The authors are grateful to Anna Labernadie, Guillaume Charri&#232;re and Patrick Delobelle for their initial contribution to this work and to Matthieu Sanchez and Fran&#231;oise Viala for their help with video filming and editing. This work has been supported by l'Agence Nationale de la Recherche (ANR14-CE11-0020-02), la Fondation pour la Recherche M&#233;dicale (FRM DEQ2016 0334894), INSERM Plan Cancer, Fondation Toulouse Cancer and Human Frontier Science Program (RGP0035/2016).

1. Preparation of Formvar-Coated Grids
<ol>
	<li>Clean electron microscopy grids with pure acetone and dry them on filter paper. Then clean a microscope slide with pure ethanol, wipe with a lens paper and remove dust with a blower.</li>
	<li>Place an ethanol-cleaned glass slide vertically in the funnel of a film casting device containing a solution of Formvar in the lower part. Cover the top of the funnel.</li>
	<li>Pump 100 mL of Formvar solution (0.5% in ethylene dichloride) with the atomizer bulb until the level reaches two thirds of the slide.</li>
	<li>Keep the slide in the solution for 1 min.</li>
	<li>Open the valve of the device to drain the Formvar solution in a steady stream. A flow rate of 10 to 15 mL/s will yield a Formvar film of 30-80 nm. The thickness of the film is determined by the concentration of the Formvar solution and by the drainage rate. 
	NOTE: The drainage rate can be controlled by venting the pressure via the air-out valve by slowly opening the valve. The faster the drainage, the thicker the film. In practice, because it is difficult to predict the film thickness from the Formvar flow rate, we prepare several batches of Formvar films and control their thickness by AFM (see Step 2).</li>
	<li>Remove the slide from the chamber and dry it delicately on filter paper to remove any liquid Formvar.</li>
	<li>Cut out a 20 mm x 50 mm rectangle from the Formvar film with a razor blade.</li>
	<li>Fill a beaker with clean distilled water and slowly plunge the slide vertically into the water to float off the film: the film should float on the water surface.</li>
	<li>Place dry acetone-cleaned grids on the film, shiny face up.</li>
	<li>Place a round glass coverslip (&#216; 12 mm) onto a few of the grids. This coverslip will serve to measure the film thickness (see Step 2).</li>
	<li>Cover a microscope slide with a white sticker resized to fit it. Dip the slide vertically at the border of the floating Formvar film until the whole film adheres to the slide. Remove excess water from the slide with filter paper and let it dry at room temperature overnight.</li>
</ol>
2. Measurement of Film Thickness
<ol>
	<li>Cut the Formvar around the coverslip with tweezers to detach it from the slide.</li>
	<li>Mark the location of the grid under the coverslip with a pen.</li>
	<li>Cut the Formvar around the marked grid to detach it from the coverslip. There will therefore be a hole in the remaining Formvar sheet, which will allow to measure the film thickness. Cover the coverslip with PBS and place it on a glass slide on the microscope.</li>
	<li>Mount a silicon nitride cantilever on the AFM glass block, making sure the golden stripe is not covered by the tip of the spring. 
	NOTE: The sensitivity and spring constant of the cantilever should have previously been calibrated in PBS.</li>
	<li>Mount the glass block onto the AFM module, and then place the module onto the microscope.</li>
	<li>Place the Formvar-coated coverslip on the observation chamber and cover it with 500 &#181;L of PBS.</li>
	<li>Scan the border of the hole in the Formvar sheet using AFM in contact mode and a 0.5 nN force set point.</li>
	<li>Use the glass coverslip surface as the reference for height measurements, and evaluate the Formvar thickness as the height of the cross-section (perform at least 10 measurements per Formvar batch).</li>
</ol>
3. Seeding Cells on Grids
<ol>
	<li>Under a sterile hood, put a strip (12 mm x 3 mm) of double-sided adhesive tape on a coverslip.</li>
	<li>Cut out a Formvar-coated grid with tweezers and put it upside-down on the adhesive strip (only the rim of the grid needs to be attached to the tape). The Formvar film needs to face the coverslip.</li>
	<li>Put this device in a culture well.</li>
	<li>Place a 10 &#181;L droplet of RPMI without FCS containing 104 macrophages differentiated from human monocytes16. 
	NOTE: Make sure not to touch the grid with the pipette tip to prevent damaging the Formvar coating.</li>
	<li>Incubate for 30 min (37 &#176;C, 5% CO2) to let cells adhere.</li>
	<li>Fill up the well with 2 mL of RPMI containing 10% FCS.</li>
	<li>Incubate the device for 2 h at 37 &#176;C and 5% CO2 to let cells adhere before AFM observation.</li>
</ol>
4. Topography Measurements of Podosome-Induced Deformations
<ol>
	<li>Stick two strips of doubled-sided adhesive tape on a glass bottom Petri dish. 
	NOTE: The distance between the two strips should be smaller than the grid diameter.</li>
	<li>Carefully detach the grid plated with macrophages from the adhesive tape.</li>
	<li>Turn the grid upside-down in order to have the cells facing the glass and stick the grid between the two adhesive strips.</li>
	<li>Fix the grid with two new strips of doubled-sided adhesive: the grid will thus be sandwiched between two adhesive strips, leaving the central area of the grid accessible to the AFM tip. 
	NOTE: Make sure that the grid is well fixed and not twisted; otherwise the AFM cantilever might not be able to approach the Formvar surface.</li>
	<li>Fill the Petri dish with 2 mL of pre-heated 37 &#176;C culture medium (RPMI with 10% FCS) supplemented with 10 mM HEPES (pH 7.4).</li>
	<li>Place the culture dish on a 37 &#176;C dish heater.</li>
	<li>Install the dish heater on the AFM stage.</li>
	<li>Let the system stabilize at 37 &#176;C.</li>
	<li>In contact mode with a 0.5 nN force, make a first global image to locate podosomes, and then scan the Formvar topography at 3 Hz with an approximately 20 nm-large pixel. 
	NOTE: Avoid scanning near the edges of the grid.</li>
</ol>

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No conflicts of interest declared.

Material properties
The choice of the material for the deformable membrane, in our case Formvar, needs to fulfill a few requirements. The material must be transparent to visible light and exhibit limited auto fluorescence to allow observations in bright field and fluorescence microscopy. The roughness of the thin film must be well below 10 nm to avoid any topographical effect on cell adhesion and to allow clear observation of the cell-induced protrusions by AFM imaging. Finall...

Animal cells interact physically with the matrix and the other cells that constitute their environment1. This is required for them to migrate, internalize bodies, acquire external information, or differentiate. In such processes, the cell must generate mechanical forces and, as numerous studies have shown over the recent years, the ability of a cell to generate forces and probe its environment influences its biological behavior, directing for instance proliferation or differentiation2,3. In turn, the measurement of cellular forces is a major aid to study the regulation of force generation and understand its implication in cell behavior and tissue fate4,5.
Recent years have witnessed the development of numerous techniques to measure the forces that a cell can exert on its environment6. The majority of these have been instrumental in revealing the traction forces that cells exert as they pull on mobile probes or a deformable substrate. However, the mechanical forces involved in protrusion into the extracellular environment suffer from a lack of measurement techniques and are to date not well characterized.
To overcome this limitation, we present a method to measure forces exerted orthogonally to the substrate. It consists in plating living cells on a thin elastic sheet that can deform in the orthogonal direction, making it possible to measure substrate deformation by the cells and deduce the forces involved. Substrate topography is measured with nanoscale resolution using atomic force microscopy and the evaluation of forces from deformation relies on the knowledge of the geometry of the protrusive cellular structures7,8,9.
Here, we describe the setup and its application to measure the forces generated by podosomes, protrusive adhesion structures formed by macrophages for their mesenchymal migration in three-dimensional environments10,11,12,13,14,15,16,17. We believe that this technique will advance the understanding of force generation and its regulation in the many cellular processes involving protrusion.

<table><tbody><tr><td>200 mesh nickel grids</td><td>Electron Microscopy Sciences</td><td>G200-Ni</td><td></td></tr><tr><td>Filter paper</td><td>Sigma-Aldrich</td><td>1001-055</td><td></td></tr><tr><td>Microscope slides</td><td>Fisher Scientific</td><td>10235612</td><td></td></tr><tr><td>White stickers 26 x 70 mm</td><td>Avery</td><td>DP033-100</td><td></td></tr><tr><td>Film casting device with valve in its outlet</td><td>Electron Microscopy Sciences</td><td>71305-01</td><td></td></tr><tr><td>Razorblades</td><td>Electron Microscopy Sciences</td><td>72000</td><td></td></tr><tr><td>Ethanol</td><td>VWR</td><td>1.08543.0250</td><td></td></tr><tr><td>Acetone</td><td>VWR</td><td>20066.321</td><td></td></tr><tr><td>Formvar 0.5% solution in ethylene dichloride</td><td>Electron Microscopy Sciences</td><td>15820</td><td></td></tr><tr><td>12 mm coverslips</td><td>VWR</td><td>631-0666</td><td></td></tr><tr><td>Inverted microscope</td><td>Carl Zeiss</td><td>Axiovert 200</td><td></td></tr><tr><td>Atomic Force Microscope</td><td>JPK Instruments</td><td>NanoWizard III</td><td></td></tr><tr><td>Temperature-controlled sample holder&amp;nbsp;</td><td>JPK Instruments</td><td>BioCell</td><td></td></tr><tr><td>Silicon nitride cantilever with a nominal spring constant of 0.01 N/m</td><td>Veeco Instruments</td><td>MLCT-AUHW</td><td></td></tr><tr><td>PBS</td><td>Gibco</td><td>14190-094</td><td></td></tr><tr><td>Double-sided adhesive tape</td><td>APLI AGIPA</td><td>118100</td><td></td></tr><tr><td>RPMI 1640</td><td>Gibco</td><td>31870-025</td><td></td></tr><tr><td>FCS</td><td>Sigma-Aldrich</td><td>F7524</td><td></td></tr><tr><td>HEPES&amp;nbsp;</td><td>Sigma-Aldrich</td><td>H0887</td><td></td></tr><tr><td>35 mm glass-bottom Petri dishes</td><td>WPI</td><td>FD35-100</td><td></td></tr></tbody></table>

protrusion force microscopy: a method to quantify forces developed by cell protrusions

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction forces have been well-characterized, but there is a lack of techniques allowing the measurement of the protrusion forces exerted by cells orthogonally to their substrate. We designed an experimental setup to measure the protrusion forces exerted by adherent cells on their substrate. Cells plated on a compliant Formvar sheet deform this substrate and the resulting topography is mapped by atomic force microscopy (AFM) at the nanometer scale. Force values are then extracted from an analysis of the deformation profile based on the geometry of the protrusive cellular structures. Hence, the forces exerted by the individual protruding units of a living cell can be measured over time. This technique will enable the study of force generation and its regulation in the many cellular processes involving protrusion. Here, we describe its application to measure the protrusive forces generated by podosomes formed by human macrophages.

The above protocol describes how to prepare the experimental setup to quantify protrusion forces applied by macrophage podosomes on a Formvar substrate. This is achieved using AFM and is illustrated in Figure 1.
When analyzing a topographical image of bulges beneath podosomes using the JPK data processing software, a third-degree polynomial fit should be subtracted from each scan line independently....

Watch this Scientific Journal Video about Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions at JoVE.com

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

Here, we detail the experimental techniques used to evaluate the protrusion forces that podosomes apply on a compliant film, from the preparation of the film to the automated analysis of topographical images.

In numerous biological contexts, animal cells need to interact physically with their environment by developing mechanical forces. Among these, traction ...

protrusion-force-microscopy-method-to-quantify-forces-developed-cell

Research

JoVE Journal

Biology

5.5K Views.  IPBS, Université de Toulouse, CNRS, UPS. Hier, detail we de experimentele technieken gebruikt om te evalueren van de uitsteeksel krachten die podosomes van toepassing op een compatibele film, vanaf de voorbereiding van de film naar de geautomatiseerde analyse van topografische beelden.

Video: Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

Mechanical properties of cells and extracellular matrix (ECM) play important roles in many biological processes including stem cell differentiation, tumor formation, and wound healing. Changes in stiffness of cells and ECM are often signs of changes in cell physiology or diseases in tissues. Hence, cell stiffness is an index to evaluate the status of cell cultures. Among the multitude of methods applied to measure the stiffness of cells and tissues, micro-indentation using an Atomic Force Microscope (AFM) provides a way to reliably measure the stiffness of living cells. This method has been widely applied to characterize the micro-scale stiffness for a variety of materials ranging from metal surfaces to soft biological tissues and cells. The basic principle of this method is to indent a cell with an AFM tip of selected geometry and measure the applied force from the bending of the AFM cantilever. Fitting the force-indentation curve to the Hertz model for the corresponding tip geometry can give quantitative measurements of material stiffness. This paper demonstrates the procedure to characterize the stiffness of living cells using AFM. Key steps including the process of AFM calibration, force-curve acquisition, and data analysis using a MATLAB routine are demonstrated. Limitations of this method are also discussed.

This paper demonstrates a protocol to characterize the mechanical properties of living cells by means of microindentation using an Atomic Force Microscope (AFM).

Mechanical  properties of cells and extracellular matrix (ECM) play important roles in many  biological processes including stem cell differentiation, ...

Bioengineering

A Direct Force Probe for Measuring Mechanical Integration Between the Nucleus and the Cytoskeleton

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous with the cytoskeleton, methods are needed to probe its mechanical behavior in adherent cells. Here, we discuss the direct force probe (DFP) as a tool to apply force directly to the nucleus in a living adherent cell. We attach a narrow micropipette to the nuclear surface with suction. The micropipette is translated away from the nucleus, which causes the nucleus to deform and translate. When the restoring force is equal to the suction force, the nucleus detaches and elastically relaxes. Because the suction pressure is precisely known, the force on the nuclear surface is known. This method has revealed that nano-scale forces are sufficient to deform and translate the nucleus in adherent cells, and identified cytoskeletal elements that enable the nucleus to resist forces. The DFP can be used to dissect the contributions of cellular and nuclear components to nuclear mechanical properties in living cells.

In this protocol, we describe a micropipette method to directly apply a controlled force to the nucleus in a living cell. This assay allows interrogation of nuclear mechanical properties in the living, adherent cell.

The mechanical properties of the nucleus determine its response to mechanical forces generated in cells. Because the nucleus is molecularly continuous ...

Biochemistry

Perturbing Endothelial Biomechanics via Connexin 43 Structural Disruption

Endothelial cells have been established to generate intercellular stresses and tractions, but the role gap junctions play in endothelial intercellular stress and traction generation is currently unknown. Therefore, we present here a mechanics-based protocol to probe the influence of gap junction connexin 43 (Cx43) has on endothelial biomechanics by exposing confluent endothelial monolayers to a known Cx43 inhibitor 2,5-dihydroxychalcone (chalcone) and measuring the impact this inhibitor has on tractions and intercellular stresses. We present representative results, which show a decrease in both tractions and intercellular stresses under a high chalcone dosage (2 &#181;g/mL) when compared to control. This protocol can be applied to not just Cx43, but also other gap junctions as well, assuming the appropriate inhibitor is used. We believe this protocol will be useful in the fields of cardiovascular and mechanobiology research.

Here, we present a mechanics-based protocol to disrupt the gap junction connexin 43 and measure the subsequent impact this has on endothelial biomechanics via observation of tractions and intercellular stresses.

Endothelial cells have been established to generate intercellular stresses and tractions, but the role gap junctions play in endothelial intercellular ...

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to understand morphogenetic processes is to directly measure cellular forces and mechanical properties in vivo during embryogenesis. Here, we present a setup of optical tweezers coupled to a light sheet microscope, which allows to directly apply forces on cell-cell contacts of the early Drosophila embryo, while imaging at a speed of several frames per second. This technique has the advantage that it does not require the injection of beads into the embryo, usually used as intermediate probes on which optical forces are exerted. We detail step by step the implementation of the setup, and propose tools to extract mechanical information from the experiments. By monitoring the displacements of cell-cell contacts in real time, one can perform tension measurements and investigate cell contacts&#39; rheology.

Probing Cell Mechanics with Bead-Free Optical Tweezers in the Drosophila Embryo

We present a setup of optical tweezers coupled to a light sheet microscope, and its implementation to probe cell mechanics without beads in the Drosophila embryo.

Morphogenesis requires coordination between genetic patterning and mechanical forces to robustly shape the cells and tissues. Hence, a challenge to ...

Developmental Biology

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called mechanotransduction. A crucial step in mechanotransduction is the transmission of forces between the external and internal environments. To transmit forces, there must be a sustained, unbroken physical linkage created by a series of protein-protein interactions. For a given protein-protein interaction, mechanical load can either have no effect on the interaction, lead to faster disassociation of the interaction, or even stabilize the interaction. Understanding how molecular load dictates protein turnover in living cells can provide valuable information about the mechanical state of a protein, in turn elucidating its role in mechanotransduction. Existing techniques for measuring force-sensitive protein dynamics either lack direct measurements of protein load or rely on the measurements performed outside of the cellular context. Here, we describe a protocol for the F&#246;rster resonance energy transfer-fluorescence recovery after photobleaching (FRET-FRAP) technique, which enables the measurement of force-sensitive protein dynamics within living cells. This technique is potentially applicable to any FRET-based tension sensor, facilitating the study of force-sensitive protein dynamics in variety of subcellular structures and in different cell types.

Here, we present a protocol for the simultaneous use of F&#246;rster resonance energy transfer-based tension sensors to measure protein load and fluorescence recovery after photobleaching to measure protein dynamics enabling the measurement of force-sensitive protein dynamics within living cells.

Cells sense and respond to physical cues in their environment by converting mechanical stimuli into biochemically-detectable signals in a process called ...

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

Fabrication and Implementation of a Reference-Free Traction Force Microscopy Platform

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their microenvironment. While many approaches exist for measuring cell-induced material strain, here we provide a methodology for monitoring strain with sub-micron resolution in a reference-free manner. Using a two-photon activated photolithographic patterning process, we demonstrate how to generate mechanically and bio-actively tunable synthetic substrates containing embedded arrays of fluorescent fiducial markers to easily measure three-dimensional (3D) material deformation profiles in response to surface tractions. Using these substrates, cell tension profiles can be mapped using a single 3D image stack of a cell of interest. Our goal with this methodology is to make traction force microscopy a more accessible and easier to implement tool for researchers studying cellular mechanotransduction processes, especially newcomers to the field.

This protocol provides instructions for implementing multiphoton lithography to fabricate three-dimensional arrays of fluorescent fiducial markers embedded in poly(ethylene glycol)-based hydrogels for use as reference-free, traction force microscopy platforms. Using these instructions, measurement of 3D material strain and calculation of cellular tractions is simplified to promote high-throughput traction force measurements.

Quantifying cell-induced material deformation provides useful information concerning how cells sense and respond to the physical properties of their ...

Traction Force Microscopy to Study B Lymphocyte Activation

Traction force microscopy (TFM) enables the measurement of forces produced by a cell on a substrate. This technique infers traction force measurements from an experimentally observed displacement field produced by a cell pulling on an elastic substrate. Here, we adapted TFM to investigate the spatial and temporal structure of the force field exerted by B cells when activated by antigen engagement of the B cell receptor. Gel rigidity, bead density, and protein functionalization must be optimized for the study of relatively small cells (~ 6 &#181;m) that interact with, and respond specifically to ligands for cell surface receptors.

Here we present a protocol used to perform traction force microscopy experiments on B cells. We describe the preparation of soft polyacrylamide gels and their functionalization, as well as data acquisition at the microscope and a summary of data analysis.

Traction force microscopy (TFM) enables the measurement of forces produced by a cell on a substrate. This technique infers traction force measurements ...

Pattern Generation for Micropattern Traction Microscopy

Micropattern traction microscopy allows control of the shape of single cells and cell clusters. Furthermore, the ability to pattern at the micrometer length scale allows the use of these patterned contact zones for the measurement of traction forces, as each micropatterned dot allows for the formation of a single focal adhesion that then deforms the soft, underlying hydrogel. This approach has been used for a wide range of cell types, including endothelial cells, smooth muscle cells, fibroblasts, platelets, and epithelial cells. 
This review describes the evolution of techniques that allow the printing of extracellular matrix proteins onto polyacrylamide hydrogels in a regular array of dots of prespecified size and spacing. As micrometer-scale patterns are difficult to directly print onto soft substrates, patterns are first generated on rigid glass coverslips that are then used to transfer the pattern to the hydrogel during gelation. First, the original microcontact printing approach to generate arrays of small dots on the coverslip is described. A second step that removes most of the pattern to leave islands of small dots is required to control the shapes of cells and cell clusters on such arrays of patterned dots. 
Next, an evolution of this approach that allows for the generation of islands of dots using a single subtractive patterning step is described. This approach is greatly simplified for the user but has the disadvantage of a decreased lifetime for the master mold needed to make the patterns. Finally, the computational approaches that have been developed for the analysis of images of displaced dots and subsequent cell-generated traction fields are described, and updated versions of these analysis packages are provided.

We describe improvements to a standard method for measuring cellular traction forces, based on microcontact printing with a single subtractive patterning step of dot arrays of extracellular matrix proteins on soft hydrogels. This method allows for simpler and more consistent fabrication of island patterns, essential for controlling cell cluster shape.

Micropattern traction microscopy allows control of the shape of single cells and cell clusters. Furthermore, the ability to pattern at the micrometer ...

Protrusion Force Microscopy: A Method to Quantify Forces Developed by Cell Protrusions

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