Cell crawling mediates collective cell migration to close undamaged

Cell crawling mediates collective cell migration to
close undamaged epithelial gaps
Ester Anona,b, Xavier Serra-Picamalb, Pascal Hersena,c, Nils C. Gauthierc, Michael P. Sheetzc,d, Xavier Trepatb,e,1,
and Benoît Ladouxa,c,1
a
Laboratoire Matière et Systèmes Complexes, Université Paris Diderot and Unité Mixte de Recherche 7057 Centre National de la Recherche Scientifique,
F-75205 Paris Cedex 13, France; bInstitute for Bioengineering of Catalonia, University of Barcelona and Ciber Enfermedades Respiratorias, 08036 Barcelona,
Spain; cMechanobiology Institute, National University of Singapore, Singapore 117411; dDepartment of Biological Sciences, Columbia University, New York,
NY 11027; and eInstitució Catalana de Recerca i Estudis Avançats, 08010 Barcelona, Spain
Fundamental biological processes such as morphogenesis and
wound healing involve the closure of epithelial gaps. Epithelial
gap closure is commonly attributed either to the purse-string
contraction of an intercellular actomyosin cable or to active cell
migration, but the relative contribution of these two mechanisms
remains unknown. Here we present a model experiment to
systematically study epithelial closure in the absence of cell injury.
We developed a pillar stencil approach to create well-defined gaps
in terms of size and shape within an epithelial cell monolayer.
Upon pillar removal, cells actively respond to the newly accessible
free space by extending lamellipodia and migrating into the gap.
The decrease of gap area over time is strikingly linear and shows
two different regimes depending on the size of the gap. In large
gaps, closure is dominated by lamellipodium-mediated cell migration. By contrast, closure of gaps smaller than 20 μm was affected
by cell density and progressed independently of Rac, myosin light
chain kinase, and Rho kinase, suggesting a passive physical mechanism. By changing the shape of the gap, we observed that lowcurvature areas favored the appearance of lamellipodia, promoting faster closure. Altogether, our results reveal that the closure
of epithelial gaps in the absence of cell injury is governed by the
collective migration of cells through the activation of lamellipodium
protrusion.
|
epithelial cell migration microfabrication
Madin-Darby canine kidney cells
| wound model assay |
A
wide variety of processes in health and disease involve the
formation and closure of epithelial gaps. In embryos, naturally occurring gaps appear at different stages of development
as a consequence of morphogenetic movements (1). A paradigmatic example is the well-studied process of dorsal closure in
Drosophila, whereby epithelial sheets migrate over the amnioserosa cell layer to seal an eye-shaped opening (2). In adults,
gaps in epithelial barriers result from dynamic tissue homeostasis, as clearly illustrated by epithelial turnover in the intestine (3).
Moreover, epithelial gaps are commonly formed during trauma
and chronic inflammatory diseases in which the epithelium is
injured and often completely denuded. A rapid healing of these
gaps is crucial to restore a functional epithelium and to prevent
further damage.
Because of the importance of the maintenance of epithelial
functions and homeostasis, many efforts have been devoted to
study gap closure, and two distinct mechanisms have emerged
(4–7). One mechanism is based on the assembly and contraction
of a multicellular acto-myosin belt lining the gap (known as
purse-string) (8), which is controlled by RhoA and its direct
regulators Rho kinase (ROCK) and myosin light chain kinase
(MLCK) (9). With a purse-string closure, the driving force is thus
provided by the contraction of the actomyosin cable around the
wound (10, 11). The second mechanism is based on cell migration mediated by lamellipodial protrusion, which is mostly regulated by Rac1 GTPase (9). In such cases, the mechanics of the
process seem less clear because the driving mechanism could be
www.pnas.org/cgi/doi/10.1073/pnas.1117814109
the pressure exerted by surrounding cells, the pulling force from
leader cells, or both (6, 7, 12).
The intricacy of the process and its regulation by the complex
family of Rho-GTPases has promoted the appearance of many
studies providing opposing roles for the different regulators (5,
13, 14). Cell–cell junctions play also an important role in the
process, because it has been suggested that the purse-string is
anchored at adherens junctions (15) or at tight junctions (16).
Ample evidence now supports each of these two mechanisms,
but their relative contribution to gap closure remains uncertain.
The controversy is also fostered by the variability in the experimental conditions used to create the gaps. The most commonly used methods to create gaps within cell monolayers are
either the classic scratch wound assay, in which a strip of cells is
mechanically removed with a pipette tip or a razor blade (17), or
laser ablation, in which single cells are destroyed by a laser pulse
(5). Both techniques are difficult to standardize because the final
size and shape of the gap depend either on the shape and velocity of the scratching utensil or on the power and focal plane of
the laser. Moreover, damage of cells during the process of wound
production releases a complex and poorly characterized mixture
of signaling molecules, death factors, and cell debris that influence the mechanisms of closure (18, 19).
How the actomyosin cable and/or lamellipodial protrusions are
activated during epithelial gap closure is unclear but may involve
secretion of soluble factors and/or mechanical tension. Up to now,
most of the literature is based on the study of wound closure,
whereby wounds are created by an aggressive method that releases
a complex and unknown mixture of debris and death factors (20,
21). The central question of epithelial gap closure is thus still
controversial and has not been addressed using well-defined
physical and geometrical conditions. To overcome these limitations, we present a unique strategy to induce well-defined gaps
within an epithelial monolayer and monitor the dynamics of epithelial gap closure in the absence of cell injury.
Results and Discussion
Dynamics of Epithelial Closure After Pillar Removal. By using a stencil
of poly-dimethylsiloxane (PDMS) micropillars (Fig. 1), we could
obtain many gaps of well-defined size and shape (Fig. 1 G–I and
Table S1). The size and shape of the pillars were varied to obtain circular pillars of different diameters, ranging from 15 to
150 μm, and squared and ellipsoidal pillars of two different sizes
Author contributions: E.A., M.P.S., X.T., and B.L. designed research; E.A., X.S.-P., X.T., and
B.L. performed research; E.A., X.S.-P., P.H., N.C.G., M.P.S., X.T., and B.L. analyzed data; and
E.A., X.T., and B.L. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission. W.M.B. is a guest editor invited by the Editorial
Board.
Freely available online through the PNAS open access option.
1
To whom correspondence may be addressed. E-mail: [email protected] or benoit.ladoux@
univ-paris-diderot.fr.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
1073/pnas.1117814109/-/DCSupplemental.
PNAS | July 3, 2012 | vol. 109 | no. 27 | 10891–10896
CELL BIOLOGY
Edited by William M. Bement, University of Wisconsin, Madison, WI, and accepted by the Editorial Board May 11, 2012 (received for review October 29, 2011)
2A
, which is the ratio of the
Rp
area A over the contour length of the interface p normalized by
half the instantaneous radius R. For a very rough interface, α ∼ 0,
whereas α ∼ 1 for a perfectly circular hole. We indeed observed
a decrease of this parameter α with time from 1 (at t = 0) down
to approximately 0.6 as the boundary became irregular owing to
the emergence of lamellipodium surrounding the gap (Fig. 2B).
According to previous studies, it was suggested that purse-string
contraction repaired small epithelial wounds (4, 5), whereas larger
wounds induced also cell crawling with formation of lamellipodia (6,
7, 23). Surprisingly, the presence of lamellipodia was observed for
all gap sizes tested from 15 up to 150 μm (Movies S1 and S2). The
formation of lamellipodia started shortly after the release of the
PDMS pillar (during the first 10 min) and were present until there
was no more available space, at which point opposing or contiguous
lamellipodia contacted and fused. In small gaps (15–30 μm), all cells
contacting the gap extended lamellipodia. For larger gaps, the
number of cells at the gap border increased, and not all of these cells
extended lamellipodia (Movie S2). Despite the presence of lamellipodia, the closure of gap was roughly isotropic, implying there was
not the so-called fingering activity (24). We then sought to analyze
the time evolution of the gap area, A(t), during epithelial closure. In
all conditions tested, the decrease of the area with time was strikingly linear with time down to a complete closure (Fig. 2C and Fig.
S3A). The trend in the decrease of the gap area as function of time
was similar for the different initial gap diameters, except for the
smallest ones (for diameters of 15 and 20 μm). As shown on Fig. 2D,
the closure time varied linearly with the size of the gap above a gap
diameter of 20 μm. By analyzing the slope of A(t), we computed the
initial radial velocity (which represents the velocity at the onset of
closure) as a function of the gap size and was found to be roughly
constant (0.3 μm/min) for areas up to 750 μm2 and then slightly
decreased for larger gaps (Fig. 2E). Consistently, the advancement
velocities of the protruding lamellipodia were found to be approximately 0.3 μm/min during the initial stage of lamellipodia formation (computed from the kymographs like Fig. S3B). Similarly, the
cell body advancement displayed the same velocity at the onset of
gap closure. Altogether, these results showed that the lamellipodium extension governed the kinetics of the mechanism of gap
closure and suggested the possibility of a size-dependent mechanism
in the dynamics of gap closure. As a comparison, we observed that
the dynamics of damage-associated gaps exhibited broader distributions due to variable initial conditions and followed exponential decay laws as a function of time (Fig. S2C). This indicates that
the presence of damaged cells or debris strongly altered the dynamics of epithelial gap closure. Interestingly, the closure dynamics
of these “wounds” are consistent with reported data on embryonic
wound healing and adult epithelial wound closure (15, 25).
To rule out an effect of differential extracellular matrix assembly that could provide directional cues to govern cell migration, we checked the status of the extracellular matrix (ECM)
organization and secretion during our experiments (Fig. S4).
Fibronectin was deposited on the substrate beneath the cells and
the pillars. After removing the pillar, fibronectin was not affected, and cells migrated over this fibronectin substrate (Fig. S4
A–D). Moreover, the staining of the overall fibronectin due to
the coating and cell secretion did not exhibit any specific pattern
(Fig. S4E). Laminin was absent in the gap area but present in the
cells as a synthetized protein, not structured in basal lamina yet
(Fig. S4G). Throughout all of the experiments, because there
was a fibronectin-based ECM roughly distributed everywhere onto
the substrate, our observations could not be attributed to ECM
remodeling underneath the monolayer as a driving mechanism.
We then analyzed the influence of cell culture density on the
progression of closure. MDCK cells are epithelial cells that can
undergo epithelial-to-mesenchymal transition (26). One could
argue, therefore, that before pillar removal cells are already in
a promigratory mesenchymal-like state, thus the protrusion of
lamellipodia and active cell migration observed would not be
a de novo response triggered by the sudden availability of free
by measuring the shape factor, α ¼
Fig. 1. Experimental design for gap patterning. (A–C) Schematic view of
the approach used for the experimental model: (A) PDMS pillar surrounded
by cells, (B) gap created upon pillar removal, (C) gap closed. (D–F) Phasecontrast micrographs of the different stages of the experimental model: (D)
PDMS pillar surrounded by cells, (E) gap created upon pillar removal (note
that the cells bordering the gap are intact), (F) gap closed by cells. (G) Array
of gaps created by using a stencil with numerous PDMS pillars. (H and I)
Microfabricated squared (H) and ellipsoidal (I) PDMS pillars with MDCK cells
cultured within the pillar stencil. (J) Assessment of cell viability by FITCdextran uptake. Note that none of the gap lining cells shows positive for
FITC-dextran. (K) Assessment of cell viability by propidium iodide internalization. (Scale bars, 10 μm in D–F, 20 μm in G–K.)
(Fig. 1 D, H, and I). The PDMS pillars are coated with a nonadhesive polymer to prevent cell attachment and are surrounded
by cells. In such a way, there are no specific adhesions between the
pillars and the bordering cells, the pillar being a mere blocking
object (Fig. S1 A and B). Madin-Darby canine kidney (MDCK)
cells are cultured in between the pillars for 15 ± 3 h. Upon careful
removal of the PDMS pillar, a gap is created within the monolayer, without tearing the bordering cells (Fig. 1 E, G, J, and K).
Additionally, we performed scratch-induced gaps either by the
removal of pillars on which cells were able to adhere (resulting in
“ripped” gaps) or by pushing pillars against a preformed epithelial
monolayer to kill the cells underneath (resulting in “crushed”
gaps) (more details in SI Materials and Methods), to compare our
model experiment with more classic scratch assays. We assessed
cell damage by using FITC-dextran (22) as well as propidium
iodide internalization by damaged cells. Both tests ascertained
the absence of damage in our pillar removal assay, whereas they
revealed cell damage in the case of ripped and crushed gaps and
in the classic scratch assay (Fig. 1 J and K and Fig. S2 A and B).
Thus, our methodology prevents the damage of cells surrounding
the pillar after removal, as opposed to what is observed during
classic scratch assays.
We first analyzed the dynamics of epithelial gap closure after
removal of circular pillars of different diameters. Video microscopy experiments upon pillar stencil removal revealed that
cells lining the gap extended lamellipodia throughout the process
of closure (Fig. 2A and Movie S1). The borders of the gap
roughened considerably after the removal of the pillar, indicating
the extension of cellular protrusions into the available free space.
We quantitatively analyzed the variations of the contour length
10892 | www.pnas.org/cgi/doi/10.1073/pnas.1117814109
Anon et al.
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Fig. 2. Dynamics of gap closure. (A) Snapshots of time-lapse video microscopy during gap closure. t = 0 is acquired right after removal of the pillar stencil. (Scale
bars, 20 μm.) (B) Evolution of roughness α at the cell–gap interface. Experiments performed with two pillar sizes, 30 and 60 μm in diameter. (C) Gap area
decrease with time, for different pillar sizes used for creating the gaps, ranging from 15 to 60 μm in diameter. Green line corresponds to gaps created with
pillars of 15 μm in diameter, orange 20, purple 30, black 40, red 50, and blue 60. Data for 80- and 150-μm pillar diameter are shown in Fig. S3. Data are reported
as mean and SD of eight gaps analyzed per each size, resulting from four independent experiments. (D) Closure time as a function of the initial gap area of
circular gaps. (E) Initial radial velocity depending on the initial gap size. (F) Effect of cell density on closure time, examined for three different gap diameters.
Cell density is indicated in the x axis, calculated from Table S2. Each graph shows the experiments from a different pillar diameter. (G) Effect of substrate
stiffness on the closure time of two different gap diameters, 20 and 50 μm. The x axis indicates the PDMS ratio used to attain different stiffnesses. No closure
stands for gaps that were not closed after 300 min.
Force Generation on Stiff Substrates Induces Epithelial Gap Closure
by Lamellipodium Activation. To verify that the activation of
a range of rigidities (Fig. S4F). Experiments performed on
PDMS substrates of Young’s modulus equivalent to glass (at
1:10, Young’s modulus is 0.5–2 MPa) showed no differences
either in dynamics or in closure time with respect to control
experiments. As the stiffness decreased, we observed a drastic
decrease of the migration speed of the cells into the gap (a 2.7fold increase in closure time of 20-μm gaps and 1.9-fold in 50-μm
gaps) (Fig. 2G). Furthermore, softer substrates (1:40 and 1:60)
prevented the final closure of the gap even after 300 min, as well
as the formation of lamellipodial protrusions (Movie S3).
Therefore, stiff substrates were needed to generate the activation
of the lamellipodium around the gap and stabilize it. Because
cells are probing substrate stiffness by applying contractile forces
(27, 29), it seems that the pulling force induced by leading cells
that exhibited lamellipodial protrusions is a key player in our
experimental model of epithelial closure.
lamellipodium formation could be the driving force of cell migration into the gap, we tested the effects of the substrate rigidity
on the dynamics of epithelial gap closure. Substrate stiffness is
known to activate cell migration, to increase cell spreading and
traction forces (27), and to govern lamellipodium dynamics (28).
We observed MDCK cell migration after pillar removal on
a PDMS substrate whose stiffness was tuned by changing the
percentage of the reticulating agent (to 1:25, 1:40, and 1:60
PDMS cross-linker:base ratio; SI Materials and Methods), and
we verified that the ECM coating was not affected within such
Universal Mechanism Drives Closure of Small Gaps. Because the
movement of epithelial cell sheets during wound closure could
exhibit a purse-string mechanism, lamellipodium-based crawling,
or both mechanisms simultaneously or at different stages (8, 23),
we explored their relative contribution in our model experiment
of epithelial gap closure. We tested the role of lamellipodial
protrusion by inhibiting Rac1 and the role of the contractile
machinery by inhibiting ROCK, MLCK, and myosin phosphorylation (Fig. S6). For each of these treatments and cell lines, we
space. To rule out this possibility, we tested different cell packing
densities, ranging from highly spread and flattened cells to the
maximal density of cells within the culture, always after confluence was reached (Fig. S5 and Table S2). For large gaps (30- and
60-μm diameters), cell density had no impact either on the closure time or on protrusive activity (Fig. 2F). In addition, this
result confirmed that the closure mechanism was not triggered by
a possible release of the internal pressure within the epithelial
cell sheet after the removal of the pillar but instead by the
lamellipodium extension. However, for the smallest gaps, there
was a decrease in the closure time as packing density increased,
further suggesting that distinct mechanisms govern the closure of
small vs. large gaps.
Anon et al.
PNAS | July 3, 2012 | vol. 109 | no. 27 | 10893
CELL BIOLOGY
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measured the closure time as a function of the initial gap size.
First, we observed that the closure time for small gaps (≤20 μm)
was insensitive to all of the pharmacological treatments mentioned above for the inhibition of Rac or Rho pathways (Fig.
3A), further suggesting that a mechanism independent of the
proposed regulators could induce the gap closure in such cases.
According to the experimental results, two regimes emerged,
depending on gap size. For small gaps (≤20 μm), we observed
different dynamics and dependency of the closure time as
a function of the gap size compared with the ones observed for
larger gaps (Figs. 2C and 3A). This strikingly universal behavior
is suggestive of a mechanism of purely physical origin. One such
mechanism could be cell spreading based on an unspecific mechanical balance between cell-substrate adhesion and cortical
tension (30). This mechanism has been shown to produce a linear dependence of spreading area with time. Moreover, it is
consistent with a decrease of closure time with higher cell density
because denser cells are more columnar and thus offer larger
lateral area for cell spreading (Fig. 2F and Fig. S5).
A
Closure time (min)
Cell Crawling Drives the Closure of Large Gaps. By contrast, the
closure time of large gaps (>20 μm) was not universal. Surprisingly, inhibition of either MLCK or ROCK had no significant
effect in gap closure progression (Fig. 3A and Movies S4 and S5).
RhoA has been described as an activator of myosin contraction
required for purse-string closure, which is in turn regulated by
MLCK and ROCK (9). Our findings thus suggest that the closure
of large gaps is not driven by purse-string contraction. To ascertain this possibility in our closure model, we investigated actomyosin distribution at the gap edge. The presence per se of
PDMS pillars for gap patterning did not trigger actin accumulation at the pillar periphery (Fig. S7 A and B). However, phalloidin staining immediately after pillar removal showed that actin
accumulated in a continuous supracellular cable-like structure at
the margins of the gap (Fig. S7 C and D). This surrounding actin
C
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Fig. 3. Mechanism of gap closure: effect of inhibitors and actomyosin distribution. (A) Closure times of the different gap sizes in control conditions
and subjected to drug treatments of the regulators (MLCK, ROCK, and Rac1
inhibition, for six gap sizes). Data points represent means, and error bars are
SEs of seven analyzed gaps. (B) Z-stack projection and x–z and y–z orthogonal projections (sections from the yellow lines), after 30 min of closure
progression Note that there is actin clustering at some cell edges and
lamellipodia. (Scale bars, 25 μm in the z-projection, 5 μm in the orthogonal
projections.) (C) Actin and phospho-MLC accumulate at the gap margin at t =
0 min but do not progress as a continuous ring as closure proceeds. (Scale
bars, 20 μm.) (D) Fold increase in the closure time of Rac-inhibited cells with
respect to control conditions for the six different gap sizes.
10894 | www.pnas.org/cgi/doi/10.1073/pnas.1117814109
cable was then disrupted as closure proceeded: the discontinuity
of the actin cable was concomitant to the formation of cell
protrusions, such as the extension of multiple lamellipodia into
the gaps (Fig. 3 B and C). Moreover, according to confocal
images in the x–z and y–z planes, it appeared that areas of actin
accumulation localized at the lateral surface of cuboidal cells,
whereas lamellipodial extension induced a flattening of the
monolayer with a more diffuse and homogeneous actin distribution (Fig. S7 E and F). Staining of phospho-MLC as a marker
for contractile myosin showed that active myosin colocalized
with the actin cable immediately after pillar removal. However,
as closure progressed, myosin in the cable accumulated only at
the margins of major actin clustering (Fig. 3C). Previous results
suggested that apoptotic cells release signals that favor the assembly of a continuous actomyosin cable all around the dying
cell to promote the extrusion of the cell from the monolayer (31).
Our findings showed that only an incomplete acto-myosin ring
could form in the absence of cell injury. This suggests that death
factors are required to develop a functional multicellular actomyosin cable.
In contrast to the inhibition of RhoA, the inhibition of Rac1
drastically slowed down the closure process of large gaps. Rac1
inhibition precluded the extension of lamellipodia and maintained a strong circularity of the gaps throughout closure (Fig. 2B
and Movie S6). The larger the gaps, the more affected the closure was by Rac1 inhibition (Fig. 3D). Interestingly, Rac inhibition did not have such a slowing-down effect in damageassociated gaps, which progressed similarly to noninhibited
damaged gaps (Fig. S2D) Taken together, our findings demonstrate a dominant role of cell crawling over purse-string closure
in the absence of cell damage. This observation supports a
recently proposed theoretical mechanical model for wound closure, in which crawling cells can close wounds without pursestring signaling, only because of their directed mechanical activity (32).
Influence of the Geometry of the Gaps. Because epithelial gap
closure occurs in various geometrical conditions in vitro as well
as in vivo, we studied the influence of curvature and shape of the
gap on the closure process. We fabricated squared and ellipsoidal-like pillars (hereafter referred as ellipsoidal pillars) of two
different sizes (Fig. 4A). Cells distributed randomly along the
gap perimeter, with no preferential alignment of cells in areas of
different curvature (Fig. S1 F–K). Live-cell microscopy showed
that, regardless of the shape of the gaps, cells extended lamellipodia throughout closure (Fig. 4C and Movies S7 and S8) and
that these lamellipodia were preferentially protruded along the
edges with the lowest curvature. We then analyzed the closure
time of squared and ellipsoidal gaps relative to circular ones.
Except in the case of the smallest square analyzed, gaps of ellipsoidal and squared shape closed systematically faster than
circular ones (Fig. 4B). This faster response might be due to the
enhancement of lamellipodial activity in regions of low curvature. A physical model had previously reported that epithelial
cells can sense and respond to different global geometric conditions by detecting the curvature of the epithelial edge at a
multicellular level (33).
Much as in the case of circular gap experiments, actin and
phospho-myosin accumulated preferentially at areas in which
lamellipodia did not protrude, thus a supracellular cable was not
continuous (Fig. 4 D–F). Hence, these results indicate that the
behavior observed in the closure of circular gaps applies also to
different gap geometries. This result is in good agreement with
previous studies showing that large wounds (i.e., lower curvature) are preferentially Rac-dependent, whereas small ones (i.e.,
larger curvature) exhibit a purse-string mechanism (5, 6).
Collective Cell Movements by Cell–Cell Junctions and Myosin Action
Provide Directional Clues. To further characterize cell rearrange-
ments during gap closure, we analyzed the cells’ shape and dynamics along the process. We tracked the trajectories of cells
Anon et al.
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Fig. 4. Effect of geometry on gap closure. (A) Squared and ellipsoidal PDMS
pillars surrounded by cells. (B) Comparison of the closure time of squared
and ellipsoidal gaps with respect to circular gaps. (C) Sequence of phasecontrast micrographs showing the progression of the closure of a squared
(Top) and ellipsoidal (Bottom) gap. (D and E) Actin (Top) and phospho-MLC
(Middle) distribution in squared (D) and ellipsoidal (E) gaps for 2 different
time points. (Bottom) Merged images. (F) Epifluorescence micrographs of
actin distribution in the closure of ellipsoidal gaps 30 min after pillar removal. (Scale bars: 20 μm.)
during closure (Fig. S8 A and B). The first row of cells experienced directed motion toward the center of the gap and moved
98% ± 20% of the gap initial radius (Fig. S8 A, B, and H). Cells
behind the leading edge showed progressively smaller and less
persistent displacements (55% for the second row, 16% for outer
cells). This result indicates that gap closure was mainly due to an
active and directed process governed by cells at the leading edge
and triggered by the presence of the free space, as previously
described in the context of collective cell migration (7, 34).
Cells at the gap edge elongated along the direction of migration as closure progressed (Fig. S8G) and acquired a wedge-like
morphology (Fig. 1F), further confirmed by the elongation of cell
nuclei as an indicator of the cell polarization (Fig. S8J). At the
closure time point, cells typically formed a rosette-like structure
Anon et al.
(Fig. 1F and Fig. S4E) that would be later dissolved through
epithelial remodeling. Interestingly, this rosette-like structure
has been observed in various situations related to the closure of
gaps (8), apoptotic cell extrusion (31), cell delamination (35),
embryonic healing (36), and in vitro wound healing (5).
To further understand the role of cell–cell communications in
the gap closure process, we used an α-catenin knock-down
MDCK cell line. As shown by Benjamin et al. (37), α-catenin
knock-down MDCK cells display increased membrane dynamics
together with higher migration rates but exhibit a lack of cadherin-mediated cell–cell adhesion. These cells, given that they do
not form proper adherens junctions, could not support the formation of continuous multicellular cable surrounding the gap
(15, 25) as well as a collective behavior mediated by cell–cell
interactions. In our experimental model, they migrated independently of their position with respect to the gap edge, in an
uncoordinated manner toward the gap center (Fig. S8 C and D
and Movie S9). Thus, they displaced greater distances than WT
MDCK cells owing to their lack of coordination (Fig. S8H). Gaps
were still closed at a rate comparable to the WT MDCK cells
(Fig. S8I). These findings demonstrate that gap closure can be
accomplished without adherens junctions.
Although the absence of a continuous supracellular acto-myosin ring did not affect the closure kinetics, direct inhibition of
myosin phosphorylation by blebbistatin caused a 1.7-fold increase in the closure time of large gaps (namely of 50 μm in
diameter) (Fig. S8I). This observation suggests that myosin may
contribute to gap closure through a mechanism that is independent of purse-string contraction. Cells treated with blebbistatin extended very broad lamellipodia with considerable
ruffling activity (Movie S10). Compared with controls, cells
moved longer distances, but their paths were not directed toward
the center of the gap (Fig. S8 E and F). Moreover, the displacement magnitude was greater (approximately 150% displacement of the initial radius) and independent of the distance
from the gap edge (Fig. S8H). Thus, the closure under blebbistatin treatment was achieved in an uncoordinated manner,
resulting in a delay in the time of closure (Fig. S8I). Myosin IIA
silencing or inhibition has previously been shown to cause increased membrane ruffling and migration speed in numerous cell
types (38). Our findings show that this phenotype is not restricted
to the single-cell level and suggest that the role of myosin IIA is
not to drive collective cell motion but to guide it.
Conclusions
We have presented a unique approach to study gap closure in
uninjured epithelia under well-defined experimental conditions.
This provides a model for naturally occurring gaps in development, avoiding possible effects of cell death in gap closure. Such
model experiments are also useful to discriminate between the
different mechanisms proposed for epithelial gap closure (5, 6,
23). By using a microfabricated stencil with an array of pillars, gaps
of precise size and shape can be patterned in parallel in an epithelial cell culture. Upon pillar removal, cells actively respond to
the free space by extending lamellipodia and crawling into the gap.
Interestingly, small gaps (≤20 μm) show no response to the
inhibition of myosin filament assembly or myosin contraction.
Moreover, closure of such small areas is dependent on the cell
density of the epithelium, because small gaps close faster in
highly packed cultures. This evidence indicates that small gaps
are closed by unspecific cell spreading. For gaps larger than
20 μm, the closure process is not altered when regulators of the
purse-string contraction are inhibited, whereas disturbance of
lamellipodial extension causes a drastic delay in closure. In addition, the intercellular actomyosin belt is lost during progression
of closure, thus pointing out a pivotal role of cell crawling in the
closure response.
To date, the cell crawling-mediated response has been mostly
referred to wounds that could be considered infinitely large (6, 7,
39). We show here that even small gaps (of dozens of micrometers in diameter) are closed by means of lamellipodial
PNAS | July 3, 2012 | vol. 109 | no. 27 | 10895
CELL BIOLOGY
B
Closure time (min)
A
extension. The mere presence of free space has been proposed as
the triggering mechanism for this response (7, 34).
In classic scratch-wound experiments, the purse-string mechanism has been found responsible for the closure of the wound.
Purse-string has also been proposed for accounting for the extrusion of apoptotic cells (31), a process clearly related to death
signaling, whereby the actomyosin cable formation is triggered
through a caspase-mediated pathway (40). Thus, evidence suggests that cell damage inflicted during the process of wound
production is promoting the purse-string mechanism by affecting
the neighboring cells. In concordance with this hypothesis, we
show here that in the absence of cell damage, purse-string is not
the dominant mechanism, but the closure is mediated by a
lamellipodial-driven crawling mechanism. In our model, the role
of a supracellular actin belt is related to the coordination of the
migrating cells toward the center of the gap, ensuring the proper
directionality and persistence of their migration.
Interestingly, our results suggest that cells extending lamellipodia act as leader cells to close the gap. Indeed, it is known that
protrusive lamellipodia are related to the mechanical probing of
the substrate. On soft substrates, either we did not observe the
formation of lamellipodia or they appear smaller and shorter in
time. As a result, cells could not close the gap. The closure
mechanism is thus associated with stabilization of protruding
lamellipodia that help to generate stronger forces at the leading
edge (28, 41). Finally, we show that squared and ellipsoidal gaps
are closed faster than circular ones. Low curvature areas promote
1. Martin P, Parkhurst SM (2004) Parallels between tissue repair and embryo morphogenesis. Development 131:3021–3034.
2. Jacinto A, Martinez-Arias A, Martin P (2001) Mechanisms of epithelial fusion and
repair. Nat Cell Biol 3:E117–E123.
3. Watson AJ, et al. (2005) Epithelial barrier function in vivo is sustained despite gaps in
epithelial layers. Gastroenterology 129:902–912.
4. Bement WM, Mandato CA, Kirsch MN (1999) Wound-induced assembly and closure of
an actomyosin purse string in Xenopus oocytes. Curr Biol 9:579–587.
5. Tamada M, Perez TD, Nelson WJ, Sheetz MP (2007) Two distinct modes of myosin
assembly and dynamics during epithelial wound closure. J Cell Biol 176:27–33.
6. Fenteany G, Janmey PA, Stossel TP (2000) Signaling pathways and cell mechanics involved in wound closure by epithelial cell sheets. Curr Biol 10:831–838.
7. Poujade M, et al. (2007) Collective migration of an epithelial monolayer in response to
a model wound. Proc Natl Acad Sci USA 104:15988–15993.
8. Bement WM, Forscher P, Mooseker MS (1993) A novel cytoskeletal structure involved
in purse string wound closure and cell polarity maintenance. J Cell Biol 121:565–578.
9. Nobes CD, Hall A (1995) Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81:53–62.
10. Kim J-H, Dooling LJ, Asthagiri AR (2010) Intercellular mechanotransduction during
multicellular morphodynamics. J R Soc Interface 7(Suppl 3):S341–S350.
11. Salbreux G, Prost J, Joanny JF (2009) Hydrodynamics of cellular cortical flows and the
formation of contractile rings. Phys Rev Lett 103:058102.
12. Trepat X, et al. (2009) Physical forces during collective cell migration. Nat Phys 5:
426–430.
13. Desai LP, Aryal AM, Ceacareanu B, Hassid A, Waters CM (2004) RhoA and Rac1 are
both required for efficient wound closure of airway epithelial cells. Am J Physiol Lung
Cell Mol Physiol 287:L1134–L1144.
14. Russo JM, et al. (2005) Distinct temporal-spatial roles for rho kinase and myosin light
chain kinase in epithelial purse-string wound closure. Gastroenterology 128:
987–1001.
15. Danjo Y, Gipson IK (1998) Actin ‘purse string’ filaments are anchored by E-cadherinmediated adherens junctions at the leading edge of the epithelial wound, providing
coordinated cell movement. J Cell Sci 111:3323–3332.
16. Florian P, Schöneberg T, Schulzke JD, Fromm M, Gitter AH (2002) Single-cell epithelial
defects close rapidly by an actinomyosin purse string mechanism with functional tight
junctions. J Physiol 545:485–499.
17. Todaro GJ, Lazar GK, Green H (1965) The initiation of cell division in a contact-inhibited mammalian cell line. J Cell Physiol 66:325–333.
18. Klepeis VE, Cornell-Bell A, Trinkaus-Randall V (2001) Growth factors but not gap
junctions play a role in injury-induced Ca2+ waves in epithelial cells. J Cell Sci 114:
4185–4195.
19. Block ER, Matela AR, SundarRaj N, Iszkula ER, Klarlund JK (2004) Wounding induces
motility in sheets of corneal epithelial cells through loss of spatial constraints: Role of
heparin-binding epidermal growth factor-like growth factor signaling. J Biol Chem
279:24307–24312.
20. McNeil PL (2002) Repairing a torn cell surface: Make way, lysosomes to the rescue. J
Cell Sci 115:873–879.
21. Woolley K, Martin P (2000) Conserved mechanisms of repair: From damaged single
cells to wounds in multicellular tissues. Bioessays 22:911–919.
10896 | www.pnas.org/cgi/doi/10.1073/pnas.1117814109
the protrusion of broad lamellipodia, but a continuous pursestring is not formed in square nor ellipsoidal gaps. Therefore,
closure of noncircular epithelial gaps also seems to be primarily
driven by lamellipodial-mediated cell crawling.
Materials and Methods
PDMS micropillars of different sizes and shapes were fabricated as previously described (41). Micropillar stencils were stuck to fibronectin-coated
glass-bottom dishes. MDCK cells were plated and allowed to grow between the pillars until confluence. Gap closure was monitored with livecell microscopy upon peeling off of the stencil, and image analysis was
performed in ImageJ. Further details on the fabrication of substrates,
inhibitors treatments, and immunofluorescence microscopy are found in SI
Materials and Methods.
ACKNOWLEDGMENTS. We thank J. Le Digabel, Y. Toyama, F. Gallet, J.-M. Di
Meglio, members of the Integrative Cell and Tissue Dynamics Laboratory at
IBEC, and members of the Mechanobiology Institute (National University of
Singapore) for fruitful discussions; A. Richert for cell culture protocols; W. J.
Nelson (Stanford University) for kindly providing the α-catenin knockdown
MDCK cell line; and the microfabrication core of MBI. Financial support was
received from the Association pour la Recherche sur le Cancer, the Association Française Contre la Myopathie, the Agence Nationale de la Recherche
[Programme Blanc 2010 (MECANOCAD)], Grant BFU2009-07595 from the
Spanish Ministry for Science and Innovation, Grant Agreement 242993 from
the European Research Council, and MBI. The research was conducted in
the scope of the International Associated Laboratory Cell Adhesion FranceSingapore. E.A. receives financial support from the Fondation pour la
Recherche Médicale.
22. Brock J, Midwinter K, Lewis J, Martin P (1996) Healing of incisional wounds in the
embryonic chick wing bud: Characterization of the actin purse-string and demonstration of a requirement for Rho activation. J Cell Biol 135:1097–1107.
23. Garcia-Fernandez B, Campos I, Geiger J, Santos AC, Jacinto A (2009) Epithelial resealing. Int J Dev Biol 53:1549–1556.
24. Reffay M, et al. (2011) Orientation and polarity in collectively migrating cell structures: Statics and dynamics. Biophys J 100:2566–2575.
25. Abreu-Blanco MT, Verboon JM, Parkhurst SM (2011) Cell wound repair in Drosophila
occurs through three distinct phases of membrane and cytoskeletal remodeling. J Cell
Biol 193:455–464.
26. Nicolás FJ, Lehmann K, Warne PH, Hill CS, Downward J (2003) Epithelial to mesenchymal transition in Madin-Darby canine kidney cells is accompanied by down-regulation of Smad3 expression, leading to resistance to transforming growth factorbeta-induced growth arrest. J Biol Chem 278:3251–3256.
27. Pelham RJ, Jr., Wang YL (1998) Cell locomotion and focal adhesions are regulated by
the mechanical properties of the substrate. Biol Bull 194:348–349, discussion 349–350.
28. Giannone G, et al. (2004) Periodic lamellipodial contractions correlate with rearward
actin waves. Cell 116:431–443.
29. Saez A, et al. (2010) Traction forces exerted by epithelial cell sheets. J Phys Condens
Matter 22:194119.
30. Cuvelier D, et al. (2007) The universal dynamics of cell spreading. Curr Biol 17:694–699.
31. Rosenblatt J, Raff MC, Cramer LP (2001) An epithelial cell destined for apoptosis
signals its neighbors to extrude it by an actin- and myosin-dependent mechanism.
Curr Biol 11:1847–1857.
32. Lee P, Wolgemuth CW (2011) Crawling cells can close wounds without purse strings or
signaling. PLoS Comput Biol 7:e1002007.
33. Mark S, et al. (2010) Physical model of the dynamic instability in an expanding cell
culture. Biophys J 98:361–370.
34. Nikolic DL, Boettiger AN, Bar-Sagi D, Carbeck JD, Shvartsman SY (2006) Role of
boundary conditions in an experimental model of epithelial wound healing. Am J
Physiol Cell Physiol 291:C68–C75.
35. Muliyil S, Krishnakumar P, Narasimha M (2011) Spatial, temporal and molecular hierarchies in the link between death, delamination and dorsal closure. Development
138:3043–3054.
36. Meghana C, et al. (2011) Integrin adhesion drives the emergent polarization of active
cytoskeletal stresses to pattern cell delamination. Proc Natl Acad Sci USA 108:
9107–9112.
37. Benjamin JM, et al. (2010) AlphaE-catenin regulates actin dynamics independently of
cadherin-mediated cell-cell adhesion. J Cell Biol 189:339–352.
38. Even-Ram S, et al. (2007) Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat Cell Biol 9:299–309.
39. Omelchenko T, Vasiliev JM, Gelfand IM, Feder HH, Bonder EM (2003) Rho-dependent
formation of epithelial “leader” cells during wound healing. Proc Natl Acad Sci USA
100:10788–10793.
40. Andrade D, Rosenblatt J (2011) Apoptotic regulation of epithelial cellular extrusion.
Apoptosis 16:491–501.
41. du Roure O, et al. (2005) Force mapping in epithelial cell migration. Proc Natl Acad Sci
USA 102:2390–2395.
Anon et al.
Supporting Information
Anon et al. 10.1073/pnas.1117814109
SI Materials and Methods
Fabrication of Poly-Dimethylsiloxane (PDMS) Pillar Stencil. To pat-
tern gaps in the cell culture, we took advantage of a stencil
containing an array of microfabricated pillars. The microfabrication of the stencils was based on a previously described
method (1). Briefly, motifs printed on a negative mask were
transferred to a silicon wafer by photolithography, followed by
a deep reactive ion etching process to obtain holes down to the
desired depth. This silicon master was silanized to facilitate the
release of the elastomer and allow reusing the master. Then,
PDMS (Sylgard 184; Dow-Corning) was poured over the silicon
template and cured at 65 °C for approximately 12 h. The stamp
of PDMS pillars can be then peeled off from the master.
Gap Patterning. PDMS pillar microstamps were stuck to fibronectin-coated glass-bottom Petri dishes by mild oxygen plasma
treatment. Coating was performed by incubating the glass with a
solution of fibronectin (Sigma) in PBS at 20 μg/mL for 1 h at 37 °C.
The sample was then submerged for 1 h in a 0.2% Pluronics
solution (BASF) (vol/vol) in PBS to prevent cell adhesion on the
pillar walls. Cells were seeded highly concentrated in a small
volume, close to the pillar stamp, to allow an even distribution of
the cells between the pillars. After 15 ± 3 h a confluent monolayer
was formed, and PDMS stencils were carefully peeled off with
tweezers. For the experiments of different cell packing densities,
the density of cells at the seeding step was varied according to
different densities degrees, but the waiting time before removing
the PDMS stencil was kept the same. On the other hand, to create
damage-associated gaps, we proceeded with two distinct experimental approaches: (i) to gently press the PDMS pillar array
against a confluent monolayer in such a way that cells beneath the
pillar are killed (“crushed gaps”); and (ii) to run the pillar removal assay without previous incubation of the pillars stencil with
pluronics in such a way that cells adhere to pillars walls and
membranes are scratched upon pillar removal (“ripped gaps”).
Cell Damage Assessment. We assessed cell damage by both propidium iodide (Sigma) and FITC-dextran (40 kDa, Sigma) uptake.
Propidium iodide labels the nuclei of damaged cells, whereas
FITC-dextran is internalized by membrane-disrupted cells.
[Y27632 (Calbiochem), ML-7 (Calbiochem), NSC23766 (Calbiochem) and blebbistatin (Sigma)] were added at 25 μM at least 2 h
before releasing the PDMS stamp to ensure proper inhibition of
the cells under the PDMS stamp.
Video Microscopy and Image Analysis. Live cell imaging was performed in a Nikon or Olympus microscope, enclosed in an incubator to maintain the samples at 37 °C and 5% of CO2
throughout the experiments. Images were acquired typically every 15 s with Metamorph software. Live cell tracking was accomplished by incubating cells with DAPI at 4 μg/mL in cell
culture media for 6 h and thorough rinsing before imaging.
Images were analyzed and processed using the software ImageJ.
Immunofluorescence Microscopy. Cells were fixed at different time
points upon pillar stencil removal with 4% paraformaldehyde
(Sigma) for 30 min, permeabilized with 0.25% Triton X-100
(Sigma) in PBS (vol/vol) for 5 min, and blocked in 1.5% BSA
(Sigma) in PBS for 30 min. Staining was performed with primary
rabbit antibody against phospho-MLC (Cell Signaling) diluted
1:200 in 2% BSA in PBS and detected with a polyclonal antirabbit conjugated with 488 Alexa Fluor (Invitrogen) at 1:100 in
PBS. Actin was visualized with Alexa Fluor 564-conjugated
phalloidin (Invitrogen) at 1:1,000 in PBS. Fibronectin and laminin
were labeled with a primary rabbit antibody against fibronectin
(Invitrogen) and a primary mouse antibody for laminin, both at
1:200, and a secondary polyclonal anti-rabbit and anti-mouse
(respectively) conjugated with 488 Alexa Fluor (Invitrogen) at
1:300. For adhesions staining, a primary antibody in rabbit for
ZO-1 (Zymed, Invitrogen) and a primary antibody in mouse for
E-cadherin (BD Transduction) were used at a dilution of 1:1,000
in 1% BSA/PBS and detected with secondary antibodies against
mouse and rabbit conjugated with 488 Alexa Fluor (Invitrogen).
Samples were mounted in Mowiol and imaged with a Nikon A1R
confocal microscope with a 60× or 100× 1.4 N.A. apochromat oil
immersion objective lens. When indicated, samples were imaged
in epifluorescence with a Deltavision microscope with a 60× 1.4
N.A. apochromat oil immersion objective lens.
(MDCK) strain II cells and stable MDCK cell line expressing actinGFP or lifeact-GFP were used. Cells were cultured in DMEM
supplemented with 10% FCS, 100 U/mL penicillin, and 100 μg/mL
streptomycin and maintained at 37 °C in a humidified atmosphere
with 5% CO2. For transfected MDCK cells, the selection antibody
geneticin was added at 2 μg/mL. α-Catenin knockdown MDCK
cells (kindly provided by James W. Nelson, Stanford University)
were maintained as the wild-type cells. Pharmacological inhibitors
Preparation of Substrates of Different Stiffness. Different PDMS
stiffness substrates were achieved by mixing PDMS at different
cross-linker:base ratios, namely 1:10 for very stiff substrates,
comparable to glass coverslips, 1:25, 1:40, and 1:60. At a 1:60 ratio,
the substrate remained mainly elastic with a Young’s modulus of
approximately 20 kPa (2). A small drop of the mixture was placed
on top of a glass coverslip, rendering a thin layer of PDMS. PDMS
substrate was cured for at least 2 h at 80 °C. The following day it
was coated with fibronectin at 20 μg/mL for 1 h at 37 °C and
proceeded with the described gap patterning protocol.
1. du Roure O, et al. (2005) Force mapping in epithelial cell migration. Proc Natl Acad Sci
USA 102:2390–2395.
2. Murrell M, Kamm R, Matsudaira P (2011) Substrate viscosity enhances correlation in
epithelial sheet movement. Biophys J 101:297–306.
Cell Culture and Drug Treatments. Madin-Darby canine kidney
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Fig. S1. Initial conditions of cells surrounding the pillars. (A and B) Immunostaining against adhesion proteins: tight junction protein ZO-1 (A) and adherens
junction protein cadherin (B). Dotted line indicates the pillar position. Note that there are no specific adhesions of the border cells with the pillar. Cells show no
preferential alignment along the gap perimeter. (C–E) Cells around circular pillar under increasing degree of confluency, from fairly spread (C) to highly dense cells
(E). (F–H) Cells are randomly distributed around ellipsoidal gaps. The differential convexity of the gap perimeter does not determine the positioning or alignment
of cells at its poles. (I–K) Again, the squared shape causes no cornering effect on the distribution of cells, which are evenly positioned. (Scale bars, 20 μm.)
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Fig. S2. Characterization of damage-associated “wound” closure experiments. (A and B) Assessment of cell damage/death in damage-associated gaps by FITCdextran incorporation (A) and propidium iodide (B). Crushed gaps result from pressing the pillar array against a confluent monolayer, and ripped gaps are
produced by the pillar removal assay without pluronic treatment. In contrast with our damage-free method, both strategies resulted in extensive cell death
around the gaps. (Scale bars, 20 μm.) (C) Decrease of area over time of the damage-free gaps (created with the pillar removal assay) (red line) vs. the damageassociated gaps (black line). The area has been normalized by the initial area to homogenize the different measurements. (D) Closure time of the damage-free
gaps in control conditions and under Rac inhibition (gray columns) with respect to damage-associated gaps under low- or high-density conditions and with Rac
inhibition (purples and red).
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Fig. S3. (A) Gap area decrease with time, for different pillar sizes used for creating the gaps, ranging from 15 to 150 μm in diameter. (B) Kymograph corresponding to the orange line of the time-lapse snapshot in the left. Frame rate was one frame every 2 s. (Scale bar, 20 μm, Left, and 3 μm, Right.)
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Fig. S4. Extracellular matrix assembly beneath cell culture. We have tested fibronectin location by two approaches: using fluorescently labeled fibronectin to
coat the substrate (A–C) and immunostaining fibronectin to observe fibronectin secretion and reorganization by cells (D–F). (A) Pillars stencil between the
epithelial culture on glass at low magnification. (B) In a close-up it can be appreciated that fibronectin is present beneath PDMS pillar. (C) Removal of the
stencil does not tear the fibronectin beneath. (D) Fibronectin and actin staining after 30 min of pillar removal and (E) at the closure time (65 min after pillar
removal). (F) Fibronectin deposition on soft PDMS substrates at 1:40 cross-linking ratio is not altered. (G) Immunostaining for visualizing laminin assembly:
laminin is not present in the gap area, whereas it is in the area covered by cells. (Scale bars, 20 μm.)
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Fig. S5. Effect of cell density on cell monolayer thickness. Central images are the mid-plane of a z-stack, and bottom and right images are orthogonal
projections of the stacks on the yellow lines. (A) Right upon reaching confluency, cells are highly spread and thus flat. (B) As cell density under the pillar stencil
increases, cells become smaller and taller. In highly packed cultures, the thickness of the layer is greater than in sparse epithelia. As can be observed in the x–z
and y–z orthogonal projections, the lateral membrane in dense cultures is larger.
Fig. S6. Dose–inhibition response curve for the different inhibitors used (NSC23766, Y-27632, and ML-7) and their vehicles (either DMSO or water). The three
concentrations tested are the most recurrently found in literature; 25 μM proved to be an adequate inhibitory concentration in our setup.
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Fig. S7. Actin distribution showed by phalloidin staining. (A) Presence of the pillar does not promote accumulation of actin surrounding the pillar. (B)
Quantification of actin fluorescence intensity in radial profiles (concentric circles of increasing diameter) at each given distance from the center of the gap. (C)
Two minutes after releasing the pillar stencil, there is some actin clustering at the gap interface. (D) Peak of actin fluorescence intensity is more defined (higher
and narrower). (E and F) Actin staining 30 min after pillar removal. As closure progresses, actin is still accumulated at the gap border where there are no
lamellipodia (E) (see y–z orthogonal projection), whereas where the actin protrudes into the lamellipodia there is no actin clustering (F). (Scale bars, 20 μm.)
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Fig. S8. Coordination of cells via cell–cell junctions and myosin action. (A–F) Trajectories of cells during the closure process. The tracks are shown after closure
is completed. The dotted lines in the phase-contrast images represent the original gap margin. (A) Cell trajectories of control experiments. (Scale bar, 20 μm.)
(B) Graph shows the trajectories (axis in micrometers) as a function of the cells’ position with respect to the gap edge (i.e., first row corresponds to cells
contacting the gap, second row refers to those behind, and consecutively). (C and D) α-Catenin knockdown MDCK cells during closure. (Scale bar, 20 μm.) (E and
F) Blebbistatin-treated cells during closure. (Scale bar, 10 μm.) Trajectories from one experiment are shown, trends being representative of three other experiments analyzed. (G) Alignment of cells at the end of closure related to their initial shape, depending on their position with respect to the gap. (H) DisLegend continued on following page
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placement of cells indicated as a percentage of the initial radius of the gap, for control, α-catenin knockdown MDCK cells, and blebbistatin-treated cells,
indicating the effective net movement of cells. (I) Closure time of 20- and 50-μm diameter gaps of α-catenin knockdown MDCK cells and cells treated with
blebbistatin with respect to control conditions. For G, H, and I, data represent means and SEs of at least three experiments. (J) Changes in nuclei circularity
during closure. Cells analyzed correspond to those contacting the gap (border cells), cells at the edge of the gap that extend lamellipodial during the closure,
and cells far from the gap (outer cells) (typically taken at the four corners of the field of view).
Table S1. Comparison of the calculated area of the pillar used to
pattern circular gaps with respect to the measured gap area right
after pillar removal
Pillar diameter (μm)
Pillar area
(μm2)
Mean gap area
(μm2)
SD of gap area
(μm2)
177
314
707
1,257
1,963
2,827
163
327
727
1,311
1,965
2,826
33
15
30
58
14
11
15
20
30
40
50
60
Experimental data shown are means and SD of a minimum of seven experiments.
Table S2. Number of cells considered for the grouping on
different degrees of cell density, always after reaching
confluence
Normalized cell density
<0.33
0.33–0.46
0.46–0.6
0.6–0.8
0.8–1
Cells/mm2
<1,200
1,200–1,700
1,700–2,200
2,200–3,000
3,000–3,700
Normalized cell density is obtained by dividing the number of cells per
mm2 by the maximal number of cells per mm2. The bin labeled as <0.33
depicts low density, and the bin 1 is the mean of the three experiments at
highest density. Data are mean and SD of at least three experiments each.
Movie S1.
Video microscopy of gap closure. Note that lamellipodia are being extended by all of the cells contacting the gap. (Scale bar, 10 μm.)
Movie S1
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Movie S2. Video microscopy of a large gap closing. Note that broad lamellipodia are extended by cells surrounding the gap. (Scale bar, 20 μm.)
Movie S2
Movie S3. Video microscopy of gap closure on 1:40 PDMS cross-linking ratio substrate. Note that closure is impeded and gap acquires noncircular shapes.
(Scale bar, 15 μm.)
Movie S3
Movie S4.
Video microscopy of gap closure under myosin light chain kinase inhibition by addition of 25 μM of ML-7. (Scale bar, 10 μm.)
Movie S4
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Movie S5. Video microscopy of gap closure under Rho kinase inhibition by addition of 25 μM of Y27632. (Scale bar, 15 μm.)
Movie S5
Movie S6.
Video microscopy of gap closure under Rac1 inhibition by addition of 25 μM of NSC23766. (Scale bar, 10 μm.)
Movie S6
Movie S7.
Video microscopy of an ellipsoidal gap closing. (Scale bar, 20 μm.)
Movie S7
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Movie S8.
Video microscopy of a squared gap closing. (Scale bar, 20 μm.)
Movie S8
Movie S9. Video microscopy of gap closure with α-catenin knockdown MDCK cells. (Scale bar, 20 μm.)
Movie S9
Movie S10.
Video microscopy of a large gap closing under myosin inhibition by addition of 25 μM of blebbistatin. (Scale bar, 20 μm.)
Movie S10
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