INSTITUTE OF PHYSICS PUBLISHING
NANOTECHNOLOGY
Nanotechnology 17 (2006) S221–S226
doi:10.1088/0957-4484/17/7/S20
Imaging of biomaterials in liquids: a
comparison between conventional and
Q-controlled amplitude modulation
(‘tapping mode’) atomic force microscopy
D Ebeling1,2 , H Hölscher1,2 , H Fuchs1,2 , B Anczykowski3 and
U D Schwarz4
1
Center for Nanotechnology (CeNTech), Gievenbecker Weg 11, 48149 Münster, Germany
Physikalisches Institut, Westfälische Wilhelms Universität Münster, Wilhelm-Klemm-Straße
10, 48149 Münster, Germany
3
nanoAnalytics GmbH, Gievenbecker Weg 11, 48149 Münster, Germany
4
Department of Mechanical Engineering, Yale University, PO Box 208284, New Haven,
CT 06520-8284, USA
2
E-mail: Hendrik.Hoelscher@uni-muenster.de
Received 22 August 2005, in final form 4 December 2005
Published 10 March 2006
Online at stacks.iop.org/Nano/17/S221
Abstract
Lambda phage DNA and DPPC thin films are imaged in liquids by atomic
force microscopy applying the amplitude modulation mode (‘tapping mode’)
with active enhancement of the Q -factor by a ‘ Q -control’ electronics. The
topography of the resulting images is compared with images obtained
without active Q -control. To enable a meaningful comparison, individual
scan lines are alternately recorded with and without Q -factor enhancement
using scan parameters optimized for each mode separately. As the major
finding, significant height differences of topographical features are observed
between the two modes. The heights measured with active Q -control are
reproducibly higher compared to the ones observed without Q enhancement.
This effect is attributed to the reduction of tip–sample forces by Q -control.
(Some figures in this article are in colour only in the electronic version)
1. Introduction
The ability of atomic force microscopy (AFM) [1] to deliver
high-resolution images of a wide variety of samples in
gases, liquids, and vacuum makes it one of the most
powerful and universally applicable tools in surface science.
While alternatives exist, for example, for the investigation
of hard, conducting samples in vacuum (such as electron
microscopy or scanning tunnelling microscopy), AFM remains
unique in its capability to map soft, insulating samples in
liquids with molecular resolution. This quality has drawn
enormous attention of molecular biologists interested in
studying biological systems in the native environment.
In many cases, the goal of all imaging efforts is to achieve
the highest possible resolution with the least distortion of the
0957-4484/06/070221+06$30.00
sample morphology, which might be altered by tip–sample
interactions. Depending on the specific systems, different
sample preparation procedures and/or imaging modes provide
the best results. For instance, high-quality molecular resolution
on macromolecules forming close-packed flat structures with
two-dimensional periodicity has been achieved by contact
mode AFM (CM-AFM) (see, e.g., [2–4] for reviews). This
is because the imaging of the undistorted structure of
biomolecules requires normal tip–sample forces lower than
100 pN, and forces as low as 10 pN have been achieved with
CM-AFM [5].
Contact mode AFM in liquids, however, has significant
drawbacks that limit its applicability to soft biological samples.
Often due to inherent thermal drift, the setpoint of the
interaction force changes continuously so that the load has to
© 2006 IOP Publishing Ltd Printed in the UK
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D Ebeling et al
S222
laser
electronics with
lock-in amplifier
photo diode
phase
amplitude
amplifier
phase
shifter
z
piezo
D+2A
d
aexc
Φ
+
function
generator
amplitude
be frequently readjusted. Worse, since lateral forces acting
during contact mode scanning are significant, only samples that
feature a considerable degree of lateral stiffness can be imaged
successfully. While this is the case for the two-dimensional
molecular crystals introduced above, the lack of resistivity to
lateral forces frequently prevents a high-resolution mapping of
most other types of biological samples [6]. For example, single
molecules supported by substrate surfaces are usually either
displaced or destroyed when imaged with CM-AFM [7].
The most commonly used approach to reduce the influence
of lateral forces is to oscillate the cantilever close to the sample
surface at a fixed driving frequency [8]. If the oscillation
amplitude of the cantilever is used as a feedback signal for
the mapping of the surface topography, this technique is
named amplitude-modulation AFM (AM-AFM) [9]. From
the viewpoint of practical applicability, it is important to note
that AM-AFM responds quickly to changes of the oscillation
amplitude if the quality factor Q of the cantilever is low. Due to
the low Q -values in liquids, AM-AFM has been the dominant
detection scheme for the imaging of biomaterials in a native
environment to date.
Practical imaging in AM-AFM mode is often complicated
by an instability in the oscillation amplitude that occurs during
the approach of the oscillating tip towards the sample [10–13].
This instability marks the transition from the regime of
attractive forces to a ‘tapping’ regime, where significant
repulsive forces are exerted between tip and sample. These
forces can be as high as several tens of nanonewtons [10, 11].
Therefore, amplitude modulation AFM is quite often denoted
as ‘tapping mode’ AFM [14] regardless of whether the
operation is actually performed in the repulsive tapping regime
or not. Even under optimized conditions, peak forces during
one oscillation in the tapping regime have to be assumed to
be one or two orders of magnitude higher than the 100 pN
threshold given above necessary for the undistorted profiling
of soft biological matter. Therefore, considerable sample
deformation has to be expected for this class of sample if
imaging is performed in the tapping regime [7, 15].
From the above discussion, we can conclude that it
is beneficial for high-resolution AM-AFM images of soft
biological matter to measure in the attractive regime, i.e.,
before the instability occurs during tip–sample approach. In
order to stay in this regime, it has been proposed to artificially
increase the Q -factor of the dynamic system (cf section 2.1),
since the instability occurs at smaller A/ A0 ratios for larger
Q -values (i.e., ‘later’ during tip–sample approach), where
A represents the actual oscillation amplitude and A0 the
oscillation amplitude of the free cantilever [16]. This makes
it easier to establish stable imaging conditions in the attractive
regime. For its practical implementation, it is used that
the dynamic response behaviour of a system containing an
oscillating cantilever can be tuned by adding an external
feedback mechanism (see, e.g., [7, 16–18]), which is most
commonly referred to as ‘ Q -control’.
However, even seven years after Q -controlled AM-AFM
was first demonstrated by Anczykowski et al [16], there
is still a debate on whether or not an enhancement of the
effective quality factor by Q -control actually leads to an
improvement of the achievable image quality, even though this
has been frequently observed [7, 19–23]. While some authors
sample
D
PID
setpoint
x-y-z-scanner
Figure 1. Sketch of the experimental set-up of an atomic force
microscope operated in amplitude-modulation mode with Q -control.
This set-up differs from an atomic force microscope driven in
conventional AM-AFM mode (‘tapping mode’) by the introduction
of an additional feedback loop consisting of a phase shifter and a
variable gain amplifier.
cannot find any noticeable improvement in image quality [6],
others report that optimizing the scan parameters led to the
same improvement in image quality as the application of
Q -control for their samples [13, 24]. It is the purpose of
the present paper to advance this discussion by reporting
on systematically performed experiments in liquids on soft,
biologically relevant ‘benchmark samples’ with and without
active Q -control. We find significantly reduced heights
of specific topographical features in data recorded with
conventional AM-AFM compared to the corresponding feature
heights if measured with active Q -control. This observation
indicates substantial sample deformation in conventional AMAFM, corroborating the theory that Q -control reduces tip–
sample interactions even in liquids.
2. Experimental details
2.1. Experimental set-up
A commercial atomic force microscope (NanoScope IIIa with
MultiMode head, Veeco Instruments Inc.) equipped with a
liquid cell for acoustic excitation was used in our experiments.
A sketch of the experimental set-up is shown in figure 1.
The movement of the cantilever is detected by a displacement
sensor based on the laser beam deflection method. During
conventional AM-AFM operation, the cantilever vibration is
driven by an external function generator, while the oscillation
amplitude A and/or the phase shift ϕ are detected by a lockin amplifier. Thereby, the external function generator supplies
not only the signal for the excitation piezo, but its output serves
also as reference for the lock-in amplifier.
For Q -controlled AM-AFM, an additional feedback
circuit consisting mainly of a phase shifter and a variable
gain amplifier is added. We used a commercial system (‘ Q control’, nanoAnalytics GmbH), which is externally added
to the existing Nanoscope IIIa electronics. The working
principle is as follows: the signal of the displacement sensor
is amplified and, after the time delay between the cantilever
Imaging of biomaterials in liquids with conventional and Q -controlled AM-AFM
without Q-Control
with Q-Control
fit (Q = 34)
fit (Qeff = 202)
normalized amplitude
1.0
0.8
0.6
0.4
0.2
0.0
30.5
31.0
31.5
32.0
32.5
33.0
driving frequency (kHz)
Figure 2. Typical resonance curves obtained with and without active
Q -control in water. The solid lines are fits to the experimental data
plotted by symbols, which result in resonance frequencies f 0 and
Q -factors of f 0 = (31.733 ± 0.001) kHz and Q eff = 202.7 ± 2.2
with active Q -control and f 0 = (31.746 ± 0.007) kHz and
Q = 33.6 ± 1.2 without Q -control, respectively.
displacement and the amplified signal has been adjusted by
a phase shifter to a preset value, subsequently added to the
cantilever excitation voltage provided by the external function
generator (see figure 1). The basic idea of the circuit is
to compensate the damping forces which are affecting the
oscillation of the cantilever. Assuming sinusoidal cantilever
oscillation and choosing a time delay corresponding to 90◦
phase shift, it can be shown that the ‘effective Q -value’ Q eff
of the system containing both the cantilever and the Q -control
electronics can be intentionally increased or decreased [25]5 .
Figure 2 illustrates the increase of the effective Q -factor in
water. Both resonance curves and all other data sets presented
in this paper were measured with silicon cantilevers (TapMulti75, BudgetSensors) featuring nominal spring constants
of cz = 3 N m−1 and eigenfrequencies of f 0 = 75 kHz. The
symbols in figure 2 represent the measurements with (circles)
and without (squares) active Q -control. The solid lines are fits
to these data points, representing the Lorentzian shape of the
resonance peak of an externally driven harmonic oscillator,
Anorm = 1
1−
f d2 2
f 02
+
f d 2
Q f0
1
,
(1)
where Anorm represents the normalized oscillation amplitude.
Fitting this equation to the experimental data points, the
obtained Q -factors are Q eff = 203 with and Q = 34 without
Q -control6.
In order to overcome the criticism expressed in [24]
that only images acquired with optimum imaging parameters
5 This is in contrast to the ‘true’ Q -value of the cantilever itself, which is an
intrinsic property of the mechanical system.
6 We would like to point out that Q -factors obtained from the thermal noise
spectra are lower, ranging typically between 1 and 5 in water without Q -control
for this type of cantilever. The reason for this discrepancy can be assigned to
the fact that resonance curves measured with a fluid cell using a mechanical
driving mechanism essentially represent a convolution of the thermal noise
spectrum with the often complex resonance spectrum of the fluid cell, which
can make the resonance peak appear ‘sharper’ than it would actually be. For
a detailed discussion of this effect see Schäffer et al [26]. For a comparison
between conventional and Q -controlled AM-AFM, however, it is important to
contrast resonance curves that have been acquired under similar conditions, as
the Q -values obtained under these conditions are the ones that determine the
dynamical behaviour of the system.
should be used if a meaningful comparison between amplitudemodulation AFM with and without active Q -control should be
attempted, we recorded all images in the so-called ‘interleave
mode’ of the NanoScope software. In this mode, it is
possible to profile the same scan line with two different sets
of parameters by sampling during an initial run with the first
settings and during a second run with the other settings. At the
same time, the Q -control electronics can be turned on or off
for each run separately. Therefore, this technique enables us to
minimize drift effects and to obtain images at identical sample
positions in AM-AFM mode with Q -control on and off. We
recorded all images with setpoints and gains optimized for the
respective imaging mode at the resonant frequency of the freely
oscillating cantilever.
2.2. Sample preparation
In order to investigate typical applications in biological
research, we used two different types of sample. Lambda
phage DNA has been chosen as an example for high-resolution
imaging, while DPPC monolayers are intended to reflect a
simplified model of biological membranes.
Lambda phage DNA (methylated Escherichia coli host
strain W3110, Sigma-Aldrich Inc.) with a nominal length
of 48 kb (≈16 µm) was obtained as lyophilized powder.
The powder was allowed to dissolve in pure water (MilliQ , Millipore GmbH) for one day before we added bufferEB (Qiagen GmbH) to the solution. Small portions of this
solution (10 µg ml−1 DNA in 10 mM buffer-EB) were then
stored at −18 ◦ C for later use. Shortly before an individual
measurement was started, the DNA solution was further diluted
to a final concentration of 1 µg ml−1 in a buffer containing
1 mM EB and approximately 6 mM NiCl2 . We added
nickel ions because it has been reported that inorganic cations
like Ni(II) enhance the binding of the DNA to the mica
substrate [27]. Immediately before imaging a droplet of about
70 µl was pipetted onto a piece of freshly cleaved mica (Plano
GmbH). Measurements of the DNA adsorbed on the substrate
were performed with the Nanoscope fluid cell, but without an
additional O-ring.
Monolayers of DPPC (L-α -dipalmitoyl-phosphatidycholine, Fluka) were prepared with the Langmuir–Blodgett
technique. The films show a lateral structure of alternating
stripes and channels, which was obtained by rapidly withdrawing a mica substrate at a low monolayer surface pressure
and constant temperature as described by Gleiche et al [28].
The stripes consist of DPPC in a liquid condensed phase (LCphase), whereas the channels between this stripes are filled
with DPPC in the liquid expanded phase (LE-phase) [29]. The
periodicity of the stripes depends on the actual conditions during the preparation of the DPPC film.
3. Results
3.1. DNA in buffer solution
Figure 3 summarizes the results obtained during imaging of
lambda phage DNA adsorbed on a mica substrate (image
size: 600 × 600 nm2 ). The two images were simultaneously
obtained during measurement in solution (EB buffer) using the
interleave mode without (a) and with active Q -control (b). The
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D Ebeling et al
a)
a)
b)
b)
2 nm
3 nm
0 nm
0 nm
1
2
with Q-Control
1
2
conventional AM-AFM
1
0
0
0
100
200
300
400
500
x-position (nm)
100
200
300
400
1
1.0
1.5
2.0
2.5
0.0
0.5
1.0
1.5
2.0
2.5
x-position (µm)
500
x-position (nm)
Figure 3. Data acquired on DNA adsorbed on mica using
amplitude-modulation AFM without ((a), left column) and with
active Q -control ((b), right column). The images (top row) were
recorded by applying the interleave mode described in the
experimental section. The arrows in the images mark the positions
where the individual scan lines displayed in the bottom row were
obtained. For better comparability, the two scan lines are shifted
along the z -axis. The maximum height of the DNA measured with
active Q -control appears to be significantly higher ((1.6 ± 0.2) nm)
compared to the value measured without Q -control ((0.9 ± 0.2) nm).
free amplitude of the cantilever (measured with a gap of 1 µm
between cantilever and sample surface) was ≈0.53 V, which
is equivalent to about 6 nm. Activating Q -control increased
the original Q -factor of Q = 30 to an effective Q -factor of
Q eff = 256. The free amplitude of the cantilever was kept
constant. The scan rate was set to 1 Hz. To keep the tip–sample
interaction as low as possible, the highest possible values were
chosen for the setpoints in both modes (0.48 V for operation
without Q -control and 0.27 V for operation with active Q control). In this context, ‘highest possible’ refers to the fact
that with setpoints higher than the chosen ones, no clearly
visible contrast could be achieved. It should additionally be
noted that the gains were also individually optimized for both
modes.
A comparison of the two resulting images obtained with
and without active Q -control reveals an increase of the
topographic contrast by using Q -control. This originates from
a corresponding increase of the apparent structural height of
the DNA in the Q -controlled image, which is illustrated in
the bottom row of figure 3 by the two scan lines taken from
the images in the top row at the positions indicated by the
arrows. By statistically averaging data obtained from several
different scan lines, we found that the apparent height of the
DNA measured without Q -control is about (0.9 ± 0.2) nm,
while it grows to (1.6 ± 0.2) nm with active Q -control.
3.2. DPPC in n-decane and water
An increase of the apparent structural height of topographical
features is also observed on mono- and bilayers of DPPC thin
films. Figure 4 shows a DPPC sample imaged in n-decane
(Sigma-Aldrich Inc.). A comparison with images recorded
in air (not shown) confirms that the nonpolar n-decane has
no visible effect on the lateral structure of the DPPC. As in
S224
0.5
x-position (µm)
0
with Q-Control
0
0.0
0
height (nm)
conventional AM-AFM
height (nm)
2
height (nm)
height (nm)
2
Figure 4. A monolayer of DPPC supported by a mica substrate and
imaged in decane. The topographical height difference between the
two terraces as measured with conventional amplitude-modulated
AFM (a) is lower compared to the one measured while the Q -control
electronics was switched on (b). The scan lines plotted in the bottom
row reveal a height difference of (0.8 ± 0.2) and (1.1 ± 0.2) nm
without and with active Q -control, respectively.
the previous example, data acquisition was carried out with an
open fluid cell.
With this setup, the Q -factor was about Q = 26 without
Q enhancement, while it grew to Q eff = 223 with active Q control. The free amplitude of the cantilever was 0.66 V for
both modes; setpoints were 0.60 V without and 0.25 V with
active Q -control. Analysis of the two images, which have been
obtained with a scan rate of 0.5 Hz, reveals a height difference
between the vertical stripes and the channels of the DPPC film
of (0.8 ± 0.2) and (1.1 ± 0.2) nm without and with active Q control, respectively.
Interestingly, measuring in water manifests a significantly
different film structure (see figure 5). We attribute this
discrepancy to the following effect: DPPC is a phospholipid
with a hydrophilic head group and two hydrophobic alkyl
chains. If the monomolecular layer gets into contact with a
polar liquid like water, the hydrophobic alkyl chains will avoid
the contact with the aqueous environment and form an internal
hydrophobic phase. This behaviour leads to a reorganization
of the DPPC monolayer to islands with bimolecular thickness.
Nevertheless, an increase of the measured film height is
also observed for the DPPC bilayers imaged in pure water.
Figures 5(a) and (b) show a 7 × 7 µm2 large area of the DPPC
sample. The data displayed in (a) were recorded applying
conventional AM-AFM with a Q -factor of 39, while the data
shown in (b) were taken with active Q -control and an increased
Q -factor of Q eff = 209. The free amplitudes were 0.67 and
0.64 V, respectively. After optimizing the gains, both images
were scanned with a setpoint of 0.5 V and a scan rate of 0.5 Hz.
As before, the two scan lines displayed in the bottom row
of figure 5 reveal a significant change in the apparent height
difference between the individual terraces if measured with
and without active Q -control. In conventional AM-AFM, the
apparent bilayer height is (3.3 ± 0.4) nm, which is significantly
less that the value of (5.2 ± 0.4) nm measured with active Q control. Note that the apparent bilayer height measured with
active Q -control is only slightly smaller than the value of 6 nm
observed in previous studies using contact mode [30, 31].
Imaging of biomaterials in liquids with conventional and Q -controlled AM-AFM
a)
b)
8 nm
0 nm
conventional AM-AFM
4
2
with Q-Control
6
height (nm)
height (nm)
6
0
4
2
0
0
2
4
6
x-position (µm)
0
2
4
6
x-position (µm)
Figure 5. DPPC bilayers on mica imaged in pure water without (a)
and with active Q -control (b). The scan lines plotted in the bottom
row were obtained at the positions marked by the arrows in the top
row images. The measured apparent bilayer height in (a) is
(3.3 ± 0.4) nm and grows to (5.2 ± 0.4) nm in (b) when Q -control is
switched on.
4. Discussion and conclusion
It is well known that the apparent topographical height
of DNA adsorbed on mica and measured by AM-AFM is
typically smaller than the theoretical value of 2 nm (see,
e.g., [22, 32–35]). While various reasons, such as molecular
adhesion with the substrate or different strengths of tip–sample
interactions on the DNA as compared to the mica substrate,
might play a role (see [22] for a more complete discussion),
it is commonly believed that this effect is mainly caused by
the elastic deformation of the soft DNA molecule due to
repulsive forces exerted by the tip. Therefore, it is plausible
that the main reason leading to the observed increase in the
apparent topographical height of the DNA molecule if imaged
with active Q -control is a reduction of tip–sample interaction
forces. This is consistent with the work of Humphris et al [20]
and Pignataro et al [22], who also observed an increase of the
apparent height of DNA on mica using Q -control in liquids and
air, respectively.
Similar arguments apply for the explanation of the change
of the apparent height the DPPC films. For LB films, the
apparent layer height as a function of the applied load was
systematically examined by Hartig et al [36]. They reported
that the correct film thickness could only be measured in
contact mode by minimizing the tip–sample force, while
measurements in AM-AFM without Q -control always led to
decreased film thicknesses. Therefore, it is interesting to
note that film thicknesses measured in the present study with
active Q -control are systematically higher than the ones found
in conventional AM-AFM mode. Again, the most likely
explanation for this result is that the application of Q -control
actually reduces the tip–sample interaction forces.
It would certainly be interesting to unambiguously
determine whether or not imaging is performed in the attractive
or repulsive regime under our optimized conditions. However,
in contrast to experiments done in ambient conditions we did
not observe an instability which marks the transition between
the ‘repulsive’ and ‘attractive’ regime [13, 40]. Nonetheless,
since adhesion forces are typically significantly reduced in
liquids, we assume that the net forces between tip and sample
are repulsive, but with significantly reduced repulsive forces
with Q -control.
Despite the positive outcome that the alteration of
the sample topography by tip–sample interactions has been
considerably reduced applying Q -control in our experiments,
we nevertheless have to review possible drawbacks and
alternatives to the Q enhancement by means of an active
feedback. First, increasing Q potentially decreases the
maximum achievable scan rates [37]. In fact, the reverse
effect was used by Sulcheck et al [38, 39]: they decreased
the Q -factor in order to increase the scan speed under ambient
conditions. In contrast, we do not observe a significant
limitation of the scan rate in liquids for a moderate increase
of the Q -factors. Activating Q -control and increasing the Q factor by a factor of 5–10 ( Q < 300), the scan rate is still in
the range of 1 Hz.
Second, it has been discussed in the literature that the
reduction of tip–sample interaction forces in air might be also
achieved by using driving frequencies slightly larger than the
resonance frequency [13]. However, a significant drawback of
this approach is that it might be difficult to determine where
exactly the excitation frequency should be chosen for best
results. We found it much more straightforward and convenient
to simply choose the maximum of the main peak and to
activate Q -control. In this case, the tip–sample forces can
be considerably decreased by driving the system at the actual
resonance frequency without a further search for an optimum
driving frequency.
In summary, we investigated atomic force microscopy in
liquids using conventional (‘tapping mode’) and Q -controlled
amplitude modulation AFM. On analysing the resulting
apparent topography of soft samples like DNA and DPPC
thin films obtained with individually optimized parameters,
we observed a significant height difference measured in the
two operation modes. The height measured with Q -control
is reproducibly higher compared to conventional AM-AFM. In
agreement with previous studies this effect can be attributed to
the reduction of tip–sample forces by Q -control.
Acknowledgments
The authors would like to acknowledge valuable discussion
with Tilman Schäffer about Q -factors in liquids. Furthermore,
we thank Oliver Panzer, Xiaodong Chen, Michael Hirtz, Lifeng
Chi, and Ulli Fischer for their assistance with the preparation
of the samples. Helpful support from Marcus Schäfer, JanErik Schmutz, and Jens Falter is gratefully acknowledged. This
project was financially support by the German Federal Ministry
of Education and Research (BMBF) (Grant No. 03N8704).
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