1. Fabrication of Deformable Polyacrylamide Hydrogels with Embedded Fluorescent Microspheres
NOTE: Directions describe polymerization of a 25 kPa hydrogel that is 22 &#956;m in diameter and approximately 66 &#956;m thick. Each or all of these parameters can be modified and directions to do so can be found in Table 1 and in the notes17.
<ol>
	<li>Activation of glass-bottom dishes or coverslips
	<ol>
		<li>Prepare the bind silane working solution for activation of a glass-bottom imaging dish or a coverslip that fits into a live-cell imaging chamber. Mix 950 &#956;L of 95% ethanol, 50 &#956;L of glacial acetic acid, and 5 &#956;L of bind silane (y-methacryloxypropyltrimethoxysilane). 
		NOTE: Using a larger bottom coverslip compared to the top coverslip will give additional room to work when preparing the gel and will facilitate positioning the glass micropipette in later steps. Also, if using a coverslip rather than a glass-bottom imaging dish, clean the coverslip as described in the following section.</li>
		<li>Activate the surface of the glass for 20 s with a corona wand and immediately overlay 50 &#956;L of the bind silane working solution. Allow the solution to dry for 10 min.</li>
		<li>Rinse two times with 95% ethanol, then two times with isopropanol and then allow the coverslips to airdry for approximately 20 min. 
		NOTE: Activated glass can be stored for up to one week in a desiccator.</li>
	</ol>
	</li>
	<li>Cleaning top coverslips
	<ol>
		<li>Clean 22 mm top coverslips by incubating in 2% HCl at 70 &#176;C for 30 min, then wash in ddH2O for 10 min two times.</li>
		<li>Incubate the coverslips in a solution of 2% cuvette cleaning concentrate in ddH2O at 50 &#176;C for 30 min, then wash in ddH2O for 10 min two times.</li>
		<li>Incubate the coverslips in ddH2O at 90 &#176;C for 30 min, then in 70% ethanol at 70 &#176;C for 10 min, and then air dry at 60 &#176;C for a minimum of 2 h. 
		NOTE: Cleaned coverslips can be stored indefinitely in a clean desiccator.</li>
	</ol>
	</li>
	<li>Fluorescent microsphere/bead deposition onto top coverslips
	<ol>
		<li>Sonicate the stock solution of fluorescent microspheres for 1 h in an ultrasonic water bath. Make a working bead solution by diluting bead stock 1:200 in 100% ethanol and sonicate again for 1 h.</li>
		<li>15 min before the bead solution has finished sonicating, thoroughly clean the coverslips by placing them vertically in a ceramic coverslip holder and treating with room-air plasma for 3 min in a tabletop plasma cleaner.</li>
		<li>To facilitate handling and prevent sliding of the coverslip during subsequent steps, place a piece of parafilm in a 60 mm Petri dish lid or a similar container. Place the coverslip in the stabilizer and lightly tap down, ensuring good contact between the parafilm and the coverslip.</li>
		<li>For a 22 mm coverslip, add 150 &#956;L of the working bead solution to the top of the coverslip. Immediately aspirate the ethanol solution off from the side of the coverslip, leaving the beads on the coverslip. Allow the coverslip to airdry. 
		NOTE: The amount of working bead solution added should be ~4 &#956;L/cm2 and can be scaled to accommodate any size coverslip.</li>
	</ol>
	</li>
	<li>Casting hydrogels with embedded fluorescent beads
	<ol>
		<li>Prepare the hydrogel solution of acrylamide and bis-acrylamide. Mix the solution according to Table 1, then add 2.5 &#956;L of 10% APS and 0.5 &#956;L of TEMED. Mix well. Immediately move to the next step. 
		NOTE: The hydrogel solution mixture can be altered to vary the Young&#8217;s modulus, or stiffness, of the hydrogel by changing the ratio of acrylamide to bis-acrylamide as shown in Table 1. These values have been verified for use in the Howe laboratory using atomic force microscopy but should be confirmed within one&#8217;s institution.</li>
		<li>Immediately after making the hydrogel solution, add a 25 &#956;L drop to the activated side of the glass-bottom dish or bottom coverslip, then immediately place the bead-coated coverslip onto the solution, bead side down. Contacting the drop with the far side of the coverslip followed by slow lowering helps avoid trapping air bubbles within the hydrogel. 
		NOTE: The height of the hydrogel should be well within the working distance of the objective lens to be used in the later experiment. A hydrogel height of 66 &#956;m works well for most systems. The size of the hydrogel can be scaled by adding more or less hydrogel solution depending on the size of the coverslip. To calculate the appropriate volume of hydrogel solution, use the equation for the volume of a cylinder, V = &#960;r2h where r is the coverslip radius and h is the desired hydrogel height. Typically, this calculation predicts with fair accuracy the actual height of the hydrogel, as measured by preparing a gel with bead-coated coverslips on both the top and bottom and using a confocal microscope to measure the distance between the two bead planes. However, it has been observed that the actual height of the hydrogel can deviate from this calculation by &#177; 20 &#956;m (e.g., depending on the thickness and manufacturer of the top glass coverslip). Direct measurement of gel height using the method described above is recommended.</li>
		<li>Allow the gel to polymerize for 30 min, then remove the top coverslip gently with forceps. Adding 50 mM HEPES pH 8.5 to the dish can facilitate removal. Wash for 5 min in 50 mM HEPES pH 8.5 three times.</li>
	</ol>
	</li>
	<li>Hydrogel activation and extracellular matrix coating
	<ol>
		<li>Activate the hydrogel surface by incubating in 0.4 mM Sulfo-SANPAH (sulfosuccinimidyl 6-(4&#39;-azido-2&#39;-nitrophenylamino) hexanoate) in 50 mM HEPES pH 8.5. Immediately expose to a UV arc lamp in an enclosed area. 
		NOTE: Protect Sulfo-SANPAH from light prior to activation. For a 400 W lamp, position the gel 10 cm away from bulb within the light box and illuminate for 100 s. The Sulfo-SANPAH solution will change from bright orange to dark brown.</li>
		<li>Wash for 5 min in 50 mM HEPES pH 8.5 three times. 
		NOTE: Hydrated gels may be stored at 4 &#176;C for up to one week.</li>
		<li>Incubate the activated hydrogel in 20 &#956;g/mL fibronectin in 50 mM HEPES pH 8.5 for 1 h at 37 &#176;C.</li>
		<li>Aspirate the fibronectin solution and wash for 5 min in phosphate-buffered saline (PBS) three times. Sterilize the hydrogel and the lid of the dish for 15 min under UV light in a tissue culture hood with a low volume of PBS. Wash once in sterile PBS. 
		NOTE: Other types of ECM protein can be used to coat the hydrogel including collagen and laminin.</li>
	</ol>
	</li>
</ol>
2. Plating cells
<ol>
	<li>Add 3 mL of media containing 21,000 cells to fill a 60 mm dish for a final cell density of ~1000 cells/cm2. Adjust the seeding density as needed to prevent crowding and allow free movement of individual cells.</li>
	<li>Allow the cells to recover at 37 &#176;C for at least 4 h and for up to 18 h before imaging. Prepare for imaging by rinsing with imaging media two times before adding imaging media. Allow the cells to equilibrate for at least 30 min before imaging. 
	NOTE: Screen media conditions in advance to determine conditions that will optimize migration in the cell line being used. For SKOV-3 cells, DMEM without phenol red, containing 20 mM HEPES and 12.5 ng/mL epidermal growth factor stimulates the most migration. Optimal conditions for Ref52 cells are Ringer&#8217;s Buffer with 10% fetal bovine serum (FBS) and 25 ng/mL platelet-derived growth factor.</li>
</ol>
3. Preparation of glass micropipette: pipette pulling and forging
<ol>
	<li>Pull 100 mm long borosilicate glass micropipettes with a 1.0 mm exterior and 0.58 mm interior diameter in a two-step process to obtain a taper over 2 mm that reduces to ~50 &#956;m in the first millimeter and extends to a long, parallel 10 &#956;m diameter tube in the last millimeter.</li>
	<li>Load pulled pipet into a microforge. Shape the pipet to have a 15 &#956;m blunted tip that is enclosed at the very end of a 250 &#956;m section bent at a ~35&#176; angle from the rest of the pipet. The approximate diameter at the bend should be around 30 &#956;m to lend strength to the tip. 
	NOTE: Taper and tip dimensions can be adjusted to properly apply desired force (see step 5). Pulling micropipettes at 65 &#176;C for the first step for 3 mm, and 60 &#176;C for the second step produces the dimensions described in step 3.1. Results using different pipet pullers may vary.</li>
	<li>Sterilize the micropipette in 70% ethanol before use.</li>
</ol>
4. Positioning the micromanipulator and the micropipette
<ol>
	<li>Remove the dish lid and load the dish onto the microscope stage and center. Use a 10X or similarly low magnification objective. Cover the media with mineral oil to prevent evaporation of the media.</li>
	<li>Inserting pulled pipet
	<ol>
		<li>Insert the pulled pipet into the micropipette sheath, pointing the hook down toward the dish. The tip of the hook will be the lowest point when lowered to the gel.</li>
		<li>Insert sheath into micromanipulator and adjust until the tip of the pipet is centered over the objective lens in both X and Y directions.</li>
		<li>Lower the pipet using coarse manipulator until it just touches the surface of the liquid.</li>
	</ol>
	</li>
	<li>Using phase contrast or brightfield, focus the microscope on the bead layer at the top of the gel. This will be the reference plane.</li>
	<li>Ensuring that the objective is in no danger of hitting the sample or stage, bring focus above the gel to find the tip of the micropipette, using small adjustments of the coarse manipulator in the X and Y directions to cast shadows on the focal plane. Only lower the micropipette when certain that the very tip of the pipet is in the field of view.</li>
	<li>Ensure that the blunted tip of micropipette is pointing down by rotating the pipet in the sheath or rotating the sheath in the micromanipulator until the tip is perpendicular to the focal plane. Repeat steps 4.4 and 4.5 as needed. Focus on the tip of the pipet.</li>
	<li>Focus back down to the top bead layer of the gel to gauge how far the pipet is from the gel surface. Focus back up to a plane that is part way between the gel and the tip of the pipet. Slowly lower the pipet to reach the intermediate focal plane.</li>
	<li>Repeat step 4.6 until very faint shadows from the tip of the micropipette can be appreciated when focusing on the hydrogel. Increase to the next highest magnification.</li>
	<li>Lower the micromanipulator until shadows and refractions of the very tip of the micropipette can be appreciated within the bead layer focal plane.</li>
	<li>Increase the magnification to that which will be used in the experiment. Lower the pipet until it hovers just above the surface of the hydrogel.</li>
</ol>
5. Calibrating the micromanipulator and force generation
<ol>
	<li>In phase or brightfield, lower the hovering micropipette to touch the surface of the hydrogel. Observe how the pipet looks upon contact with the hydrogel. Continue to lower micropipette in Z until adjustments in X and Y cause pulling and deflection of the hydrogel in those directions. Use the microspheres or nearby cells as fiduciary marks. 
	NOTE: If the micromanipulator is attached to the phase condenser arm or the bench and not the sample stage itself, always disengage the gel before moving the stage to avoid breaking the pipet or disturbing cells. If the pipet breaks, go back to step 3 and step 4.</li>
	<li>Find an area devoid of cells to engage the gel. Pull it in all directions and get comfortable with the way micromanipulation translates to deformation of the gel.</li>
	<li>Take fluorescent images of the bead field with no manipulation, with the pipet engaging the gel, and with the engaged pipet pulling the gel. Repeat this several times taking good notes regarding the tick marks on the micromanipulator, the way the pipet tip looks in phase or brightfield at each stage of pulling, and the distance the tip moves using that manipulation.</li>
	<li>Use ImageJ as previously described16,17 to calculate relative bead displacements and force applied to the beads by comparing the null bead field to the bead field without pipet engagement, the bead field with the gel engaged, and the pulled gel.</li>
	<li>To fine tune the tensional stimulus, compare force application using differing micropipette tip dimensions, distances from the cell, or distance pulled by the micromanipulator from initial point of touchdown. The effect of the micropipette tip dimension on force application gives great flexibility to the user but also demonstrates the need to generate force maps for new micropipettes, even when the dimensions and shape closely resemble previously calibrated tips.</li>
</ol>
6. Conducting the durotaxis assay
<ol>
	<li>Before performing the experiment, practice engaging the gel near a cell and observe the deformation of the cell when the micromanipulator is repositioned.</li>
	<li>Monitor a group of cells that have clear polarity and appear to be moving for 30 min to identify cells that are moving in a directed manner.</li>
	<li>Choose a cell that is moving in a single, clear direction and monitor it at the desired frame rate for an additional 30 min.</li>
	<li>If determination of forces exerted on the cell or tension exerted by the cell is desired, capture bead field images at each acquisition. If the cell changes its course of direction during monitoring, choose a different cell to monitor as this will make it difficult to determine the effect of stimulation.</li>
	<li>Engage the hydrogel approximately 50 &#956;m away from the cell. Position the pipet in front of the near side of the leading edge and move the micromanipulator such that the gel is deformed orthogonally to the cell&#8217;s direction of travel. Observe the cell over time as it responds to the acute, local gradient of stiffness. 
	NOTE: The timing provided here is effective when monitoring SKOV3 or Ref52 fibroblasts, however, the interval and overall time course should be adjusted to suit the cell type and biological event being observed. If pairing with fluorescence microscopy, pause fluorescent acquisition immediately before step 6.5., use phase contrast or brightfield to position micropipette and pull, and restart fluorescent acquisition immediately after.</li>
	<li>If the pipet slips or if the gradient is otherwise relaxed or released, find a new cell by repeating steps 6.2 and 6.3.</li>
</ol>
7. Determining durotactic migration response
<ol>
	<li>Using ImageJ19 or another image analysis program, calculate the turn angle by drawing a line between the middle of the leading edge of the cell at 0 min and 30 min post monitor (reflecting the cell&#8217;s original trajectory) and another line between the middle of the leading edge just before and 80 min after stimulation and measuring the angle between these two lines.</li>
</ol>

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The authors have nothing to disclose.

Demonstrated here is a repeatable, single-cell durotaxis assay that allows assessment of a cell&#8217;s ability to alter its migration behavior in response to acute mechanical cues. This technique can also be used in combination with fluorescence microscopy and appropriate fusion proteins or biosensors to examine subcellular signaling and cytoskeletal events within seconds of mechanical stimulation or over a longer timescale during durotactic movement. Understanding a cell&#8217;s relationship to its environment involves...

Over the past few decades, the importance of the mechanical properties of a cell&#8217;s environment has garnered increasing recognition in cell biology. Different tissues and extracellular matrices have different relative stiffnesses and, as cells migrate throughout the body, they navigate these changes, using these mechanical properties to guide them1,2,3,4,5,6,7. Cells use the stiffness of a given tissue to inform their motile behavior during processes such as development, wound healing, and cancer metastasis. However, the molecular mechanisms that allow sensation of and response to these mechanical inputs remain largely unknown1,2,3,4,5,6,7.
In order to study the mechanisms through which cells respond to physical environmental cues, the rigidity or stiffness of the substrate underlying adherent cells must be manipulated. In 2000, Chun-Min Lo, Yu-Li Wang and colleagues developed an assay8 whereby an individual cell&#8217;s motile response to changing mechanical cues could be directly tested by stretching deformable extracellular matrix (ECM)-coated polyacrylamide hydrogels on which the cells were plated. Cells exhibits a significant preference for migrating towards stiffer substrates, a phenomenon they dubbed &#8220;durotaxis.&#8221;
Since the original report in 2000, many other techniques have been employed for the study of durotaxis. Steep stiffness gradients have been fabricated by casting gels over rigid features such as polystyrene beads9 or stiff polymer posts10 or by polymerizing the substrate around the edges of a glass coverslips11 to create mechanical &#8216;step-boundaries&#8217;. Alternatively, hydrogels with shallower but fixed stiffness gradients have been fabricated by a variety of methods such as gradients of crosslinker created by microfluidic devices12,13 or side-by-side hydrogel solution droplets of differing stiffness8, or hydrogels with photoreactive crosslinker treated with graded UV light exposure to create a linear stiffness gradient14,15. These techniques have been used to great effect to investigate durotactic cellular movement en masse over time. However, typically these features are fabricated in advance of cell plating and their properties remain consistent over the course of the experiment, relying on random cell movement for sampling of mechanical gradients. None of these techniques are amenable to observation of rapid changes in cellular behavior in response to acute mechanical stimulus.
In order to observe cellular responses to acute changes in the mechanical environment, single cell durotaxis assays offer several advantages. In these assays, individual cells are given an acute, mechanical stimulus by pulling the underlying substrate away from the cell with a glass micropipette, thereby introducing a directional gradient of cell-matrix tension. Changes in the motile behavior, such as speed or direction of migration, are then observed by live-cell phase contrast microscopy. This approach facilitates direct observation of cause and effect relationships between mechanical stimuli and cell migration, as it allows rapid, iterative manipulation of the direction and magnitude of the tension gradient and assessment of consequent cellular responses in real time. Further, this method can also be used to mechanically stimulate cells expressing fluorescent fusion proteins or biosensors to visualize changes in the amount, activity, or subcellular localization of proteins suspected to be involved in mechanosensing and durotaxis.
This technique has been employed by groups who study durotaxis8,16 and is described here as it has been adapted by the Howe Laboratory to study the durotactic behavior of SKOV-3 ovarian cancer cells and the molecular mechanisms that underly durotaxis17. Additionally, a modified method is described for fabrication of hydrogels with a single, even layer of fluorescent microspheres near the cell culture surface; this facilitates visualization and optimization of micropipette-generated strain gradients and may allow assessment of cell contractility by traction force microscopy.

<table>
	<tbody>
		<tr>
			<td>Acrylamide 40 %&#160;</td>
			<td>National Diagnostic</td>
			<td>EC-810</td>
			<td></td>
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		<tr>
			<td>Ammonium Persulfate&#160;</td>
			<td>Fisher</td>
			<td>BP179-25</td>
			<td></td>
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		<tr>
			<td>BD20A High frequency generator</td>
			<td>Electro Technic Products</td>
			<td>12011A</td>
			<td>115 V -&#160;Handheld Corona Wand</td>
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		<tr>
			<td>Bind Silane (y-methacryloxypropyltrimethoxysilane) (</td>
			<td>Sigma Aldrich</td>
			<td>M6514</td>
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		<tr>
			<td>Bis-acrylamide 2%&#160;</td>
			<td>National Diagnostic</td>
			<td>EC-820</td>
			<td></td>
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		<tr>
			<td>Borosilicate glass capillaries</td>
			<td>World Precision Instruments</td>
			<td>1B100-4</td>
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		<tr>
			<td>Branson 2510 Ultrasonic Cleaner</td>
			<td>Bransonic</td>
			<td></td>
			<td>40 kHz frequency</td>
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		<tr>
			<td>Coarse Manipulator</td>
			<td>Narshige</td>
			<td>MC35A</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>DMEM without phenol red</td>
			<td>Sigma Aldrich</td>
			<td>D5030</td>
			<td></td>
		</tr>
		<tr>
			<td>Dual-Stage Glass Micropipette Puller</td>
			<td>Narshige</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Epidermal Growth Factor</td>
			<td>Peprotech</td>
			<td>AF-100-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Pharmco-aaper</td>
			<td>111000200</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (Qualified One Shot)</td>
			<td>Gibco</td>
			<td>A31606-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin&#160;</td>
			<td>EMD Millipore</td>
			<td>FC010</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluospheres Carboxylate 0.2 um&#160;</td>
			<td>Invitrogen</td>
			<td>F8810, F8807, F8811</td>
			<td></td>
		</tr>
		<tr>
			<td>Fugene 6</td>
			<td>Roche</td>
			<td>1815091</td>
			<td>1.5 ug DNA / 6uL fugene 6 per 35mm dish</td>
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		<tr>
			<td>Glacial Acetic Acid</td>
			<td>Fisher Chemical</td>
			<td>A38SI-212</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Bottom Dish</td>
			<td>CellVis</td>
			<td>D60-60-1.5-N</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Coverslip</td>
			<td>Electron Microscopy Sciences</td>
			<td>72224-01</td>
			<td>22 mm, #1.5</td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>JT Baker</td>
			<td>9535-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Hellmanex III&#160;Special cleaning concentrate</td>
			<td>Sigma Aldrich</td>
			<td>Z805939</td>
			<td>Used at 2% in ddH2O for cleaning coverslips</td>
		</tr>
		<tr>
			<td>HEPES powder</td>
			<td>Sigma Aldrich</td>
			<td>H3375</td>
			<td>Make 50mM HEPES buffer, pH 8.5</td>
		</tr>
		<tr>
			<td>Intelli-Ray 400 Shuttered UV Flood Light</td>
			<td>Uviton International</td>
			<td>UV0338</td>
			<td></td>
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		<tr>
			<td>Isopropanol</td>
			<td>Fisher Chemical</td>
			<td>A417-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Microforge</td>
			<td>Narshige</td>
			<td>MF900</td>
			<td></td>
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			<td>Micromanipulator</td>
			<td>Narshige</td>
			<td>MHW3</td>
			<td></td>
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			<td>Mineral Oil</td>
			<td>Sigma Aldrich</td>
			<td>M5904</td>
			<td></td>
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			<td>Nanopure Life Science UV/UF System</td>
			<td>Barnstead</td>
			<td>D11931</td>
			<td>ddH2O</td>
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		<tr>
			<td>Nikon Eclipse Ti</td>
			<td>Nikon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>OptiMEM</td>
			<td>Invitrogen</td>
			<td>31985062</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm M</td>
			<td>Bemis Company, Inc</td>
			<td>PM-992</td>
			<td></td>
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		<tr>
			<td>PBS</td>
			<td></td>
			<td></td>
			<td>139 mM NaCl, 2.5 mM KCl, 28.6 mM Na2HPO4, 1.6 mM KH2PO4, pH 7.4</td>
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		<tr>
			<td>Platelet Derived Growth Factor-BB (PDGF-BB)</td>
			<td>Sigma Aldrich</td>
			<td>P4056</td>
			<td></td>
		</tr>
		<tr>
			<td>Ref52</td>
			<td></td>
			<td></td>
			<td>Rat embryonic fibroblast cell line; Culture in DMEM + 10% FBS</td>
		</tr>
		<tr>
			<td>Ringer&#39;s Buffer</td>
			<td></td>
			<td></td>
			<td>134 mM NaCl, 5.4 mM KCl, 1 mM MgSO4, 2.4 mM CaCl2, 20 mM HEPES, 5 mM D-Glucose, pH 7.4</td>
		</tr>
		<tr>
			<td>SKOV-3</td>
			<td>American Type Culture Collection</td>
			<td></td>
			<td>Culture in DMEM + 10% FBS</td>
		</tr>
		<tr>
			<td>Sulfo-SANPAH&#160;</td>
			<td>Covachem</td>
			<td>&#160;12414-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Tabletop Plasma Cleaner</td>
			<td>Harrick Plasma</td>
			<td>PDC-32G</td>
			<td></td>
		</tr>
		<tr>
			<td>TEMED&#160;</td>
			<td>Sigma Aldrich</td>
			<td>T9281-50</td>
			<td></td>
		</tr>
	</tbody>
</table>

single cell durotaxis assay for assessing mechanical control of cellular movement and related signaling events

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate underlying a cell must be manipulated. While hydrogels with graded stiffness and long-term migration assays have proven useful in durotaxis studies, immediate, acute responses to local changes in substrate tension allow focused study of individual cell movements and subcellular signaling events. To repeatably test the ability of cells to sense and respond to the underlying substrate stiffness, a modified method for application of acute gradients of increased tension to individual cells cultured on deformable hydrogels is used which allows for real time manipulation of the strength and direction of stiffness gradients imparted upon cells in question. Additionally, by fine tuning the details and parameters of the assay, such as the shape and dimensions of the micropipette or the relative position, placement, and direction of the applied gradient, the assay can be optimized for the study of any mechanically sensitive cell type and system. These parameters can be altered to reliably change the applied stimulus and expand the functionality and versatility of the assay. This method allows examination of both long term durotactic movement as well as more immediate changes in cellular signaling and morphological dynamics in response to changing stiffness.

By preparing micropipettes (Figure 1) and normalizing the force generation of the pulls (Figure 2 and Figure 3) as described above, optimal durotactic conditions have been identified for multiple cell lines. Using this technique, as outlined in Figure 4, both SKOV-3 ovarian cancer cells17&#160;and Ref52 rat embryonic fibroblasts (Figure 5) move towar...

Watch this Scientific Journal Video about Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events at JoVE.com

Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events

Mechanical forces are important for controlling cell migration. This protocol demonstrates the use of elastic hydrogels that can be deformed using a glass micropipette and a micromanipulator to stimulate cells with a local stiffness gradient to elicit changes in cell structure and migration.

Durotaxis is the process by which cells sense and respond to gradients of tension. In order to study this process in vitro, the stiffness of the substrate ...

single-cell-durotaxis-assay-for-assessing-mechanical-control-cellular

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

7.9K Views.  University of Vermont Larner College of Medicine. Mechanical forces are important for controlling cell migration. This protocol demonstrates the use of elastic hydrogels that can be deformed using a glass micropipette and a micromanipulator to stimulate cells with a local stiffness gradient to elicit changes in cell structure and migration.

Video: Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events

Imaging G-protein Coupled Receptor (GPCR)-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1. This guided cell migration is referred as chemotaxis, which is essential for various cells to carry out their functions such as trafficking of immune cells and patterning of neuronal cells 2, 3. A large family of G-protein coupled receptors (GPCRs) detects variable small peptides, known as chemokines, to direct cell migration in vivo 4. The final goal of chemotaxis research is to understand how a GPCR machinery senses chemokine gradients and controls signaling events leading to chemotaxis. To this end, we use imaging techniques to monitor, in real time, spatiotemporal concentrations of chemoattractants, cell movement in a gradient of chemoattractant, GPCR mediated activation of heterotrimeric G-protein, and intracellular signaling events involved in chemotaxis of eukaryotic cells 5-8. The simple eukaryotic organism, Dictyostelium discoideum, displays chemotaxic behaviors that are similar to those of leukocytes, and D. discoideum is a key model system for studying eukaryotic chemotaxis. As free-living amoebae, D. discoideum cells divide in rich medium. Upon starvation, cells enter a developmental program in which they aggregate through cAMP-mediated chemotaxis to form multicullular structures. Many components involved in chemotaxis to cAMP have been identified in D. discoideum. The binding of cAMP to a GPCR (cAR1) induces dissociation of heterotrimeric G-proteins into G&gamma; and G&beta;&gamma; subunits 7, 9, 10. G&beta;&gamma; subunits activate Ras, which in turn activates PI3K, converting PIP2 into PIP3 on the cell membrane 11-13. PIP3 serve as binding sites for proteins with pleckstrin Homology (PH) domains, thus recruiting these proteins to the membrane 14, 15. Activation of cAR1 receptors also controls the membrane associations of PTEN, which dephosphorylates PIP3 to PIP2 16, 17. The molecular mechanisms are evolutionarily conserved in chemokine GPCR-mediated chemotaxis of human cells such as neutrophils 18. We present following methods for studying chemotaxis of D. discoideum cells. 1. Preparation of chemotactic component cells. 2. Imaging chemotaxis of cells in a cAMP gradient. 3. Monitoring a GPCR induced activation of heterotrimeric G-protein in single live cells. 4. Imaging chemoattractant-triggered dynamic PIP3 responses in single live cells in real time. Our developed imaging methods can be applied to study chemotaxis of human leukocytes.

Imaging G-protein Coupled Receptor GPCR-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

Here, we describe detailed live cell imaging methods for investigating chemotaxis. We present fluorescence microscopic methods to monitor spatiotemporal dynamics of signaling events in migrating cells. Measurement of signaling events permits us to further understand how a GPCR-signaling network achieves gradient sensing of chemoattractants and controls directional migration of eukaryotic cells.

Many eukaryotic cells can detect gradients of chemical signals in their environments and migrate accordingly 1.  This guided cell migration is referred as ...

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

F&ouml;rster resonance energy transfer (FRET) microscopy continues to gain increasing interest as a technique for real-time monitoring of biochemical and signaling events in live cells and tissues. Compared to classical biochemical methods, this novel technology is characterized by high temporal and spatial resolution. FRET experiments use various genetically-encoded biosensors which can be expressed and imaged over time in situ or in vivo1-2. Typical biosensors can either report protein-protein interactions by measuring FRET between a fluorophore-tagged pair of proteins or conformational changes in a single protein which harbors donor and acceptor fluorophores interconnected with a binding moiety for a molecule of interest3-4. Bimolecular biosensors for protein-protein interactions include, for example, constructs designed to monitor G-protein activation in cells5, while the unimolecular sensors measuring conformational changes are widely used to image second messengers such as calcium6, cAMP7-8, inositol phosphates9 and cGMP10-11. Here we describe how to build a customized epifluorescence FRET imaging system from single commercially available components and how to control the whole setup using the Micro-Manager freeware. This simple but powerful instrument is designed for routine or more sophisticated FRET measurements in live cells. Acquired images are processed using self-written plug-ins to visualize changes in FRET ratio in real-time during any experiments before being stored in a graphics format compatible with the build-in ImageJ freeware used for subsequent data analysis. This low-cost system is characterized by high flexibility and can be successfully used to monitor various biochemical events and signaling molecules by a plethora of available FRET biosensors in live cells and tissues. As an example, we demonstrate how to use this imaging system to perform real-time monitoring of cAMP in live 293A cells upon stimulation with a &beta;-adrenergic receptor agonist and blocker.

F&#246;rster resonance energy transfer (FRET) microscopy is a powerful technique for real-time monitoring of signaling events in live cells using various biosensors as reporters. Here we describe how to build a customized epifluorescence FRET imaging system from commercially available components and how to use it for FRET experiments.

F&ouml;rster resonance energy transfer (FRET) microscopy  continues to gain increasing interest as a technique for real-time monitoring  of biochemical ...

An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the neural tube in a process known as neurulation. During neurulation, morphogenesis of the mesenchyme that underlies the neural plate is believed to drive neural fold elevation. The cranial mesenchyme is comprised of the paraxial mesoderm and neural crest cells. The cells of the cranial mesenchyme form a pourous meshwork composed of stellate shaped cells and intermingling extracellular matrix (ECM) strands that support the neural folds. During neurulation, the cranial mesenchyme undergoes stereotypical rearrangements resulting in its expansion and these movements are believed to provide a driving force for neural fold elevation. However, the pathways and cellular behaviors that drive cranial mesenchyme morphogenesis remain poorly studied. Interactions between the ECM and the cells of the cranial mesenchyme underly these cell behaviors. Here we describe a simple ex vivo explant assay devised to characterize the behaviors of these cells. This assay is amendable to pharmacological manipulations to dissect the signaling pathways involved and live imaging analyses to further characterize the behavior of these cells. We present a representative experiment demonstrating the utility of this assay in characterizing the migratory properties of the cranial mesenchyme on a variety of ECM components.

The cranial mesenchyme undergoes dramatic morphogenic movements that likely provides a driving force for elevation of the neural folds1,2. Here we describe a simple ex vivo explant assay to characterize the cellular behaviors of the cranial mesenchyme during neurulation. This assay has numerous applications including being amenable to pharmacological manipulations and live imaging analyses.

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the ...

Mechanical Stimulation-induced Calcium Wave Propagation in Cell Monolayers: The Example of Bovine Corneal Endothelial Cells

Intercellular communication is essential for the coordination of physiological processes between cells in a variety of organs and tissues, including the brain, liver, retina, cochlea and vasculature. In experimental settings, intercellular Ca2+-waves can be elicited by applying a mechanical stimulus to a single cell. This leads to the release of the intracellular signaling molecules IP3 and Ca2+ that initiate the propagation of the Ca2+-wave concentrically from the mechanically stimulated cell to the neighboring cells. The main molecular pathways that control intercellular Ca2+-wave propagation are provided by gap junction channels through the direct transfer of IP3 and by hemichannels through the release of ATP. Identification and characterization of the properties and regulation of different connexin and pannexin isoforms as gap junction channels and hemichannels are allowed by the quantification of the spread of the intercellular Ca2+-wave, siRNA, and the use of inhibitors of gap junction channels and hemichannels. Here, we describe a method to measure intercellular Ca2+-wave in monolayers of primary corneal endothelial cells loaded with Fluo4-AM in response to a controlled and localized mechanical stimulus provoked by an acute, short-lasting deformation of the cell as a result of touching the cell membrane with a micromanipulator-controlled glass micropipette with a tip diameter of less than 1 &mu;m. We also describe the isolation of primary bovine corneal endothelial cells and its use as model system to assess Cx43-hemichannel activity as the driven force for intercellular Ca2+-waves through the release of ATP. Finally, we discuss the use, advantages, limitations and alternatives of this method in the context of gap junction channel and hemichannel research.

Intercellular Ca2+-waves are driven by gap junction channels and hemichannels. Here, we describe a method to measure intercellular Ca2+-waves in cell monolayers in response to a local single-cell mechanical stimulus and its application to investigate the properties and regulation of gap junction channels and hemichannels.

Intercellular communication is essential for the coordination of physiological processes between cells in a variety of organs and tissues, including the ...

Single-cell Microinjection for Cell Communication Analysis

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a hemichannel by each neighboring cell to form the gap junction. In mammalian cells, the hemichannel is formed by six connexins, monomers with four transmembrane domains and a C and N terminal within the cytoplasm. Gap junctions permit the exchange of ions, second messengers, and small metabolites. In addition, they have important roles in many forms of cellular communication within physiological processes such as synaptic transmission, heart contraction, cell growth and differentiation. We detail how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells. It is expected that in functional gap junctions, the dye will diffuse from the loaded cell to the connected cells. It is a very useful technique to study gap junctions since you can evaluate the diffusion of the fluorescence in real time. We discuss how to prepare the cells and the micropipette, how to use a micromanipulator and inject a low molecular weight fluorescent dye in an epithelial cell line.

We describe here how to perform a single-cell microinjection of Lucifer Yellow to visualize cellular communication via gap-junctions in living cells, and provide some useful tips. We expect that this paper will help everyone to evaluate the degree of cellular coupling due to functional gap junctions. Everything described here could be, in principle, adapted to other fluorescent dyes with molecular weight below 1,000 Daltons.

Gap junctions are intercellular channels that allow the communication of neighboring cells. This communication depends on the contribution of a ...

A Simplified System for Evaluating Cell Mechanosensing and Durotaxis In Vitro

The composition and mechanical properties of the extracellular matrix are highly variable between tissue types. This connective tissue stroma diversity greatly impacts cell behavior to regulate normal and pathologic processes including cell proliferation, differentiation, adhesion signaling and directional migration. In this regard, the innate ability of certain cell types to migrate towards a stiffer, or less compliant matrix substrate is referred to as durotaxis. This phenomenon plays an important role during embryonic development, wound repair and cancer cell invasion. Here, we describe a straightforward assay to study durotaxis, in vitro, using polydimethylsiloxane (PDMS) substrates. Preparation of the described durotaxis chambers creates a rigidity interface between the relatively soft PDMS gel and a rigid glass coverslip. In the example provided, we have used these durotaxis chambers to demonstrate a role for the cdc42/Rac1 GTPase activating protein, cdGAP, in mechanosensing and durotaxis regulation in human U2OS osteosarcoma cells. This assay is readily adaptable to other cell types and/or knockdown of other proteins of interests to explore their respective roles in mechanosignaling and durotaxis.

A Simplified System for Evaluating Cell Mechanosensing and Durotaxis In Vitro

Many mammalian cells preferentially migrate towards a more rigid matrix or substrate through durotaxis. The goal of this protocol is to provide a simple in vitro system that can be used to study and manipulate cell durotaxis behaviors by incorporating polydimethylsiloxane (PDMS) substrates of defined rigidity, interfacing with glass coverslips.

The composition and mechanical properties of the extracellular matrix are highly variable between tissue types. This connective tissue stroma diversity ...

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and development. Cytokinesis involves a series of events that are well coordinated in time and space. Cytokinesis involves the formation of an actomyosin ring at the division site, followed by ring constriction, membrane furrow formation and extra cellular matrix remodeling. The fission yeast, Schizosaccharomyces pombe (S.&#160;pombe)&#160;is a well-studied model system that has revealed with substantial clarity the initial events in cytokinesis. However, we do not understand clearly how different cytokinetic events are coordinated spatiotemporally. To determine this, one needs to analyze the different cytokinetic events in great details in both time and in space. Here we describe a microscopy approach to examine different cytokinetic events in live cells. With this approach it is possible to time different cytokinetic events and determine the time of recruitment of different proteins during cytokinesis. In addition, we describe protocols to compare protein localization, and distribution at the site of cell division. This is a basic protocol to study cytokinesis in fission yeast and can also be used for other yeasts and fungal systems.

The fission yeast, Schizosaccharomyces pombe&#160;is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and ...

Single-cell Microfluidic Analysis of Bacillus subtilis

Microfluidic technology overcomes many of the limitations to traditional analytical methods in microbiology. Unlike bulk-culture methods, it offers single-cell resolution and long observation times spanning hundreds of generations; unlike agarose pad-based microscopy, it has uniform growth conditions that can be tightly controlled. Because the continuous flow of growth medium isolates the cells in a microfluidic device from unpredictable variations in the local chemical environment caused by cell growth and metabolism, authentic changes in gene expression and cell growth in response to specific stimuli can be more confidently observed. Bacillus subtilis is used here as a model bacterial species to demonstrate a &#34;mother machine&#34;-type method for cellular analysis. We show how to construct and plumb a microfluidic device, load it with cells, initiate microscopic imaging, and expose cells to a stimulus by switching from one growth medium to another. A stress-responsive reporter is used as an example to reveal the type of data that may be obtained by this method. We also briefly discuss further applications of this method for other types of experiments, such as analysis of bacterial sporulation.

Single-cell Microfluidic Analysis of Bacillus subtilis

We present a method for the microfluidic analysis of individual bacterial cell lineages using Bacillus subtilis as an example. The method overcomes shortcomings of traditional analytical methods in microbiology by allowing observation of hundreds of cell generations under tightly controllable and uniform growth conditions.

Microfluidic technology overcomes many of the limitations to traditional analytical methods in microbiology. Unlike bulk-culture methods, it offers ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

Collection of Skeletal Muscle Biopsies from the Superior Compartment of Human Musculus Tibialis Anterior for Mechanical Evaluation

The mechanical properties of contracting skeletal fibers are crucial indicators of overall muscle health, function, and performance. Human skeletal muscle biopsies are often collected for these endeavors. However, relatively few technical descriptions of biopsy procedures, outside of the commonly used musculus vastus lateralis, are available. Although the biopsy techniques are often adjusted to accommodate the characteristics of each muscle under study, few technical reports share these changes to the greater community. Thus, muscle tissue from human participants is often wasted as the operator reinvents the wheel. Expanding the available material on biopsies from a variety of muscles can reduce the incident of failed biopsies. This technical report describes a variation of the modified Bergstr&#246;m technique on the musculus tibialis anterior that limits fiber damage and provides fiber lengths adequate for mechanical evaluation. The surgery is an outpatient procedure that can be completed in an hour. The recovery period for this procedure is immediate for light activity (i.e., walking), up to three days for the resumption of normal physical activity, and about one week for wound care. The extracted tissue can be used for mechanical force experiments and here we present representative activation data. This protocol is appropriate for most collection purposes, potentially adaptable to other skeletal muscles, and may be improved by modifications to the collection needle.

This technical report describes a variation of the modified Bergstr&#246;m technique for the biopsy of the musculus tibialis anterior that limits fiber damage.

The mechanical properties of contracting skeletal fibers are crucial indicators of overall muscle health, function, and performance. Human skeletal muscle ...

Single Cell Durotaxis Assay for Assessing Mechanical Control of Cellular Movement and Related Signaling Events

Summary

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Chapters in this video

Imaging G-protein Coupled Receptor (GPCR)-mediated Signaling Events that Control Chemotaxis of Dictyostelium Discoideum

FRET Microscopy for Real-time Monitoring of Signaling Events in Live Cells Using Unimolecular Biosensors

An Explant Assay for Assessing Cellular Behavior of the Cranial Mesenchyme

Mechanical Stimulation-induced Calcium Wave Propagation in Cell Monolayers: The Example of Bovine Corneal Endothelial Cells

Single-cell Microinjection for Cell Communication Analysis

A Simplified System for Evaluating Cell Mechanosensing and Durotaxis In Vitro

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Single-cell Microfluidic Analysis of Bacillus subtilis

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Collection of Skeletal Muscle Biopsies from the Superior Compartment of Human Musculus Tibialis Anterior for Mechanical Evaluation