Simon Lo Vecchio

Raghavan Thiagarajan

David Caballero

Vincent Vigon

Laurent Navoret

Raphael Voituriez

Daniel Riveline
email: riveline@unistra.fr

Vincent -5☨

Vigon

Raphaël Voituriez

Collective Dynamics of Focal Adhesions Regulate Direction of Cell Motion

à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

INTRODUCTION

The ability of cells to perform directed motion is critical to the proper function of many physiological, pathological or developmental processes [START_REF] Carmona-Fontaine | Directional cell migration in vivo: Wnt at the crest[END_REF][START_REF] Machesky | Lamellipodia and filopodia in metastasis and invasion[END_REF][START_REF] Winjnholds | Basal Cell Migration" in Regeneration of the Corneal Wound-Bed[END_REF][START_REF] Stuelten | Cell motility in cancer invasion and metastasis: insights from simple model organisms[END_REF]. Bias in cell migration have been widely studied in vitro and it has been shown that soluble gradients of specific chemicals could trigger directionality by regulating signalling pathways and reorganizing the acto-myosin cytoskeleton [START_REF] Wang | Signaling Mechanisms for Chemotaxis[END_REF][START_REF] Affolter | Signaling to Cytoskeletal Dynamics during Chemotaxis[END_REF]. This phenomenon, known as chemotaxis, remains the widespread way to explain cell directionality in vivo [START_REF] Iglesias | Navigating through models of chemotaxis[END_REF][START_REF] Roussos | Chemotaxis in cancer[END_REF][START_REF] Corcione | CCL19 and CXCL12 Trigger in Vitro Chemotaxis of Human Mantle Cell Lymphoma B Cells[END_REF]. However, these gradients are rarely demonstrated or quantified in vivo. In addition, if they are reported, they are rarely shown to be causal for cell directions. How cells can sustain directions in vivo remains therefore unclear.

Previous works showed in vitro, as a proof of a principle, that local asymmetries in cellular environment can bias cell directionality in another mode of migration named ratchetaxis [START_REF] Caballero | Protrusion fluctuations direct cell motion[END_REF][START_REF] Caballero | Ratchetaxis: Long-Range Directed Cell Migration by Local Cues[END_REF][START_REF] Caballero | The cell ratchet: interplay between efficient protrusions and adhesion determines cell motion[END_REF][START_REF] Comelles | Cells as active particles in asymmetric potentials: motility under external gradients[END_REF][START_REF] Mahmud | Directing cell motions on micropatterned ratchets[END_REF][START_REF] Sun | Asymmetric nanotopography biases cytoskeletal dynamics and promotes unidirectional cell guidance[END_REF][START_REF] Jiang | Directing cell migration with asymmetric micropatterns[END_REF][START_REF] Brückner | Stochastic nonlinear dynamics of confined cell migration in two-state systems[END_REF]. This type of behavior can lead to long-term directionality and stresses the importance of stochastic probing associated to cell protrusions, as well as the role of environment topology. Using ratchetaxis as our framework, we show that cell directionality is primarily determined by the collective behavior of its adhesion sites namely focal contactsand can be tuned and predicted using simple geometric rules describing the surrounding environment. We also define a critical nucleation area, responsible of the observed bias: below this area, protrusions retract and the motion is prohibited, and above this area, new focal adhesions nucleate in a "wave-like" manner and cell migrates. This framework is supported by a theoretical model integrating accessible parameters and fitting the rectification plots extracted from experiments.

RESULTS

Cells locally elongate protrusions, which eventually lead to adhesion and its subsequent force generation and motion as a whole. To test this interplay between cell protrusion dynamics and cell motion, we designed an assay where an array of triangular adhesive motifs over a range of gap distances were fabricated using standard microfabrication techniques (see Methods). Triangular rhodamine-labelled fibronectin (FN) motifs were patterned to guide adhesion and morphodynamics of NIH3T3 cells (see Figure 1a). Non-printed areas were passivated with poly(L-lysine) grafted poly(ethylene glycol) (PLL-g-PEG), so that cells mainly adhere on fibronectin motifs and have their shapes imposed during the spreading phase. We studied the long-term migration of NIH3T3 fibroblasts along the micro-patterns for 48 h. Cells were able to move from one motif to the neighboring motifs, and eventually exhibit persistent motion towards one particular direction (see Figure 1 b-e and Movies S1-S4). Interestingly, cells migrated faster on separated motifs than on connected ones (see Figure 1h), consistent with the fact that friction is increased with larger adhesive areas [START_REF] Palecek | Integrin-ligand binding properties govern cell migration speed through cell-substratum adhesiveness[END_REF].

For each cell moving along a lattice unit (defined as the distance between the centers of adjacent triangles), we defined a final bias, which is determined by the position of the cell at the end of its trajectory relative to its original location after 2 days of migration: the final bias for an individual cell was set as "+" if it migrated towards the tip of the motif, and "-" if it migrated towards base of the motif (see Figure 1f). Interestingly, while the bias was not significant for triangles separated by 13 µm, 16 µm and 19 µm gap distances, we found that the average bias increased and reached close to complete rectification on triangles separated by gaps of 22 µm: almost 80% of the cells moved, on average, towards the "+" direction in this condition (Figure 1f). Beyond this value, the distance exceeded the maximum extension of protrusions. We tested large gap distances (d = 45 µm), and as expected, no migration was observed (see Movie S5); cells were unable to extend protrusions and find a 'docking' site. These results show the sharp increase of rectification as a function of gap distance.

We next sought to quantify the complete trajectory of cells for each experimental condition where cells can move back and forth during 2 days. We computed the bias 𝑝 per lattice unit defined as:

𝑝 = ∑ 𝑛 + -∑ 𝑛 - ∑ 𝑛 𝑖 𝑖=+,- (1)
where ni corresponds to the number of steps performed in the i direction. This readout reports how cells behave for each gap distance independently of the total duration of a trajectory. We averaged over trajectories, and the resulting < 𝑝 > confirms the behavior observed for the total bias (see Figure 1g). For 22 µm, the total bias is maximal, and the mean bias per step is also sharply increased and reaches 0.48 (± 0.08). Distributions of 𝑝 are also informative: the distribution obtained for the line of fibronectin (75% OL) corresponds to a stochastic bistable behavior with two peaks around p = 1 and p = -1 (see Figure S1) and no preferred direction. This feature progressively vanishes and bistability is transformed into a monostable distribution with one single peak located around p = 1 for 22 µm. Altogether, these results suggest that cell micro-environment architecture and simple geometrical features are sufficient to tune bistability into an almost deterministic cell directionality, and importantly, in the absence of chemical gradients. Focal adhesions (FA) are protein complexes, connected to the actomyosin meshwork, acting as an adhesive anchor and as a local force transmitter [START_REF] Riveline | Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCKindependent mechanism[END_REF][START_REF] Balaban | Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates[END_REF]. Considering the monostable condition (22 µm) being the most simple and efficient configuration, we next asked whether FAs dynamics and distribution could be sufficient to explain rectification. Using NIH3T3 fibroblasts stably expressing GFP focal-adhesion vasodilator-stimulated phosphoprotein (VASP, see Methods and Figure S2), we recorded the dynamics of adhesion sites during motion on the micro-patterned fibronectin motifs (see Movie S6). We observed that all protrusions, as soon as they touch fibronectin regions, generate FAs (see Figure 2 and Movie S6). When the first row of FAs nucleates on the neighbouring adhesive pattern, new contacts appear along the motif in a wave-like manner (see Figure S3 and Movie S7). This indicates a collective behaviour. This phenomenon on adhesive motifs is distinct from the protrusions growing on PLL-g-PEG. In this latter case, protrusion growth stops and retracts after nucleation of FAs. This suggests that FAs dynamics could be a relevant readout for cell motion. We next tested this hypothesis quantitatively. It was shown that focal adhesions act as mechanosensors and can apply a force per unit area of 5.5 nN.µm -2 (19-21), which is conserved across cell types and substrates [START_REF] Balaban | Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates[END_REF][START_REF] Goffin | Focal adhesion size controls tension-dependent recruitment of α-smooth muscle actin to stress fibers[END_REF]. We used this relationship to test whether the force inferred from focal contacts could be predictive of motion. While the global balance of forces applied on the cell is verified at all times [START_REF] Tanimoto | A Simple Force-Motion Relation for Migrating Cells Revealed by Multipole Analysis of Traction Stress[END_REF], the spatial distribution of forces is informative of local dynamics and global cell motion. We mapped force vectors at three different relevant time-points (see Figure 2). Each focal adhesion mediates a force 𝑓 𝑖 exerted by the cell on the substrate along the long axis of the focal contacts, and its orientation points towards the interior of the cell. Its magnitude is deduced from the measured area of the FA by assuming a constant stress of 5.5 nN.µm -2 (Figure 2b). From these vectors, we extracted the total force associated with each focal contact, its amplitude and its direction along x and y axis for different cells (Figure 2c,d). Cells apply a large total force of 𝐹 ⃗ = ∑ 𝑓 ⃗ 𝑖 and is about 130 nN along the x-axis during motion, 3-fold larger than prior and after motion. To satisfy global force balance, a resistive friction force has to be taken into account.

Assuming viscous friction, the corresponding friction coefficient can be estimated as ~ 25 N.s.m -1 , also consistent with reported values of friction of ~ 30 N.s.m -1 [START_REF] Palecek | Integrin-ligand binding properties govern cell migration speed through cell-substratum adhesiveness[END_REF]. This suggests that cell motion can be captured with only focal contacts dynamics translated into local forces and could be explained quantitatively by analysis of distributions of focal contacts right and left. Since every protrusion generates focal adhesions as soon as it touches the patterns (see Figure 3a), distributions of contacts, and therefore cell direction, are expected to be critically governed by adhesive area accessible to protrusions. This was quantified by a simple phenomenological model that expresses explicitly the mean bias per step as a function of adhesive area accessible to protrusions on each side. Assuming a linear dependence of probabilities of formation of a focal adhesion with the adhesive area probed by protrusions made it possible to derive a simple expression of <𝑝> involving only measurable geometric parameters ; this however yielded a poor agreement with the plots of <𝑝> as a function of gap distance (see Figure 3b and Theory). This indicates that collective effects in focal adhesion formation, as observed (see above) must be taken into account. To do so, we introduced the nucleation area A0 (see Theory), a phenomenological parameter representing the typical area that cells need to engage in a neighboring motif to efficiently contribute to motion. For A< A0, detachment of the new adhesive zone occurs. Above this area, the nucleating zone is stable and a wave of new contacts is generated. The resulting prediction of < 𝑝 > yielded an excellent agreement with experimental values (see Figure 3b), where notably the single fitting parameter A0 was involved.

We turned back to experiments of focal contact dynamics: the nucleating area A0 obtained from the fit of 13.5 µm² indeed matched the nucleation area in experiments (see Figure 3c). This suggests that a nucleation zone is needed to sustain proper anchor on neighboring motifs, consistent with the notion that a collection of focal contacts is required to perform firm cellular anchors on neighboring motifs. Importantly, the model involves only 2 cellular parameters that can be directly measured, and no ad hoc adjustable parameter was involved to recapitulate the trend observed in experiments. This supports the relevance of our hypothesis. If the area available on both sides is greater than A0, the cell will be able to create stable anchoring zones sufficient to nucleate the subsequent rows of FAs. Therefore, the bias will be poor even if A+ >> A-(13 µm to 19 µm conditions, see Figure 3d). On the other hand, if one side goes close or below A0, the bias becomes stronger in the opposite direction. This kind of coupling between geometry and cellular features could be relevant in vivo where distributions of adhesive regions are not homogeneous (Figure 3d and ( 23)). Ratchetaxis relevance could be probed in the future in vivo, along lines recently suggested [START_REF] Bajanca | In vivo topology converts competition for cell-matrix adhesion into directional migration[END_REF]. To test this framework, anisotropy in adhesion sites need to be tracked. In addition, protrusions and cell contacts dynamics have to be acquired and correlated over time quantitatively, together with long term cell motion. These readouts are now accessible in vivo for a variety of model systems, but their simultaneous observations and correlations are not performed yet to our knowledge. New experiments will need to be conducted to establish this potential source of rectification for cell migration in vivo.

In addition to its novel way of operating, ratchetaxis encodes cellular mechanisms which convey new ways of perceiving cell motion with readouts extracted from biological physics. Cell polarity is acquired in the first steps through interactions with environment, and then a persistence in polarity is kept over time by the cell. Eventually, this translates into the mean direction taken by cells through collective effect of focal contacts. These parameters, polarity, persistence of polarity, and friction quantified through focal contacts, can be integrated in predictive models, as we reported in this study. Also, these steps and associated parameters are all stochastic, and their distributions are informative to test hypothesis. Altogether, ratchetaxis could also serve as a key phenomenon to test physical models for living matter.

NIH3T3 mouse fibroblasts (ATCC) were cultured in high glucose Dulbecco's Modified Eagle's Medium (DMEM) (Fisher Scientific, cat. 11574486) supplemented with 10% Bovine Calf Serum (BCS, Sigma, cat. 12133C) and 1% Penicillin-Streptamycin. Cells were detached from the Petri dish with Trypsin-0.25% EDTA (Fisher Scientific, cat. 11570626), centrifuged, and re-suspended in DMEM 10% BCS. About 2000 cells were seeded on micro-patterns and the non-adhering cells were washed out after 30 min of incubation at 37°C and 5% CO2. Medium was replaced by L-15 (Leibovitz Medium, Fisher Scientific, cat. 11540556) supplemented with 1% of BCS in order to reduce cell division during experiments.

Focal adhesions, named also focal contacts, were monitored by fluorescence microscopy. NIH3T3 mouse fibroblasts stably expressing a VASP-GFP vector were kindly provided by Olivier Pertz (Institute of Cell Biology, Bern University, Switzerland) [START_REF] Martin | Spatio-temporal co-ordination of RhoA, Rac1 and Cdc42 activation during prototypical edge protrusion and retraction dynamics[END_REF]. The construct is expressed in lentivirus derived from pLenti CMV MVS. The viruses were added to a culture of wild-type NIH3T3 and cells were selected after 24 h with 2 µg/mL of puromycin.

We checked that GFP-VASP was a direct and reliable readout of focal adhesions (see Methods). This was confirmed by immuno-staining cells expressing GFP-VASP with paxillin. Cells were fixed with 4% paraformaldehyde for 8 min. Then, they were treated with Triton 0.5X during 3 min. After washout, an anti-paxillin mouse primary antibody (BD transduction, cat. 610051) was used together with an anti-mouse secondary antibody grafted with Cy3 (Jackson Immuno Research, cat. 115-165-146). We observed co-localisation providing us good evidence that GFP-VASP reveals focal contactssee also results with RICM below.

Time-Lapse microscopy

Long-term phase-contrast images (48 h) were acquired using a low magnification objective (4x, N.A. 0.25) with an image acquisition frequency of 1 image each 5 min. Micro-patterns were visualised by means of a standard epifluorescence lamp (FluoArc Hg Lamp) coupled with a rhodamine fluorescence filter. A thin layer of mineral oil (Sigma, M5904) was deposited on top of the cell culture media in order to prevent medium evaporation. In Figure 1, cells were imaged using a 10x objective (N.A. 0.4). Focal contacts dynamics were monitored every 3 min with a Nikon Spinning-Disk confocal microscope with a 60x oil objective (N.A. 1.4) using the Perfect Focus System. These settings provided highly resolved images with a low phototoxicity for long-term experiments (15 h). To test whether VASP-GFP protein revealed focal contacts localisations, we used Reflection Interference Contrast Microscopy (RICM) as described in Curtis et al. [START_REF] Curtis | The mechanism of adhesion of cells to glass[END_REF]. We used a homogeneous white light source (FluoArc Hg Lamp) through a 50% dichroic and imaged interferences on the surface (see Figure S2b).

Image processing

Tracking of the cells (nucleus) was performed with ImageJ using Multiple Tracking plug-in. The generated coordinates were used to compute cell trajectories. Plots derived from data as well as statistical tests were done using OriginLab Pro (OriginLab). Force analysis made on focal contacts was performed with a custom Python code computing areas of adhesion zones, their orientations and barycenters, force vectors for each of them and total force vectors taking the rule 1 µm² corresponds to 5.5 nN [START_REF] Balaban | Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates[END_REF][START_REF] Goffin | Focal adhesion size controls tension-dependent recruitment of α-smooth muscle actin to stress fibers[END_REF]. Fits with experimental data were done through OriginLab Pro.

Theory

At low Reynolds number, force balance for each cell can be written [START_REF] Tanimoto | A Simple Force-Motion Relation for Migrating Cells Revealed by Multipole Analysis of Traction Stress[END_REF]:

∑ 𝐹 ⃗ 𝑖 = 𝜇(𝑣)𝑣 ⃑ 𝑖 ( 2
)
where 𝐹 ⃗ 𝑖 denotes the force exerted by the substrate on the cell through a given focal adhesion i, and 𝜇, which can be non-linear, effectively accounts for all friction forces resulting from the interaction of cell body with substrate. Importantly, the direction of motion is here fully determined by the knowledge of 𝐹 ⃗ 𝑖 , which can be determined quantitatively from direct imaging of focal adhesions, as argued above, and be used as a tool to predict the direction of motion.

Projecting Eq. 1 along the ratchet axis and averaging over time yields the following relation between the bias per step and mean number of focal adhesions 𝑛 + (𝑟𝑒𝑠𝑝. 𝑛 -) in the + (resp. -) direction: 𝑝 ∝ (𝑛 + -𝑛 -)/(𝑛 + + 𝑛 -). Introducing now the purely geometric quantities A+ and A-, defined as the area accessible to protrusions and FAs in each direction, we assume that 𝑛 +,-∝ 𝑛𝐴 +,-(other functional form, provided that n vanishes with A, would yield qualitatively similar results). This yields finally to:

𝑝 ∝ (𝐴 + -𝐴 -) (𝐴 + + 𝐴 -) (3)
which elucidates the dependence of the bias on geometry. Interestingly, the estimated ratio 𝐴 + 𝐴 -is always greater than 1 for protrusions lengths compatible with NIH3T3 cells and shows a monotonic behavior coupled to a strong divergence when the gap distance is close to the mean protrusion length λ (see Figure S4). When λ is large enough (i.e. close to the size of a pattern), the ratio of probed areas goes back to 1.

When we include the nucleation area A0, we used the following phenomenological expression of p:

𝑝 ∝ 𝑡𝑎𝑛ℎ ( 𝐴 + 𝐴 0 ) -𝑡𝑎𝑛ℎ ( 𝐴 - 𝐴 0 ) ∑ 𝑡𝑎𝑛ℎ ( 𝐴 𝑖 𝐴 0 ) 𝑖=+,- (4)
A0 can be thought as the critical area above which there is no significant bias because the cell has the opportunity to create a sufficient stable anchoring region to nucleate the subsequent rows of focal contacts on both sides, even if A+ >> A-(see Figure 3d and Movie S7). The value of A0 in our model corresponds to 13.5 µm², and this strikingly corresponds to a single row of focal contacts areas on patterns. Interestingly, Ai is determined with no fitting parameters from the knowledge of the mean protrusion length and the geometry of the patterns. With these minimal number of parameters, we can qualitatively reproduce observations and quantitatively predict a sharp increase of the bias for a gap of 22 mm, as observed in experiments (Figure 3c). The parameter λ, embedded into the computations of Ai, corresponds to the typical length of protrusions (essentially between 20 µm and 35 µm in our experiments for NIH 3T3). To fit our model with experiments, we used λ = 28 µm. Therefore, fitting parameters encode actual cellular quantities, and no ad hoc adjustable parameters or global pre-factors are added to fit the measures extracted from experiments. This supports the relevance of our hypothesis.

Figure 1 :

 1 Figure 1: Cell motion depends on gap distance. (a) Scheme showing the experimental setup where a cell is seeded on a microcontact printed array of asymmetric FN triangles. The gap distance is systematically increased, which determines the final direction of migration. Quantitatively, p describes the bias in directionality taking into account all steps made during the motion. (b-e) Cells are able to move from one motif to another on separated triangles, and crawl on connected ones. Time in hh:mm, scale bars: 50 µm. (f) Final bias in percentage as a function of gap distance. (g) Average p given by Eq. 1 for each condition. (h) Persistence speed in µm.h -1 . Data set: 25 < Ncells < 30 and Nbiological > 5 for 75% OL, 25% OL, 13µm, 16µm, 19µm and 22µm. Ncells = 16 for 20.5 µm. Data is shown as Mean ± S.E.

Figure 2 :

 2 Figure 2: Force inference based on focal contacts distributions correlates with motion. (a) Time-lapse images of an NIH3T3 cell stably expressing VASP-GFP. Three time points are extracted for analysis: (i) during the stationary phase, (ii) at the onset of motion, and (iii) at the end of motion. (b) Force vectors map. Each focal contact is linked to a force vector oriented in the direction of adhesion zone (in green). Its amplitude is set according to the relationship 1 µm² corresponds to 5.5 nN. The total force vector is displayed in red, its x component in blue; (c-d) Force exerted along x-axis and y-axis during the three time points. Statistical tests comparing distributions are done with a one-way ANOVA. Nbiological_repeat = 4, Ncells = 8. Scale bar = 15µm.

Figure 3 :

 3 Figure 3: The source of the bias. (a) Protrusions touching an adhesive motif imaged with fluorescence microscopy and RICM. Focal adhesions are nucleated right after contact with motifs. Arrows indicate FA on patterns. (b) Comparisons between model and experiments. The blue fit assumes mechanisms with areas probed by cells, whereas the red fit includes the nucleation area A0. Fit corresponds to A0=13.5 µm² and λ= 28 µm. (c) Experimental description of A0. From this region, subsequent rows of focal adhesions are nucleated. Arrows indicate direction of nucleation. (d) Graphical description of A0 and its influence on bias. Beyond A0, the bias is weak. If one side gets close or below A0, the bias in the opposite direction becomes strong. Time in hh:mm, scale bars = 15µm

ACKNOWLEDGMENTS

We thank O. Pertz, P. Rossolilo and A. Dupont, the Imaging Platform of IGBMC, the Riveline Lab. for constructs and discussions. This study with the reference ANR-10-LABX-0030-INRT has been also supported by a French state fund through the Agence Nationale de la Recherche under the frame programme Investissements d'Avenir labelled ANR-10-IDEX-0002-02. SLV is supported by the University of Strasbourg. R.T. was an IGBMC International PhD Programme fellow supported by LabEx INRT funds. D.R. acknowledges support from ciFRC Strasbourg, the University of Strasbourg, CNRS and Foundation Cino del Duca.

Conflicts of interest: none.

Materials and Methods:

Micro-contact printing Fibronectin asymmetric motifs were patterned using standard microcontact printing protocol [START_REF] Théry | Adhesive micropatterns for cells: a microcontact printing protocol[END_REF][START_REF] Maout | Directing cell migration on flat substrates and in confinement with microfabrication and microfluidics[END_REF]. Briefly, polydimethylsiloxane (PDMS, 1:10 w/w cross-linker:prepolymer) (Sylgard 184 kit, Dow Corning, cat. DC184-1.1) stamps were replicated from SU-8 molds fabricated by standard UV-photolithography. After cleaning in ethanol 70%, stamps were rendered hydrophilic by oxygen plasma activation (Diener Electronic, cat. ZeptoB) for 30 s. Then, stamps were incubated for 1h with a 10 µg/mL rhodamine-labelled fibronectin solution (Cytoskeleton, cat. FNR01-A) and dried under a stream of nitrogen at room temperature for about 5 min. Next, stamps were placed in contact on top of a glass coverslip functionalized by vapour phase for 1 h with 3-(mercapto)propyltrimethoxysilane (FluroChem). A 50 g weight was placed on the top of the stamp during 30 min to ensure a constant and isotropic pressure during stamping. After release, the coverslip was cleaned in PBS 1x. The non-patterned areas were passivated with a 0.1 mg/mL solution of PLL-g-PEG (in 1 mM HEPES pH 7.4, SuSoS AG, cat. SZ33-15) for 20 min at room temperature. Finally, the patterned glass coverslips were again rinsed in PBS 1x prior cell deposition.

Cell Culture


Alka Bhat

Linjie Lu

Chen-Ho Wang

Simon Lo Vecchio

Riccardo Maraspini

Alf Honigmann

Daniel Riveline

How to orient cells in microcavities for high resolution imaging of cytokinesis and lumen formation

Keywords: Biological Physics, Microfabrication, Super-resolution STED, Cytokinesis, Cell Polarization, Organoids

Imaging dynamics of cellular morphogenesis with high spatial-temporal resolution in 3D is challenging, due to the low spatial resolution along the optical axis and photo-toxicity. However, some cellular structures are planar and hence 2D imaging should be sufficient, provided that the structure of interest can be oriented with respect to the optical axis of the microscope. Here, we report a 3D microfabrication method which positions and orients cell divisions very close to the microscope coverglass. We use this approach to study cytokinesis in fission yeasts and polarization to lumen formation in mammalian epithelial cells. We show that this method improves spatial resolution on range of common microscopies, including super-resolution STED. Altogether, this method could shed new lights on self-organization phenomena in single cells and 3D cell culture systems.

Introduction

Cells have been cultured traditionally on Petri dishes [START_REF] Petri | A small modification of Koch's Plate Method[END_REF] where they can grow easily. In addition, adherent cell culture facilitates microscopy observation within single or a couple of focal planes. This 2D approach has become the standard method to culture and to observe cells and multicellular systems, in basic research, in drug discovery, and in medicine for diagnosis.

However, this method is not optimal for many reasons [START_REF] Abbott | Biology's new dimension[END_REF] . Cells are artificially 'forced' to spread on planar and flat surfaces, potentially inducing artificial shapes and pathological signaling pathways compared to environments encountered in physiological conditions [START_REF] Weaver | Reversion of the malignant phenotype of human breast cells in three-dimensional culture and in vivo by integrin blocking antibodies[END_REF] [START_REF] Anders | Disruption of 3D tissue integrity facilitates adenovirus infection by deregulating the coxsackievirus and adenovirus receptor[END_REF]. Cells evolve more physiologically in 3D environments, and classical Petri dishes may not reproduce faithfully this condition. However, 3D cell culture methods, in which cells are typically embedded in hydrogels, pose new challenges to our standard microscope observation techniques. In 3D and sometime also in 2D cell culture many structures/organelles will be ill-oriented with respect to the plane of observation for cells. Placing these organelles within the focal plane of objectives could lead to gain in resolutions in space and in time. These limitations have generated the need to design 3D structures closer to physiological conditions.

In this context, microfabrication has allowed to prepare 3D environments which are compatible with cell growth and organ designs (Sontheimer-Phelps, Hassell, & Ingber, 2019) [START_REF] Eyckmans | 3D culture models of tissues under tension[END_REF]. Shapes in 3D are generated with micrometer resolution using protocols which can be performed with simple training (see Figure 1). We reported since 2009 [START_REF] Riveline | Devices and methods for observing the cell division[END_REF] [START_REF] Wollrab | Devices and methods for observing eukaryotic cells without cell wall[END_REF] [START_REF] Riveline | Methods for observing cells with cell wall or invertebrate embryos with oblong eggshell[END_REF] [START_REF] Wollrab | Ordering Single Cells and Single Embryos in 3D Confinement: A New Device for High Content Screening[END_REF] the preparation of cavities, where cells can be placed vertically with respect to the coverglass. In these 3D configurations, cell division occurs also in a direction perpendicular to the long axis of the cell, but structures such as cytokinetic rings can be seen with unprecedented resolutions. In standard coverslips, rings are perpendicular to the plane of acquisition of objectives: each plane of image capture gives two points, and full rings visualization requires multiple z-stacks acquisitions and 3D reconstruction. In contrast, in these cavities, cytokinetic rings can be seen 'en face' in a single focal plane of acquisition (see Figure 2). Any close to planar cellular structures with such orientations can in principle be imaged with this device and this method.

In this paper, we describe our different procedures and strategies to orient cell structures with 3D microfabrication. We also go one step further and prepare 3D cavities, which are optimized for super-resolution microscopy. We benchmark our system by imaging the cytokinetic rings in fission yeast and in mammalian cells, as well as the polarization and lumen formation in mammalian MDCK cells after the first cell division. The gain in resolution due to the positioning close to the coverglass and the orientation along the optical axis is significant in time and in space (see Figure 3). For example, clusters of myosin could be resolved with this method with resolution of 200nm with conventional confocal microscopy. They were visualized either by inverted microscope or by having cells imaged directly from below with no interface (Figure 3B). Through-hole cavities allowed us the usage of super-resolution STED microscopy for cytokinetic rings and lumen (Figure 3B.ii and Figure 4) with high NA objectives and minimal optical aberrations. The gain in temporal resolution is also important since organelles can be imaged within sub-seconds in their planes of evolution (see Figure 3C). For example, we reported thanks to this method that myosin clusters were still during cytokinesis in the cell framework. This unexpected observation and its analysis and comparisons with theory and other systems led to the conclusions that myosin clusters generate stress which is associated to the ring constriction in mammalian cells. This supported the notion that myosin clusters dynamics and their self-organizations can serve as generic readouts in cells and in model systems to track and predict changes in shapes [START_REF] Wollrab | Still and rotating myosin clusters determine cytokinetic ring constriction[END_REF]. Altogether, these results show that cells inserted vertically in cavity can resolve spatial and temporal cues not accessible so far.

We report below the steps for preparation of 3D micro-patterns by taking the following outline.

We start by describing the design and preparation of masks (I), and continue by the fabrication of motifs (II and III). We then explain how to place cells within cavities (V). Troubleshooting is reported in part VI.

I.

Design of mask for optimized cavity preparation.

Masks will serve as support to expose and protect resins from light during the process. In their designs, motifs and inter-distances need to be carefully rationalized.

For each cellular system, cavity dimensions need to be matched to cell dimensions. The first step consists of measuring the distribution of cell diameter right after trypsinization for mammalian cells, and plating cells on a coverslip coated with fibronectin (typically 10g/ml for 1 hour at room temperature). Distributions for cell diameter can be very sharp like for fission yeasts, and broader for mammalian cells. The mean value is a good starting point for selecting the disk dimension to be prepared on masks.

In addition to cavity diameter, the inter-distance between motifs is important. Inter-distance determines number of cavities per sample. In addition, as reported below, low inter-distance secures a satisfactory filling percentage of cells in cavities. However, this inter-distance should not be too low in order to secure firm attachment of the elastomer stamp. A rule of a thumb consists in taking inter-distance at least twice the cavity size.

After the designed motifs are printed and ready on a photomask, microfabrication process of the motifs follows. The mask prepared in our case had diameters of 17µm through 22µm with an interdistance of 30µm for mammalian cells or 5µm for fission yeast. Photomasks can be designed using software such as Clewin (Freeware) or Autocad®. Disks are plain or transparent depending on the type of photoresist used in the next steps.

II. Microfabrication of motifs on Si wafer.

After the photomask is ready, a light sensitive epoxy-based material (photoresist) is used to obtain motifs on Si wafer. The photoresist (SU-8 in our case) is spin-coated on the wafer with specific coating time and speed to obtain the target height for cavities. In this part, we report SU-8 pillars which are fabricated on Si wafers.

The average diameters for HeLa and MDCK cells were measured to be ∼20μm and ∼15μm

respectively. Therefore, the chosen diameter/height in our case were 22μm/25μm and 17μm/12μm

for Hela cells and MDCK cells respectively. The preparation of cavities for fission yeasts is reported in this reference [START_REF] Riveline | Methods for observing cells with cell wall or invertebrate embryos with oblong eggshell[END_REF] with special conical shapes to secure firm anchorage of cells. Steps for photolithography:

Materials

1. To ensure that the silicon wafer is contaminant free, it is first cleaned with acetone. It is then dried with a nitrogen stream and followed by similar treatment with ethanol and drying process.

Solvent cleaning ensures complete removal of oils and organic residues from the wafer surface.

After solvent cleaning, it is advisable to heat up the wafers at 200°C for 10-20 minutes for complete removal of humidity from the surface of wafer.

2. To obtain the targeted height, pour the photoresist (SU-8 2025 for 25µm, and SU-8 2010 for 10µm) on the wafer and remove air bubbles with the help of a plastic dropper. Set a two-step spinning process on the spin-coater. Spin at 500rpm for 10s with an acceleration of 100 rpm/s during the first step to homogenize the layer over the wafer for attaining both 25µm and 12µm height. For a height of 25µm, spin the wafer containing resin at 3000rpm for 30s with acceleration of 300 rpm/s for the second step (WS-650-23 Spin Coater; Laurell Technologies corporation, centrifuge and rotor details kept throughout for microfabrication). For a height of 12µm, spin the wafer containing the resin (SU-8 2010) at 3500rpm for 30s with acceleration of 1000rpm/s. This should result in a homogeneous photoresist layer of the required thickness.

3. After obtaining the photoresist evenly spread, pre-bake has to be done to ensure a firm attachment of the photoresist to the wafer. This step has to be performed for 5min at 95°C.

4. After pre-bake, make a firm contact between the wafer and the mask containing the respective motifs. A defective contact will lead to ill-defined motifs. After tight contact with the mask, cross-link the photoresist by exposing the wafer to UV irradiation with a dose of 150 mJ/cm 2 .

The time of exposure will depend on the power of the device (e.g. mask aligner) for the UV light exposure (see Figure 1A.i). 5. As soon as the exposure is over, follow a post-bake of 1min at 65°C and then 5min at 95°C. An image of the mask should appear on the SU-8 photoresist coating.

Figure 1: Microfabrication and preparation of cavities. A. Microfabrication of a silicon master with photoresist SU-8 (A.; (i)) to obtain SU-8 pillars (A.; (ii)). Un-crosslinked PDMS is poured over the silicon master (A.; (iii)) and peeled off to give PDMS cavities (A.; (iv)). B. PDMS cavity preparation with polymer (PDMS) layer at the bottom by first silanizing the PDMS cavities with TMCS (B.; (i)) followed by spincoating a drop of PDMS (B.; (ii)) to obtain uniform thin layer and crosslinking at 65°C O/N. The assembly is plasma-bonded with a glass coverslip and pressed uniformly (B.; (iii)). Crosslinked PDMS is peeled off from the assembly revealing cavities with PMDS layer at the bottom (B.; (iv)). C. PDMS cavities without polymer layer at

6. After post-bake, develop the structures by immersing them in the SU-8 developer solution, while gently agitating the container for around 4min. This will strip off the non-crosslinked resin, leaving behind the motifs. Finally, rinse the surface with 2-propanol to remove the leftover photoresist.

7. White left-over seen over the wafers during 2-propanol treatment is an indicator for residual un-crosslinked resin. Repeat step 6 until rest of the non-crosslinked resin is stripped off the wafer completely.

8. Rinse with 2-propanol and dry the wafer containing motifs using a nitrogen air stream. 9. After the silicon wafers are ready with desired motifs, do a 'hard bake' step at ~150°C for 10 minutes. This is useful for annealing any surface cracks that may have appeared after development; this step is relevant to all layer thicknesses. Structures fabricated in such a way have excellent thermal and mechanical stability (see Figure 1A.ii).

10.

After obtaining the motifs, cover the wafers with PDMS (crosslinker: base [1:9]) (see

Figure 1A.iii).
11. Remove all air bubbles through desiccation, and finally cure it overnight at 65°C on a leveled surface.

12.

Once the elastomer is cured, peel off the PDMS block containing the desired cylindrical pillars with appropriate height (see Figure 1A.iv).

III. Process for cavity preparation

Cavities are prepared using polymers such as PDMS (heat-crosslinking polymer) or NOA-74 (UVcrosslinking polymer). Different steps are taken to achieve cavities with a thin polymer layer at the bottom, as well as through-hole cavities with no bottom polymer layer. (i) Cavities with polymer layer at the bottom:

Materials

PDMS base and the curing agent are mixed in a ratio of 9:1 (v/v). This liquid PDMS is poured over the silicon master obtained via photolithography (for HeLa and MDCK cells), and silicon master is obtained via deep reactive ion etching (for fission yeast, [START_REF] Riveline | Methods for observing cells with cell wall or invertebrate embryos with oblong eggshell[END_REF].

1. Liquid PDMS poured over the silicon wafers is degassed (removing trapped air inside the liquid polymer) and cured at 65°C overnight.

2. After curing by heat, the PDMS layer is peeled off (see Figure 1A.iv) from the wafer surface.

This peeled off layer is considered as Master 1 with holes in the PDMS block. This is used as replica mold to fabricate pillars used as final motifs for the preparation of cavity.

3. Next, PDMS block with pillars is plasma activated and subsequently passivated with chlorotrimethyl silane (TMCS) under a desiccator for 10min.

4. Small amount of uncured liquid PDMS (~50µl) is poured over the motifs with subsequent spincoating (and vacuum to hold the PDMS block) with 500rpm for 10s (to homogeneously distribute the liquid PDMS drop) 1700rpm for 45s to obtain the preferred height (~ 40µm)

(see Figure 1B.ii). (KW-4A Spin coater; SPI supplies, centrifuge and rotor details kept throughout for Section III (i)). This thickness will be corresponding to the polymer layer at the bottom of glass coverslip.

5. This combination of uncured polymer over PDMS block (containing pillars) is then incubated at 65°C overnight for curing.

6. Cured PDMS block and glass coverslip #0 (100µm thickness) are activated under oxygen plasma. In addition, PDMS block is bonded with the glass coverslip (coated side down). This configuration is pressed for several seconds to ensure tight bonding between them (see Figure 1B.iii).

7. This coverslip-PDMS 'sandwich' is incubated at 65°C overnight. Longer exposure to heat helps the bonding to be stronger between PDMS and coverslip.

8. Next, coverslip is held on both sides of the PDMS block, while peeling it off carefully. This avoids the risk of coverslip breakage during the process. 9. Thin bottom layer of PDMS on the coverslip (~ 40µm) with cavities on top is revealed (see Figure 1B.iv) and Figure 3A.i).

Even though cells are oriented in the right orientation with a good match between cell and cavity dimensions, an elastomer layer at the bottom decreases resolution. This leads to loss in fluorescence as well as in spatial resolution.

A typical example is shown with cytokinetic ring of HeLa cells observed using such cavities with polymer layer at the bottom and an inverted setup: the ring is blurred (see Figure 3A.iia-b). This blurring effect is due to distortion of light caused by this thin PDMS layer at the bottom. Therefore, for such configurations, it is better to visualize these structures with an upright microscope (see Figure 3A.iiia-b) and this allowed to reveal regularly spaced (0.8m) myosin clusters around the cytokinetic ring (Wollrab et al., 2016). Visualizing cells in such configuration does not only increase spatial resolution, but also temporal resolution for capturing fast dynamics (e.g. for cytokinesis in HeLa cells; see Figure 3C.i-ii).

To be able to visualize structures with higher resolutions, cavities without any polymer layer at the bottom (see Figure 1C.iv and Figure 3B.i) are fabricated and cells are visualized using STED super-resolution microscopy (see Figure 3B.iia-b). (Wollrab, 2016) (ii) Cavities without polymer layer at the bottom.

Cells directly 'touching' the glass surface -without any polymer layer at the bottom -have a significant advantage to gain in resolutions during image acquisition. In addition, this configuration also provides higher freedom of objective working distance as well as the ease in using both upright and inverted microscope setups. Through-hole cavities are achieved with both polymers mentioned earlier, PDMS and NOA-74.

1. Steps from section III (i) steps 1 and 2 are repeated to obtain Master 1 (PDMS block with holes).

2. To obtain PDMS pillars (final motifs for cavity preparation), Master 1 is activated under oxygen plasma. Next, it is passivated with chlorotrimethyl silane (TMCS) under a desiccator for 20-30 mins (see Figure 1B.i). Post silanization PDMS is poured over the Master 1 (with holes), degassed and cured at 65°C overnight.

3. After curing, PDMS is peeled off the Master 1 (PDMS with holes) revealing PDMS block with pillars of target height.

4. To obtain cavities without PDMS layer at the bottom, first, blocks of PDMS with pillars (0.5cm X 0.5cm) are cut and sonicated (5min) with 70% ethanol to remove any dirt and residual silane.

5. These PDMS blocks (with pillars) are used for the preparation of cavities for fission yeast cells, as well as through-hole NOA-74 cavities for HeLa and MDCK cells.

6. Surfaces of PDMS pillars are activated under oxygen plasma, along with the glass coverslip on which pillars are to be bonded.

7. Activated PDMS block with pillars is now turned (activated side down) and put on top of the activated coverslip. The assembly is carefully pressed to ensure proper bonding. Inadequate boding would lead to non through-hole cavities (see Section VI and Figure 5D). Excessive strong bond will result in blocked cavities (see Section VI and Figure 5C). Care is taken that the pressure does not tilt these pillars (see Section VI, Figure 1C.ii and Figure 5A).

8. The assembly is then incubated for passivation with TMCS in a desiccator for 20-30 mins.

Passivation is done to prevent irreversible bonding of PDMS pillars with the introduced polymer (NOA-74). 9. Post-passivation: the polymer (NOA-74) is introduced as a drop on one side of the assembly.

Capillary action pulls the polymer into the assembly (see Figure 1C.iii). Speed of polymer flow throughout the assembly is dependent on height of the pillars as well as viscosity of the polymer. For example, NOA-74 is less viscous than PDMS and will 'travel' through the assembly quicker and more efficiently than PDMS. Desiccator can be used for 5-7 min for efficient and homogeneous filling of pillars, removing any air bubble trapped inside the assembly.

10. Polymer NOA-74 incubated assembly is then placed for curing (crosslinking) inside a chamber with UV-A source for minimum 4-6 hours. The same procedures 9-11 can be used to prepare PDMS cavities as well with no bottom layers.

A typical example of cells inside such configuration, imaged with an inverted STED superresolution microscope, is shown where myosin can be observed at higher resolution (see IV. Choosing the optimal polymer for the process: advantages and drawbacks of different polymers

As described above, two types of polymers, PDMS and NOA-74 are used. There are advantages and drawbacks for each polymer type. For imaging cells inside cavities, it is important to use a biocompatible polymers [START_REF] Sia | Microfluidic devices fabricated in Poly(dimethylsiloxane) for biological studies[END_REF]) [START_REF] Masters | Easy Fabrication of Thin Membranes with Through Holes. Application to Protein Patterning[END_REF].

Also, polymer viscosity can be critical to prepare cavities with small heights. Such configuration would require shorter PDMS pillars, plasma-bonded to coverslip. Incubation of this assembly with a comparatively lower viscosity polymer (80-95cP; NOA-74) is much easier than a polymer with higher viscosity (3500cP; PDMS). If a high viscosity polymer is required, polymer flooding with desiccation is needed. This helps to secure the process with an even distribution of the polymer.

Next, to obtain optimal resolution, the cured polymer with refractive index closer to glass coverslip (~ 1.52) must be chosen. This minimizes distortion of light due to passage through the polymer layer. Refractive indices of both PDMS (~1.41) and NOA-74 (~1.52) are close to glass, with NOA-74 being a better candidate.

Cured PDMS polymer forms strong attachment with glass coverslip. Such property is preferred when robust and long-lasting and/or permanent attachment with coverslip are required. If the polymer layer is to be detached from the surface, polymers with low modulus (NOA-74) are preferred. This allows complete detachment of the layer from the substrate, while keeping its original structure after sonication (10min).

V. Cell seeding and visualization

We next report the last steps prior observations:

Mammalian cells (HeLa and MDCK):

1. Small rectangular piece of PDMS is attached to the coverslip (with cavities) with a drop of uncrosslinked PDMS. This assembly is incubated inside the oven (65°C) for 4h. Such PDMS block will serve as a holder to handle the cavities with help of tweezers in the subsequent steps.

2. Cavities are plasma activated under oxygen plasma.

3. Cavities further immersed in fibronectin (10g/ml) for 4h at room temperature.

4. Cylindrical PMMA piece is introduced inside 50ml tube.

5. Culture media is introduced into the tube until it forms a thin layer over the PMMA piece. ((i); scale bar 40µm) leads to wrong orientation of dividing cells with respect to the visualization plane ((ii); scale bar 10 µm). [START_REF] Wollrab | Active gels in vivo : Patterns and dynamics in cytokinetic rings and their functions in cell divison[END_REF].

6. After the centrifugation steps, cavities are carefully removed and put into a metal holder with selected culture medium, and the sample is ready for acquisition.

VI. Troubleshooting for cavity preparation

We list below typical issues during protocols:

 Tilted PDMS pillars leading to tilted cavities due to pressing during plasma bonding Excessive pressing during plasma bonding (see section III. (ii), step 7.) of coverslip with PDMS pillars can result in tilted pillars (see Figure 5A). Consequently, cells and/or structures under observations would not appear at the correct focus and orientation, leading to poor image resolution.

 Inhomogeneous cavity distribution due to trapped air bubble

In order to achieve homogeneous (NOA-74) prepolymer distribution, one should make sure to introduce the prepolymer on one side of the assembly as a small drop (~10-20l) and left on the bench for the capillary action to fill the whole assembly. In contrast, polymer introduced from more than one sides could trap air bubble (see section III. (ii), step 9 and Figure 5B).

 Blocked cavities with PDMS left behind

In case of excessive strong bond between coverslip and PDMS pillars (see section III. (ii), step 7.), peeling of PDMS pillars (after NOA-74 crosslinking) may leave PDMS blocked inside the cavities leading to obstructed cavities (see Figure 5C).

 Polymer layer at the bottom inadequate plasma activation leading to poor bonding

Poor plasma bonding between coverslip and PDMS pillars (see section III. (ii), step 7.) could lead to insertion of NOA-74 prepolymer below PDMS pillars leading to non-through-hole cavities (see Figure 5D).

 Wrong cavity size

Cavities should be designed according to the measured, average cell diameter of cells (see Section I). Failing to do so would lead to either horizontal (see Figure 5E.i) or tilted cells (see Figure 5E.ii) inside cavities. This would prevent optimized microscopy and correct orientation of cells and structures under study.

VII. Conclusions

This article reports strategies to prepare cavities optimized for orientation of cellular structures into the high-resolution plane of microscopes. Many cellular structures could be placed in the 'right' plane with our method. In fact, the usage of 3D environments for studying cells appear to be the natural way to reproduce situations encountered in physiological conditions. The cost is minimal as well as the need for specialized equipment. It is easy to foresee numerous applications of this new point of view in a literal sense to reveal new structures and dynamics with relevance for basic research and for medical applications.

  Figure 1: Microfabrication and preparation of cavities. A. Microfabrication of a silicon master with photoresist SU-8 (A.; (i)) to obtain SU-8 pillars (A.; (ii)). Un-crosslinked PDMS is poured over the silicon master (A.; (iii)) and peeled off to give PDMS cavities (A.; (iv)). B. PDMS cavity preparation with polymer (PDMS) layer at the bottom by first silanizing the PDMS cavities with TMCS (B.; (i)) followed by spincoating a drop of PDMS (B.; (ii)) to obtain uniform thin layer and crosslinking at 65°C O/N. The assembly is plasma-bonded with a glass coverslip and pressed uniformly (B.; (iii)). Crosslinked PDMS is peeled off from the assembly revealing cavities with PMDS layer at the bottom (B.; (iv)). C. PDMS cavities without polymer layer at the bottom is prepared using UV crosslinking polymer NOA-74 by first using the silanized PDMS master 1 (B.; (i)) with holes and pouring non crosslinked polymer over it, and the sample is allowed to polymerize at 65°C O/N. After polymerization, the PDMS mold with pillars is peeled off from the assembly (C; (i)) and plasma bonded to a glass coverslip (C.; (ii)). The assembly is exposed to TMCS for silanization. Next, the assembly is incubated with (iii)) and polymerized under UV-A light for minimum 4 hours. PDMS mold with pillars is peeled off carefully from the glass coverslip revealing through-hole cavities (C.; (iv)). Same process can be used to obtain both PDMS and NOA through-hole cavities.



    PDMS (Base and crosslinker; Sylgard 184 Silicone Elastomer)

Figure 2 :

 2 Figure 2: Orienting cells lead to optimal resolutions in space for a variety of cellular systems. A. Fission yeast cytokinetic ring (tagged-myosin regulatory light chain RLC) visualized on a flat surface (glass coverslip), single plane imaging shows two points in the 'xy' plane corresponding to the ring (A. (i); scale bar 3µm). Reconstitution of ring with 'z' planes projections showing blurred tilted ring (A. (i); scale bar 3µm). Reconstitution of 'yz' planes, (A. (iib); scale bar 3µm) showing a blurred ring and with sectional cut 'en face' (green: pxl1 red for the ring: blankophor for the cell wall). Vertical cells(A. (iiia); scale bar 3µm) allow to reveal myosin clusters around the ring perimeter[START_REF] Riveline | Devices and methods for observing the cell division[END_REF]. Also new structures appear such as arms, and concentric rings (A. (iiib); scale bar 3µm) (green: Pxl1 and red: Rlc1). B. Mammalian cell (HeLa) cytokinetic ring (tagged-myosin heavy chain MHC) visualized on a flat surface (B. (i); scale bar 10µm) with 'xy' plane showing two point accumulations. Reconstitution of 'z' planes projections showing blurred tilted ring (B. (iia); scale bar 10µm). Reconstitution of 'yz' planes, (B. (iib); green: myosin; scale bar 5µm) showing a deformed blurred ring with maximal projection and with sectional cut 'en face'. To visualize the full ring perimeter, cells are oriented vertically with respect to the plane of visualization inside cavities (B. (iii); scale bar 5µm), revealing myosin clusters around the ring perimeter. C. Lumen formation in MDCK doublets, cultured on glass coverslip (with Matrigel) after 24h visualized by confocal microscopy in single plane, showing two points accumulation (light blue:ZO1; lumen). Reconstitution of 'yz' planes showing a blurred and deformed lumen contour with sectional cut 'en face' for MDCK doublets. (C.; (ii)). Gain in information as well as in resolution is achieved after seeding cells inside through-hole NOA-74 cavities showing resolved and undistorted lumen contour (light blue: ZO1, magenta: E-cadherin, green: podocalyxin) Scale bars 3µm.

Figure 3 :

 3 Figure 3: Gain in spatial and temporal resolutions in cavities; an example myosin clusters of cytokinetic ring. A. PDMS cavities with polymer (PDMS) layer at the bottom (A; (i) Scale bar 10 µm) is used. Synchronized cells are seeded inside cavities and visualized with an inverted set-up (A. (ii)) showing a blurred cytokinetic ring (tagged-MHC) (A. (iia); scale bar 5µm); inset (red dashed lines) zoomed view showing a blurred portion of the ring (A. (iib); scale bar 3µm; Wollrab, 2014). To better resolve the cytokinetic ring inside a cavity with a PDMS layer at the bottom, imaging is done with an upright microscope (A. (iii)) avoiding the bottom polymer layer, revealing evenly spaced myosin clusters around the cytokinetic ring perimeter (A. (iiia); scale bar 5µm); inset (iib; red dashed lines; scale bar 1µm) zoomed view revealing the distance between clusters to be ~0.8µm. For further gain in resolution and optimized set-up usage, NOA-74 cavities without polymer layer at the bottom (B. (i) Scale bar 10 µm) are prepared. Synchronized cells are seeded inside cavities and visualized with an inverted setup (B. (ii)) using STED microscopy. Visualization of cytokinetic ring revealing individual myosin clusters (B. (iia); scale bar 5µm); inset (iib; red dashed lines; scale bar 1µm) zoomed view revealing myosin cluster inter-distance to be ~0.2 µm. C. Temporal resolution in cytokinetic ring (cluster) dynamics achieved by following the ring closure over time at high frame rate (10s), overlay of 25 frames, (C. (i)) shows clusters moving radially (still clusters) throughout the ring closure (C. (ii)) scale bar 5µm)(Wollrab, 2016)

11.

  After curing, pillars are carefully peeled off the surface of coverslip. Through-hole NOA-74 cavities are revealed, directly attached to the glass coverslip without any polymer layer at the bottom (see Figure1C.iv and Figure3B.i).

Figure

  Figure 3B.iia-b) revealing myosin clusters spaced at ~0.2m intervals. Another example of oriented structures such as 'lumen' can also be visualized inside similar (PDMS) cavities without polymer layer at the bottom. (see Figure1C (iv) and Figure 4C; STED)

Figure 4: Visualisation 'en face' of MDCK lumen at the two cell stages in cavities with STED microscopy.

	A. Sketch of the lumen initiation over
	time. Zoom shows the cell-cell
	adhesion interface and some of the
	key molecular components involved
	in organising the lumen. (B; (i))
	Point spread function of confocal
	and 2D-STED, showing the 10 fold
	increase in XY resolution for STED
	but axial resolution is low. (B; (i))
	Illustration of MDCK doublets inside
	through-hole NOA-74 cavities with
	lumen (inset; red). (C) Confocal and
	STED images of MDCK doublets
	cultured in through-hole NOA-74
	cavities for 24h. Cells, which divided
	in the axial direction, allow to
	resolve the cell-cell adhesion
	interface with super-resolution at
	different stages of lumen expansion.
	Note that E-cadherin adhesion
	clusters are clearly resolved in
	lateral domain in the STED image.
	Additionally, the apical membrane
	protein podocalyxin appears to form
	protrusions in the STED image,
	which is consistent with micro-villi
	formation on the apical membrane.

Acknowledgements:

We thank the Riveline Lab and the Imaging Platform of IGBMC for discussions. This study was supported by the Human Frontier Science Program number RGP0050/2018. A.B. and SLV were also funded by the University of Strasbourg. This work with the reference ANR-10-LABX-0030-INRT has been supported by a French state fund through the Agence Nationale de la Recherche under the frame programme Investissements d'Avenir labelled ANR-10-IDEX-0002-02. C-H.W and R.M. were funded by the Max Planck Society (ELBE fellowship) and by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) -Project Number 112927078 -TRR 83.