to high-resolution imaging, Proc. Natl. Acad. Sci. USA 97(2000) 3809–3813.
[76] T. Fukuma, M. Kimura, K. Kobayashi, K. Matsushige, H. Yamada, Development of low noise cantileverdeﬂection sensor for multienvironment frequency-modulation atomic force microscopy, Rev. Sci. Instrum.
76 (2005) 053704.
[77] J. Kokavecz, Z. Tóth, Z.L. Horváth, P. Heszler, Á Mechler, Novel amplitude and frequency demodulation
algorithm for a virtual dynamic atomic force microscope, Nanotechnology 17 (2006) S173–S177.
[78] R. Hillenbrand, M. Stark, R. Guckenberger, Higher-harmonics generation in tapping-mode atomic-force
microscopy: insights into the tip–sample interaction, Appl. Phys. Lett. 76 (2000) 3478–3480.
[79] R.W. Stark, W.M. Heckl, Higher harmonics imaging in tapping-mode atomic-force microscopy, Rev. Sci.
Instrum. 74 (2003) 5111–5114.
[80] S. Crittenden, A. Raman, R. Reifenberger, Probing attractive forces at the nanoscale using higher-harmonic
dynamic force microscopy, Phys. Rev. B 72 (2005) 235422.
[81] J. Legleiter, M. Park, B. Cusick, T. Kowalewski, Scanning probe acceleration microscopy (SPAM) in ﬂuids:
mapping mechanical properties of surfaces at the nanoscale, Proc. Natl. Acad. Sci. USA 103 (2006) 4813–
4818.
[82] W.-J. Chang, J.-C. Hsu, T.-H. Lai, Inverse calculation of the tip–sample interaction force in atomic force
microscopy by the conjugate gradient method, J. Phys. D: Appl. Phys. 37 (2004) 1123–1126.
[83] M. Stark, R.W. Stark, W.M. Heck, R. Guckenberger, Inverting dynamic force microscopy: from signals totime-resolved interaction forces, Proc. Natl. Acad. Sci. USA 99 (2002) 8473–8478.
[84] O. Sahin, Harnessing bifurcations in tapping-mode atomic force microscopy to calibrate time-varying tip–
sample force measurements, Rev. Sci. Instrum. 78 (2007) 103707.
[85] O. Shahin, S. Magonov, C. Su, C.F. Quate, O. Solgaard, An atomic force microscope tip designed tomeasure time-varying nanomechanical forces, Nat. Nanotechnol. 2 (2007) 507–514.
[86] J.H. Kindt, G.E. Fantner, J.A. Cutroni, P.K. Hansma, Rigid design of fast scanning probe microscopes
using ﬁnite element analysis, Ultramicroscopy 100 (2004) 259–265.
[87] T. Ando, T. Uchihashi, N. Kodera, A. Miyagi, R. Nakakita, H. Yamashita, K. Matada, High-speed atomic
force microscopy for capturing the dynamic behavior of protein molecules at work, e-J. Surf. Sci. Nanotech.
3 (2005) 384–392.
[88] M.J. Rost, L. Crama, P. Schakel, E. van Tol, G.B.E.M. van Velzen-Williams, C.F. Overgauw, H. ter Horst,
H. Dekker, B. Okhuijsen, M. Seynen, A. Vijftigschild, P. Han, A.J. Katan, K. Schoots, R. Schumm, W. vanLoo, T.H. Oosterkamp, J.W.M. Frenken, Scanning probe microscopes go video rate and beyond, Rev. Sci.
Instrum. 76 (2005) 053710.
[89] B. Anczykowski, J.P. Cleveland, D. Krüger, V. Elings, H. Fuchs, Analysis of the interaction mechanisms indynamic mode SFM by means of experimental data and computer simulation, Appl. Phys. A 66 (1998)
S885–S889.
[90] S. Gao, L.F. Chi, S. Lenhert, B. Anczykowski, C.M. Niemeyer, M. Adler, H. Fuchs, High quality mappingof DNA–protein complexes by dynamic scanning force microscopy, Chem. Phys. Chem. 6 (2001) 384–388.
[91] J. Tamayo, A.D.L. Humphris, R.J. Owen, M.J. Miles, High-Q dynamic force microscopy in liquid and its
application to living cells, Biophys. J. 81 (2001) 526–537.
[92] N. Kodera, H. Yamashita, T. Ando, Active damping of the scanner for high-speed atomic force
microscopy, Rev. Sci. Instrum. 76 (2005) 053708.
[93] G. Schitter, F. Allgöwer, A. Stemmer, A new control strategy for high-speed atomic force microscopy,
Nanotechnology 15 (2004) 108–114.
[94] S.K. Hung, E.-T. Hwu, I.-S. Hwang, L.-C. Fu, Postﬁtting control scheme for periodic piezoscanner driving,
Jpn. J. Appl. Phys. 45B (2006) 1917–1921.
[95] A. Stemmer, G. Schitter, J.M. Rieber, F. Allgöwer, Control strategies towards faster quantitative imaging
in atomic force microscopy, Eur. J. Contr. 11 (2005) 384–395.
[96] Q. Zou, K.K. Leang, E. Sadoun, M.J. Reed, S. Devasia, Control issues in high-speed AFM for biologicalapplications: collagen imaging example, Asian J. Control 6 (2004) 164–178.
[97] Y. Wu, Q. Zou, Iterative control approach to compensate for both the hysteresis and the dynamics eﬀects ofpiezo actuators, IEEE Trans. Contr. Syst. Technol. 15 (2007) 936–944.

432

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437[98] G.M. Clayton, S. Tien, A.J. Fleming, S.O.R. Moheimani, S. Devasia, Inverse-feed-forward of chargecontrolled piezopositioners, Mechatronics 18 (2008) 273–281.
[99] T. Fukuma, Y. Okazaki, N. Kodera, T. Uchihashi, T. Ando, High resonance frequency force microscopy
scanner using inertia balance support, Appl. Phys. Lett. 92 (2008) 243119.
[100] J. Schiener, S. Witt, M. Stark, R. Guckenberger, Stabilized atomic force microscopy imaging in liquids
using second harmonic of cantilever motion for setpoint control, Rev. Sci. Instrum. 75 (2004) 2564–
2568.
[101] T. Uchihashi, N. Kodera, H. Itoh, H. Yamashita, T. Ando, Feed-forward control for high-speed AFM
imaging of biomolecules, Jpn. J. Appl. Phys. 45 (2006) 1904–1908.
[102] W. Han, S.M. Lindsay, T. Jing, A magnetically driven oscillating probe microscope for operation in liquids,
Appl. Phys. Lett. 69 (1996) 4111–4113.
[103] G.R. Jayanth, Y. Jeong, C.-H. Menq, Direct tip-position control using magnetic actuation for achieving
fast scanning in tapping mode atomic force microscopy, Rev. Sci. Instrum. 77 (2006) 053704.
[104] F.L. Degertekin, B. Hadimioglu, T. Sulchek, C.F. Quate, Actuation and characterization of atomic force
microscope cantilevers in ﬂuids by acoustic radiation pressure, Appl. Phys. Lett. 78 (2001) 1628–1630.
[105] A.G. Onaran, F.L. Degertekin, B. Hadimioglu, Tapping mode and elasticity imaging in liquids using
an atomic force microscope actuated by acoustic radiation pressure, Appl. Phys. Lett. 80 (2002) 4063–
4065.
[106] A.G. Onaran, F.L. Degertekin, A ﬂuid cell with integrated acoustic radiation pressure actuator for atomic
force microscopy, Rev. Sci. Instrum. 76 (2005) 103703.
[107] N. Umeda, S. Ishizaki, H. Uwai, Scanning attractive force microscope using photothermal vibration, J.
Vac. Sci. Technol. B 9 (1991) 1318–1322.
[108] O. Marti, A. Ruf, M. Hipp, H. Bielefeldt, J. Colchero, J. Mlynek, Mechanical and thermal eﬀects of laser
irradiation on force microscope cantilevers, Ultramicroscopy A 42–44 (1992) 345–350.
[109] D. Ramos, J. Tamayo, J. Mertens, M. Calleja, Photothermal excitation of microcantilevers in liquids, J.
Appl. Phys. 99 (2006) 124909.
[110] S.S. Verbridge, L.M. Bellan, J.M. Parpia, C. Craighead, Optically driven resonance of nanoscale ﬂexural
oscillators in liquid, Nano Lett. 6 (2006) 2109–2114.
[111] H. Yamashita, T. Uchihashi, N. Kodera, A. Miyagi, D. Yamamoto, T. Uchihashi, T. Ando, Tip–sample
distance control using photo-thermal actuation of a small cantilever for high-speed atomic force
microscopy, Rev. Sci. Instrum. 78 (2007) 083702.
[112] G.E. Fantner, P. Hegarty, J.H. Kindt, G. Schitter, G.A.G. Cidade, P.K. Hansma, Data acquisition system
for high speed atomic force microscopy, Rev. Sci. Insturm. 76 (2005) 026118.
[113] G.E. Fantner, G. Schitter, J.H. Kindt, T. Ivanov, K. Ivanova, R. Patel, N. Holten-Andersen, J. Adams, P.J.
Thurner, I.W. Rangelow, P.K. Hansma, Components for high speed atomic force microscopy,
Ultramicroscopy 106 (2006) 881–887.
[114] S.N. Magonov, J.P. Cleveland, V. Elings, D. Denley, M.H. Whangbo, Tapping-mode atomic force
microscopy study of the near-surface composition of a styrene–butadiene–styrene triblock copolymer ﬁlm,
Surf. Sci. 389 (1997) 201–211.
[115] G. Bar, Y. Thomann, M.-H. Whangbo, Characterization of the morphologies and nanostructures of blendsof poly(styrene)-block-poly(ethene-co-but-1-ene)-block-poly(styrene) with isotactic and atactic polypropylenes by tapping-mode atomic force microscopy, Langmuir 14 (1998) 1219–1226.
[116] R. Garcı́a, R. Magerle, R. Perez, Nanoscale compositional mapping with gentle forces, Nat. Mater. 6 (2007)
405–411.
[117] D.M. Czajkowsky, M.J. Allen, V. Elings, Z. Shao, Direct visualization of surface charge in aqueous
solution, Ultramicroscopy 74 (1998) 1–5.
[118] N.F. Martı́nez, R. Garcı́a, Measuring phase shifts and energy dissipation with amplitude modulation
atomic force microscopy, Nanotechnology 17 (2006) S167–S172.
[119] T. Uchihashi, T. Ando, H. Yamashita, Fast phase imaging in liquids using a rapid scan atomic force
microscope, Appl. Phys. Lett. 89 (2006) 213112.
[120] M. Stark, R. Guckenberger, Fast low-cost phase detection setup for tapping-mode atomic force
microscopy, Rev. Sci. Instrum. 70 (1999) 3614–3619.
[121] T. Fukuma, K. Kobayashi, K. Matsushige, H. Yamada, True molecular resolution in liquid by frequencymodulation atomic force microscopy, Appl. Phys. Lett. 86 (2005) 193108.
[122] T. Fukuma, M.J. Higgins, S.P. Jarvis, Direct imaging of lipid-ion network formation under physiologicalconditions by frequency modulation atomic force microscopy, Phys. Rev. Lett. 98 (2007) 106101.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

433[123] T. Fukuma, S.P. Jarvis, Development of liquid-environment frequency modulation atomic force
microscope with low noise deﬂection sensor for cantilevers of various dimensions, Rev. Sci. Instrum. 77(2006) 043701.
[124] Y. Martin, C.C. Williams, H.K. Wickramasinghe, Atomic force microscope–force mapping and proﬁling
on a sub 100-Å scale, J. Appl. Phys. 61 (1987) 4723–4729.
[125] A.L. Weisenhorn, M. Egger, F. Ohnesorge, S.A.C. Gould, S.-P. Heyn, H.G. Hansma, R.L. Sinsheimer,
H.E. Gaub, P.K. Hansma, Molecular-resolution images of Langmuir–Blodgett ﬁlms and DNA by atomic
force microscopy, Langmuir 7 (1991) 8–12.
[126] P.K. Hansma, J.P. Cleveland, M. Radmacher, D.A. Walters, P.E. Hillner, M. Bezanilla, M. Fritz, D. Vie,
H.G. Hansma, C.B. Prater, J. Massie, L. Fukunaga, J. Gurley, V. Elings, Tapping mode atomic force
microscopy in liquids, Appl. Phys. Lett. 64 (1994) 1738–1740.
[127] C.A.J. Putman, K.O.V. der Werf, B.G.D. Grooth, N.F.V. Hulst, J. Greve, Tapping mode atomic force
microscopy in liquid, Appl. Phys. Lett. 64 (1994) 2454–2456.
[128] F.J. Giessibl, A direct method to calculate tip–sample forces from frequency shifts in frequency-modulation
atomic force microscopy, Appl. Phys. Lett. 78 (2001) 123–125.
[129] F.J. Giessibl, H. Bielefeldt, S. Hembacher, J. Mannhart, Calculation of the optimal imaging parameters forfrequency modulation atomic force microscopy, Appl. Surf. Sci. 140 (1999) 352–357.
[130] F.J. Giessibl, S. Hembacher, H. Bielefeldt, J. Mannhart, Subatomic features on the silicon (1 1 1)–(7  7)
surface observed by atomic force microscopy, Science 289 (2000) 422–425.
[131] M. Guggisberg, M. Bammerlin, Ch. Loppacher, O. Pfeiﬀer, A. Abdurixit, V. Barwich, R. Bennewitz, A.
Baratoﬀ, E. Meyer, H.-J. Güntherodt, Separation of interactions by noncontact force microscopy, Phys.
Rev. B 61 (2000) 11151–11155.
[132] T. Fukuma, K. Kobayashi, K. Matsushige, H. Yamada, True atomic resolution in liquid by frequencymodulation atomic force microscopy, Appl. Phys. Lett. 87 (2005) 034101.
[133] T. Fukuma, K. Kobayashi, H. Yamada, K. Matsushige, Surface potential measurements by the dissipative
force modulation method, Rev. Sci. Instrum. 75 (2004) 4589–4594.
[134] N. Burnham, R.J. Colton, Measuring the nanomechanical properties and surface forces of materials using
atomic force microscopy, J. Vac. Sci. Technol. A 7 (1989) 2906–2913.
[135] D.J. Müller, D. Fotiadis, S. Scheuring, S.A. Müller, A. Engel, Electrostatically balanced subnanometer
imaging of biological specimens by atomic force microscope, Biophys. J. 76 (1999) 1101–1111.
[136] F.J. Giessibl, M. Tortonese, Self-oscillating mode for frequency modulation noncontact atomic force
microscopy, Appl. Phys. Lett. 70 (1997) 2529–2531.
[137] F.J. Giessibl, Principle of nc-AFM, in: S. Morita, R. Wiesendanger, E. Meyer (Eds.), Noncontact Atomic
Force Microscopy, Springer, New York, 2000, pp. 11–46.
[138] M. Ojima, A. Arimoto, N. Chinone, T. Gotoh, K. Aiki, Diode laser noise at vide frequencies in optical
videodisc players, Appl. Opt. 25 (1986) 1404–1410.
[139] A. Arimoto, M. Ojima, N. Chinone, A. Oishi, T. Gotoh, N. Ohnuki, Optimum conditions for the highfrequency noise reduction method in optical videodisc players, Appl. Opt. 25 (1986) 1398–1403.
[140] T. Ando, N. Kodera, Y. Naito, T. Kinoshita, K. Furuta, Y.Y. Toyoshima, A high-speed atomic force
microscope for studying biological macromolecules in action, Chem. Phys. Chem. 4 (2003) 1196–1202.
[141] T. Ando, T. Uchihashi, N. Kodera, A. Miyagi, R. Nakakita, H. Yamashita, M. Sakashita, High-speed
atomic force microscopy for studying the dynamic behavior of protein molecules at work, Jpn. J. Appl.
Phys. 45 (2006) 1897–1903.
[142] R.E. Cheney, M.K. O’ Shea, J.E. Heuser, M.V. Coelho, J.S. Wolenski, E.M. Espreaﬁco, P. Forscher, R.E.
Larson, M.S. Mooseker, Brain myosin-V is a two-headed unconventional myosin with motor activity, Cell
75 (1993) 13–23.
[143] F.S. Espindola, D.M. Suter, L.B.E. Partata, T. Cao, J.S. Wolenski, R.E. Cheney, S.M. King, M.S.
Mooseker, The light chain composition of chicken brain myosin-Va: calmodulin, myosin-II essential lightchains, and 8-kDa dynein light chain/PIN, Cell Motil. Cytoskel. 47 (2000) 269–281.
[144] T. Sakamoto, I. Amitani, E. Yokota, T. Ando, Direct observation of processive movement by individual
myosin V molecules, Biochem. Biophys. Res. Commun. 272 (2000) 586–590.
[145] A.A.C. Nascimento, R.E. Cheney, S.B.F. Tauhata, R.E. Larson, M.S. Mooseker, Enzymatic characterization and functional domain mapping of brain myosin-V, J. Biol. Chem. 271 (1996) 17561–17569.
[146] H. Koide, T. Kinoshita, Y. Tanaka, S. Tanaka, N. Nagura, G. Meyer zu Hörste, A. Miyagi, T. Ando,
Identiﬁcation of the single speciﬁc IQ motif of myosin V from which calmodulin dissociates in the presenceof Ca2+, Biochemistry 45 (2006) 11598–11604.

434

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437[147] S.A. Burgess, M.L. Walker, H. Sakakibara, P.J. Knight, K. Oiwa, Dynein structure and power stroke,
Nature 421 (2003) 715–718.
[148] A. Yildiz, J.N. Forkey, S.A. McKinney, T. Ha, Y.E. Goldman, P.R. Selvin, Myosin V walks hand-overhand: single ﬂuorophore imaging with 1.5-nm localization, Science 300 (2003) 2061–2065.
[149] J.N. Forkey, M.E. Quinlan, M.A. Shaw, J.E.T. Corrie, Y.E. Goldman, Three-dimensional
structural dynamics of myosin V by single-molecule ﬂuorescence polarization, Nature 422 (2003)
399–404.
[150] D.M. Warshaw, G.G. Kennedy, S.S. Work, E.B. Krementsova, S. Beck, K.M. Trybus, Diﬀerential labelingof myosin V heads with quantum dots allows direct visualization of hand-over-hand processivity, Biophys.
J. 88 (2005) L30–L32.
[151] S. Syed, G.E. Snyder, C. Franzini-Armstrong, P.R. Selvin, Y.E. Goldman, Adaptability of myosin V
studied by simultaneous detection of position and orientation, EMBO J. 25 (2006) 1795–1803.
[152] S. Burgess, M. Walker, F. Wang, J.P. Sellers, H.D. White, P.J. Knight, J. Trinick, The prepower stroke
conformation of myosin V, J. Cell Biol. 159 (2002) 983–991.
[153] K. Braig, Z. Otwinowski, R. Hegde, D.C. Boisvert, A. Joachimiak, A.L. Horwich, P.B. Sigler, The crystal
structure of the bacterial chaperonln GroEL at 2.8 Å, Nature 371 (1994) 578–586.
[154] Z. Xu, A.L. Horwich, P.B. Sigler, The crystal structure of the asymmetric GroEL–GroES–(ADP)7chaperonin complex, Nature 388 (1997) 741–750.
[155] S.G. Burston, N.A. Ranson, A.R. Clarke, The origins and consequences of asymmetry in the chaperonin
reaction cycle, J. Mol. Biol. 249 (1995) 138–152.
[156] O. Yifrach, A. Horovitz, Nested cooperativity in the ATPase activity of the oligomeric chaperonin GroEL,
Biochemistry 34 (1995) 5303–5308.
[157] G.H. Lorimer, Protein folding. Folding with a two-stroke motor, Nature 388 (1997) 720–722.
[158] H.S. Rye, S.G. Burston, W.A. Fenton, J.M. Beechem, Z. Xu, P.B. Sigler, A.L. Horwich, Distinct actions of
cis and trans ATP within the double ring of the chaperonin GroEL, Nature 388 (1997) 792–798.
[159] H.S. Rye, A.M. Roseman, S. Chen, K. Furtak, W.A. Fenton, H.R. Saibil, A.L. Horwich, GroEL–
GroES cycling: ATP and nonnative polypeptide direct alternation of folding-active rings, Cell 97 (1999)
325–338.
[160] A. Azem, M. Kessel, P. Goloubinoﬀ, Characterization of a functional GroEL14(GroES7)2 chaperonin
hetero-oligomer, Science 265 (1994) 653–656.
[161] M. Schmidt, K. Rutkat, R. Rachel, G. Pfeifer, R. Jaenicke, P. Viitanen, G. Lorimer, J. Buchner, Symmetriccomplexes of GroE chaperonins as part of the functional cycle, Science 265 (1994) 656–659.
[162] A. Engel, M.K. Hayer-Hartl, K.N. Goldie, G. Pfeifer, R. Hegerl, S. Muller, A.C.R. Da Silva, W.
Baumeister, F.U. Hartl, Functional signiﬁcance of symmetrical versus asymmetrical GroEL–GroES
chaperonin complexes, Science 269 (1995) 832–836.
[163] M.K. Hayer-Hartl, J. Martin, F.U. Hartl, Asymmetrical interaction of GroEL and GroES in the ATPase
cycle of assisted protein folding, Science 269 (1995) 836–841.
[164] H. Grallert, J. Buchner, Review: a structural view of the GroE chaperone cycle, J. Struct. Biol. 135 (2001)
95–103.
[165] H. Taguchi, T. Ueno, H. Tadakuma, M. Yoshida, T. Funatsu, Single-molecule observation ofproteinprotein interactions in the chaperonin system, Nat. Biotechnol. 19 (2001) 861–865.
[166] S.A. Darst, M. Ahlers, P.H. Meller, E.W. Kubalek, R. Blankenburg, H.O. Ribi, H. Ringsdorf, R.D.
Kornberg, Twodimensional crystals of streptavidin on biotinylated lipid layers and their interactions withbiotinylated macromolecules, Biophys. J. 59 (1991) 387–396.
[167] S.-T. Yau, B.R. Thomas, P.G. Vekilov, Molecular mechanisms of crystallization and defect formation,
Phys. Rev. Lett. 85 (2000) 353–356.
[168] D. Yamamoto, T. Uchihashi, N. Kodera, T. Ando, Anisotropic diﬀusion of point defects in twodimensional crystal of streptavidin observed by high-speed atomic force microscopy, Nanotechnology, 19(2008) 384009.
[169] A.C. Ku, S.A. Darst, C.R. Robertson, A.P. Gast, R.D. Kornberg, Molecular analysis of two-dimensional
protein crystallization, J. Phys. Chem. 97 (1993) 3013–3016.
[170] S.-W. Wang, C.R. Robertson, A.P. Gast, Molecular arrangement in two-dimensional streptavidin crystals,
Langmuir 15 (1999) 1541–1548.
[171] S.J. Demarest, M. Martinez-Yamout, J. Chung, H. Chen, W. Xu, H.J. Dyson, R.M. Evans, P.E. Wright,
Mutual synergistic folding in recruitment of CBP/p300 by p160 nuclear receptor coactivators, Nature 415(2002) 549–553.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

435[172] Y. Minezaki, K. Homma, A.R. Kinjo, K. Nishikawa, Human transcription factors contain a high fractionof intrinsically disordered regions essential for transcriptional regulation, J. Mol. Biol. 359 (2006) 1137–
1149.
[173] A. Miyagi, Y. Tsunaka, T. Uchihashi, K. Mayanagi, S. Hirose, K. Morikawa, T. Ando, Visualization of
intrinsically disordered regions of proteins by high-speed atomic force microscopy, Chem. Phys. Chem. 9(2008) 1859–1866.
[174] R. Belotserkovskaya, S. Oh, V.A. Bondarenko, G. Orphanides, V.M. Studitsky, D. Reinberg, FACT
facilitates transcription-dependent nucleosome alteration, Science 301 (2003) 1090–1093.
[175] D. Reinberg, R.J. Sims III, de FACT of nucleosome dynamics, J. Biol. Chem. 281 (2006) 23297–23301.
[176] T. Shimojima, M. Okada, T. Nakayama, H. Ueda, K. Okawa, A. Iwamatsu, H. Handa, S. Hirose,
Drosophila FACT contributes to Hox gene expression through physical and functional interactions withGAGA factor, Gene. Dev. 17 (2003) 1605–1616.
[177] C. Rivetti, M. Guthold, C. Bustamante, Scanning force microscopy of DNA deposited onto mica:
equilibration versus kinetic trapping studied by statistical polymer chain analysis, J. Mol. Biol. 264 (1996)
919–932.
[178] G.S. Manning, The persistence length of DNA is reached from the persistence length of its null isomerthrough an internal electrostatic stretching force, Biophys. J. 91 (2006) 3607–3616.
[179] H. Kojima, A. Ishijima, T. Yanagida, Direct measurement of stiﬀness of single actin ﬁlaments with andwithout tropomyosin by in vitro nanomanipulation, Proc. Natl. Acad. Sci. USA 91 (1994) 12962–12966.
[180] M. Radmacher, M. Fritz, J.P. Cleveland, D.A. Walters, P.K. Hansma, Imaging adhesion forces andelasticity of lysozyme adsorbed on mica with the atomic force microscope, Langmuir 10 (1994) 3809–3814.
[181] M. Tachibana, H. Koizumi, K. Kojima, Eﬀect of intracrystalline water on longitudinal sound velocity intetragonal hen-egg-white lysozyme crystals, Phys. Rev. E 69 (2004) 051921.
[182] R. Afrin, M.T. Alam, A. Ikai, Pretransition and progressive softening of bovine carbonic anhydrase II asprobed by single molecule atomic force microscopy, Protein Sci. 14 (2005) 1447–1457.
[183] A.D.L. Humphris, J.K. Hobbs, M.J. Miles, Ultrahigh-speed scanning near-ﬁeld optical microscopy capableof over 100 frames per second, Appl. Phys. Lett. 83 (2003) 6–8.
[184] A.D.L. Humphris, M.J. Miles, J.K. Hobbs, A mechanical microscope: high-speed atomic force microscopy,
Appl. Phys. Lett. 86 (2005) 034106.
[185] M. Yokokawa, C. Wada, T. Ando, N. Sakai, A. Yagi, S.H. Yoshimura, K. Takeyasu, Fast-scanning atomic
force microscopy reveals the ATP/ADP-dependent conformational changes of GroEL, EMBO J. 25 (2006)
4567–4576.
[186] M. Yokokawa, S.H. Yoshimura, Y. Naito, T. Ando, A. Yagi, N. Sakai, K. Takeyasu, Fast-scanning atomic
force microscopy reveals the molecular mechanism of DNA cleavage by ApaI endonuclease, IEE Proc.
Nanobiotechnol. 153 (2006) 60–66.
[187] N. Crampton, M. Yokokawa, D.T.F. Dryden, J.M. Edwardson, D.N. Rao, K. Takeyasu, S.H. Yoshimura,
R.M. Henderson, Fast-scan atomic force microscopy reveals that the type III restriction enzyme EcoP15I iscapable of DNA translocation and looping, Proc. Natl. Acad. Sci. USA 104 (2007) 12755–12760.
[188] M. Kobayashi, K. Sumitomo, K. Torimitsu, Real-time imaging of DNA–streptavidin complex formation insolution using a high-speed atomic force microscope, Ultramicroscopy 107 (2007) 184–190.
[189] S.F. Zhang, P. Rolfe, G. Wright, W. Lian, A.J. Milling, S. Tanaka, K. Ishihara, Physical and biologicalproperties of compound membranes incorporating a copolymer with a phosphorylcholine head group,
Biomaterials 19 (1998) 691–700.
[190] P. Vadgama, Surface biocompatibility, Annu. Rep. Prog. Chem., Sect. C: Phys. Chem. 101 (2005) 14–52.
[191] E. Sackmann, Supported membranes: scientiﬁc and practical applications, Science 271 (1996) 43–48.
[192] I. Reviakine, A. Brisson, Formation of supported phospholipid bilayers from unilamellar vesicles
investigated by atomic force microscopy, Langmuir 16 (2000) 1806–1815.
[193] S. Scheuring, D.J. Müller, P. Ringler, J.B. Heymann, A. Engel, Imaging streptavidin 2D crystals on
biotinylated lipid monolayers at high resolution with the atomic force microscope, J. Microsc. 193 (1999)
28–35.
[194] I. Reviakine, A. Brisson, Streptavidin 2D crystals on supported phospholipid bilayers: toward constructing
anchored phospholipid bilayers, Langmuir 17 (2001) 8293–8299.
[195] M.E. Price, R.M. Cornelius, J.L. Brash, Protein adsorption to polyethylene glycol modiﬁed liposomes fromﬁbrinogen solution and from plasma, Biochim. Biophys. Acta 1512 (2001) 191–205.
[196] A. Eing, A. Janshoﬀ, H.-J. Galla, C. Block, C. Steinem, Quantiﬁcation of the Raf–C1 interaction with solidsupported bilayers, Chem. Biol. Chem. 3 (2002) 190–197.

436

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437[197] A. Raudino, F. Castelli, Interaction of globular proteins with mixed lipid vesicles – a thermodynamic studyof the lipid lateral phase separation, J. Therm. Anal. 37 (1991) 1877–1886.
[198] A. Knoll, R. Magerle, G. Krausch, Tapping mode atomic force microscopy on polymers: where is the true
sample surface? Macromolecules 34 (2001) 4159–4165.
[199] Q. Zhang, O.K.C. Tsui, B. Du, F. Zhang, T. Tang, T. He, Observation of inverted phases in poly(styrene-bbutadiene-b-styrene) triblock copolymer by solvent-induced order-disorder phase transition, Macromolecules 33 (2000) 9561–9567.
[200] D. Raghavan, M. VanLandingham, X. Gu, T. Nguyen, Characterization of heterogeneous regions inpolymer systems using tapping mode and force mode atomic force microscopy, Langmuir 16 (2000) 9448–
9459.
[201] D. Raghavan, X. Gu, T. Nguyen, M. VanLandingham, A. Karim, Mapping polymer heterogeneity
using atomic force microscopy phase imaging and nanoscale indentation, Macromolecules 33 (2000)
2573–2583.
[202] A. Noy, C.H. Sanders, D.V. Vezenov, S.S. Wong, C.M. Lieber, Chemically-sensitive imaging in tapping
mode by chemical force microscopy: relationship between phase lag and adhesion, Langmuir 14 (1998)
1508–1511.
[203] B.B. Sauer, R.S. McLean, R.R. Thomas, Tapping mode AFM studies of nano-phases on ﬂuorinecontaining polyester coatings and octadecyltrichlorosilane monolayers, Langmuir 14 (1998) 3045–3051.
[204] R.W. Stark, G. Schitter, A. Stemmer, Tuning the interaction forces in tapping mode atomic force
microscopy, Phys. Rev. B 68 (2003) 85401.
[205] X. Chen, C.J. Roberts, J. Zhang, M.C. Davies, S.J.B. Tendler, Phase contrast and attraction–repulsion
transition in tapping mode atomic force microscopy, Surf. Sci. 519 (2002) 593–598.
[206] M. Stark, C. Möller, D.J. Müller, R. Guckenberger, From images to interactions: high-resolution phaseimaging in tapping-mode atomic force microscopy, Biophys. J. 80 (2001) 3009–3018.
[207] J. Legleiter, T. Kowalewski, Insights into ﬂuid tapping-mode atomic force microscopy provided bynumerical simulations, Appl. Phys. Lett. 87 (2005) 163120.
[208] K. Saito, T. Aoki, T. Aoki, T. Yanagida, Movement of single myosin ﬁlaments and myosin step size on an
actin ﬁlament suspended in solution by a laser trap, Biophys. J. 66 (1994) 769–777.
[209] T. Fukuma, J.I. Kilpatrick, S.P. Jarvis, Phase modulation atomic force microscope with true atomic
resolution, Rev. Sci. Instrum. 77 (2006) 123703.
[210] J. Schiener, S. Witt, M. Hayer-Hartl, R. Guckenberger, How to orient the functional GroEL-SR1 mutant
for atomic force microscopy investigations, Biochem. Biophys. Res. Commun. 328 (2005) 477–483.
[211] J.E. Sader, S.P. Jarvis, Accurate formulas for interaction force and energy in frequency modulation force
spectroscopy, Appl. Phys. Lett. 84 (2004) 1801–1803.
[212] L. Herbette, C.A. Napolitano, R.V. McDaniel, Direct determination of the calcium proﬁle structure for
dipalmitoyllecithin multilayers using neutron diﬀraction, Biophys. J. 46 (1984) 677–685.
[213] C. Altenbach, J. Seelig, Ca2+ binding to phosphatidylcholine bilayers as studied by deuterium magnetic
resonance. Evidence for the formation of a Ca2+ complex with two phospholipid molecules, Biochemistry23 (1984) 3913–3920.
[214] T.R. Hermann, A.R. Jayaweera, A.E. Shamoo, Interaction of europium(III) with phospholipid vesicles asmonitored by laser-excited europium(III) luminescence, Biochemistry 25 (1986) 5834–5838.
[215] H. Binder, O. Zschörnig, The eﬀect of metal cations on the phase behavior and hydration characteristics ofphospholipid membranes, Chem. Phys. Lipids 115 (2002) 39–61.
[216] M.L. Berkowitz, D.L. Bostick, S. Pandit, Aqueous solutions next to phospholipid membrane surfaces:
insights from simulations, Chem. Rev. 106 (2006) 1527–1539.
[217] S. Garcia-Manyes, G. Oncins, F. Sanz, Eﬀect of ion-binding and chemical phospholipid structure on thenanomechanics of lipid bilayers studied by force spectroscopy, Biophys. J. 89 (2005) 1812–1826.
[218] R.A. Böckmann, A. Hac, T. Heimburg, H. Grubmüller, Eﬀect of sodium chloride on a lipid bilayer,
Biophys. J. 85 (2003) 1647–1655.
[219] S.A. Pandit, D. Bostick, M.L. Berkowitz, Molecular dynamics simulation of a dipalmitoylphosphatidylcholine bilayer with NaCl, Biophys. J. 84 (2003) 3743–3750.
[220] P.K. Hansma, B. Drake, O. Marti, S.A. Gould, C.B. Prater, The scanning ion-conductance microscope,
Science 243 (1989) 641–643.
[221] L. Ying, A. Bruckbauer, D. Zhou, J. Gorelik, A. Shevchuk, M. Lab, Y. Korchev, D. Klenerman, The
scanned nanopipette: a new tool for high resolution bioimaging and controlled deposition of biomolecules,
Phys. Chem. Chem. Phys. 7 (2005) 2859–2866.

T. Ando et al. / Progress in Surface Science 83 (2008) 337–437

437[222] A.I. Shevchuk, G.I. Frolenkov, D. Sanchez, P.S. James, N. Freedman, M.J. Lab, R. Jones, D. Klenerman,
Y.E. Korchev, Imaging proteins in membranes of living cells by high-resolution scanning ion conductancemicroscopy, Angew. Chem. Int. Ed. Engl. 45 (2006) 2212–2216.
[223] C. Stroh, H. Wang, R. Bash, B. Ashcroft, J. Nelson, H. Gruber, D. Lohr, S.M. Lindsay, P. Hinterdorfer,
Single-molecule recognition imaging microscopy, Proc. Natl. Acad. Sci. USA 101 (2004) 12503–12507.
[224] P. Hinterdorfer, Y.F. Dufrêne, Detection and localization of single molecular recognition events using
atomic force microscopy, Nat. Meth. 3 (2006) 347–355.
[225] Y. Sako, T. Yanagida, Single-molecule visualization in cell biology, Nat. Rev. Mol. Cell Biol. 4 (2003) SS1–SS5.
[226] M.I. Wallace, J.E. Molloy, D.R. Trentham, Combined single-molecule force and ﬂuorescence measurements for biology, J. Biol. 2 (2003) 4.
[227] P.V. Cornish, T. Ha, A survey of single-molecule techniques in chemical biology, ACS Chem. Biol. 2 (2007)
53–61.
[228] R. Kassies, K.O. Van Der Werf, A. Lenferink, C.N. Hunter, J.D. Olsen, V. Subramaniam, C. Otto,
Combined AFM and confocal ﬂuorescence microscope for applications in bio-nanotechnology, J. Microsc.
217 (2005) 109–116.
[229] K. Meller, C. Theiss, Atomic force microscopy and confocal laser scanning microscopy on the cytoskeletonof permeabilised and embedded cells, Ultramicroscopy 106 (2006) 320–325.
[230] T. Kodama, H. Ohtani, H. Arakawa, A. Ikai, Development of confocal laser scanning microscope/atomic
force microscope system for force curve measurement, Jpn. J. Appl. Phys. 43 (2004) 4580–4583.
[231] A. Noy, T.R. Huser, Combined force and photonic probe microscope with single molecule sensitivity, Rev.
Sci. Instrum. 74 (2003) 1217–1221.
[232] D.-C. Kim, R. Kim, H.-J. Kim, J. Kim, D.-H. Park, H.-S. Kim, J. Joo, Raman study of polymer–metal
hybrid nanotubes using atomic force/confocal combined microscope, Jpn. J. Appl. Phys. 46 (2007) 5556–
5559.
[233] M.S. Anderson, W.T. Pike, A Raman-atomic force microscope for apertureless-near-ﬁeld spectroscopy andoptical trapping, Rev. Sci. Instrum. 73 (2002) 1198–1203.
[234] T. Ichimura, N. Hayazawa, M. Hashimoto, Y. Inouye, S. Kawata, Application of tip-enhanced microscopy
for nonlinear Raman spectroscopy, Appl. Phys. Lett. 84 (2004) 1768–1770.
[235] T. Ichimura, N. Hayazawa, M. Hashimoto, Y. Inouye, S. Kawata, Tip-enhanced coherent anti-stokes
Raman scattering for vibrational nanoimaging, Phys. Rev. Lett. 92 (2004) 220801.
[236] M.S.Z. Kellermayer, Á Karsai, A. Kengyel, A. Nagy, P. Bianco, T. Huber, Á Kulcsár, C. Niedetzky, R.
Proksch, L. Grama, Spatially and temporally synchronized atomic force and total internal reﬂection
ﬂuorescence microscopy for imaging and manipulating cells and biomolecules, Biophys. J. 91 (2006) 2665–
2677.
[237] L. Peng, B.J. Stephens, K. Bonin, R. Cubicciotti, M. Guthold, A combined atomic force/ﬂuorescencemicroscopy technique to select aptamers in a single cycle from a small pool of random oligonucleotides,
Microsc. Res. Techniq. 70 (2007) 372–381.
[238] T. Yamada, R. Afrin, H. Arakawa, A. Ikai, High sensitivity detection of protein molecules picked up on aprobe of atomic force microscope based on the ﬂuorescence detection by a total internal reﬂection
ﬂuorescence microscope, FEBS Lett. 569 (2004) 59–64.
[239] A. Aydinli, A. Serpengüzel, D. Vardar, Visible photoluminescence from low temperature depositedhydrogenated amorphous silicon nitride, Solid State Commun. 98 (1996) 273–277.
[240] V. Lulevich, C. Honig, W.A. Ducker, An atomic force microscope tip as a light source, Rev. Sci. Instrum.
76 (2005) 123704.
[241] D.J. Wolford, B.A. Scoot, J.A. Reimer, J.A. Bradley, Eﬃcient visible luminescence from hydrogenatedamorphous silicon, Physica B 117–118 (1982) 920–922.
[242] F.L. Degertekin, A.G. Onaran, M. Balantekin, W. Lee, N.A. Hall, C.F. Quate, Sensor for direct
measurement of interaction forces in probe microscopy, Appl. Phys. Lett. 87 (2005) 213109.
[243] A.G. Onaran, M. Balantekin, W. Lee, W.L. Hughes, B.A. Buchine, R.O. Guldiken, Z. Parlak, C.F. Quate,
F.L. Degertekin, A new atomic force microscope probe with force sensing integrated readout, active tip,
Rev. Sci. Instrum. 77 (2006) 023501.