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From: Daisuke Yamamoto, Takayuki Uchihashi, Noriyuki Kodera, Hayato
Yamashita, Shingo Nishikori, Teru Ogura, Mikihiro Shibata, and Toshio Ando,
High-Speed Atomic Force Microscopy Techniques for Observing Dynamic
Biomolecular Processes In Nils G. Walter editor:
METHODS IN ENZYMOLOGY, Vol. 475,
Burlington: Academic Press, 2010, pp.541-564.
ISBN: 978-0-12-381482-1© Copyright 2010, Elsevier Inc.
Academic Press.

Author's personal copy
C H A P T E R

T W E N T Y

High-Speed Atomic ForceMicroscopy Techniques for
Observing Dynamic Biomolecular
Processes
Daisuke Yamamoto,*,† Takayuki Uchihashi,*,† Noriyuki Kodera,*,†Hayato Yamashita,* Shingo Nishikori,‡ Teru Ogura,†,‡Mikihiro Shibata,* and Toshio Ando*,†Contents
1. Introduction
2. Survey of Requirements for High-Speed Bio-AFM Imaging
3. Substrate Surfaces
3.1. Bare mica surface
3.2. Mica surface-supported planar lipid bilayers
3.3. Streptavidin 2D crystals as substrates
4. Control of Diffusional Mobility
5. Protein 2D Crystals as Targets to Study
5.1. Streptavidin 2D crystals
5.2. p97 2D crystals
5.3. Bacteriorhodopsin 2D crystals
6. Low-Invasive Imaging
7. UV Flash-Photolysis of Caged Compounds
8. Cantilever Tip
References

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Abstract
Atomic force microscopy (AFM) enables direct visualization of single-protein
molecules in liquids at submolecular resolution. High-speed AFM further makes
it possible to visualize dynamic biomolecular processes at subsecond* Department of Physics, Kanazawa University, Kakuma-machi, Kanazawa, Japan
Core Research for Evolutional Science and Technology (CREST) of the Japan Science and Technology
Agency, Sanban-cho, Chiyoda-ku, Tokyo, Japan{
Department of Molecular Cell Biology, Institute of Molecular Embryology and Genetics, Kumamoto
University, Kumamoto, Japan{

Methods in Enzymology, Volume 475
ISSN 0076-6879, DOI: 10.1016/S0076-6879(10)75020-5# 2010 Elsevier Inc.

All rights reserved.

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resolution. However, dynamic imaging of biomolecular processes imposes various requirements on ‘‘wet techniques’’ and imaging conditions, which are often
different from those for static imaging. This chapter first surveys the imposed
requirements, then focuses on practical techniques associated with dynamic
imaging, highlighting the preparation of substrate surfaces, and presents
examples of the use of these techniques.

1. Introduction
Single-molecule visualization of functional behavior of proteins has
been performed using fluorescence microscopy (e.g., Goldman, 2009; Jooet al., 2008; Park et al., 2007). This approach has enriched our knowledge ofdynamic processes of proteins. However, even using fluorescence microscopy tools with a spatial resolution beyond the diffraction limit (Huang
et al., 2009), it is impossible to observe protein molecules themselves since
only single featureless fluorescent spots are observable. Therefore, we needto acquire additional information to bridge the gap between the behavior offluorescent spots and the actual behavior of labeled protein molecules.
Atomic force microscopy (AFM) allows for the direct observation ofindividual protein molecules in solutions, at submolecular resolution.
However, its imaging rate is too low to trace their dynamic behaviors.
Rapidly moving molecules cannot be imaged at all. In the last decade,significant efforts have been carried out to increase the imaging rate of
biological AFM (bio-AFM) (e.g., Ando et al., 2001, 2008; Fantner et al.,2006; Viani et al., 1999). Only increasing the scan speed of AFM would
not be difficult, if we ignore potential damage to the sample or disturbanceof biomolecular processes. Very precise and fast control of the tip-sample
distance is mandatory for observing delicate dynamic biomolecular processes. To achieve high-speed and low-invasive AFM imaging, various
techniques have been devised (e.g., Kodera et al., 2005, 2006). As a result,some dynamic events and structural changes in proteins have been successfully visualized (Ando et al., 2006; Miyagi et al., 2008; Shibata et al.,2010; Yamamoto et al., 2008; Yamashita et al., 2009). As comprehensive
descriptions for realizing the instrumentation are already given elsewhere(Ando et al., 2008), we focus here on other critical aspects of dynamic
AFM imaging.
Static AFM imaging of biological samples has been performedby many researchers, and various methods for this imaging have accumulated.
On the other hand, dynamic AFM imaging of biological samples is rather
new, and hence, methods and scan conditions specifically required

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High-Speed Atomic Force Microscopy Techniques543

for dynamic imaging are not yet found in the literature. Here, we introduce
methods useful for dynamic AFM imaging, highlighting the preparation ofsubstrate surfaces.

2. Survey of Requirements for High-Speed
Bio-AFM Imaging
A substrate surface, on which a sample is placed, plays an important rolein AFM imaging. It should be flat enough so that we can easily identify themolecules of interest deposited on it. To observe dynamically acting protein
molecules under physiological conditions, the substrate surface must not
firmly bind the molecules so that their physiological function is retained.
However, the sample–surface interaction is essential to avoid too fast Brownian motion, particularly for single-protein molecules isolated completelyfrom other molecules. Dynamic AFM imaging requires much more definedconditions for the substrate surface than static AFM imaging does.
We also have to consider requirements for observing dynamic protein–protein interactions. Supposing that one species of protein dynamicallyinteracts with a protein counterpart, only one but not the other can be
immobilized to a substrate surface. If both are surface bound, they havealmost no chance to interact with each other. Consequently, selective
protein attachment to a surface is required. A dynamic protein–protein
interaction implies that their association is weak. Tapping mode AFM issuitable for imaging fragile samples because little lateral force is exerted tothe sample, while the tip taps the sample. This tapping should not disturb
delicate protein–protein interactions. Ideally, we have to achieve a very
weak tapping force without slowing the imaging speed.
Protein molecules exhibit Brownian motion alongside physiologically
relevant dynamic conformational changes. In many cases, the purpose ofobserving the dynamic behavior of proteins is to detect physiologically
relevant changes in protein conformation. We therefore have to be able
to distinguish the two types of movement. Special means sometimes have tobe implemented for this distinction, as well as a means to impede Brownian
motion.
Biological membranes are very fluidic. When membrane proteins are
not assembled into a large ensemble or anchored to an immobile entity, theydiffuse rapidly in a membrane, so that there is no way to observe theirstructure even using high-speed AFM with the capacity to image at
33 frames/s for a scan range of 240  240 nm2 (Yamashita et al., 2007).
In this case, we need some means to slow down diffusion.
In the following sections, several solutions for the difficulties mentioned
above are presented and their effects are exemplified.

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3. Substrate Surfaces
For AFM imaging, bare surfaces of mica and highly oriented polygraphite (HOPG), as well as gold-coated surfaces of these substrates orsilicon have often been used. In aqueous solution, proteins do not attach
to HOPG except for rare cases. Gold-coated surfaces are typically used for
preparing self-assembled monolayers of molecules containing a thiol. Proteins can be immobilized to these monolayers when the constituting
monolayer molecules also possess a reactive or functional group. Since thepreparation methods and characterization of these monolayers are welldescribed elsewhere (Hinterdorfer and Dufrêne, 2006; Liu et al., 2008),
we here only cover bare mica surfaces, mica surface-supported planar lipid
bilayers, and streptavidin 2D crystals formed on lipid bilayers.

3.1. Bare mica surface
Mica (natural muscovite or synthetic fluorophlogopite) has frequently beenused as a substrate owing to its surface flatness at the atomic level over a large
area. It has a net negative charge and is therefore quite hydrophilic. A baremica surface adsorbs various proteins by electrostatic interactions. Except inrare cases (e.g., GroEL attachment in an end-up orientation), the orientation of adsorbed proteins is not unique, and the selective attachment of a
specific species of protein is not expected. A mica surface is useful forobserving the dynamic processes of a single species of isolated protein or
membrane proteins in native membrane fragments. We can control theaffinity for a specific protein by varying the ionic strength, pH, or by adding
divalent cations such as Mg2þ. Divalent cations can link negatively charged
proteins to a negatively charged mica surface.
Mica disks are prepared from a mica sheet with thickness of <0.1 mm by
cutting holes using a sharp puncher. Serrated edge formation, which often
accompanies partial cleavage of interlayer contacts in the disk, should be
avoided. Hydrodynamic pressure produced by rapid scanning of the sample
stage induces vibrations of the disk through movement of the cleaved sites.
For high-speed imaging, the disk should be small (1–2 mm in diameter) to
avoid generation of too large a hydrodynamic pressure (Ando et al., 2002).
The following handling protocol is recommended:
1. Glue a mica disk to a sample stage using epoxy and wait until it has dried(1–2 h).
2. Press a Scotch tape to the surface of the mica disk and then smoothly
remove the tape from the mica. The top layer of the mica will be
removed with the tape, which can be checked by inspecting the surfaceof the removed tape.

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3. Place a sample solution on the freshly cleaved mica disk surface for
1–3 min, and then rinse with an appropriate buffer solution.
4. Judge the affinity of the sample for the mica surface by AFM imaging.
5. Find an appropriate solution condition by repeated imaging using different solutions (an example is shown in Fig. 20.1 for myosin V). Whenthe affinity is too weak, reduce the ionic strength or pH, or change theconcentration of divalent cations.

A

B

C 400

600 mMy (nm)

300

1

200

300 mM100

1

0

100 mM

02

2

Root-mean-squaredisplacement (nm)

D
300

3

200 300x (nm)

400

500(from top to bottom lines)
600 mM
500 mM
300 mM
200 mM
100 mM

200
100
0

3

100

0

0.2

0.4

0.6

Time (s)Figure 20.1 Ionic strength dependent diffusion of MyoV-HMM (tail-truncated myosin V) on bare mica surface. (A, B) Typical successive AFM images taken at 100 mM(A) and 600 mM KCl (B). Imaging rate: 102 ms/frame, scan area: 300  300 nm2 with100  100 pixels, scale bar: 100 nm. The number in each frame represents theframe number. The observation buffer is 20 mM imidazole–HCl (pH 7.6), 2 mM
MgCl2, 1 mM EGTA, and 100 or 600 mM KCl. (C) Typical trajectories of MyoVHMM measured in 100, 300, or 600 mM KCl-containing solutions. (D) Root-meansquare displacement (xrms) as a function of time. Each point was calculated
pﬃﬃﬃﬃﬃﬃﬃﬃ fromthe trajectories of 100 molecules and well fitted by an equation xrms ¼ 4Dt , where
D is the diffusion coefficient and t is the time elapse. The values of D (nm2/s) at respective
D200
KClconcentrationsare
D100
mM ¼ 79  10,mM ¼ 213  26,
D300 mM ¼ (0.99  0.01)  104, D500 mM ¼ (1.69  0.04)  104, and D600 mM ¼
(2.73  0.09)  104.

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3.2. Mica surface-supported planar lipid bilayers
Mica surface-supported planar lipid bilayers are useful for attaching proteinsto their surfaces with a controlled affinity or selectivity. A membrane surface
with zwitterionic polar head groups such as phosphatidyl choline (PC) andphosphatidyl ethanolamine (PE) is known to resist protein adsorption(Vadgama, 2005; Zhang et al., 1998). Various lipids with functional groups(e.g., biotin and Ni-NTA) on their polar heads are commercially available.
They enable specific attachment of biotin- or His-tag-conjugated proteinsto planar lipid bilayers. For electrostatic attachment, negatively charged
head groups including phosphatidylserine (PS), phosphoric acid (PA), or
phosphatidylglycerol (PG) and positively charged head groups such as
trimethylammoniumpropane (TAP) and ethylphosphatidylcholine (EPC)
are available. Unlike mica surfaces, the surface charge density can easily becontrolled by mixing charged and neutral lipids in different ratios as long as
no phase separation occurs.
3.2.1. Choice of alkyl chains
Dioleoyl-phosphatidyl-choline (DOPC) is useful for the preparation of 2D
crystals of streptavidin and His-tag-labeled proteins when it is used inconjunction with biotinylated lipids and Ni-NTA-containing lipids,
respectively (Reviakine and Brisson, 2001; Scheuring et al., 1999). DOPC
contains an unsaturated hydrocarbon in each of the two alkyl chains, whichcauses the bending of the chains and therefore weakens the interaction
between neighboring DOPCs. This weak interaction lowers the phasetransition temperature of DOPC to 20  C, thereby affording considerable fluidity to the planar bilayer at room temperature and facilitating theformation of 2D protein crystals. Such densely packed proteins do not
diffuse easily. Dipalmitoyl-phosphatidyl-choline (DPPC) contains no
unsaturated hydrocarbons in the alkyl chains. Therefore, its phase-transition
temperature is high (41  C) and it is suitable for preparing planar lipid
bilayers with low fluidity. For example, when planar bilayers are formed
using DPPC at high temperature (60  C) together with a suitable fractionof DPPE-biotin, streptavidin sparsely attached to the surface hardly diffusesat room temperature. When DOPE-biotin is used together with DPPC, thesparsely attached streptavidin diffuses at a moderate rate (Fig. 20.2).
3.2.2. Preparation of planar lipid bilayers
For the preparation of planar lipid bilayers on a mica surface, two methods
are commonly used; the Langmuir–Blodgett method and the vesicle fusion
method. In both cases, uniform and smooth surfaces of lipid bilayers can beobtained. Here, we focus on the latter method, because of its simplicity andlack of a requirement for any expensive instruments and skills. Supported

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B
100
5

1

5

1

4
3

1

4
3

2

0 ms

5
4

3

2

154.5 ms

2

1
3

0

309 ms

0

D
100

3
4
0 ms

7

2
5 8
6

7

1
3
4

176 ms

2
58

7

1
3

4

58

352 ms

3

2

4 5

7

8

6

6

6

100

x (nm)1y (nm)

2

5

4

C
1

2

y (nm)A00x (nm)

100

Figure 20.2 Mobility of lipid molecules monitored by diffusion of attached streptavidin molecules and its dependence on lipid compositions. Successive AFM images of
streptavidin on lipid bilayers of DPPC þ DPPE-biotin (9:1, w/w) and DPPC þ
DOPE-biotin (9:1, w/w) are shown in (A) and (C), respectively. Imaging rate:
154.5 ms/frame (A), 176 ms/frame (C), scan area, 200  200 nm2 with 100  100
pixels, scale bar: 40 nm. The apparent difference in the size of streptavidin molecules
between (A) and (C) is due to different cantilever tips. Trajectories of streptavidin
molecules observed in (A) for  15.6 s and (C) for  4.6 s are plotted in (B) and (D),respectively. The numbers beside trajectories in (B) and (D) correspond to the numbered streptavidin molecules in (A) and (C), respectively.

planar lipid bilayers are formed from small unilamellar vesicles by direct
deposition of the solution on a freshly cleaved mica surface. Planar lipid
bilayer formation proceeds through: (i) adsorption of small unilamellar
vesicles on the mica surface, (ii) rupture of the vesicles, and (iii) fusion of
the resulting small membrane patches with juxtaposed lipid membranes.
Although protocols for preparing lipid vesicles and supported lipid bilayers
can be found elsewhere, we describe here a procedure routinely used in ourlaboratory.
1. Dissolve each lipid compound in chloroform or in a mixture of chloroform, methanol, and water (follow the instructions provided by the
manufacturers).
2. Mix lipid solutions at a desired ratio in a glass test tube.
3. Dry the organic solvent under a stream of N2 gas. During this procedure,warm up the test tube by gripping it with your hand to facilitate solvent
evaporation.

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4. To ensure full evaporation, leave the test tube in a desiccator underaspirator vacuum for more than 30 min.
5. Add a buffer solution to the test tube (typical final concentration oflipids, 0.125 mM) and vortex it. If necessary, sonicate the test tube using
an ultrasonic bath to disperse the lipids in the medium. At this stage,
multilamellar lipid vesicles are formed. The lipid suspension is divided
into small aliquots (100 ml) and stored at 80  C.
6. To obtain small unilamellar vesicles, sonicate the multilamellar vesicle
suspension (typically 100 ml) with a tip sonicator at intervals of 1 s with aduty ratio of  0.5 until the suspension becomes transparent (typically nomore than 30 cycles).
7. Place a drop of the small unilamellar vesicle solution on a freshly cleaved
mica surface and incubate it for 30 min in a sealed container, while
maintaining high humidity in the container to avoid sample drying by
stuffing therein a piece of paper damped with water. Subsequently, rinse
the sample with an appropriate buffer solution.
8. When the gel–liquid crystal transition temperature of the sample ishigher than room temperature, warm up the sample in the sealedcontainer slightly above the transition temperature for 15 min.
9. Examples for lipid compositions (buffer conditions) are given below.
The mixing ratios of lipids should be changed depending on the samples
and dynamic events to be visualized.
 highly fluidic bilayers with biotin; DOPC:DOPS:biotin-capDOPE ¼ 7:2:1 (w/w) [10 mM HEPES–NaOH (pH 7.4), 150 mM
NaCl and 2 mM CaCl2];
 low fluidic bilayers with biotin; DPPC:biotin-cap-DPPE ¼ 9:1 (w/w)[10 mM HEPES–NaOH (pH 7.4), 150 mM NaCl and 2 mM CaCl2];
 slightly fluid bilayers with biotin; DPPC:biotin-cap-DOPE ¼ 9:1(w/w) [10 mM HEPES–NaOH (pH 7.4), 150 mM NaCl and 2 mM
CaCl2];
 highly fluidic bilayers with Ni-NTA; DOPC:DOPS:DOGS-NTA(Ni) ¼ 7:2:1 (w/w) [10 mM HEPES–NaOH (pH 7.4), 150 mM
NaCl and 2 mM CaCl2];
 low fluidic bilayers with Ni-NTA; DPPC:DOGS-NTA(Ni) ¼ 9:1(w/w) [50 mM Tris–HCl (pH 8.0), 50 mM KCl and 3 mM MgCl2];
 low fluidic bilayers with positively charged head groups; DPPC:DPTAP ¼ 7:3 (w/w) or only DPTAP (distilled water); low fluidic bilayers with negatively charged head groups; DPPG
100% [10 mM HEPES–NaOH (pH 7.4), 150 mM NaCl and 2 mM
CaCl2];
 low fluidic bilayers with negatively charged head groups formed on
positively charged lipid bilayers; DPPA 100% (distilled water).

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3.2.3. Planar lipid bilayers as substrates
3.2.3.1. Electrostatic immobilization Since the mica surface is negatively
charged, positively charged lipid bilayers can easily be formed on thesurface. Depending on the sample to be imaged, an appropriate positivecharge density has to be found by repeated imaging with various positive toneutral lipid ratios (examples are shown in Fig. 20.3A–C). Negatively
charged lipid bilayers can be formed in a wide area on a mica surface
A

B

186

C

86

187
87

D
Avidin DPPA

188
88

DPPC:DPTAP = 7:3

Mica

125

126

127

Figure 20.3 Electrostatic adsorption of proteins on charged lipid bilayers. (A) AFM
image of actin filament paracrystal formed on positively charged lipid bilayers (DPPC:DPTAP ¼ 7:3, w/w). The white arrowhead indicates an exposed mica surface. Scalebar, 100 nm. (B, C) High-speed AFM images showing the effect of positive-charge
surface density on the mobility of attached MyoV-HMM. The lipid compositions are(B) DPPC:DPTAP ¼ 7:3 (w/w) and (C) only DPTAP. The leftmost AFM images show
the surface of lipid bilayers before adding MyoV-HMM. Imaging rate, 101 ms/frame;
scale bar, 100 nm. The number in each frame represents the frame number. MyoVHMM was hardly visible (B) due to the large mobility, whereas it was visible (C) due tothe low mobility. (D) AFM images of negatively charged lipid bilayers formed on
positively charged lipid bilayers. The leftmost AFM image was taken before adding
avidin (a positively charged protein). The white arrowhead, diamond, and asterisk
represent the surfaces of mica, positively charged lipid bilayers, and negatively charged
lipid bilayers, respectively (imaging rate: 418 ms/frame, scale bar: 100 nm).
The remaining successive AFM images were taken after adding avidin (imaging rate:
175 ms/frame, scale bar: 100 nm). Avidin molecules are only observed on thenegatively charged surfaces of mica and DPPA (see the upper schematic). The observation buffer contains 20 mM imidazole–HCl (pH 7.6), 25 mM KCl, 2 mM MgCl2, and
1 mM EGTA.

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using DPPG (transition temperature, Tm, 41  C). Since the negative chargeof DPPG is not positioned at the distal end of the polar head, the bilayer
surface weakly adsorbs positively charged proteins. Thus far, negatively
charged lipid bilayers have not been formed on a mica surface using
DPPA (Tm ¼ 67  C; the negative charge is positioned at the distal end of
the polar head) or DPPS (Tm ¼ 54  C). Another means is to form negatively charged lipid bilayers on positively charged bilayers. Although notcompletely successful yet, at least small patches of DPPA bilayers can beformed by this means (Fig. 20.3D).
3.2.3.2. Immobilization via specific interaction Specific association pairs
with strong affinities, such as (biotin and streptavidin) and (His-tag and NiNTA), can be used for selective immobilization of proteins. When relatively
large molecules mediate the immobilization of a protein on lipid bilayers, theattached mediator would make it difficult to identify the immobilized protein. When streptavidin is used as a mediator, the use of streptavidin 2D
crystals is recommended (see Section 3.3). Here, we show only an example ofHis-tag-conjugated protein immobilization. p97 is a hexameric ATPase of
the AAA family and involved in cellular processes (more descriptions of p97
are given later). As shown in Fig. 20.4, recombinant p97 of which each
subunit was conjugated with a His-tag was immobilized on a Ni-NTAcontaining lipid bilayer. p97 was attached to the bilayer surface in a specificorientation due to the specific-site conjugation of a His-tag to each subunit.

3.3. Streptavidin 2D crystals as substrates
2D crystals of streptavidin formed on biotin-containing lipid bilayers meetvarious requirements for substrate surfaces used in dynamic AFM imaging.
Streptavidin is comprised of four identical subunits, each of which specifically

100 nm

20 nm

Figure 20.4 p97 attached on Ni-NTA-containing lipid bilayer through His-tag conjugated at the N-terminus of each subunit. (A) Segregation of Ni-NTA lipid in bilayer(p97 are densely packed at bright regions but absent in dark regions). (B) Ring structureof p97 (C-terminus D2 domain surface), z-scales: (A) 20 nm, (B) 14 nm.

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binds to one biotin molecule with a strong affinity (Ka  1013 M 1)(Green, 1990). In 2D crystals, two biotin-binding sites face the solution, andtherefore, can bind to biotinylated samples to be imaged (Darst et al., 1991;
Reviakine and Brisson, 2001). Since biotinylated Ni-NTA is commerciallyavailable, the surface can also bind to His-tag conjugated recombinant proteins. The surface roughness of a specific type of 2D crystal is very small, as
mentioned later. Importantly, streptavidin is resistant to nonspecific binding ofmany proteins (Green, 1990), which safeguards the surface-bound proteins
against dysfunction (Heyes et al., 2004) and allows the selective surface attachment of one species of protein in a multicomponent system. In addition, it
allows protein attachment in a controlled orientation through the biotin or
His-tag conjugation sites in the protein.
3.3.1. Preparation of streptavidin 2D crystals
On highly fluidic planar lipid layers containing a biotinylated lipid, streptavidin self-assembles into three distinct crystalline arrangements of C222, P2,
and P1 symmetries, depending on the crystallization conditions such as pH(Wang et al., 1999) and ionic strength (Ratanabanangkoon and Gast, 2003),
as described in the following. Note that all the 2D crystals have P2 symmetry (Yamamoto et al., 2009) but we followed previous reports in thenotation for the symmetry groups.
1. Crystallization buffer solutions: (C222 crystal) 10 mM HEPES–NaOH(pH 7.4), 150 mM NaCl, and 2 mM CaCl2; (P2 crystal) 10 mM MES–NaOH (pH 5.8), 450 mM NaCl, and 2 mM CaCl2; (P1 crystal) 10 mM
acetate–NaOH (pH 4.0), 450 mM NaCl, and 2 mM CaCl2.
2. Prepare the mica-supported lipid planar bilayers containing biotin as
mentioned in Section 3.2.2.
3. After washing the lipid planar bilayers with a crystallization buffer
solution, place 0.1–0.2 mg/ml streptavidin in the crystallization buffer
solution on the lipid bilayer and incubate at room temperature in a sealedcontainer (keeping 100% relative humidity is a key to the successful
crystallization with fewer defects).
4. After incubating at least for 2 h, wash out excess streptavidin with the
crystallization buffer solution used.
5. If necessary, chemically fix the streptavidin crystals by applying a 10 mM
glutaraldehyde-containing crystallization solution and incubate for5 min. Then, quench the reaction using 20 mM Tris–HCl mixed in
the crystallization buffer.
Figure 20.5 (top) shows high-magnification AFM images of the three
types of streptavidin crystals. Among the three, the P1 crystal shows thesmallest surface roughness (Fig. 20.6C, bottom; RMS roughness 0.14 nm)
and the highest stability (Yamamoto et al., 2009). Even when tapped with an
oscillating cantilever with relatively strong force, the crystal structure isAuthor's personal copy
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A

B

C

20 nm

20 nm

20 nm

10 nm

1 nm

1 nm

1 nm

10 nm

10 nm

Figure 20.5 High-magnification AFM images of the three types of streptavidin crystals (Yamamoto et al., 2009). The space group symmetries of the crystals are (A) C222,(B) P2, and (C) P1. Top: AFM images, according to previous reports (P2 is a correct
notation for all the crystals). Bottom: Surface profile along the lines indicated in the AFM
images in top. The AFM images were obtained at an imaging rate of 1 s/frame with256  256 pixels.

unchanged, and hence, no chemical fixation is necessary. However, the P1
crystal often shows line cracks. The C222 crystal shows high uniformity
over a wide area but similarly to the P2 crystal, its stability is lower than thatof the P1 crystal, and therefore, chemical fixation is sometimes necessary. Its
chemical fixation does not significantly affect the affinity of streptavidin forbiotin.
3.3.2. Use of streptavidin 2D crystals as substrates
Supposing that a protein to be imaged is relatively rigid and does not possess
flexible chains, its biotinylated species immobilized on a streptavidin 2D
crystal surface would not exhibit rapid Brownian motion even when only
one biotin per molecule is introduced. If it rapidly rotates around thebiotinylated site due to a flexible linker contained in the biotin compound
used, we can use reactive dibiotin compounds for less mobility. However,when a protein to be imaged contains a flexible moiety and is biotinylated
only at one site, it certainly exhibits too rapid Brownian motion to be
imaged (examples are shown in Fig. 20.6 for calmodulin (CaM) and tailtruncated myosin V (MyoV-HMM)). This rapid motion can be stopped byintroducing biotin to the protein at more than two sites (Fig. 20.6C). Thus,
the streptavidin 2D crystals are excellent substrates for oligomerized proteinsas exemplified below but their usefulness is limited for monomeric proteins
with flexible polypeptide chains.
We can specifically place a protein on streptavidin 2D crystals in a
desired orientation when the protein is biotinylated at designed sites. For
example, the D490C GroEL mutant, in which biotin is conjugated to theAuthor's personal copy
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A

79

80

81

1927

1928

1929

B

C
95

96

97

Figure 20.6 Biotinylated proteins attached onto streptavidin 2D crystal surfaces.
(A) Successive AFM images of biotinylated calmodulin (CaM) attached on P1 crystalof streptavidin. The images do not reveal a typical dumbbell-like appearance of CaM
because one of the two globes connected with a flexible central linker is not biotinylated. Scale bar, 20 nm; imaging rate: 991 ms/frame, scan area: 100  100 nm2 with256  256 pixel, z-scale: 4.5 nm. The observation buffer is 20 mM imidazole–HCl (pH7.6), 100 mM KCl, 2 mM MgCl2, 1 mM EGTA. (B, C) Effect of the number ofconjugated biotin per molecule on AFM image of MyoV-HMM immobilized on
streptavidin 2D crystal. MyoV-HMM-containing biotinylated CaM was immobilizedon the C222 crystal of streptavidin. Scale bar: 20 nm, imaging rate: 487 ms/frame, scanarea: 100  100 nm2 with 128  128 pixels, z-scale: 8.0 nm. The observation buffer
was 20 mM Tris–HCl (pH 6.8), 100 mM KCl, 2 mM MgCl2, 1 mM EGTA, and 5 mM
DTT. (B) MyoV-HMM having biotinylated CaM only at one of the two necks. It does
not show the typical molecular shape because the unbiotinylated head moves rapidly
and even the biotinylated head rotates rapidly around the biotin-linker. (C) MyoVHMM with biotinylated CaM at both necks.

Cys-490 located at the outer surface of the equatorial domain, can attach to
the C222 crystal surface in a side-on orientation (Fig. 20.7) (Yamamoto
et al., 2009). In this orientation, both rings of GroEL are accessible to GroES
floating in bulk solution. GroES never attaches to the crystal surface even

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0s

6s

*

8s

10 s

*
*

50 nm

Figure 20.7 AFM images of the GroEL–GroES complex (Yamamoto et al., 2009). The
D490C GroEL mutant was biotinylated at the outer surface of the equatorial domain,
and was immobilized in a side-on orientation on a streptavidin C222 crystal that is
chemically fixed with glutaraldehyde. The GroEL–GroES complex was formed andimaged in the buffer containing 1 mM GroES and 1 mM ADP. The asterisks indicate
GroES bound to one of the GroEL rings. The AFM images were obtained at an imaging
rate of 2 s/frame with 200  200 pixels, z-scale: 19 nm.

when the concentration is increased to the 10 mM range, allowing for theobservation of dynamic events of association/dissociation of the GroEL/
GroES complex. Figure 20.7 (asterisks) shows such a dynamic event in thepresence of ADP.

4. Control of Diffusional Mobility
Proteins that are weakly and sparsely attached to a substrate surface
often exhibit rapid Brownian motion. Even when densely packed on highly
fluidic planar lipid bilayers, some proteins hardly form 2D crystals due to
weak association or repulsion between protein molecules. Noncrystalline
proteins diffuse due to the fluidity of the lipid bilayers, making it difficult tocapture an image. Depending on the situation, we need a specific means toimpede the rapid diffusion, as detailed in the following.
1. Brownian motion of a protein that is attached to a surface by electrostaticinteraction can be impeded by strengthening the interaction using a low
ionic strength solution (Fig. 20.1) or a more charged surface(Fig. 20.3A). This approach is easy and effective for a large number ofsamples as the stronger electrostatic association usually does not hamper
protein function. However, when it does, there are no ideal means toslow down the Brownian motion. One compromise is to place, with anappropriate surface density, small particles on the surface that have arelatively strong affinity for the surface. Diffusion of the protein molecules is slowed down by the contact with the particles. Using this means,
we succeeded to visualize the step process of myosin V (MyoV) along
actin tracks.

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A

B

C

Figure 20.8 (A) An AFM image of the Annexin A-V two-dimensional crystal on lipid
bilayer. The lipid bilayer is composed of 65% DOPC, 34% DOPS, and 1% biotincapped DOPE (w/w). The circular dark areas correspond to the single defects. (B) The
streptavidin molecules are trapped in the large defects of the Annexin crystal andobserved as bright spots. (C) Streptavidin molecules rapidly diffuse on the lipid bilayer
without Annexin and therefore they were observed as spike noises. For all images, thescan area is 200  200 nm and the buffer solution is 10 mM HEPES–NaOH (pH 6.8),
20 mM NaCl, 150 mM KOAc, 2 mM MgCl2, and 2 mM CaCl2.

2. When a protein hardly forms 2D crystals on a highly fluidic lipid bilayer,change the pH to reduce the electrostatic repulsion (an example isshown below), or place the noncrystalline protein on low fluidic lipid
bilayers as exemplified in Fig. 20.2.
3. For membrane proteins embedded in native lipid membranes, theirrapid diffusion sometimes can be impeded using Annexin (Ichikawa
et al., 2006). Annexins are a protein family that bind to negatively
charged phospholipids in the presence of calcium and assemble as trimers(Richter et al., 2005). Annexin A-V forms 2D crystals on PS-containing
lipid membranes (Fig. 20.8A). Using this property, Annexin A-V can beused to trap membranes’ proteins in the defect sites of an Annexin 2D
crystal. Although not native membrane proteins, streptavidin molecules
are demonstrated to be trapped in the defects of the Annexin A-V crystal
formed on very fluidic lipid bilayers (Fig. 20.8B). Without Annexin
A-V, no image of streptavidin molecules can be taken due to rapid
diffusion (Fig. 20.8C).

5. Protein 2D Crystals as Targets to Study
5.1. Streptavidin 2D crystals
As mentioned in Section 3.2.3, the C222 and P2 crystals of streptavidin are
not stable. Monovacancy defects are formed by increasing the tapping force
exerted from an oscillating tip. The defects in the C222 crystal move in theAuthor's personal copy
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A

17.0 s

21.0 s

26.0 s

B
150

30 nm
30.5 s36.0 s44.5 sY (nm)

120
90
60
30
b

0
a

C

D

0s

2s

0

4s

30

60 90
X (nm)

120 150

6s

b
a

40 nm

Figure 20.9 Diffusion of monovacancy defects in streptavidin C222 crystal observedby high-speed AFM (Yamamoto et al., 2008). (A) High-speed AFM images of thestreptavidin 2D crystal and monovacancy defects migrating in the crystal. The monovacancy defects in the crystal are enclosed by the dashed squares and circles. The arrows
indicate the direction of the lattice vectors of the crystal. Successive images wereobtained at an imaging rate of 0.5 s/frame with 200  200 pixels. (B) Trajectories ofthe monovacancy defects obtained from the successive images (A). Closed squares andcircles correspond to defects enclosed by the open squares and circles in (A), respectively. (C) Schematic of streptavidin C222 crystal. The closed circles indicate biotinbound subunits of streptavidin in the crystal, while the open circles correspond tobiotin-unbound subunits. (D) Fusion of monovacancy defects in the C222 crystal.
Two monovacancy defects migrate independently in the crystal at 0–2 s, and thenfused to form a divacancy defect at 4 s. The resultant divacancy defect continued tomigrate in the crystal (4–6 s). The successive images were obtained at an imaging rate of
0.5 s/frame with 200  200 pixels.

plane, and this diffusion exhibits anisotropy correlated with the two crystallographic axes in the orthorhombic C222 crystal (Fig. 20.9A and B)(Yamamoto et al., 2008); one axis (a-axis) is comprised of contiguous
biotin-bound subunit pairs whereas the other axis (b-axis) is comprised ofcontiguous biotin-unbound subunit pairs (Fig. 20.9C). The diffusivity
along the b-axis is approximately 2.4 times larger than that along the aaxis. This anisotropy is ascribed to the difference in the free energy of
association between the biotin-bound and not biotin-bound subunit–
subunit interaction. A preferred intermolecular contact occurs between
the not biotin-bound subunits. The difference in the intermolecular-binding free energy between the two types of subunit pairs is estimated from theratio of diffusion constants to approximately 0.52 kcal/mol. AnotherAuthor's personal copy
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dynamic behavior observed for point defects is the fusion of two point
defects into a larger defect, which occurs much more frequently than thefission of a point defect into smaller defects (Fig. 20.9D). The diffusivity ofpoint defects increases with increasing defect size. Fusion and higher diffusivity of larger defects are suggested to be involved in the mechanism of
defect-free crystal formation.

5.2. p97 2D crystals
A multifunctional AAA ATPase, p97, is involved in cellular processes suchas membrane fusion (Hetzer et al., 2001; Rabouille et al., 1998) and extraction of proteins from the endoplasmic reticulum for cytoplasmic degradation ( Jarosch et al., 2002; Ye et al., 2001). Each subunit of the hexameric
p97 contains two AAA domains (N-terminal D1 and C-terminal D2) and aflexible N-domain (Huyton et al., 2003). Its conformational changes during
the ATPase cycle have been studied using cryo-EM and single-particle
analysis in the presence of analogues of ATP hydrolysis products (Rouiller
et al., 2002). It has been suggested that the relative positioning of the ringshaped D1 and D2 domains changes and the N-domain moves in a direction
perpendicular to the ring planes. We sought to prepare 2D crystals of p97 on
Ni-NTA-containing lipid bilayers using a p97 with His-tag at the Nterminus of each subunit. It did not form crystals at neutral pH, but formeda crystal at acidic pH. Interestingly, p97 placed in an end-up orientation
showed fluctuations in the height even in the absence of nucleotides(Fig. 20.10), suggesting that the N-domain assumes two conformations inequilibrium. This conformational equilibrium probably shifts to one of thetwo states depending on the intermediate in the ATPase cycle, which islikely a process needed to extract proteins from the endoplasmic reticulum.

0s

1s

2s

3s

50 nm

Figure 20.10 High-speed AFM images of two-dimensional crystal of p97 having Histag at the N-terminus, which was formed on Ni-NTA-containing highly fluidic lipid
bilayer. The crystal was formed under acidic pH (pH 5.6). The height of p97 fluctuates
relative to other molecules in the crystal (arrowheads). Successive images wereobtained at an imaging rate of 0.5 s/frame with 256  256 pixels.

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0.6 s

2.1 s

Figure 20.11 Dynamics at the crystal edge of bacteriorhodopsin in purple membrane(Yamashita et al., 2009). The AFM images were taken at 3.3 frames/s (scale bar,
10 nm). The bR molecules encircled by the black dotted lines (at 0.6s) indicate
newly bound bR trimer. The white triangles (at 2.1 s) indicate the previously boundtrimers.

5.3. Bacteriorhodopsin 2D crystals
In the purple membrane (PM) of Halobacterium salinarum, bacteriorhodopsin(bR) monomers associate to form a trimeric structure, and the trimers
arrange in a hexagonal lattice (Henderson et al., 1990). The 2D crystals ofbR and any crystals in general are in dynamic equilibrium with the constituents at the interface between crystal and liquid. High-speed AFM
movies of the crystal edge of PM directly revealed this dynamic equilibrium(Fig. 20.11). Binding of trimeric bR predominates (82%), whereas bindingof dimeric and monomeric bR is much less frequent, indicating that trimers
are preformed in the noncrystalline region. The residence time of newly
bound bR trimers at the crystal edge depends on the number of interaction
sites. Within the 2D bR crystal, a trimer can interact with the surrounding
trimers through six sites, whereas the number of interaction sites at the
crystal edge is reduced to 1, 2, or 3, depending on the binding position of a
newly bound trimer. From the ratio of the average residence times (0.19 and0.85 s) of two bound species containing two or three bonds, the elementary
association energy is estimated to about 1.5kBT, which corresponds to0.9 kcal/mol at 300 K (Yamashita et al., 2009).

6. Low-Invasive Imaging
In tapping mode high-speed AFM, the lateral force exerted from an
oscillating cantilever tip to a sample is very small even when an image istaken within 30 ms (scan lines, 100). High-speed AFM for biologicalsamples requires small cantilevers with a high resonant frequency (1 MHz)and a relatively small spring constant (0.1–0.2 N/m). With high-frequency
oscillations, the cantilever tip makes contact with a sample for only a veryAuthor's personal copy
High-Speed Atomic Force Microscopy Techniques559

short time (100 ns). During this virtually instantaneous contact, thesample stage is displaced only by <0.2 nm as long as the scan rangeis < 300 nm, and hence, deformation of the sample in the lateral direction
is negligible. On the other hand, the vertical force (tapping force) should
be reduced by combining the various protocols and devices described in
the following. This is mandatory for visualizing delicate biomolecular
processes.
1. The free oscillation amplitude of a cantilever is optimized so that thetapping force is reduced and the feedback operation is not slowed down
significantly. For very delicate samples, the amplitude has to be set at< 2 nm, sacrificing the feedback speed to some extent.
2. The amplitude set point should be set close to the free oscillation
amplitude (> 90% of the free oscillation amplitude). Tip parachuting,which often occurs under this condition at the steep downhill regions of
the sample, can be avoided using a dynamic proportional-integralderivative (PID) controller (Kodera et al., 2006). In this controller, thegain parameters can be adjusted automatically and dynamically depending on the cantilever oscillation amplitude.
3. The free oscillation amplitude of a cantilever has to be maintained
precisely constant. Otherwise, the tip-sample distance (hence, thetapping force) changes with time even when the cantilever amplitude
is maintained during imaging at the set point. Since the difference
between the free oscillation amplitude and the set point is less than
0.2–0.3 nm under a very weak tapping force, even small drifts toward
smaller free oscillation amplitude easily result in complete tip-sample
detachment. This problem can be solved by a compensator for drift in
the free oscillation amplitude (Kodera et al., 2006). This compensator
controls the AC voltage applied to a cantilever-excitation piezoactuatorto maintain a constant second-harmonic amplitude of the cantilever.

7. UV Flash-Photolysis of Caged Compounds
UV flash-photolysis of caged compounds is useful to detect minutestructural changes of proteins induced by the interaction with chemicals
such as ATP and calcium. Detected structural changes that are synchronizedwith the flash illumination can be concluded to be caused by the action of
the uncaged chemical, facilitating distinction from Brownian motion. After
irradiating a small area around the imaging area by a UV flash, the concentration of uncaged compound at the imaging area quickly decreases by rapid
diffusion of the chemical. Therefore, repeated observation of structuralchanges is possible when they are reversible.

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For a successful observation, the following precautions are required
because UV illumination is prone to disturb the oscillation of a cantileveror to displace the sample stage by thermal expansion.
1. Use high-frequency weak UV pulses (0.1 mJ/pulse) instead of a single
shot of strong flash. A Nd:YVO4 355 nm laser with a repetition frequency >50 kHz is commercially available.
2. Apply laser pulses while the y-scan toward the starting point of imaging
is performed (without x-scan) immediately after acquisition of a frame.
3. Withdraw the sample stage from the cantilever tip during the y-scan.
Using a calcium-binding protein, CaM, attached to the neck region of
MyoV, we here demonstrate high-speed AFM imaging combined withflash-photolysis of caged calcium. Each neck region of MyoV comprisedof 6 IQ motifs to which six light chains (5 CaM and an essential light chain)
are attached. It is known that CaM attached to the second IQ dissociates in
the presence of Ca2þ at micromolar concentrations (Koide et al., 2006). To
facilitate the observation of structural changes induced in MyoV by CaM
dissociation, MyoV was weakly attached to an aminosilane-coated mica
surface on which the neck regions can extend (Fig. 20.12A). On each neckregion, three small globes corresponding to three light chains are visible, but
the other three light chains face the mica surface. Just after the illuminationby 200 UV pulses with 50 kHz, one of the motor domains (a large globe)
located at the distal end of the head partially dissociates from the adjacentneck region (Fig. 20.12B), but the three globes on the neck remain visible.
Probably one of the three light chains facing the mica surface dissociatedfrom the neck. The partially dissociated motor domain was observed to belinked to the neck region through a thin structure. This behavior was rather
rare among many observations. The observations mostly showed shrinkage–0.46 s+0.46 s

A

B

30 nm

Figure 20.12 Structural change in myosin V upon uncaging calcium from cagedcalcium. Calcium was released from caged-calcium at 0 s. (A) Before UV illumination,(B) after UV illumination. The AFM images were taken at 464 ms/frame.

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High-Speed Atomic Force Microscopy Techniquesof the neck region after UV illumination, suggesting folding of an exposed
IQ motif after dissociation of CaM.

8. Cantilever Tip
Finally, high spatial resolution is also important in high-speed AFM.
The resolution is mainly determined by the apex radius and the aspect ratioof a cantilever tip, in addition to the performance of the instrument. Thusfar, small cantilevers with a sharp tip are not commercially available. Pieceby-piece attachment of a carbon nanotube to a cantilever tip is possible (Daiet al., 1996). However, this laborious attachment cannot be routinely
performed. Instead, we have employed electron beam deposition (EBD)
to grow a carbon tip on the original tip (Wendel et al., 1995). A piece ofphenol crystal (sublimate) is placed in a small container with small holes( 0.1 mm diameter) in the lid. The container is placed in a scanningelectron microscope chamber and small cantilevers are placed above theholes. A spot-mode electron beam is focused onto each original tip, which
produces a needle on the original tip at a growth rate of 17 nm/s.
The newly formed carbon tip has an apex radius of 15–25 nm(Fig. 20.13A and B) and is sharpened by plasma etching in argon or oxygengas, which decreases the apex radius to 4–5 nm (Fig. 20.13C).

A

B

C

r ~ 5 nmr ~ 17 nm

1mm
100 nm

100 nmFigure 20.13 (A) An SEM image of an EBD tip grown from phenol as a vapor source.
The deposition time was 90 s. The electron energy, working distance, and aperture sizeof the FE-SEM during the deposition were 20 kV, 4 mm, and 30 mm, respectively. (B)The apex radius of the grown tip is about 17 nm. (C) After etching in Ar gas for 8 minusing a plasma etcher with the power of 16 W, the apex radius is reduced to about 5 nm.

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