Supplementary Materials for

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1 Supplementary Materials for Adhesion Functions in Cell Sorting by Mechanically Coupling the Cortices of Adhering Cells Jean-Léon Maître, Hélène Berthoumieux, Simon Frederik Gabriel Krens, Guillaume Salbreux, Frank Jülicher, Ewa Paluch, * Carl-Philipp Heisenberg * *To whom correspondence should be addressed. heisenberg@ist.ac.at (C.-P.H.); paluch@mpicbg.de (E.P.) This PDF file includes: Supplementary Text Materials and Methods Figs. S1 to S10 Legends for movies S1 to S23 Tables S1 to S12 References Published 23 August 2012 on Science Express DOI: /science Other Supplementary Material for this manuscript includes the following: (available at Movies S1 to S23

2 Supplementary material Supporting model 3 Equilibrium shape of a cell doublet 3 Cell triplet: Determination of the cortex tension at the cell- cell interface 7 Discussion of the model assumptions 11 Mechanical behavior of cell- cell and cell- medium interfaces 11 Line tension at the cell- cell contact edge 14 Supporting materials and methods 17 mrna and morpholino injections 17 Plasmids 17 Transgenic zebrafish lines 18 Dual pipette aspiration assay (DPA) 19 Cell preparation 19 Micropipette preparation 19 Micromanipulator set up 20 Separation force measurements 20 Cell triplet assay 24 Immunostaining 24 Cell sorting in vitro 26 Whole embryo live imaging 27 Supplementary figures 28 Figure S1. Contact size in progenitor cell doublets. 28 Figure S2. Localization of core components of the Cadherin- adhesion complex at the cell- cell contact in progenitor cell doublets. 29 1

3 Figure S3. Accumulation of core components of the Cadherin- adhesion complex at the cell- cell contact edge in progenitor cell doublets. 30 Figure S4. Actin, Myl12.1 and Cdh2 localization at the cell- cell contact in progenitor cell doublets. 32 Figure S5. Influence of Cdh2 cytoskeletal anchoring on progenitor separation force Fs. 33 Figure S6. Influence of Cdh2 cytoskeletal anchoring on progenitor cell- cell contact radius Rc. 34 Figure S7. Germ layer progenitor cell sorting in vitro. 35 Figure S8. Myl12.1 localization in germ layer progenitor cells in vivo. 36 Figure S9: Schematic illustration of a homotypic doublet. 37 Figure S10: Schematic illustration of a cell triplet shape before and after separation. 38 Supporting legends 39 Supporting tables 44 Supporting references 57 2

4 Supporting model The formation and expansion of a cell- cell contact is thought to depend primarily on cell adhesion and cortex tension (6, 16). Several theoretical models have been developed to describe the role of cell adhesion and/or cortex tension in cell- cell contact formation (1, 17, 18), the generation of tissue surface tension (2, 19), and cell sorting (4). Moreover, various biophysical techniques have been used to analyze the adhesive and/or tensile properties of single cells and relate those properties to the adhesiveness of their contacts (10, 20-23) and their sorting behavior (12, 24). However, studies that combine biophysical experimentation with theoretical modeling to elucidate the function of cell adhesion and cortex tension in cell- cell contact formation and cell sorting are still sparse. To better understand the role of adhesion and tension in cell- cell contact formation, we studied the formation and separation of cell- cell contacts between zebrafish progenitor cells. We first developed a theoretical description of the shape of a doublet of adhering cells, which yields the ratio of surface tensions at the cell- cell and cell- medium interfaces of the doublet. We then designed an experiment to directly measure cortex tension at the cell- cell contact zone, using triplets of adhering cells. Together, these measurements allowed us to unravel the respective contributions of adhesion and cortex tension to contact formation. Equilibrium shape of a cell doublet In this first part, we develop a physical description of the equilibrium shape of a homotypic cell doublet (fig. S9). For zebrafish progenitors, a stationary contact shape is typically reached within 5 minutes after the two cells have been put into contact (Fig. 1B and supplementary Movies S1-3). The shape of a cell is primarily determined by the cellular actomyosin cortex, a thin layer of cross- linked actin filaments connected to the cell plasma membrane (25). At 3

5 the spatial scale of cells, the actomyosin cortex behaves like a viscoelastic material, with a characteristic relaxation timescale from elastic to viscous behavior of about 1 min (26, 27). On the timescale typical for stationary progenitor cell- cell contact formation, the cortex thus behaves as a sheet under tension generated by myosin activity, and the cells can be described as fluid objects with a surface tension. The observation of a decrease in cortical myosin intensity at the cell- cell interface compared to the cell- medium interface suggests that tension is decreased in the contact area (Fig. 1E) (6). We thus introduced two tensions: Tcm, the cortex tension at the cell- medium interface, and Tcc, the cortex tension at the cell- cell contact. We further assumed that cell- cell contact expansion is promoted by chemical interactions between Cadherin trans- membrane adhesion molecules. The contribution of the formation of adhering trans- bonds to the global energy of the system is captured by a chemical energy Echem. The total energy of the system can then be written as the sum of the mechanical and chemical energies: E = 2TcmAcm + 2TccAcc + Echem 2PV [1] where Acm is the cell- medium interface area, Acc the cell- cell contact zone area, and Echem the chemical energy of the system. P is a Lagrange multiplier that imposes the conservation of the cell volume V. To determine an explicit expression of Echem, we considered a simplified mechanism for the formation of Cadherin- Cadherin adhesion bonds. We further assumed that single Cadherin molecules freely exchange between the cell- cell and the cell- medium interfaces according to the reactions:! C cm! C cc [2]! C cm! C cc [3] 4

6 where C! cm and C! cm denote the Cadherin molecules outside of the adhesion zone and C! cc and C! cc denote single Cadherins inside the adhesion zone on cell 1 and cell 2 respectively. A recent study suggests that Cadherins not engaged in Cadherin- Cadherin trans- bonds do not interact with the actomyosin cortex i (28). We thus assumed that the free energy of species C cm i and C cc (with i = 1, 2) was the same, setting the equilibrium constants of the reactions [2] and [3] to 1. Inside the adhesion zone, Cadherins from the two contacting cells can interact and form trans bonds according to the following mechanism: k!" C!! cc + C cc C! [4] k!"" where C2 denotes a trans- bond Cadherin complex and kon and koff the association and dissociation rate constants of this complex, respectively. The total number of Cadherin molecules, N, at the cell surface is assumed to be constant and equal for each cell. The chemical energy of the system Echem can then be written as: Echem = 2Ncmμcm + 2Nccμcc + N2μ2 [5] where Ncm, Ncc and N2 are the number of species Ccc, Ccm and C2, and μcc, μcm and μ2 their respective chemical potentials. Conservation of matter imposes N = Ncm + Ncc + N2. The chemical potential μi, (i = cm, cc, 2) of species Ci is written!! as μi = μ i + kbt ln Ci, where μ i is the standard chemical potential of each species, Ci their surface concentration, kb the Boltzmann constant, and T the temperature. The equilibrium shape of the cell doublet can then be obtained by minimizing the total energy given in Eq. [1]. The shape of each cell is a spherical cap of 5

7 radius R and contact angle θ (fig. S9). By minimizing the total energy E with respect to Ncc, N2, R, θ, one obtains: 2γ cm cos(θ) = 2γ cc ω with ω = k b TC! [6] C cm = C cc, C! C cc! = k on k off [7] P =!γ cm R [8] Equation [6] corresponds to the force balance at the contact projected on the y- axis (fig. S9). In order to account for the entropic contribution from free diffusion of unbound Cadherins, we introduced corrected cell- medium and cell- cell cortex tensions: γcm = Tcm kbtccm and γcc = Tcc kbtccc. The trapping of cadherins engaged in trans- bonds in the contact zone generates an adhesive tension of magnitude ω, promoting the expansion of the contact. Eq. [7] corresponds to the law of mass action applied to chemical reactions [2-4]. Eq. [8] corresponds to Laplace s law at the cell- medium interface. From Eq. [6], we obtained equation [1] given in the main text. This relation can be rewritten as cos(θ) = 1 γ cm! γ cc γ cm ω!γ cm, illustrating that two factors control the size of the cell- cell contact: the difference in cortex tensions between the cell- medium and the cell- cell interfaces γcm γcc, and the adhesion tension ω = kbtc2. When both of these factors vanish, the contact angle θ vanishes and no contact is formed. By measuring the contact angle θ for ectoderm, mesoderm and endoderm homotypic doublets (materials and methods), we were able to extract the ratio progenitor type. We found that the ratio γ i!γ cm, with γi = 2γcc ω for each γ i!γ cm was smaller for ectoderm cells than for endoderm and mesoderm doublets (Fig. 1B and supplementary Movies S1-3). 6

8 Cell triplet: Determination of the cortex tension at the cell- cell interface In order to disentangle the relative contributions of the difference in cortex tensions between the cell- medium and cell- cell interfaces γcm - γcc, and of the adhesion tension ω, to progenitor cell- cell contact formation, we designed an approach to directly measure cortex tension at the cell- cell interface. To this end, we analyzed shape changes in the contact region upon separation of a cell- cell contact, using a dual- micropipette assay (DPA) to separate adhering cells (29, 30). We reasoned that upon contact separation, the former contact zone, where cortex tension is expected to be lower than at the cell- medium interface (Fig. 1E), should deform and assume a curvature dictated by cortex tension in this region, γcc. As cell aspiration into micropipettes may perturb cellular mechanical properties, such as tension (31-33), we used linear aggregates of three cells (cell triplets) and analyzed the deformation of the middle cell, which was not in contact with a micropipette. When put into contact, the three cells reached a steady- state configuration within 5 minutes (supplementary Movies S4-6). Two of the cells in the aggregate were then separated (materials and methods). After contact separation, the former adhesion zone of the middle cell deformed and adopted a spherical cap shape with increased curvature, indicative of reduced cortex tension at the cell- cell contact as compared to the cell- medium interface. About 10 seconds after detachment, a stationary shape was reached and remained stable for about 1 minute, after which the bulge retracted (supplementary Movies S4-6). To extract the ratio of surface tensions at the cell- cell and the cell- medium interface γcc/γcm for each type of germ layer progenitors, we analyzed the shapes adopted by the cell triplets during contact separation. We first derived the force balance equations setting the steady- state shape of 7

9 the cell triplet before contact separation. A schematic description of the geometry of the cell triplet before separation is given in fig. S10A. Strong micropipette aspiration has been shown to affect cell surface tension (31). We thus assumed that the aspirated cells had a surface tension γs different from the tension γcm of the cell- medium interface of the non- aspirated cell in the middle of the triplet, with γs being dominated by the tension of the plasma membrane (32-34). We further assumed that the deformation of the cell- medium cortex of the middle cell remains negligible and thus that its cortex tension is equal to γcm. Introducing the radius of an aspirated cell, the contact radius R b c, and the contact angle θb (fig. S10A), we then wrote the force balance equation at the edge of the contact. The projection of this force balance equation on the long axis of the cell triplet yields the ratio of the cortex tension of the aspirated cell over the cortex tension of the middle cell at the cell- medium interface: γ s γ cm = R b R c b!"# θ b!!"#!! R c b R b [9] We then derived the force balance equations describing the shape of the cells after contact separation. A schematic description of the stationary two- cell shape reached 10 seconds after detachment of the third cell is given in fig. S10B. During the short time after separation we assumed that the cell cortex behaves as an elastic material under contractile tension. The two side cells and the former cell- cell interface of the middle cell, now forming the bulge, adopted spherical shapes (supplementary Movies S4-6), and were characterized by effective surface tensions γs, γi and γcc, respectively, while elastic effects were neglected compared to surface tension. In contrast, the surface of the middle cell was strongly curved near the basis of the bulge. This local change in curvature indicates that a normal force is required in the force balance at the bulge basis, which cannot be accounted for by surface tension 8

10 alone, which would exclusively generate tangential forces. Therefore, elastic stresses play a role and we described the cell- medium interface of the middle cell as an elastic shell. For more details of the model assumptions, see discussion at the end of this section. Based on these assumptions, we wrote the force balance equation at the edge of the remaining cell- cell contact, projected on the x- axis of the cell doublet, and the pressure balance equation between the two non- separated cells. From these two relations, we extracted the ratios of cortex tensions: γ i =!"#(θa ) γ s!"#(θ i ) [10] γ cc γ s = R b R a γ i R b γ s R i [11] where R a is the radius of the side cell, Ri is the curvature radius of the remaining contact, θ a is the contact angle, and Rb is the radius of the bulge. The angle θi, defined in fig. S10B, has the following expression: θ i = π + sin!! R c a R a θ a sin!! R c a R i, where R c a is the radius of the remaining contact. Using Eqs. [9-11], we then extracted the ratios of tensions γ i γ s, γ cc γ s and γ s γ cm from the geometrical parameters describing cell shape that we measured during the cell separation experiment: (R b, R c b, θb, R a, R c a, Ri, θa). The ratios γ i γ s and γ cc γ s were computed for ectoderm, mesoderm and endoderm cell triplets and allowed us to deduce the ratio of tensions at the cell- cell and cell- medium interfaces γ cc γ cm (Fig. 1D and Table S3). The decrease in cortex tension at the cell- cell interface compared to the cell- medium interface was more pronounced in the case of ectoderm progenitors (~50 %) than in the case of endoderm or 9

11 mesoderm progenitors (~30 %). From Eq. [6], we then obtained ω!γ cm = γ cc γ cm cos (θ) and calculated the ratio of the adhesion tension to twice the tension at ω the cell- medium interface for ectoderm, mesoderm and endoderm!γ cm doublets (Fig. 1D and Table S3). For all three germ layers, the adhesion tension ω was found to be negligible compared to the difference in cortex tension γcm - γcc. This indicates that the parameter controlling the contact size in different germ layers (Fig.1B) is primarily the ratio of cortex tensions at the cell- cell and cell- medium interfaces γ cc γ cm, rather than the adhesion tension ω. In other words, ectoderm doublets form larger contacts than mesoderm or endoderm cells because the relative decrease of cortex tension in the cell- cell contact zone γ cm! γ cc γ cm is more pronounced in ectoderm doublets. Finally, to confirm that the adhesion tension ω is negligible compared to the difference in cortex tension γcm γcc, we estimated the trans- bond concentration that would theoretically be needed at the contact zone in order for adhesion energy to contribute 10 % of the mechanical energy resulting from the difference in cortex tensions at the contact, i. e. ω!γ cm = 0.1 γ cm! γ cc γ cm. Using the expression of ω given by Eq. [6], T = 300 K, and a value γcm = 50 pn.µm 1 (previously determined for progenitor cells (12)), we found a surface concentration of adhesion complexes of about bonds.µm 2. This value is considerably higher than the density of ~10 3 bonds.µm - 2 suggested by previous measurements (35). This further supports our conclusion that adhesion tension ω does not play a major mechanical role is setting the size of a cell- cell contact. Taken together, our experiments and theory suggest that variations in the concentration of Cadherins engaged in trans- bonds among germ layer progenitors have little, if any, influence on the observed differences in contact 10

12 size. Instead, the size of a cell- cell contact appears to be determined by the extent of decrease in cortical tension at the contact zone. Discussion of the model assumptions Mechanical behavior of cell- cell and cell- medium interfaces In our analysis of the cell triplet experiments, we assumed that the deformation of the former cell- cell interface (magenta on fig. S10B) and the cell- medium interface (black on fig. S10B) of the middle cell is controlled by different mechanical properties. This assumption was motivated by our experimental observations that the cell- cell interface contains little actomyosin cortex compared to the cell- medium interface (Fig. 1E), and that the cell- cell interface displays a strong deformation after contact separation (Fig. 1C and Movie S4). More specifically, on short timescales, the deformation of a viscoelastic cortex under contractile tension depends both on cortex elasticity and on myosin generated tension (25). In the following, we show that both bending and stretching elasticity can be neglected when describing the deformation of the cell- cell interface after separation. As a result, a Laplace law governed by tension determines the shape of the bulge growing at the former cell- cell interface, whereas elasticity can significantly contribute to the behavior of the cell- medium interface. Before contact detachment, the cell- cell interface is flat and circular. We thus modeled this part of the cortex as a circular elastic plate of radius R b c (~ 5 µm), thickness h, elastic modulus E and a contractile tension γcc. Upon separation of the two adhering cells, the pressure difference across the plasma membrane at the former cell- cell interface is increased from almost zero (only a small pressure difference between the two cells is to be expected as indicated by the flat interface) up to the cell- medium pressure difference. In contrast, the pressure difference across the plasma membrane at the cell- 11

13 medium interface does not change. The change in pressure difference at the cell- cell interface, ΔP, causes the cortex at the cell- cell interface to strongly bend and stretch (see Movie S4). To address the contribution of bending elasticity to the deformation of the cell- cell interface, we described the deformation of the cell- cell interface as the deformation of an elastic plate under a normal load using the non- linear von Karman equations (36). Depending on the order of magnitude of applied forces and on the material properties of the elastic plate, the response can be dominated by tension or bending elasticity (37, 38). The appropriate description depends on the value of a control parameter k1: k! = ΔP E!! R c b h!! [12] For large values of k1 (k1 15), corresponding to a large load or/and a thin cortex, the effect of bending becomes negligible compared to the effect of tension (37). The deformation and forces exerted in the rim of the bulge depend on the boundary conditions at the circumference of the plate. In our case, it is reasonable to assume that the plate boundaries are free to rotate. Assuming a ratio of 20 between the radius of the interface R c b and the thickness of the cortex h (consistent with previous estimates of cortex thickness (39)), and with the pressure difference given by the Laplace law, ΔP =!γ cm R, at the cell medium interface (with γcm ~ 50 pn.µm- 1 and R ~ 10 µm), k1 15 corresponds to cortical Young modulus values: E 600 Pa [13] Previous measurements indicate that the elastic modulus of a prominent actin cortex is of the order of ~1000 Pa (25, 40). Our observation that the cortex is 12

14 considerably weaker at the site of bulge formation indicates that assuming a large value for k1 (k1 15) is appropriate, and that we can thus neglect the bending energy. Consequently, bending moments and the resulting normal forces at the bulge basis are dominated by the tension and elasticity of the cell- medium interface at the margin of the bulge. To address the contribution of stretching to the deformation of the cell- cell interface, we assumed that the contact zone deforms elastically after separation. The average additional tension created by the stretching due to the deformation of the bulge γel is given by γel Eh( S/S), where S/S is the amplitude of the deformation. This elastic tension γel would contribute to the total tension of the deformed bulge γ cc: γ cc = γcc + γel [14] where γcc is the active tension created by myosin activity, taken equal before and after deformation. Because the bulge is spherical, Laplace s law implies that the total tension is uniform within the bulge, justifying that we only consider average tensions. In the analysis of the triplet experiment described above (see Eq. [11]), we estimated the total tension of the deformed bulge γ cc from the application of Laplace law and the balance of pressure between the two non- separated cells. To estimate how stretching elasticity affects our estimate of the adhesion tension ω, we now consider a cell doublet at steady state (Fig. 1A). From Eq. [6], we obtain: 13

15 ω = - 2 γcm cosθ + 2γ cc - 2γel [15] We then used the estimation of γ cc to extract the adhesion tension neglecting the role of cortex elasticity at the contact, ω0 = - 2 γcm cosθ + 2γ cc. When taking into account the role of the cortical elasticity, the estimate of the adhesion tension would then be modified to: ω = ω0 2γel [16] Eq. [16] shows that the presence of an elastic tension γel in the bulge would further decrease the estimate of the adhesion tension ω. Because the adhesion tension ω has to be positive and the value of ω0 that we obtained is not significantly different from zero (Fig. 1D), we conclude that (1) the value of ω remains negligible compared to cortex tension, (2) γel brings a small contribution to the total tension of the bulge. Indeed ω > 0 implies 2γel < ω0. Because ω0 << γcm (Fig. 1D), this implies γel << γcm. Thus, we conclude that the elastic contribution due to stretching of the cortex during the growth of the bulge can be neglected. Taken together, these considerations allow us to conclude that neither bending nor stretching elasticity has a significant contribution to the deformation of the cell- cell interface of the middle cell of a cell triplet after contact separation. Therefore, Eqs. [9-11] can be used to accurately calculate the ratio of tension between the cell- cell and cell- medium interface γcc/γcm. Line tension at the cell- cell contact edge An accumulation of actin and myosin- 2 (Myl12.1) at the margin of the cell- cell contact in progenitor cell doublets and triplets could in principle give rise to a 14

16 line tension working against cell- cell contact formation. However, while we observed a ring- like localization of Cdh1/2, Ctnnb, Ctnna and actin at the contact margin, no such localization was found for Myl12.1 (Fig. 2C). This suggests that there is no prominent actomyosin ring present at the cell- cell contact margin exerting significant line tension. To more directly estimate the influence of line tension on cell- cell contact formation, we calculated the expected change in adhesion tension ω due to a line tension generated by a local accumulation of the cortex at the contact edge. A local cortex thickening would result in a line tension Λ contributing to the force balance at the contact (Fig. 1A) by a factor 2Λ/Rc. The line tension resulting from an accumulation of the cortex over h = 500 nm (which is likely to be an overestimate as no accumulation is observed with fluorescent probes Fig. 2C, fig. S3) can be approximated by Λ ~ γcmh, where γcm is the cortex tension at the cell- medium interface. Using the values of γcm ~ 50 pn.µm - 1 and Rc ~ 5 µm extracted from the experiments, the resulting contribution of the line tension to the force balance would be Λ/Rc ~ 5 pn.µm - 1. The resulting change in ω due to the presence of a line tension Λ would be given by: ω new γ cm = ω old γ cm + 2Λ γ cm R c [15]!Λ γ cm R c < ~0.2 [16] Therefore, a line tension resulting from a cortex accumulation at the contact edge would only lead to a small change in ω/γcm (Fig. 1D) and therefore would not change our conclusion that adhesion tension is negligible compared to cortex tension. 15

17 Also note that a line tension would not substantially affect our estimation of the tension ratio γcc/γcm in the triplet experiments. Indeed, the extraction of the ratio γcc/γcm relies on the calculation of several intermediate tension ratios: The ratio γs/γcm of the side cell tension over the cell- medium interfacial tension is extracted from the force balance equation at the contact projected in the direction normal to the contact. The contribution of a line tension vanishes in this direction. Therefore, the calculation of γs/γcm is independent of line tension. The ratio γi/γs of the cell- cell interfacial tension (of the remaining cell- cell contact in the triplet) over side cell tension. In this case, the plasma membrane of the side cell is likely to be strongly tensed because of pipette aspiration (see also above), and we therefore assumed that plasma membrane tension dominates at these interfaces (32, 33). Finally, we used the Laplace law to estimate the ratio γcc/γi. This relation remains unchanged in the presence of line tension. We conclude that the presence of line tension is unlikely to modify the obtained value of γcc/γcm. 16

18 Supporting materials and methods mrna and morpholino injections Zebrafish embryos were induced to consist of one germ layer progenitor cell type only by micro- injection of 1- cell stage embryos with: 100 pg lefty1 mrna for ectoderm, 100 pg cyclops together with 2 ng casanova morpholino (MO, 5 - GCATCCGGTCGAGATACATGCTGTT - 3, GeneTools) for mesoderm, or 50 pg casanova mrna to obtain endoderm (12). To visualize Cdh2, Ctnnb1, Ctnna or filamentous actin, 100 pg of cdh2- egfp, egfp- ctnnb1, ctnna- egfp or 50 pg of LifeAct- RFP (41) mrna were injected in 1- cell stage embryos. To substitute endogenous Cdh1 with controlled amounts of Cdh2, 2 ng cdh1mo (5 - TAAATCGCAGCTCTTCCTTCCAACG - 3, GeneTools) together with 100 pg of cdh2- egfp or 50 pg cdh2δcyto- egfp mrna were injected in 1- cell stage embryos. Synthetic mrna was produced by using the SP6 mmesage mmachine kit (Ambion). Plasmids Tagging of Xenopus Ctnnb1 (42) was performed using Gateway technology (Invitrogen). The following primers were used to flank the gene with attb1 and attb2 overhangs: 5 - GGGGACCACTTTGTACAAGAAAGCTGGGTA TACAGATCGGTGTCAAAC - 3 and 5 - GGGGACAAGTTTGTACAAAAAAGCAGGCTT AATGGCTACCCAGTCTGAC - 3. The PCR product was recombined with pdonr221 (Lawson #208) and the resulting entry- clone was recombined with the pcsnter- egfp destination vector (Lawson #223) to obtain the pcs- egfp- Ctnnb1 expression plasmid. 17

19 Tagging of zebrafish Ctnna (43) was performed by employing Gateway technology (Invitrogen) using the following primers: 5 - GGGGACAAGTTTGTACAAAAAAGCAGG CTTAATGACGAGCATTAACACT - 3 and 5 - GGGGACCACTTTGTACAAGAAAGCTGGGT AAATACTATCCATAGCTTT - 3. The resulting entry clones were recombined with pcsdest2 (Lawson #444) and p3e egfp (Lawson #440) to generate the pcs- Ctnna- egfp expression plasmid. Tagging of zebrafish Cdh2 and Cdh2Δcyto was performed by generating a Gateway entry- clone containing the full- length zebrafish cdh2 gene with attb1 and attb2 overhangs using the following primers: 5 - GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGTACCCCTCCGGAGGC - 3 and 5 - GGGGACCACTTTGTACAAGAAAGCTGGGTAGTCGTCGTTACCTCCGTA - 3 (corresponding to cdh2), and 5 - GGGGACAAGTTTGTACAAAAAAGCAGG CTTAATGACGAGCATTAACACT - 3 and 5 - GGGGACCACTTTGTACAAGAAAGCTGGGT AAATACTATCCATAGCTTT- 3 (corresponding to cdh2δcyto). The PCR products were then recombined with pdonr221 (Lawson #208) to generate Entry- clones, which in turn were used to recombine with pcsdest2 (Lawson #444) and p3e egfp (Lawson #440) to obtain the expression plasmids pcs- Cdh2- egfp and pcs- Cdh2Δcyto- egfp. Transgenic zebrafish lines The Tol2/Gateway technology was used (44) to generate transgenic zebrafish lines ubiquitously expressing an egfp or mcherry tagged version of Myl12.1 The zebrafish non- muscle myosin regulatory light chain coding sequence for the myl12.1 gene was amplified with gene specific primers (5 - GGGGACAAGTTTGTACAAAAAAGCAGGCTTAATGTCGAGCAAACGCGCC- 3 and 5 - GGGGACCACTTTGTACAAGAAAGCTGGGTAATCGTCTTTGTCTTTGGC- 3 ) from a poly- dt primer cdna library, generated from shield- stage TL embryos. The myl12.1 gene- product was subcloned into the gateway pdonr221 18

20 (Lawson #208), and the resulting entry clone was recombined with pdesttol2pa2 (Chien #394), pse β- actin promoter (Chien #299) and either p3e egfp (Lawson #440) or p3e mcherry (Lawson #441) to produce respectively ptol2- β- actin::myl12.1- egfp or ptol2- β- actin::myl12.1- mcherry. This plasmid was co- injected in wild type TL zebrafish embryos with mrna encoding a transposase (Invitrogen). Individual positive carriers were selected and out- crossed to wild type TL fish over 2 generations to select for stable single- copy genetic integrations. Dual pipette aspiration assay (DPA) Cell preparation 3-6 embryos at 5 hpf were dechorionated in Danieau s medium (58 mm NaCl, 700 µm KCl, 400 µm MgSO4, 600 µm Ca(NO3)2 and 5 mm Hepes at ph 7.2) and transferred to 1 ml CO2- independent DMEM/F12 (Invitrogen, complemented with L- Glut, 15 mm Hepes and 100 U/mL penicillin plus streptomycin, adjusted at ph 7.5, sterilized using 0.45 µm pore filters, and preheated to 28 C). Embryos were mechanically dissociated into single cells by mild shaking, and yolk proteins were removed by 2 successive wash steps with 1 ml fresh medium followed by centrifugation at 100 G for 2 min. Dissociated cells were seeded on a passivated glass bottom dish (Mattek). Passivation was performed by incubation with heat inactivated FBS (Invitrogen) for 17 min at RT. Micropipette preparation Glass capillaries (World Precision Instrument, TW100-3) were pulled automatically using a micro- needle puller (Sutter Instrument, P- 97 Flaming Brown micropipette puller), using a customized program (Ramp +3-5, Pull 50, Velocity 50, Time 50 and Pressure 500) to produce capillaries with a 1 cm long taper. The taper of the capillary was cut to an opening of ~ 3.5 µm internal radius and bent to a ~ 45 angle using a micro- forge (Microdata 19

21 instrument, MFG- 5). The micropipettes were passivated with heat inactivated FBS (Invitrogen) for 7 min at RT, followed by washing with PBS using a syringe (World Precision Instrument, Microfil MF28G- 5). Micromanipulator set up The micropipettes were connected to a Microfluidic Flow Control System (Fluigent, Fluiwell), with negative pressure ranging from Pa, a pressure accuracy of 7 Pa and change rate of 200 Pa.s - 1 on two independent channels. The microfluidic setup was mounted on two micromanipulators (Eppendorf, Transferman Nk2). Micropipette movement and pressure were controlled via a custom- programmed Labview (National Instruments) interface. To manipulate cells, ~ 20 Pa negative pressure was used. Separation force measurements Methods The separation force Fs was determined as previously described (29). After bringing cells into contact, cell doublets were left to adhere for a given contact time. The adhering cells were then grabbed by two glass micropipettes (holding and probing pipette) on the opposite sides of the cell- cell contact. The holding micropipette was used to firmly hold one cell with a fixed pressure ranging from Pa depending on the strength of the probed cell- cell contact. The probing micropipette was used to apply stepwise increasing pressures ranging from Pa with step- sizes between 10 and 500 Pa to the other cell. After each pressure step, the micropipettes were pulled apart at 20 µm.s - 1 in an attempt to separate the contacting cells. Once the applied pressure in the probing pipette was sufficiently high to separate the contacting cells, Fs could be calculated from the final pressure and the pressure of the last failed separation using Fs = π Rp 2 (Pn- 1 +Pn)/2, with Rp being the micropipette radius, Pn the pressure applied by the probing micropipette during the separating step, and Pn- 1 the pressure applied by the 20

22 probing micropipette during the pulling step preceding the separating step. The number of pressure steps was usually kept between 1 and 3. Cell separations were imaged on two microscope setups (1) Zeiss Axiovert inverted microscope equipped with a dry Zeiss LD PlanNeoFluar 40X, 0.6 NA Ph2 Korr, a cooled charge- coupled device CoolSnap HQ (Photometrics), and a heating box set to 28 C; (2) Leica SP5 confocal microscope equipped with a resonant scanner, using 488 nm and 561 nm laser lines for simultaneous fluorescent and bright- field imaging. Imaging was performed using a Leica 20X, 0.7 NA objective. Temperature in the dish was kept constant at 28 C by a heating sample holder. Images were acquired using Metamorph software on the Axiovert setup or the Leica Application Suite on the Leica SP5 setup. Cell doublet geometry (contact angles, contact size, cells radii) was characterized using FIJI (FIJI Is Just ImageJ) using images in which cells are in focus and the contact size is stable for over 10 s. To visualize core components of the Cadherin adhesion complex during progenitor cell- cell contact separation, we expressed egfp- tagged versions of Cdh2, Ctnnb1 and Ctnna, previously shown to link Cadherins to the cortical cytoskeleton, and, more generally, regulate membrane- to- cortex attachment (16, 28, 45, 46). We visualized exogenous Cdh2 as a proxy for endogenous Cdh1 localization, since exogenous Cdh2 could functionally replace endogenous Cdh1 in our DPA assay (Figs. 3B,C), and fluorescently tagged versions of Cdh1 itself did not efficiently localize to the cell- cell contact of progenitor cells and were unable to functionally replace endogenous Cdh1. We also expressed a C- terminal truncated version of Cdh2 (Cdh2Δcyto), which has previously been shown to be unable to anchor to the cytoskeleton, but still efficiently localize to cell- cell junctions (47) and form trans- bonds (48, 49). 21

23 When comparing Cdh2 and Cdh2Δcyto expressing cell doublets, fluorescence intensity at the cell- cell contact was used to normalize the separation force Fs measurements as follows: the mean intensity (Ic) of exogenous Cdh2 expression at cell- cell contacts was measured using FIJI. Ic was then used to normalize the measured Fs for the different cell- cell contacts to their respective Cdh2- egfp expression levels. The normalized Fs/Ic of Cdh2- egfp expressing ectoderm homotypic doublets was subsequently used as a reference to which Fs/Ic of Cdh2Δcyto- egfp expressing ectoderm doublets, and Cdh2- egfp or Cdh2Δcyto- egfp expressing endoderm doublets were compared. Interpretation of separation forces The DPA assay was performed under dynamic loading, and the measured separation force Fs depends on experimental settings such as the pulling velocity and on the mechanical properties of the cells. In the limit where the pulling velocity is very slow, separation would be quasi- static and a fluid description would be used (50, 51). However, in order to avoid mechanosensitive responses from the cells, we performed our experiments in a fast pulling regime (pulling velocity of 20 µm.s - 1 ) such that the actual cell- cell contact separation occurs in less than 1 s. During this time, the cytoskeleton behaves elastically and the cell- cell contact does not reorganize. Indeed, the contact radius did not change visibly during pulling in our experiments (Movie S7 shows the separation process imaged at 1 frame per s). Such a situation is best captured by the separation of two elastic solids. A simple model for such a situation has been proposed (11). In the framework of an elastic description of the cell response during the separation process (11), the separation force Fs as a function of the number of adhesive bonds N is bounded from above by a linear function F lim s = a N, where a is a coefficient depending on the elastic properties of the system, the 22

24 velocity of pulling and the strength of individual adhesion bonds. This upper limit is the exact value of the separation force in the absence of cooperative effects during unbinding. Assuming that adhesive bonds are located at the periphery of the contact (Fig. 2C), F lim s can be rewritten as F lim s = 2 π a ρ Rc, where ρ is the line density of adhesive bonds and Rc is the contact radius. The higher separation force of ectoderm doublets in our experiments (Fig. 2A) could therefore be due to a higher contact radius Rc, line density of adhesive bonds ρ and/or other cell properties (contained in a). To exclude the possibility that higher separation forces in ectoderm doublets are solely due to higher contact sizes, we computed the ratio of separation force to contact radius Fs/Rc. Since this ratio was still higher for ectoderm doublets than for mesoderm and endoderm doublets (Fig. 2B), we concluded that ρ and/or a are not equal between the different progenitor cell types. This conclusion would not change assuming cooperative effects between bonds, since in such case Fs would scale with log (N) and thus the influence of the radius on the force would be smaller (11). Consequently, dividing the separation force by Rc would in this situation overestimate, rather then underestimate the effect of the radius on the force. Progenitor cells in the gastrulating embryo in vivo frequently make contact to multiple cells rather than one cell as in cell doublets in vitro. To test whether cells in contact with multiple cells behave differently in respect to our separation- force measurements compared to cells in contact with one other cell (cell doublets), we analyzed separation forces in cell triplets, where the middle cell is in contact with two side cells. Separation forces for both ectoderm and endoderm homotypic triplets at 1 min contact time were indistinguishable from the forces measured when separating doublets ((mean ± SD): ecto doublet (15.2 ± 4.4; n = 9) nn, ecto triplet (14.6 ± 6.6; n = 9) nn, endo doublet (1.7 ± 1.3; n = 7) nn and endo triplet (2.1 ± 2.8; n = 7) nn). This suggests that cells in contact with multiple (two) cells show similar separation forces than cells in contact with just one other cell. 23

25 Finally, the method to quantify cell adhesion in our previous study (Single Cell Force Spectroscopy, SCFS; (12)) differs from the method we used to quantify cell- cell separation forces in our present study (DPA), and consequently the values obtained with these two methods are not directly comparable. The main difference between these two methods is that in the case of SCFS, progenitor cells are brought into contact through constant force or distance, while in the DPA experiments, cells can freely form their cell- cell contacts. It is therefore conceivable that the separation forces FS recorded with those two methods differ because of differences in cell- cell contact formation. Cell triplet assay When contacting cells are separated, the former cell- cell contact directly after separation deforms. The curvature of the deformation depends on the surface tension ratio between the cell- medium and former cell- cell interface. However, the degree of this deformation can be affected by aspiration of the cell into the micropipette. To exclude this effect, we used cell triplets instead of doublets, since in triplets the middle cell is not aspirated and thus provides a more reliable readout for the deformation at the former cell- cell contact after separation. In the triplet assay, cells were separated after 5 min contact time with a single separation step using aspiration pressures ranging from Pa. Cell triplet geometry (cells radii, contact angles, contact sizes, contact bulge radius, contact bulge height, contact bulge opening, contact bulge angles, tongue lengths) was characterized both before and after separation using FIJI. Immunostaining Single progenitor cells were prepared as described for the DPA assay and allowed to adhere to an uncoated glass bottom dish for 30 min. Cells were 24

26 then fixed with 2% PFA (Sigma) in culture medium for 10 min at RT, washed twice with PBS to remove the PFA, and incubated in PBS with 0.1% Triton X100 from Merck (PBT) for 1 min at RT to permeabilize the plasma membrane. PBT was subsequently replaced with blocking solution consisting of PBT with 1% DMSO (Sigma) and 10% goat serum for 45 min at RT before primary antibodies diluted in blocking solution were added over- night at 4 C. Primary antibodies were washed three times with PBS for 1 min at RT and secondary antibodies diluted in blocking solution were added for 30 min at RT, followed by three washes with PBS for 1 min at RT to remove the antibodies. Immuno- labeled cells were imaged on an Olympus - IX71 inverted microscope equipped with spinning disc scan head Yokogawa CSU10 and Olympus UPlanSApo 100x 1.4 NA Oil, using 488 nm and 561 nm laser lines with an optical slicing of 0.5 µm. Resulting stacks of images were rotated using Bitplane Imaris to obtain cross sections of cell- cell contacts between cell doublets. Using FIJI, contact ring intensities were measured along the cell- cell contact margin using a 1 µm thick line whereas contact disc intensities were measured inside the cell- cell contact margin (Fig. 2C). Alternatively, fluorescence intensity was measured along the cell edge, across the cell- cell contact using a 1 µm thick line (fig. S4). Antibody staining of whole embryos was performed as described for single cells with the following modifications: embryos were fixed at 5 hpf in 4% PFA over- night at 4 C, washed three times in PBS for 5 min at RT to remove the PFA, and permeabilized by incubating them in PBT for 1 h at RT. Embryo dechorionation was performed in PBT, and embryos were incubated in blocking solution for 5 h at 4 C. Primary antibodies were added over- night at 4 C and then washed away three times with PBST for 5 min at RT. Secondary antibodies diluted in blocking solution were added for 3 h at RT and then washed away three times with PBST for 5 min at RT. Immuno- labeled embryos were imaged on an upright Leica SP5 confocal microscope equipped with a Leica 25x 0.95 NA dipping lens, using 488 nm and 561 nm laser lines. 25

27 The following antibodies were used: α- catenin (Transduction lab C21620, at 1:1000), β- catenin (Sigma C2081, at 1:500), Cdh1 (MPI- CBG (#174), at 1:200) and fluorophore- conjugated secondary antibodies (Molecular Probes A , A , A , A , A , A , A , A , at 1:250). F- Actin was stained using Rhodamine- conjugated Phalloidin (Invitrogen, at 1:200) that was incubated together with the secondary antibody. Cell sorting in vitro Micro- injected zebrafish embryos were kept at 31 C until they were dissociated at sphere- stage (4 hpf), as previously described (52). Cell sorting experiments were performed as previously described (53), with the following modifications: micro- molds were used with wells containing a diameter of ~ 400 µm and a height of ~ 800 µm, and were produced from a PDMS negative (Microtissues) following the supplier s guidelines. The 2% agarose micro- mold gel was equilibrated for at least 4 h with 4 ml of CO2- independent DMEM/F12 medium (Invitrogen) at 31 C. Homogenously mixed cell populations in equal ratios were seeded and cell sorting was recorded in 8 to 9 of the 256 micro- wells for at least 5 h in 3D over time at 28.5 C by acquiring 4 µm- spaced z- stacks of the aggregate in 2 or 3 channels at 5 min time- intervals, using a Leica SP5 confocal microscope equipped with a Leica 25x 0.95 NA dipping lens. After 4-5 h in culture, the degree of cell sorting was quantified by the decrease in the number of cells of one type at the aggregate- to- medium interface. This decrease was calculated by dividing the ratio of the total cell number of the 2 cell types by the ratio of the cell number of the 2 cell types at the aggregate- to- medium interface (N A total/n B total)/(n A surface/n B surface), with N being the number of cells in the analyzed focal plane, and A and B denoting the two different cell types. 26

28 Whole embryo live imaging Transgenic embryos expressing hras- egfp or Myl12.1- egfp, and embryos transiently expressing Cdh2- egfp were imaged on a Zeiss LSM 710 NLO upright microscope, equipped with an objective heater, which was set to 28 C and mounted on a Zeiss C- Apochromat 40x, 1.2 NA dipping lens. Two- photon microscopy was used to record high- resolution time- lapse movies in deep tissues with the multiphoton laser (Coherent, Chameleon) set to 900 nm. 27

29 Supplementary figures Figure S1. Contact size in progenitor cell doublets. Cell- cell contact radii Rc as a function of time and binned over 50 s are shown as mean ± SEM for ectoderm (red; n = 58), mesoderm (green; n = 40) and endoderm (blue; n = 47) homotypic progenitor cell doublets (supplementary Movies S1-3; see statistics in supplementary Table S9). 28

30 Figure S2. Localization of core components of the Cadherin- adhesion complex at the cell- cell contact in progenitor cell doublets. Fluorescence intensity at the cell- cell contact periphery (ring) and inner region of the contact (disc) in ectoderm (ecto) and endoderm (endo) homotypic doublets stained with α- Cdh1, α- Ctnnb1, α- Ctnna antibodies and phalloidin. Intensities of Cdh1 (n = 10/17/36), Ctnnb1 (n = 11/16/33), Ctnna (n = 24/16/20) and Actin (n = 10/16/12) are normalized to the respective mean values of ring intensities in ecto doublets. Error bars show the standard deviation (SD) as percentage of the mean values. 29

31 Figure S3. Accumulation of core components of the Cadherin- adhesion complex at the cell- cell contact edge in progenitor cell doublets. Ratio of fluorescence intensities at the contact edge (ring) to the inner region (disc) at cell- cell contacts of ectoderm (ecto, red; n = 10/11/24/10/8) mesoderm (meso, green; n = 17/16/16/16/7) and endoderm (endo, blue; n = 36/33/20/12/8) progenitor cell doublets. 30

32 31

33 Figure S4. Actin, Myl12.1 and Cdh2 localization at the cell- cell contact in progenitor cell doublets. F- Actin (top), Myl12.1- mcherry (middle) and Cdh2- egfp (bottom) localization at the contact edge in homotypic progenitor cell doublets. As outlined in the schematic diagram (top), fluorescence intensities were measured along the cell- medium and cell- cell interface edge, normalized to the maximum intensity along this line, and plotted as mean ± SEM. In ectoderm (ecto, red), mesoderm (meso, green) and endoderm (endo, blue) homotypic doublets, intensities of Cdh2- egfp (n = 17/13/16), F- Actin (n = 18/17/16) and Myl12.1- mcherry (n = 17/13/16) were measured. Scale bar = 5 µm. 32

34 Figure S5. Influence of Cdh2 cytoskeletal anchoring on progenitor separation force Fs. Separation force Fs for ectoderm (red) and endoderm (blue) homotypic progenitor cell doublets expressing Cdh2- egfp (n = 17/20) or Cdh2 lacking its cytoplasmic tail and fused to egfp (Cdh2Δcyto- egfp; n = 20/14) at 5 min contact time. Values are plotted as mean ± SEM and normalized to the Cdh2 expression level at the measured contact and at the contact of ectoderm doublets (supplementary Movies S12-15; see statistics in supplementary table S10). 33

35 Figure S6. Influence of Cdh2 cytoskeletal anchoring on progenitor cell- cell contact radius Rc. Cell- cell contact radius Rc in ectoderm (red) and endoderm (blue) homotypic progenitor cell doublets expressing Cdh2- egfp (n = 17/20) or Cdh2 lacking its cytoplasmic tail and fused to egfp (Cdh2Δcyto- egfp; n = 20/14) at 5 min contact time. Values are plotted as mean ± SEM and normalized to the Cdh2 expression level at the measured contact and at the contact of ectoderm doublets (supplementary Movies S12-15; see statistics in supplementary Table S11). 34

36 Figure S7. Germ layer progenitor cell sorting in vitro. Cell sorting efficiency for the experiments shown in Fig. 4A determined by dividing the ratio of the total cell number of the two cell types by the ratio of the cell number of the two cell types at the aggregate- to- medium interface (N A total/n B total)/(n A surface/n B surface). N, number of cells in the analyzed focal plane; A and B denote the different cell types undergoing sorting (see statistics in supplementary Table S12). 35

37 Figure S8. Myl12.1 localization in germ layer progenitor cells in vivo. The intensity ratio between the cell- cell interface (Icc) and the cell- interstitial space interface (Icm) are plotted for epiblast (epi, ectoderm progenitors; n = 15) and hypoblast cells (hypo, mesoderm and endoderm progenitors; n = 12). 36

38 Figure S9: Schematic illustration of a homotypic doublet. Contacting cells in a doublet are modeled as fluid spherical caps with associated surface tensions γcm at the cell- medium interfaces and γcc and ω at the cell- cell interface. γcm and γcc arise mainly from the contractility of the actomyosin cortex whereas ω is the adhesion tension, which negatively contributed to the overall surface tension at the cell- cell interface (see also Fig. 1A). These three surface tensions control the shape of the cell doublet reflected by the contact angle θ. R is the cell radius. 37