Erik Jonsson School of Engineering and Computer Science
2013-10-04
Nanomechanical Imaging of Soft Samples in Liquid
Using Atomic Force Microscopy
UTD AUTHOR(S): Majid Minary-Jolandan
©2013 AIP Publishing LLC
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Nanomechanical imaging of soft samples in liquid using atomic force microscopy
Majid Minary-Jolandan and Min-Feng Yu
Citation: Journal of Applied Physics 114, 134313 (2013); doi: 10.1063/1.4824080
View online: http://dx.doi.org/10.1063/1.4824080
View Table of Contents: http://scitation.aip.org/content/aip/journal/jap/114/13?ver=pdfcov
Published by the AIP Publishing
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JOURNAL OF APPLIED PHYSICS 114, 134313 (2013)
Nanomechanical imaging of soft samples in liquid using atomic force
microscopy
Majid Minary-Jolandan1,a) and Min-Feng Yu2
1
Department of Mechanical Engineering, The University of Texas at Dallas, Richardson, Texas 75080, USA
Department of Aerospace Engineering, Georgia Institute of Technology, Atlanta, Georgia 30332, USA
2
(Received 24 July 2013; accepted 16 September 2013; published online 4 October 2013)
The widely used dynamic mode atomic force microscopy (AFM) suffers severe sensitivity
degradation and noise increase when operated in liquid. The large hydrodynamic drag between
the oscillating AFM cantilever and the surrounding liquid overwhelms the dissipative tip-sample
interaction forces that are employed for nanomechanical imaging. In this article, we show that the
recently developed Trolling-Mode AFM based on a nanoneedle probe can resolve nanomechanical
properties on soft samples in liquid, enabled by the significantly reduced hydrodynamic drag
between the cantilever and the liquid. The performance of the method was demonstrated by
C 2013 AIP Publishing LLC.
mapping mechanical properties of the membrane of living HeLa cells. V
[http://dx.doi.org/10.1063/1.4824080]
INTRODUCTION
As an indispensable nanoscale tool for experimental
studies, atomic force microscope (AFM) has been widely
applied across diverse scientific fields for materials characterization and surface engineering, providing nanoscale
spatial-resolution in mapping and patterning surfaces and
realizing pico-Newton sensitivity sensing of nanoscale interactions.1 By employing a nanoscale probe tip to physically
sense the surface of interest, one of the most significant benefits of AFM, over other characterization techniques, is that
it provides a direct means to access and interrogate the nanoscale site of interest mechanically, electrically, and/or optically by simply integrating the required functionalities into
the AFM tip.
This direct access to the sample of interest with AFM is
of significant value to biological studies as well, especially
at the cellular level. Various elegant methods2–13 based on
the scanning probe techniques, such as active Q-control, frequency modulation, and the scanning ion conductance microscopy (SICM), have been developed in the past decades
for the sole purpose of improving the imaging resolution of
biological samples, particularly mammalian cells, in a liquid
medium. However, simply due to the extreme mechanical
softness of such cells (on the order of several kPa or less in
membrane elasticity, much softer than some other cell types
with a cell wall, such as bacteria),3,14–17 it remains to be a
major challenge for AFM to image live mammalian cells and
achieve high resolution while avoiding excessive deformation. Interested reader may refer to the images of living cells
obtained in conventional AFM imaging mode in the following literature, where the cell membrane is often excessively
deformed and the underlying cytoskeleton is visible in the
acquired images.3,14,18–20 Being able to directly image and
resolve mechanical heterogeneities and microscale and
a)
Author to whom correspondence should be addressed. Electronic mail:
majid.minary@utdallas.edu
0021-8979/2013/114(13)/134313/6/$30.00
nanoscale features on a living cell membrane and to visualize the dynamics of the membrane proteins and lipid rafts
and their functional activities in vitro could facilitate answering many outstanding biological questions of cell and membrane physiology.21–23 In order to image such samples with
AFM, an AFM probe must exert as a minimal force as possible (down to several tens of pico-Newton) to avoid significantly deforming the cell membrane during imaging scan.
For this reason, the tapping mode (also called the dynamic
mode) AFM has been the operational mode of choice for
imaging biological samples.15,24–27
The tapping mode AFM, however, suffers a major problem when used for imaging biological samples in liquid, i.e.,
the low sensitivity of the AFM probe. In this mode, the AFM
probe is immersed in liquid to access the sample, and since
the hydrodynamic energy dissipation experienced by an
oscillating object scales with its characteristic dimensions,28
the relatively bulky AFM probe experiences large hydrodynamic drag. The Q-factor (defined as the ratio of the energy
stored in the resonator and the energy dissipated per cycle)
of the dynamic operation of the AFM probe drops from
around several hundred in air to the order of 1 in
liquid.29–32 The force applied onto the sample in the tapping
mode AFM is roughly proportional to the needed excitation
force on the AFM cantilever described by kA/Q, where k is
the spring constant, A is the free oscillation amplitude, and Q
is the Q-factor of the AFM cantilever.33 For a specific
dynamic operation at the designated oscillation amplitude, a
low Q-factor means high minimal tapping force for AFM
imaging. To more effectively minimize this tapping force, it
is desirable to have an “intrinsically” high Q-factor in the
dynamic system when operated in liquid.
One of the most explored methods for enhancement of
the Q-factor of AFM probe in liquid is the active Qcontrol,6,33–37 where an external feedback-control circuitry
dynamically boosts the resonance oscillations and tunes down
the non-resonance oscillations of the cantilever. Effective Qfactors over one thousand have been demonstrated using
114, 134313-1
C 2013 AIP Publishing LLC
V
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M. Minary-Jolandan and M.-F. Yu
active Q-control.38–42 However, the active Q-control does not
fundamentally change the damping “intrinsically” experienced by the oscillating AFM probe in liquid. Rather, it has
been shown that it amplifies the thermal noise and thus the
random thermal fluctuation forces experienced by the AFM
cantilever.17,43
In addition to the topographical imaging, mapping
viscoelastic surface properties of soft samples in liquid using
AFM without imposing significant deformation on the sample remains also a challenging task. The success of the highresolution viscoelastic mapping intimately depends on the
quality of the topographical image. If the sample is significantly deformed during imaging, the resolution in viscoelastic mapping is significantly degraded. For example, the
commonly used force modulation AFM method is incapable
of achieving high-resolution mapping and can only provide
overall viscoelastic properties by significantly deforming the
sample.44 Similarly, AFM-based nanoindentation of cells
provides bulk elastic or viscoelastic properties of the cell and
does not provide a continuous map of the surface.45,46 The
imaging contrast in nanomechanical mapping of sample surface in AFM is the result of dissipative interaction between
the probe tip and the sample surface.15 This interaction is often very small and can be resolved only if there is a significant spatial variation in the surface properties, unless a
sensitive probe is employed.
J. Appl. Phys. 114, 134313 (2013)
The overall damping experienced by an AFM cantilever
system can be simply minimized by reducing its interaction
cross-section with the fluid environment. To realize this
advantage while maintaining the general flexibility of an
AFM system for imaging biological samples in liquid, we
recently presented a theoretical and experimental study
based on a resonating nanowire partially submerged in liquid
and showed that such a system can achieve an intrinsic high
Q-factor dynamic AFM in liquid. We showed that this
method, termed the “Trolling-Mode,” shows 2–3 orders of
magnitude improvement in sensitivity over the current
dynamic AFM methods in liquid and maintains the harmonic
dynamic response of the cantilever in air, thus significantly
reduces the minimal imaging tapping force while improving
the signal-to-noise ratio performance of the dynamic AFM
system, Fig. 1(a).28
In this article we show that, in addition to the topographical imaging, this method is capable of mapping nanomechanical properties of soft samples in liquid. We show that
such a system with an intrinsically high Q-factor can more
sensitively differentiate the energy dissipations contributed
specifically by the tip-sample interactions without the overwhelming influence of the hydrodynamic contributions, and
thus provide high-resolution dynamic mapping of material
properties of the sample surface in liquid. This is enabled by
the significantly reduced energy dissipation between the
FIG. 1. (a) Schematic of the “Trolling-Mode” AFM setup. A solid and mechanically stable nanoneedle functions as an extend tip to scan a sample submerged
under a shallow liquid medium. The cantilever system is kept in air and hence, it retains its high intrinsic Q-factor performance. (b) SEM image shows a
60 lm long and 600 nm-diameter Pt nanowire directly deposited onto the AFM cantilever with the meniscus-confined electro-deposition technique. The
insets show the base of the nanoneedle on the cantilever (left) and the tip of the nanoneedle sharpened down to a radius of 25 nm with FIB milling (right).
The nanowire was grown in angle of 128 to compensate for the imaging angle in AFM. (c) The pull-in and pull-out force response of the nanoneedle probe
into/from water/air interface vs. arbitrary time-lapse. The first drop in the deflection response is the pull-in force and the larger drop is the pull-out force after
breaking the meniscus. (d) Acquired dynamic responses of the nanoneedle-probe in air before immersion into the liquid, and after immersions of the nanoneedle into liquid by 10 lm showing that the resonance frequency does not change. The Q-factor of the cantilever system is 450 in air and 110 in liquid. (e)
Schematic shows that when the soft cell membrane is imaged with a large tapping force, the membrane is deformed and topography is compromised. (f) When
the soft cell membrane is imaged with a smaller force, the membrane deformation is minimal and the true topography of the cell membrane, including protein
domains, could be imaged.
134313-3
M. Minary-Jolandan and M.-F. Yu
AFM probe and the surrounding liquid. To demonstrate the
performance of the method, we provide high-resolution
images of the topography, amplitude, and phase images
obtained on the membrane of living HeLa cells. The results
reveal the highly heterogeneous nature of the membrane of a
living cell, both in terms of topographical features and nanomechanical properties.
MATERIALS AND METHODS
We fabricated the nanoneedle AFM probes with the
meniscus-confined electrodeposition technique.47,48 Metallic
nanoneedles with desired diameters and lengths were
directly deposited onto commercial tapping mode AFM cantilevers having a nominal force constant of 6 N/m. For this
study, the results were obtained using Pt nanoneedles having
an aspect ratio of 100, a length of 60 lm, and a diameter of
600 nm. The tip of the nanoneedle was then sharpened
with focused ion beam (FIB) milling to attain a tip radius of
curvature of 15–25 nm (Fig. 1(b), inset). The sample and the
AFM cantilever system were enclosed in a humidity chamber49 under a 100% relative humidity to limit water evaporation. Static deflection response of the cantilever was
monitored using a signal access module in a LabView
program.
RESULTS
To image samples in liquid and map their nanomechanical properties, the interaction of the nanoneedle probe and
the liquid should be fully controlled and characterized.50 For
this purpose, we used an external LabView program to register the cantilever deflection as the nanoneedle proceeded toward the liquid surface, submerged into the liquid, and was
retracted from its surface. Figure 1(c) shows the typical
snap-in and snap-back behavior and the corresponding
changes in the deflection (force) of the cantilever as the
nanoneedle advances into and retracts out of the air/water
interface. The snap-in is the result of the instability of the
cantilever as the tip of the nanoneedle senses the long-range
van der Waals attractive forces between the water surface
and the tip. When the nanoneedle is inside the liquid, the
force on the cantilever can be calculated according to
F ¼ pcdcos ðhÞ. This equation relates the surface tension of
a liquid to the applied force on a sample with diameter d
(diameter of the nanoneedle). In this equation, c is the surface tension of the liquid and ðhÞ is the contact angle
between the nanoneedle (Pt) and water. When the probe is
retracted from the air/water interface, the deflection force
increases as the contact angle decreases to zero at the point
of meniscus break-down. At the point of meniscus breakdown the deflection force is maximum, F ¼ pcd, for h ¼ 0.
Experimentally obtained value of 120 nN is in a good
agreement with the calculated value from surface tension
equation (135 nN).
In liquid, the apparent mass of the cantilever increases
due to the added mass of the liquid oscillating along with the
cantilever. We oscillated the nanoneedle probe in the air and
after submerging the nanoneedle into liquid. Dynamic
responses of the probe in air and in liquid are shown in Fig.
J. Appl. Phys. 114, 134313 (2013)
1(d). The results show that the harmonic behavior of the cantilever was near perfectly preserved without the appearance
of any spurious peaks in liquid. In addition, there was no significant shift in the resonance frequency of the cantilever
indicating the minimal effect of the added mass onto the
nanoneedle, as expected based on the small size of the nanoneedle compared with the cantilever dimensions. This is an
indication that the cantilever in Trolling-Mode is operating
at near in-air conditions while the nanoneedle can image
samples in liquid.
The Q-factor was obtained for each curve by fitting the
resonance curve with a Lorentzian function of a single harmonic oscillator. The Q-factor was 450 in air and 110
when the nanoneedle was inserted into water by 10 lm.
Since the average tapping force hFts i applied onto the sample
in dynamic mode AFM is proportional to the inverse of the
Q-factor (hFts i / Qk ½A20 A2sp 1=2 ),17 enhancing the intrinsic
Q-factor in liquid from 1 in conventional tapping mode to
100 in Trolling-Mode results in at least two orders of magnitude reduction in the applied force on the sample. In addition to the Q-factor, the hydrodynamic dissipative forces
also decrease by 2–3 orders of magnitude in this technique.
The hydrodynamic damping coefficient in tapping mode
AFM in liquid is related to the dimensions of the cantilever,
cL ¼ q p4 W 2 LxL C00 , where q is the density of the liquid, W
and L are the width and length of the cantilever, and C00 is
the hydrodynamic function.29,30 For a typical AFM cantilever, the damping coefficient cL due to the hydrodynamic
interaction is estimated to be 8 106 Ns=m. For the
nanoneedle probe system, the dissipation is due to Stokes’
2pgL
)51 and the nanomeniscus dissipations
drag (cd ln ðL=rÞ0:72
(cm 2prg hlnd ðd=aÞ).52,53 In here, g is the dynamic viscosity of
the liquid, L and r are the length and the radius of the nanoneedle, respectively, hd is the dynamic contact angle
between liquid and the nanoneedle, d is the evanescent decay
length, and a is the liquid molecule size. The total damping
is then estimated to be 5 108 Ns/m, which is two orders
of magnitude smaller than that in the conventional tapping
mode operation.
Two-order of magnitude enhancement in Q-factor and
two-order of magnitude reductions in dissipative forces are
critical for gentle imaging of soft samples in liquid and mapping their nanomechanical properties. Schematic in Fig. 1(e)
shows that if a soft sample, such as a living cell, is imaged
under large scanning forces, the membrane of the cell is
deformed conformal to the underlying and more rigid cytoskeleton of the cell; hence, the obtained topography and the
nanomechanical properties are compromised. While if the
tapping force in minimized and intrinsic dissipations of the
system are reduced, un-deformed topography and hence
nanomechanical surface image properties could be obtained
(Fig. 1(f)).
After characterization of the behavior of the probe in
air/liquid interface, we imaged individual living HeLa cells
under physiological conditions. The acquired images from
an individual living HeLa cell in PBS (phosphate buffered
saline) are shown in Figure 2. The images were acquired at a
free oscillation amplitude of <20 nm, a set-point of 80%,
134313-4
M. Minary-Jolandan and M.-F. Yu
J. Appl. Phys. 114, 134313 (2013)
clearly resolve the nanomechanical heterogeneity on the cell
membrane. The resolved structures on the cell membrane are
several microns in length and a few hundred nanometers in
height. They clearly have different mechanical properties
compared with the surrounding area on the membrane. We
speculate that these regions may be protein domains on the
cell membrane, which are mechanically more rigid than the
lipid bilayer. Another interesting feature is the appearance of
an elliptical region in the phase image in Fig. 2(c). It seems
that this region is the area above the cell nucleus, which has
different mechanical properties than the rest of the cell, and
provides a strong contrast in the phase image. Figs. 2(g) and
2(h) show a line-scan along the dashed line shown in Fig.
2(c). These line-scans show that the area covered by the cell
shows higher height and an overall larger phase shift with
respect to the surrounding substrate, indication of the more
viscoelastic nature of the living cells.
Figures 3(a)–3(c) show three consecutive 10 lm2 topography image scans over the same area on the membrane. The
topographic images, besides showing the reproducible presence of some microscale features (protrusions having a
height of 50–400 nm above the membrane in the images,
presumably membrane proteins, or lipid domains) and their
partially confined presence in their preferred regions on the
membrane surface, showed also the dynamic or mobile nature of some other surface features evidenced by their relative displacements from their original locations in those
consecutively acquired images.
FIG. 2. (a) and (d) topography, (b) and (e) amplitude, and (c) and (f) phase
images of the whole cell and high magnification of the squared area in (b),
respectively. The delineated area in (c) shows the traces of cell nucleus
resolved in the phase image. RMS roughness obtained from image analysis
in (d) is about 114 nm. The phase and amplitude image, which are related
to nanomechanical properties, show that the cell membrane is highly heterogeneous at the nanoscale. (g) and (h) are line-profile of the height and phase
marked by the vertical dashed line in (c). The area covered by the cell shows
higher height and an overall larger phase shift with respect to the surrounding substrate, indicated by the difference in the horizontal dashed lines.
and a scan rate of 1 Hz. Topography images of the cells were
mapped following the natural membrane contour and showed
no obvious deformation of the cell evidenced by the lack of
ostensible protrusions due to the presence of the cytoskeleton
structures (Fig. 2(a)). Amplitude and phase image in
dynamic AFM are related to the local elastic and viscoelastic
properties of the surface.15,26 Figs. 2(b) and 2(c) show the
amplitude and the phase images of the cell membrane,
respectively. Due to the significantly reduced hydrodynamic
damping, our technique enabled differentiating the delicate
tip-sample interactions in different regions even on a soft
cell membrane, instead of being overwhelmed by phase
noises from the dominant hydrodynamic damping as in the
conventional AFM imaging method in liquid. These images
show that the cell membrane is highly heterogeneous in its
mechanical properties at the nanoscale.
Figs. 2(d)–2(f) are the higher magnification topography,
amplitude, and phase images of the area marked with the
square in Fig. 2(b). These high magnification images more
FIG. 3. (a)–(c) are three consecutive 10 lm 10 lm topography images
acquired at the same area on the cell membrane on a living HeLa cell in
physiological conditions. Images overall show the reproducibility of the
imaging procedure. As the cell membrane is dynamic, there is apparent
domain-movement in consecutives images. (d) and (e) are higher magnifications images of the areas enclosed in boxes in (a). (f) Shows the distribution
of the height of the domains in (a), which is a linear distribution with a peak
value of 411 nm.
134313-5
M. Minary-Jolandan and M.-F. Yu
Figures 3(d) and 3(e) show higher magnification images
of the areas boxed in Fig. 3(a). Image analysis was performed on various features in acquired images. Due to the
tip/sample convolution effect in AFM, features appear wider
in lateral direction than their actual size. We measured the
size of the features at full-height and in half-height of the
peak for each feature. On average, feature sizes as small as
43.8 nm on full height and 103 nm on half-height were
resolved in the images. Figure 3(f) shows the distribution of
the height of the domains in Fig. 3(a), which is a linear distribution with a peak value of 411 nm.
DISCUSSION
The membrane of a living cell, especially the cancerous
type HeLa cell, is recognized to be extremely soft, having an
effective elastic modulus in the 1 kPa range. Considering the
importance of membrane domains in cell physiology, developing methods to image their topographical and nanomechanical properties are critical. AFM has the potential to achieve
this goal, however despite much progress, high resolution
imaging of soft samples in liquid using AFM has been elusive.
In previous AFM imaging studies of cells with the dynamic
mode AFM, the cells were often severely deformed by the
tapping force from the AFM tip during the imaging process.14
The technique presented in this article shows a great
potential for simultaneous mapping of topographical and
nanomechanical properties on the cell membrane. This is
enabled by the significantly reduced hydrodynamic damping
in our technique enabling differentiation of the delicate tipsample interactions in different regions on a soft cell membrane, instead of being overwhelmed by phase noises from
the dominant hydrodynamic damping as in the conventional
AFM imaging method in liquid. Another advantage of the
Trolling-Mode is that it can be combined with other AFM
techniques, such as high-speed AFM, frequency modulation,
and multi-frequency to obtain fast dynamic processes at the
cell membrane, such as lateral motion of lipid rafts and protein domains. Combining this method with optical methods,
such as fluorescent microscopy, would be also a powerful
method for biological studies at the single cell level. The presented method requires further development to achieve higher
resolution and more sensitive performance. Resolution of the
obtained images could be improved by using smaller diameter nanoneedles to even further reduce the intrinsic dissipation
of the system to enable more sensitive mapping of nanomechanical surface properties.
Based on equation hFts i / Qk ½A20 A2sp 1=2 , the applied
force on the sample has an inverse relationship with the quality
factor, and a direct relationship with the spring constant of the
cantilever. Decreasing the free amplitude and the set-point amplitude decreases the applied force on the sample. In dynamic
AFM operation, the spring constants are often >5 N/m, since
the soft cantilevers used for contact mode do not generate reliable dynamic response. By reducing the spring constant, the
applied force could be further reduced on the sample.
However, for biological samples, such as living cells, reducing
the oscillation amplitude does not necessarily result in
improvement in image quality due to loss in sensitivity.
J. Appl. Phys. 114, 134313 (2013)
CONCLUSIONS
In conclusion, we demonstrate the potential of the
Trolling-Mode AFM for mapping nanomechanical properties
on a cell membrane. The technique preserves the harmonic
dynamic response of the cantilever and has an intrinsic high
Q-factor in liquid without the need of active Q-control.
Enabled by the small interaction cross-section with the surrounding liquid, it introduces minimal mechanical disturbance to the surrounding liquid and the sample. These
combinations result in inherently high signal-to-noise ratio
performance of the system for acquiring high-resolution
AFM images and higher sensitivity to damping associated
with tip-sample interactions in liquid environment. The
acquired images reveal the highly non-uniform and heterogeneous structure of the cell membrane and the existence of
near membrane structures.
ACKNOWLEDGMENTS
The work is supported by NSF Grant Nos. CMMI0726878 and CBET-0731096.
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