Polarizing cytoskeletal tension to induce leader cell formation during collective cell migration
© Rausch et al.; licensee Springer. 2013
Received: 15 October 2013
Accepted: 12 November 2013
Published: 22 November 2013
The collective migration of cells is fundamental to epithelial biology. One of the hallmarks of collective behavior in migrating cohesive epithelial cell sheets is the emergence of so called leader cells. These cells exhibit a distinct morphology with a large and highly active lamellipodium. Although it is generally accepted that they play a crucial part in collective migration, the biophysical factors that regulate their formation remain unknown.
Here we show that a geometry-based cue like local variation of curvature of the collective’s perimeter is capable of triggering leader cell formation and promoting enhanced motility at defined positions. Remarkably, the extent of this effect scales with the magnitude of the curvature.
Cytoskeletal tension was found to be important for geometry induced leader cell formation, as cells treated with tension reducing agents appeared less sensitive to local curvature variation. Accordingly, traction force microscopy revealed an increased level of shear stress at highly curved positions even before the cell migration had actually started, indicating the presence of a collective polarization induced by the geometry of the confinement.
Together our findings suggest that high curvature leads to locally increased stress accumulation, mediated via cell-substrate interaction as well as via cytoskeleton tension. The stress accumulation in turn enhances the probability of leader cell formation as well as cell motility. This work defines the importance of geometric cue such as local curvature in the collective migration dynamics of epithelial cells and thus shows implications for the biophysical regulation of epithelium during wound healing, embryonic development, and oncogenesis.
KeywordsCollective migration Leader cell Geometric constraint Traction force MDCK cells
A hallmark of collective migration is the appearance of so called leader cells. These cells exhibit a distinct morphology with a large and highly active lamellipodium . In a classical wound-healing scenario these cells emerge at the leading edge of the migrating cell ensemble, where a fraction of cells acquires the leader cell characteristics early in the migration process [4, 5].
Leader cells have been suggested to play an active role in guiding the collective forward [6–9] accompanied by high tension within the collective . As numerous cells behind the leading edge add considerably to the total traction force exerted on the substrate, leader cells might also merely happen to be in the leading position without adding a substantial share to the migration process [11, 12]. Yet, leader cell formation has been consistently observed in collective cell migration, and thus it is generally accepted that leader cells play a crucial part in the migration process. Although their origin is assumed to be governed by a variety of chemical and physical signals, the basic biophysical factors underlying leader cell generation could not be conclusively identified .
In order to probe the role of the diverse and often conflicting signals, several recent studies have aimed at reducing the complexity of this multiparametric problem by creating well-defined model systems. In order to control the size and shape of cell ensembles, a very successful system has emerged making use of confinements of different kinds [4, 14–17]. This enables quantitative studies by reducing the complexity of the cell collective system by allowing the control of pivotal parameters of cell collectives, namely size, cell density and general shape . Shape is described best by length and curvature of the cell collective’s perimeter. As a matter of fact, controlling merely the parameter of curvature has been shown to reproduce experimental behavior of finger formation typically involved in leader cell formation in a computational model .
Moreover, several experimental studies indicate that the probability of leader cell formation might be enhanced by convex boundaries of the cell collective [20, 21] similar to what has been shown for the directed migration of single cells . Additional results highlighting the role of geometry for diverse physiological processes have been derived from experiments with spatially confined cell clusters. Cell collectives patterned on adhesive islands preferentially extended new lamellipodia from their corners . Also, a strong correlation of geometry and cell proliferation was observed, revealing that the latter can be an active regulator of tissue growth . Taken together, these findings hint towards curvature being a general parameter underlying biological and especially migration processes.
The question how local curvature in a confined setting effects leader cell formation in a subsequently triggered collective cell migration has not yet been resolved. We aim for a conclusive understanding of this crucial parameter and the underlying mechanisms involved. For this purpose we designed experiments that enabled us to largely emphasize local curvature as a mechanical cue in comparison to other factors. Therefore we developed a novel micro-stencil technique in order to precisely control the cell collective’s area and its global as well as local perimeter curvature. We used two dimensional epithelial cell sheets on fibronectin coated surfaces in order to uncouple and analyze this particular parameter in a very well defined experimental setting. This allowed us to gain quantitative data by focusing on the role of curvature of the cell collective’s perimeter on leader cell formation.
This work shows that local variation in curvature of the cell collective’s perimeter correlates with locally increased motility, leader cell formation and traction stress. We identified the organization of the actin cytoskeleton and its collective polarization as a possible candidate for relying information about physical stress accumulation which in turn increases the probability of leader cell formation.
To obtain geometrically well-defined cell collectives, we employed micro-stencils made of polydimethylsiloxane (PDMS). Stencil masks were fabricated in an adapted soft lithography process . In short, SU-8 25 negative photo resist (MicroChem, Newton, MA, USA) was spincoated on a 2“ silicon wafer (Si-Mat, Kaufering, Germany) in a clean room facility, prebaked on a hot plate, illuminated for 12 sec in Mask Aligner MBJ4 (Suess MicroTec Lithography, Munich, Germany) and baked again on a hot plate. To remove non-irradiated SU8 resist, wafers were bathed in SU-8 Developer mr-Dev600 (Microresist Technology, Berlin, Germany) and then treated with 1H,1H,2H,2H-Perfluorooctyl-trichlorosilane to reduce adhesiveness. A sandwich consisting of the wafer with photoresist structures, 0.5 mL of uncured PDMS, a piece of parafilm, a piece of paper and a glass slide was put into a custom made molding press to obtain uniform pressure distribution. The assembly was put into a compartment dryer at 65°C for 100 min to allow PDMS polymerization. PDMS membrane thickness of 50–60 μm was achieved regularly. To prevent cell adhesion, stencil masks were incubated in a solution of Pluronic F-127 (Sigma Aldrich, 2% w/v in deionized water) for 30 minutes prior to use.
MDCK II cells were seeded on fibronectin coated surfaces partially blocked by micro-stencils. They were maintained in Minimum Essential Medium Eagle supplemented with 5% FBS, 2 mM L-glutamine, 10U mL-1 penicillin and 10 μg mL-1 streptomycin. The average density of cells compromising a single collective was about 3600 cells/mm2, or 350 cells per collective.
Time lapse image acquisition was performed on an inverted Observer microscope (Zeiss, Germany) directly after removal of the micro-stencils. Phase contrast images of at least 95 individual collectives distributed into at least two independent experiments for each stencil type used were acquired every 5 min using a 10x objective. Coordinates and timepoints of leader cell formation were determined by hand. All other data analysis were performed with Matlab (Mathworks, Germany).
Inhibition experiments were conducted with Blebbistatin and Y-27632 to reduce cytoskeleton tension. Drugs were added to the medium 1 hour before start of the experiment in a concentration of 50 μM (Blebbistatin) or 30 μM (Y-27632). During experiments, i.e. after removal of the stencil mask, cells were maintained in standard cell culture medium supplied with 5 μM blebbistatin or 3 μM Y-27632, respectively. For control experiments cell collectives were incubated for one hour in Opti-MEM containing DMSO (1 μL per mL of medium) before the stencil mask was removed. The experiment was then conducted in standard cell culture medium.
Traction force microscopy was carried out as previously described  on polyacrylamide (PAA) substrates with a Young’s modulus of about 23kPa, in which fluorescent 500 nm carboxylated polystyrene beads were embedded as position markers. To ensure cell attachment PAA was covalently functionalized with fibronectin. Bead displacements were tracked following a Matlab adaptation of the algorithm developed by Crocker and Grier . Subsequently a regularized Fourier-transform traction cytometry was employed to calculate the traction  in each independent cell collective of which 17 were superimposed to calculate the average stress distribution. For all traction field reconstructions the regularization parameter, which effectively filters out high frequency noise, was kept constant.
Cell stainings were performed on fixed and permeabilized cells with the primary antibody, rabbit monoclonal [clone Y113] to Paxillin (ab32084, Abcam, Germany), followed by anti-rabbit secondary antibody tagged with the fluorescent dye Alexa Fluor 488 (Invitrogen, Germany), and with DNA-binding dye 4′,6-diamidino-2-phenylindole (DAPI, 1 ug/ml in PBS, Invitrogen, Germany). Visualization of the actin cytoskeleton was done by adding TRITC-labeled phalloidin at the secondary incubation step, if required.
Analysis of the actin belt was based on the computation of the angular distribution of stained actin within an approximately 4 μm wide region along the boundary of the cell collectives.
The significance in all experiments was determined using the Mann–Whitney-Wilcoxon test.
Contraction of the colony monolayer was simulated using a two-dimensional continuum model that has been introduced previously by Edwards and Schwarz . In this model, an isotropic and homogeneous active stress is introduced into the elastic equations for a thin elastic sheet which in turn is coupled to an elastic foundation. For a given geometry, this model is solved numerically with Finite Element Methods (FEM) in Comsol Multiphysics. The model has two free parameters, the coupling constant κ and the contractile pressure σ con . As input for the model fitting we used the derived mean displacement field and reconstructed traction pattern. From the model the traction can be calculated by T = κu, while u is the calculated model displacement, which depends on both σ con and κ. The parameters were optimized by sampling, fitting once the data of the spike shaped pattern. Here, we adjusted the parameters in such a way that a best agreement with measured displacement and reconstructed traction pattern was achieved. More details on the methods described in this section can be found in the supplementary information (Additional file 1).
3 Results and discussion
3.1 Migration assay of geometrically well defined epithelial cell collectives
We sought to derive quantitative information on the influence of curvature on collective cell migration driven by the formation of leader cells. For this purpose we developed a micro-stencil technique to reproducibly create cell collectives with well-defined geometrical shapes. The essential part of the micro-stencils is a thin PDMS-membrane with precisely defined holes that can be placed on any adhesive surface. Initially the cells form a monolayer covering the area inside the holes in the geometry induced by the PDMS stencil. Removal of the stencil triggers free migration of cells on the freshly uncovered substrate. Since cells are not damaged in this process, the experimental conditions are better controlled in comparison to classical scratch wounding assays, where local necrotic damage might influence cell behavior.
Next we investigated if the increase in local curvature using collective geometry with hemispherical protrusions alters the behavior of the collective. Again we analyzed the angular position of leader cell appearance, this time in such a way that the center of the protrusions was located at an angle of 45° to an arbitrarily fixed line. We observed an enhanced probability of leader cell formation at the angles corresponding to the positions of hemispherical protrusions (Figure 3B). Interestingly, the formation of leader cells in this case was also slightly delayed on average. There were still leader cells forming at every time point of the experiment, but only one third of the total number formed during the first 90 minutes (Additional file 4: Figure S4).
Finally, we used collective geometry with triangular protrusions, which introduced an even higher local curvature into the system at the tip of the triangle. We found that in this setting, the probability of leader cells forming at the protrusions is again increased almost twofold with respect to the hemispherical design (Figure 3C). This increase can directly be attributed to the higher local curvature, since the only difference to the former setting is the different geometry yielding an increased curvature at the tip of the triangle. In this scenario more than half of the leader cells formed during the first 90 minutes of the experiment (Additional file 5: Figure S5).
The average number of leader cells forming over 6 hours per collective was determined to be 10.0 ± 0.2 for circular collectives, 8.5 ± 0.2 for collectives with hemispherical protrusions and 10.4 ± 0.2 for collectives with spike protrusions. This shows that not all of the leader cells were emerging at one of the protrusions. However, we intentionally placed the protrusions at low proximity in order to have them separated spatially far enough as to avoid interference between them. Yet, these results clearly show that the actual position of leader cell appearance changes from a random distribution towards a distribution with enhanced probability at positions of higher local curvature and that this effect scales with the magnitude of the local curvature.
Furthermore, we analyzed the displacement speed of the collective’s perimeter over time. In addition to the earlier observation of increased probability for leader cell formation at spike protrusions, we also observed that the radial velocity of migration of the perimeter at these positions is higher during the early stage of migration (i.e. during first two hours of migration, Figure 2D). A linear fit to the data of the first 90 minutes shows an average velocity at the spike protrusions of 0.24 μm min-1 (filled quares in Figure 2D) which is significantly higher than the average velocity of 0.18 μm min-1 (gray circles in Figure 2D) at the normal curved regions. The local increase in velocity correlates directly with the enhanced formation of leader cells at these positions indicating that they indeed play an important role for the migration process by locally enhancing the outward directed migration speed. A fit to the data of perimeter displacement of a completely circular collective without protrusions of enhanced curvature yielded an average cluster expansion velocity of only 0.14 μm min-1 (white circles in Figure 2D). This velocity of 0.14 μm min-1 at collectives without protrusions is slower than the velocity of 0.18 μm min-1 within the 0° to 10° angular section of collectives with protrusions, although the local curvatures are identical in these both cases. Thus, the increase in curvature not only increases the velocity at positions of high local curvature, but also leads to a general velocity increase across the whole collective even at positions without increased curvature.
Taken together our results show a clear correlation between increased probability of leader cell formation and increased local curvature resulting in turn in a locally enhanced migration velocity of the cell collective. Thus, leader cell formation in fact plays an active role in the collective migration process.
3.2 Role of intracellular tension in geometry induced leader cell formation
Previous studied indicate that cell monolayers exist in a state of tensile stress [11, 32, 33]. Additionally, it has been shown that extracellular compressive stress such as one imposed by the hydrostatic pressure plays a role in leader cell formation . This observation led us to the question if an increase in local perimeter curvature is actually accompanied by an increased local tension or stress level as a reaction of the cell collective. To test this hypothesis, we treated cell collectives with blebbistatin and Y-27632 which are known to reduce the intracellular tension. Blebbistatin inhibits myosin II activity  and thus cell motility. The pyridine derivative Y-27632 is known to inhibit the Rho-associated protein kinase (p160ROCK) pathway, which in turn directly decreases actomyosin-mediated contractile tension . We used these drugs in concentrations that have recently been shown to not compromise the stress induced leader cell formation .
The addition of DMSO as a vehicle for both drugs was shown not to influence the experimental conditions in an independent set of control experiments (Figure 4A).
In conclusion these results offer complementary data to our previous finding that the probability of leader cell formation scales with the intracellular tension. A slightly enhanced probability still remains at the highly curved regions even after drug treatment. Nonetheless, the magnitude of the effect clearly shows that in fact cytoskeletal tension plays an important role in the formation of leader cells.
3.3 Curvature dependent increase in local traction force at the cell-substrate interface
This evidence shows that cells within the protrusions with high local curvature exert strong pulling forces on the substrate even before onset of the outward directed migration process. In contrast, there is no such pronounced accumulation of traction stress present near or at the normal curved perimeter, ruling out the possibility that this effect is simply due to the proximity to the collective’s perimeter itself.
Since much of the cellular traction is known to be transmitted through sites of focal adhesion, we further looked into the distribution and orientation of focal adhesion points at different locations of the collectives. Focal adhesions play a major role in connecting cells with the substrate in order to exert forces, and they are involved in transmitting mechanical forces as well as regulatory signals. We stained cell collectives with antibodies against the focal adhesion complex protein Paxillin to gain further information on the mechanical stress state between cells and substrate. The necessary fixation was done directly after removal of the stencil mask, allowing no time for major reorganization of focal adhesions which takes at least several minutes . At the cell collectives perimeter we found a striking difference in the orientation of focal adhesion points between areas of normal curvature and areas of protrusions with high curvature. The focal adhesions in the normal curved regions are oriented tangential to the perimeter of the collective. In contrast, the focal adhesions in the area of a spike protrusion are oriented radially with respect to the center of the collective (Additional file 7: Figure S7). This correlates well to the results of the traction force microscopy experiments.
We further investigated the relevance of local curvature for the appearing stress distribution within the collective using a two-dimensional continuum model which has been introduced previously by Edwards and Schwarz . Here, the cell collective was represented by a homogeneous contractile layer of an elastic, isotropic material which is elastically coupled to the substrate. This represents the fact that contractile forces are generated and transmitted throughout the entire cell monolayer while each individual cell feels its interaction with the underlying substrate locally. The two free model parameters colony contractility σ con and the substrate coupling constant κ have been fitted regarding only the two values for maximal measured mean displacement and traction of all experiments as well as their respective geometry (Figure 5C). As values we derived κ = 130pN/μm and σ con = 3.8pN/μm. Although the model represents a coarse-grained situation, generic differences in traction magnitude based on the initial geometry are clearly evident. The model predicts both the homogeneously distributed elevated traction at the rim and the pronounced stress distribution in the protrusions. In particular, it predicts a difference in the average stress distribution of about 50 Pa between the highly curved protrusion at 45° (Figure 5C) and the circular part at 0° (Figure 5D) in good agreement with the experimental results.
In summary these results show that the local increase in curvature is sensed by the cell collective and converted into a local accumulation of traction force at the cell-substrate interface. This change in cell induced stress distribution is also reflected by the reorientation of the focal adhesions which link the cytoskeleton to the substrate. When we analyzed this situation in a numerical continuum model, we found that contractile forces in conjunction with geometry are sufficient to explain the accumulation of traction stress in regions of high local curvature. This agreement clearly shows that regarding stress distribution, cells within the cell collectives act in a collective manner rather than as single cells.
3.4 Role of pluricellular actin belt for leader cell formation
Stress fibers consisting of actomyosin bundles play an important role for mechanotransduction in diverse cellular systems . Enhanced mechanical stress levels in the cells favor the formation of stress fibers. Since our results from traction force microscopy strongly indicate that local accumulation of traction stress favors the formation of leader cells, we decided to look into the distribution of actin filaments in more detail.
In multicellular systems, connected actin cytoskeletons transmit intercellular stress over length scales larger than a single cell . A multicellular actin belt, comprised of actin stress fibers lining the perimeter of cell collectives, has been reported previously in many in vivo and in vitro systems [40, 41]. This structure plays a major role in the purse string wound closure and in multicellular migration [4, 5, 7]. The purse string model describes wound healing of mainly small wounds with concave boundaries by formation of an actomyosin cable and subsequent contraction of the actin belt at the wound borders. It is believed that the actin bundles (also called actin cables) play a role for force transmission to distribute arising forces more evenly among cells of differing activity .
In contrast to the purse string model, which is assumed for small concave wounds, our observation is based on structures with convex boundaries. However, in the experiments presented here the actin belt might also be involved in force transmission, similar to a purse string model where contractile stress is mediated by actomyosin bundles. The break in the actin belt structure at the very tip of the highly curved spike protrusions is formed before the barrier is lifted, i.e. before migration starts. It is reasonable to assume that this might facilitate the outgrowth of cells in the subsequent migration process, and thus result in higher incidences of leader cell formation.
In order to study the collective behavior of migrating epithelial cell sheets, we designed PDMS-membranes that induce well-controlled geometrical constraints on the cell collectives. Thus we were able to compare the effect of different local curvatures of the perimeter on the collective’s migration behavior. The evidence described here clearly shows that the collective is polarized in terms of mechanical stress even before the onset of the migration process. This stress polarization is directly translated to an increased probability of leader cell formation and thus higher migration velocity. This is evident by a break in the actin belt, the inhomogeneous force distribution map, as well as the orientation of focal contacts and is further supported by complementary data from a mathematical model.
The distribution of actin stress fibers is known to give insights into the state of tension in cells. We observed a prominent pluricellular actin belt encompassing the whole collective. These actin bundles might give a possible explanation on how cells transmit curvature information over many cell lengths. Since they span the whole cell collective’s perimeter they might be in a tensile state resembling the local curvature situation. The mechanotransducting effect of these bundles might thus gather strain information and transmit it to the tip point where the curvature and thus local strain is too high to maintain the structural integrity. Accordingly, at these tips we observed a gap in the actin rich structure giving the tip cells an advantage during the onset of migration. They are thus more prone to become leader cells due to less obstructive force of the actin bundles.
Similarly, the traction stress distribution is sensitive to the geometry of the collective. The accumulation of traction stress shows that these cells undergo what we call stress polarization. Here, we discovered a considerable accumulation of stress pointing towards the center of the collective in sections of the perimeter with high local curvature. This is further supported by the finding that focal adhesions in highly curved protrusions are oriented radially.
The initial collective polarization might give cells at the positions of high local curvature a head start when the migration process is initiated by removal of the physical barrier. This explains the increased migration velocity and the significantly enhanced probability of leader cell formation at highly curved protrusions in comparison to perimeters with lower local curvature. The increased probability of leader cell formation scales with the magnitude of the curvature increase and is consistent with our finding that by inhibiting cytoskeletal tension through the addition of drugs the angular distribution of leader cell formation is broadened drastically. In this case the information that a portion of the collective is exposed to higher tension due to curvature can no longer be transmitted sufficiently enough between cells and the prepolarizing effect is mostly lost.
In general, woundhealing scenarios can roughly be divided into two major categories depending on the wound size. One is the closure of very small wounds, which has been shown to take place by an actively contracting actomyosin cable at the wound perimeter . The other mechanism, which can be observed at larger wounds, is the active collective migration of cells towards each other for a reepithelialization of the denuded wound surface . Our results suggest that in our experimental framework both mechanisms contribute to a collective migration scenario: The active migration of monolayers of cells in response to increased magnitudes of curvature is supported by a multicellular actin structure playing a role in information transmission before the actual active migration takes place.
It is reasonable to assume that in an in vivo situation the reaction to local curvature is also beneficial for invading tumor cell clusters. There, a small local difference in the rigidity of the surrounding tissue might easily lead to a formation of a locally slightly higher curved region in the tumor surface area. This in turn enhances migrational activity towards the weaker part of the surrounding tissue.
Our results show that plain geometrical parameters, in this case curvature variation, have a major impact on collective cell migration in spite of the inherent complexity of the living system.
The authors thank Christiane Antoni for technical assistance and Tabea Mundinger for proof reading and valuable discussion. This work was supported by the Max-Planck Society. JPS is the Weston Visiting Professor at the Weizmann Institute of Science. JPS and USS are members of the Heidelberg cluster of excellence CellNetworks. This work was supported by CellNetworks EcTop2, the grass-root project of the Max Planck Institute for Intelligent Systems, and the MechanoSys-grant of the BMBF. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
- Streichan SJ, Valentin G, Gilmour D, Hufnagel L: Collective cell migration guided by dynamically maintained gradients. Phys Biol 2011,8(4):045004.View ArticleGoogle Scholar
- Friedl P, Hegerfeldt Y, Tusch M: Collective cell migration in morphogenesis and cancer. Int J Dev Biol 2004,48(5–6):441–449.View ArticleGoogle Scholar
- Martin P: Wound Healing–Aiming for Perfect Skin Regeneration. Science 1997,276(5309):75–81.View ArticleGoogle Scholar
- Poujade M, Grasland-Mongrain E, Hertzog A, Jouanneau J, Chavrier P, Ladoux B, Buguin A, Silberzan P: Collective migration of an epithelial monolayer in response to a model wound. Proc Natl Acad Sci 2007,104(41):15988–15993.View ArticleGoogle Scholar
- Omelchenko T, Vasiliev JM, Gelfand IM, Feder HH, Bonder EM: Rho-dependent formation of epithelial “leader” cells during wound healing. Proc Natl Acad Sci 2003,100(19):10788–10793.View ArticleGoogle Scholar
- Khalil AA, Friedl P: Determinants of leader cells in collective cell migration. Integr Biol 2010,2(11–12):568–574.View ArticleGoogle Scholar
- Lim JI, Sabouri-Ghomi M, Machacek M, Waterman CM, Danuser G: Protrusion and actin assembly are coupled to the organization of lamellar contractile structures. Exp Cell Res 2010,316(13):2027–2041.View ArticleGoogle Scholar
- Vitorino P, Meyer T: Modular control of endothelial sheet migration. Genes Dev 2008,22(23):3268–3281.View ArticleGoogle Scholar
- Aigouy B, Lepelletier L, Giangrande A: Glial chain migration requires pioneer cells. J Neurosci 2008,28(45):11635–11641.View ArticleGoogle Scholar
- Reffay M, Petitjean L, Coscoy S, Grasland-Mongrain E, Amblard F, Buguin A, Silberzan P: Orientation and polarity in collectively migrating cell structures: statics and dynamics. Biophys J 2011,100(11):2566–2575.View ArticleGoogle Scholar
- Trepat X, Wasserman M, Angelini T, Millet E, Weitz D, Butler J, Fredberg J: Physical forces during collective cell migration. Nat Phys 2009,5(6):426–430.View ArticleGoogle Scholar
- Farooqui R, Fenteany G: Multiple rows of cells behind an epithelial wound edge extend cryptic lamellipodia to collectively drive cell-sheet movement. J Cell Sci 2005,118(Pt 1):51–63.View ArticleGoogle Scholar
- Trepat X, Fredberg JJ: Plithotaxis and emergent dynamics in collective cellular migration. Trends Cell Biol 2011,11(21):638–646.View ArticleGoogle Scholar
- Vedula SRK, Leong MC, Lai TL, Hersen P, Kabla AJ, Lim CT, Ladoux B: Emerging modes of collective cell migration induced by geometrical constraints. Proc Natl Acad Sci 2012,109(32):12974–12979.View ArticleGoogle Scholar
- Anon E, Serra-Picamal X, Hersen P, Gauthier NC, Sheetz MP, Trepat X, Ladoux B: Cell crawling mediates collective cell migration to close undamaged epithelial gaps. Proc Natl Acad Sci 2012,109(27):10891–10896.View ArticleGoogle Scholar
- Ng MR, Besser A, Danuser G, Brugge JS: Substrate stiffness regulates cadherin-dependent collective migration through myosin-II contractility. J Cell Biol 2012,199(3):545–563.View ArticleGoogle Scholar
- Hirashima T, Hosokawa Y, Lino T, Nagayama M: On fundamental cellular processes for emergence of collective epithelial movement. Biol Open 2013,2(7):660–666.View ArticleGoogle Scholar
- Nelson CM, Jean RP, Tan JL, Liu WF, Sniadecki NJ, Spector AA, Chen CS: Emergent patterns of growth controlled by multicellular form and mechanics. Proc Natl Acad Sci 2005,102(33):11594–11599.View ArticleGoogle Scholar
- Mark S, Shlomovitz R, Gov NS, Poujade M, Grasland-Mongrain E, Silberzan P: Physical model of the dynamic instability in an expanding cell culture. Biophys J 2010,98(3):361–370.View ArticleGoogle Scholar
- Rolli CG, Nakayama H, Yamaguchi K, Spatz JP, Kemkemer R, Nakanishi J: Switchable adhesive substrates: revealing geometry dependence in collective cell behavior. Biomaterials 2012,33(8):2409–2418.View ArticleGoogle Scholar
- Tse JM, Cheng G, Tyrrell JA, Wilcox-Adelman SA, Boucher Y, Jain RK, Munn LL: Mechanical compression drives cancer cells toward invasive phenotype. Proc Natl Acad Sci 2012,109(3):911–916.View ArticleGoogle Scholar
- Jiang X, Bruzewicz DA, Wong AP, Piel M, Whitesides GM: Directing cell migration with asymmetric micropatterns. Proc Natl Acad Sci 2005,102(4):975–978.View ArticleGoogle Scholar
- Brock A, Chang E, Ho C-C, LeDuc P, Jiang X, Whitesides GM, Ingber DE: Geometric determinants of directional cell motility revealed using microcontact printing. Langmuir 2003,19(5):1611–1617.View ArticleGoogle Scholar
- Duffy DC, McDonald JC, Schueller OJA, Whitesides GM: Rapid Prototyping of Microfluidic Systems in Poly(dimethylsiloxane). Anal Chem 1998,70(23):4974–4984.View ArticleGoogle Scholar
- Aratyn-Schaus Y, Oakes PW, Stricker J, Winter SP, Gardel ML: Preparation of complaint matrices for quantifying cellular contraction. J Vis Exp 2010, 46:2173.Google Scholar
- Crocker JC, Grier DG: Methods of digital video microscopy for colloidal studies. J Colloid Interface Sci 1996,179(1):298–310.View ArticleGoogle Scholar
- Sabass B, Gardel ML, Waterman CM, Schwarz US: High Resolution Traction Force Microscopy Based on Experimental and Computational Advances. Biophys J 2008,94(1):207–220.View ArticleGoogle Scholar
- Edwards CM, Schwarz US: Force localization in contracting cell layers. Phys Rev Lett 2011,107(12):128101.View ArticleGoogle Scholar
- Weijer CJ: Collective cell migration in development. J Cell Sci 2009,122(Pt 18):3215–3223.View ArticleGoogle Scholar
- Rørth P: Collective Cell Migration. Annu Rev Cell Dev Biol 2009,25(1):407–429.View ArticleGoogle Scholar
- Friedl P, Gilmour D: Collective cell migration in morphogenesis, regeneration and cancer. Nat Rev Mol Cell Biol 2009,10(7):445–457.View ArticleGoogle Scholar
- Tambe DT, Hardin CC, Angelini TE, Rajendran K, Park CY, Serra-Picamal X, Zhou EH, Zaman MH, Butler JP, Weitz DA, Fredberg JJ, Trepat X: Collective cell guidance by cooperative intercellular forces. Nat Mater 2011,10(6):469–475.View ArticleGoogle Scholar
- Kim JH, Serra-Picamal X, Tambe DT, Zhou EH, Park CY, Sadati M, Park J-A, Krishnan R, Gweon B, Millet E, Butler JP, Trepat X, Fredberg JJ: Propulsion and navigation within the advancing monolayer sheet. Nat Mater 2013,12(9):856–863.View ArticleGoogle Scholar
- Straight AF, Cheung A, Limouze J, Chen I, Westwood NJ, Sellers JR, Mitchison TJ: Dissecting temporal and spatial control of cytokinesis with a myosin II Inhibitor. Science 2003,299(5613):1743–1747.View ArticleGoogle Scholar
- Uehata M, Ishizaki T, Satoh H, Ono T, Kawahara T, Morishita T, Tamakawa H, Yamagami K, Inui J, Maekawa M, Narumiya S: Calcium sensitization of smooth muscle mediated by a Rho-associated protein kinase in hypertension. Nature 1997,389(6654):990–994.View ArticleGoogle Scholar
- Pelham RJ, Wang YL: Cell locomotion and focal adhesions are regulated by substrate flexibility. Proc Natl Acad Sci 1997,94(25):13661–13665.View ArticleGoogle Scholar
- Butler JP, Tolić-Nørrelykke IM, Fabry B, Fredberg JJ: Traction fields, moments, and strain energy that cells exert on their surroundings. Am J Physiol Cell Physiol 2002,282(3):C595-C605.View ArticleGoogle Scholar
- Berginski ME, Vitriol EA, Hahn KM, Gomez SM: High-resolution quantification of focal adhesion spatiotemporal dynamics in living cells. PLoS ONE 2011,6(7):e22025.View ArticleGoogle Scholar
- Tojkander S, Gateva G, Lappalainen P: Actin stress fibers - assembly, dynamics and biological roles. J Cell Sci 2012,125(8):1855–1864.View ArticleGoogle Scholar
- Bement WM, Mandato CA, Kirsch MN: Wound-induced assembly and closure of an actomyosin purse string in Xenopus oocytes. Curr Biol 1999,9(11):579–587.View ArticleGoogle Scholar
- Martin P, Lewis J: Actin cables and epidermal movement in embryonic wound healing. Nature 1992,360(6400):179–183.View ArticleGoogle Scholar
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