Journal of Cell Science
Vinculin tension distributions of individual stress
fibers within cell–matrix adhesions
Ching-Wei Chang and Sanjay Kumar*
Department of Bioengineering, University of California Berkeley, Berkeley, CA 94720, USA
*Author for correspondence (firstname.lastname@example.org)
Accepted 17 April 2013
Journal of Cell Science 126, 3021–3030
? 2013. Published by The Company of Biologists Ltd
Actomyosin stress fibers (SFs) enable cells to exert traction on planar extracellular matrices (ECMs) by tensing focal adhesions (FAs) at
the cell–ECM interface. Although it is widely appreciated that the spatial and temporal distribution of these tensile forces play key roles
in polarity, motility, fate choice, and other defining cell behaviors, virtually nothing is known about how an individual SF quantitatively
contributes to tensile loads borne by specific molecules within associated FAs. We address this key open question by using femtosecond
laser ablation to sever single SFs in cells while tracking tension across vinculin using a molecular optical sensor. We show that
disruption of a single SF reduces tension across vinculin in FAs located throughout the cell, with enriched vinculin tension reduction in
FAs oriented parallel to the targeted SF. Remarkably, however, some subpopulations of FAs exhibit enhanced vinculin tension upon SF
irradiation and undergo dramatic, unexpected transitions between tension-enhanced and tension-reduced states. These changes depend
strongly on the location of the severed SF, consistent with our earlier finding that different SF pools are regulated by distinct myosin
activators. We critically discuss the extent to which these measurements can be interpreted in terms of whole-FA tension and traction
and propose a model that relates SF tension to adhesive loads and cell shape stability. These studies represent the most direct and high-
resolution intracellular measurements of SF contributions to tension on specific FA proteins to date and offer a new paradigm for
investigating regulation of adhesive complexes by cytoskeletal force.
Key words: Cell mechanics, Mechanotransduction, Stress fibers, Focal adhesions, Laser ablation, Fluorescence resonance energy transfer
Over the past decade, it has become clear that the application of
tensile force by cells on their surroundings plays key roles in cell
shape determination, motility, and fate decisions (Dupont et al.,
2011; Fu et al., 2010; Levental et al., 2009; Prager-Khoutorsky
et al., 2011). Mammalian cells cultured on solid supports
generate these tensile forces in part through contractile stress
fibers (SFs), which are assemblies of filamentous actin (F-actin),
actin-binding proteins, and myosin motors. SFs contribute to
cytoskeletal pre-stress by anchoring into focal adhesions (FAs) at
the cell–extracellular matrix (ECM) interface, which in turn
enables the cell to generate traction against its microenvironment
(Deguchi et al., 2006; Etienne-Manneville and Hall, 2002;
Hotulainen and Lappalainen, 2006). While the molecular
mechanisms through which FAs sense and transmit tensile
forces between SFs and the ECM remain controversial, it is
known that some FA proteins can undergo load-dependent
conformational changes, which can trigger biochemical events
critical to downstream signaling such as phosphorylation and
protein–protein binding (Cohen et al., 2006; Hyto ¨nen and Vogel,
2008; Sawada et al., 2006; Vogel, 2006). Thus, understanding
cell–ECM mechanotransduction requires insight into how tensile
forces generated by single SFs are spatially distributed across
these molecular mechanosenors within FAs, where they may
ultimately contribute to downstream signaling.
We and others have found femtosecond laser nanosurgery to be
a powerful tool for measuring the contractile properties of single
SFs in living cells (Colombelli et al., 2009; Colombelli et al.,
2006; Kumar et al., 2006; Lele et al., 2006; Rauzi et al., 2008;
Ronchi et al., 2012; Russell et al., 2009; Tanner et al., 2010). In
this method, a cellular SF is visualized and irradiated with a
femtosecond laser, resulting in SF scission and retraction. The
retraction dynamics yield quantitative information about the
viscoelastic properties of the fiber, and the resulting change in
cell shape lends insight into the contribution of the targeted SF to
cell shape stability. We have previously used this approach to
demonstrate that the mechanical properties of a SF depend
strongly on whether the fiber is located at the cell center or
periphery (Tanner et al., 2010), and that this is related to the
differential control of these two SF populations by the myosin
activators Rho-associated kinase (ROCK) and myosin light chain
kinase (MLCK), respectively (Katoh et al., 2011; Katoh et al.,
2007; Totsukawa et al., 2004). Moreover, disruption of peripheral
contraction, which has led us to speculate that central and
peripheral SFs distribute their tensile loads differently across the
cell–ECM interface and that this in turn gives rise to their
differential contributions to cell shape stability. This could
potentially be accompanied by and reflected in changes in tension
across specific mechanosensors within individual FAs.
Genetically encoded probes, particularly those based on
fluorescence resonance energy transfer (FRET), have now
made it possible to directly visualize the activation of specific
cellular signal transduction pathways in a spatially and
temporally defined fashion. Most notably, Grashoff et al.,
recently developed a FRET sensor of tension-induced strain
Journal of Cell Science
within the FA protein vinculin, which is capable of sensing
piconewton (pN) loads in migrating cells (Grashoff et al., 2010).
In this vinculin tension sensor (VinTS), the vinculin head and tail
domains are separated by an elastic wormlike chain peptide that
can stretch under tension and is flanked by FRET donor and
acceptor fluorophores (mTFP1 and venus, respectively). Tension-
dependent deformation of this molecule alters the FRET signal in
a graded and dynamic way, enabling visualization of tension
across vinculin within individual FAs throughout the cell.
In this study, we combine this VinTS with femtosecond laser
nanosurgery to determine how single SFs distribute their tensile
loads across vinculin molecules within FAs. We find that while
compromise of a single SF leads to overall reductions in vinculin
tension, it surprisingly does not do so in a spatially or temporally
uniform fashion. Specifically, we find that incision of a single SF
decreases vinculin tension within some FAs while increasing it
on others, and that these changes depend strongly on the
directional alignment of the SF and FA. Moreover, individual
FAs unexpectedly undergo highly dynamic transitions between
vinculin tension-enhanced and tension-reduced states. These
findings differ significantly for central versus peripheral SFs and,
if reflective of tension on the entire FA, may explain the dramatic
difference in how each SF population contributes to cell shape
Mapping tension distributions of single stress fibers
To determine how individual SFs transmit and distribute tensile
introduced a vinculin tension sensor (VinTS) to U373 MG
human glioblastoma cells stably transduced with mCherry
Lifeact (Tanner et al., 2010), severed single stress fibers with
femtosecond laser nanosurgery (Kumar et al., 2006; Tanner et al.,
2010) and tracked the resulting evolution of the mCherry and
VinTS donor and FRET fluorescence intensities (Fig. 1).
Consistent with our previous studies, SF irradiation led to
retraction of the severed ends of the targeted SF (top row), and
inspection of the donor and FRET intensity (middle rows)
revealed that the VinTS localizes to FAs as expected. Because
the FRET intensity also depends on factors unrelated to donor–
acceptor separation (Chang and Mycek, 2012; Chang et al., 2007;
Chang et al., 2009; Zhong et al., 2007), such as the amount of
localized VinTS, we created FRET ratio maps in which we
masked the FRET intensity to show only FAs and normalized this
intensity by the donor intensity (bottom row). In these maps, an
increasing FRET ratio corresponds to an increasing FRET signal
(actual degree of energy transfer) or decreasing tension, which
can in turn be tracked on an FA-by-FA basis. These FRET ratio
maps show that, vinculin tension changes were not restricted to
FAs in direct contact with the targeted SFs as we had expected
but rather were delocalized throughout the entire cell. While
some FAs exhibited significant vinculin tension reduction
(increased FRET ratio), other FAs did not.
Correlating FRET response with contractile activity and
The VinTS FRET sensor reads out tension-induced strain across
the vinculin molecule, which motivated us to investigate the
sensitivity of the FRET signal to wholesale reductions in cellular
contractile force and the predictive value of these molecular
strain changes for traction stresses associated with the entire FA.
To first determine if VinTS FRET signals change in expected
ways following attenuation of contractile signals known to
enhance cellular traction force, we measured total VinTS FRET
ratios following modest pharmacological inhibition of the
myosin activators ROCK (Y-27632) and MLCK (ML-7). These
Fig. 1. Mapping tension distributions of single stress fibers. The
panels depict representative mCherry-Lifeact (top row), vinculin
tension sensor (VinTS) donor (second row), VinTS FRET (third
row), and FRET ratio (bottom row) images immediately before
(0 min) and after (2 min and 4 min) laser ablation. The SF ablation
site is indicated by the arrow in the mCherry image at 0 min. In the
FRET ratio maps, all vinculin signal outside of the FAs depicted was
masked out to facilitate analysis (see the Materials and Methods).
The hotspots in the FRET ratio maps depict regions of increased
FRET ratios following SF ablation, reflecting tension reduction. The
contrast and brightness of all fluorescence images were optimized for
clarity of presentation (see the Materials and Methods). Scale bar:
Journal of Cell Science 126 (14)3022
Journal of Cell Science
treatments moderately reduced cell spread area, SF assembly, and
VinTS localization to FAs without completely disassembling FAs
(not shown). As expected, and as reported previously (Grashoff
et al., 2010), we observed increased VinTS FRET ratio in live cells
within 1 hour (supplementary material Fig. S1), illustrating a
qualitative correlation between the VinTS signal and myosin-
induced traction forces. To determine whether the VinTS FRET
response correlates with local traction at the cell–ECM interface,
we performed traction force microscopy on VinTS-expressing
cells (supplementary material Fig. S2). These measurements
revealed a modest inverse linear correlation between the average
traction force experienced at a VinTS-positive adhesion and the
average FRET ratio within that adhesion (R250.0733, P50.00029
against the null hypothesis of no correlation), with high VinTS
FRETratios uniquely associated
Importantly, however, FAs with low FRET ratios (i.e. high
apparent vinculin tension) were associated with a diverse range of
traction forces, even though the mean traction force for these
adhesions was comparatively high. This may be a consequence of
the relatively high noise floors of the two measurements (both of
which are based on fluorescence readouts) or the possibility that
tension transmitted to vinculin may be dissipated by other FA
proteins and not always reflect whole-FA traction. We therefore
conservatively chose to interpret our measurements in terms of
tension across vinculin.
FA tension falls with SF disruption and depends on SF
Because SFs generate contractile force, one would expect that
disruption of a single SF would reduce the total level of tension
across vinculin within all FAs throughout the cell. To test this, we
severed individual SFs and measured the total FRET ratio signal
across all cellular FAs as a function of time (Fig. 2), initially
restricting our analysis to FAs clearly visible throughout the
entire time course of the experiment (‘4-Image Tracking’). As
expected, the mean FRET ratio increased following SF photo-
disruption, indicating a reduction in overall FA-based vinculin
tension compared to non-irradiated control cells. Given our
earlier finding that central and peripheral SFs exhibit different
viscoelastic properties and contribute differently to cell shape
stability (Tanner et al., 2010), we reasoned that each SF
subpopulation might also contribute differently to overall levels
of vinculin tension within FAs. Indeed, when we segregated the
data according to SF location (Fig. 2A,B), we found that
peripheral SF ablation induced a much greater overall reduction
in vinculin tension than central SF ablation. The dynamics of
these tension changes differed between these two groups as well,
with the peripheral SF FRET ratio evolving over longer time
scales than the central SF FRET ratio changes. As expected, these
characteristic responses were not observed in cells transfected
with a control vinculin FRET sensor lacking the tail domain
(supplementary material Fig. S3).
An important limitation of this analysis is that it requires that a
given FA be clearly identifiable at each of the four time points of
the experiment. However, the reality is that some FAs are lost to
tracking because of changes in focus, the low signal-to-noise ratio,
and other factors. To determine whether exclusion of these FAs
artificially influenced our result, we also tracked FAs and analyzed
FRET ratios in pairs of temporally consecutive images (‘2-Image
Tracking’), which enabled inclusion of the FAs that would have
been excluded in the longer-term analysis (Fig. 2C). The FRET
ratios were analyzed separately in each time interval because an
FA visible in a given temporally adjacent set of images was not
always visible in the preceding or subsequent set of images. For
comparison, we also applied this same analysis to the set of FAs
visible through all four time points (Fig. 2B). This refined analysis
revealed qualitatively similar dynamics, with the cell evolving to a
lower-tension steady-state upon SF disruption and peripheral SFs
contributing more to this tension reduction than central SFs.
SF disruption can either increase or decrease tension
across vinculin within a given FA
The previous results show that photo-disruption of an SF reduces
total FA-based vinculin tension in the cell, which in turn depends
Fig. 2. SF incision reduces total tension in cellular FAs. (A) Overall mean normalized FRET ratio versus time for ‘4-Image Tracking’, i.e. including only
FAs that were clearly visualized throughout all four time points of the experiment. The FRET ratio values were normalized to the corresponding FRET ratio at
time 0 min. The statistics are based on 546 focal adhesions from 17 cells for peripheral SF (PSF) ablation, 376 FAs from 10 cells for central SF (CSF) ablation,
and 164 FAs from 8 cells for non-irradiated control cells. The increase in the control FRET ratio is presumably due to fluorophore photobleaching that was not
fully corrected. [See supplementary material Fig. S3 for vinculin tail-less sensor (Grashoff et al., 2010) results]. (B) Changes in overall mean normalized FRET
ratios across adjacent time points for the set of FAs considered in A (see the text). The FRET ratio changes were normalized to the corresponding FRET ratio at
the first time point of each two-minute interval. (C) Changes in overall mean normalized FRET ratios for FAs that could be tracked across two temporally
consecutive images (‘2-Image Tracking’). As in B, FRET ratio changes were normalized to the corresponding FRET ratio at the first time point of each
two-minute interval. The statistics are based on 681–716 FAs from 17 cells for peripheral SF ablation, 458–485 FAs from 10 cells for central SF ablation, and
226–232 FAs from 8 cells for controls. *P,0.05 compared with corresponding control;+P,0.05 for CSF vs PSF;#P,0.05 compared with the corresponding ‘0 to
2 min’ group. In all cases, the data are means6s.e.m. and statistical comparisons were performed using two-tailed Student’s t-tests.
Tension distributions of stress fibers3023
Journal of Cell Science
on the location of the SF. As we further considered the
relationship between SF geometry and FA tension, we noted
from the FRET ratio maps (Fig. 1) that the FRET signal of a
given FA appeared to depend on the degree to which the FA was
aligned with the severed SF. This is consistent with past
observations that FAs typically align in the direction of net
applied tension, which would be expected to coincide with the
orientation of one or more attached SFs (Goffin et al., 2006; Tan
et al., 2003). To quantify this observation, we defined an
orientation angle between the severed SF and the long axis of
each FA in the cell (Fig. 3A–C) and used this coordinate system
to further explore the geometric distribution of SF tension
(Fig. 3D,E). For control cells in which no SF was severed, we
used the long axis of the cell itself as the reference, which in
practice is frequently parallel to the most prominent cellular SFs.
Equipped with this coordinate system, we plotted FRET ratio
changes within each individual FA as a function of its angle of
orientation to the severed SF (Fig. 4A–C, binned into box-and-
whisker plots). Here, each data point represents the FRET ratio
change for a single FA, with points falling above and below the
horizontal axis (zero normalized FRET ratio change) experiencing
reduced and enhanced vinculin tension, respectively. We thus
and lower than the 5th percentile of the corresponding control
distribution to be tension-reduced and tension-enhanced FAs,
abbreviated as TR and TE, respectively). Approximately 26% of
all FAs fell within the tension-reduced category, and this
subpopulation of FAs was not restricted to the FAs projected to
be in direct contact withtheablated SFs (see below and Fig. 4E for
quantitative analysis). Surprisingly, ,12% of FAs fell within
the tension-enhanced category, reflecting a population of FAs
experiencing increased tension upon SF ablation. In other words,
even though ablation of a single SF reduces overall vinculin
tension in FAs throughout the cell (Fig. 2), these tension
reductions reflectthe net result
redistribution in which tension across vinculin falls in some FAs
and increases in others.
SFs distribute tension in a spatially non-uniform fashion
To understand the dependence of vinculin tension reduction and
enhancement on FA orientation, we considered the angular
distributions of the FRET ratio changes from one time point to
Fig. 3. Coordinate system for measuring relative orientations of SFs and FAs. (A) Schematic of coordinate system. We defined FA angles as the
angles between the long axis of each FA and the axial orientations of the cell (green solid arrow) or the severed SF (yellow solid arrow). The cell-referenced
angle system is also shown (dotted arrow and labels). For clarity, only the image of the SF network (mCherry-Lifeact) is shown. (B,C) Examples of FA
orientations showing their long and short axes. (D) Cell with FA orientations (arrows) overlaid upon a FRET intensity image, with region enclosed by white square
zoomed and shown in (E). The angle of a selected FA is illustrated as h with the severed SF axis as the reference. The colors in (B–E) reflect arbitrary units
of VinTS FRET channel fluorescence intensity. White scale bars: 50 mm; yellow scale bars: 1 mm.
Fig. 4. SFs distribute tension to FAs in an angle-dependent manner.
(A–C) Distributions of normalized FRET ratio changes in individual FAs
following SF ablation. The FRET ratio changes were calculated from 2-
image FA tracking (Fig. 2C) over the indicated time intervals, and then
plotted as a function of the angle between FA long axis and the reference
axis (SF axis for ablated cells and cell long axis for non-ablated controls, as
shown in Fig. 3A). The corresponding central and peripheral SF-ablation
subpopulations are shown in supplementary material Fig. S8. The FRET
ratio changes were normalized to the corresponding FRET ratio at the first
time point of each two-minute interval. The horizontal box lines indicate
25th, 50th and 75th percentiles, the whisker ends indicate 5th and 95th
percentiles, and the solid dots indicate mean values. Each data point
represents one FA. (D–F). Time-dependent histograms of angles between FA
and SF orientations for (D) all FAs, (E) tension-reduced (TR) FAs, and
(F) tension-enhanced (TE) FAs. The curves are the corresponding Gaussian
curve fits. The histograms for central and peripheral SF-ablation
subpopulations are shown in supplementary material Fig. S9. (G–I). Breadth
of tension distributions as a function of time and SF location. (G) Gaussian
widths for all FAs, with the red, green, and blue curves (all, tension-reduced,
and tension-enhanced) corresponding to the Gaussian fits shown in D–F,
respectively. (H,I) Gaussian widths for (H) peripheral SF (PSF) and (I)
central SF (CSF) subpopulations.
Journal of Cell Science 126 (14)3024
Journal of Cell Science
another (Fig. 4A–C). At the earliest time interval following SF
ablation (Fig. 4A), the distribution exhibited a broad peak around
0˚, indicating that these tension changes were preferentially
concentrated in FAs co-aligned with the SF, which are most
likely to be found near the ends of the SF. The significant breadth
of the peak further indicates that these tension changes are also
delocalized across FAs located throughout the cell, which was
evident from the FRET ratio maps (Fig. 1). Over time (Fig. 4B,C),
the angle-dependenceof the
achievement of a new tensile steady-state. Importantly, this
distributions to the maximal amplitude, indicating that the zero-
angle FRET ratio peak was not simply a sampling artifact
associated with the high number of FAs found at this angle
(supplementary material Fig. S5) (Moore and McCabe, 2002).
To further quantify and understand the dynamics of this angle
specificity, we asked whether FAs exhibiting strong changes
in vinculin tension are preferentially enriched in specific
orientations relative to the ablated SF. To do this, we
constructed histograms of FA number as a function of
orientation angle for all FAs, vinculin tension-reduced FAs and
vinculin tension-enhanced FAs (Fig. 4D–F, with tension-reduced
FAs and tension-enhanced FAs defined as described above and as
depicted in supplementary material Fig. S4 and Table S1). Here,
the total FA histogram (Fig. 4D) serves as a reference, where
distributions narrower or wider than this indicate an angle-
specificity close to or away from the targeted SF orientation,
respectively. Analysis of this angle-specificity, which we
quantified as the width of the corresponding Gaussian fit
(Fig. 4G), revealed two important features of the tension-
reduced and tension-enhanced populations. First, the angle-
specificities of both populations were significantly more
dynamic than the overall FA population, which displayed a
relatively constant distribution width throughout the time course
(Fig. 4G, circles). This constant distribution also illustrates that
FA orientations were relatively stable throughout the time
course, with minimal rotation. Second, the magnitude of the
distribution widths were significantly narrower for the tension-
enhanced and tension-reduced populations, particularly at early
time points, indicating that both subpopulations are most likely
to be found in FAs co-aligned with the SF and presumably found
near SF ends.
Central and peripheral SFs produce different vinculin
As mentioned earlier, central and peripheral SFs contribute
differently to cell shape stability (Tanner et al., 2010), which
suggests that these two SF subpopulations spatially distribute
tension in distinct ways and that this might be reflected in
differential vinculin tension delocalization following ablation of
each subpopulation of SFs. To test this hypothesis, we plotted the
distribution widths of these two subpopulations separately
(Fig. 4H,I). Within 2 min of peripheral SF ablation (Fig. 4H),
tension-reduced and especially tension-enhanced FAs were
distributed much more narrowly in angular orientation than all
FAs. This indicates that the majority of vinculin tension changes,
particularly enhancements, occurred in FAs oriented along the
axis of the targeted SF following photo-disruption. This strongly
contrasted with the angle specificity dynamics following central
SF ablation (Fig. 4I), in which tension-enhanced FAs (inverted
triangles) were initially distributed much more broadly than
either the total (circles) or tension-reduced FA (upright triangles)
populations. We reasoned that the qualitatively symmetric but
opposite kinetics displayed by vinculin within tension-enhanced
and tension-reduced FA populations might be due to exchange of
tension between different subsets of FAs, potentially mediated by
physical connections within the cellular SF network.
Vinculin undergoes highly dynamic tension transitions
following SF ablation
If tension indeed can be transferred between FAs through the SF
network following SF ablation, one would expect to observe time
dependent changes in the tensile state of vinculin within
individual FAs. To explore this, we followed the FRET ratio
for individual FAs following SF ablation and mapped the
trajectory of each FA through the three tensile states (non-
tension-reduced/tension-enhanced, tension-reduced, and tension-
enhanced) over the three time intervals examined above. (Fig. 5;
in these representations, the intensity of the color reflects the
number of FAs in a given category for a given time interval, and
the width of the transition line reflects the number of FAs
transitioning between two specific states from one time interval
to another). As expected from our earlier analysis (Fig. 4A), most
FAs were in the non-tension-reduced/tension-enhanced category
and remained so for the entire observation period. However, the
Fig. 5. FAs undergo dynamic tension transitions following SF ablation. Tension state transition plots for (A) all SFs, (B) peripheral SF (PSF), and (C) central
SF (CSF) subpopulations. FAs were analyzed from 4-image tracking data depicted in Fig. 2A,B. In these diagrams, each of the nine tiles represents the category
into which a given FA falls [tension-reduced (TR), tension-enhanced (TE), non-tension-reduced/tension-enhanced] for a given time interval, with the color
representing the number of FAs (see scale on right-hand side). The lines between temporally adjacent tiles reflect FAs transitioning from one state to another, with
the thickness of each line indicating the number of FAs undergoing that particular transition (which can include remaining in the same state). Scale bar thickness:
50 FAs (note variation of scale between panels).
Tension distributions of stress fibers3025
Journal of Cell Science
smaller subset of FAs in the tension-enhanced and tension-
reduced states predominantly
transitions between the three states. In cases where non-
dynamics, more transitioned from the non-tension-reduced/
tension-enhanced state to the tension-reduced than to the
tension-enhanced state, consistent with our earlier finding that
SF ablation induces an overall tension reduction (Fig. 2A).
Moreover, for the peripheral SF population (Fig. 5B), the
number of tension-reduced FAs continuously decreased, again in
agreement with our previous finding (Fig. 2A), while those
of tension-enhanced FAs, and non-tension-reduced/tension-
enhanced FAs continuously increased (indicated by the color
scale), despite the fact that ablation of a peripheral SF leads to
overall reductions in tension (Fig. 2A). A quantitative analysis of
these tensional transitions indicated that the most significant
(Fig. 5B,C) lay in the percentage of adhesions that begin in the
tension-reduced state at one time interval and then transition to
the tension-enhanced state in the subsequent time interval (43.5%
and 43.9% for central SFs for the earlier and later transitions
versus 25.0% and 35.4% respectively for peripheral SFs).
This could potentially be explained by greater mechanical
‘communication’ between FAs following central SF photo-
disruption, which would also explain why the numbers of both
tension-reduced and tension-enhanced FAs were more stable for
the central SF population (Fig. 5C) than the peripheral SF
population. In other words, tension dissipated by compromise
of a central SF is redistributed to vinculin within a broader
comparatively few tensional transitions are observed.
To gain insight into the relationship between these transitions
and myosin-dependent contractile activity, we pharmacologically
pre-treated cells with the MLCK inhibitor ML-7 and the ROCK
inhibitor Y-27632 prior to SF ablation (supplementary material
Fig. S6). While some trends were preserved (e.g. decreasing
numbers of tension-reduced FAs but increasing numbers
of tension-enhanced and non-tension-reduced/tension-enhanced
FAs along the time course; see 1st column, supplementary
material Fig. S6), both treatments enhanced the total number of
transitions following SF ablation, suggesting that mechanical
communication between FAs is sensitive to motor activity within
the actomyosin cytoskeleton.
To conceptually unify all of these findings, we devised a
structural model in which central SFs are more physically
interconnected and mutually reinforced than peripheral SFs due
to the presence of transverse actomyosin structures that link
central SFs into a cohesive network. This model is supported by
between central SFs but comparatively few between peripheral
SFs (Fig. 6A–D; supplementary material Fig. S7 illustrates our
ability to sever single central SFs). In our model, these lateral
connections explain the differences in tension redistribution and
shape stability associated with compromise of central and
Fig. 6. Structural model of SF tension distribution across cellular FAs. (A–C) Confocal fluorescence images depicting actin cytoskeleton (phalloidin). Red
boxes highlight regions where lateral interconnections between central SFs are clearly visible. Arrows indicate peripheral SFs, which exhibit these lateral
connections to a much lesser extent. (D) Live-cell image illustrating actomyosin cytoskeletal structure (mCherry-Lifeact, red) and FAs [overlaid VinTS donor
(blue) and FRET (green) channels]. White dotted arrows indicate FAs that are associated with the transverse actomyosin structures. Scale bar: 20 mm.
(E–G) A simplified model for tension re-distribution after (F) central and (G) peripheral SF photo-disruption, following disruption of the mechanical equilibrium
depicted in (E). FAs are represented by green ovals, with a darker green color indicating higher tension experienced by that FA. Blue arrows represent the involved
force vectors along SFs, with stronger force reflected by thicker arrows. The red solid or dashed lines depict the SFs. The ablation site is indicated by an ‘X’. In
this model, the interconnected central SFs re-direct tension to a broader array of FAs with a diversity of cellular locations and orientations, which preserves cell
shape after SF ablation. Conversely, peripheral SFs, which are not as structurally coupled into the total SF network, distribute dissipated tension to a narrower
segment of cellular FAs with similar orientations. Thus, peripheral SF disruption is more likely to trigger FA rupture and cellular contraction. TE, tension-
enhanced FA; TR, tension-reduced FA.
Journal of Cell Science 126 (14)3026
Journal of Cell Science
peripheral SFs. We specifically speculate that these lateral
connections provide physical conduits for the dissipation of
released tension to vinculin mechanosensors localized with
multiple FAs (Fig. 6E–G). If the tensile state of vinculin
reflects tension in the entire FA (an assumption discussed at
length below), disruption of a central SF leads to robust
redistribution of the released tension to FAs throughout the
entire cell and thus is not strongly accompanied by compromise
of cell shape, whereas disruption of a peripheral SF produces a
relatively narrow spatial redistribution of tension and thus is
accompanied by FA rupture and catastrophic collapse of cell
shape. In both cases, the frequency of these transitions increases
in the setting of myosin inhibition. While the mechanism of this
myosin dependence remains unclear, one possibility is that the
reduced actomyosin sliding velocity associated with myosin
inhibition reduces viscous dissipation of tension upon SF
retraction (Besser and Schwarz, 2007; Colombelli et al., 2009),
which in turn frees this tension to be redistributed throughout the
In this study, we have elucidated how tensile forces generated by
single SFs are temporally and spatially distributed to vinculin
mechanosensors located within cell–matrix adhesions by using
femtosecond laser nanosurgery and a FRET-based tension sensor.
Our results demonstrate that following SF photo-disruption,
vinculin localized to FAs throughout the cell can undergo both
tension reduction and enhancement. The magnitude of vinculin
tension change in an FA depends strongly on the co-alignment
of the FA and disrupted SF, and the spatial and temporal
redistribution of vinculin tension following incision differs
dramatically for central and peripheral SFs. Specifically, incision
tension within FAs throughout the cell and a highly dynamic set of
tension transitions, which is consistent with the observation that
cell structure is relatively stable to photo-disruption of this SF
It is important to note that our measurements are based on the
use of a sensor that tracks tension borne by a specific FA
molecule, vinculin, which populates the FA in the context of both
endogenous vinculin and the multitude of other proteins that
localize to FAs (Kanchanawong et al., 2010). It remains unclear
whether this readout may be used as a surrogate for tension
within the entire FA or traction force generated by an FA against
the ECM. Several lines of evidence support a correlation between
these three quantities: First, pharmacologic inhibition of
contractility is accompanied by an increase in VinTS FRET
ratio across the whole cell (supplementary material Fig. S1).
Second, the pattern in which SFs distribute vinculin tension in
this study qualitatively resembles the pattern with which SFs are
known to distribute traction forces (Kumar et al., 2006). Third,
we observe a gross correlation between VinTS FRET signal and
traction force for a given adhesion (supplementary material Fig.
S2). Nonetheless, this last correlation is a comparatively modest
one, with most FAs generating small VinTS FRET signals and
being associated with a broad range of traction force values. As
discussed earlier, this may be a consequence of the relatively
high noise floor in the FRET measurements (leading to
overrepresentation of low-FRET adhesions) or the possibility
that changes in sensor tension do not always reflect the tensile
state of the whole adhesion. This second notion is consistent with
the very real possibility that tension across vinculin is highly
regulated within FAs and may vary in non-intuitive ways
with overall FA tension, e.g. through altered protein–protein
interactions or maturation state. We do not regard our study as
definitively resolving this question either way, and we suspect
that revisiting these measurements with higher-resolution TFM
measurements, improved FRET pairs (Lam et al., 2012), and
analogous sensors based on other FA proteins should help clarify
Nonetheless, it is valuable to consider the implications for
cellular structure and mechanics if these measurements indeed
reflect alterations in whole-FA tension and cell–ECM traction.
Our findings would challenge the common notion that the tension
generated by a given SF is concentrated primarily near its points
of attachment. More generally, our finding that stress fiber
network connectivity plays a role in intracellular force
redistribution supports and lends direct mechanistic insight into
the emerging idea that specific cytoskeletal elements can rapidly
transmit mechanical information across specific positions within
the cell. For example, application of stress to integrin-based
adhesions can deform organelles (Hu et al., 2005) and activate
mechanotransductive signals at distal cellular locations less than
one second after imposition of force, in contrast to canonical
receptor–ligand signaling (Na et al., 2008; Wang et al., 2009).
Our findings hint that the physical ‘wiring’ of the SF network
may directly shape the speed and directionality of this
transmission process, with different routes channeling varying
degrees of tension to different adhesive nodes at the cell–matrix
An important feature of our model (again, assuming that
vinculin tension reflects whole-FA tension and traction) is that
lateral connections between SFs provide a mechanism to broadly
distribute tension released from a severed SF and thus maintain
cellular structural integrity. This may be placed in context of past
studies of SF network structure and its possible role in cellular
tension transmission (Hotulainen and Lappalainen, 2006; Hwang
and Barakat, 2012; Rossier et al., 2010; Tondon et al., 2012). In
particular, SFs have been broadly classified into three different
categories: transverse arcs, dorsal SFs, and ventral SFs.
anteroposterior axis of the cell, physically couple and may later
evolve into ventral stress fibers, and do not closely associate with
FAs. Our model would predict that these transverse arcs
function to mechanically couple other SFs and distribute load.
Interestingly, the transverse structures in the cells considered here
sometimes do appear to anchor into FAs (Fig. 6D), suggesting
that these SFs may be in the process of evolving into ventral SFs.
High-resolution live-cell time-lapse imaging on SF assembly and
evolution in these experiments may lend insight into this
possibility. Moreover, recent modeling efforts strongly support
a role for SFs aligned orthogonally to the direction of force
application in the rapid and robust transmission of force across
the cell (Hwang and Barakat, 2012); in our system, the applied
force is the tension released upon stress fiber incision, and the
orthogonal population of SFs is the transverse SFs. Thus,
different SF subpopulations may play diverse and previously
unappreciated mechanical roles in maintaining shape stability
through the differential transmission of tensile information.
More specifically for our studies, central SF photo-disruption
causes spontaneous vinculin tension enhancement in FAs
with various orientations due to force transmission through the
Tension distributions of stress fibers3027
Journal of Cell Science
inter-connection of the targeted SF and the transverse SFs
(Fig. 6E,F), which facilitates tension transmission to a broader
area and hence dissipates the mechanical disturbance more
quickly. Therefore, the high resistance of the cell to structural
changes following central SF ablation relative to peripheral SF
ablation (Tanner et al., 2010) can be attributed not only to
differences in initial tension (Fig. 2) but also to differences in the
physical connectivity of those SFs to the transverse SF network.
Another unexpected and very interesting discovery revealed by
our studies is the highly dynamic transitions in vinculin tension
enhancement/reduction states following SF ablation (Fig. 5). If
our FRET measurements reflect whole-FA tension and traction
changes, this would provide additional molecular-scale evidence
that FAs mutually transmit force to one another, potentially as
part of a shape-stabilization strategy. Importantly, while both
peripheral and central SFs exhibit these transitions, peripheral
SFs do so to a much lesser extent, and appear to concentrate those
tension changes in a more localized pool of FAs. Somewhat
unexpectedly, ML-7 and Y-27632 pre-treatments of cells
increased the number of transitions. We hypothesize that this
effect is related to the viscous drag of actin–myosin sliding, as
described in previous mechanical models of SFs (Besser and
Schwarz, 2007; Colombelli et al., 2009). Specifically, when SFs
are severed in the absence of myosin inhibition, sliding and SF
retraction occur rapidly, and the resulting viscous drag dissipates
tensile force that would otherwise be transmissible to other SFs
and adhesions. When myosin is inhibited, sliding velocity and
viscous drag fall, and the dissipated tension is more easily
transmitted throughout the SF network. Importantly, this does not
imply that the magnitude of transmitted tension is greater upon
myosin inhibition, only that the number of transition events
increases. Related to (and consistent with) this framework, recent
numerical simulations (Besser et al., 2011) of post-ablation SF
dynamics reveal damped oscillations in the ends and interior of
the severed fiber, which in turn derives from the competing
influences of SF elasticity and frictional coupling to the
cytoplasm. While the technical demands of our combined
ablation/FRET measurements did not allow us to directly
visualize these oscillations in our system, it is conceivable that
these oscillations may be contributing to the vinculin tension
transitions we observe at individual FAs. The plausibility of this
hypothesis could be tested in the future via construction of
multiscale models that incorporate SF viscoelastic properties, SF
network architecture, and FA-based mechanosensing, as well
as experimental studies with myosin mutants with defined
Our studies also raise a number of important open questions for
future investigation. The first is the overall utility of tension
across vinculin as a readout of whole-FA tension and traction,
which as described above could be addressed with higher-
resolution measurements and a broader palette of optical sensors.
Second, it would be informative to determine how SF tension
distributions depend on the organization and anisotropy of the
overall SF network, which can now be controlled to some extent
by micropatterning (Ronchi et al., 2012; The ´ry, 2010). By
dictating FA geometry and thus the length and orientation of
intervening SFs, such measurements may also enable elucidation
of length–tension relationships for SFs, which have thus far been
difficult to clearly measure in living cells. These methods may
also enable more precise control of SF thickness, which could
et al.,2009) and
potentially make it possible to understand whether the thickness
can alter the roles of SFs at different locations. Finally, by
molecular biosensors, this study introduces a significant
methodological advance that may be broadly applicable for
quantitatively studying mechanochemical coupling between load-
bearing cytoskeletal elements and the signaling complexes into
which they attach, such as insertion of actin bundles into cadherin
complexes (Borghi et al., 2012; le Duc et al., 2010). This strategy
may also potentially be generalized to analogous interactions
between load-bearing elements in tissue (e.g. ECM fibers) and
their cognate sites of anchorage (e.g. integrin-based adhesions).
Clearly, progress in this area will be greatly aided by continued
methodological advances in photoablation technology and the
development of new optogenetic force sensors.
Materials and Methods
Cell culture, transduction, and transient transfection
U373 MG human glioblastoma cells were stably transduced with mCherry-Lifeact
(Tanner et al., 2010) and then transiently transfected with a previously described
vinculin tension sensor (VinTS) (Grashoff et al., 2010) (Addgene, Plasmid ID#:
26019) using polyethyleneimine-based transfection (Tinsley et al., 2004; Zhang
et al., 2004). The transfected cells were plated on 35 mm 1.5 coverslip-bottomed
dishes (0.16–0.19 mm, MatTek Corporation, Ashland, MA) coated with
fibronectin at a density of 10 mg/cm2, and then cultured at 37˚C supplied with
5% CO2 until imaging. Cultures were imaged at low (,40%) confluence to
minimize or eliminate cell–cell contacts that may play a role in cellular responses
after SF photo-disruption.
Confocal imaging and photo-disruption of single SFs
experiments were performed on a Zeiss LSM 510 Meta confocal microscope
equipped with a MaiTai Ti:sapphire femtosecond laser (Spectra Physics, Newport
Beach, CA) based on previously published protocols (Kumar et al., 2006; Tanner
et al., 2010). 35 mm dishes with fluorescent live cells were placed on a plate heater
(Warner Instruments) set at 37˚C during imaging. mCherry-Lifeact images
(excitation: 543 nm, emission: 565–615 nm), VinTS FRET channel images
(excitation: 458 nm, emission: 533–587 nm), and VinTS donor channel images
(excitation: 458 nm, emission: 469–501 nm) were obtained by one-photon
confocal imaging with a 406 water-dipping objective (NA50.8). For SF photo-
disruption, the femtosecond laser was used at 770 nm for one iteration, resulting in
energy deposition of 1–2 nJ on a single stress fiber. All images were acquired with
5126512 pixels and pixel size50.41 mm. For presentation purposes, the contrast
and brightness of fluorescence images were optimized using ImageJ.
imaging,and SF photo-disruption
FRET ratio mapping
VinTS FRET and donor channel images were first background-subtracted and
corrected for photobleaching, which was implemented by taking the overall
fluorescence intensity of each image excluding the background and the identified
FA regions (see below), and using this overall intensity value to normalize each
image to its corresponding image at the first time point. Note that this approach
assumes that the majority of cytoplasmic VinTS is subject to photobleaching but
experiences no dramatic tension change. FA FRET ratio maps were then generated
by taking the ratio of the FRET channel signals to the VinTS donor channel
signals, using the same identified FA masks (see below).
Traction force microscopy
Traction force microscopy was performed according to our previous studies
(MacKay et al., 2012; Sen et al., 2009) using fibronectin-coated 30 kPa
Technologies, Cat. No.: F8816, F8807, and F8799). Traction maps were first
obtained using Fourier transform traction cytometry (Butler et al., 2002) and then
subjected to an FA mask in order to isolate traction forces at adhesions [adapted
from (Gardel et al., 2008; Plotnikov et al., 2012; Stricker et al., 2011; Stricker et al.,
Single FA identification, tracking, and morphometric/orientational analysis
FAs in VinTS fluorescence intensity images were identified using the previously
described ‘water algorithm’ (Grashoff et al., 2010; Zamir et al., 1999) with merger
size520 pixels (3.36 mm2) and FA reject size55 pixels (0.84 mm2). Briefly, after
background subtraction, all pixels were sorted from highest to lowest intensity, and
each pixel was inspected one-by-one to determine if it belonged to a certain FA. If
Journal of Cell Science 126 (14)3028
Journal of Cell Science
the pixel was not in contact (8-point connectivity) with any previous pixels, it
would be assigned to a new FA; otherwise, if the pixel was in contact with only
one previously assigned FA, it would be assigned to that FA. When a pixel was in
contact with more than one previously assigned FAs, these FAs would be merged
to form a new FA if the size of at least one of these FAs was less than the merger
size; otherwise, the pixel would be assigned to the brightest FA, with no merger.
This process continued until each pixel was assigned to an FA, and then all FAs
with sizes less than the reject size would be removed.
After FA identification, images were shifted, if necessary, relative to the image
at the first time point to remove any unexpected xy-plane displacement and/or
drifting during imaging. Tracking was then applied to the identified FAs using a
previously reported IDL particle tracking method (Cheezum et al., 2001; Crocker
and Grier, 1996). In our FA tracking, the maximum displacement was set to 10
pixels (4.1 mm) and no disappearing of FA was allowed at any time point.
FA orientation was then determined with a gradient-based structure tensor (also
known as the second-moment matrix) method (Zamir et al., 1999) for those
successfully identified and tracked FAs. After determining the two ellipse axes of
an FA (Fig. 3B,C) the orientation of the FA was assigned to be the longer axis with
the upward direction (the vector in the 1st or 2nd quadrants in the Cartesian
coordinate system). The angle of orientation was then reassigned to be within 90˚
and 290˚relative to either the severed SF or the cell orientation (see below), both
in the upward direction as well (Fig. 3A).
For cell orientation (long axis) determination, we first considered all vectors in
1˚increments in the upward (positive y) direction passing through the cell center of
mass, and then calculated the sum of squares of the shortest distances of all pixels
in the cell to the vector. The orientation was determined to be the vector that
produced the minimum sum.
C-.W.C. was responsible for conception and design, collection and
assembly of data, data analysis and interpretation, and manuscript
writing. S.K. was responsible for obtaining financial support,
conception and design, data analysis and interpretation, manuscript
writing, and final approval of manuscript.
This work was supported by the National Institutes of Health
[Director’s New Innovator Award 1DP2OD004213 and Physical
Sciences–Oncology Center Award 1U54CA143836 to S.K.]; and the
National Science Foundation [CAREER Award CMMI 1055965 to
S.K.]. Deposited in PMC for release after 12 months.
Supplementary material available online at
Besser, A. and Schwarz, U. S. (2007). Coupling biochemistry and mechanics in cell
adhesion: a model for inhomogeneous stress fiber contraction. New J. Phys. 9, 425.
Besser, A., Colombelli, J., Stelzer, E. H. and Schwarz, U. S. (2011). Viscoelastic
response of contractile filament bundles. Phys. Rev. E Stat. Nonlin. Soft Matter Phys.
Borghi, N., Sorokina, M., Shcherbakova, O. G., Weis, W. I., Pruitt, B. L., Nelson,
W. J. and Dunn, A. R. (2012). E-cadherin is under constitutive actomyosin-
generated tension that is increased at cell-cell contacts upon externally applied stretch.
Proc. Natl. Acad. Sci. USA 109, 12568-12573.
Butler, J. P., Tolic ´-Nørrelykke, I. M., Fabry, B. and Fredberg, J. J. (2002). Traction
fields, moments, and strain energy that cells exert on their surroundings. Am. J.
Physiol. 282, C595-C605.
Chang, C. W. and Mycek, M. A. (2012). Quantitative molecular imaging in living cells
via FLIM. In Reviews in Fluorescence (ed. C. D. Geddes), pp. 173-198. New York,
Chang, C. W., Sud, D. and Mycek, M. A. (2007). Fluorescence lifetime imaging
microscopy. Methods Cell Biol. 81, 495-524.
Chang, C. W., Wu, M., Merajver, S. D. and Mycek, M. A. (2009). Physiological
fluorescence lifetime imaging microscopy improves Fo ¨rster resonance energy transfer
detection in living cells. J. Biomed. Opt. 14, 060502.
Cheezum, M. K., Walker, W. F. and Guilford, W. H. (2001). Quantitative comparison
of algorithms for tracking single fluorescent particles. Biophys. J. 81, 2378-2388.
Cohen, D. M., Kutscher, B., Chen, H., Murphy, D. B. and Craig, S. W. (2006). A
conformational switch in vinculin drives formation and dynamics of a talin-vinculin
complex at focal adhesions. J. Biol. Chem. 281, 16006-16015.
Colombelli, J., Pepperkok, R., Stelzer, E. H. K. and Reynaud, E. G. (2006). [Laser
nanosurgery in cell biology]. Med. Sci. (Paris) 22, 651-658.
Colombelli, J., Besser, A., Kress, H., Reynaud, E. G., Girard, P., Caussinus, E.,
Haselmann, U., Small, J. V., Schwarz, U. S. and Stelzer, E. H. (2009).
Mechanosensing in actin stress fibers revealed by a close correlation between force
and protein localization. J. Cell Sci. 122, 1665-1679.
Crocker, J. C. and Grier, D. G. (1996). Methods of digital video microscopy for
colloidal studies. J. Colloid Interface Sci. 179, 298-310.
Deguchi, S., Ohashi, T. and Sato, M. (2006). Tensile properties of single stress fibers
isolated from cultured vascular smooth muscle cells. J. Biomech. 39, 2603-2610.
Dupont, S., Morsut, L., Aragona, M., Enzo, E., Giulitti, S., Cordenonsi, M.,
Zanconato, F., Le Digabel, J., Forcato, M., Bicciato, S. et al. (2011). Role of YAP/
TAZ in mechanotransduction. Nature 474, 179-183.
Etienne-Manneville, S. and Hall, A. (2002). Rho GTPases in cell biology. Nature 420,
Fu, J., Wang, Y. K., Yang, M. T., Desai, R. A., Yu, X., Liu, Z. and Chen, C. S.
(2010). Mechanical regulation of cell function with geometrically modulated
elastomeric substrates. Nat. Methods 7, 733-736.
Gardel, M. L., Sabass, B., Ji, L., Danuser, G., Schwarz, U. S. and Waterman, C. M.
(2008). Traction stress in focal adhesions correlates biphasically with actin retrograde
flow speed. J. Cell Biol. 183, 999-1005.
Goffin, J. M., Pittet, P., Csucs, G., Lussi, J. W., Meister, J. J. and Hinz, B. (2006).
Focal adhesion size controls tension-dependent recruitment of alpha-smooth muscle
actin to stress fibers. J. Cell Biol. 172, 259-268.
Grashoff, C., Hoffman, B. D., Brenner, M. D., Zhou, R., Parsons, M., Yang, M. T.,
McLean, M. A., Sligar, S. G., Chen, C. S., Ha, T. et al. (2010). Measuring
mechanical tension across vinculin reveals regulation of focal adhesion dynamics.
Nature 466, 263-266.
Hotulainen, P. and Lappalainen, P. (2006). Stress fibers are generated by two distinct
actin assembly mechanisms in motile cells. J. Cell Biol. 173, 383-394.
Hu, S. H., Chen, J. X., Butler, J. P. and Wang, N. (2005). Prestress mediates force
propagation into the nucleus. Biochem. Biophys. Res. Commun. 329, 423-428.
Hwang, Y. and Barakat, A. I. (2012). Dynamics of mechanical signal transmission
through prestressed stress fibers. PLoS ONE 7, e35343.
Hyto ¨nen, V. P. and Vogel, V. (2008). How force might activate talin’s vinculin binding
sites: SMD reveals a structural mechanism. PLOS Comput. Biol. 4, e24.
Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W.,
Hess, H. F. and Waterman, C. M. (2010). Nanoscale architecture of integrin-based
cell adhesions. Nature 468, 580-584.
Katoh, K., Kano, Y. and Ookawara, S. (2007). Rho-kinase dependent organization of
stress fibers and focal adhesions in cultured fibroblasts. Genes Cells 12, 623-638.
Katoh, K., Kano, Y. and Noda, Y. (2011). Rho-associated kinase-dependent
contraction of stress fibres and the organization of focal adhesions. J. R. Soc.
Interface 8, 305-311.
Kumar, S., Maxwell, I. Z., Heisterkamp, A., Polte, T. R., Lele, T. P., Salanga, M.,
Mazur, E. and Ingber, D. E. (2006). Viscoelastic retraction of single living stress
fibers and its impact on cell shape, cytoskeletal organization, and extracellular matrix
mechanics. Biophys. J. 90, 3762-3773.
Lam, A. J., St-Pierre, F., Gong, Y., Marshall, J. D., Cranfill, P. J., Baird, M. A.,
McKeown, M. R., Wiedenmann, J., Davidson, M. W., Schnitzer, M. J. et al.
(2012). Improving FRET dynamic range with bright green and red fluorescent
proteins. Nat. Methods 9, 1005-1012.
le Duc, Q., Shi, Q. M., Blonk, I., Sonnenberg, A., Wang, N., Leckband, D. and de
Rooij, J. (2010). Vinculin potentiates E-cadherin mechanosensing and is recruited to
actin-anchored sites within adherens junctions in a myosin II-dependent manner.
J. Cell Biol. 189, 1107-1115.
Lele, T. P., Pendse, J., Kumar, S., Salanga, M., Karavitis, J. and Ingber, D. E.
(2006). Mechanical forces alter zyxin unbinding kinetics within focal adhesions of
living cells. J. Cell. Physiol. 207, 187-194.
Levental, K. R., Yu, H., Kass, L., Lakins, J. N., Egeblad, M., Erler, J. T., Fong, S. F.,
Csiszar, K., Giaccia, A., Weninger, W. et al. (2009). Matrix crosslinking forces
tumor progression by enhancing integrin signaling. Cell 139, 891-906.
MacKay, J. L., Keung, A. J. and Kumar, S. (2012). A genetic strategy for the dynamic
and graded control of cell mechanics, motility, and matrix remodeling. Biophys. J.
Moore, D. S. and McCabe, G. P. (2002). Introduction to the Practice of Statistics. New
York, NY: W. H. Freeman.
Na, S., Collin, O., Chowdhury, F., Tay, B., Ouyang, M. X., Wang, Y. X. and Wang,
N. (2008). Rapid signal transduction in living cells is a unique feature of
mechanotransduction. Proc. Natl. Acad. Sci. USA 105, 6626-6631.
Plotnikov, S. V., Pasapera, A. M., Sabass, B. and Waterman, C. M. (2012). Force
fluctuations within focal adhesions mediate ECM-rigidity sensing to guide directed
cell migration. Cell 151, 1513-1527.
Prager-Khoutorsky, M., Lichtenstein, A., Krishnan, R., Rajendran, K., Mayo, A.,
Kam, Z., Geiger, B. and Bershadsky, A. D. (2011). Fibroblast polarization is a
matrix-rigidity-dependent process controlled by focal adhesion mechanosensing. Nat.
Cell Biol. 13, 1457-1465.
Rauzi, M., Verant, P., Lecuit, T. and Lenne, P. F. (2008). Nature and anisotropy of
cortical forces orienting Drosophila tissue morphogenesis. Nat. Cell Biol. 10, 1401-
Ronchi, P., Terjung, S. and Pepperkok, R. (2012). At the cutting edge: applications
and perspectives of laser nanosurgery in cell biology. Biol. Chem. 393, 235-248.
Rossier, O. M., Gauthier, N., Biais, N., Vonnegut, W., Fardin, M. A., Avigan, P.,
Heller, E. R., Mathur, A., Ghassemi, S., Koeckert, M. S. et al. (2010). Force
generated by actomyosin contraction builds bridges between adhesive contacts.
EMBO J. 29, 1055-1068.
Tension distributions of stress fibers3029
Journal of Cell Science
Russell, R. J., Xia, S. L., Dickinson, R. B. and Lele, T. P. (2009). Sarcomere
mechanics in capillary endothelial cells. Biophys. J. 97, 1578-1585.
Sawada, Y., Tamada, M., Dubin-Thaler, B. J., Cherniavskaya, O., Sakai, R.,
Tanaka, S. and Sheetz, M. P. (2006). Force sensing by mechanical extension of the
Src family kinase substrate p130Cas. Cell 127, 1015-1026.
Sen, S., Dong, M. and Kumar, S. (2009). Isoform-specific contributions of alpha-
actinin to glioma cell mechanobiology. PLoS ONE 4, e8427.
Stricker, J., Sabass, B., Schwarz, U. S. and Gardel, M. L. (2010). Optimization of
traction force microscopy for micron-sized focal adhesions. J. Phys. Condens. Matter
Stricker, J., Aratyn-Schaus, Y., Oakes, P. W. and Gardel, M. L. (2011).
Spatiotemporal constraints on the force-dependent growth of focal adhesions.
Biophys. J. 100, 2883-2893.
Tan, J. L., Tien, J., Pirone, D. M., Gray, D. S., Bhadriraju, K. and Chen, C. S.
(2003). Cells lying on a bed of microneedles: an approach to isolate mechanical force.
Proc. Natl. Acad. Sci. USA 100, 1484-1489.
in stress fiber mechanics in living cells with laser nanosurgery. Biophys. J. 99, 2775-2783.
The ´ry, M. (2010). Micropatterning as a tool to decipher cell morphogenesis and
functions. J. Cell Sci. 123, 4201-4213.
Tinsley, R. B., Vesey, M. J., Barati, S., Rush, R. A. and Ferguson, I. A. (2004).
Improved non-viral transfection of glial and adult neural stem cell lines and of
primary astrocytes by combining agents with complementary modes of action.
J. Gene Med. 6, 1023-1032.
Tondon, A., Hsu, H. J. and Kaunas, R. (2012). Dependence of cyclic stretch-induced
stress fiber reorientation on stretch waveform. J. Biomech. 45, 728-735.
Totsukawa, G., Wu, Y., Sasaki, Y., Hartshorne, D. J., Yamakita, Y., Yamashiro,
S. and Matsumura, F. (2004). Distinct roles of MLCK and ROCK in the regulation
of membrane protrusions and focal adhesion dynamics during cell migration of
fibroblasts. J. Cell Biol. 164, 427-439.
Vogel, V. (2006). Mechanotransduction involving multimodular proteins: converting
force into biochemical signals. Annu. Rev. Biophys. Biomol. Struct. 35, 459-488.
Wang, N., Tytell, J. D. and Ingber, D. E. (2009). Mechanotransduction at a distance:
mechanically coupling the extracellular matrix with the nucleus. Nat. Rev. Mol. Cell
Biol. 10, 75-82.
Zamir, E., Katz, B. Z., Aota, S., Yamada, K. M., Geiger, B. and Kam, Z. (1999).
Molecular diversity of cell-matrix adhesions. J. Cell Sci. 112, 1655-1669.
Zhang, C., Yadava, P. and Hughes, J. (2004). Polyethylenimine strategies for plasmid
delivery to brain-derived cells. Methods 33, 144-150.
Zhong, W., Wu, M., Chang, C. W., Merrick, K. A., Merajver, S. D. and Mycek,
M. A. (2007). Picosecond-resolution fluorescence lifetime imaging microscopy: a
useful tool for sensing molecular interactions in vivo via FRET. Opt. Express 15,
Journal of Cell Science 126 (14)3030