We thank Mrs. Rebecca Alvarado for the support in the protocol video production and Mr. Stephen Casale for constructive criticism on the manuscript. We thank the colleagues from the Department of Cellular Biophysics, Max Planck Institute for Medical Research for the helpful discussions. The financial support from the Max-Planck-Gesellschaft to E.A.C.-A. and the Deutsche Forschungsgemeinschaft (DFG SFB1129, Projektnummer 240245660, P15 to E.A.C.-A. and P4 to U.S.S.; DFG EXC 2082/1-390761711 to U.S.S.) is also greatly acknowledged. J.B. thanks the Carl Zeiss Foundation for financial support. E.A.C.-A., C.S. and U.S.S. acknowledge funding through the Max Planck School Matter to Life supported by the German Federal Ministry of Education and Research (BMBF). C.S. is supported by the European Research Council (Consolidator Grant PHOTOMECH, no.101001797).

1. Preparation of methacrylated coverslips
NOTE: Glass coverslips are methacrylated to covalently link PA hydrogels and therefore prevent their detachment during the incubation with cells. Microscope glass coverslips are used to achieve high image quality.
<ol>
	<li>To prepare a 300 mL solution, take a clean 400 mL glass beaker and place it inside a fume hood.</li>
	<li>Add 10 mL of double distilled water (ddH2O) into the beaker. Add 18.75 mL of acetic acid (&#8805;99.8% pro analysis, p.a.), 18.75 mL of 3-(Trimethoxysilyl)propyl methacrylate and 252.5 mL of ethanol (&#8805;99.8% p.a.) using a serological pipette. Pour the solution into a crystallizing dish.</li>
	<li>Clean 24 mm round glass coverslips with precision wipes. Place the coverslips in a custom-made polytetrafluorethylene rack.</li>
	<li>Immerse the rack into the solution and incubate for 15 min at room temperature (RT).</li>
	<li>Take the rack and rinse all coverslips with ethanol (99.8% p.a.). Dry the coverslips under air flow. 
	NOTE: Methacrylated coverslips can be stored for up to 1 month at RT.</li>
</ol>
2. Micropatterning of extracellular matrix proteins on glass coverslips
NOTE: Glass coverslips are first coated with a layer of molecules, comprising of protein and cell repellent. This layer is then removed using a photopatterning technique, to allow the subsequent deposition of extracellular matrix proteins in micropatterned regions.
<ol>
	<li>Take a 15 mm round glass coverslip and place it on a Petri dish. Use a diamond pen to mark the upper side of the coverslip. Then, proceed to cleaning via oxygen plasma treatment at 0.4 mbar and 200 W for 2 min.</li>
	<li>Pipette 100 &#181;L of 0.01% poly-L-lysine solution on the surface of each coverslip. Incubate for 30 min at RT. Wash the coverslips with 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) pH 8.5. Remove any excess liquid but keep the surface wet.</li>
	<li>Pipette 100 &#181;L of 50 mg/mL poly(ethylene glycol) methyl ether succinimidyl valeric acid (mPEG-SVA) in 10 mM HEPES pH 8.5 on the surface of each coverslip. Incubate for 1 h at RT. Rinse the coverslips with 10 mM HEPES pH 8.5 and dry off under air flow.</li>
	<li>Add 2 &#181;L of UV-sensitive photoinitiator (e.g., PLPP-gel) followed by 40 &#181;L of ethanol (&#8805;99.8% p.a.) onto the surface. To obtain a homogeneous distribution of the gel and ethanol on the surface, gently tilt the Petri dish back and forth. Wait for 5 min for the solution to polymerize.</li>
	<li>Place a single coverslip inside a 35 mm Petri dish with a 20 mm hole at the bottom. This allows the laser beam coming from underneath to directly reach the glass surface.</li>
	<li>Turn on the microscope and a light patterning module (working with this device is only permitted after a safety introduction from the laser safety officers). Calibrate the UV-A laser on the 20x air objective. Put the sample with the photoinitiator on the stage. Adjust the focus on the surface of the glass and turn on the focus control. Load and lock the pre-drawn pattern.
	<ol>
		<li>Prepare the pattern using a graphics software like Inkscape. Ensure that the pattern file does not exceed the DMD dimensions (1824 x 1140 px, which corresponds to 552 x 325 &#181;m on 20 x lens (0.28 &#181;m/px)). 
		NOTE: It is recommended to use the DMD size (1824 x 1140 pixels) as the pattern file size, as this will ensure no error during the patterning. For example, do not produce a pattern file with 1830 x 1130 pixels dimension.</li>
		<li>For the purpose of pattern transfer to polyacrylamide, ensure that the patterning is done only up to 60%-70 % of the radius from the center of the glass to the edge.</li>
		<li>For patterning a single cell, use a diameter of 50-100 &#181;m with the distance of 100 &#181;m between patterns and a diameter of 150-300 &#181;m for a cell group. The appropriate pattern size highly depends on the typical cell size, and it has to be sufficiently large to allow cells to adhere.</li>
	</ol>
	</li>
	<li>Start the patterning by UV dose of 30 mJ/mm2. The patterning duration depends on the number of patterns. Making a single pattern takes approximately 1 s.</li>
	<li>After completing the patterning step, rinse the surface with phosphate-buffered saline (PBS) three times. Incubate the sample with 100 &#181;L of 25 &#181;g/mL fibronectin dissolved in 1x PBS and 25 &#181;g/mL fibrinogen Alexa488 conjugate in PBS for 1 h at RT. For other ECM proteins, find the optimal concentrations completely covering the patterned regions.</li>
	<li>Rinse the surface with 1x PBS three times. Ensure that the patterned glass is immediately used for glass-PA transfer. Store the patterned glass in PBS at RT during the hydrogel preparation.</li>
</ol>
3. Fabrication of patterned polyacrylamide hydrogels
NOTE: Polyacrylamide hydrogels are prepared, including oxidized N-hydroxyethyl acrylamide (HEA) to present reactive aldehyde groups for the covalent binding of matrix proteins on the surface. Additionally, a dual-layer approach to embed fluorescent beads only on the top layer of the hydrogel is used to improve the recording of bead displacement during traction force microscopy experiments.
<ol>
	<li>Put the methacrylated glass coverslips into a Petri dish.</li>
	<li>To prepare fresh oxidized HEA solution, add 9.55 mL of double-distilled water into a 15 mL centrifuge tube. Then, add 0.5 mL of HEA, and 42 mg of sodium(meta)periodate to get 10 mL of the solution. Continuously stir the solution in the dark for 4 h.</li>
	<li>Mix acrylamide, bis-acrylamide, and double-distilled water according to Table 1, to get 10 mL of stock solution hydrogel (do it under the fume hood, monomers are neurotoxic). Degas and close the lid to an airtight seal. 
	NOTE: The stock solution can be kept in aliquots for up to 1 year at 4 &#176;C. Always characterize the Young&#39;s modulus of the hydrogel prior to use.</li>
	<li>Prepare a dual-layered PA hydrogel (do it under the fume hood, monomers are neurotoxic).
	<ol>
		<li>Start with the bottom layer by gently mixing 99.3 &#181;L of stock solution, 0.5 &#181;L of 1% ammonium persulfate (APS), and 0.2 &#181;L of tetramethylethylenediamine (TEMED) in a 1.5 mL tube. Take 10 &#181;L from the solution and pipette it dropwise on the center of the methacrylated coverslip.</li>
		<li>Carefully place a 15 mm round coverslip on the droplet and wait for 45 min for polymerization. Gently detach the top coverslip with a scalpel.</li>
		<li>For the top layer, gently mix 93.3 &#181;Lof stock solution with 1 &#181;L of fresh oxidized HEA solution, 5 &#181;L of fluorescent beads (corresponding to three beads per square micron for beads of 200 nm sized beads), 0.5 &#181;L of 1% APS, and 0.2 &#181;L of TEMED in a 1.5 mL tube. Take 5 &#181;L from the solution and pipette it dropwise on the center of the already polymerized bottom layer.</li>
		<li>Gently place the micropatterned coverslip on the droplet. Wait for 45 min at RT for the droplet to polymerize. Make sure the patterned side touches the droplet. Gently detach the coverslip with a scalpel.</li>
	</ol>
	</li>
	<li>Glue the 24 mm coverslips to the bottom of custom-drilled 6-well plates using two-component silicone-glue. After 5 min, add PBS to the wells.</li>
	<li>Characterize the Young&#39;s modulus of the polyacrylamide hydrogel samples by atomic force microscopy (AFM).
	<ol>
		<li>Mount a spherical tip cantilever on the AFM holder.</li>
		<li>Calibrate each cantilever by acquiring a force-distance curve (3-4 V setpoint) on a hard substrate (e.g., glass or mica), extrapolate the cantilever sensitivity, retract from the surface, and perform a thermal tune to get their spring constant.</li>
		<li>Place the calibrated cantilevers over the hydrogel sample and acquire force curves in PBS buffer, using the following parameters: 20-30 nN force setpoint, 5 or 10 &#181;m/s approach and retraction speed, 5 &#181;m ramp size. 
		NOTE: An inverted optical microscope, equipped with an air 40x objective and a green fluorescent protein (GFP) filter was used to visualize and target ECM-micropatterned areas within the PA hydrogel samples.</li>
		<li>Plot the acquired force versus distance curves with the AFM analysis software.</li>
		<li>Find the contact point and convert force versus distance curves into force versus indentation curves.</li>
		<li>Fit the indentation part of the curve with Hertz model (or a suitable mechanics model in function of the tip geometry), using a Poisson ratio between 0.2 and 0.5.</li>
	</ol>
	</li>
</ol>
4. Local release of cell traction forces by UV-A laser illumination (LUVI-TFM)
NOTE: UV-A laser is applied to release traction forces in cells localized in defined regions of the hydrogels. The UV-A laser (i.e., &#955; = 375 nm solid-state laser, &#60;15 mW) is a class 3B laser. The unshielded laser beam is dangerous to the eyes and often for the skin. Reflected and scattered light and radiation can be dangerous. Moreover, UV radiation may cause skin cancer. Working with devices is only permitted after safety introduction from the laser safety officers.
<ol>
	<li>Aspirate the PBS from the wells.</li>
	<li>Seed 3 x 106 cells per 6-well plate in growth medium. For fibroblasts, use high glucose Dulbecco&#39;s Modified Eagle Medium (DMEM) containing L-glutamine and supplemented with 10% Foetal Bovine Serum (FBS) and 1% penicillin/streptomycin. Incubate the cells overnight at 37 &#176;C/5% CO2. Wash samples to remove floating cells before starting microscopy.</li>
	<li>Turn on the microscope incubation chamber and gas supply to obtain a temperature of 37 &#176;C and 5% CO2 atmosphere.</li>
	<li>Turn on the microscope and the laser patterning module. If necessary, re-calibrate the UV-A laser on the 20x air objective using the Leonardo software. Place the custom-made 6-well plate with the samples on the stage. Adjust focus on the surface of the PA.</li>
	<li>Setup the illumination pattern and mark the cells of interest. Refocus on the surface of the hydrogel. Acquire the image of the cells in the brightfield channel. Switch to the Cy5 channel (far-red filter, 650 nm excitation, 670 nm emission) and take an image of beads in the deformed state.</li>
	<li>Shift to the laser channel. Turn on the laser for 3 min. In our particular setup, this corresponded to a light dose of 6,000 mJ/mm2.</li>
	<li>Switch to the Cy5 channel and acquire an image of the beads in the undeformed state. 
	NOTE: For experiment using mouse embryonic fibroblasts, wait for 15 min post-exposure to record the reference/undeformed image (See the Representative Results section). It might be different for other type of cells. Repeat steps 4.5-4.7 again for another cell(s) of interest.</li>
	<li>To measure the elevated oxidative stress after UV-A illumination, use the commercially available reagent (CellRox). CellRox is non-fluorescent in the reduced state and, when oxidized by reactive oxygen species, fluoresces with an emission maximum of ~665 nm.</li>
	<li>According to the provided protocol, add 5 &#181;M reagent into the imaging media and incubate for 30 min at 37 &#176;C/5% CO2. Then, wash the cells 3x with warm 1x PBS and replace the imaging media. Record the Cy5 signal by fluorescence imaging before and after UV-A illumination using a similar light exposure.</li>
</ol>
5. Image processing, particle image velocimetry and calculation of traction forces
NOTE: Cell traction forces are recorded by analyzing displacement of fluorescent beads and calculated using image analysis tools based on particle image velocimetry.
<ol>
	<li>Open fluorescent beads images, before (i.e., deformed) and after laser (i.e., undeformed) using Fiji. Merge two images as one stack (Image | Stacks | Images to Stack).</li>
	<li>Correct lateral drift between the two images using StackReg plugin (P. Th&#233;venaz, Swiss Federal Institute of Technology Lausanne). Change images to 8 bit (Image | Type | 8 bit) and re-scale to 1024 x 1024 pix (Image | Scale). Save as an Image Sequence (TIFF format) with the reference image always at the beginning.</li>
	<li>Acquire the displacement vector field using a cross-correlation technique19 from the PIV-field as implemented in the OpenPIV project.
	<ol>
		<li>Ensure that the relaxed (undeformed) image and the deformed image are covered with search windows of size wR and wD in pixels. For each search window, define a cross-correlation function using the following formula: 
		<img alt="Equation 1" src="/files/ftp_upload/63121/63121eq01.jpg" /> 
		Here, R(i,j) and D(i,j) describe the intensity fields of the two images truncated to the selected search window and subsequently mirror padded. <img alt="Equation 2" src="/files/ftp_upload/63121/63121eq02.jpg" /> and <img alt="Equation 3" src="/files/ftp_upload/63121/63121eq03.jpg" /> describe their mean value. The window sizes are chosen to be wR = 32 and wD = 32 and an overlap of 70% is used between the neighboring search windows.</li>
		<li>For each search window determine a displacement vector <img alt="Equation 4" src="/files/ftp_upload/63121/63121eq04.jpg" /> that optimizes the cross-correlation function and assign to the center position of the search window. Subpixel accuracy is obtained using a Gaussian fit around the maxima region as described20.</li>
		<li>Find and remove ambiguous displacement vectors. Displacement vectors corresponding to search windows where the ratio between two highest local maxima of the cross-correlation function below a threshold of 1.5 are considered as ambiguous21.</li>
		<li>Discard displacement vectors that fail the normalized median test for a residual threshold of 1.522.</li>
		<li>(Optional) Correct the remaining lateral drift by subtracting a fixed vector <img alt="Equation 5" src="/files/ftp_upload/63121/63121eq05.jpg" /> from all displacements. This is done because the displacement in a selected area far away from the cell vanishes.</li>
		<li>Interpolate the resulting displacement vector field to a regular grid with spacing of 4 px of the input images using a cubic polynomial interpolation. Missing datapoints are filled using a smooth bivariate spline extrapolation. A displacement vector in pixels is related to a local deformation by the pixel ratio of the image (0.33 &#181;m/px for 20x air lens).</li>
		<li>(Optional) Mirror pad the image to reduce ringing artifacts in the subsequent analysis.</li>
		<li>Multiply the deformation field by a Tukey window function to eliminate the edge effect due to the drift correction, with a parameter of 0.2-0.3. In this experiment, the micropatterned area which is in the center of FOV is of interest.</li>
	</ol>
	</li>
	<li>Reconstruct the traction using regularized Fourier Transform Traction Cytometry (FTTC)18. As regularization we use the simplest reasonable choice, namely, 0th order Tikhonov (L2) regularization23,24,25. An optimal regularization parameter &#955; is chosen such that it minimizes the Generalized Cross Validation (GCV) function defined by26: 
	<img alt="Equation 6" src="/files/ftp_upload/63121/63121eq06.jpg" /> 
	Here&#160;&#964;&#955; is a stacked vector containing the x and y components of the reconstructed traction field for the given value of &#955; for all Fourier sampling modes and G is the linear operator mapping traction fields to their corresponding displacement field, as defined for FTTC. The sum is running over vector components. ui are the x and y components of the displacement field for all Fourier sampling modes. GCV is widely used in the field of ill-posed inverse problems and is a good alternative to the L-curve criterion often used in TFM. A Lanzcos filter is used to reduce artifacts introduces due to the limited frequency space and the displacement field upsampling. The traction calculation depends on the Young&#39;s modulus of the PA (e.g., 11.3 kPa) and Poisson&#39;s ratio (e.g., for PA in the range between 0.2 and 0.5 depending on crosslinker concentration).</li>
</ol>

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S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calculation+of+forces+at+focal+adhesions+from+elastic+substrate+data:+the+effect+of+localized+force+and+the+need+for+regularization.">Calculation of forces at focal adhesions from elastic substrate data: the effect of localized force and the need for regularization.</a> Biophysical Journal. 83 (3), 1380-1394 (2002).</li><li>Huang, Y., Gompper, G., Sabass, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Bayesian+traction+force+microscopy+method+with+automated+denoising+in+a+user-friendly+software+package.">A Bayesian traction force microscopy method with automated denoising in a user-friendly software package.</a> Computer Physics Communications. 256, 107313 (2020).</li><li>Vedadghavami, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manufacturing+of+hydrogel+biomaterials+with+controlled+mechanical+properties+for+tissue+engineering+applications.">Manufacturing of hydrogel biomaterials with controlled mechanical properties for tissue engineering applications.</a> Acta Biomaterialia. 62, 42-63 (2017).</li><li>Huth, S., Sindt, S., Selhuber-Unkel, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Automated+analysis+of+soft+hydrogel+microindentation:+Impact+of+various+indentation+parameters+on+the+measurement+of+Young's+modulus.">Automated analysis of soft hydrogel microindentation: Impact of various indentation parameters on the measurement of Young's modulus.</a> PLoS One. 14 (8), 0220281 (2019).</li><li>Oria, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Force+loading+explains+spatial+sensing+of+ligands+by+cells.">Force loading explains spatial sensing of ligands by cells.</a> Nature. 552 (7684), 219-224 (2017).</li><li>Buxboim, A., Rajagopal, K., Brown, A. E., Discher, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+deeply+cells+feel:+methods+for+thin+gels.">How deeply cells feel: methods for thin gels.</a> Journal of Physics. Condensed Matter. 22 (19), 194116 (2010).</li><li>Sen, S., Engler, A. J., Discher, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Matrix+strains+induced+by+cells:+Computing+how+far+cells+can+feel.">Matrix strains induced by cells: Computing how far cells can feel.</a> Cell and Molecular Bioengineering. 2 (1), 39-48 (2009).</li><li>Arnold, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+cell+polarization+and+migration+by+a+gradient+of+nanoscale+variations+in+adhesive+ligand+spacing.">Induction of cell polarization and migration by a gradient of nanoscale variations in adhesive ligand spacing.</a> Nano Letters. 8 (7), 2063-2069 (2008).</li><li>Galluzzi, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atomic+force+microscopy+methodology+and+AFMech+Suite+software+for+nanomechanics+on+heterogeneous+soft+materials.">Atomic force microscopy methodology and AFMech Suite software for nanomechanics on heterogeneous soft materials.</a> Nat Communications. 9 (1), 3584 (2018).</li></ol>

The authors have nothing to disclose.

In this protocol, we describe the preparation of micropatterned PA hydrogels containing fluorescent beads, which are used as fiducial markers for TFM studies. Our approach is based on three steps: 1) preparation of dual-layered PA hydrogels; 2) micropatterning of ECM proteins and their transfer onto the hydrogel surface; 3) use of patterned near-UV light for TFM. The experimental setup to analyze cell tractions to the substrate requires the use of linear elastic materials with known stiffness values to calculate the forces related to displacement of the fluorescent beads26. PA hydrogels are easy to prepare, the stiffness can be easily tuned, and they are commonly used for rigidity sensing and TFM18,28. However, to obtain reproducible polymerization times and homogeneous polymerization of the entire hydrogel, attention should be paid to storing conditions and time for the reagents, e.g., APS should be kept in a desiccator to avoid losing its activity; TEMED should be protected from direct light. The use of oxidized HEA allows the covalent binding of matrix proteins on the hydrogel surface, which might be advantageous to achieve the formation of a full and stable protein layer. The oxidized HEA solution should be prepared fresh every time PA hydrogels are fabricated. The dual-layered PA hydrogel offers three main advantages: 1) it provides an alternative way to reproducibly get a homogenous distribution of fiducial beads near the hydrogel surface, without the need for making the gel extremely thin (i.e., &lt;20 &#181;m). Control over hydrogel thickness is crucial for obtaining accurate measurements with TFM. When the elastic substrate is too thin, strong adherent cells such as fibroblasts may sense and mechanically respond to the underlying rigid glass substrate29,30. Thick hydrogels make the image acquisition for the force reconstruction more challenging. Moreover, many microscopes will not have sufficient space to accommodate them, considering the additional thickness of the glass substrate used to attach the hydrogel, unless ultrathin microscopy slides are used. 2) In the dual-layered PA hydrogel, the homogenous distribution of fiducial beads near the surface of the PA hydrogel is achieved without using a centrifuge, but rather with a simple incubation of precise amounts of hydrogel solutions and fluorescent beads. High bead density is of significant advantage when performing the PIV analysis because it increases the resolution of the traction forces and the signal-to-noise ratio without the need for confocal microscopy. 3) Confining beads in a thin layer close to the cell-material interface enables imaging of traction forces with epifluorescence microscopes as well as confocal microscopes. When preparing the hydrogel, the user should make sure that it firmly adheres to the bottom glass before proceeding with the subsequent steps of the protocol. We recommend following the incubation time indicated for the polymerization of the hydrogel layers, as it might be difficult to remove the glass on top of the surface without damaging the hydrogel surface.
The most well-known techniques to measure elastic properties are AFM, nanoindentation, tensile tests, and rheometry. However, nanoindentation induces very high strains on the materials that can affect the determination of elastic properties. Tensile tests and rheometry on the other hand are macroscopic measurement techniques, whereas cells interact on a microscopic scale31,32. AFM allows measurements at the microscale with reduced strains under physiological conditions. The reliability of AFM measurements can be adversely affected if experimental details are missing (e.g., indentation force and speed) or insufficient data is recorded27. Huth et al. describes an algorithm to extract Young's moduli from AFM data, which emphasizes keeping measurement details constant27. This algorithm offers a precise and reliable determination of Young's moduli and was used for our experiments. Additionally, we measured many curves on samples fabricated on different days and gained very similar results (variation of mean values of ca. 1-2 kPa). This shows that the stiffness of our gels can be reliably predicted.
In this protocol, we use a photopatterning module to create micropatterned regions on glass, which are then transferred to the hydrogel surfaces. The micropatterning shown in this protocol is based on DMD (digital micro-mirror device)-based maskless near-UV lithography (&#955; = 375 nm)7. A DMD consists of a large number of micromirrors on a chip. A single pixel corresponds to a single micro-mirror. The pixelated pattern image file from a computer is projected through DMD and focused on the surface using an objective lens. For micropatterning of proteins, the focused laser light is used to cleave repellent polymer brushes with the help of a photoinitiator. Afterwards, the exposed regions are filled with ECM proteins. This maskless ablation method offers great flexibility in designing new patterns, as it does not rely on the use of a photomask. Designing and applying a pattern is very easy as it only takes several minutes using a freeware like Inkscape. However, the number of patterns and sample produced in a short amount of time is a major drawback, as this method can only be used to pattern a single substrate each time. The photopatterning module uses a near UV solid-state laser source that emits several milliwatts. The unshielded laser beam is dangerous to the eyes and skin. Reflected and scattered light and radiation can also be dangerous. Handling must be accompanied by a safety instruction from laser officers. The most critical step in the protocol when using a photopatterning module is to ensure that during the micropatterning and the ablation the laser is properly focused on the surface. A consistent illumination dose (intensity multiplied by time) of UV-light during patterning is dependent on how the laser is focused on the surface. A weak intensity on the surface due to poor focus may result in the failure of ECM transfer to the hydrogel surface resulting in no cells becoming attached to the hydrogel.
In TFM experiments, after the initial imaging of adherent cells and the fiducial markers, cells are released from the PA hydrogel by trypsin treatment to record their relaxed state. A drawback in performing this step is handling the sample on the microscopy stage. Without a perfusion chamber, opening the lid of the dish, aspirating the medium, rinsing, and pipetting trypsin solution represent a challenge for beginners and experienced users. In fact, these procedures are a major source of the drift in xyz axes, resulting in loss of position and focus. Our local-UV illumination protocol makes TFM a more accessible technique for beginners. It should be noted that we used a commercially available microscopy-based maskless photopatterning module, but in principle any UV-A lighting system could be used, eventually in combination with a mask to protect regions of the substrates, where cell traction forces should not be recorded. Exposing cells with a significant dose of low wavelength visible light (e.g., the violet light that excites DAPI emission) will lead to increasing oxidative stress, which can result in phototoxicity and cell death. Therefore, this technique can be applied even in an epifluorescence microscope without a UV laser module. However, as the intensity is much weaker, it will be much easier to accomplish TFM with the enzymatic treatment in this case.
With LUVI-TFM it is possible to use the same sample for several measurements because of the local detachment of single cells or small groups of cells. However, attention should be paid to the selection of cells to detach, to avoid recording forces from neighboring cells. Thus, for single cell measurements in absence of micropatterns, crowded regions should be avoided; for measurements on single micropatterned cells, the patterns should be designed such that the distance between single structures is at least twice the diameter of the patterned area. We also recommend not using cells from adjacent patterns in sequence, but rather sampling them from distant regions on the substrate. Our measurement is well conducted on an epifluorescence microscope equipped with a focus control feature and a 40 x air NA = 0.9 lens, where the working distance of the lens is sufficiently long and the rapid movement from one well to another is highly tunable. The targeted detachment of cells for TFM applications is effective for measuring cellular force of a single cell or of an entire small cell cluster (e.g., up to 300 &#181;m in diameter). Using our setup, we observed cell rounding and detachment for UV treatment of single cells (Figure 3A), whereas detachment seldom occurred for small cell clusters. This might lead to an underestimation of cell traction forces. For larger cell clusters, enzymatic detachment is recommended, since the users should use a 20x air objective to image the entire cluster and a focus control feature is needed. As the depth of focus of the 20x air lens is much longer, the handling will not be as critical as the higher magnification objective. The user should ensure that the focus is correctly set for the ablation of cells, since the illumination dose is dependent on the laser focus on the surface. While we are aware of the possible limitations in using LUVI-TFM in cell collectives because of possible mechanical interactions with neighboring untreated cells, this aspect could actually turn out to be useful for studies on the mechanics of targeted cell extrusion, e.g., from epithelial monolayers.
In conclusion, with our TFM approach combined with light induced release of micropatterned cells, we provide a robust and high-throughput protocol to measure cell adhesion forces. The versatility of this method could be further exploited using microscopy and imaging setup aimed at improving resolution and sensitivity.

When interacting with their extracellular environment, adherent cells exert forces that are mainly mediated by integrin-based focal adhesions connecting the extracellular matrix (ECM) to the actin cytoskeleton. Focal adhesions are multiprotein assemblies that are centered around the binding of integrins to ECM-proteins like fibronectin and collagen. Integrin clustering and growth of focal adhesions is crucial not only for the establishment of a mechanically stable connection, but also for recruitment of other focal adhesion proteins, including those that activate the RhoA pathway for the regulation of cellular contractility1. The RhoA-dependent contractility of the actin cytoskeleton allows cells to spread and migrate on the underlying ECM, and also to sense its stiffness2. The distribution of traction forces strongly depends on cells&#39; spread area and shape, both of which rely on matrix properties and therefore impact cytoskeletal organization, eventually forming a closed feedback loop between matrix mechanics and cytoskeletal organization3,4.
Surface micropatterning techniques allow a defined control of cell shape by creating micron-sized regions presenting ECM adhesive proteins; according to the size of these regions, single cells, or groups of cells that adhere to the micropattern5. ECM proteins can be patterned on glass substrates by different approaches such as microcontact printing, photo-patterning or laser-patterning6. The use of UV light (&#955; = 185 or 375 nm) combined with surface antifouling strategies offers the flexibility for designing different shapes and sizes and immobilization of multiple protein types with a high precision near surface edges7,8. The regions coated with protein repellent chemicals such as polyethylene glycol (PEG) are protected with either a chromium photomask or a digital mirror device (DMD)-based maskless lithography system. The patterns in the masks allow the exposure to UV light of regions that will then be patterned with ECM proteins. The patterning of hydrogel surfaces with ECM proteins requires a transfer step to remove proteins from the glass surface and crosslink them to the patterns. Alternatively, patterning on soft materials can be achieved by first coating the photomask with a repellent protein, followed by burning the unmasked regions using deep UV illumination. Since deep UV generates ozone and makes the surface reactive for protein binding, the unmasked regions are coated with ECM proteins and finally the gel is polymerized directly on top of the unmasked regions9,10.
The patterned hydrogels can be used to perform TFM, a technique that measures cell forces at the cell-material interface11. In 2D-TFM, one uses the flat surface of a thick polymer film, in which marker beads have been embedded to track deformations12,13,14,15. In order to extract displacement vectors, it is essential to combine two images, one of the deformed state and one reference image without deformations. The two images are then mapped onto each other with image processing. At high marker density, this is usually done with Particle Image Velocimetry (PIV), which is a well-established method to reconstruct hydrodynamic flow. At low marker density and in 3D-TFM, this is usually done with Particle Tracking Velocimetry (PTV), which includes specific features of the experimental data set. An example of a computationally cheaper alternative is optical flow, such as the Kanade-Lucas-Tomasi (KLT) algorithm16. In the case of hydrogel substrates, fluorescent beads are usually embedded at high density during the material polymerization, and images are recorded before and after cell release upon enzymatic detachment. Enzymatic detachment of adherent cells, e.g., by trypsinization, leads to simultaneous release of cellular tractions from all cells on the hydrogels, making it difficult to obtain a detailed analysis from a high number of cells.
Here we present a protocol to prepare micropatterned hydrogels to control cell shape and localization and a method to efficiently measure traction forces in sequence using UV to detach individual cells from the substrate. For traction force measurements, we present a technique to produce dual-layered PA hydrogels, where fluorescent beads are embedded only in the top layer, which increases their density and reduces their vertical spread. Combining UV-mediated release of cellular traction forces with micropatterning makes it possible to obtain spatial control over cell detachment (e.g., of single cells without affecting the adhesion of other cells in the region of interest), provided there is a sufficient distance between patterned cells. Cellular traction is then reconstructed using the most efficient and reliable method for 2D-TFM, namely, Fourier Transform Traction Cytometry (FTTC) with regularization17,18.

<table><tbody><tr><td>3-(Trimethoxysilyl)propyl methacrylate</td><td>#440159</td><td>Sigma Aldrich</td><td></td></tr><tr><td>Acetic acid</td><td>#33209</td><td>Honeywell</td><td></td></tr><tr><td>Acrylamide 40 %</td><td>#1610140</td><td>Bio-Rad</td><td>CAUTION : toxic, work under a fume hood</td></tr><tr><td>AFM cantilever</td><td>#CP-CONT-BSG-A-G</td><td>NanoAndMore</td><td>5 &amp;mu;m spherical tip</td></tr><tr><td>AFM system</td><td>#NanoWizard3</td><td>JPK</td><td>Coupled to optical microscope equipped with 40x air objective and GFP filter</td></tr><tr><td>Ammonium persulfate</td><td>#A3678</td><td>Sigma</td><td></td></tr><tr><td>Bis-acrylamide 2 %</td><td>#1610142</td><td>Bio-Rad</td><td>CAUTION : toxic, work under a fume hood</td></tr><tr><td>Camera sCMOS</td><td>#C11440-42U30</td><td>Hamamatsu</td><td></td></tr><tr><td>Camera sCMOS</td><td>#ANDORZYLA4.2</td><td>Oxford Instrument</td><td></td></tr><tr><td>CellRox</td><td>#C10422</td><td>Thermo Fisher</td><td></td></tr><tr><td>Custom incubator chamber</td><td></td><td>EMBLEM</td><td></td></tr><tr><td>Dental glue</td><td>#1300 100</td><td>Picodent</td><td></td></tr><tr><td>DMEM</td><td>#41965</td><td>Thermo Fisher</td><td></td></tr><tr><td>Epifluorescence microscope</td><td>#Eclipse Ti2E</td><td>Nikon</td><td></td></tr><tr><td>Epifluorescence microscope</td><td>#Axiovert200</td><td>Zeiss</td><td></td></tr><tr><td>Ethanol</td><td>#9065.3</td><td>Carl Roth</td><td></td></tr><tr><td>FBS South America</td><td>#S181T</td><td>Thermo Fisher</td><td></td></tr><tr><td>Fibrinogen</td><td>#F13191</td><td>Invitrogen</td><td>Alexa488 conjugate</td></tr><tr><td>Fibronectin</td><td>#F1141</td><td>Sigma</td><td></td></tr><tr><td>Fluorescent beads 200 nm</td><td>#F8805</td><td>Invitrogen</td><td>Carboxylated (365/415)</td></tr><tr><td>Fluorescent beads 200 nm</td><td>#F8848</td><td>Invitrogen</td><td>Carboxylated (505/515)</td></tr><tr><td>Fluorescent beads 200 nm</td><td>#F8810</td><td>Invitrogen</td><td>Carboxylated (580/605)</td></tr><tr><td>Fluorescent beads 200 nm</td><td>#F8807</td><td>Invitrogen</td><td>Carboxylated (660/680)</td></tr><tr><td>Glass coverslip 15 mm round</td><td>#41001115</td><td>Assistent</td><td></td></tr><tr><td>Glass coverslip 24 mm round</td><td>#41001124</td><td>Assistent</td><td></td></tr><tr><td>Microscope Objective</td><td>#MRH08230</td><td>Nikon</td><td>20x air NA 0.45</td></tr><tr><td>Mouse Embryonic Fibroblasts</td><td>#CRL-2991</td><td>ATCC</td><td></td></tr><tr><td>mPEG-SVA</td><td>#MPEG-SVA-5000</td><td>Laysan Bio</td><td></td></tr><tr><td>N-Hydroxyethyl acrylamide</td><td>#697931</td><td>Aldrich</td><td></td></tr><tr><td>Plasma cleaner</td><td>#100-E</td><td>TePla</td><td></td></tr><tr><td>PLPP gel</td><td>#PLPPgel</td><td>Alveole</td><td></td></tr><tr><td>Poly-L-lysine</td><td>#P4823</td><td>Sigma Aldrich</td><td></td></tr><tr><td>Poly(L-lysine)-graft-poly(ethylene glycol)</td><td>#PLL(20-g[3.5]-PEG(2)</td><td>SuSoS</td><td></td></tr><tr><td>Sodium(meta) periodate</td><td>#S1878</td><td>Sigma Aldrich</td><td></td></tr><tr><td>TEMED</td><td>#17919</td><td>Thermo Scientific</td><td></td></tr><tr><td>UV Patterning module</td><td>#PRIMO</td><td>Alveole</td><td></td></tr><tr><td>UVO cleaner</td><td>#342-220</td><td>Jelight</td><td></td></tr></tbody></table>

control of cell adhesion using hydrogel patterning techniques for applications in traction force microscopy

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat soft substrates that deform under cell traction (2D-TFM). TFM relies on the use of linear elastic materials, such as polydimethylsiloxane (PDMS) or polyacrylamide (PA). For 2D-TFM on PA, the difficulty in achieving high throughput results mainly from the large variability of cell shapes and tractions, calling for standardization. We present a protocol to rapidly and efficiently fabricate micropatterned PA hydrogels for 2D-TFM studies. The micropatterns are first created by maskless photolithography using near-UV light where extracellular matrix proteins bind only to the micropatterned regions, while the rest of the surface remains non-adhesive for cells. The micropatterning of extracellular matrix proteins is due to the presence of active aldehyde groups, resulting in adhesive regions of different shapes to accommodate either single cells or groups of cells. For TFM measurements, we use PA hydrogels of different elasticity by varying the amounts of acrylamide and bis-acrylamide and tracking the displacement of embedded fluorescent beads to reconstruct cell traction fields with regularized Fourier Transform Traction Cytometry (FTTC).
To further achieve precise recording of cell forces, we describe the use of a controlled dose of patterned light to release cell tractions in defined regions for single cells or groups of cells. We call this method local UV illumination traction force microscopy (LUVI-TFM). With enzymatic treatment, all cells are detached from the sample simultaneously, whereas with LUVI-TFM traction forces of cells in different regions of the sample can be recorded in sequence. We demonstrate the applicability of this protocol (i) to study cell traction forces as a function of controlled adhesion to the substrate, and (ii) to achieve a greater number of experimental observations from the same sample.

The PA hydrogels were polymerized on methacrylated glass coverslips, which present reactive groups for the covalent linkage of the PA. In this way, when placing the hydrogels in an aqueous solution, their detachment from the substrate was prevented (Figure 1A-D). To obtain a high density of fiducial markers near the hydrogel surface, we developed the preparation of a dual-layered PA hydrogel. The bottom layer, without any fluorescent beads, was polymerized on the methacrylated coverslip. Subsequently, another layer containing beads was polymerized on top of the bottom layer (Figure 1E-H), replacing the need for sedimentation, which is often used to achieve bead distribution in a single focal plane and increase bead density. To achieve control over cell shape during TFM measurements, we produced micropatterned PA hydrogels via direct transfer of fibronectin-patterned microstructures from a glass coverslip to the pre-polymerized PA (Figure 1F-F&#39;). The addition of 1% non-conventional crosslinker (oxidized HEA) in the top hydrogel layer solution provides aldehyde groups that covalently bind to amine groups of fibronectin.
We prepared PA hydrogels, which are approximately 60 &#181;m in thickness and confined fluorescent beads in the upper layer close to the hydrogel surface. This type of hydrogel is a suitable substrate for imaging cellular tractions with inverted microscopes and thick enough (minimum thickness to perform TFM has been reported to be 20-30 &#181;m)18 to prevent any impact of the glass substrate. Localization of the fluorescent beads was imaged by confocal microscopy (Figure 1I). To visualize protein transfer on the hydrogel, we used fluorescently labeled ECM proteins and imaged them by epifluorescence microscopy (Figure 1J). We prepared micropatterned circles of fibronectin with a diameter of 100 &#181;m (left side) and 50 &#181;m (right side) to allow adhesion of groups of cells or single cells, as shown in the overlay of bright field images of adherent cells, and fluorescently labeled fiducial beads and fibronectin micropatterns. On a different sample, the adhesion response of single cells to micropatterned fibronectin was validated by indirect immunofluorescence microscopy to image the localization of focal adhesion proteins such as paxillin (Figure 1K).
Both ECM protein coating and hydrogel stiffness are crucial parameters for cell adhesion26. To characterize the mechanical properties of our PA hydrogels, we performed nanoindentation experiments by AFM27, coupled with an epifluorescence microscope. Hydrogel substrates of three different stiffnesses were prepared and tested with our setup (Figure 2 and Table 1). Sphere-shaped AFM cantilevers were cyclically approached and retracted from unpatterned PA hydrogels and fibrinogen conjugated to Alexa488-micropatterned areas, while recording force-distance curves. Subsequently, contact point evaluation and application of Hertz model allowed us to estimate the Young&#39;s modulus (E) of each sample. ECM micropatterning did not alter the Young&#39;s modulus, which remained comparable to each of the unpatterned PA hydrogels.
To measure traction forces exerted by adherent cells on the substrate, we developed an experimental setup for reference based TFM carried out on a widefield epifluorescence microscope equipped with near UV laser module (UV-A 6000 mJ/mm2) to release cell tractions (Figure 3A). As the laser beam illumination can be spatio-temporally controlled, not only is it possible to selectively expose a single cell or a cell cluster with a high laser dose, but also and more importantly, the disruptive intermediate step of cell removal from the entire sample using a digestive enzyme like trypsin is no longer necessary.Illuminating cells with such a high laser dose induced an elevated oxidative stress leading to cell death (Figure 3B). Cell death yielded the release of tractions from the substrate, as indicated by the bead displacement (illustrated in Figure 3A). We combined the UV-A illumination with a light patterning module to selectively illuminate micron-sized regions of the PA hydrogels (Figure 3B-C). In this way it is possible to release the tractions of micropatterned cell clusters (Figure 3C,F) or single cells (Figure 3B). Importantly, at our time scales the mechanical properties of the ECM patterned hydrogels were not significantly affected by UV exposure (Figure 2C). Increase in oxidative stress is shown in Figure 3D,E. The traction forces were reconstructed by using regularized FTTC with a regularization parameter chosen by Generalized Cross Validation (Figure 3B,F). The release of forces for single cells occurred over a period of 15 min, and the result of the LUVI-TFM is comparable to the trypsin-based TFM (Figure 3G-H).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63121/63121fig01.jpg" /> 
Figure 1: Scheme of patterned fibronectin-substrate preparation for TFM. (A) The solution for the bottom layer is pipetted on a methacrylated surface. (B) A smaller clean coverslip is carefully placed on the droplet. (C) The gelation time is 45 min. (D) The top coverslip is detached. The bottom layer is ready. (E) The solution for the top layer is pipetted on the hydrogel. (F) The patterned coverslip is carefully placed on the droplet. (F&#39;) The micropatterning of adhesive proteins on glass is produced by maskless near UV-lithography, and then transferred from glass to PA hydrogel. The free amine groups of adhesive proteins, e.g., fibronectin, bind covalently to aldehyde groups on the PA surface. (G) The gelation time is 45 min. (H) The top coverslip is detached. The patterned PA hydrogel is ready. (I) A 3D representation of a confocal xyz micrograph showing a high density of fiducial beads near the surface. (J) A micrograph of fluorescently labeled fibronectin (magenta) on the surface of PA with embedded fluorescent beads (green), overlayed with a bright field image of cells. (K) Indirect immunofluorescence microscopy imaging of a single cell adhering to a circle-shaped fibronectin micropattern (100 &#181;m). Nucleus (blue), paxillin (red). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63121/63121fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63121/63121fig02.jpg" /> 
Figure 2: Characterization of sample mechanical properties by AFM. (A) AFM nanoindentation experimental setup. A sphere-shaped cantilever is used to probe ECM micropatterns before or after UV treatment, and the non-patterned regions of dual layered PA (5% Acrylamide, 0.3% Bis-acrylamide) areas. (B) Representative force versus indentation curve for ECM micropattern (black). Hertz fit (red) is used to calculate the Young&#39;s modulus (E) of the sample. (C) Mechanical measurements for non-patterned dual layered PA areas (no ECM), ECM micropattern and ECM micropattern after UV-A exposure. The bars show mean &#177; S.E.M. (Brown-Forsythe and Welch ANOVA test). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63121/63121fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63121/63121fig03.jpg" /> 
Figure 3. Local UV-A illumination TFM (LUVI-TFM) enables local traction force measurements inside a large field of view. (A) Scheme of local UV-A illumination TFM. Cells are treated with a high dose of UV-A laser to get the undeformed (reference) image. (B) The brightfield image of a single cell before UV-A laser illumination. Middle: The brightfield image of a single cell after UV-A laser illumination. Right: Traction force of a single MEF adhering to a fibronectin coated PA hydrogel (E = 5.74 kPa) illuminated with a 50 &#181;m diameter UV-light beam (red dashed line). (C) Left: Recording of traction forces of a cluster of mouse embryonic fibroblasts (MEF) adhering to micropatterned fibronectin (circle, 100 &#181;m diameter). The cluster was illuminated with a 200 &#181;m diameter UV-light beam (red dashed line). Right: Recording of traction forces of a cluster of mouse embryonic fibroblasts (MEF) adhering to micropatterned fibronectin (circle, 300 &#181;m diameter). The cluster was illuminated with a 300 &#181;m diameter UV-light beam (red dashed line). Scale bar = 200 &#181;m. (D,E) An increase in oxidative stress in cells indicated by the increased Cy5 signal intensity after a high UV-A laser dose is detected by fluorescence microscopy. The oxidative stress leads to cell death.&#160;Scale bar = 200 &#181;m. (F) Left: The stress heat map from a cluster of mouse embryonic fibroblasts (MEF) adhering to micropatterned fibronectin (circle, 100 &#181;m diameter). Right: The stress heat map from a cluster of mouse embryonic fibroblasts (MEF) adhering to micropatterned fibronectin (circle, 300 &#181;m diameter). (G)&#160;MEF cells adhering to 100 &#181;m fibronectin micropatterns slowly release their tractions after UV-A exposure. Full release is achieved after 15 min (the reference image was then taken 15 min post exposure). (H) A comparison with the conventional trypsin based-TFM for MEF cells adhering to 300 &#181;m diameter patterned-fibronectin. The result of UV-A illumination (6,000 mJ/mm2) followed by 15 min waiting is non-significantly different from the conventional trypsin treatment (i.e., 0.05% for 20 min). The bars show mean S.E.M. (two-tailed Student&#39;s t-test, ****P &#60; 0.0001, * P &#60; 0.05, ns P &#8804;0.5). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/63121/63121fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Acrylamide (%)</td>
			<td>Bis-acrylamide (%)</td>
			<td>Acrylamide from 40 % stock solution (ml)</td>
			<td>Bis-acrylamide from 2 % stock solution (ml)</td>
			<td>Water (ml)</td>
			<td>Young&#8217;s modulus (kPa)</td>
		</tr>
		<tr>
			<td>4</td>
			<td>0.1</td>
			<td>1</td>
			<td>0.5</td>
			<td>8.5</td>
			<td>5,74 &#177; 0,53</td>
		</tr>
		<tr>
			<td>5</td>
			<td>0.15</td>
			<td>1.25</td>
			<td>0.75</td>
			<td>8</td>
			<td>9,69 &#177; 0,68</td>
		</tr>
		<tr>
			<td>5</td>
			<td>0.3</td>
			<td>1.25</td>
			<td>1.5</td>
			<td>7.25</td>
			<td>11,33 &#177; 1,06</td>
		</tr>
	</tbody>
</table>
Table 1: Hydrogel stock solution mixtures and resulting elasticity
All data are deposited in the Open Access Data Repository of The Max Planck Society (Edmond) and can be accessed through the following address:&#160;https://edmond.mpdl.mpg.de/imeji/collection/JTu8PlWqpbymN9Qf

Watch this Scientific Journal Video about Control of Cell Adhesion using Hydrogel Patterning Techniques for Applications in Traction Force Microscopy at JoVE.com

Control of Cell Adhesion using Hydrogel Patterning Techniques for Applications in Traction Force Microscopy

Near-UV lithography and traction force microscopy are combined to measure cellular forces on micropatterned hydrogels. The targeted light-induced release of single cells enables a high number of measurements on the same sample.

Traction force microscopy (TFM) is the main method used in mechanobiology to measure cell forces. Commonly this is being used for cells adhering to flat ...

control-cell-adhesion-using-hydrogel-patterning-techniques-for

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

5.5K Views.  Max Planck Institute for Medical Research. Near-UV lithography and traction force microscopy are combined to measure cellular forces on micropatterned hydrogels. The targeted light-induced release of single cells enables a high number of measurements on the same sample.

Video: Control of Cell Adhesion using Hydrogel Patterning Techniques for Applications in Traction Force Microscopy

Preparation of Complaint Matrices for Quantifying Cellular Contraction

The regulation of cellular adhesion to the extracellular matrix (ECM) is essential for cell migration and ECM remodeling. Focal adhesions are macromolecular assemblies that couple the contractile F-actin cytoskeleton to the ECM. This connection allows for the transmission of intracellular mechanical forces across the cell membrane to the underlying substrate. Recent work has shown the mechanical properties of the ECM regulate focal adhesion and F-actin morphology as well as numerous physiological processes, including cell differentiation, division, proliferation and migration. Thus, the use of cell culture substrates has become an increasingly prevalent method to precisely control and modulate ECM mechanical properties.
To quantify traction forces at focal adhesions in an adherent cell, compliant substrates are used in conjunction with high-resolution imaging and computational techniques in a method termed traction force microscopy (TFM). This technique relies on measurements of the local magnitude and direction of substrate deformations induced by cellular contraction. In combination with high-resolution fluorescence microscopy of fluorescently tagged proteins, it is possible to correlate cytoskeletal organization and remodeling with traction forces.
Here we present a detailed experimental protocol for the preparation of two-dimensional, compliant matrices for the purpose of creating a cell culture substrate with a well-characterized, tunable mechanical stiffness, which is suitable for measuring cellular contraction. These protocols include the fabrication of polyacrylamide hydrogels, coating of ECM proteins on such gels, plating cells on gels, and high-resolution confocal microscopy using a perfusion chamber. Additionally, we provide a representative sample of data demonstrating location and magnitude of cellular forces using cited TFM protocols.

In this video, we demonstrate the experimental techniques used to fabricate compliant, extracellular matrix (ECM) coated substrates suitable for cell culture, and which are amenable to traction force microscopy and observing effects of ECM stiffness on cell behavior.

The regulation of cellular adhesion to the extracellular matrix (ECM) is essential for cell migration and ECM remodeling.  Focal adhesions are ...

Creating Adhesive and Soluble Gradients for Imaging Cell Migration with Fluorescence Microscopy

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as growth factors. Here, we outline a method to create opposing gradients of adhesive and soluble cues in a microfluidic chamber, which is compatible with live cell imaging. A copolymer of poly-L-lysine and polyethylene glycol (PLL-PEG) is employed to passivate glass coverslips and prevent non-specific adsorption of biomolecules and cells. Next, microcontact printing or dip pen lithography are used to create tracks of streptavidin on the passivated surfaces to serve as anchoring points for the biotinylated peptide arginine-glycine-aspartic acid (RGD) as the adhesive cue. A microfluidic device is placed onto the modified surface and used to create the gradient of adhesive cues (100% RGD to 0% RGD) on the streptavidin tracks. Finally, the same microfluidic device is used to create a gradient of a chemoattractant such as fetal bovine serum (FBS), as the soluble cue in the opposite direction of the gradient of adhesive cues.

A method for the assembly of adhesive and soluble gradients in a microscopy chamber for live cell migration studies is described. The engineered environment combines antifouling surfaces and adhesive tracks with solution gradients and therefore allows one to determine the relative importance of guidance cues.

Cells can sense and migrate towards higher concentrations of adhesive cues such as the glycoproteins of the extracellular matrix and soluble cues such as ...

Preparation of Hydroxy-PAAm Hydrogels for Decoupling the Effects of Mechanotransduction Cues

It is now well established that many cellular functions are regulated by interactions of cells with physicochemical and mechanical cues of their extracellular matrix (ECM) environment. Eukaryotic cells constantly sense their local microenvironment through surface mechanosensors to transduce physical changes of ECM into biochemical signals, and integrate these signals to achieve specific changes in gene expression. Interestingly, physicochemical and mechanical parameters of the ECM can couple with each other to regulate cell fate. Therefore, a key to understanding mechanotransduction is to decouple the relative contribution of ECM cues on cellular functions.
Here we present a detailed experimental protocol to rapidly and easily generate biologically relevant hydrogels for the independent tuning of mechanotransduction cues in vitro. We chemically modified polyacrylamide hydrogels (PAAm) to surmount their intrinsically non-adhesive properties by incorporating hydroxyl-functionalized acrylamide monomers during the polymerization. We obtained a novel PAAm hydrogel, called hydroxy-PAAm, which permits immobilization of any desired nature of ECM proteins. The combination of hydroxy-PAAm hydrogels with microcontact printing allows to independently control the morphology of single-cells, the matrix stiffness, the nature and the density of ECM proteins. We provide a simple and rapid method that can be set up in every biology lab to study in vitro cell mechanotransduction processes. We validate this novel two-dimensional platform by conducting experiments on endothelial cells that demonstrate a mechanical coupling between ECM stiffness and the nucleus.

We present a new polyacrylamide hydrogel, called hydroxy-PAAm, that allows a direct binding of ECM proteins with minimal cost or expertise. The combination of hydroxy-PAAm hydrogels with microcontact printing facilitates independent control of many cues of the natural cell microenvironment for studying cellular mechanostransduction.

It is now well established that many cellular functions are regulated by interactions of cells with physicochemical and mechanical cues of their ...

A Novel Method for Localizing Reporter Fluorescent Beads Near the Cell Culture Surface for Traction Force Microscopy

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When the substrate is coated with an extracellular matrix protein, cells adhere to the gel and apply forces, causing the gel to deform. The deformation depends on the cell traction and the elastic properties of the gel. If the deformation field of the surface is known, surface traction can be calculated using elasticity theory. Gel deformation is commonly measured by embedding fluorescent marker beads uniformly into the gel. The probes displace as the gel deforms. The probes near the surface of the gel are tracked. The displacements reported by these probes are considered as surface displacements. Their depths from the surface are ignored. This assumption introduces error in traction force evaluations. For precise measurement of cell forces, it is critical for the location of the beads to be known. We have developed a technique that utilizes simple chemistry to confine fluorescent marker beads, 0.1 and 1 &#181;m in diameter, in PA gels, within 1.6 &#956;m of the surface. We coat a coverslip with poly-D-lysine (PDL) and fluorescent beads. PA gel solution is then sandwiched between the coverslip and an adherent surface. The fluorescent beads transfer to the gel solution during curing. After polymerization, the PA gel contains fluorescent beads on a plane close to the gel surface.

Traditional techniques for fabricating polyacrylamide (PA) gels containing fluorescent probes involve sandwiching a gel between an adherent surface and a glass slide. Here, we show that coating this slide with poly-D-lysine (PDL) and fluorescent probes localizes the probes to within 1.6 &#181;m from the gel surface.

PA gels have long been used as a platform to study cell traction forces due to ease of fabrication and the ability to tune their elastic properties. When ...

Light-mediated Formation and Patterning of Hydrogels for Cell Culture Applications

Click chemistries have been investigated for use in numerous biomaterials applications, including drug delivery, tissue engineering, and cell culture. In particular, light-mediated click reactions, such as photoinitiated thiol&#8722;ene and thiol&#8722;yne reactions, afford spatiotemporal control over material properties and allow the design of systems with a high degree of user-directed property control. Fabrication and modification of hydrogel-based biomaterials using the precision afforded by light and the versatility offered by these thiol&#8722;X photoclick chemistries are of growing interest, particularly for the culture of cells within well-defined, biomimetic microenvironments. Here, we describe methods for the photoencapsulation of cells and subsequent photopatterning of biochemical cues within hydrogel matrices using versatile and modular building blocks polymerized by a thiol&#8722;ene photoclick reaction. Specifically, an approach is presented for constructing hydrogels from allyloxycarbonyl (Alloc)-functionalized peptide crosslinks and pendant peptide moieties and thiol-functionalized poly(ethylene glycol) (PEG) that rapidly polymerize in the presence of lithium acylphosphinate photoinitiator and cytocompatible doses of long wavelength ultraviolet (UV) light. Facile techniques to visualize photopatterning and quantify the concentration of peptides added are described. Additionally, methods are established for encapsulating cells, specifically human mesenchymal stem cells, and determining their viability and activity. While the formation and initial patterning of thiol-alloc hydrogels are shown here, these techniques broadly may be applied to a number of other light and radical-initiated material systems (e.g., thiol-norbornene, thiol-acrylate) to generate patterned substrates.

We describe a sequential process for light-mediated formation and subsequent biochemical patterning of synthetic hydrogel matrices for three-dimensional cell culture applications. The construction and modification of hydrogels with cytocompatible photoclick chemistry is demonstrated. Additionally, facile techniques to quantify and observe patterns and determine cell viability within these hydrogels are presented.

Click chemistries have been investigated for use in numerous biomaterials applications, including drug delivery, tissue engineering, and cell culture. In ...

Patterning the Geometry of Human Embryonic Stem Cell Colonies on Compliant Substrates to Control Tissue-Level Mechanics

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that correspond to primary germ layer formation during embryogenesis. Thus, these cells represent a powerful tool with which to examine the mechanisms that drive early human development. We have developed a method to culture human embryonic stem cells in confined colonies on compliant substrates that provides control over both the geometry of the colonies and their mechanical environment in order to recapitulate the physical parameters that underlie embryogenesis. The key feature of this method is the ability to generate polyacrylamide hydrogels with defined patterns of extracellular matrix ligand at the surface to promote cell attachment. This is achieved by fabricating stencils with the desired geometric patterns, using these stencils to create patterns of extracellular matrix ligand on glass coverslips, and transferring these patterns to polyacrylamide hydrogels during polymerization. This method is also compatible with traction force microscopy, allowing the user to measure and map the distribution of cell-generated forces within the confined colonies. In combination with standard biochemical assays, these measurements can be used to examine the role mechanical cues play in fate specification and morphogenesis during early human development.

Extracellular matrix ligands can be patterned onto polyacrylamide hydrogels to enable the culture of human embryonic stem cells in confined colonies on compliant substrates. This method can be combined with traction force microscopy and biochemical assays to examine the interplay between tissue geometry, cell-generated forces, and fate specification.

Human embryonic stem cells demonstrate a unique ability to respond to morphogens in vitro by self-organizing patterns of cell fate specification that ...

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 Microscopy Integrated with Microfluidics for Chemotactic Collective Migration

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical gradient, known as chemotaxis, plays an important role in development, the immune response, wound healing, and cancer metastasis. While chemotaxis modulates the migration of single cells as well as collections of cells in vivo, in vitro research focuses on single-cell chemotaxis, partly due to the lack of the proper experimental tools. To fill that gap, described here is a unique experimental system that combines microfluidics and micropatterning to demonstrate the effects of chemical gradients on collective cell migration. Furthermore, traction microscopy and monolayer stress microscopy are incorporated into the system to characterize changes in cellular force on the substrate as well as between neighboring cells. As proof-of-concept, the migration of micropatterned circular islands of Madin-Darby canine kidney (MDCK) cells is tested under a gradient of hepatocyte growth factor (HGF), a known scattering factor. It is found that cells located near the higher concentration of HGF migrate faster than those on the opposite side within a cell island. Within the same island, cellular traction is similar on both sides, but intercellular stress is much lower on the side of higher HGF concentration. This novel experimental system can provide new opportunities to studying the mechanics of chemotactic migration by cellular collectives.

Collective cell migration in development, wound healing, and cancer metastasis is often guided by the gradients of growth factors or signaling molecules. Described here is an experimental system combining traction microscopy with a microfluidic system and a demonstration of how to quantify the mechanics of collective migration under biochemical gradient.

Cells change migration patterns in response to chemical stimuli, including the gradients of the stimuli. Cellular migration in the direction of a chemical ...

Controlled Strain of 3D Hydrogels under Live Microscopy Imaging

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized in vitro stretching methods. Various available systems use 2D substrate-based stretchers, while the accessibility of 3D techniques to strain soft hydrogels, is more restricted. Here, we describe a method that allows external stretching of soft hydrogels from their circumference, using an elastic silicone strip as the sample carrier. The stretching system utilized in this protocol is constructed from 3D-printed parts and low-cost electronics, making it simple and easy to replicate in other labs. The experimental process begins with polymerizing thick (&#62;100 &#956;m) soft fibrin hydrogels (Elastic Modulus of ~100 Pa) in a cut-out at the center of a silicone strip. Silicone-gel constructs are then attached to the printed-stretching device and placed on the confocal microscope stage. Under live microscopy the stretching device is activated, and the gels are imaged at various stretch magnitudes. Image processing is then used to quantify the resulting gel deformations, demonstrating relatively homogenous strains and fiber alignment throughout the gel&#8217;s 3D thickness (Z-axis). Advantages of this method include the ability to strain extremely soft hydrogels in 3D while executing in situ microscopy, and the freedom to manipulate the geometry and size of the sample according to the user&#8217;s needs. Additionally, with proper adaptation, this method can be used to stretch other types of hydrogels (e.g., collagen, polyacrylamide or polyethylene glycol) and can allow for analysis of cells and tissue response to external forces under more biomimetic 3D conditions.

The presented method involves uniaxial stretching of 3D soft hydrogels embedded in silicone rubber while allowing live confocal microscopy. Characterization of the external and internal hydrogel strains as well as fiber alignment are demonstrated. The device and protocol developed can assess the response of cells to various strain regimes.

External forces are an important factor in tissue formation, development, and maintenance. The effects of these forces are often studied using specialized ...

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 ...