1. A new integrated system combining atomic force microscopy
and micro-electrode array for measuring the mechanical
properties of living cardiac myocytes
Jose F. Saenz Cogollo & Mariateresa Tedesco &
Sergio Martinoia & Roberto Raiteri
Published online: 1 April 2011
# Springer Science+Business Media, LLC 2011
Abstract In this paper we present a new experimental set-up
which combines the surface characterization capabilities of
atomic force microscopy at the sub-micrometer scale with
non-invasive electrophysiological measurements obtained by
using planar micro-electrode arrays. In order to show the
potential of the combined measurements we studied the
changes in cell topography and elastic properties of cardiac
muscle cells as during the contraction-relaxation cycle. The
onset of each beating cycle was precisely identified by the use
of the extracellular potential signal, allowing us to combine
nanomechanical measurements from multiple cardiomyocyte
contractions in order to analyze the time-dependent variation
of cell morphology and elasticity. Moreover, by estimating the
elastic modulus at different indentation depths in a single
location on the cell membrane, we observed a dynamic
mechanical behavior that could be related to the underlying
myofibrillar structure dynamics.
Keywords Atomic force microscopy. Micro-electrode
arrays . Cardiac myocytes . Transverse stiffness .
Nanoindentation . Elasticity
1 Introduction
During each cardiac cycle, the sub-cellular structures of
cardiac myocytes undergo significant changes that define
the contractile properties of the heart muscle and largely
determine the physiological and pathological aspects of the
cardiac function (Parker and Ingber 2007). Dynamic
mechanical properties of the myocardium have been studied
in great details at the organ and tissue level (Miyaji et al.
1998; Shishido et al. 1998; Konofagou et al. 2002;
Kolipaka et al. 2009). However, in relevant pathological
conditions, like heart failure, a deeper knowledge of sub-
cellular and molecular myocardial properties is needed in
order to understand the origin of the clinical and hemody-
namic features of the disease and to develop appropriated
therapies (Borbely et al. 2005; Hamdani et al. 2008; van
Dijk et al. 2008).
The recent advances in micro- and nano-technology
allowed to study the axial mechanical properties of cardiac
myocytes in relation to force generation at the whole cell
level (Nishimura et al. 2004; Borbely et al. 2005; Iribe et al.
2007), however the study of myocardial transverse
mechanical properties, important in the active force
generation (Halperin et al. 1987), has been limited mainly
to chemically stabilized conditions in order to avoid the
experimental and data analysis challenges associated with
actively beating cells (Hofmann et al. 1997; Mathur et al.
2001; Lieber et al. 2004). Atomic force microscopy (AFM)
can be used to study the transverse sub-cellular mechanical
properties of living cells in general and cardiac myocytes in
particular, as pioneered by Radmacher and coworkers
(Radmacher, 1997; Hofmann et al. 1997). It allows the
application and the measurement of very low forces with
nanometer spatial resolution and minimal disruption of the
cell membrane (Kuznetsova et al. 2007). A recent study
made by Azeloglu and Costa (Azeloglu and Costa 2010)
showed the feasibility of using AFM to quantify the
dynamic elastic properties of actively beating cardiomyo-
cytes at the sub-cellular level. This was achieved by
synchronizing optical video microscopy recordings of the
J. F. Saenz Cogollo :M. Tedesco :S. Martinoia :R. Raiteri (*)
Department of Biophysical and Electronic Engineering—DIBE,
Università di Genova,
Via Opera Pia 11a,
16145 Genova, Italy
e-mail: rr@unige.it
Biomed Microdevices (2011) 13:613–621
DOI 10.1007/s10544-011-9531-9

2. motion (contraction cycle) of a compact layer of cells with
AFM operation. Because different cells in the same culture
dish can contract with different timing and dynamics and
since it is not possible to identify a clear initiation of each
single cycle from the optical image, a low temporal
resolution could be obtained even when high-speed CCD
imaging was employed, hence only a rough separation of
the systolic (cell contraction) from the diastolic (cell
relaxation) measurement was reported.
Here we propose to use, as a reliable time reference for
the beating cycle, the cardiomyocyte action potential (AP).
It is known that voltage-clamp methods allow a local and
precise recording of the AP, however they greatly affect the
cell mechanics and damage the sub-membranous cytoskel-
eton and its connections to the actin cortex (Merkel et al.
2000; Charras et al. 2004). Voltage sensitive dyes allow fast
two-dimensional electrical recordings, however they are
sensible to motion artifacts and are toxic for the cell, which
limits the duration of the experiment. On the other hand, it
is possible to perform extracellular recording of the AP
from contracting cardiac myocytes without interfering with
cells motility or producing undesirable side effects when
the cells are grown on glass surfaces with integrated micro-
electrode arrays (MEAs) (Egert and Meyer 2005). By
providing non-invasive, simultaneous multi-site, extracel-
lular recording from cardiac myocytes, the MEAs allow the
recording of the electrical activity and avoid mechanical or
chemical alterations of the cell physiology. A first attempt
of coupling MEA with AFM was proposed to study
morphology profile changes in neuroblastoma cultured
cells when they are electrically stimulated (Shenai et al.
2004).
In this paper, we report the development of a novel
AFM-MEA platform that allows to measure minimal
changes in the morphology and in the mechanical proper-
ties of living cardiac myocytes while recording their
extracellular potential (Fig. 1). In the development of the
AFM-MEA platform, we decided to mechanically adapt
commercially available devices and couple them with an
optical microscope in order to be able to position the
AFM tip onto any location of the MEA with an
acceptable level of electrical and mechanical noise during
the electrophysiological and AFM measurement, respec-
tively. We also included a perfusion system and a
temperature control, necessary for maintaining suitable
environment for cell viability. Next, we integrated the
AFM and MEA signals into a single acquisition software
with a common user interface for ensuring the synchro-
nization of the measurements. Finally we applied this
integrated set-up to perform nanoindentation measure-
ments onto single cardiac myocytes while measuring
extracellular potential changes. This approach allowed us
to quantify and follow, for the first time to the best of
our knowledge, the changes in cardiomyocyte mechanical
properties during the contraction-relaxation cycle with
high time and spatial resolution.
2 Materials and methods
2.1 Coupling the AFM with the MEA
The AFM-MEA platform is based on a commercial
atomic force microscope, (Agilent 5500, Agilent Tech-
nologies, Tempe, AZ) equipped with a scanner capable
of 100×100 μm scan range in X-Y and 7 μm scan range
in Z direction. The scanner has X-Y-Z position sensors in
order to compensate artifacts in its movements such has
creep, non linearity, etc. We employed commercial
MEAs (Multi-Channel Systems, Reutlingen, Germany)
consisting in 60 planar microelectrodes (30 μm diameter
electrodes and 200 μm pitch distance in an 8×8 config-
uration) embedded onto a glass surface. The contact pads
and tracks are made of transparent Indium Tin Oxide
Fig. 1 (a) Schematic illustration of the combined AFM-MEA approach. (b) bright field image of an AFM cantilever with the tip indenting a
beating cardiomyocyte near a planar microelectrode (scale bar: 30 μm). The white cross indicates the position of the tip on the cantilever
614 Biomed Microdevices (2011) 13:613–621

3. (ITO) whereas surface of the electrodes is covered with a
thin layer of titanium nitride (TiN).
The AFM head sits onto a custom-made MEA stage
while both are positioned onto an inverted optical micro-
scope (Olympus IX70, Olympus, Tokyo, Japan). The
custom-made MEA stage has micromanipulators that,
together with the step motors of the AFM head, allow the
X-Y-Z positioning of the AFM scanner over the MEA (see
Fig. 2); another sample stage allows X-Y movements of the
whole system over the objective of the optical microscope.
The MEA stage is also equipped with four 1.25 W adhesive
heater mats and one RTD temperature sensor in order to
keep a constant temperature inside the MEA chamber.
2.2 Data acquisition
A simplified block diagram of the AFM-MEA platform is
shown in Fig. 3. From the custom-made MEA connector a
68-pin high-grade cable links the microlectrodes to a 64-
channel filter amplifier (Multi-Channel Systems, Reutlin-
gen, Germany) with a gain of 1,000 and a band-pass
between 10 Hz and 10,000 Hz. The 60 amplified and
filtered signals are acquired together with the AFM
cantilever deflection signal and the X-Y-Z position of the
piezo by a 64-channel Data Acquisition Card (model
NI6071E, National Instruments, Austin, TX). Custom
routines written in LabView (National Instruments, Austin,
TX) and Matlab (MathWorks, Natick, MA) were used for
the data acquisition and the signal analysis respectively. By
using a threshold based real-time peak detection algorithm
for the recorded electrical signal, a high speed Ethernet link
between the computer that acquires the signals and the
AFM controller, and a specific library for controlling the
AFM software, it was possible to synchronize the AFM
operation with the cells contraction-relaxation timing.
2.3 Cell culture
Cardiac myocytes were isolated from 16 to 19 days rat
embryos, purified by differential attachment technique, and
then plated and grown as confluent monolayer on the MEA
surface as described elsewhere (Rapila et al. 2008). Briefly,
the hearts were dissected out and transferred to ice-cold
buffer solution containing 100 mM NaCl, 10 mM KCl,
1.2 mM KH2PO4, 4 mM MgSO4, 50 mM Taurine, 20 mM
Glucose, 10 mM Hepes, pH 7.0. Ventricles were separated
from atria. The ventricular tissue, minced into 1–3 mm3
pieces, was enzimatically digested in 0.2% pancreatin and
0.2% collagenase type II and diluted in the same buffer
solution already used during the previous step. Tissue
fragments were incubated at 37°C for 15–18 min and the
cell suspension, after a gentle mechanical dissociation, was
transferred into a conical centrifuge tube with nutrient
medium DMEM and 10% FBS to block enzymatic activity.
This cycle of digestion was repeated twice; afterward all the
cell suspension collected during the two cycles was
centrifuged for 8 min at 1,000 rpm. The pellet was re-
suspended in DMEM-M199 (4:1), 6% HS, 4% FBS, 1%
Glutamax, 10 μg•ml−1
Gentamycin, and pre-plated onto a
Petri dish (100 mm diameter) for 2 h a 37°C in 5% CO2
humidified incubator in order to separate cardiac fibroblast
cells from cardiomyocytes, which have a slower time of
adhesion and thus remained in the culture medium
suspension. Finally, ventricular myocytes were collected,
centrifuged, re-suspended in culture medium, and 3.104
–
6.104
cells were plated onto the MEA electrodes area,
Fig. 2 (a) Simplified drawing of the AFM-MEA platform. Micro-
manipulators in the MEA stage together with the step motors of AFM
head allow the precise X-Y-Z positioning of the AFM scanner over the
MEA device, while X-Y movements of the optical microscope stage
allow positioning the whole system over the microscope objective. (b)
Photograph of the AFM-MEA platform mounted onto the inverted
optical microscope
Biomed Microdevices (2011) 13:613–621 615

4. which had been previously sterilized and incubated with a
solution of laminin at a concentration of 50 μg/ml. All
reagents used for cell culture preparation were from
Invitrogen or Sigma-Aldrich. Experiments were done after
3 days in vitro (DIV).
2.4 Experimental protocol
Repeated force versus Z-displacement curves (force
curves) (Butt et al. 2005) were performed and the region
of the curves after contact (indentation curve) was
considered in order to measure the changes in cell stiffness
during the cardiomyocyte beating cycle. Rectangular AFM
cantilevers (model CSC38, MikroMash, Tallinn, Estonia)
with 0.03 N/m nominal spring constant and conical tip
were used. Before each experiment the deflection sensi-
tivity of the AFM photodetector was determined by taking
force curves against a flat and hard surface, while the
actual spring constant of the cantilever was calculated
using the Sader method (Sader et al. 1995). The selection
criteria of the cells to be probed were the following: cell
position close or on top of an electrode, visible spontane-
ous beating activity, and high signal-to-noise ratio of the
recorded electrical signal. We usually indented the central
region of the cell for 60 s with force curves taken at
intervals of ∼400 ms, with a maximum applied load of
15 nN, and a constant vertical speed of nearly 130 μm/s.
The resulted indentation time was typically less than
30 ms. We always verified that the tip completely detached
from the cell membrane after each loading/unloading
cycle. All experiments were carried out at a controlled
37°C temperature and fresh culture medium was contin-
uously perfused at 100 μl/min through the measurement
chamber using a peristaltic pump.
2.5 Indentation data analysis
AFM raw cantilever deflection and Z position signals were
used to identify and separate the loading and unloading region
of each indentation event. For each force curve the point of
contact (i.e. the Z-piezo displacement corresponding to the tip
touching the surface at zero applied force) was automatically
identified by fitting a bi-domain linear-quadratic function
(linear fitting before contact and quadratic fitting after contact)
by least squares optimization (see Fig. 4). The calculated
point of contact was used to calculate the change in the
height of the cell and to obtain the indentation curve. In
Fig. 3 Block diagram of the
AFM-MEA platform. Signals
from the 60 micro-electrodes are
filtered, amplified, and acquired
together with the signals from
the AFM, using an A/D card.
The acquisition software runs a
real-time peak-detection algo-
rithm that allows the synchroni-
zation, via a fast Ethernet link to
the AFM controller, of the AFM
indentations with the cell elec-
trical activity
Fig. 4 Typical AFM force curve obtained by pressing against a
cardiac myocyte. Only the part corresponding to the tip approaching
the cell and indenting it is shown. A bi-domain linear-quadratic fitting
enables the automatic detection of the tip-cell contact point
616 Biomed Microdevices (2011) 13:613–621

5. order to characterize the mechanical properties of the cell
under study we estimated the transverse dynamic elastic
modulus based on the analysis proposed by Oliver and Parr
(1992) and described elsewhere (VanLandingham et al.
2001); such value can be considered only an estimation
since one expects viscoelastic and heterogeneous properties
of the cardiac myocytes. Briefly, loading or unloading parts
of the indentation curves (we did not observe hysteresis
between loading and unloading and between successive
indentations due to plastic deformations) were fitted using a
spline (see Fig. 5(c)) in order to calculate the slope of the
indentation curve dPðhÞ
dh (i.e. the contact stiffness) at different
indentation depths. The real part E(h) of the cell elastic
modulus can be estimated by:
EðhÞ ¼
1
2
ffiffiffi
p
p
ffiffiffiffiffiffiffiffiffiffi
AðhÞ
p ð1 À u2
Þ
dPðhÞ
dh
ð1Þ
where A(h) is the projected tip contact area at indentation h, P
(h) is the instantaneous force and υ is the Poisson’s ratio. A
value of υ=0.5 was used assuming the cell as incompressible.
During the analysis, each indentation curve was tagged
with the delay with respect to the last spontaneous extracel-
lular AP peak measured at the nearest microelectrode, taking
into account the indentation contact time. Data obtained from
all indentations taken onto a single cell during an interval of
60 s were pooled together to obtain the traces of cardiomyo-
cyte elasticity versus beating cycle timing.
3 Results and discussion
We first verified that the combined AFM-MEA set-up does
not compromise the capability neither of the AFM to image
and probe the mechanical properties with a high spatial and
force resolution, nor the capability of the MEA to record small
extracellular potentials. We therefore quantified the increase
in the electrical and mechanical noise in the new platform with
respect to the standard configurations of both AFM and MEA.
By using the same cantilever, the mechanical noise of the
cantilever in contact with a silicon surface increased from
0.39 nm (RMS) when the AFM was used stand-alone, to
5.6 nm (RMS) when coupled with the MEA and the inverted
optical microscope, and while in contact with a microelec-
trode of the array where the electric potential was being
recorded. On the other hand, the average background
electrical noise from microelectrode recordings increased
from 7 μV (RMS) in stand-alone configuration to 26 μV
(RMS) while scanning the electrode surface with the AFM tip.
Since the measured extracellular potentials from high density
cultures of cardiac myocytes usually ranged from 50 μV to
1,000 μV or more, the signal to noise ratio was enough for
unambiguous detection of AP peaks (see, as an example, the
electrical signals in Figs. 5(b) and 6).
During the experiments with populations of cardiac
myocytes grown onto MEAs, we observed a regular
spontaneous electrical activity as expected for a confluent
monolayer of cells (Clay and DeHaan 1979); fluctuations in
Fig. 5 Representative indentation curves obtained from three consec-
utive indentation events during the same contraction-relaxation cycle.
(a) applied load as a function of time. (b) extracellular potential
recorded from the nearest electrode as a function of time. (c) plots of
the resulting loading versus indentation curves. Each curve (symbols)
is tagged with the delay with respect to the last spontaneous
extracellular AP peak and is fitted with a spline (continuous line)
which was used to calculate the contact stiffness (i.e. slope of the
curve) at different indentation depths
Biomed Microdevices (2011) 13:613–621 617

6. the AP periodicity were less than 2% over 60 s. As already
observed by Domke and co-workers (Domke et al. 1999)
when the AFM tip is brought and maintained into contact
with the membrane of one cardiomyocyte at a constant
applied force, it is possible to follow “morphological”
changes of the cell during the contraction-relaxation cycle
by monitoring the AFM Z-position signal, as shown in
Fig. 6(a–b). As discussed later this “morphology” signal
can be actually affected by changes in cell stiffness and
lateral movements of the underlying cell structures. Graphs
in Fig. 6 clearly show that the measured Z-position signal
(Fig. 6(a) and (b)) are triggered by the electrophysiological
Fig. 6 (a and b) simultaneous AFM Z position measured on two
different cardiomyocytes at 3 DIV, and MEA extracellular potential
signals (respectively c and d). The Z position signals (a and b) were
recorded with the tip in contact and applying a constant force of 0.5 nN
over the cell membranes
Fig. 7 (a) Graphical representation of changes of cardiomyocyte
height and elasticity on a line over the cell (yellow points in image
(b)) during the beating cycle. The colors of the facets represent the
elastic modulus calculated at a constant indentation depth of 195 nm.
Represented heights and elasticities are values averaged over multiple
cycles. (b) bright field image showing the positions (yellow dots)
where indentations were performed. The white arrow indicates the
“position along the cell” value in graph (a) the to 40 μm. The black
lines delineating the cell shape and nucleus are illustrative and does
not necessary correspond with real borders. The AFM cantilever and
the recording round microelectrode are visible in the upper part of the
image
618 Biomed Microdevices (2011) 13:613–621

7. activity (Fig. 6(c) and (d)) which can then therefore be used
to precisely relate the morphological and mechanical
measurements with the beating cycle timing. Usually, after
each AP peak, a positive contraction pulse is followed by a
resting period; however, on some cells, contraction wave-
forms can show multiple peaks as in Fig. 6(b).
In order to map changes in cell height and elasticity
within a single cardiac myocyte, we used the AFM-MEA
setup to perform repetitive indentations with different
delays with respect to the extracellular AP peak (see
Fig. 5(a)). By using the tip-cell contact points calculated
from each force curve, we could obtain a better estimation
of the changes in cell height than the one obtained from the
Z position while in contact with the cell membrane (Jiao
and Schaffer 2004). From the analysis of the indentation
curves we could also estimate the spatiotemporal variations
in cell elasticity during the contraction-relaxation cycle.
As an example, height changes and elasticity values of a
representative cardiomyocyte as a function of the beating
cycle timing and of the position along a defined profile are
shown in Fig. 7. For each point along the profile, 150
indentation curves were taken in 60 s. The beating rate for
this particular cell was 2.56±0.03 Hz. In order to visualize
the uncertainty related to the reported average value, graphs
in Fig. 8 report elasticity and height average values and
standard deviations of the 150 curves taken at a single
location over the cell nucleus (the one corresponding to the
white arrow in Fig. 8(b)). The graph in Fig. 8(b) compares
the height changes calculated from the point of contact of
each force curve with the continuous Z-piezo movement
when keeping the tip in contact with an applied force of
0.5 nN. The difference in the plotted “morphology” profiles
can be ascribed to the fact that changes in cell stiffness and
lateral movements of the underlying cell structures can
affect the force probed by the AFM tip and thus induce
changes in the Z-piezo movement profile.
During analysis we were concerned about the possibility
that the elasticity measurement could be affected by a
penetration of the tip through the cell membrane at high
applied loads. We therefore verified that no sudden jumps
or discontinuities in the contact region of the loading curves
were present. This confirmed that our measurements, taken
at Z-velocities higher than 100 μm/s, did not disrupt the
cell membrane even at the maximum applied loads of
15 nN, as one would expect from the values of force vs
indentation velocity needed to generate cell lyses reported
by Hategan and co-workers (Hategan et al. 2003).
We also performed experiments applying forces up to
200 nN and observed induced electrical responses with
forces greater than 40 nN (data not shown), which could
result from the activation of mechano-sensitive ion chan-
nels or from the creation of (reversible) micro-pores in the
membrane.
From the graphs of Fig. 8 it can be observed that
changes in height are closely related with changes in
transverse elasticity which can be ascribed to the activa-
tion of the actin-myosin crossbridges (Azeloglu and Costa
2010). Usually, few milliseconds after the onset of the AP,
the rise in the free intracellular Ca2+
concentration
switches the contractile machinery and then, as the action
potential ends, the Ca2+
ions are transported out of the
cytosol, allowing relaxation to occur. One could expect
that this results in a single contraction-then-relaxation
sequence as exemplified in Fig. 6(a). However, in some
preparations we observed contraction waveforms that
differ from this behavior by not-presenting single well
defined contraction pulses as illustrated in Figs. 6(b) and
8. Such “double-contraction” pattern could be an indicator
of an altered or not synchronous contraction sequence due
to an inhomogeneous sarcomere shortening (Sarai et al.
2002), and could be similar to the abnormal contraction-
Fig. 8 Height changes and elasticity values of a cardiomyocyte as a
function of the delay from the AP during the beating cycle.
Measurements were taken in a single point over the cell’s nucleus
(corresponding to the white arrow in Fig. 7b). (a) dynamic transverse
elastic modulus calculated at an indentation depth of 195 nm. (b)
changes in cell height calculated from the contact point of each force
curve (connected points, average values ± standard deviations) and
measured from the Z-displacement with the tip continuously in contact
applying a constant force (continuous line)
Biomed Microdevices (2011) 13:613–621 619

8. relaxation patterns observed in pathological conditions
(Maltsev et al. 1998) and chemically altered states
(Gorelik et al. 2002).
In order to analyze further the changes in elasticity
during the observed “double-contraction” pattern, we
evaluated elasticity values at different indentation depths
and constructed a color map representation of the Z-axis
(depth) dependency of cell elasticity at a single location that
is presented in Fig. 9(a). A non homogeneous elasticity
distribution along the Z axis can be observed during the
contraction peak, while, during the “resting” period at the
end of the cycle, the cell is less stiff and more homoge-
neous. The fact that stiffer regions appear and disappear and
do not seem to move up or down during the cycle, supports
the idea that they are produced by the activation-
inactivation of contracting structures (myofibrils) in the
interior of the cell.
The presented results show the ability to precisely
identify the onset of the beating cycle using the extracel-
lular potential signal and to combine AFM mechanical
measurements from multiple cardiomyocyte contractions in
order to analyze the time-dependant variation of the cell
morphology and elasticity. A previous study has already
shown that the elasticity of the cell varies continuously
through the beating cycle (Azeloglu and Costa 2010), but
our approach is the first capable to resolve the time
evolution of the contraction-relaxation cycle with enough
temporal resolution to identify the state mechanical state of
the cell at a specific delay after the onset of the AP without
interfering with cells motility or producing undesirable side
effects.
4 Conclusions
We have presented a novel experimental platform based on
a combined AFM and MEA set-up for performing
simultaneous mechanical and electrical measurements of
cell networks dynamics. The set-up is based on a
commercial AFM instrument and adaptations and modifi-
cations of commercial MEA system components. We used
this platform for measure the changes in morphology and
elasticity of actively beating cardiac myocytes from rat
embryos. The combination of the AFM with the MEAs
does not compromise the ability neither of the AFM to
image and probe the mechanical properties of cells with a
high spatial and force resolution, nor of the MEA to record
small extracellular potentials. Going beyond of just putting
together separate mechanical and electrical measurements
this combined approach allowed us to use the extracellular
potential signal to precisely identify the onset of the beating
cycle allows and combine AFM mechanical measurements
from multiple cardiomyocyte contractions for analyzing the
time-dependant variation of the cell morphology and
elasticity. By estimating the elastic modulus at different
indentation depths from a single location on the cell we
observed a dynamic mechanical behavior that can be
related with the underlying myofibrillar structure dynamics.
Such new capabilities, together with the possibility to
include inmunofluorescence analysis and live calcium
imaging, could allow to study unclear aspects of coupled
mechano-electrical phenomena like cardiac excitation-
contraction coupling (Bers 2002) and cardiac mechano-
electric feedback (Ravens 2003).
Fig. 9 (a) Color representation of the elasticity calculated at a single
point onto a cardiac myocyte as a function of indentation depth and
time during the beating cycle. (b) Depth dependency of the elastic
modulus at 0.09 s (vertical dotted line in (a)) and 0.39 s after the onset
of the extracellular AP
620 Biomed Microdevices (2011) 13:613–621

9. Acknowledgements The authors would like to thank Fillipo Sante
for the support in the construction of the mechanical framework, and
the students Salvatore Saporito, Valentina Romani and Greta Badino
who participated at some points of the developments and experiments.
One of the authors (JFS) was supported through the Programme
Latin America, scholarship No. E07D401181CO).
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