ARTICLES PUBLISHED ONLINE: 1 DECEMBER 2013 | DOI: 10.1038/NMAT3814

Epithelial bridges maintain tissue integrity during collective cell migration Sri Ram Krishna Vedula1† , Hiroaki Hirata1 , Mui Hoon Nai1 , Agustí Brugués2 , Yusuke Toyama1,3 , Xavier Trepat2 , Chwee Teck Lim1,4 * and Benoit Ladoux1,5 * The ability of skin to act as a barrier is primarily determined by the efficiency of skin cells to maintain and restore its continuity and integrity. In fact, during wound healing keratinocytes migrate collectively to maintain their cohesion despite heterogeneities in the extracellular matrix. Here, we show that monolayers of human keratinocytes migrating along functionalized micropatterned surfaces comprising alternating strips of extracellular matrix (fibronectin) and non-adherent polymer form suspended multicellular bridges over the non-adherent areas. The bridges are held together by intercellular adhesion and are subjected to considerable tension, as indicated by the presence of prominent actin bundles. We also show that a model based on force propagation through an elastic material reproduces the main features of bridge maintenance and tension distribution. Our findings suggest that multicellular bridges maintain tissue integrity during wound healing when cell–substrate interactions are weak and may prove helpful in the design of artificial scaffolds for skin regeneration.

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ound healing is a fundamental biological process that involves an overlapping cascade of events such as inflammation, tissue formation and tissue remodelling1 . Despite the common function of re-establishing tissue integrity, wound healing can vary depending on the context in which the process occurs. For instance, wound healing in embryos is associated with rapid tissue repair and results in scarless restoration of normal tissue. In contrast, wound healing in adults is slower and often results in scarring2 . An in-depth understanding of the various factors regulating wound healing is hence necessary to gain a better insight into tissue repair and regeneration3 . Such an insight could also pave the way for designing better alternatives that can overcome the present limitations in the field of skin tissue engineering for promoting satisfactory skin regeneration, for example in the treatment of burns4 . A critical component of the wound healing response is re-epithelialization, wherein directed collective migration of keratinocytes restores the continuity of the epidermal barrier5 . Defects in re-epithelialization are associated with chronic nonhealing wounds, such as venous stasis, diabetic and pressure ulcers6 . During re-epithelialization, collectively migrating keratinocytes encounter an extracellular matrix (ECM) that is heterogeneously distributed within the wound bed. Previous studies suggest that collective cell movements are affected by the physical properties of their ECM environment as they migrate through heterogeneous tissues in terms of topography, porosity and stiffness7 . Such heterogeneities in the spatial distribution of ECM offer two distinct micromechanical challenges for migrating cells. First, cells are forced to migrate under a wide range of geometrical constraints depending on the biological process involved. For example, ‘chains’ of cancer cells invading the ECM during metastasis are subjected to highly confined geometries8 . The second important

hurdle that migrating cells have to overcome is to progress over regions that are spatially discontinuous in terms of the distribution of ECM proteins9 . Such non-adhesive regions do not provide cells with stable ‘foot-holds’ for propelling themselves forward. How cells respond to these two challenges has been well explored in the context of single cells10,11 . However, the migratory behaviour of a single epithelial cell is very different from that of a cell within a monolayer12–15 owing to the role of intercellular adhesion, a key regulator of many biological processes in epithelial cells16,17 . Hence, it is not only difficult but in many instances erroneous to extrapolate single-cell responses to multicellular systems. In the context of the collective migration of epithelial skin cells, it seems particularly important to understand how these cells respond to external physical constraints, migrate over heteregeneous ECM, and integrate the resulting mechanical tension over multiple cells to maintain tissue integrity and restore a continuous epithelial cell sheet. We have recently developed an in vitro migration assay using microcontact printing (µCP) that allows us to systematically investigate the influence of geometrical constraints on collective cell migration15 . Here, using a similar approach, we have studied the collective behaviour of human keratinocyte cell sheets migrating on micropatterned geometries comprising alternating adherent and non-adherent strips. We show that they form pluricellular suspended epithelial ‘bridges’ over non-adhesive regions. Even though different types of intercellular bridge have been described previously in a variety of cell types18 including cytonemes, tunnelling nanotubes and tubular epithelial bridges19–21 , the architecture and mechanism of formation of epithelial bridges presented here are significantly different from the ones described in previous studies. These multicellular structures require the build-up of large-scale tension driven by actomyosin contractility and its

1 Mechanobiology

Institute, National University of Singapore, 117411, Singapore, 2 Institut de Bioenginyeria de Catalunya (IBEC), ICREA, and Facultat de Medicina—Universitat de Barcelona, 08028 Barcelona, Spain, 3 Department of Biological Sciences, National University of Singapore and Temasek Life Sciences Laboratory, 117543, Singapore, 4 Department of Biomedical Engineering and Department of Mechanical Engineering, National University of Singapore, 117576, Singapore, 5 Institut Jacques Monod (IJM), CNRS UMR 7592 and Université Paris Diderot, Paris 75013, France. † Present address: L’oreal Research and Innovation, #06-06, Neuros, 8A Biomedical Grove, 138648, Singapore. *e-mail: [email protected]; [email protected] NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

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Figure 1 | Keratinocytes migrating on microcontact-printed fibronectin patterns form multicellular suspended epithelial bridges. a, Schematic outlining the in vitro migration assay on fibronectin patterns. Details are provided in the main text. b, Keratinocytes migrating from the reservoir (RES) into the 10-µm-wide fibronectin strips form multicellular suspended epithelial bridges over the non-adhesive regions between the fibronectin strips. c, MDCK epithelial cells do not form well-developed epithelial bridges but some ‘nascent’ bridge-like structures can be observed at the junction of the fibronectin strip and the reservoir (arrow). d, Basal confocal sections of epithelial bridges immunostained for paxillin (top panel) and phospho-paxillin (bottom panel) showing focal adhesions (arrows). The inset in the top panel (magenta) shows F-actin in the region bounded by the red square. White dotted lines represent the concave edge of the epithelial bridge. e, Schematic showing patterned three-dimensional PDMS substrates with raised features (∼13 µm from surface) stamped with fibronectin (red). f, SEM image of keratinocytes showing the formation of suspended epithelial bridges between raised PDMS ridges. The inset shows a magnified view of the epithelial-bridge region enclosed by the red square. g, ‘End-on’ view of an epithelial bridge on the PDMS ridges. h, Formation of a ‘hole’ (arrow) in the epithelial bridge during cell division. Scale bars, 50 µm (b,c,h) and 20 µm (d,f,g).

transmission through bundles of actin connected to intercellular adhesion sites, especially adherens junction proteins.

Formation of epithelial bridges Migrating keratinocyte monolayers form suspended multicellular bridges on patterned substrates. A schematic of the experimental set-up is shown in Fig. 1a. The fibronectin pattern used for microcontact printing consists of a large rectangular ‘reservoir’ joined to ∼10-µm-wide strips separated by ∼120 µm. Initially, a block of polydimethyl siloxane (PDMS) was placed over the pattern 88

allowing the HaCaT cells to form a monolayer that was confined to the ‘reservoir’ region of the pattern (Fig. 1a, refer to ref. 15 for details). After the removal of the PDMS barrier, cells at the leading front of the monolayer migrated into the fibronectin strips. Surprisingly, as the cells migrated into the strips, the monolayer also advanced over non-adhesive regions in between the strips (Fig. 1b and Supplementary Video 1). As cells cannot adhere to the non-adhesive regions between the fibronectin strips, we reasoned that the monolayer was suspended between adjacent fibronectin strips forming ‘epithelial bridges’. Accumulating evidence suggested NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

NATURE MATERIALS DOI: 10.1038/NMAT3814 that keratinocytes formed suspended cell layers above the substrate. At first, we confirmed that cells did not escape from the other sides of the reservoir (those that were not continuous with the strips) within similar timescales to the ones observed for the formation of the epithelial bridges, suggesting that neither ECM produced by the cells nor the proliferative pressure from within the epithelial monolayer could explain the presence of cells over areas devoid of fibronectin. Then, we observed that the migration of another epithelial cell line (MDCK) on similar patterns did not exhibit any epithelial bridges even though small ‘fin’-like cellular extensions could be observed at the base of the fibronectin strips (arrow in Fig. 1c and Supplementary Video 2). Basal confocal sections of epithelial bridges immunostained for paxillin (Fig. 1d, top panel), phospho-paxillin (Fig. 1d, bottom panel) and α5 β1 integrin (Supplementary Fig. 1a) showed that focal adhesions were largely restricted to the fibronectin pattern, suggesting that epithelial bridges were not adhering to the substrate. Final evidence came from scanning electron microscopy (SEM) images of cell monolayers that were allowed to advance over patterned PDMS substrates (stamps that were used for µCP, Fig. 1e). In contrast to µCP Petri dishes, the patterns (reservoir and the lines) on these three-dimensional substrates are raised (∼13 µm high) from the surface. Fibronectin was deposited only on these raised features using a flat block of fibronectin-coated PDMS (Fig. 1e). SEM images clearly showed that the advancing cell monolayer was indeed suspended between the fibronectin-coated ridges (Fig. 1f,g). During the course of migration, we occasionally observed the formation of ‘holes’ within the cell sheet as a consequence of cell division (arrows in Fig. 1h and Supplementary Video 1). The integrity of a freely migrating monolayer (Methods) remained intact and no ‘holes’ were observed during the entire period of observation. The appearance of such ‘holes’ at the particular areas not only confirms the formation of epithelial bridges but also indicates that those structures are submitted to high tension. Moreover, the morphology of the epithelial bridges is also highly reminiscent of the concave actin-based structures formed by single cells adhering to fibronectin patterns that have been shown to be highly contractile in nature9,22,23 . Here, because the spacing between the fibronectin strips in our experimental setup (∼120 µm) was much larger than the size of individual cells (∼30 µm), the epithelial bridges were formed by multiple cells spanning across adjacent fibronectin strips, and thus the tension should be transmitted through neighbouring cells. Taken together, these findings demonstrate that epithelial bridges are pluricellular units suspended between the fibronectin strips and could be sustained by actin contractility. The formation of epithelial bridges relies on the pulling forces exerted by migrating cells. We first reasoned that the formation of epithelial bridges was driven by the pulling forces exerted by cells migrating on the fibronectin strips. Thus, we analysed the overall migration speed of the leading edge. First, we compared the migration characteristics of keratinocytes on micropatterns with the migration of a monolayer in a free geometry (Methods and Supplementary Video 3). Under such conditions, the leading cells confined to the fibronectin strips migrated slower than the monolayer (Fig. 2a,b). Moreover, whereas the displacement of the monolayer showed a continuous and linear progression with time (Fig. 2a,b, blue), the displacement of cells on the fibronectin strips reached a plateau within the same period of observation (∼48 h; Fig. 2a,b, red). Our results are contrary to previous studies done on single human epidermal keratinocytes10 as well as collectively migrating MDCK epithelial cell sheets15 , where it has been observed that confined geometries increase the overall migration speed of cells. This suggests that cells on fibronectin patterns pull on cells forming the epithelial bridges, which in turn slow down NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

ARTICLES their migration up to a saturation regime. These pulling forces are thus balanced by the cellular resistive forces at the back, which leads to a high mechanical tension within the epithelial bridges transmitted over multiple cells. We also observed a reduced displacement of epithelial bridges relative to the adherent cells along the strips (Fig. 2a,b, black). However, they both reached a plateau after roughly 30 h (Fig. 2a,b). Consequently, we then analysed the influence of geometry on the formation and maintenance of the epithelial bridges. First, we varied the spacing between the fibronectin strips to 200 and 400 µm while keeping the width of the fibronectin strip constant (∼10 µm, Fig. 2c,d). Interestingly, under such conditions, the migration of cells onto the fibronectin strips was limited to short distances from the reservoir (Fig. 2a, green and Supplementary Video 4). This suggests that there exists a critical length scale (∼200 µm) at which the formation of epithelial bridges is inhibited. However, cells on the strips pulled a small portion of the cell sheet along with them, leading to the formation of ‘fin’-like bridge structures (arrow in Fig. 2c,d, right panel) at the junction of the fibronectin strip with the reservoir. Then we checked whether epithelial-bridge formation depends on the number of pulling cells. To do so, we increased the width of the fibronectin strips up to 250 µm (from the original 10 µm) keeping the same spacing between the strips (∼120 µm, Fig. 2e). We found that migration of epithelial bridges as well as cells on the strips increased considerably (Fig. 2f,g, red and black, and Supplementary Video 5). Together, these results confirm our hypothesis that epithelial bridges form as a result of tensile forces transmitted from the pulling cells on the fibronectin strips.

Integrity of epithelial bridges Epithelial bridges are under tension. Our previous observations suggest that epithelial bridges are subjected to tension. Studies on single cells plated on micropatterned ECM dots suggest that the relationship between the curvature of membrane bridges and tension within them is influenced by the spacing between the anchor points22 . We found that the curvature (Methods) of the concave leading front of the multicellular epithelial bridge quickly increased when the cell sheet migrated from the reservoir into the region between the fibronectin lines (Fig. 3a). Once formed, the curvature fluctuated over time but otherwise remained fairly constant, suggesting that the epithelial bridge was indeed under continuous tension. Previous studies have shown that many types of cell remodel their actin network and develop well-formed stress fibres when subjected to external mechanical tension24–26 . We reasoned that if the epithelial bridges were indeed under tension, the actin network within the cells constituting the epithelial bridges would undergo significant remodelling. Staining with phalloidin revealed a network of F-actin bundles suspended between the two fibronectin strips (Fig. 3b). Moreover, the cell sheet seemed thinner (in the z-direction) at the concave leading front (Fig. 3c, top panel) compared with the region of epithelial bridge farther away from the leading front (Fig. 3c, bottom panel). Cells at the concave leading edge of the epithelial bridge were elongated (whereas those away from the edge of the epithelial bridge were more isotropic in shape and direction), as shown by nucleus staining as an indicator of cell orientation27 (arrows, Fig. 3d). This suggests that the concave front was probably under much more tension than the rest of the epithelial bridge. Basal confocal sections also showed thick bundles of actin traversing multiple cells within the body of the epithelial bridges (arrows, Fig. 3e). Cells within the epithelial bridges cannot form focal adhesions, so we reasoned that the observed bundles of actin were anchored to intercellular adhesion sites. As adherens junctions (of which E-cadherin is an important member) are known to play a key role in the formation and maintenance of intercellular adhesion, cells were co-immunostained for E89

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Figure 2 | Migration dynamics of keratinocytes under different geometrical confinements. a, Displacement with time of a keratinocyte monolayer (blue) compared with that of epithelial bridges (black) and cells on 10-µm-wide fibronectin strips separated by either 120 µm (red) or 400 µm (green). Displacement of cells on the fibronectin strips and displacement of epithelial bridges show saturation towards the end of the observation period. b, The velocity of the monolayer is higher than the velocity of epithelial bridges and cells on fibronectin strips. c,d, Cells migrating on 10-µm-wide strips that are spaced 200 µm (c) or 400 µm (d) apart form small ‘fin’-shaped bridges (arrow, right panel). e, Cells migrating on 250-µm-wide fibronectin strips that are spaced 120 µm apart form epithelial bridges. f,g, Displacement (f) and velocity (g) of epithelial bridges (black) and cells on the 250-µm-wide fibronectin strips (red) are higher than those of cells on 10-µm-wide fibronectin strips. Error bars show standard error of mean. Scale bars, 50 µm (c,d,e).

cadherins (Fig. 3f) and actin (Fig. 3g). Interestingly, in the epithelial bridges three distinct staining patterns of E-cadherins were observed depending on their location in the z-plane. In basal confocal sections, a ‘stretched’ or ‘punctate’ staining morphology oriented perpendicular to the intercellular junction was observed (arrow, Fig. 3f). Bundles of F-actin were found to be concentrated in close proximity to these ‘stretched’ junctions (arrow, Fig. 3f, inset). Intermediate confocal sections showed a ‘linear’ 90

morphology oriented parallel to the intercellular junction (arrow, Supplementary Fig. 1b, left panel). These linear E-cadherin junctions co-localized with cortical actin (arrow, Supplementary Fig. 1b, right panel). Apical confocal sections showed a slightly ‘stretched’ staining morphology (arrow, Supplementary Fig. 1c, left panel) and some actin bundles (arrow, Supplementary Fig. 1c, right panel) although they were not as obvious as those observed in the basal regions. Previous studies have shown that the NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

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Figure 3 | Epithelial bridges are subjected to considerable tension. a, Change in the curvature of the leading edge of epithelial bridges with time. Error bars represent standard error of mean. b, Z-projection image of staining for actin shows a network of F-actin bundles suspended between the fibronectin strips (dashed red lines). c, The upper panel shows an x–z section of the leading edge of the epithelial bridge (along the solid white line in b), and the lower panel shows an x–z section of the epithelial bridge away from the leading edge (along the dashed white line in b). Red lines represent fibronectin strips. d, Nuclei at the leading edge are elongated along the curvature of the epithelial bridge (arrows). e, Basal confocal section of epithelial bridges shows actin stress fibres traversing multiple cells (arrows). f,g, Basal confocal sections of epithelial bridges co-immunostained for E-cadherin (f) and actin (g) reveal the stretched morphology of E-cadherins (arrows). Inset: magnified view of the epithelial bridge enclosed by the white rectangle showing actin bundles (magenta) inserting into E-cadherins. Scale bars, 10 µm (c), 20 µm (b,d–g).

‘stretched’ staining pattern of E-cadherins resulted from myosincontractility-mediated tensile forces acting on adherens junctions through actin filaments28,29 . In contrast to epithelial bridges, cells in the reservoir showed a linear staining pattern of E-cadherins in the basal sections (arrow, Supplementary Fig. 2a, left panel). Furthermore, although actin stress fibres were observed in the basal regions (arrow, Supplementary Fig. 2a, right panel), unlike within epithelial bridges they neither concentrated close to E-cadherins nor spanned multiple cells. This suggested that these actin stress fibres were most probably inserting into focal adhesions. The staining morphology of E-cadherin and actin in the apical and intermediate regions of the intercellular adhesion, on the other hand, was similar to that observed in the epithelial bridges (Supplementary Fig. 2b,c). Again, these observations point out that epithelial bridges are being subjected to constant tension, presumably by forces transmitted from cells on the fibronectin strips through the actin bundles and adherens junctions. Myosin-II-mediated contractility is required for the formation of epithelial bridges. We studied the influence of myosinII-mediated contractility on the formation and maintenance of epithelial bridges. We first inhibited myosin II by adding 50 µM blebbistatin to the culture medium immediately after NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

the removal of the PDMS barrier. Despite cells migrating into the fibronectin strips, epithelial-bridge formation was markedly inhibited, suggesting that myosin contractility was indeed required for the formation of epithelial bridges (Supplementary Fig. 3a and Video 6). We also noted that blebbistatin significantly altered cell morphology. Cells were more spread out and showed enhanced membrane ruffling (Supplementary Video 6). In the second set of experiments, blebbistatin was added to the culture medium after the formation of epithelial bridges (∼48 h after allowing the cells to migrate into the strips). We found that relaxation induced by blebbistatin was most noticeable at the concave leading edge of the epithelial bridge (Supplementary Fig. 3b and Video 7), further supporting our previous observation that the edge of the bridge was indeed under much higher tension than the rest of the epithelial bridge. Staining for actin showed destruction of the actin cables within the epithelial bridge (Supplementary Fig. 3c) as well as thickening of the epithelial bridge (in the z-direction, Supplementary Fig. 3d) when compared with the untreated bridges (Fig. 3c). To further investigate how inhibition of myosin II contractility affected the staining pattern of actin and E-cadherins within epithelial bridges, we co-immunostained cells for E-cadherins and actin after treating the epithelial bridges with 50 µM blebbistatin for 1 h. Basal confocal sections showed complete destruction of actin bundles in epithelial bridges as well as on the 91

ARTICLES strips and reservoir regions (Supplementary Fig. 4). Interestingly, whereas the ‘stretched’ morphology of E-cadherins (observed in the basal section of epithelial bridges and apical sections throughout the monolayer) was disrupted, the ‘linear’ E-cadherins comprising the intermediate region of the adherens junctions remained largely intact in epithelial bridges as well as over the fibronectin strips and reservoir. These results suggest that the ‘stretched’ morphology of E-cadherins is indeed a result of force exerted on the junctions through the actin filaments mediated by myosin II contractility. However, contrary to previous observations on single cells (where inhibition of myosin II led to a complete collapse of the bridge regions onto the fibronectin strips)9 , the integrity of the epithelial bridges remained largely unperturbed. Taken together, these results suggested that whereas myosin contractility was required for the formation of epithelial bridges, the overall integrity of the bridges seems to be largely regulated by the intermediate pool of Ecadherins that form stable ‘linear’ junctions. Role of adherens junctions in the formation and maintenance of epithelial bridges. We further reasoned that the integrity of epithelial bridges is regulated by a balance between the tension within the bridge and the ability of the intercellular adhesion complex to withstand high tension. To test this, we modified the strength of intercellular adhesion and tension within the bridge using lowCa2+ medium and calyculin A, respectively. Cell migration as well as epithelial-bridge formation was inhibited when cells were cultured and allowed to migrate in low-Ca2+ medium (Supplementary Fig. 5a and Video 8). On the other hand, switching to low-Ca2+ medium after the formation of epithelial bridges resulted in relaxation of the bridges (Supplementary Fig. 5b and Video 9). Increasing the tension within the epithelial bridges by addition of 20 nM calyculin A (a phosphatase inhibitor that increases myosin contractility) resulted in complete disintegration of the cell sheet (Supplementary Video 10). However, epithelial bridges remained intact in the presence of low-dose (1 nM) calyculin A (Supplementary Video 11). Furthermore, switching to low-Ca2+ medium containing 1 nM calyculin resulted in the formation of holes within epithelial bridges (arrow in Supplementary Fig. 5c, right panel and Supplementary Video 12). Together, these results suggest that although the intercellular adhesion is significantly weakened in low-Ca2+ medium, it is still able to maintain the integrity of the epithelial bridges. However, subjecting the weakened intercellular adhesion to increased tension can lead to disruption of the intercellular adhesion, leading to collapse of the epithelial bridges. Although these experiments strongly point towards the role of adherens junctions in the formation and maintenance of epithelial bridges, the wide range of effects that Ca2+ has on cellular processes precluded us from directly correlating the importance of adherens junctions in the formation of epithelial bridges under these conditions. To specifically investigate the role of adherens junctions in the formation of epithelial bridges, we generated HaCaT cells in which α-catenin was stably knocked down (α-HaCaT cells) by introducing short hairpin RNA (shRNA). The knockdown was confirmed by western blotting (Supplementary Fig. 5d). α-catenin is a well-established cytoplasmic adaptor molecule localizing at adherens junctions that has recently been shown to act as a mechanosensor30 . Knockdown of α-catenin has been shown to have a profound effect on the strength of adherens junctions. As observed in other epithelial cell types31 , knockdown of α-catenin resulted in loss of the ability of HaCaT cells to migrate collectively in a cohesive manner. In contrast to HaCaT cells expressing shRNA with a scrambled sequence that formed epithelial bridges (arrow, Supplementary Fig. 5e), α-HaCaT cells did not migrate collectively and were unable to form epithelial bridges when allowed to migrate on the fibronectin patterns (Supplementary Fig. 5f and Video 13). Instead, they formed chains of loosely adherent cells that migrated 92

NATURE MATERIALS DOI: 10.1038/NMAT3814 in an individualistic manner (Supplementary Fig. 5f). In several instances, single cells stretched across non-adhesive regions (either between two adjacent fibronectin strips or between the reservoir and a fibronectin strip forming long tethers (arrow, Supplementary Fig. 5f). These tethers are highly reminiscent of tunnelling nanotubes described previously19 . It is highly likely that such ‘singlecell bridges’ lay the foundation for the development of elaborate and complicated multicellular epithelial bridges. However, absence of strong intercellular cohesiveness in the α-HaCaT cells probably prevents the maturation of such nascent epithelial bridges. Together, these results suggest that adherens junctions are essential for the formation as well as integrity of epithelial bridges. Epithelial bridges exhibit elastic-like behaviour. On the basis of our experimental study, accumulating evidence favours a mechanism based on the pulling forces exerted by cells adhering on the fibronectin strips that promote the bridge formation. This situation is reminiscent of physical systems where localized forces induce the distortion of a deformable object such as a liquid contact line32 or an elastic membrane33 . As the presence of actin bundles and the ‘stretched’ staining pattern of E-cadherins provided evidence for tension within the epithelial bridges, we assumed that our system could be described by a longitudinal deformation of an elastic membrane by a point force acting perpendicular to the membrane edge (Fig. 4a). At equilibrium, the deformation of the membrane along the fibronectin strip (uy ) and perpendicular to the strip (ux ; ref. 33) is given by:    F 1−υ ux = πE e      1+υ F F  x uy = +2 ln πE e πEe a where F represents a localized force normal to the edge, υ is Poisson’s ratio, E is Young’s modulus, e is the thickness of the membrane or the monolayer and a is a cutoff length. This simple mechanical model captures the observed mechanism. The profile of the bridge as well as the ‘fin’ formation by cells migrating into the 10 µm strips spaced 400 µm apart indeed corresponds to a logarithmic profile (Fig. 4b,c). Indentation studies performed on an atomic force microscope using a spherical bead (Methods) suggest that Young’s modulus of a monolayer of HaCaT cells is 2 ± 0.7 kPa, in agreement with previous studies34 . However, reported values in the literature for keratinocytes vary considerably from 2 to 100 kPa (refs 35, 36). Considering an average thickness of the epithelial bridges as ∼3 µm (ref. 35; Fig. 3c), our model predicts the forces exerted by cells on the strips to be ∼250 nN. This model not only confirms that epithelial bridges are indeed subjected to tensile forces transmitted from pulling cells on the fibronectin strips, but also estimates such forces. The force values given by our model were confirmed by traction force microscopy37 (TFM). Large traction forces directed downwards towards the reservoir were observed at the leading-cell front on the fibronectin strips (Fig. 4d, inset). Stresses of the order of ∼1 kPa were registered at the leading-cell front (Fig. 4d, white arrows). Considering that these regions of high stresses have an average area of ∼300 µm2 , the estimated force exerted by the leading-cell front is of the order of ∼300 nN, which is in good agreement with the values predicted by our elastic materiallike model and those previously reported for keratinocyte cell clusters38 . We then tested our model based on the treatment of epithelial bridges as elastic materials by performing laser ablation studies on well-formed bridges. In contrast to previous ablation studies performed on finger-like structures in migrating NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

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Figure 4 | Modelling epithelial-bridge formation as deformation of an elastic membrane. a, Schematic of the deformation of an elastic membrane by localized force acting normal to its edge. b, Contour of the ‘fin’-shaped bridges (red line) formed by cells migrating on 10-µm-wide fibronectin strips that are separated by 400 µm. c, Representative graph showing a logarithmic fit of the contour. d, Colour-coded quiver plot showing the direction and magnitude of stresses exerted by cells migrating on 10-µm-wide fibronectin strips. Inset: magnified view of the region bounded by the white rectangle. e, Laser ablation of the leading-cell front on the fibronectin strip (arrow, left panel) results in immediate elastic recoil of the epithelial bridge (right panel). Black dotted lines are a guide to the eye to show the position of the cell front on the fibronectin strip before and immediately after laser ablation. 1y represents the immediate displacement of the cell front after laser ablation. f, A kymograph along the blue line shown in e. The red dashed line traces the edge of the cell front on the fibronectin strip to guide the eye. Scale bars, 50 µm in d,e and 1 s in f.

monolayers (where the ablated edges retracted in 15–30 min; ref. 39), ablation of the leading-cell front on the fibronectin strip was associated with immediate relaxation of epithelial bridges, confirming their predominantly elastic nature as well as the existence of tension within them (Fig. 4e and Supplementary Video 14). The rapid relaxation is evident from the kymograph taken along the length of the fibronectin strip (Fig. 4f). Furthermore, the elastic recoil of the cell sheet along the length of the fibronectin strip (1y, Fig. 4e,f) immediately after laser ablation was found to be 38.1 ± 6.7 µm (n = 4 bridges). Assuming the value for cell-sheet stiffness, ∼6 mN m−1 , used before (2 kPa by 3 µm), the forces exerted by the leading-cell front on the fibronectin strip equal ∼230 nN, which matches well with the force predicted by our model. On the other hand, epithelial bridges pretreated with blebbistatin (50 µM for ∼1 h before ablation) did not show significant recoil, confirming that myosin-II-mediated contractility is indeed necessary for maintaining tension within the epithelial bridges (Supplementary Video 15). We thus reasoned that the formation of epithelial bridges should reflect the mechanical properties of the cell sheets and, in particular, how strong cells within these epithelial sheets are connected to one another by intercellular junctions to maintain the integrity of the tissue. Interestingly, such behaviour is shared by primary human keratinocytes (Fig. 5a, left panel) and corneal epithelial cells (Fig. 5a, right panel) but not by other cell types such as 3T3 fibroblasts40 and simple epithelial cells (for example MDCK). Consequently, we NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

mapped the velocity field (Fig. 5b) and its vorticity (Fig. 5c) using particle image velocimetry (PIV) to characterize tissue fluidity by cell–cell rearrangements for both HaCaT and MDCK cell sheets. We indeed observed that keratinocytes exhibited homogeneous distributions of the velocity field (Fig. 5b, right panel) and vorticity (Fig. 5c, right panel), whereas MDCK cells showed heterogeneous landscapes for both velocity and vorticity even far away from the leading edge of the monolayer (Fig. 5b,c, left panels). These differences also were reflected in the traction-force fields of migrating MDCK and HaCaT cell sheets: HaCaT cells exhibited large traction forces at the leading front with a homogeneous distribution of the forces far away from the edge (Fig. 5d, right panel), whereas MDCK cells showed a rough distribution of the forces even far behind the leading front (Fig. 5d, left panel)13 . These results confirm that keratinocytes presented a more elastic-like behaviour than MDCK, allowing the formation of epithelial bridges in good agreement with our model. Keratinocytes migrating on fibronectin strips generate pulling forces that lead to the buildup of high tension transmitted to cells in the reservoir through intercellular contacts within the epithelial bridges. In contrast, the more fluid-like behaviour of MDCK induces a dissipation of the tension exerted by leading cells through cell–cell rearrangements.

Outlook In contrast to the classical view of wound closure and collective cell migration, we demonstrate the ability of epithelial skin 93

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Figure 5 | Comparison of cell–cell rearrangements within HaCaT and MDCK monolayers using PIV. a, Epithelial bridges are also formed by primary human skin keratinocytes (left) and corneal epithelial cells (right) migrating on micropatterned substrates. b, Velocity fields within migrating HaCaT (right) and MDCK (left) monolayers obtained using PIV. c, Heat map of angular velocity (rad h−1 ) within HaCaT (right) and MDCK (left) showing higher velocity gradients in MDCK monolayers. d, Heat map of the stress landscape of migrating MDCK (left) and HaCaT (right) monolayers obtained using traction force microscopy. All scale bars, 50 µm.

cells to form unanticipated multicellular structures that bridge non-adhesive areas. As these cells are specialized in wound repair, the physiological relevance of epithelial-bridge formation clearly seems to be a means to ensure proper tissue integrity and restore a homogeneous epithelial barrier regardless of the ECM conditions. These pluricellular structures require force transmission and coordination over length scales much larger than individual cells, and probably involve a large-scale mechanosensing mechanism mediated by actin cytoskeleton41 . During wound healing, keratinocytes are faced with the challenge of migrating over regions devoid of ECM. The ability of keratinocytes to form 94

epithelial bridges could help them to advance over regions devoid of ECM, resulting in accelerated wound closure. It is noteworthy that epithelial-bridge formation has indeed been observed in vivo and ex vivo during wound healing of epidermis and corneal epithelium42–44 . Recently, such bridges have also been described by other groups in the context of generating engineered skin tissue45 . The concave edge of the epithelial bridges is highly reminiscent of actin purse-string-mediated re-epithelialization observed in chick embryos46 . Indeed, it has been suggested that the mechanism of re-epithelialization during wound healing (whether lamellipodial crawling or contraction of actin cable) is regulated by distribution NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

NATURE MATERIALS DOI: 10.1038/NMAT3814 of ECM (ref. 47). Interestingly, decreased cell–substrate adhesion (due to hypomorphic mutations in specific integrins) results in separation of apposed epidermal sheets and formation of blisters in wings of Drosophila48,49 . Similarly, the skin of mice deficient in specific integrins shows disorganized matrix distribution and blisters at the dermal–epidermal junction50 . In light of our results, these studies not only provide strong evidence for the existence of epithelial bridges in vivo but also suggest that the physiological function of these structures is to maintain tissue integrity when the cell–substrate interactions are either absent or poorly developed. In the context of tissue engineering and repair, skin substitutes represent a prospective source of advanced therapy in combating severe skin wounds. At present, bioengineered skin models do not fully replicate the natural properties of uninjured skin. To restore the skin barrier, skin cells must migrate collectively through heterogeneous ECM scaffolds, and are then subjected to high mechanical tension. Our findings provide unanticipated modes of collective cell migration to preserve tissue integrity during wound healing and could help in designing artificial scaffolds for skin models. As formation and maintenance of epithelial bridges seems to be tightly coupled to various cell- and tissue-level variables such as intercellular adhesion, force generation and transmission, ECM geometry and possibly the inherent stiffness of the cell monolayer, they can be used for assessing changes in these variables in high-throughput assays. Such assays could also have potential applications in investigating changes in the mechanical properties of epithelial cells in various diseases and during ageing.

Methods Cell culture and reagents. HaCaT cells (spontaneously immortalized primary human keratinocytes) were purchased from Cell Lines Service. They were maintained in DMEM supplemented with 10% fetal bovine serum and antibiotics. Early passage cells (∼10 passages) were used for the experiments. Primary human keratinocytes (kindly provided by the Advanced Skin Research Centre, L’oreal, Singapore) and corneal epithelial cells (HCEC, Life Technologies) were maintained in keratinocyte serum-free medium. They were trypsinized and re-suspended in normal culture medium before seeding onto the patterns. Low-calcium medium (∼0.03 mM) was prepared by adding 1% fetal bovine serum and antibiotics to calcium-free DMEM (Sigma). Blebbistatin (Tocris) and cytochalasin D (Sigma) were dissolved in dimethylsulphoxide before diluting into culture medium. Antibodies against α-catenin and β-actin were purchased from Sigma Chemical. Generation of α-catenin knockdown HaCaT cells. To generate retroviruses expressing shRNA specific to human catenin alpha1, the target sequence 50 -GACTTAGGAATCCAGTATA-30 was inserted into the pSUPER.puro retroviral vector. For control, a scrambled sequence 50 -ATAGTCACAGACATTAGGT-30 was introduced. The shRNA-containing vector was co-transfected with the pE-ampho vector into HEK293T cells using the GeneJuice transfection reagent (Novagen). Supernatants containing viral particles were collected 48 h after the transfection, filtered through 0.45-µm syringe filters, and used for infection into HaCaT cells in the presence of 8 µg ml−1 Polybrene (Sigma). Infected HaCaT cells were selected with 1.5 µg ml−1 puromycin. Silencing of α-catenin expression was confirmed by western blotting (Supplementary Fig. 5d). Microcontact printing and model wound assay. Wafers containing the patterns were fabricated using photolithographic techniques as described previously51 . The master wafers were silanized and used for preparing PDMS stamps. Briefly, PDMS (Sylgard 184, Dow Corning) was mixed in a ratio of 1:10, degassed and poured over the wafers. After curing at 80 ◦ C for 2 h, stamps were peeled off. A mixture of fibronectin and a small amount of Cy3-conjugated fibronectin was used to ink the stamps. For migration studies, microcontact printing (µCP) was performed on plastic Petri dishes (non-culture-treated, Greiner). For characterizing the migration of a keratinocyte monolayer, a model wound assay was used. A thick PDMS slab was cast in a Petri dish and a large rectangular well (∼3 × 1.5 cm) was cut with a sharp scalpel such that the Petri dish was exposed at the base of the PDMS well. The Petri dish was plasma-cleaned and a large drop of fibronectin was added inside the well to coat the base of the well. After one hour of incubation, the rest of the PDMS was blocked with 0.2% Pluronics (Sigma). After washing thoroughly with phosphate-buffered saline (PBS), a portion of the well was blocked with a PDMS slab, and cells were seeded in the exposed region of the well. After reaching confluence, the PDMS slab was removed, allowing the cells to migrate into the fibronectin-coated free space. NATURE MATERIALS | VOL 13 | JANUARY 2014 | www.nature.com/naturematerials

ARTICLES Live-cell imaging and analysis. PDMS blocks were gently removed when cells reached confluence in the reservoir. The dishes were rinsed gently with culture medium to remove any floating cells and debris, and fresh medium was added before imaging. For live-cell imaging, images were acquired (every 4 min for migration studies and every 1 min for drug treatment studies) using a ×10 objective on a Biostation (Nikon) at 37 ◦ C and 5% humidified CO2 . The Biostation uses red light illumination for imaging, eliminating the phototoxicity of blebbistatin. Analysis of images was performed using ImageJ. For computing the curvature, a central strip of 50 µm of the epithelial bridge was fitted to a circle using the three-point ROI plug-in for image J. The curvature was defined as the inverse of the radius of the fitted circle. PIV analysis was performed using the MatPIV package52 implemented in MATLAB. Velocity gradients were calculated using the curl function in MATLAB as described previously15 . Immunofluorescence staining. For confocal imaging of immunofluorescence-stained samples, patterns were printed on a thin layer of PDMS that was spin-coated over a glass-bottom Petri dish (Iwaki). After the formation of bridges, cells were fixed in 4% PFA, and permeabilized in 0.1% Triton X-100. For staining of actin alone, cells were incubated with Alexa-488-labelled phalloidin for 30 min. For co-staining E-cadherins and actin, following fixation and permeabilization, cells were incubated in blocking solution (3% BSA in 0.1% Triton X-100) for 1 h. They were then incubated in mouse anti-E-cadherin antibody (1:100 diluted in blocking solution) for 1 h, washed with PBS and incubated in Alexa-488-labelled goat anti-mouse antibody. Finally, they were incubated in Alexa-634-labelled phalloidin and DAPI (1 µg ml−1 ) for 30 and 5 min, respectively, and washed with PBS. Immunostaining for paxillin and phospho-paxillin was performed using a similar protocol. Samples were imaged while immersed in PBS to prevent any mounting- or drying-induced artefacts. Confocal sections (1 µm thick) were acquired on a Nikon (C1si) or Leica (SP5) using either a ×60 or ×100 objective and processed in ImageJ to enhance contrast. SEM. Cells were seeded on patterned PDMS stamps that were used for µCP. In contrast to µCP Petri dishes, the patterns (reservoir and the lines) in this case were raised (∼13 µm high) from the surface. Fibronectin was microcontact printed only on these raised features using a flat block of fibronectin-coated PDMS. After the formation of epithelial bridges, cells were fixed in 3% glutaraldehyde for 10 min, washed in PBS and dehydrated in increasing concentrations of alcohol (50, 70, 90 and 100% ethyl alcohol for 10 min each). The alcohol was drained off and the sample was critical point dried. Finally, the samples were sputter-coated with gold and imaged using a scanning electron microscope (Jeol). TFM. Soft silicone elastomer substrates for TFM were prepared as described previously with some modifications53 . CyA and CyB components were mixed in a 1:1 ratio, spin-coated on a glass-bottom dish (Iwaki) at 500 r.p.m. for 1 min and cured at 80 ◦ C for 2 h (elastic modulus ∼ 8 kPa). The substrate was silanized using a 5% solution of aminopropyl triethoxysilane (Sigma) in ethyl alcohol for 5 min. Subsequently, carboxylated green fluorescent beads (100 nm, Invitrogen) were diluted in deionized water (1:500) and added to the substrate. After incubation for 5 min, the substrates were washed with deionized water to remove loosely bound beads. To pattern these soft substrates, we used a modified microcontact printing method described recently54 . Briefly, the fibronectin pattern was first printed on a watersoluble polyvinyl alcohol (Sigma Aldrich) membrane and then transferred to the soft substrate. A solution of 0.2% Pluronics was added to dissolve the membrane and block the non-fibronectin-coated regions. ‘Force-loaded’ images (with cells) of the beads were first obtained using a ×20 objective. The ‘Null-force’ or cell-free image was obtained after lysing the cells using an SDS lysis buffer. Computation of stresses was performed using a custom-written MATLAB code as described previously55 . Atomic force microscopy. HaCaT monolayers grown on Petri dishes stamped with fibronectin were mounted on the stage of an atomic force microscope (Nanowizard II BioAFM, JPK instruments AG). Indentation was performed with a polystyrene bead of ∼4.5 µm in diameter attached to the end of a cantilever (k = 0.03 N m−1 , Novascan Technologies) using a force of 3 nN at 1 Hz. Young’s modulus was extracted by fitting the indentation curves with a Hertz model using JPK Data Processing Software (JPK instruments AG). Laser ablation studies. Laser ablation was performed using an ultraviolet laser (355 nm, Minilite II, Continuum, USA) mounted on a confocal microscope (Nikon A1R) using a ×40 water-immersion objective. Images were acquired at ∼15 frames per second and analysis of the relaxation was performed in ImageJ.

Received 19 December 2012; accepted 17 October 2013; published online 1 December 2013

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Acknowledgements The authors thank C. Gay, J-B. Fournier, A. J. Kabla, R-M. Mège, J-M. di Meglio and W. James Nelson for helpful discussions. The authors would also like to thank M. Ashraf and S. Vaishnavi for the microfabrication and C. Xi for the illustrations. Financial support from the Agence Nationale de la Recherche (ANR 2010 BLAN 1515 awarded to B.L.), the Human Frontier Science Program (grant RGP0040/2012) and the Mechanobiology Institute (Team project funding) is gratefully acknowledged. B.L. acknowledges the Institut Universitaire de France (IUF) for its support. The research was conducted in the scope of the International Associated Laboratory Cell Adhesion France Singapore (CAFS). X.T. acknowledges financial support from the Spanish Ministry for Economy and Competitiveness (BFU2012-38146), and the European Research Council (Grant Agreement 242993).

Author contributions S.R.K.V., B.L. and C.T.L. designed research, S.R.K.V., M.H.N. and Y.T. performed experiments, H.H. contributed new reagents, S.R.K.V., A.B., H.H., X.T. and B.L. analysed data, S.R.K.V. and B.L. wrote the paper, and B.L. and C.T.L. oversaw the project. All authors read the manuscript and commented on it.

Additional information Supplementary information is available in the online version of the paper. Reprints and permissions information is available online at www.nature.com/reprints. Correspondence and requests for materials should be addressed to C.T.L. or B.L.

Competing financial interests The authors declare no competing financial interests.

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