LETTERS
PUBLISHED ONLINE: 14 FEBRUARY 2010 | DOI: 10.1038/NNANO.2010.7

High-speed atomic force microscopy shows
dynamic molecular processes in photoactivatedbacteriorhodopsin
Mikihiro Shibata1, Hayato Yamashita1, Takayuki Uchihashi1,2, Hideki Kandori3 and Toshio Ando1,2 *Dynamic changes in protein conformation in response toexternal stimuli are important in biological processes, but it
has proved difﬁcult to directly visualize such structuralchanges under physiological conditions1–10. Here, we showthat high-speed atomic force microscopy7 can be used to visualize dynamic changes in stimulated proteins. High-resolution
movies of a light-driven proton pump, bacteriorhodopsin11,12,
reveal that, upon illumination, a cytoplasmic portion of eachbacteriorhodopsin monomer is brought into contact with adjacent trimers. The bacteriorhodopsin–bacteriorhodopsin interaction in the transiently formed assembly engenders both
positive and negative cooperative effects in the decay kinetics
as the initial bacteriorhodopsin recovers and, as a consequence,
the turnover rate of the photocycle is maintained constant, on
average, irrespective of the light intensity. These results
conﬁrm that high-resolution visualization is a powerfulapproach for studying elaborate biomolecular processes underrealistic conditions.
The biological function of a protein is closely associated with its
ability to undergo structural changes. In many cases, these structuralchanges are triggered by external stimuli such as pH, temperature,
ligand binding, mechanical stress and light. Although direct realspace and real-time visualization of such changes is a straightforward approach to understanding the dynamic molecular processes
involved, the lack of suitable techniques has been a barrier.
Atomic force microscopy (AFM) is a versatile technique withwhich to image proteins in liquids at submolecular resolution, but
its poor temporal resolution has meant that only static or slow
time-lapse images of proteins have been obtained1–5. In the last
decade, various efforts have been made to increase the scan speedof AFM6–9. As a result, single protein molecules showing
Brownian motion have been captured on video with a temporal resolution of 30 ms (the highest achieved to date; ref. 10). However,dynamic visualization of physiologically relevant conformationalchanges in proteins has been difﬁcult because the tip–sample interaction tends to interfere with the physiological functions. To solve
this problem, a new method has recently been developed thatallows fast and precise control of the tip–sample distance with aminimum load to the sample7. Using such methods, we report
here the ﬁrst example of dynamic imaging of a functioning
biological sample.
Bacteriorhodopsin (bR) represents a well-known case of theassociation between stimulus-triggered structural dynamics andbiological function11,12, and its direct visualization has long been agoal. Bacteriorhodopsin contains seven transmembrane a-helices(termed A–G) enclosing the chromophore retinal13,14. In the photocycle, a series of spectral intermediates, designated J, K, L, M, N andO, occur in that order12. The light-induced conformational changes
in bR have been investigated by various methods15–25, leading to aconsensus that the proton channel at the cytoplasmic surface is
opened by the tilting of helix F away from the protein
centre21,23,24. Sass et al. reported helix F displacement of 0.1 nm
in the late M state, based on X-ray diffraction of the three-dimensional crystal of wild type (WT)21. However, a larger structuralchange in bR was reported in the electron crystallography study of
the D96G, F171C, F219 L triple mutant of bR: displacement ofhelix F by 0.35 nm, away from the centre of the protein23. An electron crystallography study of the F219 L mutant further reported
that helices E and F tilt away from the centre of the protein, followed
by a shift of the E–F loop by 0.3 nm (ref. 24), resulting in largescale conformational changes in the M and N states18,19,25.
Nevertheless, a consensus on the conformational changes has not
yet been reached. As for other proteins, knowledge of the structuraldynamics of bR is limited because its structural data have been accumulated using static and ensemble averaging approaches. Direct anddynamic visualization using high-speed AFM should provide a
better overview of the conformational changes of bR.
We used the D96N bR mutant, which has a longer photocycle(10 s) than the WT (10 ms), but retains its proton pumping
ability26. Figure 1a presents successive images of D96N at the cytoplasmic surface, captured at 1 frame per second (fps; Supplementary
Movie S1). Individual bR molecules can be seen clearly, as reported
in earlier pioneering studies for the unphotolysed state3–5. Upon
illumination with green light, bR drastically changes its structure(compare images at 1 and 2 s) and returns to the unphotolysedstate in a few seconds after light-off, an outcome that is reproducible
in repeated dark–illumination cycles (Supplementary Movie S1).
We analysed the ‘mass centre’ positions of the individual monomers imaged during the dark–illumination cycles. Note that the‘mass centre’ for each bR monomer was calculated from thecorresponding surface area and height distribution in the image(see Methods). All the activated bR monomers exhibited displacements of their centre of mass, with an average displacement of
0.69+0.15 nm (Supplementary Fig. S1, in the green region). In
Fig. 1b, the trajectories of the mass centre positions during thedark–illumination cycles are superimposed on the AFM image inthe unphotolysed state (red, blue and black marks). They show
highly centrosymmetric trajectories, oriented outward from thetrimer centres.
The top and bottom images in Fig. 1c show magniﬁed images
obtained before and during illumination, respectively. The prominent protrusion in the AFM topographs reﬂects interhelical loop
structures on the cytoplasmic surface4,27. Compared with theatomic model of the a-helical cytoplasmic ends in the unphotolysed

1

Department of Physics, Kanazawa University, Kanazawa 920-1192, Japan, 2 CREST/JST, Tokyo 102-0075, Japan, 3 Department of Frontier Materials, Nagoya
Institute of Technology, Nagoya 466-8555, Japan. * e-mail: tando@kenroku.kanazawa-u.ac.jp

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LETTERS

NATURE NANOTECHNOLOGY DOI: 10.1038/NNANO.2010.7
a
5 nm

0s

1.0 s

3.0 s

2.0 s

c
5 nm

E
D C F
G
B
A
C

5 nm

d

D
E

G F

E
DC F
G
BA

1

2

E
D C F
G
B
A

Dark

Light

θ

A

B C

D
E

G F

0.0

100

Abs. change at 410 nm (%)

B
A

Ratio of activated molecules (%)

b

1.0
nm

80
60

100
80
60
40
20

7 8

9

0

40

0

20

50

8

9

50

100
Time (s)7100Time (s)

150

200

0
0

150

200

Figure 1 | High-speed AFM images of the cytoplasmic surface of D96N under dark or illuminated conditions. a, Successive AFM images of D96N bR
adsorbed onto a mica surface in 10 mM Tris-HCl (pH 7) and 300 mM KCl. Frame rate, 1 fps; pixel size, 200  200 pixels (Supplementary Movie S1). A bR
trimer is highlighted by the white triangle. The green bars indicate illumination of 532-nm green light at 0.5 mW. b, Traces (red, blue and black marks) of themass centre positions under dark–illumination cycles are superimposed on the image of D96N in the dark. The photo-induced movement of bR includes
counterclockwise rotation (7.4+2.28) around the trimer centre. c, Surface maps of the magniﬁed images in the dark (upper panel) and under illumination(bottom panel). The position of each trimer centre is denoted by white dots. A monomer in the dark is indicated by a single white arrow. Under illumination,
the topography of the monomer splits into major and minor protrusions, as indicated by green arrows 1 and 2, respectively. d, Decay after ﬂash-illuminationof the activated state at different pH values (7, 8 or 9), detected by high-speed AFM. The exponential decay constants (t ) at pH 7, 8 and 9 are 6.7+0.10 s(the total number of analysed bR molecules, nT ¼ 320; the number of different molecules analysed, nm ¼ 52), 25+0.25 s (nT ¼ 373, nm ¼ 65), and48+0.59 s (nT ¼ 214, nm ¼ 120), respectively. The inset shows the absorbance change at 410 nm (that is, decay of the M-intermediate) after ﬂashillumination of D96N measured at various values of pH (7, 8 or 9). The exponential decay constants (t ) at pH 7, 8 and 9 are 3.4+0.025 s, 14+0.034 s
and 33+0.080 s, respectively.

state14 (A–G in Fig. 1b,c), the protruding areas around helices E andF shift outwards from the trimer centre under illumination.
Therefore, the protrusion movement visualized here is ascribed tothe displacement of the E–F loop. As a result of E–F loop displacement, the protrusion is divided into two sections (green arrows 1
and 2 in Fig. 1c), and the minor protrusion (green arrow 2) likely
corresponds to the position around helices A and B. Note that theoverall position of each bR molecule does not change because
of indiscernible alterations at the extracellular surface(Supplementary Fig. S2 and Movie S5).
To ensure that the conformational changes observed here are not
artifacts such as tip-force-induced structural alterations4, we
measured the decay of the active state after ﬂash illumination at
various pH values (Fig. 1d). UV-visible spectroscopy26 has shownthat the lifetime of the M-intermediate of D96N and WT is prolonged at alkaline pH. In agreement with this fact (inset, Fig. 1d)is the observation here with D96N that the decay detected byhigh-speed AFM showed a strong pH dependence (Fig. 1d). Using
WT at pH 7 and pH 10, this ﬁnding was further conﬁrmed(Fig. 2, Supplementary Movies S2,S3). Because of the very fast
photocycle, no conformational changes were detected in WT at
pH 7, whereas at pH 10, conformational changes similar to thoseof D96N were observed, although the activated state decays much
faster than that of D96N.
The activation of D96N was observed within 1 s immediately
after illumination, but was not observed after light-off, even at alkaline pH (data not shown). A cysteine accessibility study with D96N
indicated displacement of the E–F loop in the M state19. Therefore,
the high-speed AFM imaging of D96N did capture the lightinduced ground-to-M-state transition. Because the AFM observation was performed with the samples on a mica surface, wecannot exclude a possible effect of the surface. Despite this uncertainty, the longer decay constant observed using AFM than thatobserved optically suggests that the activated state detected by

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209

LETTERS
M1
M2
Distance from initial position (nm)Distance from initial position (nm)

M2
M3
M4
M5
M6
M7
M8
M9
M10
0

M3
M4
M5
M6
M7
M8
M9
M10

1.5 nm30

60
90Time (s)

120

150

0

30

60
90
Time (s)120

150

Figure 2 | Displacement of mass centre positions for WT at thecytoplasmic surface. a,b, Ten typical bR monomers at pH 7 (a) and pH 10(b) are analysed. The AFM movies for WT at pH 7 and pH 10 are shown in
Supplementary Movies S2 and S3, respectively. The green regions show
periods of green light illumination (532 nm, 0.5 mW). Conformationalchanges in WT are not detected at pH 7, because the photocycle is too fast,
whereas at pH 10, they were repeatedly detected because the photocycle isprolonged at alkaline pH29.

AFM may contain an additional state in which the Schiff base isreprotonated (during M decay) but the protein conformation has
not yet returned to the ground state.
As a result of the outward displacement of the E–F loop, nearestneighbour bR monomers, each belonging to a different adjacent
trimer, transiently assemble. Here, we use a new designation,‘trefoil’, for the triad of nearest-neighbour monomers to distinguishit from the original trimer. Notably, this transient assembly in a
trefoil alters the decay kinetics of the activated state(Supplementary Movie S4). Figure 3a shows the time course of
the displacement of the mass centres for six bR monomers belonging to two different trefoils observed at different light intensities(Mn1–Mn3; n indicates different trefoils) (Supplementary
Fig. S3). Under weak illumination, it is mainly the case that only
one monomer in each trefoil is activated (M13 and M22 in the left
panel of Fig. 3a). When only one monomer is activated in a
trefoil, it decays with a time constant of 7.3+0.58 s (the top panelof Fig. 3b). In contrast, under stronger illumination, two or threemonomers within a trefoil tend to be activated together.
Interestingly, the decay of each monomer markedly depends on
the order of its activation. The monomer that is activated latest
among the activated monomers in the trefoil (blue arrows in theright panel of Fig. 3a) decays with a shorter time constant of
2.0+0.16 s (the middle panel of Fig. 3b). On the other hand, thedecay kinetics of the early activated monomers (M12 and M22 in
the right panel of Fig. 3a) does not follow a single exponential(the bottom panel of Fig. 3b), and the averaged decay time lengthensbStronger

M11

M11

M12

M12
Distance from initial position (nm)Distance from initial position (nm)Weaker

M13

M21

M22

Number of molecules (N)

a

M13

M21

M23

1.5 nm20
40
Time (s)

60

40
30
20
10
0
0

5 10 15 20 25 30 35 40 45
Time (s)0

5 10 15 20 25 30 35 40 45
Time (s)0

5 10 15 20 25 30 35 40 45
Time (s)

80
60
40
20
0

M22

M23

0

Number of molecules (N)

b

M1

0

20
40
Time (s)60

Number of molecules (N)a

NATURE NANOTECHNOLOGY DOI: 10.1038/NNANO.2010.7

40
30
20
10
0

Figure 3 | Cooperative effects on the decay kinetics in D96N. a, Displacement of mass centre positions as a function of time measured for six bRmonomers forming two different trefoils (Mn1–Mn3; n indicates different trefoils) at pH 7 under different light intensities (Supplementary Fig. S3 and Movie
S4). The green regions correspond to the illumination, and their shade level indicates the relative light intensity (0.007 and 0.5 mW). The blue arrows
indicate the conformational changes of the monomers (M13, M21 and M23) that are activated subsequent to the activation of other monomers in therespective trefoils. b, The top ﬁgure shows the decay of activated bR monomers (as depicted by the red circle) while the other monomers (the grey circles)
within the respective trefoils are not in the activated state. The exponential decay constant (t ) is 7.3+ 0.58 s (nT ¼ 279, nm ¼ 55). On the other hand, when
one or two nearest-neighbour molecules (as depicted by green circles in the middle ﬁgure) in a trefoil are already activated, the bR monomer activated latest(as depicted by the blue circles in the middle ﬁgure) decays faster. The exponential time constant (t ) is shortened to 2.0+0.16 s (nT ¼ 228,nm ¼ 55). In contrast, the early activated molecules within a trefoil (as depicted by green circles in the bottom ﬁgure) show a non-exponential broad
distribution of decay time with an average of 13.3 s (nT ¼ 194, nm ¼ 55).
210

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NATURE NANOTECHNOLOGY DOI: 10.1038/NNANO.2010.7

Number of molecules (N)80

Weaker light intensity

60

40

20

0
0

5

10

15

20

25

30

35

40

45

LETTERSAFM contains both the M-state and subsequent states.
Importantly, the cooperative effects we found are caused by bR–bR interactions within a trefoil, not within a trimer.
As demonstrated above, direct and real-space visualization of the
dynamically changing structure of stimulated proteins provides a
straightforward way of elucidating how protein molecules function.
We can expect that a large number of dynamic biomolecular
processes will be directly visualized in the near future, leading to
new ﬁndings inaccessible using other approaches. With further
advances in high-speed AFM techniques, even molecular processes
on membranes of live cells will be directly visualized. The present
study is the ﬁrst step towards achieving these expected innovations
in biological research.

Time (s)Methods
Number of molecules (N)60

Sample preparation. Purple membranes containing either WT or D96N mutant
were isolated from Halobacterium salinarum as described previously30. The samples
were suspended in a solution containing 10 mM Tris-HCl (pH 7, 8, 9 or 10) and
300 mM KCl.

Stronger light intensity

40

20

0
0

5

10

15

20

25

30

35

40

45

Time (s)Figure 4 | Decay rate of activated D96N under different light intensities.
The upper ﬁgure shows the decay of activated molecules under weakillumination (0.004–0.09 mW). Although the frequency is low, two bR
monomers in a trefoil are activated together, the events of which are alsocounted in this analysis. The exponential decay constant (t ) is 5.4 + 0.34 s(nT ¼ 429, nm ¼ 55). The bottom ﬁgure shows the decay of activatedmolecules under relatively strong illumination (0.23 or 0.5 mW). The
exponential decay constant (t ) is 6.1+0.37 s (nT ¼ 416, nm ¼ 55).

to 13 s. This observation indicates that the early activatedmonomers do not return to the ground state as long as the adjacentmonomers within the trefoil are in the active state.
In addition, Fig. 4 shows decay of the activated D96N under weakor strong light intensities. Note that all the activated molecules werecounted regardless of their activation order and of the number ofactivated monomers in a trefoil. The distributions of the activatedstate lifetime are well ﬁt to single exponential functions underboth weak and strong light illumination, and their decay constants
are almost identical, despite the different light intensities. This canbe explained by the coexistence of the positive and negative cooperativity effects elicited by bR–bR interactions within a trefoil (Fig. 3b).
Because of the coexistence and the bipolar nature, the two cooperative effects cancel out one another on averaging, resulting in a constant decay rate independent of light intensity. Under a conceivableassumption that the conformational change turnover of an activatedbR monomer is tightly coupled with proton transfer by the bR
monomer, the average rate of proton pumping by the activated bR
would be conserved independently of light intensity. As a result of
the cancellation effect, the cooperative effects can never be detectedby ensemble averaging approaches. Thus, the cooperative effects aredifferent from those proposed from absorption spectroscopic
studies in which fast and slow decays of the M-state were detectedthat depend on light intensity, while the total turnover rate was independent of light intensity28,29. This previous report is consistent with
our presumption that the activated state detected by high-speed

High-speed AFM observation. The laboratory-built high-speed AFM apparatus isan extensively improved version of the previously reported one6. See a
comprehensive review7 of instrument development, including recent technicaladvances. AFM images were acquired in the tapping mode. To detect cantileverdeﬂection, we used an optical beam deﬂection detector equipped with an infraredlaser (980 nm). The laser beam was focused onto a small cantilever using a 50
objective lens. The cantilever (Olympus) was 6–7 mm long, 2 mm wide and 90 nm
thick, with a spring constant of 0.1–0.2 N m21. Its resonant frequency and quality
factor in aqueous solution were 1 MHz and 2, respectively. In the AFM imaging,
the free oscillation amplitude was 1 nm and the set-point amplitude was 90% of
the free amplitude. The tapping force estimated was less than 30 pN. An amorphous
carbon tip was grown on the original tip by electron-beam deposition. The tip length
was adjusted to 1 mm, and the tip apex was sharpened by plasma etching underargon gas (4 nm in radius). AFM observations were performed under solution atroom temperature. For illumination of the bR sample, a green laser (532 nm) was
irradiated through the 50 objective lens. The intensity measured at the exit of theobjective lens was varied in the range 0.004–0.5 mW.
Data analysis of mass centre position. The mass centre positions of imaged bR
monomers were calculated by the following procedures. First, a whole AFM image
was segmented into each monomer region by a watershed algorithm. Second, thegreyscale of each segment was digitized using a threshold determined by a fuzzyentropy algorithm. The pixels with greyscales above threshold were deﬁned as thoseof a monomer (region of interest; ROI). Finally, the centre of mass of a monomer
was calculated for the ROI using the pixel positions and the greyscales. Note that thecentre of mass analysed here is not the real centre of mass of the protein, but that of
the AFM image of the monomer.

Received 1 September 2009; accepted 8 January 2010;
published online 14 February 2010

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Acknowledgements
This work was supported by Japan Science and Technology Agency for Core Research for
Evolutional Science and Technology (T.A.), Grants-in-Aids for Scientiﬁc Research fromJapan Society for the Promotion of Science (JSPS) (no. 15101005; T.A.) and from the
Ministry of Education, Culture, Sports, Science and Technology, Japan (no. 19042009, T.U.;
no. 20108014, H.K.), and Research Fellowships of JSPS for Young Scientists (M.S. andH.Y.). The authors thank J.K. Lanyi, S.P. Balashov and L.S. Brown for commentson the draft.

Author contributions
T.A., H.K. and M.S. conceived and designed the experiments. T.A. and T.U. developed thehigh-speed AFM instrument. T.U., M.S. and H.Y. performed the experiments. T.U. and
M.S. analysed the data. T.A., T.U. and M.S. co-wrote the paper. All authors discussed theresults and commented on the manuscript.

Additional information
The authors declare no competing ﬁnancial interests. Supplementary informationaccompanies this paper at www.nature.com/naturenanotechnology. Reprints andpermission information is available online at http://npg.nature.com/reprintsandpermissions/.
Correspondence and requests for materials should be addressed to T.A.

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