We thank Margaret Gardel for generously allowing the use of the rheometer. We acknowledge support from the NIH (1R15GM116082) that enabled this work.

1. Fabrication of Soft Silicone Substrate
<ol>
	<li>Weigh out 1.75 g of the A component and 1.75 g of the B component (A:B = 1:1) from the soft silicone elastomer kit using (polystyrene) weighing trays.</li>
	<li>Add the A component to the B component in the weighing tray and mix them together for 5 min using an appropriate applicator stick.</li>
	<li>Add the above mixture to a 35 mm Petri dish. Allow the mixture to spread evenly across the Petri dish for a couple of minutes. 
	Note: The choice of the Petri dish diameter and the amount of soft silicone will determine the soft silicone thickness. Here, the thickness will be around 3.5 mm; more about choosing the elastomer thickness in the Discussion section.</li>
	<li>Place the Petri dish with the silicone mixture, with the lid off, in a vacuum chamber for 15 min to remove any air bubbles. During this time, pre-heat a hot plate to 70 &#176;C.</li>
	<li>Once the hot plate reaches 70 &#176;C, place a glass slide on it and then place the Petri dish with the silicone mixture on the glass slide. Let the silicone cure at 70 &#176;C for 30 min. Do not place the polystyrene dish directly on the hot plate, as the Petri dish may melt.</li>
</ol>
2. Coupling of Fluorescent Microbeads to the Soft Silicone
<ol>
	<li>Place the cured soft silicone (in the uncovered Petri dish) in a deep UV chamber (an enclosure with a deep UV lamp of light wavelengths of 185 and 254 nm). Expose the soft silicone sample (~5 - 10 cm away from the UV lamp) to deep UV light for 5 min.
	<ol>
		<li>While the silicone is being exposed to deep UV light, proceed with steps 2.2 - 2.6 below. After the deep UV exposure, degas the deep UV chamber for at least 5 min before retrieving the sample.</li>
	</ol>
	</li>
	<li>In the meantime, weigh out 19 mg of 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) in a 1.5 mL microcentrifuge tube and add 500 &#956;L of deionized water (DI) water to it. Dissolve the EDC by gently shaking the tube.</li>
	<li>In a separate 1.5 mL microcentrifuge tube, weigh out 11 mg of N-hydroxysulfosuccinimide (sulfo-NHS), add 500 &#956;L of DI water to it, and dissolve the sulfo-NHS by gently shaking the tube. Then, combine the EDC and sulfo-NHS solutions in a single microcentrifuge tube.</li>
	<li>To this EDC/sulfo-NHS solution, add 30 &#956;L of 0.44 &#956;m diameter red (or any other fluorescence color based on the filter cubes available in the fluorescence microscope) carboxylate modified fluorescent microbeads (with a 1% w/v stock concentration).</li>
	<li>To the EDC/sulfo-NHS/bead mixture, add 0.02 mg of collagen I (from a rat tail, stock concentration of 4 mg/mL in 0.02 M acetic acid) to obtain a concentration of about 0.02 mg/mL.</li>
	<li>Vortex the EDC/NHS/bead/collagen I mixture briefly to ensure that the beads are evenly dispersed throughout, before coupling.</li>
	<li>Pipette 1 mL of the EDC/NHS/bead/collagen I mixture on a piece of parafilm placed on top of another shallow, flat lid (of smaller diameter). Invert the Petri dish with soft silicone on this mixture so that the soft silicone surface contacts the mixture but does not directly touch the surface of the smaller Petri dish lid below. To raise the inverted Petri dish, use one or two glass slides under either side of the inverted Petri dish spacers. 
	Note: Please refer to Figure 1 to see how step 2.7 is performed.</li>
	<li>Cover the sample with aluminum foil and incubate it at room temperature for 30 min.</li>
	<li>Remove the Petri dish with soft silicone and set it upright (silicone-side up).</li>
	<li>Wash the soft silicone surface with phosphate-buffered saline (PBS) by adding 2 mL of PBS (pH 7.4) to the dish. Let it sit for a couple of minutes. Aspirate off the PBS and wash the silicone again with 2 mL of PBS. Let the silicone cure further for about a day. For that reason, place the soft silicone sample in PBS at 37 &#176;C overnight.</li>
</ol>
3. Measurement of Silicone Stiffness with Sphere Indentation using a Widefield Fluorescence Microscope
<ol>
	<li>Retrieve the Petri dish with soft silicone and ensure that it contains at least 1 mL of PBS to have the silicone surface several mm below the liquid surface.</li>
	<li>Using pointed tweezers, drop five 1 mm zirconium sphere indentors on the soft silicone. Immerse the spheres into the liquid medium and drop them, away from the edges of the silicone layer and at least 5 indentor diameters away from the location of the other indentors. 
	Note: When dropped above the liquid surface, the spheres may fail to enter the liquid medium (float) due to the liquid medium&#8217;s surface tension.</li>
	<li>Place the Petri dish with soft silicone on the microscope stage so that it is possible to image through the Petri dish base.</li>
	<li>Using phase imaging with a 10X objective (such as a dry 10X objective of NA 0.30), locate and bring a sphere indentor into focus.</li>
	<li>Take a phase image of a part or the whole of the indentor and save this image. Use a tile scan if available. If the indentor has any visible defects, discard and replace it with another indentor.</li>
	<li>Under live phase imaging, pan to the left of the indentor&#8217;s edge so that the left edge of the frame is at least a distance of ~1.5 R from the indentor center. Ensure that the center of the indentor remains visible on the right side, close to the right edge of the image frame. Take a phase image and save it.</li>
	<li>Switch the microscope light source to the illumination for the red fluorescent channel. With the x- and y-coordinates unchanged (the x-y position of the indentor center within but near the right edge of the frame), focus down (decrease Z) until the red fluorescent microbeads under the sphere indentor&#8217;s center just go out of focus.</li>
	<li>Take a z-stack with an image for every z-increment of 0.5 &#181;m till the microbeads in the top layer of the silicone far from the indentor (near the left edge of the imaging frame) go out of focus.</li>
	<li>Repeat steps 3.4 - 3.8 with the other indentors on the sample.</li>
</ol>
4. Calculating the Silicone&#8217;s Stiffness (Young&#8217;s Modulus)
<ol>
	<li>Open the phase image of the indentor using ImageJ, click on the line tool, and measure the indentor&#8217;s diameter in pixels. Click and hold on a point on the indentor edge, move the cursor to a diametrically opposite point on the edge and note the length in pixels displayed on the status bar of the ImageJ main window before releasing the cursor.
	<ol>
		<li>Ensure that the unit of length is set to pixels by clicking Analyze | Set Scale and checking the Unit of length.</li>
		<li>Convert the indentor&#8217;s radius in pixels to &#956;m by taking into account the objective magnification and the CCD camera pixel size (R in &#956;m = R in pixels x the CCD camera pixel size in &#956;m / the objective magnification).</li>
	</ol>
	</li>
	<li>Open the red channel z-stack of microbead images (if the microbeads are red fluorescent) in ImageJ by clicking on File | Import | Image Sequence and select any image in the stack and click OK to open the stack. 
	Note: F1 is the frame number at which the microbeads under the indentor center are in the best possible focus and F2 is the frame number at which the microbeads (at a region of ~1.5 R away from the bead center) near the left edge of the frame are in the best possible focus. The z-difference between the two frames is the indentation depth &#948;.
	<ol>
		<li>Using the line tool in ImageJ, draw a line across a well-defined microbead in the image. Click on Analyze | Plot Profile and click on the Live button to obtain the updated line scan intensity across the bead while selecting different frames. The frame that gives the highest value of the maximum intensity can be chosen as the frame in focus.</li>
		<li>Since the z-increment between the frames in the z-stack is 0.5 &#956;m, calculate the indentation depth in &#956;m as &#948; = (F2-F1) x 0.5.</li>
	</ol>
	</li>
	<li>Calculate the force exerted on the gel by the indentor due to its weight (minus the opposing buoyant force), that is, the indentation force F, as the volume of the indentor x (the density of the indentor - the density of the liquid medium) x the acceleration due to gravity. Use the equation F = (4/3) x 3.142 x (R3) x (&#961;indentor&#173; - &#961;medium) x g where R is the radius of the indentor, &#961;indentor is the density of the indentor, &#961;medium is the density of the liquid medium and g is the acceleration due to gravity (9.81 m/s2). Express all quantities on the right-hand side in SI units to obtain F (in N).</li>
	<li>Calculate the Young&#8217;s modulus (E) of the silicone using a modified11 Hertz model12 equation: 
	<img alt="Equation 1" src="/files/ftp_upload/57797/57797eq1.jpg" /> 
	Where: 
	c = a correction factor that modifies the Hertz model expression that follows it; 
	v = Poisson&#8217;s ratio of the silicone gel (taken as 0.5 as for incompressible materials7); 
	F = the indentation force; 
	R = the indentor radius; and 
	&#948; = the indentation depth. 
	Express all quantities on the right-hand side in SI units to obtain E in Pa.
	<ol>
		<li>Calculate the correction factor c as follows3: 
		<img alt="Equation 2" src="/files/ftp_upload/57797/57797eq2.jpg" /> 
		Where: 
		<img alt="Equation 3" src="/files/ftp_upload/57797/57797eq3.jpg" /> 
		<img alt="Equation 4" src="/files/ftp_upload/57797/57797eq4.jpg" />; and 
		<img alt="Equation 5" src="/files/ftp_upload/57797/57797eq5.jpg" />. 
		It should be noted that this correction factor is specifically to be used only when the soft silicone adheres well to the Petri dish (or stiff silicone) underneath it (which is the case here).</li>
		<li>Calculate the height h of the soft silicone layer based on the amount of silicone added and the Petri dish diameter. Alternatively, obtain h directly by determining the z-coordinate of the top and bottom surfaces of the silicone layer by phase imaging (minor impurities come into focus at either surface). Note that for a large h (h2 &#62; R&#948;), the correction factor c is close to 1.</li>
	</ol>
	</li>
	<li>Repeat steps 4.1 - 4.4 for each indentor. Average the Young&#8217;s modulus obtained from each indentor to obtain the mean Young&#8217;s modulus for the silicone sample.</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+movement+is+guided+by+the+rigidity+of+the+substrate.">Cell movement is guided by the rigidity of the substrate.</a> Biophysical Journal. 79, 144-152 (2000).</li><li>Discher, D. E., Janmey, P., Wang, Y. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue+cells+feel+and+respond+to+the+stiffness+of+their+Substrate.">Tissue cells feel and respond to the stiffness of their Substrate.</a> Science. 310, 1139-1143 (2005).</li><li>Kandow, C. E., Georges, P. C., Janmey, P. A., Beningo, K. 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The authors have nothing to disclose.

While the sphere indentation method is easy to implement, careful attention must be paid to the choice of indentor and the thickness of the soft silicone sample. The equation used to calculate the Young&#39;s modulus is valid under a set of conditions11and these are typically satisfied when the thickness of the silicone sample is &#62; 10% of the indentor radius and &#60; ~13x the indentor radius. We found that a silicone thickness of 5 - 10x the indentor radius was a good choice, wherein the samp...

Cells in soft tissues reside in a micro-environment whose stiffness is in the kilopascal range1, in contrast to tissue culture dishes whose stiffness is several orders of magnitude higher. Early experiments with cells on extracellular matrix protein-coated soft substrates showed that the substrate stiffness influences how cells move on as well as adhere to the extracellular matrix beneath2,3. In fact, the substrate stiffness fundamentally influences the cell function4&#160;in a manner similar to pervasive biochemical signals. Polyacrylamide gels (coated with extracellular matrix proteins) are (water-permeating) hydrogels that have been extensively used as cell culture substrates for mechanobiology studies5. Polydimethylsiloxane (PDMS), the most common silicone (polysiloxane), has been widely used as a stiff silicone with megapascal-range stiffness for micron-scale fabrication6. More recently, soft silicone substrates with stiffness in the more physiologically relevant kilopascal range have been employed as cell culture substrates for mechanobiology studies7,8.
Several methods have been used to measure the stiffness of flexible substrates, including atomic force microscopy, macroscopic deformation of whole samples upon stretching, rheology, and indentation using spheres and spherically tipped microindentors9. While each technique has its own advantages and disadvantages, indentation with a sphere is an especially simple yet fairly accurate method that only requires the access to a widefield fluorescence microscope. Indentation with a metallic sphere has been used to measure the stiffness of hydrogels in prior work3,9,10. Early work that demonstrated the importance of substrate stiffness to cell movement utilized this method to determine hydrogel substrate stiffness3. More recently, confocal microscopy has also been used for an elegant characterization10.
Here, we present a step-by-step protocol for preparing a soft silicone substrate, coupling fluorescent beads (and an extracellular matrix protein such as collagen I) just to the top surface, imaging an indenting sphere and the top surface using phase and fluorescence imaging, respectively, and finally analyzing the images to compute the Young&#39;s modulus of the silicone substrate. The soft silicone substrate prepared in this manner can be readily used for traction force microscopy experiments. The use of stiff silicone (instead of a Petri dish) as the base for the soft silicone also enables mechanobiology studies using an external stretch. Where warranted, practical considerations necessary for avoiding possible complications are also indicated.

<table><tbody><tr><td>CY 52-276 A/B silicone elastomer kit</td><td>&amp;nbsp;Dow Corning</td><td>CY 52-276</td><td>Store at room temperature</td></tr><tr><td>Thermo Scientific Pierce EDC</td><td>Fisher Scientific</td><td>PI22980</td><td>Store at -20&amp;deg;C</td></tr><tr><td>Thermo Scientific&amp;nbsp;Pierce Sulfo-NHS crosslinker</td><td>Fisher Scientific</td><td>PI-24510</td><td>Store at 4&amp;deg;C</td></tr><tr><td>Carboxyl fluorescent pink particles, 0.4-0.6 &amp;micro;m, 2 mL</td><td>Spherotech, Inc.</td><td>CFP-0558-2</td><td>Store at 4&amp;deg;C, do not freeze</td></tr><tr><td>1.0 mm Acid washed Zirconium beads</td><td>OPS Diagnostics LLC</td><td>BAWZ 1000-250-33</td><td></td></tr><tr><td>Deep UV chamber with ozone evacuator</td><td>Novascan Technologies, Inc.</td><td>PSD-UV4, OES-1000D</td><td></td></tr><tr><td>Wide field fluorescence microscope</td><td>Leica Microsystems</td><td>DMi8</td><td></td></tr><tr><td>Collagen I, from rat tail</td><td>Corning</td><td>354236</td><td>Stock concentration = 4 mg/ml; store at 4&amp;deg;C</td></tr><tr><td>ImageJ-NIH</td><td>N/A</td><td>N/A</td><td>public-domain software</td></tr></tbody></table>

stiffness measurement of soft silicone substrates for mechanobiology studies using a widefield fluorescence microscope

Soft tissues in the human body typically have stiffness in the kilopascal (kPa) range. Accordingly, silicone and hydrogel flexible substrates have been proven to be useful substrates for culturing cells in a physical microenvironment that partially mimics in vivo conditions. Here, we present a simple protocol for characterizing the Young's moduli of isotropic linear elastic substrates typically used for mechanobiology studies. The protocol consists of preparing a soft silicone substrate on a Petri dish or stiff silicone, coating the top surface of the silicone substrate with fluorescent beads, using a millimeter-scale sphere to indent the top surface (by gravity), imaging the fluorescent beads on the indented silicone surface using a fluorescence microscope, and analyzing the resultant images to calculate the Young's modulus of the silicone substrate. Coupling the substrate's top surface with a moduli extracellular matrix protein (in addition to the fluorescent beads) allows the silicone substrate to be readily used for cell plating and subsequent studies using traction force microscopy experiments. The use of stiff silicone, instead of a Petri dish, as the base of the soft silicone, enables the use of mechanobiology studies involving external stretch. A specific advantage of this protocol is that a widefield fluorescence microscope, which is commonly available in many labs, is the major equipment necessary for this procedure. We demonstrate this protocol by measuring the Young's modulus of soft silicone substrates of different elastic moduli.

Using the protocol detailed above, we prepared soft silicone in a 35 mm Petri dish, cured it at 70 &#176;C for 30 min and coupled fluorescent microbeads (and collagen I) to the top surface as schematically depicted in Figure 1. Deep UV has been used previously for the eventual protein coupling to substrates13. Note that (I) the curing conditions used here are specific to this soft silicone and (II) the indentation measurement is pe...

Watch this Scientific Journal Video about Stiffness Measurement of Soft Silicone Substrates for Mechanobiology Studies Using a Widefield Fluorescence Microscope at JoVE.com

Stiffness Measurement of Soft Silicone Substrates for Mechanobiology Studies Using a Widefield Fluorescence Microscope

Substrates with stiffness in the kilopascal-range are useful to study the response of cells to physiologically relevant micro-environment stiffness. Using just a widefield fluorescence microscope, the Young's modulus of soft silicone gels can be determined using an indentation with a suitable sphere.

Soft tissues in the human body typically have stiffness in the kilopascal (kPa) range. Accordingly, silicone and hydrogel flexible substrates have been ...

stiffness-measurement-soft-silicone-substrates-for-mechanobiology

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JoVE Journal

Bioengineering

6.8K Views.  Old Dominion University. Ondergronden met stijfheid in het kilopascal-bereik zijn nuttig zijn te onderzoeken van de reactie van cellen op fysiologisch relevante stijfheid van de micro-omgeving. Met behulp van slechts een widefield fluorescentie Microscoop, kan de Youngs modulus van zachte siliconen gel worden bepaald met behulp van een inkeping met een geschikt gebied.

Video: Stiffness Measurement of Soft Silicone Substrates for Mechanobiology Studies Using a Widefield Fluorescence Microscope

Micropipette Aspiration of Substrate-attached Cells to Estimate Cell Stiffness

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell proliferation, differentiation, migration and cell-cell interactions. The two key parameters of cellular biomechanics are cellular deformability or stiffness and the ability of the cells to contract and generate force. Here we describe a quick and simple method to estimate cell stiffness by measuring the degree of membrane deformation in response to negative pressure applied by a glass micropipette to the cell surface, a technique that is called Micropipette Aspiration or Microaspiration.
Microaspiration is performed by pulling a glass capillary to create a micropipette with a very small tip (2-50 &mu;m diameter depending on the size of a cell or a tissue sample), which is then connected to a pneumatic pressure transducer and brought to a close vicinity of a cell under a microscope. When the tip of the pipette touches a cell, a step of negative pressure is applied to the pipette by the pneumatic pressure transducer generating well-defined pressure on the cell membrane. In response to pressure, the membrane is aspirated into the pipette and progressive membrane deformation or "membrane projection" into the pipette is measured as a function of time. The basic principle of this experimental approach is that the degree of membrane deformation in response to a defined mechanical force is a function of membrane stiffness. The stiffer the membrane is, the slower the rate of membrane deformation and the shorter the steady-state aspiration length.The technique can be performed on isolated cells, both in suspension and substrate-attached, large organelles, and liposomes.
Analysis is performed by comparing maximal membrane deformations achieved under a given pressure for different cell populations or experimental conditions. A "stiffness coefficient" is estimated by plotting the aspirated length of membrane deformation as a function of the applied pressure. Furthermore, the data can be further analyzed to estimate the Young's modulus of the cells (E), the most common parameter to characterize stiffness of materials. It is important to note that plasma membranes of eukaryotic cells can be viewed as a bi-component system where membrane lipid bilayer is underlied by the sub-membrane cytoskeleton and that it is the cytoskeleton that constitutes the mechanical scaffold of the membrane and dominates the deformability of the cellular envelope. This approach, therefore, allows probing the biomechanical properties of the sub-membrane cytoskeleton.

Here we describe a quick and simple method to measure cell stiffness. The general principle of this approach is to measure membrane deformation in response to well-defined negative pressure applied through a micropipette to the cell surface. This method provides a powerful tool to study biomechanical properties of substrate-attached cells.

Growing number of studies show that biomechanical properties of individual cells play major roles in multiple cellular functions, including cell ...

Measuring the Mechanical Properties of Living Cells Using Atomic Force Microscopy

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

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

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

Simple Polyacrylamide-based Multiwell Stiffness Assay for the Study of Stiffness-dependent Cell Responses

Currently, most of the in vitro cell research is performed on rigid tissue culture polystyrene (~1 GPa), while most cells in the body are attached to a matrix that is elastic and much softer (0.1 &#8211; 100 kPa). Since such stiffness mismatch greatly affects cell responses, there is a strong interest in developing hydrogel materials that span a wide range of stiffness to serve as cell substrates. Polyacrylamide gels, which are inexpensive and cover the stiffness range of all soft tissues in the body, are the hydrogel of choice for many research groups. However, polyacrylamide gel preparation is lengthy, tedious, and only suitable for small batches. Here, we describe an assay which by utilizing a permanent flexible plastic film as a structural support for the gels, enables the preparation of polyacrylamide gels in a multiwell plate format. The technique is faster, more efficient, and less costly than current methods and permits the preparation of gels of custom sizes not otherwise available. As it doesn&#8217;t require any specialized equipment, the method could be easily adopted by any research laboratory and would be particularly useful in research focused on understanding stiffness-dependent cell responses.

Here, a method that enables quick, efficient, and inexpensive preparation of polyacrylamide gels in a multiwell plate format is described. The method does not require any specialized equipment and could be easily adopted by any research laboratory. It would be particularly useful in research focused on understanding stiffness-dependent cell responses.

Currently, most of the in vitro cell research is performed on rigid tissue culture polystyrene (~1 GPa), while most cells in the body are attached to a ...

Live Cell Imaging during Mechanical Stretch

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in their capability for measuring responses in real time in live cells or viable tissue. A protocol was developed with the use of a cell actuator to distend live cells grown on or tissues attached to an elastic substrate while imaging with confocal and atomic force microscopy (AFM). Preliminary studies show that tonic stretching of human bronchial epithelial cells caused a significant increase in the production of mitochondrial superoxide. Moreover, using this protocol, alveolar epithelial cells were stretched and imaged, which showed direct damage to the epithelial cells by overdistention simulating one form of lung injury in vitro. A protocol to conduct AFM nano-indentation on stretched cells is also provided.

A novel imaging protocol was developed using a custom motor-driven mechanical actuator to allow the measurement of real time responses to mechanical strain in live cells. Relevant to mechanobiology, the system can apply strains up to 20% while allowing near real-time imaging with confocal or atomic force microscopy.

There is currently a significant interest in understanding how cells and tissues respond to mechanical stimuli, but current approaches are limited in ...

An Optimized O9-1/Hydrogel System for Studying Mechanical Signals in Neural Crest Cells

Neural crest cells (NCCs) are vertebrate embryonic multipotent cells that can migrate and differentiate into a wide array of cell types that give rise to various organs and tissues. Tissue stiffness produces mechanical force, a physical cue that plays a critical role in NCC differentiation; however, the mechanism remains unclear. The method described here provides detailed information for the optimized generation of polyacrylamide hydrogels of varying stiffness, the accurate measurement of such stiffness, and the evaluation of the impact of mechanical signals in O9-1 cells, a NCC line that mimics in vivo NCCs.
Hydrogel stiffness was measured using atomic force microscopy (AFM) and indicated different stiffness levels accordingly. O9-1 NCCs cultured on hydrogels of varying stiffness showed different cell morphology and gene expression of stress fibers, which indicated varying biological effects caused by mechanical signal changes. Moreover, this established that varying the hydrogel stiffness resulted in an efficient in vitro system to manipulate mechanical signaling by altering gel stiffness and analyzing the molecular and genetic regulation in NCCs. O9-1 NCCs can differentiate into a wide range of cell types under the influence of the corresponding differentiation media, and it is convenient to manipulate chemical signals in vitro. Therefore, this in vitro system is a powerful tool to study the role of mechanical signaling in NCCs and its interaction with chemical signals, which will help researchers better understand the molecular and genetic mechanisms of neural crest development and diseases.

Detailed step-by-step protocols are described here for studying mechanical signals in vitro using multipotent O9-1 neural crest cells and polyacrylamide hydrogels of varying stiffness.

Neural crest cells (NCCs) are vertebrate embryonic multipotent cells that can migrate and differentiate into a wide array of cell types that give rise to ...

Developmental Biology

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the physical properties of the nucleus contribute significantly to the overall biomechanical behavior of cells under physiological and pathological conditions. For example, in migrating neutrophils and invading cancer cells, nuclear stiffness can pose a major obstacle during extravasation or passage through narrow spaces within tissues.1 On the other hand, the nucleus of cells in mechanically active tissue such as muscle requires sufficient structural support to withstand repetitive mechanical stress. Importantly, the nucleus is tightly integrated into the cellular architecture; it is physically connected to the surrounding cytoskeleton, which is a critical requirement for the intracellular movement and positioning of the nucleus, for example, in polarized cells, synaptic nuclei at neuromuscular junctions, or in migrating cells.2 Not surprisingly, mutations in nuclear envelope proteins such as lamins and nesprins, which play a critical role in determining nuclear stiffness and nucleo-cytoskeletal coupling, have been shown recently to result in a number of human diseases, including Emery-Dreifuss muscular dystrophy, limb-girdle muscular dystrophy, and dilated cardiomyopathy.3 To investigate the biophysical function of diverse nuclear envelope proteins and the effect of specific mutations, we have developed experimental methods to study the physical properties of the nucleus in single, living cells subjected to global or localized mechanical perturbation. Measuring induced nuclear deformations in response to precisely applied substrate strain application yields important information on the deformability of the nucleus and allows quantitative comparison between different mutations or cell lines deficient for specific nuclear envelope proteins. Localized cytoskeletal strain application with a microneedle is used to complement this assay and can yield additional information on intracellular force transmission between the nucleus and the cytoskeleton. Studying nuclear mechanics in intact living cells preserves the normal intracellular architecture and avoids potential artifacts that can arise when working with isolated nuclei. Furthermore, substrate strain application presents a good model for the physiological stress experienced by cells in muscle or other tissues (e.g., vascular smooth muscle cells exposed to vessel strain). Lastly, while these tools have been developed primarily to study nuclear mechanics, they can also be applied to investigate the function of cytoskeletal proteins and mechanotransduction signaling.

We present two independent, microscope-based tools to measure the induced nuclear and cytoskeletal deformations in single, living adherent cells in response to global or localized strain application. These techniques are used to determine nuclear stiffness (i.e., deformability) and to probe intracellular force transmission between the nucleus and the cytoskeleton.

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the ...

Biology

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record forces curves on 1000 cells in 4 hours automatically. To maintain a 4 hour analysis time, the number of force curves per cell is reduced to 9 or 16. The method combines a Jython based program and a strategy for assembling cells on defined patterns. The program, implemented on a commercial Bio-AFM, can center the tip on the first cell of the array and then move, automatically, from cell to cell while recording force curves on each cell. Using this methodology, it is possible to access the biophysical parameters of the cells such as their rigidity, their adhesive properties, etc. With the automation and the large number of cells analyzed, one can access the behavior of the cell population. This is a breakthrough in the Bio-AFM field where data have, so far, been recorded on only a few tens of cells.

Automation of Bio-Atomic Force Microscope Measurements on Hundreds of C. albicans Cells

This protocol aims to automate AFM measurements on hundreds of microbial cells. First, microbes are immobilized into PDMS stamp microstructures and then force spectroscopy measurements are performed automatically on hundreds of immobilized cells.

The method presented in this paper aims to automate Bio-AFM experiments and the recording of force curves. Using this method, it is possible to record ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Measurement of Liver Stiffness Using Atomic Force Microscopy Coupled with Polarization Microscopy

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell behavior such as cell function, differentiation, and motility. However, as these processes are not homogeneous throughout the whole organ, it has become increasingly important to understand changes in the mechanical properties of tissues on the cellular level.
To be able to monitor the stiffening of collagen-rich areas within the liver lobes, this paper presents a protocol for measuring liver tissue elastic moduli with high spatial precision by atomic force microscopy (AFM). AFM is a sensitive method with the potential to characterize local mechanical properties, calculated as Young&#39;s (also referred to as elastic) modulus. AFM coupled with polarization microscopy can be used to specifically locate the areas of fibrosis development based on the birefringence of collagen fibers in tissues. Using the presented protocol, we characterized the stiffness of collagen-rich areas from fibrotic mouse livers and corresponding areas in the livers of control mice.
A prominent increase in the stiffness of collagen-positive areas was observed with fibrosis development. The presented protocol allows for a highly reproducible method of AFM measurement, due to the use of mildly fixed liver tissue, that can be used to further the understanding of disease-initiated changes in local tissue mechanical properties and their effect on the fate of neighboring cells.

We present a protocol to measure the elastic moduli of collagen-rich areas in normal and diseased liver using atomic force microscopy. The simultaneous use of polarization microscopy provides high spatial precision for localizing collagen-rich areas in the liver sections.

Matrix stiffening has been recognized as one of the key drivers of the progression of liver fibrosis. It has profound effects on various aspects of cell ...

Stiffness Measurement of Soft Silicone Substrates for Mechanobiology Studies Using a Widefield Fluorescence Microscope

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