Probing the viscoelastic behavior of cultured airway smooth muscle cells with atomic force microscopy: stiffening induced by contractile agonist.
ABSTRACT Complex rheology of airway smooth muscle cells and its dynamic response during contractile stimulation involves many molecular processes, foremost of which are actomyosin cross-bridge cycling and actin polymerization. With an atomic force microscope, we tracked the spatial and temporal variations of the viscoelastic properties of cultured airway smooth muscle cells. Elasticity mapping identified stiff structural elements of the cytoskeletal network. Using a precisely positioned microscale probe, picoNewton forces and nanometer level indentation modulations were applied to cell surfaces at frequencies ranging from 0.5 to 100 Hz. The resulting elastic storage modulus (G') and dissipative modulus (G'') increased dramatically, with hysteresivity (eta = G''/G') showing a definitive decrease after stimulation with the contractile agonist 5-hydroxytryptamine. Frequency-dependent assays showed weak power-law structural damping behavior and universal scaling in support of the soft-glassy material description of cellular biophysics. Additionally, a high-frequency component of the loss modulus (attributed to cellular Newtonian viscosity) increased fourfold during the contractile process. The complex shear modulus showed a strong sensitivity to the degree of actin polymerization. Inhibitors of myosin light chain kinase activity had little effect on the stiffening response to contractile stimulation. Thus, our measurements appear to be particularly well suited for characterization of dynamic actin rheology during airway smooth muscle contraction.
- SourceAvailable from: Andreas Janshoff[Show abstract] [Hide abstract]
ABSTRACT: Mechanical phenotyping of cells by atomic force microscopy (AFM) was proposed as a novel tool in cancer cell research as cancer cells undergo massive structural changes, comprising remodelling of the cytoskeleton and changes of their adhesive properties. In this work, we focused on the mechanical properties of human breast cell lines with different metastatic potential by AFM-based microrheology experiments. Using this technique, we are not only able to quantify the mechanical properties of living cells in the context of malignancy, but we also obtain a descriptor, namely the loss tangent, which provides model-independent information about the metastatic potential of the cell line. Including also other cell lines from different organs shows that the loss tangent (G″/G') increases generally with the metastatic potential from MCF-10A representing benign cells to highly malignant MDA-MB-231 cells.Open Biology 05/2014; 4(5). · 4.56 Impact Factor
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ABSTRACT: Plant and animals have evolved different strategies for their development. Whether this is linked to major differ-ences in their cell mechanics remains unclear, mainly because measurements on plant and animal cells relied on independent experiments and setups, thus hindering any direct comparison. In this study we used the same micro-rheometer to compare animal and plant single cell rheology. We found that wall-less plant cells exhibit the same weak power law rheology as animal cells, with comparable values of elastic and loss moduli. Remarkably, microtubules primarily contributed to the rheological behavior of wall-less plant cells whereas rheology of animal cells was mainly dependent on the actin network. Thus, plant and animal cells evolved different molecular strategies to reach a comparable cytoplasmic mechanical core, suggesting that evolutionary convergence could include the internal biophysical properties of cells.Biophysical Journal 11/2014; 107. · 3.83 Impact Factor
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ABSTRACT: We propose a magnetic force modulation method to measure the stiffness and viscosity of living cells using a modified AFM apparatus. An oscillating magnetic field makes a magnetic cantilever oscillate in contact with the sample, producing a small AC indentation. By comparing the amplitude of the free cantilever motion (A0) with the motion of the cantilever in contact with the sample (A1), we determine the sample stiffness and viscosity. To test the method, the frequency-dependent stiffness of 3T3 fibroblasts was determined as a power law ks(f) = α + β(f/f[combining macron])(γ) (α = 7.6 × 10(-4) N m(-1), β = 1.0 × 10(-4) N m(-1), f[combining macron] = 1 Hz, γ = 0.6), where the coefficient γ = 0.6 is in good agreement with rheological data of actin solutions with concentrations similar to those in cells. The method also allows estimation of the internal friction of the cells. In particular we found an average damping coefficient of 75.1 μN s m(-1) for indentation depths ranging between 1.0 μm and 2.0 μm.Soft Matter 03/2014; 10(13):2141-9. · 4.15 Impact Factor
Probing the Viscoelastic Behavior of Cultured Airway Smooth Muscle
Cells with Atomic Force Microscopy: Stiffening Induced by
Benjamin A. Smith,* Barbara Tolloczko,yJames G. Martin,yand Peter Gru ¨tter*
*Department of Physics, Nanoscience & Scanning Probe Microscopy Group, andyDepartment of Medicine, Meakins Christie Laboratories,
McGill University, Montreal, Quebec, Canada
microscope, we tracked the spatial and temporal variations of the viscoelastic properties of cultured airway smooth muscle cells.
Elasticity mapping identified stiff structural elements of the cytoskeletal network. Using a precisely positioned microscale probe,
picoNewton forces and nanometer level indentation modulations were applied to cell surfaces at frequencies ranging from 0.5 to
showed weak power-law structural damping behavior and universal scaling in support of the soft-glassy material description of
cellular biophysics. Additionally, a high-frequency component of the loss modulus (attributed to cellular Newtonian viscosity)
increased fourfold during the contractile process. The complex shear modulus showed a strong sensitivity to the degree of actin
polymerization. Inhibitors of myosin light chain kinase activity had little effect on the stiffening response to contractile stimulation.
Thus,ourmeasurementsappeartobeparticularly wellsuitedfor characterizationofdynamicactinrheologyduringairwaysmooth
Complex rheology of airway smooth muscle cells and its dynamic response during contractile stimulation involves
Airway smooth muscle (ASM) cells have unique structural
and mechanical properties that set them apart from skeletal
muscle and nonmuscle cells (Murphy, 1994; Small, 1995).
The complex mechanics and contraction of ASM tissues,
being both length and history dependent, are largely
responsible for airway narrowing and airway hyperrespon-
siveness in asthmatic subjects (Brusasco and Pellegrino,
2003; Fredberg, 2004; Ma et al., 2002; Woolcock et al.,
1984). To explain these pathological phenomena, inves-
tigators have studied ASM microstructure (Daniel, 1988;
Kuo et al., 2003), cellular complex microrheology (An et al.,
2002; Fabry et al., 2001a, 2001b; Hubmayr et al., 1996), and
the biochemical pathways activated during contractile events
(Gerthoffer and Gunst, 2001; Morgan and Gangopadhyay,
2001; Pfitzer, 2001; Schramm and Grunstein, 1992; Somlyo
and Somlyo, 2003). The prospect of understanding the link
among these observations has been a difficult challenge.
An emerging description of ASM rheology is that of the
soft-glassy material hypothesis, which does not rely on
specific molecular mechanisms but embodies the concepts of
rigidity as well as the ability of cells to flow and remodel
(Fabry et al., 2001a; Fabry and Fredberg, 2003; Sollich,
ratio of the dissipative to the elastic components of the
complex shear modulus) and the dependence of the complex
intracellular environment. Structural disorder in the cell
explainstheabsence ofanycharacteristic timeconstantofthe
complex rheology or specific mode of deformation. Meta-
stability is understood as weak confinement of structural
elements in the cell, in that the energy of stochastic forces is
comparable to confinement potentials, and thus structural
rearrangements are probabilistic events. In this way the
rigidity of the cell acquires a weak (power-law) dependence
on frequency of perturbation, as stress relaxation can occur
over long time scales and is progressively less probable at
higher frequencies. If the source of random forces is non-
thermal in nature (e.g., ATP hydrolysis by motor proteins or
dynamic actin filaments), it is possible for the cell to rapidly
modulate its mechanical state.
The applicability of this description as an integrative
framework encompassing the diversity of underlying
mechanisms involved in ASM contractile events has recently
been reviewed (Gunst and Fredberg, 2003). In this
framework, the onset of contractile processes is described
by a rapid increase in microscale agitations (interpreted from
a transient increase in hysteresivity), force and stiffness
increase, and then gradual freezing or solidification (de-
creased hysteresivity). The two main elements within the
cell’s mechanical structure that are implicated in ASM
contractile processes are actomyosin cross-bridge cycling
and actin filament polymerization. Increased actomyosin
cross-bridge activity is initiated by receptor stimulated
Submitted June 17, 2004, and accepted for publication January 10, 2005.
Address reprint requests to Benjamin A. Smith, E-mail: bsmith@physics.
? 2005 by the Biophysical Society
2994Biophysical JournalVolume 88 April 2005 2994–3007
elevation of the concentration of intracellular calcium ions,
which promotes myosin light chain phosphorylation via
calmodulin and myosin light chain kinase (MLCK). This
activation results in force production and sliding of adjacent
actin and myosin filaments (Huxley and Hanson, 1954) and
is widely accepted as the key step in smooth muscle con-
traction (Kamm and Stull, 1986; Murphy, 1980). The rapid
cycling of cross-bridges is proposed to explain the transient
increase in hysteresivity, and the subsequent decay is attrib-
uted to the formation of latch-state binding of myosin heads
to actin filaments (Fredberg et al., 1996; Murphy, 1994).
Enhanced actin polymerization and dynamic remodeling is
observed during the onset of contraction (An et al., 2002;
Hirshman and Emala, 1999; Mehta and Gunst, 1999). The
actin cytoskeleton is suggested to provide a fortified envi-
ronment for tension development and to strengthen mem-
brane contactsofthe contractile apparatus, thus playing akey
role in force transmission to the extracellular matrix and
in sensing and adapting to mechanical forces (Gunst et al.,
Measurable quantities, like tension, stiffness, hysteresiv-
ity, and noise temperature, are all integrative parameters,
typically sensitive to both actomyosin activity and actin
polymerization (An et al., 2002; Jones et al., 1999; Mehta
and Gunst, 1999). What is required is a technique to
precisely target individual mechanisms with a mechanical
probe. This may only be possible if there is a spatial seg-
regation of nonmyosin-associated actin and the actomyosin
contractile apparatus in the ASM cell. There is evidence to
suggest that such a distinction between contractile (a-) and
cytoskeletal (b-) actin in ASM is present (Small, 1995)
where cytoskeletal actin may be selectively localized to
membrane-associated structures. However, this segregation
is still a point of dispute (Song et al., 2000; Stromer et al.,
Atomic force microscopy (AFM; Binnig et al., 1986) is
established as a versatile tool for imaging and measuring
nanoscale elastic properties of living cells (A-Hassan et al.,
1998; Radmacher et al., 1996), with sensitivity to underlying
cytoskeletal elements such as actin filaments (Rotsch and
Radmacher, 2000). Recent advances have measured com-
plex (viscoelastic) rheology (Alcaraz et al., 2003; Mahaffy
et al., 2000). In this study, we employ AFM to map the
mechanical structure and explore the frequency-dependent
complex rheology of ASM cells in culture. We observe
stiffening events induced by a known contractile agonist and
test the sensitivity of these measurements to MLCK inhi-
bitors and actin polymerization dynamics. We also compare
our work with recent studies of ASM rheology and
contractile stiffening in tissue strips (Fredberg et al., 1996)
and in cultured cells by using magnetic twisting cytometry
(MTC; An et al., 2002; Fabry et al., 2001a). AFM allows for
versatile control over the spatial localization of the measure-
ment probe and the mechanical stimulus with the goal of
separating characteristic effects of actomyosin interactions
from cytoskeletal rearrangements.
MATERIALS AND METHODS
Cell culture and treatments
Trachealsmoothmuscle cells from7- to 9-week-oldmale Fisher rats (Harlan
Sprague Dawley, Wallkerville, MD) were isolated and cultured according to
previously described methods (Tolloczko et al., 1995). Briefly, the cells
were enzymatically dissociated with 0.05% elastase type IV and 0.2%
collagenase type IV and cultured in 1:1 Dulbecco’s modified Eagle’s
medium, Ham’s F12 medium supplemented with 10% fetal bovine serum,
0.224% NaHCO3 and 100 U/ml penicillin, 100 mg/ml streptomycin and
25 mg/ml amphotericin in the presence of 5% CO2. Cell culture reagents
were purchased from GIBCO (Mississauga, Ontario, Canada). Confluent
first or second passage cells were rendered quiescent by incubation in
medium containing 0.5% fetal bovine serum for 4 days before experiments.
Confirmation of a smooth muscle phenotype was based on typical
morphology, positive smooth muscle specific a-actin staining, negative
keratin staining, and contractile responses to agonists.
For AFM experiments, cells were mounted on the microscope stage in
;300 ml of Hanks’ balanced salt solution (Invitrogen Canada, Burlington,
Ontario, Canada), pH 7.3, at room temperature. Time-course measurements
were made for 10–12 min before and for 15 min after addition of 10 mM
contractile agonist 5-hydroxytryptamine (5-HT; serotonin). To inhibit
MLCK, cells were preincubated with 1 mM wortmannin or 5 mM
ride (ML-7) for 10 and 25 min, respectively (Nakanishi et al., 1992; Saitoh
et al., 1987). To cause actin depolymerization, actin capping agent
cytochalasin-D was added to the cells after treatment with 5-HT for
15 min and the measurements continued for 10–15 min. Cytochalasin-D,
wortmannin, and ML-7 were obtained from Sigma-Aldrich Canada
(Toronto, Ontario, Canada).
Measurement of cell contraction
Cells used for contraction measurements were plated on homologous cell
substrate (Tao et al., 2003) and rendered quiescent by serum deprivation at
;70% confluence. The cells were incubated for 10 min with 1 mM
wortmannin or for 25 min with 5 mM ML-7 or with the appropriate vehicle
(DMSOor ethylalcohol, respectively). Images of cells stimulated with 5-HT
were acquired at the rate of one image per minute using a video camera
(Hamamatsu Photonics, Hamamatsu City, Japan) mounted on a microscope
equipped with Nomarski optics (Nikon Diaphot, Nikon, Tokyo, Japan) and
PTI (Photon Technology International, Princeton, NJ) software. The length
of the cells was measured before and 10 min after stimulation. Each
experimental group consisted of 31–37 cells. Data are expressed as
percentage of cell length decrease. Negative values indicate increase in
Atomic force microscopy
Measurements were performed with a Bioscope AFM equipped with a
G-type scanner, Nanoscope IIIa control electronics, and software version
4.43r8 (Digital Instruments, Veeco Metrology Group, Santa Barbara, CA).
Silicon nitride triangular microlevers (TM Microscopes, Veeco) with
a nominal spring constant of k ¼ 0.01 N/m were used for both force
volume and complex modulus measurements. For the latter, a 4.5 mm
diameter polystyrene bead (Polysciences, Warrington, PA) was fixed to the
tip of each probe with UV light curable adhesive (Elecro-lite, Danbury, CT)
using a custom procedure. The deflection sensitivity of the optical lever was
calibrated before and after each experiment by measuring the slope of the
Smooth Muscle Cell Mechanics by AFM2995
Biophysical Journal 88(4) 2994–3007
contact region of a force-distance curve acquired on a clean glass cover slip
in Hanks’ balanced salt solution.
Elasticity mapping via force volume
In this AFM imaging mode (supported by Nanoscope control software,
version 4.23 b6 or higher), the cantilever is scanned vertically to obtain
a force-distance curve at each point in an x-y array over the cell surface. The
probe indents the surface with a predetermined maximum force relative to
baseline on each approach (relative trigger mode) then is retracted again
before moving laterally to the next point in the array. The height (z-piezo
scannervoltage) at which the maximum force is reached is used as the height
image value. The inverse slope of the indentation region of each force curve
is measured to first order by taking the force value at a fixed height above
the maximum-force height for that curve. These values are plotted as the
elasticity map, providing a qualitative measure of the variations of the in-
verse local elastic modulus of the cell surface. Parameters used for the force
volume imaging were: 1 mm z-scan, 64 data points per curve, 64 3 64 array,
10 Hz vertical scan rate, 14 Hz lateral scan rate, and ;0.4 nN maximum
force (although ;0.1 nN was attributed to the hydrodynamic force on the
probe). The total time required for each image was 15 min.
Indentation modulation, complex
to indent the surface at a specified location on each cell. Although the lateral
resolution of the force volume imaging is not maintained, the use of beaded
probes provides a better assessment of the cells rheology by averaging over
a larger area of the cell surface with well-defined contact geometry. Also,
destructive deformation (Dimitriadis et al., 2002). The relative trigger mode
was used to control the maximum force applied on each indentation. Force
curves were acquired at a much slower rate (0.317 Hz, 1.5 mm z-scan, 4096
data points). At this scan speed, the hydrodynamic force on the lever was
insignificant compared to the maximum loads used (;0.3 nN). We did not
rely on force curve analysis for quantitative evaluation of complex cellular
rheology but instead used the indentation modulation procedure described
below. The only parameter obtained from force curves (example in Fig. 1 A)
each approach (i.e., at the point of maximum indentation force) and each
retraction from the cell surface. During the indentation delay, a sinusoidal
oscillation was added to the vertical scan control voltage, with small
lock-in amplifier (SR 830, Stanford Research Systems, Sunnyvale, CA) was
used for drive signal generation and measurement of the resulting amplitude
and phase of the deflection signal. These outputs were sampled with a 12-bit
data acquisition board (PCI-6024E, National Instruments, Dallas, TX) and
recorded on a computer for further analysis.
Time-course profiles were constructed from repeated indentations with
modulation at constant frequency (typically 10 Hz). Frequency-dependent
measurements were acquired during periods of stable mechanical response
of the cell. On alternate indentations, the modulation frequency was toggled
between the time-course (control) frequency and other frequencies used (in
random order), such that the control frequency was used before and after
each measurement at another frequency. In this way small but persistent
shifts in the response to the control frequency were used to scale mea-
surements at other frequencies relative to the average response at control
frequency modulations. These changes were typically ,10%, but in those
cases where larger changes occurred, the corrected values were consistent
with cases in which the changes were small.
The hydrodynamic drag on the lever was calibrated for each probe used
(Alcaraz et al., 2002). These measurements were made by modulating the
probe (at 50 Hz) at different heights (0.2–2 mm) above the cell surface at the
same location and with the same amplitude (5–8 nm) as used for the
indentations experiments. The phase response of the AFM z-piezo was
calibrated by bringing a stiff probe (k ¼ ;42 N/m, Olympus etched Si
tapping-mode probe) into contact with a clean glass cover slip (in air),
modulating the z-piezo at all frequencies used in this study, and recording
the phase of the lever deflections relative to the drive signal. The frequency-
dependent phase shift was subtracted off of all other phase measurements.
The Hertzian contact mechanics model (Hertz, 1882; Johnson, 1987) for
a spherical indenter relates the force experienced by the probe (F) to the
indentation depth (d) for elastic deformations of linear, homogeneous
samples of infinite thickness. It is commonly used to extract values of elastic
modulus from AFM indentation data. We used the first term of the Taylor
expansion of this model to compute the complex shear modulus, G*, of the
cell from the small amplitude indentation oscillations, d1, (Alcaraz et al.,
2003; Mahaffy et al., 2000)
where n is the Poisson ratio of the cell, R is the radius of the spherical
probe, d0is the operating indentation, and E0the zero frequency value of the
elastic modulus obtained from the slow approach force curve. E1is the
3ð1 ? n2Þ
a beaded AFM probe (at a rate of 0.317 Hz) such that it approaches, indents,
retracts, and releases from a cell’s surface. The inset is an illustration of
a typical signal applied to the vertical scanner to perform indentation
AFM probe above a cell surface. The fit to the scaled spherical drag function
(see text) and its extrapolation to surface indentation (q) are shown. The
inset shows the linear dependence of the hydrodynamic drag on modulation
frequency, and the purely viscous nature of this drag (H9 is the in-phase
elastic component which is negligible compared to the out-of-phase viscous
(A) Typical force curve obtained from vertical scanning of
2996Smith et al.
Biophysical Journal 88(4) 2994–3007
frequency-dependent elastic modulus related to G* by E1¼ 2(1 1 n)G* for
a continuous medium (Landau and Lifshitz, 1986). The indentation depth is
calculated as d ¼ z ? d, were z is the controlled vertical position of the base
of the probe (measured as positive values extending down from the point of
contact between the bead and the cell surface) and d (¼ F/k) is the measured
deflection of the lever. Note that d1and F1¼ 2(Rd0)1/2E1d1/(1 ? n2) are both
complex values having both amplitudes and phases relative to the vertical
drive oscillation, thus E1and G* have both real (in-phase) and imaginary
(90? out-of-phase) components. The Poisson ratio is taken to be 0.5 for
incompressible materials (Mahaffy et al., 2004).
Further extension of the Hertz model to include the effects of finite
sample thickness and adherence to the substrate required a detailed
knowledge of the cells thickness (Dimitriadis et al., 2002; Mahaffy et al.,
2004). AFM topography of nonconfluent cells and optical focal plane
measurements of confluent cells estimate cell thickness at h ¼ ;3 mm. For
this thickness and the indentation depths used (;100 nm), rheology
coefficients could be overestimated by as much as 20% ([Rd0]1/2/h).
Correction for the hydrodynamic drag forces experienced by the probe
moving through the surrounding liquid was done by calculating the
hydrodynamic drag function iH$ ¼ F1/d1¼ ib(h)f (b is the drag factor, i is
the imaginary unit, i2¼ ?1, and f is the oscillation frequency) for different
heights (h) above the cell and extrapolating to h ¼ 0 (Alcaraz et al., 2002).
As a model for the drag factor dependence on h, we used the scaled spherical
function with an added constant to account for the nonzero drag on probe
oscillations far from the surface: b(h) ¼ 6phaeff2/(h 1 heff) 1 b0, where aeff
and heffare the effective radius and height of the lever, respectively, h is the
liquid viscosity, and b0is the drag far from the cell surface. For these
measurements, without contact or adhesion to the cell surface, the real
component of the drag was negligible and the imaginary component scaled
linearly with frequency, as expected for purely viscous drag. An example of
a fit of the imaginary component of the drag to the functional height
dependence of a scaled spherical oscillating probe with nonzero decay at
long distance is shown in Fig. 1 B (modulation frequency 50 Hz). This fit
was used to determine a value for the drag factor at zero height, b(0). The
hydrodynamic drag on the lever was subtracted from complex shear
modulus measurements (at any frequency).
Finally, we calculate the complex shear modulus of the cell as
G?¼ G91iG$ ¼
1 ? n
Here the complex shear modulus has been separated into its real (storage
modulus, G9) and imaginary (loss modulus, G$) components and has
only a weak dependence on the operating indentation and probe radius
(G*;(Rd0)?1/2). For time-course plots of G9 and G$ we define a parameter
k0¼ k(1 ? n)/4(Rd0)1/2, and calculate only the ratios G9/k0and G$/k0so that
any uncertainty associated with k (probe spring constant), n, R, and d0are
not included. This is justified because only d0changes throughout the course
of each measurement and these variations have little impact on variations of
G*. We also calculate the loss tangent or hysteresivity (h ¼ G$/G9), which
provides a measure of solidlike (h ? 1) or fluidlike (h ? 1) behavior of the
cell, and where dependence on k, n, R, and d0is eliminated (and thus the
common errors associated with determination of the contact point, control of
the maximum force, and uncertainty associated with Hertzian contact
mechanics). Values of G9, G$, and h reported in the text (and in Table 1) are
averages over 5 min of individual indentation measurements (including
calculated values of k0), then averaged among all cells tested, reported as
mean 6 SE. Percent change of responses (R) relative to baseline (B) values
are calculated as (R ? B)/B 3 100% for each cell then averaged within the
population, where values in the stimulated or treated states are 5 min
averages taken between 5 and 10 min after addition of the drug. Statistical
significance of differences between reported means was determined by
a two-sample t-test at a level of p , 0.05.
Frequency-dependent cellular mechanics model
To describe the functional dependence of G9 and G$ on oscillation
frequency, we used the power-law structural damping model (Fabry et al.,
2001a; Fredberg and Stamenovic, 1989; Hildebrandt, 1969):
where the structural damping coefficient ? h h is not an independent parameter
butis relatedto the power-law exponenta by ? h h¼tan(pa/2), G0isa modulus
scalefactor,f0isa frequency scalefactor,Gdenotesthe gammafunction,and
2) is close to unity for small values of a and is ignored in interpretations and
discussions, although included in data fits. This model implies that the loss
modulusconsistsofacomponent(withscalefactor ? h hG0)thatiscoupledtothe
interpreted as frictional damping. Additionally, the loss modulus contains
this term, the structural damping constant (? h h) is not equivalent to the
Measurements of G9 and G$ versus f were fit to this model with a global
Corp., Northampton, MA), determining values of a and m for the cells in
unstimulated, stimulated, and drug treated states. In fitting individual data
uniquely. However, multiple data sets (cells under multiple treatments) were
G0that are common to all cellular states (similar to the analysis of Fabry
et al., 2001a).
G?¼ G0ð11i? h hÞ
Gð1 ? aÞcosðpa=2Þ1imf;
Force volume imaging was performed on a confluent culture
of ASM cells for two different lateral scan sizes (40 mm and
TABLE 1Population average complex rheology of ASM cells under various treatments
5.1 6 0.4
21 6 2
7.2 6 0.6
5.1 6 0.4
4.1 6 0.5
4.4 6 0.6
Baseline (n ¼ 16)
5-HT (n ¼ 11)
Cytochalasin-D (n ¼ 5)
Wortmannin (n ¼ 5)
Wortmannin 1 5-HT (n ¼ 5)
ML-7 (n ¼ 7)
ML-7 1 5-HT (n ¼ 7)
*Measured at 10 Hz modulation.
yParameters from fits of frequency-dependent rheology.
1.2 6 0.2
2.7 6 0.5
0.5 6 0.1
1.3 6 0.3
2.4 6 0.2
0.6 6 0.2
1.1 6 0.3
0.32 6 0.05
0.52 6 0.09
0.25 6 0.05
0.35 6 0.09
0.34 6 0.07
0.20 6 0.04
0.26 6 0.06
0.27 6 0.03
0.20 6 0.02
0.53 6 0.05
0.25 6 0.06
0.14 6 0.02
0.40 6 0.06
0.26 6 0.04
0.120 6 0.005
0.055 6 0.005
0.160 6 0.009
0.069 6 0.002
0.158 6 0.009
0.101 6 0.004
Smooth Muscle Cell Mechanics by AFM 2997
Biophysical Journal 88(4) 2994–3007
5 mm; Fig. 2). The height under constant force profile for the
larger scan (Fig. 2 A) clearly shows one cell with three other
neighboring cells contacting each other at the cell bound-
aries. The elasticity profile shows that the thick central region
(likely the nucleus) is softer than the surrounding regions of
the cell (roughly a fourfold difference in force-indentation
slope) and a nodal fiber network that spans the entire area of
the cell surface. This microstructure is even more striking in
the smaller scan of the perinuclear region (Fig. 2, C and D).
The fibrils are not aligned with any particular direction in
the cell, but form a network of interconnected nodes. The
to the surrounding cellular material (.2-fold difference in
compliance). This rich elasticity contrast on the submicron
length scale is part of the motivation for using a large
(4.5 mm) bead on the AFM cantilever as an indentation probe
for single-location measurements, providing some local av-
eraging of the cell’s shear modulus.
Although the force volume imaging provided a qualitative
view of the spatial variations of the cells’ mechanical
structure, it was not useful for quantitative assessment of
elasticity, lacking both in accuracy and temporal resolution.
The major limitation was the presence of viscous forces on
the AFM probe, as well as adhesion to the cell surface. The
hydrodynamic drag force on the probe and the viscoelastic
nature of the cell caused a significant hysteresis between the
approach and retraction in the indentation region of the force
curves (Fig. 1 A). This made measurement of elastic modulus
via slope analysis, or even a fit to Hertzian indentation
model, highly convoluted with viscosity effects. Averaging
approach and retraction curves in an attempt to eliminate
viscous forces from the analysis (Radmacher et al., 1996)
was complicated by surface adhesion artifacts. Many kinetic
processes involved in ASM contraction occur within minutes
(Gunst and Fredberg, 2003), but sustained contractions can
last for 15 min or longer and therefore it was possible to
obtain force volume maps (15 min per image) before and
during contractile activation with 10 mM 5-HT. Some
structural rearrangements were observed near the cell
periphery as well as subtle changes in structure and apparent
elasticity contrast throughout the cell; however, these effects
were difficult to quantify and distinguish from random vari-
ations (data not shown).
Complex modulus time-course response
The AFM probe was positioned onto an ASM cell surface in
the region between the nucleus and the periphery using the
Bioscope optics (403 objective). This was the region where
the highest detail in the cytoskeletal network was observed in
the force volume images. Indentation modulation (at 10 Hz)
was used to compute the storage modulus (G9), loss modulus
(G$), and loss tangent (h ¼ G$/ G9). The elastic storage
modulus values are typically in the range 0.5–3 kPa (G9 ¼
1.25 6 0.20 kPa, n ¼ 16, 5 min averages), with dissipative
loss moduli one fifth to one third of these values (h ¼ 0.27 6
0.03) although in some cases as large as 0.5G9. Addition of
the contractile agonist 5-HT (10 mM) causes a dramatic
increase in G9 (150 6 30% increase relative to baseline
values, n ¼ 11), which is often paralleled by a similar
increase in G$ (67 6 15%). Two examples are shown in Fig.
3. The loss tangent time courses tend to be much less noisy
than the time courses for G9 and G$, indicative of coupling
between elastic and dissipative processes within the cell (the
structural damping hypothesis). The time-course plots of h
show a definitive decrease after addition of 5-HT (?28 6
5%), interpreted as a shift toward solidlike behavior.
These changes in G9, G$, and h all occur within the first
few minutes after addition of 5-HT, sometimes within the
first 30 s (one or two data points). However, the amplitude
and kinetics of the contractile stiffening varies among cells.
Typically cells with larger G9 and smaller h baseline values
responded less than cells with smaller G9 and larger h
baseline values, as if there is an upper threshold stiffness (or
lower threshold for the loss tangent) beyond which the cell
cannot stiffen any further. The threshold for G9 seemed to be
a confluent culture of rat tracheal smooth muscle cells using a constant
maximum applied force of ;0.4 nN (height color scale spans 1 mm in A and
0.3 mm in C; dark is stiff and light is soft in the elasticity maps). A and B
covers an entire apical surface of a cell and portions of three neighboring
cells. C and D are of the area within the square marked in A. These images
reveal a stiff fibrous network that is highly nodal and spans the entire cell,
although the nuclear region appears to provide a very soft background (large
light region in the top left quarter of B). The higher resolution images in C
and D clearly identify the nodes as well as their interconnecting fibers as the
structures of mechanical rigidity within the cell.
Force volume height (A, C) and elasticity (B, D) profiles of
2998Smith et al.
Biophysical Journal 88(4) 2994–3007
;2.5 kPa and ;0.15 for h, although this has not been
rigorously quantified. Cells with baseline G9 values
significantly above this stiffness threshold were eliminated
from the analysis. Another significant heterogeneity in the
response profiles was that for some cells the stiffness
increased to a peak value then decayed slowly (over ;5 min)
toward baseline levels, but for other cells (the majority) the
stiffness increases were unimodal, reaching a plateau that did
not decay within the measurement time span.
When two modulation amplitudes, 2.5 and 6.2 nm, were
used on alternating indentations (Fig. 3, C and D), the results
for G9, G$, and h at these two amplitudes are indistinguish-
able in both the baseline and stimulated states. This suggests
that the modulations used fall within a linear deformation
regime, in support of the use of only the first Taylor
expansion term of the Hertz model (Eq. 1), so that results do
not depend on the precise choice of modulation amplitude.
Baseline values of G9 and G$ often exhibit an increase
from the onset of indentation probing (Fig. 3, especially Fig.
3 C). This stiffening occurs mainly in the first minute of
probing, decaying to a gradual increase (barely detectable) or
flat-line response after that. This implies evidence for
a mechanosensitive response of the ASM cells to the
mechanical perturbation from the AFM probing. For this
reason we probed each cell continuously for 8–10 min before
adding 5-HT, to establish a relatively stable baseline level of
the storage and loss moduli.
Effects of cytochalasin-D and MLCK inhibitors
The dependence of the ASM complex modulus on the
polymerized actin content in the contractile state was tested
with the actin capping agent, cytochalasin-D (30 mM) after
stimulation with 5-HT. A dramatic decrease in both the
storage and loss moduli was observed within 3 min after
addition of cytochalasin-D, indicative of cellular softening or
relaxation (Fig. 4). The hysteresivity exhibits a large increase
from ;0.2 to 0.53 over a 5-min period after the onset of the
softening response. This is interpreted as a shift toward
fluidlike behavior. The average values of G9, G$, and h from
all cells tested for baseline, 5-HT stimulated, and cytocha-
lasin-D treated states are presented in Table 1. With
cytochalasin-D present, storage modulus values drop to
a level not only well below the contractile level, but also
significantly below the baseline level (n ¼ 5). The contractile
stiffening and solid- versus liquidlike behavior of ASM cells
is clearly sensitive to the degree of actin polymerization.
Inhibitors of MLCK (wortmannin) and ML-7 were used
to test the sensitivity of ASM baseline rheology and response
to contractile stimulation to the activity of actomyosin cross-
bridge cycling. Results are listed in Table 1. Wortmannin
treatments had no effect on baseline rheology as the storage
and loss moduli average values after treatment (n ¼ 5) were
not significantly different than control baseline values (n ¼
16). Furthermore, 5-HT resulted in a stiffening response that
(A and C) and hysteresivity (B and D)
time-course measurements on two dif-
ferent cells, stimulated with the con-
tractile agonist 5-HT (serotonin). The
solid symbols in A and C are the elastic
storage moduli and the open symbols
are the dissipative loss moduli obtained
from repetitive indentation modulation
coefficient, k0¼ k(1 ? n)/4(Rd0)1/2).
After an initial increase, likely a mecha-
modulus reaches a relatively stable
baseline before stimulation. 5-HT in-
duces a dramatic stiffening response
and reduced hysteresivity with variable
kinetics. Two different modulation
amplitudes were used in C and D (6.2
nm: squares for baseline and circles
after 5-HT; 2.5 nm: triangles pointing
up for baseline and pointing down for
which indicates that cell rheology was
probed in a linear deformation regime.
Complex shear modulus
Smooth Muscle Cell Mechanics by AFM2999
Biophysical Journal 88(4) 2994–3007
was not significantly different in amplitude or kinetics for
wortmannin treated cells than for control cells. Loss tangent
values showed a large decrease after 5-HT stimulation in
wortmannin treated cells similar to those for 5-HT stimulated
control cells. Therefore, a 5-HT-induced contractile stiffen-
ing and transition toward solidlike behavior was still present
in wortmannin treated cells.
Cells treated with ML-7 had significantly smaller storage
to untreated cells. Due to the length of time needed for ML-7
treatment, we were not able to track the time course of this
decrease in G9. After stimulation of these cells with 5-HT, G9
values increased, although not to the same level as was
reached in control cells after 5-HT. Loss tangent values were
larger for ML-7 treated cells and showed significant decrease
after 5-HT stimulation. Thus, ML-7 has a softening (liquefy-
ing) effect on ASM cells but does not prevent the stiffening
(solidifying) effect of 5-HT stimulation.
To illustrate this stiffing response, independent of baseline
levels, we calculated the percent response to 5-HT relative
to baseline values for each cell, then averaged the percent
responses for control, wortmannin treated, and ML-7 treated
cells (Fig. 5). Although some slight differences in G9, G$, or
h mean response to 5-HT are observed between control and
treated cells (for example, a decrease in G9 percent response
from 150% in control cells to 104% in MLCK inhibited
cells), these changes were not statistically significant.
Optical measurements of cell contraction confirmed that
the MLCK inhibitory treatments used were sufficient to
block 5-HT-induced cell shortening (Table 2). Contraction
was reduced by the ML-7 vehicle (ethanol control response
is half the DMSO control), but was substantially blocked by
both wortmannin and ML-7.
To investigate further the details of ASM rheology and the
changes in cellular mechanics involved in contractile
activation, we studied the frequency dependence of the
complex shear modulus. Results show a weak power-law
dependence of G9 on deformation frequency (Fig. 6 A,
baseline conditions, n ¼ 10). The behavior of G$ is more
low frequencies (parallel to G9) but having a much stronger
dependence at higher frequencies. This complex behavior is
well fit by the power-law structural damping model with
additional Newtonian viscosity, described above (Eq. 2). The
provides values for the power-law exponent a ¼ 0.120 6
arbitrary choice for this single data-set fit).
lasin-D, on the complex shear modulus and hysteresivity of an ASM cell
after stimulation with 5-HT. The severe decrease in storage (G9) and loss
(G$) modulus, to levels well below baseline, indicate that dynamic actin
polymerization is largely responsible for maintenance of cellular rigidity in
the contractile and resting states. The large increase in hysteresivity (h)
characterizes the inhibition of actin polymerization as a transition toward
fluidlike behavior of the cell.
Example of the effects of the actin capping agent, cytocha-
baseline) calculated for each cell tested and averaged for control and MLCK
inhibited conditions. Although mean values are reduced, there is no sig-
nificant inhibition of the contractile responses (P . 0.05).
Relative responses to stimulation (percent difference from
MLCK inhibitors or appropriate vehicle
5-HT-evoked contraction of ASM cells treated with
% contraction:20.7 6 2.7
?2.4 6 2.6 9.7 6 2.70.85 6 0.07
*Vehicle for wortmannin.
yVehicle for ML-7.
3000Smith et al.
Biophysical Journal 88(4) 2994–3007
This assay was repeated after the stimulation of the ASM
cells with contractile agonist, 5-HT, and inhibitory drug
treatments (Fig. 6 B). The parameters a and m varied among
treatments and are listed in Table 1. The power-law exponent
showed anticovariance with the level of stiffness. Contractile
stimulation, with or without MLCK inhibitors present,
increased G9 at all frequencies tested and decreased the
power-law exponent (slope in Fig. 6 B). Treatments that
decreased stiffness, such as actin depolymerization with
cytochalasin-D or MLCK inhibition with ML-7 (although
not with wortmannin) resulted in stronger power-law
frequency dependence (increased a). This behavior was fit
well using the constraint of a universal coordinate (f0, G0) at
much higher frequency than accessible with current
technique, where all data fits for G9 converge (Fig. 6 C).
The global nonlinear regression (lines in Fig. 6, B and C)
determined log(f0) ¼ 9.0 6 0.7 and log(G0) ¼ 0.97 6 0.05
(or f0¼ 109Hz and G0¼ 9.3 kPa). Values of a were
identical to those found if data sets were fit individually for
each treatment (not shown). The Newtonian viscosity coef-
ficient varied among treatments but without consistent cor-
relation with any other parameter of the model. It increased
by ;4-fold under stimulation with 5-HT and remained
slightly elevated after relaxation with cytochalasin-D. Unlike
the changes in stiffness and a, m showed no significant
variation in the presence of MLCK inhibitors (wortmannin
The force volume images presented here provide maps of
spatial variations of cultured ASM cells’ mechanical
structure under constant applied normal force (;0.4 nN).
With this level of force the cells plasma membrane is
expected to be easily deformed (Evans, 1989) but is rarely
punctured (as observed by inspection of our force curves).
Indentation depths are typically 50–100 nm but can be as
large as 200 nm when probing a very soft region of the cell
(i.e., the nucleus). This allowed us to probe the underlying
(yet still relatively superficial) cytoskeleton of the ASM
cells. The observed cross-linked fiber network, with no
striations or alignment with any particular direction in the
cell, is a hallmark of smooth muscle microstructure (Small,
1995). The stiff nodes we observe appear very similar to the
dense bodies or the membrane-associated dense plaques that
microscopy (Ashton et al., 1975; Draeger et al., 1990; Gunst
and Tang, 2000). Our observations are more likely of dense
plaques due to the superficial nature of the surface inden-
tations used. These structures are critical for maintaining the
structural integrity of the cells as well as being heavily
involved in mechanical signaling and contraction. Dense
plaques are known to be associated with actin filaments and
interconnected by the more stable intermediate filaments,
whereas cytosolic dense bodies connect actomyosin contrac-
tile fibers to the actin filament cytoskeleton (Draeger et al.,
1990). The lateral resolution in AFM measurements on soft
surfaces is limited mainly by the tip-surface convolution
during indentation and the decay of the resulting stress field
within the cell (Charras et al., 2001). A good estimate for the
resolution for our probes and indentation level is 50–80 nm.
The distance between individual measurements in Fig. 2 B
is 78 nm, so we are measuring features at the scale of our
resolution. This is comparable to, if not better than, the reso-
of living cells. Our force-based imaging demonstrates the
ability of AFM to identify specifically mechanical structures
in ASM cells and gives an appropriate length scale for the
variations in cellular mechanics.
AFM force volume imaging provides additional informa-
tion contained in the elasticity contrast. The enhanced
stiffness of the fiber network relative to the surrounding
with an additional Newtonian viscosity component (Eq. 2). The weak power-law behavior of the elastic modulus, spanning the entire frequency range tested,
supports the hypothesis that cells behave as soft-glassy materials close to the glass transition (a ¼ 0). (B) Frequency-dependent complex moduli under all
treatment conditions, with a global fit to Eq. 2. The power-law exponent and Newtonian viscosity varied among treatments (fit parameters giving in Table 1).
(C) Extrapolation of the storage moduli (G9) fits to the universal coordinate (f0¼ 109Hz and G0¼ 9.3 kPa).
(A) Frequency dependence of the complex modulus of untreated ASM cells. Data are well fit by the weak power-law structural damping model
Smooth Muscle Cell Mechanics by AFM3001
Biophysical Journal 88(4) 2994–3007
cellular material supports the concept that it is the stress
bearing structure of the cell. Within the tensegrity model for
cellular biomechanics (Ingber, 1997), the fiber stiffness is
related not only to its intrinsic rigidity but to existing
prestress in the fiber network, recently confirmed for ASM
cells (Stamenovic et al., 2004; Wang et al., 2002). Modeling
of adherent cell poking experiments (Coughlin and Stame-
novic, 2003) show that prestress in the peripheral actin
cytoskeleton provides the key resistance to indentation at
forces comparable to those used here. The soft-glassy
material model involves structural elements that are in close
interaction (confinement) yet still undergo nonequilibrium
remodeling events (Fabry and Fredberg, 2003; Sollich,
1998). In cells, the identity of these structural elements is not
known, but it seems reasonable that (for ASM cells) the stiff
nodes we observe are potential candidates. Force volume
mapping could be used to target individual elements and test
the malleability of the interconnecting fibers, which likely
define the confinement interactions.
We characterized the microscale viscoelasticity of cultured
ASM cells with AFM indentation modulation in the fibrous
perinuclear region. The complex shear modulus, measured in
response to nanoscale oscillatory perturbations, exhibited
soft-glassy rheology. The elastic (storage) modulus scaled as
a weak power-law (G9 ; f0.12060.005) and the loss modulus,
;1/5 G9 at low frequencies (in agreement with the structural
damping relation ? h h¼ tan(pa/2) ¼ 0.191), scaled with the
same power-law dependence on frequency. Weak power-law
behavior of the loss modulus, coupled to the elastic modulus,
is referred to as frictional damping, as opposed to viscous
damping which has a much stronger dependence on
frequency. This behavior has been observed from micro-
rheology measurements of ASM and other cell types with
MTC (Fabry et al., 2001a) and in lung epithelial cells by
AFM measurements (Alcaraz et al., 2003). The structural
damping relation between G9 and G$ is also observed in
tissue-level rheology (Fredberg and Stamenovic, 1989) and
suggests that the elastic and dissipative processes in the cell
are fundamentally linked at the level of the stress bearing
element. An example of such a linkage was postulated using
actomyosin cross-bridge formation and stretching as the
energy storage process and their release as the energy loss
process (Fredberg et al., 1996). In this way the ratio between
G9 and G$ is independent of the number of actomyosin
bridges and only on their cycling rate. Analogously, one
could describe a structural damping mechanism based on
dynamic actin polymerization, where the addition of G-actin
monomers to F-actin filaments acts to resist deformation
(storing elastic energy) and is coupled to the energy loss
process of depolymerization, or removal of actin monomers
Scale-free rheology is an indication of disordered mechan-
ical structure in that the cell cannot be modeled by a defined
set of elastic springs and dissipative sinks yielding a finite set
of characteristic deformation frequencies. Rather, the cells
demonstrate a continuous distribution of relaxation time
constants, and thus there are no distinguishing inflections in
the shear modulus frequency spectrum. The power-law
exponent has been attributed to an effective ‘noise’
(metastable) system existing close tothe glass transition(a ¼
of soft-glassy rheology. Rather, these properties have been
described with a model constitutive equation (Sollich, 1998),
reasoning that mechanical properties are not a result of
specific molecular interactions, but of a higher level of
structural organization (confined structural elements with
a heterogeneous distribution of confinement energies). The
effective noise temperature is expected to be an integrative
function of many forms of molecular agitations, the foremost
of which being cross-bridge cycling and dynamic actin
polymerization. This was confirmed in our study by the
sensitivity of baseline mechanics to both cytochalasin-D and
chain phosphorylation are typically observed in unstimulated
ASM (Kamm and Stull, 1986).
At high frequencies, the loss modulus, G$, contains
a component that is fit by a linear dependence on f and is
attributed to Newtonian viscosity of the cells. Accurate
measurement of this term relies directly on the calibration of
phenomena have the same functional behavior. For this
reason we measured the lever drag and its decay with
increasing height from the cell surface for each probe used.
Errors in this method could arise from hydrodynamic
couplingbetween thelever/fluid motions andthecellsurface.
In this case, simply subtracting the drag calibration from
rheology measurements may not be appropriate as this
effect that depends on cell viscoelasticity. Our calibrations
varied considerably between probes (;30%) and different
surfaces (;10%), but no consistent dependence on surface
compliance was identified (tested over glass, cells, and cells
stimulated with 5-HT). Improved calibration techniques and
modeling of hydrodynamic effects are desired. Interpretation
of Newtonian viscosity in terms of molecular mechanisms is
also difficult. It characterizes the degree of purely dissipative
particle interactions in a fluid, without reference to the
elements involved or specific chemical bond formation/
rapid reshaping, flow, and internal motions in the cell, as it is
has commonly been treated as a constant property of the
3002Smith et al.
Biophysical Journal 88(4) 2994–3007
have shown that this is not true for ASM contractile events
(further discussion below).
Baseline measurements made in this study may not
represent the unperturbed state of the ASM cells. Activation
of a mechanotransduction mechanism is suspected in re-
sponse to the mechanical stimulation of the AFM probing,
often observed as an initial stiffening response over the first
minute or so of the complex modulus measurements. Cyclic
mechanical strain applied to ASM cultures via flexible
substrate techniques has been shown to increase cellular
contractile proteins and force production, Ca21sensitivity,
cytoskeletal and focal adhesion reorganization, and stiffness
(Smith et al., 2000, 2003a), but these changes are observed
after many days of mechanical stimulation. Recently, local-
ized mechanical stress applied to ASM cells with MTC has
been shown to induce actin accumulation and stiffening over
the time scale of our experiments (;30 min; Deng et al.,
2004). Possible mechanical sensing mechanisms have been
proposed for ASM involving tyrosine phosphorylation of
paxillin and focal adhesion kinase (Tang et al., 1999) and
activation of RhoA (Smith et al., 2003b), also implicated in
regulating contractility (Gunst et al., 2003). AFM has been
shown to activate mechanotransduction signals in the form
2001). ASM cells experience mechanical strains in vivo
during breathing, but reproducing physiologically relevant
mechanical conditions for ASM rheological studies is non-
trivial (Latourelle et al., 2002). We have demonstrated the
potential for study of rapid mechanosensitive stiffening in
ASM using AFM, as a mechanical probe which does not rely
on specific membrane-receptor attachment (as does MTC).
Stiffening response to stimulation
We observed dynamic stiffening of ASM cells in response to
the contractile agonist 5-HT. The fractional increase in
storage modulus (150 6 30%) is paralleled by a similar, but
unequal increase in the loss modulus (67 6 15%). The
coupling is in keeping with the structural damping hy-
pothesis for cellular mechanics, but the unbalanced response
is indicative of a more fundamental change of the mechanical
ordering state of the cell. Structural damping predicts that the
hysteresivity parameter (loss tangent, h ¼ G$/G9) is a
constant index of the coupling between storage and loss
processes. After stimulation, the definitive decrease in
hysteresivity (?28 6 6%) signifies the increased prevalence
of the elastic, solidlike nature of the cell. A model proposed
to explain decreased hysteresivity of ASM tissue after con-
tractile stimulation is based on conversion of actomyosin
bridges from the rapidly cycling phosphorylated cross-
bridge state to the more stable latch-state when bound
myosin heads become dephosphorylated (Fredberg et al.,
1996; Murphy, 1994). However, we do not observe the fast
transient increase in h at the onset of stiffening that was
observed in the measurements published by Fredberg et al.
(first 20–30 s, preceding the slow definitive decrease), inter-
preted as the increased cross-bridge cycling from the mostly
inactive baseline state. Transient increases in hysteresivity
have also been observed in MTC experiments (Fabry et al.,
2001b), although the amplitude was smaller and decay was
within 10–15 s. It is possible that a similar transient occurs
before our first indentation after stimulation, but this is
unlikely because the complex modulus usually does not
respond until at least 50–100 s after addition of 5-HT. This
apparent discrepancy is an indication that we are probing
a fundamentally different mechanism of ASM contractile
stiffening. Alternatively, the absence of the hysteresivity
transient could be due to a difference in cross-bridge kinetics
in rat ASM compared to human ASM such as used for the
studies of Fredberg et al. and Fabry et al. (Lecarpentier et al.,
Actomyosin activity is also a common mechanism for
describing the actuation of ASM contraction. It is therefore
intriguing that inhibition of myosin phosphorylation through
MLCK had very little effect on the stiffening response. Both
of the inhibitory agents, wortmannin and ML-7, at the doses
induce phosphorylation andthus activationof myosinmotors
Our observation that contractile shortening is blocked by the
treatments in inhibiting MLCK activity. A recent MTC study
reported a stiffening response of cultured ASM to 5-HT,
by a similar reduction of baseline stiffness), but complete
ablation of the response required a dose of 20 mM (An et al.,
2002). We do not discount the possibility that our MLCK
inhibitory treatments were only partially effective and that
further study of dose-dependent effects is warranted. How-
ever, our observation that ML-7 (5 mM) reduced baseline
stiffness by 50%, yet did not significantly alter the relative
response to 5-HT (Fig. 5), suggests that there was some
sensitivity to myosin activity; but the stiffening response we
measured probed a cellular mechanical process that is largely
the vehicle used for ML-7 delivery (ethanol) was responsible
for the decrease in baseline stiffness as it reduced the
shortening response by 50%.
The conflict of our results with those of An et al. may be
partly due to differences in the measurement techniques. In
MTC, Arg-Gly-Asp (RGD)-containing peptides are used to
couple the measurement beads directly to the cell surface
integrin receptors, key components of focal adhesions or
dense plaques in ASM and known to be the sites of mech-
anical coupling from the extracellular matrix to the in-
tracellular contractile fibers (Burridge et al., 1988; Draeger
et al., 1989; Gunst and Tang, 2000; Schoenwaelder and
Burridge, 1999). Therefore, it is not surprising that stiffness
changes measured with MTC are dominated by the activity
of actomyosin contractile elements. Integrin receptor binding
Smooth Muscle Cell Mechanics by AFM 3003
Biophysical Journal 88(4) 2994–3007
is also known to stimulate a number of biochemical signals
that could have pronounced effects on cellular mechanics
(Meyer et al., 2000; McNamee et al., 1996; Plopper and
Ingber, 1993). The use of chemically nonspecific AFM
probes avoids this potential effect. One can also reason that
the geometry and direction of the applied force in MTC
(lateral twists) and AFM (vertical oscillations) has a signif-
icant influence on the mechanical structures being probed.
These arguments require a detailed examination of the
induced intracellular strains and stress transmission (Charras
et al., 2001), which are particularly difficult to quantify/
model for MTC (Coughlin and Stamenovic, 2003).
Complete frequency-dependent assays before and after
contractile stimulation confirmed the description of contrac-
tile stiffening as a transition toward solidlike glassy rheology
damping to the observed changes in complex rheology.
Further support for the soft-glassy framework is provided
by the extrapolation of our frequency-dependent results to
the universal coordinate (f0, G0) common to all treatments,
leaving the noise temperature (a 1 1) as the key determinant
of the cells’ mechanical state. However, our observation that
by Newtonian viscosity, is dynamic between baseline and
stimulated states is unique. Newtonian viscosity may be
useful in distinguishing between contractile events (myosin
dependent) and other stiffening processes, as its dynamics
during stimulation seems to be blocked by MLCK inhibitors
(unlike the stiffening response). Highlighted by the behavior
under wortmannin treatment, the variations in Newtonian
viscosity response appear to be independent of changes in
stiffness. This supports our claim that Newtonian viscosity
dynamics are not an artifact of hydrodynamic coupling at the
cell surface (stiffness-dependent effect discussed above).
ML-7 treatment also inhibited the 5-HT-induced Newtonian
viscosity increase (but not the stiffness increase). Although
a mechanism for such processes is unclear, we have shown
that Newtonian viscosity, as well as the power-law exponent
(noise temperature), are critical in characterizing ASM
Dynamic actin polymerization
The results presented here point to dynamic actin polymer-
ization as a good candidate for a myosin-independent
stiffening response, due to the high sensitivity to modulation
capping agent, is suspected to only affect actin that is in
a state of dynamic polymerization-depolymerization and not
stable actin filaments. Thus, it can be reasonably stated that
the mechanical integrity of the cells as probed by AFM
surface indentations, relies to a large extent on dynamic
cycling, or treadmilling, of actin filaments (Wegner, 1976),
also known to generate propulsive force in motile cells
(Mogilner and Oster, 2003). Actin filaments of the
contractile apparatus in smooth muscle may be dynamic
(i.e., cytochalasin-D sensitive; Tseng et al., 1997), thus
interpretation in terms of nonmyosin-associated actin de-
pends also on the MLCK inhibitor tests discussed above.
Increased actin polymerization has been previously
observed during contractile events in ASM (An et al.,
2002; Herrera et al., 2004; Hirshman and Emala, 1999;
Mehta and Gunst, 1999). These and subsequent studies
implicate a number of signaling and regulatory molecules of
actin polymerization, which are not directly dependent on
myosin light chain phosphorylation, and myosin ATPase
activity. Gerthoffer and Gunst (2001) review the evidence
for contractile agonist stimulation of tyrosine phosphoryla-
tion of focal adhesion kinase, c-Src, and paxillin, which lead
to actin remodeling through the Rho (or Rac) family of
GTPases and the 27-kDa heat shock protein. Since these
pathways are also involved in mechanotransduction, they
are stimulated by tension development during contraction
and thus are partially dependent on cross-bridge cycling and
force production. Membrane attachments of the actin cyto-
skeleton (via vinculin and talin) are altered and strengthened
during contractile activation (Gunst and Fredberg, 2003).
Other actin regulatory pathways may involve activation or
modulation of phosphoinositides, Arp2/3, profilin, cofilin,
filamin, or a-actinin, as known from studies of nonmuscle
cells (Pollard et al., 2000; Stossel et al., 2001; Yin and
The functional role of dynamic actin polymerization and
remodeling during contraction of ASM is suggested to be
necessary for optimal energy utilization during contraction
enhance force transmission from the contractile apparatus to
the extracellular matrix (and thus to neighboring cells) by
strengthening the region surrounding focal adhesions (Gunst
and Fredberg, 2003; Janmey, 1998; Kuo and Seow, 2004).
mechanisms may enhance the cytoskeletal-membrane junc-
tion, strengthening specifically at sites of high tension or
strain. Similarity of the viscoelastic characteristics of the
contractile apparatus, the connective layer beneath the
membrane, and the extracellular matrix is of critical
importance for cooperative, coordinated muscle contraction
1989). Therefore, this study stands as an important step in
Atomic force microscopy provides a unique tool to probe the
nanoscale mechanical structure and complex rheology at the
surface of living cells, with the ability to track variations of
these properties in the space, time, and frequency domains.
We show that force volume imaging can identify mechanical
3004Smith et al.
Biophysical Journal 88(4) 2994–3007
structures of interest for modeling cytoskeletal mechanics.
The indentation modulation technique allowed for measure-
ment of the elastic and dissipative components of the
complex shear modulus, which increased dramatically with
contractile stimulation and showed a definitive shift in
fundamental mechanical behavior. We presented evidence
implicating actin polymerization and not actomyosin activity
as the dominant mechanism actuating the stiffening re-
sponse. Thus, actin dynamics has a prevalent role in near-
surface mechanics of ASM cells during contraction. The
frequency dependence of the complex modulus under all
treatments supports the soft-glassy rheology model of
cellular mechanics with an additional term describing the
Newtonian viscosity (or purely fluid behavior) at high fre-
quencies. The soft-glassy behavior of ASM cells, governed
largely by the level of molecular agitations (noise temper-
ature) is the key determinant of the cells’ mechanical state.
We revealed significant changes in the Newtonian viscosity
term that vary independently of the soft-glassy properties
(showing stronger correlation with actomyosin activity).
Thus, with further study of frequency-dependent ASM cell
surface rheology, it should be possible to distinguish features
of actin polymerization and cross-bridge interactions during
cellular mechanical events. Measurement and characteriza-
tion of these phenomena require a precision nanoscale probe
with picoNewton force sensitivity and detailed analysis of
frequency-dependent complex rheology, as is possible with
We greatly appreciate the technical assistance of Jamilah Saeed.
We greatly appreciate the financial support granted by the Natural Science
and Engineering Council of Canada (NSERC Discovery Grant and
NanoInnovation Platform) as well as the Canadian Institute for Health
Research (CIHR No. MOP-10381).
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C. A. Schoenenberger, and J. H. Hoh. 1998. Relative microelastic
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