Nanoscale Forces during Confined Cell Migration - ACS Publications

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Nanoscale forces during confined cell migration Emma Desvignes, Anais Bouissou, Adrian Laborde, Thomas Mangeat, Amsha Proag, Christophe Vieu, Christophe Thibault, Isabelle Maridonneau-Parini, and Renaud Poincloux Nano Lett., Just Accepted Manuscript • DOI: 10.1021/acs.nanolett.8b02611 • Publication Date (Web): 20 Sep 2018 Downloaded from http://pubs.acs.org on September 21, 2018

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Nanoscale forces during confined cell migration

Emma Desvignes1, Anaïs Bouissou2, Adrian Laborde1, Thomas Mangeat3, Amsha Proag3, Christophe

Vieu1,

Christophe

Thibault1,*,

Isabelle

Maridonneau-Parini2,#,

Renaud

Poincloux2,,#*

1

LAAS-CNRS, Université de Toulouse, CNRS, INSA, Toulouse, France;

2

Institut de

Pharmacologie et de Biologie Structurale, Université de Toulouse, CNRS, UPS, France; 3

LBCMCP, Centre de Biologie Intégrative, Université de Toulouse, CNRS, UPS, France.

#

These authors share senior authorship.

*Correspondence to: [email protected] and [email protected]

ACS Paragon Plus Environment

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Abstract: In vivo, immune cells migrate through a wide variety of tissues, including confined and constricting environments. Deciphering how cells apply forces when infiltrating narrow areas is a critical issue that requires innovative experimental procedures. To reveal the distribution and dynamics of the forces of cells migrating in confined environments, we designed a device combining microchannels of controlled dimensions with integrated deformable micropillars serving as sensors of nanoscale subcellular forces. First, a specific process composed of two steps of photolithography and dry etching was tuned to obtain micrometric pillars of controlled stiffness and dimensions inside microchannels. Second, an image analysis workflow was developed to automatically evaluate the amplitude and direction of the forces applied on the micropillars by migrating cells. Using this workflow, we show that this micro-device is a sensor of forces with a limit of detection down to 64 pN. Third, by recording pillar movements during the migration of macrophages inside the confining microchannels, we reveal that macrophages bended the pillars with typical forces of 0.3 nN and applied higher forces at the cell edges than around their nuclei. When the degree of confinement was increased, we found that forces were redirected from inwards to outwards. By providing a micro-device that allows the analysis of force direction and force magnitude developed by confined cells, our work paves the way for investigating the mechanical behavior of cells migrating though 3D constricted environments.

Key Words: confined migration, cellular forces, pillars, microfluidics, microchannels, macrophages


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Immune cell migration is essential for the delivery of protective immune responses to tissues, but also contributes to a wide variety of diseases 1-3. Macrophages are immune cells that migrate in all body tissues to fulfil microbicide and tissue repair functions, whatever the composition, structure or stiffness of the tissues, which range from highly dense (bone, cancer, fibrosis) to sparse environments (mesentery, interstitial tissue). When encountering dense environments, macrophages dig narrow tunnels thanks to proteolytic digestion of the extracellular matrix. Macrophages are constrained inside these tunnels, as revealed by their elongated and protrusive morphologies and their squeezed nuclei

4, 5

. In order to advance

towards therapeutic control of macrophage migration, it is now crucial to understand the mechanisms used by macrophages to infiltrate tissues. However, most studies of cell migration have only tackled the issue of two-dimensional (2D) migration, which significantly differs from migration within a tridimensional (3D) constricting environment 6-8. One of the key differences between 2D and 3D migration is the mechanical role of the nucleus. Indeed, due to its size and stiffness, the nucleus is the main limiting organelle slowing down cell migration through constrictions

9, 10

, whereas it is not a barrier for 2D migration.

Noteworthily, nuclear envelope deformability depends on the density and proportion of the different isoforms of lamins which form the intermediate filaments of nuclei

11

. To trigger

nucleus deformation, the Arp2/3 complex mediates perinuclear actin nucleation that allows disruption of the nuclear lamina and passage through constrictions

12

. However, if the

constriction is too small (from 2 to 7 µm², depending on the cell type), the nucleus cannot be deformed enough to go through, and the migration process is blocked 13. For cell progression in constrained areas, the forces described to deform nuclei are in the nano-Newton range and they cause physical reorganization and alteration of the chromatin and even lead to breakage of the nuclear envelope and cell death 14-18. To further understand the mechanics


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of cell migration in 3D environments, it is important to develop new tools to evaluate the forces applied during confined migration, and in particular to decipher the mechanisms involved in nuclear deformability when cells pass through constrictions.

Since the seminal work of Harris et al., who introduced the first method to reveal cellgenerated forces

19

, the development of traction force microscopy based on deformation

measurements on elastomeric substrates

20, 21

or on arrays of flexible micropillars

22, 23

has

greatly improved our understanding of 2D cell migration mechanisms. However, how cells apply forces when they migrate through confined environments remains insufficiently explored. Of note, a model has been proposed to explain how non-adherent cells are able to move in tridimensional environments. Malawista et al. first demonstrated that adhesiondeficient neutrophils could migrate when confined between two glass surfaces and proposed that these cells use a chimneying mode of migration 24. To be able to migrate by chimneying, cells that cannot adhere and generate traction forces probably apply outward pressure forces to their environment 25. However, this mechanism has still to be elucidated.

Our goal is to understand how macrophages apply forces in order to migrate through confining environments. To investigate the mechanics of confined cells, we developed an in vitro device, which combines microchannels of controlled dimensions with micropillars serving as sensors of nanoscale forces. This method allows the real-time evaluation of force amplitude and direction in the nuclear area and at the cell extremities, and reveals that macrophages redirect forces from inwards to outwards when the degree of confinement is increased.


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Fabrication of a micro-device to evaluate nanoscale forces during confined migration. To evaluate the forces generated by cells during migration in confined environments, we developed a microfluidic device composed of two arrays of parallel 6 µm- or 12 µm-wide and 11 µm-high microchannels made of polydimethylsiloxane (PDMS). Microchannels from each group were connected together to the same 1.8 mm-large inlet reservoirs at their ends. To estimate cell forces, microchannels were equipped with cylindrical pillars attached to the top of the microchannel (Figure 1A). From beam theory, the relation between the bending force F applied at the top of a cylindrical pillar and its deflection d is given by equation (1), where dp and hp are the diameter and height of the pillar, respectively, and E the Young’s modulus, i.e. about 2 MPa in the case of PDMS 26.

=

(1)

Considering that migrating cells generally apply forces in the nano-Newton range 23, 27, 28, we therefore designed pillars with a diameter and height of 2 µm and 9 µm, respectively. This way, pillars would have an estimated stiffness of 6.5 nN/µm, and a 1 nN force would induce a displacement of 154 nm, which is compatible with the measurement of pillar deflection by optical microscopy.

The fabrication process of the microdevice is based on casting PDMS over a silicon (Si) mold exhibiting negative structures of microchannels and pillars and sealed over a glass coverslip (Figure S1A). A first issue for the Si mold fabrication behind this process was the production of 9 µm high pillars in the center of 6 µm-wide and 11 µm-high microchannels, so that the


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pillars are free to move but high enough to minimize the space between the free extremity of the pillars and the surface of the coverslip, ensuring future contacts with migrating cells. To achieve this dimensional control, the etching of the pillar and the microchannel structures on the Si mold required two steps of photolithography and dry etching, using sequentially two different masks and specific etching times for pillars and microchannels (see experimental procedure section).

The second microfabrication challenge consisted in unmolding the pillars while keeping them upright. Indeed, unmolding the device in open air caused the random collapse of the pillars (Figure S1B, left). To solve this issue, the PDMS stamp was unmolded in ethanol, then placed during 1 h on a glass coverslip with the microchannels facing the glass surface until complete ethanol evaporation. Surprisingly, pillars initially straight when immersed in ethanol (Figure S1A, step 6), were bent on one side of the channel after air drying (Movie S1 and Figure S1B, middle image). The reservoirs connecting the microchannels were then punched, which eventually led to a rise of the pillars (Figure 1 D-G and Figure S1B, right image). Finally, the stamp was sealed on a new glass coverslip after plasma treatment of both surfaces.

Using this procedure, we managed to produce in a reproducible way pillars with a 1.8 ± 0.1 µm diameter and an 8.4 ± 0.8 µm height inserted in 11.2 ± 0.7 µm-high microchannels (Figure S1C). The distance of 2.8 µm between the pillars and the glass coverslip allows cells to migrate through microchannels with mandatory interaction with pillars.


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To be able to convert pillar displacements into cell forces, we experimentally evaluated the stiffness of the microfabricated pillars using Atomic Force Microscopy (AFM). Measurements were performed by scanning pillars inside a microchannel so that the cantilever was in contact with the top of the pillar during the scan (Figure 1B). By recording the vertical deflection of the cantilever, it is then possible to extract a force/displacement curve and extract a spring constant k=7.1 ± 1.9 nN/µm (mean +/- SD, 12 pillars measured) (Figure 1C). Of note, this value is close to the value of 5.9 nN/µm derived from classical beam theory (equation 1), with Young’s modulus of PDMS: E = 2.05 MPa 26, dp = 1.8 µm and hp= 8.4 µm as measured by Scanning Electron Microscopy (SEM).

Pillar deformation by migrating macrophages enables force measurements inside microchannels Human monocyte-derived macrophages were seeded in the reservoirs and after 48 h, we evaluated whether cells migrating inside microchannels bent the pillars along their trajectory (Figure 2A-C). SEM and deconvolution fluorescence microscopy of fixed cells revealed that macrophages were in contact with each side of the microchannels and interacted with several micropillars at the same time (Figure 2A,B, Figure S2 and Movie S2,3). We then performed time-lapse experiments to observe living macrophages inside the microchannels. Macrophages migrated inside microchannels with an average velocity of 12 ± 4 µm/h and 10 ± 7 µm/h for 12 and 6 µm-wide microchannels, respectively (Figure 2C, Movie S4,5), which is of the same order of magnitude as macrophages migrating through 3D collagen gels 5

. Zooming on the nucleus revealed that nuclei were squeezed by the pillars (Figure 2D). In

order to evaluate the degree of cellular confinement as a function of microchannel size, we measured the nuclear aspect ratio of macrophages migrating in 12 and 6 µm-wide


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microchannels. In 12 µm-wide microchannels, the nuclear aspect ratio was slightly higher than in cells plated on glass coverslips, indicating moderate constraints. In 6 µm-wide microchannels, nuclei were mostly elongated, with an aspect ratio increased two-fold compared to 12 µm-wide microchannels (Figure 2E,F and Figure S3).

We then assessed whether macrophages could move pillars while migrating inside microchannels through the development of a dedicated automated analysis procedure. This consisted, sequentially, in the registration of time-lapse movies, the segmentation of cells and nuclei and the tracking of pillars to measure displacements and angles relative to the microchannel axis (Figure S4). As an internal reference, we plotted the positions of the pillars not in contact with macrophages. The positions of the control pillars obtained by automated tracking exhibited fluctuations with a standard deviation of 9 nm. These fluctuations indicate the level of accuracy of our registration process. The force corresponding to a displacement of 9 nm through equation (1) is 64 pN (see Figure 2G). Measurements of the displacements of the pillars in contact with cells exhibited a larger distribution than that of control pillars, with a median of 47 nm, which corresponds to a force of 332 pN (Figure 2G).

Such force values are of the same order of magnitude as migration forces measured on 2D substrates 22, 27, 29. Of note, equation (1) assumes that cells only apply a punctual force at the tip of the pillars, but we cannot exclude that applied forces extend over a larger area. To evaluate how force distribution could affect force evaluation, we performed mechanical simulations and compared pillar deflections when forces are applied at the tip of the pillars or on larger surfaces. Figure S5A-B shows a 2.4-fold decrease in deflection values when forces are uniformly applied on the total length of a pillar, as compared to a tip-loaded force.


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For the rest of the manuscript, we use the simple analytical equation (1) to evaluate cellular forces, although it under-estimates cell forces . Altogether, these observations demonstrate that this micro-device is able to record the forces generated by cells migrating in an environment with a controlled degree of confinement.

Cell force landscape and dynamics Having validated the force measurement procedure inside confining microchannels, we questioned whether macrophages apply different forces at distinct cellular locations and if these forces vary with the degree of cell confinement.

We measured the displacement over time of pillars located either at cell protrusions or close to the nucleus, both in the weak confinement condition (12 µm-wide microchannels) and the strong confinement condition (6 µm-wide microchannels). To perform this analysis, we classified pillars in two categories, distinguishing the pillars located in the nuclear area, from those in the extra-nuclear area (Figure 3C). Time-lapse measurements showed that macrophages applied forces intermittently, with values fluctuating between noise level and 2 nN. Furthermore, pillar displacement in the nuclear area seemed generally lower than at cell edges for both 6 and 12 µm-wide microchannels (Figure 3A,B, compare pillars p2 and p5 to p1, p3, p4 and p6). We then plotted as a function of the micro-channel width, the maximal force recorded either in nuclear or extra-nuclear areas. We observed that the degree of confinement did not influence maximal cell forces. We also noticed that the maximal forces exerted in the extra-nuclear areas turned out to be significantly higher than the forces in the


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nuclear areas (Figure 3D), suggesting that force generation around the nucleus or at the cell edges involves different mechanisms.

Macrophage force orientation depends on confinement. Next, we questioned whether cell confinement influences the direction of the force exerted on pillars. Tracking pillar displacements revealed that pillars located at the edges of cells migrating in 12 µm-wide microchannels displayed inward movements whereas pillars in 6 µm-wide microchannels showed outward movements (Figure 4A). We performed automated measurements of the orientation of pillar displacements for at least 33 pillars per condition. The frequency of the orientation of displacements of extra-nuclear pillars confirmed predominant inward pillar movements in 12 µm-wide microchannels, and outward movements in 6 µm-wide microchannels (Figure 4B). The analysis of force orientation in the nuclear area showed that in 6 µm-wide microchannels, predominant outward pillar movements relatively to the nucleus were observed (Figure 4B). On the contrary, no specific orientation of the pillar movements was noted in 12 µm-wide microchannels (Figure 4B). This can be explained by the fact that, in 12 µm-wide microchannels, nuclei located around the pillars (see Figure 2A), and not on one side of the pillars as in 6 µm-wide microchannels (see Figure 2B).

In conclusion, we present a microdevice that allows the automated evaluation at the subcellular level of the amplitude and orientation of the nanoscale tangential and orthogonal forces generated by living macrophages moving in a confined environment. Our results evidence that the degree of cell confinement does not affect the amplitude of forces but their direction, and suggest that cells sense and respond to confinement by redirecting


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the forces they applied on their environment, from inwards to outwards (Figure 4C). This redirection of cellular forces is reminiscent of the chimneying migration mechanism, as hypothesized by Malawista et al.

25

, except that in our system, macrophages adhere to a

fibronectin-coated environment whereas chimneying was defined as a non-adhesive mechanism of migration. It would be interesting to verify whether macrophages redirect forces the same way when they migrate in non-adhesive conditions. This capability can bring new insights to complement our understanding of how cells apply forces during confined migration, which has been the subject of only very few studies. By confining cells on arrays of pillars, Raman et al. could evaluate the amplitude of traction, but not outward forces inside microchannels and proposed that cell forces decrease with the degree of confinement 29

. Based on the results we have described in this paper, it is tempting to speculate that the

decrease of these traction forces was the result of a change in the vector direction of the force rather than a decrease of its modulus. This would be consistent with our results showing that confinement reduces inward traction forces in favor of outward forces. Our measurements are also in agreement with the finding that leukemia cells, when confined between two non-adhesive polyacrylamide gels, apply forces against the surfaces of the confining gels 30. More recently, Molino et al. followed cells migrating inside microchannels and going around oil droplets that they deformed with mechanical stress evaluated at 500 ± 100 Pa

31

. This stress value does not conflict with the magnitude of the forces

measured in this paper if we consider that cells apply forces to micrometric areas on the pillars, giving rise to stresses around 1 kPa (1 nN/µm2).

This device also made it possible to observe how confinement influenced nucleus deformation and interaction with the environment. It will provide a means to decipher the


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molecular mechanisms involved in the mechanical response of the nucleus and in the generation of inward and outward forces during confined migration.

EXPERIMENTAL PROCEDURES Si mold fabrication Microchannels are divided in 2 groups of different sizes (54 micro-channels of 6 µm-width and 49 micro-channels of 12 µm-width). The master mold is fabricated by processing a 4 inch-silicon wafer. 22 different designs of the microdevices are accommodated on this wafer, each microdevice being composed of 103 micro-channels (3 mm-long) and 4 reservoirs, and each micro-channel being equipped with micro pillars (1 pillar every 15 µm or 12 µm for the 6 and 12 µm-wide micro-channels, respectively). Two sequences of UV photolithography and pattern transfer by Reactive Ion Etching were sequentially conducted in order to define the topographical microstructures of the mold corresponding to the pillars and the micro-channels. The cavities of the mold corresponding to the future PDMS pillars were created during the first stage and the microchannels during the second stage. For each photolithography step, the substrate was cleaned and activated using O2 plasma treatment (5 min, 80 W). For the first sequence, the wafer was then baked at 100°C for 15 min before hexamethyldisilazane (Sigma Aldrich) surface coating to promote the adhesion of the resist to the wafer. A positive photoresist film (ECI 3012, CIPEC Spécialités) was spin-coated on the wafer during 30 s (3600 rpm speed, 5000 rpm/s acceleration) to obtain a thickness of 1.1 µm. The resist was exposed with UV light (20 mW/cm2 at 405 nm) for 11 s in vacuum contact with an optical aligner (Suss Microtec, MA 150), followed by post-exposure bake (60 s at 110°C). The wafer was then developed for 25 s in MF CD 26 (CIPEC Spécialités) and the


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silicon wafer was dry etched during 243 s using the resist as an etch mask inside an inductively coupled plasma Alcatel - AMS 4200 cluster tool. Deep reactive-ion etching was based on a sequence of Si etching steps by SF6 (2 s) and passivation by C4F8 (3.5 s). For the second sequence, the same process was used for transferring the patterns of the micro-channels by dry-etching during 251 s. The fine adjustment of the two etching times allowed a control of the appropriate height of the pillars relatively to the height of the micro-channels. Finally, the silicon mold was treated with perfluorodecyltrichlorosilane (Sigma Aldrich) in vapor phase to allow unmolding of the casted PDMS replica.

Production of polydimethylsiloxane micro-fluidic devices The prepolymer (Sylgard 184) and the cross-linking agent (Sylgard 184) were mixed at a 10:1, casted on the full wafer of Si and degassed under primary vacuum for a few minutes. The PDMS was solidified at 100°C for 60 min and unmolded manually in ethanol (Figure S1). Individual micro-devices were then isolated by cutting the PDMS replica with a razor blade. Microdevices were then placed on a glass coverslip and left to dry at room temperature. Of note, the ethanol/air meniscus pushed and collapsed the pillars when progressing into the channels. The punching of the reservoirs allowed us to create inlet reservoirs and resulted in the rise of the pillars (Figure S1). An O2 plasma treatment (90 s, 50 W) was used to activate both a glass coverslip (24 mm-diameter, Marienfeld) and the surface of the PDMS microdevices. After bonding the PDMS chip to the glass coverslip, devices were sterilized for 1 h at 100°C. The devices were immersed in Phosphate Buffered Saline (PBS) (Gibco), then a PBS solution containing 0.5 mg/mL of fibronectin (F-2006, Sigma), an extracellular matrix protein present in all body tissues that plays an important role in macrophage adhesion and


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migration, was injected for 1 h in the micro-channels. Finally, the micro-channels were washed three times with PBS.

Evaluation of pillar stiffness by Atomic Force Microscopy AFM measurements were performed using silicon nitride cantilevers (MLCT, BRUKER) with a nominal spring constant of 20 nN/µm mounted on a NanoWizard III AFM (JPK Instruments) coupled to an inverted optical microscope (Axiovert 200, Carl Zeiss). The cantilever sensitivity and spring constant were calibrated before each experiment with the JPK Instruments software using the thermal noise method.

Finite Element Simulations Simulations of pillar deformation by cell forces were performed using the Structural Mechanics module of COMSOL Multiphysics 4.3 software (COMSOL France). Briefly, a cylindrical pillar of Young’s modulus 2.05 MPa, Poisson’s ratio 0.45, density 965 kg/m³, diameter 1.86 μm and height 8.42 µm was clamped at its base. Cell forces were modelled as 1 nN forces orthogonal to the pillar exerted on a hemi-cylindrical surface covering the half lateral surface of the pillar, placed at the tip of the pillar and of height varying from 1 to 8.42 µm (see Figure S5). Meshing used COMSOL default tetrahedral configuration with sizes determined so that the smallest distance spanned at least two tetrahedrons.

Cell culture and manipulation


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Human monocytes were isolated from the blood of healthy donors and differentiated into macrophages as previously described 5. After 7 days of differentiation human macrophages were stained during 30 min with Cell Tracker Green CMFDA dye (5 ng/µL, C7025, ThermoFisher) and Hoechst (20 µg/mL, 33342, ThermoFisher) and trypsinized (Invitrogen). 104 cells resuspended in 10 µL of RPMI medium containing fetal calf serum (0.5%) were injected in each reservoir of the micro device and incubated at 37°C in humidified 5% CO2 for 24 h before image acquisition.

Time-lapse imaging Time-lapse experiments were performed in differential interference contrast (DIC) microscopy and confocal fluorescence microscopy using the 60x oil objective (1.35 NA, UPlanSAPO) of an inverted spinning disk microscope (Olympus, IX83) equipped with an EMCCD camera (iXon Ultra, Andor), thermoregulation and a humidified CO2 (5%) controller, and driven by Andor IQ3 software. Images were acquired every 60 s for at least 1 h. For each time point, DIC images of the free tip of pillars as well as Hoechst and Cell Tracker fluorescent images were acquired. At the end of the experiment, cells were lysed during 30 min with 0.5 % SDS (Sigma Aldrich) and an image of unstrained pillars was acquired (Figure S4). We verified that nucleus staining with Hoechst did not have any toxic effect during the time of recording, nor affected cell forces.

Scanning electron microscopy


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Cells migrating inside microchannels were fixed with 4% glutaraldehyde (Sigma Aldrich) for 20 minutes, followed by baths of 25, 50, 75 and 100% ethanol diluted in water. Samples were let to dry overnight at room temperature and PDMS devices were unmolded from the glass coverslip. Samples were sputtered with a 50 nm-thick gold layer (Precision Etching and Coating Systems, Gatan 682) and observed by SEM (Hitachi S-4800). Cells were pseudocolored with Photoshop (Adobe).

Immunofluorescence Cells were fixed inside the device with 3.7% PFA and 15 mM sucrose (Fisher Scientific), permeabilized with 2% Triton X-100 and saturated with BSA (3% in PBS) for 2 h at room temperature. Cells were incubated overnight with Texas-Red phalloidin (Thermo Fisher Scientific) and DAPI (Sigma Aldrich) in PBS containing 3% BSA. 70 images were acquired every 250 nm with an 60x oil objective (NA 1.4, Nikon) mounted on a Nikon Eclipse Ti-E and a Hamamatsu Orca flash 4.0 LT sCMOS. Images were de-convolved with Huygens Professional version 16.10 (Scientific Volume Imaging, The Netherlands), using the CMLE algorithm, with SNR: 20, 10 iterations. For nuclear aspect ratio measurements, nuclei were segmented with ImageJ, and aspect ratio was measured as the ratio of major-to-minor axis length.

Automated image processing and analysis DIC images were first registered with the Template matching ImageJ plugin, using as a reference an area where control pillars are not in contact with cells. Hoechst and Cell Tracker time series were aligned based on this first registration. Pillars were then tracked with the


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Trackmate ImageJ plugin

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, using the LoG detector and an estimated blob diameter of 10

pixels. Using a homemade image processing software written in R (Figure S4), objects marked with Hoechst and Cell Tracker were segmented to determine the positions of nuclei and cells along time. Pillar positions were classified into five groups: (i) reference pillars not in contact with cells, (ii-iii) pillars in the nuclear area, among which the Y coordinate was below (ii) or above (iii) the nucleus center of mass, (iv-v) pillars contacting the extra-nuclear area of the cell that were positioned to the left (iv) or the right (v) of the nucleus. A second registration was then applied to the pillars by subtracting the average position of the reference pillars (Figure S4B). Pillar populations (ii) and (iii) were pooled after applying a 180° rotation to the (iii) pillars and pillars (iv) and (v) were pooled after applying a 180° rotation to the (iv) pillars (Figure S4C, D). At the end of the experiment the cells were lysed with detergent (SDS 0.5 %) and imaged after 30 min to obtain the original position of the pillars without the cell. The displacement of the pillars is measured as follows. The origin position of each pillar was defined as the mean of its positions at all the times when it is not in contact with a cell, including the image acquired after cell lysis. The relative pillar displacements (d) were given by the following equation (2), where X, Y are the coordinates of the pillar at a given time and X0, Y0 its origin positions, = ( − ) + ( − ) (2) The direction of the pillar movement was obtained by calculating the angle between the axis of the microchannel (x axis) and the in-plane vector of coordinates (X-X0, Y-Y0) (Figure S4E).

Statistical analysis


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All box-and-whisker plots show the median, lower and upper quartiles (box). The variance over different macrophages was similar between each condition. Statistical significance was assessed using the Wilcoxon-Mann-Whitney test. In all cases * corresponds to p < 0.05, ** to p < 0.01 and *** to p < 0.001.

AUTHOR CONTRIBUTIONS C.V., C.T., I.M.P. and R.P. designed the study. E.D., A.B., C.V., C.T., I.M.P. and R.P. designed the experiments. E.D. performed most experiments. E. D. and A. L. fabricated the wafer mold. T.M. deconvolved fluorescence microscopy images. E.D., A.P., C.V., C.T., I.M.P. and R.P. interpreted the results and wrote the manuscript with input from the other authors.

SUPPLEMENTAL INFORMATION Figures detailing the fabrication process and the image analysis pipeline, scanning electron micrographs of contacts between macrophages and pillars, figures showing the morphology of macrophage nuclei inside microchannels, as well as the influence of force distribution on pillar displacement. Movies showing pillar bending by an air bubble during micro-device drying, macrophage morphology and migration inside micro-channels.

ACKNOWLEDGEMENTS The authors are grateful to Karine Pingris and Myriam Ben Neji for macrophage preparation and to Edouard Barker for critical reading of the manuscript. The authors also acknowledge the clean room facility of the LAAS-CNRS and the IPBS-TRI imaging facility. This work has been supported in part by l’Agence Nationale de la Recherche (ANR14-CE11-0020-02), la


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Fondation pour la Recherche Médicale (FRM DEQ2016 0334894), INSERM Plan Cancer, Fondation Toulouse Cancer and Human Frontier Science Program (RGP0035/2016). This work was partly supported by LAAS-CNRS micro and nanotechnology platform, member of the French RENATECH network.

COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests.

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14. Aureille, J.; Belaadi, N.; Guilluy, C. Mechanotransduction Via the Nuclear Envelope: A Distant Reflection of the Cell Surface. Curr Opin Cell Biol 2017, 44, 59-67. 15. Petrie, R. J.; Koo, H.; Yamada, K. M. Generation of Compartmentalized Pressure by a Nuclear Piston Governs Cell Motility in a 3d Matrix. Science 2014, 345, 1062-1065. 16. Guilluy, C.; Osborne, L. D.; Van Landeghem, L.; Sharek, L.; Superfine, R.; Garcia-Mata, R.; Burridge, K. Isolated Nuclei Adapt to Force and Reveal a Mechanotransduction Pathway in the Nucleus. Nat Cell Biol 2014, 16, 376-381. 17. Denais, C. M.; Gilbert, R. M.; Isermann, P.; McGregor, A. L.; te Lindert, M.; Weigelin, B.; Davidson, P. M.; Friedl, P.; Wolf, K.; Lammerding, J. Nuclear Envelope Rupture and Repair During Cancer Cell Migration. Science 2016, 352, 353-358. 18. Raab, M.; Gentili, M.; de Belly, H.; Thiam, H. R.; Vargas, P.; Jimenez, A. J.; Lautenschlaeger, F.; Voituriez, R.; Lennon-Dumenil, A. M.; Manel, N., et al. Escrt Iii Repairs Nuclear Envelope Ruptures During Cell Migration to Limit DNA Damage and Cell Death. Science 2016, 352, 359-362. 19. Harris, A. K.; Wild, P.; Stopak, D. Silicone Rubber Substrata: A New Wrinkle in the Study of Cell Locomotion. Science 1980, 208, 177-179. 20. Dembo, M.; Wang, Y. L. Stresses at the Cell-to-Substrate Interface During Locomotion of Fibroblasts. Biophys J 1999, 76, 2307-2316. 21. Plotnikov, S. V.; Pasapera, A.; Sabass, B.; Waterman, C. M. Force Fluctuations within Focal Adhesions Mediate Ecm Rigidity Sensing to Guide Directed Cell Migration. Mol Biol Cell 2012, 23. 22. Balaban, N. Q.; Schwarz, U. S.; Riveline, D.; Goichberg, P.; Tzur, G.; Sabanay, I.; Mahalu, D.; Safran, S.; Bershadsky, A.; Addadi, L., et al. Force and Focal Adhesion Assembly: A Close Relationship Studied Using Elastic Micropatterned Substrates. Nat. Cell Biol. 2001, 3, 466-472. 23. Tan, J. L.; Tien, J.; Pirone, D. M.; Gray, D. S.; Bhadriraju, K.; Chen, C. S. Cells Lying on a Bed of Microneedles: An Approach to Isolate Mechanical Force. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 14841489. 24. Malawista, S. E.; de Boisfleury Chevance, A.; Boxer, L. A. Random Locomotion and Chemotaxis of Human Blood Polymorphonuclear Leukocytes from a Patient with Leukocyte Adhesion Deficiency-1: Normal Displacement in Close Quarters Via Chimneying. Cell Motil Cytoskeleton 2000, 46, 183-189. 25. Heuze, M. L.; Vargas, P.; Chabaud, M.; Le Berre, M.; Liu, Y. J.; Collin, O.; Solanes, P.; Voituriez, R.; Piel, M.; Lennon-Dumenil, A. M. Migration of Dendritic Cells: Physical Principles, Molecular Mechanisms, and Functional Implications. Immunol Rev 2013, 256, 240-254. 26. Johnston, I. D.; McCluskey, D. K.; Tan, C. K. L.; Tracey, M. C. Mechanical Characterization of Bulk Sylgard 184 for Microfluidics and Microengineering. Journal of Micromechanics and Microengineering 2014, 24. 27. Ricart, B. G.; Yang, M. T.; Hunter, C. A.; Chen, C. S.; Hammer, D. A. Measuring Traction Forces of Motile Dendritic Cells on Micropost Arrays. Biophys J 2011, 101, 2620-2628. 28. Marelli, M.; Gadhari, N.; Boero, G.; Chiquet, M.; Brugger, J. Cell Force Measurements in 3d Microfabricated Environments Based on Compliant Cantilevers. Lab Chip 2014, 14, 286-293. 29. Raman, P. S.; Paul, C. D.; Stroka, K. M.; Konstantopoulos, K. Probing Cell Traction Forces in Confined Microenvironments. Lab Chip 2013, 13, 4599-4607. 30. Yip, A. K.; Chiam, K. H.; Matsudaira, P. Traction Stress Analysis and Modeling Reveal That Amoeboid Migration in Confined Spaces Is Accompanied by Expansive Forces and Requires the Structural Integrity of the Membrane-Cortex Interactions. Integr Biol (Camb) 2015, 7, 1196-1211. 31. Molino, D.; Quignard, S.; Gruget, C.; Pincet, F.; Chen, Y.; Piel, M.; Fattaccioli, J. On-Chip Quantitative Measurement of Mechanical Stresses During Cell Migration with Emulsion Droplets. Sci Rep 2016, 6, 29113. 32. Tinevez, J. Y.; Perry, N.; Schindelin, J.; Hoopes, G. M.; Reynolds, G. D.; Laplantine, E.; Bednarek, S. Y.; Shorte, S. L.; Eliceiri, K. W. Trackmate: An Open and Extensible Platform for SingleParticle Tracking. Methods 2017, 115, 80-90.


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FIGURE LEGENDS Figure 1. PDMS micro-channels equipped with deformable pillars. (A) PDMS devices are composed of two groups of 6 and 12 µm-wide microchannels connected on both ends to reservoirs containing cells (arrows). w, hc, dp and hp correspond to the micro-channel width and height, and pillar diameter and height, respectively. (B) Pillar stiffness was measured by scanning pillars with an AFM so that the tip contacts the top of the pillar. (C) Typical plot of the force induced by pillar flexion, obtained by recording the vertical deflection of the cantilever just after contact with a pillar. (D-G) Scanning electron micrographs of a 12 µm- (D,F) and a 6 µm- (E,G) wide microchannels equipped with cylindrical pillars of diameter 1.8 µm. Scale bars, 5 µm.

Figure 2. Pillar displacement during macrophage confined migration. (A,B) De-convolved fluorescence microscopy images of macrophages fixed inside a 12 µm(A) and 6 µm- (B) wide micro-channels, immunostained for F-actin (red), vinculin (green) and the nucleus (blue), with explicative schematics. (C) Stills of a living macrophage (nucleus in blue stained with Hoechst, cytoplasm in green stained with Cell Tracker) migrating in a 6 µm-wide micro-channel equipped with pillars. (D) Enlargement of the nucleus shown in (C). Note that the nucleus is deformed close to the pillar (white dotted line). (E) Nuclear aspect ratio is defined as the ratio between the nucleus major and minor axes. (F) Aspect ratio of the nuclei of cells plated on a 2D glass coverslip and inside 6 and 12 µmwide micro-channels. At least 27 nuclei were analyzed per condition.


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(G) Box-plots of the recorded and computed displacements of control pillars and pillars in contact, with cells with the corresponding forces. At least 34 pillars were analyzed per condition. Scale bars, 10 µm.

Figure 3. Force dynamics and amplitude. (A,B) Upper panels. Images of macrophages migrating inside a 12 (A) and 6 (B) µm-wide micro-channels, stained with Cell Tracker (green) and Hoechst (blue). Lower panels. Displacements of the pillars over time and corresponding force amplitudes. Each graph corresponds to the pillar indicated in the upper frames. Note that pillars p2 and p5, located in the nuclear area, exhibit smaller displacements than pillars located in extra nuclear areas. (C) Scheme of a macrophage inside a microchannel. Pillars are classified in two groups: the ones in the nuclear area (blue) and those in the extra-nuclear area (black). (D) Box-plots of the maximal pillar displacements and corresponding forces observed in extra-nuclear (EN) and nuclear (N) areas in 6 and 12 µm-wide micro-channels. At least 6 cells from 3 donors were analyzed per condition. Scale bars, 10 µm.

Figure 4. Direction of the force exerted during macrophage migration in micro-channels. (A) Polar coordinate plots showing the direction of the displacements of each pillar over time and the direction of the corresponding forces (same pillars as in Figure 3A-B). (B) Polar coordinate plots depicting the frequency of the direction along which pillar are displaced in 6 and 12 µm-wide micro-channels and in nuclear and extra-nuclear areas. When necessary, rotations were applied to position extra-nuclear pillars at the right of the nucleus and nuclear pillars below nuclei (see Figure S2). At least 7 pillars were analyzed per condition.


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(C) Typical orientations of the forces applied by macrophages migrating in semi-confined (left) and confined (right) environments. Under weak confinement, macrophages apply inward forces at the cell protrusions, whereas they apply outward forces under strong confinement.


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Graphical abstract. 77x39mm (300 x 300 DPI)


Nanoscale Forces during Confined Cell Migration - ACS Publications

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