Atomic Force Microscop y in Biology
In the last decade the atomic force microscope
(AFM) (Binnig et al.
, 1986) has become a powerful tool
in structural biology. The unique possibility of acquiring
the topography of biological samples at high resolution
under physiological conditions, i.e., in buffer
solution, at room temperature, and under normal pressure,
makes this instrument outstanding. The high
signal-to-noise ratio of AFM topographs has not only
allowed the observation of whole cells, chromosomes,
nucleic acids, and proteins, but also the tracking of
conformational changes of biomolecules during the
exertion of their function (Engel et al.
, 1999; Engel and
Mtiller, 2000). Furthermore, time-lapse AFM imaging
has enabled the monitoring of dynamic changes in the
conformation, association, and functional state of individual
proteins (Stolz et al.
The principle of the AFM is relatively simple: A
sharp tip mounted at the end of a flexible cantilever is
raster scanned over a sample surface in a series of
horizontal sweeps. The bending of the cantilever caused
by the probe-sample interaction is detected by the
deflection of a laser beam that is focused onto the end
of the cantilever and reflected into an optical detector.
This signal, termed deflection signal, is coupled to a
servo system that moves the sample vertically to maintain
the cantilever deflection at a constant value. The
surface topography is then reconstructed from the vertical
movement of the scanner. In this imaging mode,
called contact mode, the probing tip always touches
the surface with a constant force during scanning. An
alternative and widely used imaging mode in biology
is the tapping mode. Here, the AFM tip is oscillated
rapidly in the vertical direction while scanning the
sample. Oscillation of the tip reduces frictional forces,
thereby minimizing artifactual deformation and displacement
of the sample. Therefore, the tapping mode
is used frequently to image the surface topography of
weakly immobilized biomolecules, e.g., single proteins
and fibrils. However, for imaging of biological membranes,
the highest resolution so far obtained has been
in contact mode.
This article focuses on the application of contact
mode AFM to acquire subnanometer resolution structural
information of membrane proteins from membrane
specimens in liquid. Most of these proteins are
fragile structures comparable to a submerged sponge.
To prevent their damage by the scanning tip, soft cantilevers
with spring constants of 0.01-0.1N/m must be
used, and scanning must be performed with minimal
applied force to the tip (≤100 pN). An additional crucial
factor for successful high-resolution imaging is the
correct adjustment of the imaging buffer (Müller et al.
1999). By optimizing these factors that minimize the
force experienced by the sample, lateral resolutions
down to 0.41nm and vertical resolutions down to
0.10 nm have been achieved on biological membranes
in solution (Stahlberg et al.
, 2001). However, application
of higher forces can be of advantage to perform precise
and controlled "dissections" of biological samples by
manipulation with the AFM tip (Fotiadis et al.
|FIGURE 1 Bacteriorhodopsin with its retinal
(arrowhead). The extracellular (top) and the
(bottom) of BR with their prominent
loops and termini are indicated.
This illustration of BR was
calculated using the coordinates
of Kimura et al. (1997)
and the visualization program DINO
In this article, contact mode AFM is illustrated using
membranes that contain the heptahelical protein
bacteriorhodopsin (BR). This 26-kDa integral protein
acts as a light-driven proton pump in Haloarchaea
(Oesterhelt and Stoeckenius, 1973). Photoisomerization
of the chromophore from all-trans
initiates the unidirectional translocation of one proton
across the cell membrane (reviewed by Oesterhelt et al.
1992; Lanyi, 1997). This establishes an electrochemical proton gradient across the membrane that can then be
used for ATP synthesis and other energy-requiring
processes in the cell. BR forms trimers and highly
ordered two-dimensional (2D) trigonal crystal lattices
= 6.2nm, γ = 60°) in the native
membrane of the bacterium Halobacterium salinarum
these crystalline patches are termed purple membrane
because of their color (Blaurock and Stoeckenius,
1971). The flatness of these crystalline membrane
patches makes them very suitable for AFM. Highresolution
three-dimensional structures of BR (Fig. 1)
have been determined by electron and X-ray diffraction
(for a review, see Cartailler and Luecke, 2003). In
BR, the prosthetic group retinal (see Fig. 1; arrowhead)
is bound covalently by a protonated Schiff base to
K216 of helix G of the protein (the seven transmembrane
α helices are generally denoted A to G). With the
AFM it is the protruding domains, i.e., the termini and
connecting loops between the helices, that are visualized,
not the helices themselves that are embedded in
the lipid bilayer. On the cytoplasmic side, the major
protrusions of BR are the loop connecting the transmembrane
α helices A and B and that connecting E
and F (AB and EF loops; Fig. 1), of which the EF loop
appears to be highly flexible (Müller et al.
, 1995a). On
the extracellular side the protruding B-C interhelical
loop (BC loop; Fig. 1) forms a β hairpin and is fairly
The procedures described for BR in this article constitute
a basis for sample preparation and application
of AFM on other biological membranes.
II. MATERIALS AND
A. Materials: Preparation of Mica Supports
Polished ferromagnetic stainless steel disks of 11mm
diameter (manufactured by the internal workshop
services of the Biozentrum, Basel, Switzerland)
Teflon sheets of 0.25 mm thickness (Maag Technic AG,
Mica sheets with a thickness between 0.3 and 0.6 mm
(Mica House, 2A Pretoria Street, Calcutta 700 071,
"Punch and die" set from Precision Brand Products
Inc. (2250 Curtiss Street, Downers Grove, IL 60515)
Ethanol [purity: >96% (v/v)]
Loctite 406 superglue (KVT König, Dietikon,
Araldite Rapid: Two-component epoxy glue from
Ciba-Geigy, Basel, Switzerland
Scotch tape (3M AG, Rüschlikon, Switzerland)
B. Materials: Bacteriorhodopsin and Buffers
Sodium azide (NAN3
, Fluka Cat. No. 71289)
, Merck Cat. No. 1.08382.2500]
Magnesium chloride hexahydrate (MgCl2·
Cat. No. 63064)
Potassium chloride (KCl, Merck Cat. No. 1.04936.1000)
Purple membranes of H. salinarum
Stock suspension of purple membrane fragments:
0.25mg/ml in double-distilled water containing
(stored at 4°C and protected from unnecessary
A commercial multimode AFM equipped with a
120-µm scanner (j-scanner) and a liquid cell
(Digital Instruments, Veeco Metrology Group, Santa
micro cantilevers of 100 and
200 µm length, and nominal spring constants of 0.08 and 0.06N
/m from Olympus Optical Co. Ltd.,
Tokyo, Japan, and from Digital Instruments, Veeco
Metrology Group, respectively.
A. Preparation of Mica Supports for Sample
B. Preparing Bacteriorhodopsin for AFM
- Punch Teflon disks of 0.5in. and mica disks of
0.25 in. diameter using the "punch and die" set and
- Clean the steel and Teflon disks with ethanol and
- Glue a Teflon disk centrally on a steel disk using the
Loctite 406 superglue and allow the glue to dry.
- Glue a mica disk centrally on the Teflon surface
of the Teflon-steel disk with the Araldite twocomponent
- Let the supports dry overnight.
- Adsorption buffer: 20mM Tris-HCl (pH 7.8), 150mM KCl
- Imaging buffer for the extracellular side (ES-imaging
buffer): 20mM Tris-HCl (pH 7.8), 150mM KCl,
- Imaging buffer for the cytoplasmic side (CS-imaging
buffer): 20mM Tris-HCl (pH 7.8), 150mM KCl
|FIGURE 2 Schematic diagram of the atomic force
setup for imaging in liquid. The piezo scanner
moves the sample in xyz directions under the fixed
cantilever (marked by an asterisk).
C. Operation of the AFM
- Dilute and mix 3µl of purple membrane stock
solution with 30µl of adsorption buffer in an Eppendorf
- Cleave the mica by applying Scotch tape to the
upper surface and then peeling off until a new smooth
intact mica surface is exposed. This new surface is
clean and molecularly flat, suitable for deposition of
the specimen to be imaged.
- Pipet 33µl of the diluted purple membranes on
the hydrophilic surface of the freshly cleaved mica
- Allow the sample to adsorb for 15 to 30min.
- Wash away purple membrane fragments that are
not firmly attached to the mica by removing approximately
two-thirds of the fluid volume from the mica
surface and readding the same volume of the desired imaging buffer. Repeat this washing procedure at least
- Transport the specimen support and its associated
fluid onto the piezo scanner of the AFM.
- Place the AFM head with its mounted fluid cell
and cantilever on the scanner (see Fig. 2).
- Make sure that the space between the mica
surface and the cantilever-fluid cell contains enough of
the corresponding imaging buffer to avoid drying
of the specimen during the imaging experiment. An
experiment may last several hours.
- Align the laser spot onto the cantilever and the
reflecting beam into the photodiode with the appropriate
laser alignment screws. Caution: It is very important
not to put any reflective objects into the laser
trajectory in order to avoid reflection of the laser light
into your eyes! Additionally, wear appropriate protection
All measurements are carried out under ambient
pressure and at room temperature.
These steps assume basic knowledge of AFM
- Let the instrument equilibrate thermally.
- Set the scan size to zero to prevent specimen
deformation and contamination of the tip after
- Initiate engagement process, i.e., permit the tip to
approach the surface.
- As soon as the tip is engaged, and prior to scanning
the surface, set the operating point of the instrument
to forces below 100pN, i.e., minimal force.
- Calibrate the deflection sensitivity of the cantilever
using the force calibration menu of the control
- Start imaging and keep the forces during scanning
as small as possible by correcting manually for
- Optimize the feedback parameters of the system,
i.e., the integral gain and the proportional gain, by
increasing their values. If the feedback loop starts to
oscillate, introducing noise, reduce these gains until
the noise goes away.
- Record two images of 512 by 512 pixels simultaneously
at low magnification (frame size >1µm), one
showing the height signal in the trace direction and the
other the deflection signal in the retrace direction. Set
the scan speed to two to four lines per second. Crystalline
structures are usually recognized more easily in
the deflection signal image than in the height image.
- Find and centre a suitable purple membrane
fragment. Zoom in and set the scan speed to four to
six lines per second. Record images of the height
signals in both the trace and retrace directions at high
magnification (frame size <1µm). Comparison of the
trace and the retrace images allows deformation of
the sample in the fast scan direction to be detected.
Such deformations can be minimized by working at
minimal force. At such high magnifications, the scan
range of the z piezo can be reduced to avoid limitation
of the axial z resolution that might otherwise occur
because of rounding errors during (16-bit) digitalisation
of the analogue signal (AD conversion).
The following sections explain and help understand
the features observed on the topographies acquired
during the AFM experiment.
A. Morphology of Purple Membranes
Figure 3 shows a typical low-magnification topograph
of purple membrane fragments adsorbed on
mica. The diameter of the BR crystals varies between
0.5 and 1.5µm. The number of adsorbed membrane
fragments depends on the adsorption buffer and
time and on the concentration of the purple membrane
suspension deposited on the mica. It is advantageous
not to adsorb membranes too densely on the mica
surface; this decreases the probability to contaminate the AFM tip. Two different surfaces can be discerned:
The extracellular side of purple membrane is fairly flat
(Fig. 3; arrows), whereas the cytoplasmic side is characterized
by protruding bumps of 10-30nm in diameter
(Fig. 3; arrowheads) (Müller et al.
, 1995b, 1996).
A feature that can be used to differentiate further
between the two sides is the small difference in thickness
seen when scanning in CS-imaging buffer. Purple
membranes exposing the cytoplasmic side appear
slightly thinner than those exposing the extracellular
side. As described by Müller and Engel (1997), pH and
electrolyte concentration affect the apparent height
measured between the mica and the cytoplasmic or the
extracellular side of the purple membrane. This results
from different surface charge densities.
B. The Extracellular Surface of Purple
|FIGURE 3 AFM images of purple membrane at low magnification: height (A) and deflection (B) signals
recorded in trace and retrace directions, respectively. Membrane patches exposing the cytoplasmic side
are decorated by small debris, can be distinguished from patches exposing the flat extracellular
The membrane patches have a height of ~6 nm when imaged in CS-imaging buffer.
Vertical brightness ranges: 12nm (A) and 1nm (B).
At high magnification, the appearance of the extracellular
side of BR is characterized by protrusions
extending 0.5 ± 0.1 nm out of the lipid bilayer (Fig. 4A
and inset). The β hairpin in the loop connecting the
transmembrane α helices B and C of BR constitutes the
main portion of the observed protrusion.
C. Estimating the Resolution of AFM
The resolution of a micrograph containing a regular
structure can be estimated from its power spectrum,
i.e., its 2D Fourier transformation image. This was calculated
from the topograph of the extracellular side of
BR (Fig. 4A) and is displayed in Fig. 4B. Most software
packages delivered with atomic force microscopes
enable calculation of power spectra from recorded
AFM topographs. Various public domain programs
such as NIH image or ImageJ, both from the National
Institutes of Health (Bethesda, MD), or SXM from the
University of Liverpool (Liverpool, United Kingdom),
have such algorithms implemented and can be downloaded
for free (NIH image: http://rsb.info.nih.gov/
nih-image/; ImageJ: http://rsb.info.nih.gov/ij/; and
|FIGURE 4 (A) The extracellular side of purple membrane at high magnification. For clarity, the area
around the contoured BR trimer was enlarged and is displayed in the inset (frame size: 17 × 17 nm). (B) Power
spectrum calculated by Fourier analysis from the image in A. The (3, 10) diffraction spot (small circle) is
marked and represents a resolution of 0.46 nm. The broken circle represents the 1-nm resolution limit.
brightness range: 0.8 nm (A and inset in A).
The larger the distance of the discernable spots from
the origin in a power spectrum, the higher the resolution
of the topograph. Here, spots extend beyond the
1-nm resolution limit (see broken circle in Fig. 4B). To
calculate the resolution at a selected diffraction spot,
Eq. (1) can be used. This equation is applicable to all
lattice types, e.g., for trigonal, hexagonal, square, or
orthorhombic lattices, which occur frequently in native and reconstituted 2D crystals of membrane proteins
(for further reading on crystallography, see Misell and
where δ is resolution; r-
(r raised to bar) is vector from origin to the diffraction
spot in Fourier space; a
are basic lattice
vectors in real space; γ is the angle between the basic
lattice vectors; and h
are Miller indices of the diffraction
Example: The encircled spot (h, k
) = (3, 10) in Fig. 4B
corresponds to a lateral resolution of 0.46nm assuming
the lattice parameters of BR (a = b = 6.2nm, γ = 60°).
D. The Cytoplasmic Surface of Purple
|FIGURE 5 Force-dependent topography of the
surface of a purple membrane. The area
above the broken line was
recorded at minimal force
applied to the AFM stylus (≤100pN). At
this force, the
flexible EF loop is fully extended, whereas at a stylus
force of ~200pN (area below the broken line) the loop is
away by the scanning tip and is therefore almost
anymore. For clarity, areas
(frame size: 7 × 7nm) around the contoured
recorded at minimal force (broken circle; inset top
and at a force of ~200 pN (solid circle; inset bottom left)
enlarged as insets. Occasionally, defective
BR trimers with a
monomer missing can be found
(arrowheads). Vertical brightness
range: 1.2 nm.
Imaging of the cytoplasmic surface of BR is force
dependent because of the flexible EF loop (Müller et
al., 1995a). At minimal force (≤100pN) applied by the
AFM stylus (Fig. 5; area above the broken line and
inset top left), the fully extended EF loops can be discerned
as protrusions of 0.8 ± 0.2nm height. These become less prominent or even disappear if the
loading force is increased to ~200pN (Fig. 5; area
below the broken line and inset bottom left). At this
force, the AB loops become visible with a height of
0.6 ± 0.1nm above the lipid bilayer. The advantage of
such force-induced conformational changes is that
shorter loops otherwise covered by the bigger ones can
A. Damping of Vibrations
For high-resolution AFM imaging, a vibrationally
and acoustically isolated setup of the microscope is
crucial. Antivibration and damping tables or lead platforms
suspended by bungee cords offer excellent
vibration damping. Acoustic isolation of the AFM can
be achieved efficiently by installing a vacuum bell jar
around the microscope.
B. Adjustment of Buffer for High-Resolution
Topographs of this membrane protein with a lateral
resolution of 0.41nm (Stahlberg et al.
, 2001) can be
recorded reproducibly with the AFM, provided the
imaging buffer is adjusted correctly (Müller et al.
and the force applied to the tip is minimized. Under
nonoptimal imaging conditions, even the smallest
force that is adjustable by the instrument can be too
high, leading to deformation of the biomolecules, e.g.,
concealing the EF-loop in BR (Fig. 5, lower portion).
The effective interaction force acting between the AFM
stylus and the specimen is determined by the force
applied to the stylus, the electrostatic repulsion, and
the van der Waals attraction between the two surfaces.
By adjusting pH and ionic strength of the imaging
buffer, van der Waals attraction and electrostatic repulsion
between tip and sample can be balanced. The
best imaging conditions are determined by recording
and analyzing force-distance curves between tip and
sample in different buffers. Conditions that yield force
curves showing a small repulsion are ideal for highresolution
imaging. Under these conditions the tip is
assumed to surf on a cushion of electrostatic repulsion,
thereby minimizing the deformation experienced by
the biomolecules. This screening method revealed the
two slightly different imaging buffers for BR mentioned
earlier, i.e., CS- and ES-imaging buffer. For
further reading on this topic, see Müller et al.
where buffer conditions for different biological
samples are discussed.
C. Tip Effects and Artefacts
At this time, no commercial AFM tips are available
with ideal point probes and perfect geometries in the
subnanometer range. Therefore, tip effects and artefacts
arising from the tip geometry are unavoidable
and have to be considered when interpreting AFM
topographies (for further reading on tip effects and
artefacts, see Xu and Arnsdorf, 1994; Schwarz et al.
1994). Tip effects occur when the probe does not have
a single, small, and sharp interaction area with the
sample. This leads to an AFM image that represents a
convolution of the sample features with the tip shape.
To be sure of having acquired the correct surface structure
and not an artefact, the same surface topography
of the object being investigated has to be reproduced
several times using different tips from different
batches. Tip artefacts can also be identified by changing
the direction in which the AFM tip scans the
sample (scan angle), as artefacts will rotate correspondingly
(Xu and Arnsdorf, 1994). Ordered structures,
e.g., mosaic 2D crystals, that are differently
oriented with respect to the scan direction of the AFM
tip, are also good indicators for tip artefacts. Ideally,
the building blocks of the crystal, e.g., the BR trimer,
should look the same in the differently oriented crystals.
Finally, structures of samples that have been
determined by other methods, e.g., the structure of BR
by electron and X-ray crystallography, can be used to
further compare and confirm AFM data.
Figure 6 displays topographs of the extracellular
side of BR recorded with an artefact-free tip (Fig. 6A)
together with images acquired with artefacteous tips
(Figs. 6B-6D). Tentative explanations of the observed
artefacts are omitted because the statements would be
too speculative. We restrict ourselves to a comparison
of artefacteous surfaces with the nonartefacteous one.
Compared to Fig. 6A, the depression at the three-fold
symmetry axis of the trimer in Fig. 6B is completely
missing. The trimer seems to consist of a single
plateau. The BR trimers in Fig. 6C have lost their trigonal
shapes and resemble tetramers. In Fig. 6D the
BR trimer is distorted, displaying a Y shape. The
lipid areas that separate the neighbouring BR trimers
cannot be resolved by the artefacteous tip, and instead
ridges of constant height are seen (broken straight
VI. CONCLUDING REMARKS
|FIGURE 6 Tip-induced effects and artefacts on the extracellular surface of bacteriorhodopsin. (A) Artefact-
free and (B-D) artefacteous topographies of BR. In B the characteristic central depression at the threefold
axis of the BR trimer is absent. The central trimer in C resembles a tetramer. In D the protrusions are
not clearly separated, but connected along the broken line. In all images the central trimers are marked by
broken circles. The frame sizes in A to D are 17 nm. The heights are 0.8 nm (A), 1.3 nm (B), 0.8 nm (C), and
0.8 nm (D).
This article presented procedures for sample preparation
and AFM imaging of native 2D crystals of
bacteriorhodopsin. The experience gained will enable
the readers to perform similar AFM experiments with
other biological membranes.
This work was supported by the Swiss National
Research Foundation, the M. E. Müller Foundation, the
Swiss National Center of Competence in Research
(NCCR) "Structural Biology," and the NCCR
"Nanoscale Science." The authors are indebted to Dr.
Ansgar Philippsen for Fig. 1 and to Professors Dieter
Oesterhelt (Max-Planck-Institut fiir Biochemie, Martinsried,
Germany) and Georg Büdt (Forschungszentrum
Jülich, Jülich, Germany) for kindly providing us
with BR. The authors acknowledge Dr. David Shotton
for constructive comments on the manuscript.
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