The authors would like to acknowledge the Interdisciplinary Innovation Initiative Program, University of Illinois, grant 12035. S.K. was funded at UIUC from National Science Foundation (NSF) Grant 0965918 IGERT: Training the Next Generation of Researchers in Cellular and Molecular Mechanics and BioNanotechnology.&#160;This work was supported in part by the grant from the nanoBIO node of the National Science Foundation ( ECC-1227034).

Fabricating and functionalizing PA gels of varying stiffnesses with fluorescent microspheres embedded near the cell culture surface.
1. Functionalizing the Top Glass Cover Slips
<ol>
	<li>Clean glass cover slips (#1.0, 12 mm diam.) with soap and water, followed by ethanol to remove extraneous dust.</li>
	<li>Place glass cover slips on a grated surface (i.e. pipette tip holder) such that they are not touching to facilitate ease of interaction with the coverslips.</li>
	<li>Coat the entire surface of the cover slips with Poly-D-Lysine (0.1 mg/ml) for 1 hr (Figure 1A).</li>
	<li>During this time, perform a 1:10,000 dilution of the colloid solution of 0.1 &#956;m diameter, red fluorescent microspheres with deionized (DI) water to obtain a particle density of approximately 1 microsphere per 20 &#956;m2 on the gel surface. See Figure 2 for the results of various dilutions. This dilution can be modified to meet the need of specific experiments.</li>
	<li>Place the diluted solution in an ultrasonic water bath for 30 min.</li>
	<li>After 1 hr, use tweezers to carefully lift each cover slip and blow dry with air. Return the dry cover slips to the grated surface.</li>
	<li>Remove the diluted colloid solution from the ultrasonic bath and pipette 150 &#956;l onto each cover slip. Leave for 10 min (Figure 1B).</li>
	<li>Use tweezers to carefully lift each cover slip and blow dry with air. Return the dry cover slips to the grated surface and store in the dark until ready to use.</li>
</ol>
2. Preparing PA Gel Directly on Glass Bottom Petri Dishes
<ol>
	<li>Preheat hotplate to 100 &#176;C.</li>
	<li>Lay out the desired number of glass bottom Petri dishes (35 mm dish with 14 mm micro-well, #1.0) on a flat surface in a chemical fume hood.</li>
	<li>Cover the glass portion of each Petri dish micro-well with 97% 3-aminopropyl-trimethoxysliane (3-APTES) for 7 min for chemical activation. Take caution to avoid inadvertent dripping of the 3-APTES to the surface of the plastic in the Petri dish to avoid degradation of the polystyrene.</li>
	<li>After 7 min, fill the Petri dish with DI water and dispose into waste container.</li>
	<li>Repeat step 2.4 3x&#160;for each dish, and then shake the Petri dish to remove extra water. Place the Petri dishes on the hot plate until the glass portion is dry.</li>
	<li>Remove the Petri dishes from the hot plate and return to a flat surface in a chemical fume hood.</li>
	<li>In a chemical fume hood, make a solution of 0.5% glutaraldehyde and cover the glass portion of each Petri dish well with the solution for 30 min. Take caution to avoid inadvertent dripping of the glutaraldehyde to the surface of the plastic in the Petri dish to avoid degradation of the plastic.</li>
	<li>After 30 min, fill the Petri dish with DI water and dispose into waste container to rinse and remove the glutaraldehyde.</li>
	<li>Repeat step 2.4 3x&#160;for each dish, and then shake the Petri dish to remove extra water. Place the Petri dishes on the hot plate until the glass portion is dry.</li>
	<li>Before mixing the components of the PA gel solution, move the functionalized glass slides into the chemical fume hood such that they are easily accessible, allowing for the quick sandwiching of the gel with the glass bottom Petri dishes after mixing the gel solution.</li>
	<li>In a 15 ml centrifuge tube, mix 40% bisacrylamide, 2% acrylamide, and acrylic acid in immediate succession in the concentrations listed in Table 1 (adapted from published protocol10) to achieve the desired matrix elasticity.</li>
	<li>Add 100 mM HEPES, 10% ammonium persulfate, and TEMED in quantities listed in Table 1 corresponding to desired matrix elasticity to complete the gel solution.</li>
	<li>Immediately pipette 15 &#956;l of gel solution onto the center of the glass portion of the petri dishes.</li>
	<li>Immediately pick up a functionalized glass cover slip with tweezers.
	<ol>
		<li>Flip the glass cover slip over such that the fluorescent beads are on the side making contact with gel solution.</li>
		<li>Lay the cover slip gently on top of the now-liquid PA gel such that the functionalized side is in contact with the gel (Figure 1C). Note: For best results, a second person is recommended for the role of adding the cover slip in order to avoid possibility of partial polymerization while pipetting liquid PA gel solution onto multiple Petri dishes.</li>
	</ol>
	</li>
	<li>Flip all Petri dishes over to assist with avoiding gravity effects on fluorescent nanoparticles polymerizing into lower levels of the PA gel.</li>
	<li>Wait for at least 35 min, or until the stock solution of PA gel has visibly polymerized in its centrifuge tube.</li>
	<li>Flip the Petri dishes back over and fill them with PBS to assist with removing the cover slip.</li>
	<li>Carefully make contact with the glass portion of the Petri dish and the outline of the cover slip, using tweezers to scrape the circumference of the cover slip. Perform several cycles until the cover slip is dislodged. Remove the cover slip and dispose in a proper sharps waste container.</li>
	<li>After removing all cover slips, fluorescent beads will have transferred to the gel (Figure 1D). Cover the PA gels completely with PBS, place the Petri dish lid on each dish, and store at 4 &#176;C.</li>
</ol>
3. Functionalizing PA gel with Fibronectin
<ol>
	<li>Prepare the following premixed solutions as described in an established protocol11: Soak solution (137 mM NaCl, 5% (v/v) glycerol) and 2x conjugation buffer (0.2 M 2-(N-morpholino)ethanesulfonic acid (MES), 10% (v/v) glycerol, pH 4.5).</li>
	<li>Use a vacuum pump in a biological hood to remove all PBS from the glass-bottom dishes containing the PA gels.</li>
	<li>Pipette soak solution onto each gel such that the gel is completely submerged. Incubate at room temperature for at least 1 hr.</li>
	<li>Warm 1-ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC) and N-hydroxysulfosuccinimide (NHS) to room temperature.</li>
	<li>Mix 10x solutions of EDC (150 mM, 19 mg/ml in DI water) and NHS (250 mM, 29 mg/ml in DI water).</li>
	<li>Mix 1 part 10x EDC, 1 part 10x NHS, 3 parts DI water, and 5 parts 2x conjugation buffer.</li>
	<li>Use a vacuum pump in a biological hood to remove the soak solution. Make sure all fluid is removed from the gel surface.</li>
	<li>Add enough NHS/EDC solution to cover the gel surface and fill the glass-bottom well of the Petri dish (150&#8211;250 &#956;l). Incubate at room temperature for 30 min in the dark.</li>
	<li>Thaw fibronectin at room temperature. Once thawed, mix sterile DI water to create a 50 &#956;g/ml fibronectin solution.</li>
	<li>Use a vacuum pump in a biological hood to remove the NHS/EDC solution. Make sure all fluid is removed from the gel surface.</li>
	<li>Add 150 &#956;l of fibronectin solution to each gel. Incubate at room temperature for 35 min to allow for attachment of fibronectin.</li>
	<li>After 35 min, add PBS to each Petri dish and store at 4 &#176;C for up to 2 weeks.</li>
</ol>
4. Traction Force Experiments
<ol>
	<li>Warm cell media, PBS, and trypsin to 37 &#176;C in a water bath.</li>
	<li>Rinse gels 5x with sterile PBS, aspirating the PBS in between rinses, and leave covered in the hood.</li>
	<li>Add 1 ml trypsin per 25 cm2 to flask containing cells. After cells are lifted from flask, dilute the trypsin with cell media and count the cells. Based on gel surface area, determine the number of cells required per gel for a final cell seeding density of 3,000 cells/cm2. Dilute or concentrate suspension such that 150 &#956;l of the cell-media mixture contains this number of cells. Aliquot 150 &#956;l of cell suspension on to each gel.</li>
	<li>Place the Petri dishes containing cells in an incubator for 30 min. Then, carefully remove the Petri dishes and fill the remainder of the Petri dish with media (approximately 2 ml) such that the surface of the dish is completely submerged.</li>
	<li>Place the Petri dishes back in the incubator until image acquisition.</li>
	<li>Prepare the microscope and data acquisition system for imaging: insert the 40x water immersion objective, insert the Differential Interference Contrast (DIC) prism, select the mCherry or equivalent fluorescent filter, and turn on the environmental chamber.</li>
	<li>When prepared for imaging, remove one Petri dish from the incubator and place it gently on the microscope stage. Remove the Petri dish lid for DIC imaging.</li>
	<li>Locate a single cell and capture a single still image of the cell in DIC.</li>
	<li>Without moving the microscope stage, switch the imaging mode to fluorescence. Focus on the fluorescent microspheres and record an image of the microspheres.</li>
	<li>Carefully remove cell media from the Petri dish with a pipette and add 0.05% trypsin-EDTA.</li>
	<li>Image the microspheres under the cell after the cells have detached.</li>
	<li>Use particle image velocimetry (PIV) analysis in ImageJ12 to compute the displacement field due to cellular forces.</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> Biophys. J. 79 (1), 144-152 (2000).</li><li>Aratyn-Schaus, Y., Oakes, P. W., Stricker, J., Winter, S. P., Gardel, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Complaint+Matrices+for+Quantifying+Cellular+Contraction.">Preparation of Complaint Matrices for Quantifying Cellular Contraction.</a> JoVE. , (2010).</li><li>Kandow, C. E., Georges, P. C., Janmey, P. A., Beningo, K. 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A., Boehm, H., Li, B., Vogel, V., Spatz, J. P., Watt, F. M., Huck, W. T. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extracellular-matrix+tethering+regulates+stem-cell+fate.">Extracellular-matrix tethering regulates stem-cell fate.</a> Nature Materials. 11, 642-649 (2012).</li><li>Wong, J. Y., Leach, J. B., Brown, X. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Balance+of+chemistry,+topography,+and+mechanics+at+the+cell-biomaterial+interface:+Issues+and+challenges+for+assessing+the+role+of+substrate+mechanics+on+cell+response.">Balance of chemistry, topography, and mechanics at the cell-biomaterial interface: Issues and challenges for assessing the role of substrate mechanics on cell response.</a> Surface Science. 570 (1-2), 119-133 (2004).</li><li>Mih, J. D., Sharif, A. S., Marinkovic&#769;, A., Symer, M. M., Tschumperlin, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Multiwell+Platform+for+Studying+Stiffness-Dependent+Cell+Biology.">A Multiwell Platform for Studying Stiffness-Dependent Cell Biology.</a> PLoS ONE. 6 (5), e19929 (2011).</li><li>Butler, J. P., Tolic&#769;-N&#248;rrelykke, I. M., Fabry, B., Fredberg, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Traction+fields,+moments,+and+strain+energy+that+cells+exert+on+their+surroundings.">Traction fields, moments, and strain energy that cells exert on their surroundings.</a> Am J Physiol Cell Physiol. 282, 595-605 (2001).</li><li>Tolic&#769;-N&#248;rrelykke, I. M., Butler, J. 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The authors declare that they have no competing financial interests.

When using this technique, it is essential that the bead solution is properly diluted and the beads are chosen based on desired diameter, PA gel substrate stiffness, and the size scale of the phenomena that is explored in the desired experiment.
Caution should be taken when diluting the bead solution prior to functionalizing top glass cover slips. The spacing between beads on the gel surface can be altered by changing the dilution factor of the colloid solution. Diluting the solution too much ...

The mechanical interaction of a living cell with its local environment has commonly been studied using PA gels. These substrates rely on a simple, well-characterized protocol established by Dembo and Wang in 19971. One of the main advantages of these substrates is that their stiffness may be tuned by modifying the concentrations of specific components of the gel solution. This provides a desirable platform to study cells&#8217; interaction with environments of different rigidities. When PA gels are coated with extracellular matrix (ECM) proteins, cells adhere to them, generating force. As a result of cell force, the gel deforms as an elastic body. This deformation depends on the magnitude of the force applied by the cells and the elastic properties of the gel. Various studies have employed PA gels to investigate cellular traction forces.
In one variation of PA gel fabrication, fluorescent microspheres (beads) are embedded throughout the gel to quantify cell traction forces on gels of different rigidities2. Upon cell force application, the beads displace from their initial location following gel deformation. The deformation field is measured from the individual bead displacements. This deformation field is utilized with elasticity theory and the elastic properties of the gel to compute the traction forces. These measurements provide insight as to how cells mechanically sense and interact with their local microenvironment3.
In many widely used PA gel fabrication protocols, the beads are intermixed throughout the PA gel in its liquid, unpolymerized state. A fully polymerized PA gel contains fluorescent beads throughout its volume. When computing cell traction forces, beads most near the gel surface (cell-substrate interface) are monitored. The displacements of these beads are assumed to occur on the cell culture surface for simplicity in the force calculation. The actual location of the beads within the depth of the gel is disregarded. However, in an elastic medium (such as PA gel), a bead closer to a point of force application will move more than a bead that is further away from the point. Thus, treating the displacement of a point (at the bead location) distal from the surface as that at the surface results in an underestimation of cellular tractions. The degree of error depends on the distance of the bead from the surface. The error cannot be estimated without the knowledge of the location of the bead.
The need for a simple method to confine beads very near the cell culture surface has been addressed by a few techniques. One way is to increase the density of beads throughout the whole gel such that there are a sufficient number of beads in the top focal plane to measure motion very near the surface. Another technique involves building a confocal imaging chamber for live cell imaging such that the light from only the beads in the upper-most focal plane is collected4. A different method involves overlaying an extremely thin layer of PA gel containing beads on top of an already polymerized gel without beads5. A drawback of each of these techniques is that the precise location of the beads within the gel is not known. This introduces error into the calculation of the displacement field of the beads, and thus the calculation of cell forces. Another technique involves conjugation of beads to the top surface of an already polymerized PA gel using Sulfo-SANPAH6. This technique ensures the beads are indeed only on the top of the PA gel, but the extent to which they are embedded in the depth of the gel is unknown. This could potentially create a local topography for the cells, which could alter cell behavior, as prior work has suggested that cells can sense force several microns away7. Recently, a technique for patterning PA gels with 1 &#956;m diameter fluorescent fibronectin dot markers in a regularized array was established8. In this case, the depth of the fluorescent markers is known, and is essentially zero, as the fibronectin pattern is indirectly printed on the gel surface. However, this method does not provide a continuous environment on which cells can attach, as the ECM protein is limited to 1 &#956;m diameter dots. A method for fully integrating tracker beads within PA gels and confining them to a known location very near the surface has yet to be established.
Here, we develop a technique to constrain sub-&#956;m to &#956;m diameter fluorescent beads to a focal plane very near the cell culture surface within PA gel. A gel is typically cured by sandwiching unpolymerized liquid gel solution between two glass plates. One of the plates is functionalized so that the gel strongly adheres to it. The other is unfunctionalized and is removed after the gel cures. We modify this removable glass surface by coating it with a layer of beads. Upon sandwiching the liquid gel between the functionalized and the bead coated glass surface, the beads transfer to the gel while it is curing. This limits the distance of the beads&#8217; integration into the gel within 1.6 &#181;m of the surface. Glass-bottom Petri dishes are used as the adherent surface on which the gel is cured. To form a flat top gel surface during polymerization, a circular glass cover slip is used to sandwich the gel with the glass-bottom Petri dish. Prior to gel fabrication, the top glass cover slip is coated with poly-D-lysine (PDL), yielding a positive surface charge. The PDL is blown off with compressed air, and a solution of beads in water is deposited on the cover slips. We utilize carboxylated fluorescent microbeads, which carry a negative charge, and interact with the positively charged surface created by treatment with PDL. After blowing the bead solution off the coverslip with compressed air, a single layer of beads remains electrostatically coupled to the dry cover slip. The PDL coating does not affect the adhesiveness of the glass to the gel surface, as the glass slides are undamaged and removed from the PA gel fully intact.
The glass-bottom Petri dishes are made adherent by treatment with 97% 3-aminopropyl-trimethoxysliane and 0.5% glutaraldehyde. PA gels of desired rigidities are created by mixing appropriate concentrations of bisacrylamide and acrylamide via a standard procedure9. A droplet of the gel solution is pipetted onto the glass-bottom Petri dish. The glass cover slip containing the beads is used to sandwich the gel with the Petri dish. When the gel is cured, the top cover slip is removed leaving the beads embedded in the PA gel within 1.6 &#181;m from the surface.

<table border="0" cellpadding="0" cellspacing="0" width="502">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Name</td>
			<td style="width:128px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Reagents</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:255px;">97% 3-Aminopropyl-trimethoxysliane (APTES)</td>
			<td>Sigma-Aldrich</td>
			<td>281778</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">70% Glutaraldehyde</td>
			<td>Polysciences, Inc.</td>
			<td>111-30-8</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">1M HEPES buffer solution</td>
			<td>Sigma-Aldrich</td>
			<td>83264</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">40% Acrylamide</td>
			<td>Sigma-Aldrich</td>
			<td>A4058</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">2% Bisacrylamide</td>
			<td>Sigma-Aldrich</td>
			<td>M1533</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:255px;">0.1 um Fluorescent microspheres (mCherry)</td>
			<td>Invitrogen</td>
			<td>F8801</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Fibronectin, Human, 1mg</td>
			<td>BD Biosciences</td>
			<td>354008</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Ammonium Persulfate</td>
			<td>Bio-RAD</td>
			<td>161-0700</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:255px;">Tetramethylethylenediamine (TEMED)</td>
			<td>Bio-RAD</td>
			<td>161-0801</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Poly-D-Lysine</td>
			<td>Millipore</td>
			<td>A-003-E</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:255px;">1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC)&#160;</td>
			<td>Thermo Scientific</td>
			<td>22980</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">N-hydroxysulfosuccinimide (NHS)</td>
			<td>Thermo Scientific</td>
			<td>24500</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">.05% Trypsin-EDTA (1X)</td>
			<td>Life Technologies</td>
			<td>25300-054</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">NaCL</td>
			<td>Sigma-Aldrich</td>
			<td>S9888</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Acrylic Acid&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>147230</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Glycerol</td>
			<td>Sigma-Aldrich</td>
			<td>G7757</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:255px;">2-(N-morpholino)ethanesulfonic acid (MES)</td>
			<td>Sigma-Aldrich</td>
			<td>M3671</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;"></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Materials</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:255px;">35 mm Glass bottom dish with 14 mm micro-well #1 cover glass</td>
			<td>In Vitro Scientific</td>
			<td>D35-14-1-N</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:255px;">Glass cover slips, 12 mm diam.</td>
			<td>Ted Pella, Inc.</td>
			<td>26023</td>
		</tr>
	</tbody>
</table>

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.

Confocal imaging was used to determine that the beads were indeed underneath the gel surface and to quantify their precise location within the gel depth. Fluorescent beads of a different wavelength than those inside the gel were allowed to settle on the surface, and the distance between the embedded fluorescent nanoparticles and those on the surface was calculated using a centroid identification algorithm. The location of the bead on the gel surface serves as a reference for the top surface of the gel&#8212;where cells a...

Watch this Scientific Journal Video about A Novel Method for Localizing Reporter Fluorescent Beads Near the Cell Culture Surface for Traction Force Microscopy at JoVE.com

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

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 ...

a-novel-method-for-localizing-reporter-fluorescent-beads-near-cell

Research

JoVE Journal

Bioengineering

13.6K vistas.  University of Illinois at Urbana-Champaign.  Las técnicas tradicionales para la fabricación de poliacrilamida (PA) geles que contienen sondas fluorescentes implican intercalando un gel entre una superficie adherente y un portaobjetos de vidrio. Aquí, mostramos que el revestimiento de esta diapositiva con poli-D-lisina (PDL) y sondas fluorescentes localiza las sondas dentro de 1,6 m de la superficie del gel. 

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

Self-reporting Scaffolds for 3-Dimensional Cell Culture

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting 3D cellular construct can often be more relevant to studying the molecular events and cell-cell interactions than similar experiments studied in 2D. To create effective 3D cultures with high cell viability throughout the scaffold the culture conditions such as oxygen and pH need to be carefully controlled as gradients in analyte concentration can exist throughout the 3D construct. Here we describe the methods of preparing biocompatible pH responsive sol-gel nanosensors and their incorporation into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds along with their subsequent preparation for the culture of mammalian cells. The pH responsive scaffolds can be used as tools to determine microenvironmental pH within a 3D cellular construct. Furthermore, we detail the delivery of pH responsive nanosensors to the intracellular environment of mammalian cells whose growth was supported by electrospun PLGA scaffolds. The cytoplasmic location of the pH responsive nanosensors can be utilized to monitor intracellular pH (pHi) during ongoing experimentation.

Biocompatible pH responsive sol-gel nanosensors can be incorporated into poly(lactic-co-glycolic acid) (PLGA) electrospun scaffolds. The produced self-reporting scaffolds can be used for in&#160;situ monitoring of microenvironmental conditions whilst culturing cells upon the scaffold. This is beneficial as the 3D cellular construct can be monitored in real-time without disturbing the experiment.

Culturing cells in 3D on appropriate scaffolds is thought to better mimic the in vivo microenvironment and increase cell-cell interactions. The resulting ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Surface Potential Measurement of Bacteria Using Kelvin Probe Force Microscopy

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. Kelvin probe force microscopy (KPFM) is a module of atomic force microscopy (AFM) that measures the contact potential difference between surfaces at the nano-scale. The combination of KPFM with AFM allows for the simultaneous generation of surface potential and topographical maps of biological samples such as bacterial cells. Here, we employ KPFM to examine the effects of surface potential on microbial adhesion to medically relevant surfaces such as stainless steel and gold. Surface potential maps revealed differences in surface potential for microbial membranes on different material substrates. A step-height graph was generated to show the difference in surface potential at a boundary area between the substrate surface and microorganisms. Changes in cellular membrane surface potential have been linked with changes in cellular metabolism and motility. Therefore, KPFM represents a powerful tool that can be utilized to examine the changes of microbial membrane surface potential upon adhesion to various substrate surfaces. In this study, we demonstrate the procedure to characterize the surface potential of individual methicillin-resistant Staphylococcus aureus USA100 cells on stainless steel and gold using KPFM.

Here, we present a protocol explaining the use of Kelvin probe force microscopy as a tool for generating high resolution nano-scale surface potential maps. This tool was applied to assess the role of surface potential on the binding capacity of microorganisms to substrate surfaces.

Surface potential is a commonly overlooked physical characteristic that plays a dominant role in the adhesion of microorganisms to substrate surfaces. ...

Atomic Force Microscopy Imaging and Force Spectroscopy of Supported Lipid Bilayers

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient magnification to examine membrane substructures and even individual molecules. AFM can act as a force probe to measure interactions and mechanical properties of membranes. Supported lipid bilayers are conventionally used as membrane models in AFM studies. In this protocol, we demonstrate how to prepare supported bilayers and characterize their structure and mechanical properties using AFM. These include bilayer thickness and breakthrough force. 
The information provided by AFM imaging and force spectroscopy help define mechanical and chemical properties of membranes. These properties play an important role in cellular processes such as maintaining cell hemostasis from environmental stress, bringing membrane proteins together, and stabilizing protein complexes.

We describe a protocol for preparation of supported lipid bilayers and its characterization using atomic force microscopy and force spectroscopy.

Atomic force microscopy (AFM) is a versatile, high-resolution imaging technique that allows visualization of biological membranes. It has sufficient ...

A High-throughput Cell Microarray Platform for Correlative Analysis of Cell Differentiation and Traction Forces

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly controlled, high-throughput engineered system for measuring the impact of arrayed biochemical signals on cell differentiation. Recent efforts using cell microarrays have demonstrated their utility for combinatorial studies in which many microenvironmental factors are presented in parallel. However, these efforts have focused primarily on investigating the effects of biochemical cues on cell responses. Here, we present a cell microarray platform with tunable material properties for evaluating both cell differentiation by immunofluorescence and biomechanical cell&#8211;substrate interactions by traction force microscopy. To do so, we have developed two different formats utilizing polyacrylamide hydrogels of varying Young&#39;s modulus fabricated on either microscope slides or glass-bottom Petri dishes. We provide best practices and troubleshooting for the fabrication of microarrays on these hydrogel substrates, the subsequent cell culture on microarrays, and the acquisition of data. This platform is well-suited for use in investigations of biological processes for which both biochemical (e.g., extracellular matrix composition) and biophysical (e.g., substrate stiffness) cues may play significant, intersecting roles.

Cell differentiation is regulated by a host of microenvironmental factors, including both matrix composition and substrate material properties. We describe here a technique utilizing cell microarrays in conjunction with traction force microscopy to evaluate both cell differentiation and biomechanical cell&#8211;substrate interactions as a function of microenvironmental context.

Microfabricated cellular microarrays, which consist of contact-printed combinations of biomolecules on an elastic hydrogel surface, provide a tightly ...

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 ...

Traction Force Microscopy to Study B Lymphocyte Activation

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

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

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

Simultaneous Quantification of Selected Kynurenines Analyzed by Liquid Chromatography-Mass Spectrometry in Medium Collected from Cancer Cell Cultures

The kynurenine pathway and the tryptophan catabolites called kynurenines have received increased attention for their involvement in immune regulation and cancer biology. An in vitro cell culture assay is often used to learn about the contribution of different tryptophan catabolites in a disease mechanism and for testing therapeutic strategies. Cell culture medium that is rich in secreted metabolites and signaling molecules reflects the status of tryptophan metabolism and other cellular events. New protocols for the reliable quantification of multiple kynurenines in the complex cell culture medium are desired to allow for a reliable and quick analysis of multiple samples. This can be accomplished with liquid chromatography coupled with mass spectrometry. This powerful technique is employed in many clinical and research laboratories for the quantification of metabolites and can be used for measuring kynurenines.
Presented here is the use of liquid chromatography coupled with single quadrupole mass spectrometry (LC-SQ) for the simultaneous determination of four kynurenines, i.e., kynurenine, 3-hydroxykynurenine, 3-hydroxyanthranilic, and xanthurenic acid in the medium collected from in vitro cultured cancer cells. SQ detector is simple to use and less expensive compared to other mass spectrometers. In the SQ-MS analysis, multiple ions from the sample are generated and separated according to their specific mass-to-charge ratio (m/z), followed by the detection using a Single Ion Monitoring (SIM) mode.
This paper draws the attention on the advantages of the reported method and indicates some weak points. It is focused on critical elements of LC-SQ analysis including sample preparation along with chromatography and mass spectrometry analysis. The quality control, method calibration conditions and matrix effect issues are also discussed. We described a simple application of 3-nitrotyrosine as one analog standard for all target analytes. As confirmed by experiments with human ovary and breast cancer cells, the proposed LC-SQ method&#160;generates reliable results and can be further applied to other in vitro cellular models.

Described here is a protocol for the determination of four different tryptophan metabolites generated in the kynurenine pathway (kynurenine, 3-hydroxykynurenine, xanthurenic acid, 3-hydroxyanthranilic acid) in the medium collected from cancer cell cultures analyzed by liquid chromatography coupled with a single quadrupole mass spectrometry.

The kynurenine pathway and the tryptophan catabolites called kynurenines have received increased attention for their involvement in immune regulation and ...

Application of Atomic Force Microscopy to Detect Early Osteoarthritis

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in elasticity can propagate or trigger the onset of major diseases like cancer or osteoarthritis (OA). Atomic force microscopy (AFM) has emerged as a strong tool to qualitatively and quantitatively characterize the biomechanical properties of specific biological target structures on a microscopic scale, measuring forces in a range from as small as the piconewton to the micronewton. Biomechanical properties are of special importance in musculoskeletal tissues, which are subjected to high levels of strain. OA as a degenerative disease of the cartilage results in the disruption of the pericellular matrix (PCM) and the spatial rearrangement of the chondrocytes embedded in their extracellular matrix (ECM). Disruption in PCM and ECM has been associated with changes in the biomechanical properties of cartilage. In the present study we used AFM to quantify these changes in relation to the specific spatial pattern changes of the chondrocytes. With each pattern change, significant changes in elasticity were observed for both the PCM and ECM. Measuring the local elasticity thus allows for drawing direct conclusions about the degree of local tissue degeneration in OA.

We present a method to investigate early osteoarthritic changes at the cellular level in articular cartilage by using atomic force microscopy (AFM).

Biomechanical properties of cells and tissues not only regulate their shape and function but are also crucial for maintaining their vitality. Changes in ...