Name: a novel method for localizing reporter fluorescent beads near the cell culture surface for traction force microscopy
Uploaded: 2014-09-16T13:00:00.000+00:00
Duration: 13 min 30 s
Description: 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

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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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+and+Patterning+Polyacrylamide+Substrates+Functionalized+with+N-Hydroxysuccinimide.">Characterizing and Patterning Polyacrylamide Substrates Functionalized with N-Hydroxysuccinimide.</a> Cell and Mol. Bioengineering. 6 (3), 299-309 (2013).</li><li>Tseng, Q., Duchemin-Pelletier, E., Deshiere, A., Balland, M., Guillou, H., Filhol, O., Th&#233;ry, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+organization+of+the+extracellular+matrix+regulates+cell&#8211;cell+junction+positioning.">Spatial organization of the extracellular matrix regulates cell&#8211;cell junction positioning.</a> PNAS. 109 (5), 1506-1511 (2012).</li><li>Trappmann, B., Gautrot, J. E., Connelly, J. T., Strange, D. G. T., Li, Y., Oyen, M. L., Cohen Stuart, M. 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. P., Chen, J., Wang, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+and+temporal+traction+response+in+human+airway+smooth+muscle+cells.">Spatial and temporal traction response in human airway smooth muscle cells.</a> Am J Physiol Cell Physiol. 283, 1254-1266 (2002).</li><li>Atanackovic, T., Guran, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Theory of Elasticity for Scientists and Engineers. , (2000).</li></ol>

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 will reduce the number of beads that couple to the cover slip and ultimately reduce the number of beads in the gel. Diluting the solution too little can yield too high a density of beads in the gel such that they are in contact and cannot be distinguished from one another. While it is often impossible to completely avoid the presence of nanoparticle aggregation, the application of ultrasonic waves to the diluted solution effectively minimizes nanoparticle aggregation. We have found that a 10,000 fold dilution of the colloid solution utilized in this protocol gives a bead concentration of 3.64 x 108 beads/ml, and yields a desirable particle density of 0.05 beads/&#956;m2 for 0.1 &#956;m diameter beads. A 1:20,000 dilution (1.82 x 108 beads/ml) yields an average of 52 &#956;m2 per particle. Figure 2 shows images of the surface of gels with different dilutions. It should be noted that there is variability in the number of beads that transfer to the gel. This is due to the variability associated with removing the bead solution from the cover slip with compressed air during functionalization. In general, we have found that increasing the dilution factor by at least a factor of two consistently results in at least a twofold increase in the area per bead.
It is also critical to choose a bead size (diameter) that will result in full immersion of the bead inside the gel upon PA polymerization. Gel stiffness affects the location of beads as shown in Figure 5A. For 0.1 &#956;m diameter beads, the bead depth increases with decreasing stiffness. This could be a result of the inverse relationship between pore size and PA gel stiffness13. For all of the three stiffness gels tested, the beads are consistently localized inside the gel, and within 1 &#956;m of the gel surface as illustrated in Figure 8A. The average location of 1 &#956;m beads within the gel also varies with stiffness. They are embedded deeper (approximately 1.25 &#956;m) into the 1 kPa gels. For the 10 and 40 kPa gels, the location of the top of 1 &#956;m diameter beads ranges from approximately 0.5 &#956;m above to 0.5 &#956;m below the gel (Figure 5) surface, which means the beads occasionally protrude through the top of the gel (Figure 8B). This is detrimental because it introduces topography for the cells, and topography is known to affect cell-substrate interactions14. Thus, it is necessary to be mindful of choosing beads for this technique when preparing gels of a specific rigidity. Of the two bead sizes utilized to characterize this method, 0.1 &#181;m diameter proves the optimum size. Other bead sizes may be utilized in with this technique, but should be calibrated using confocal microscopy as we describe here, to understand the precise depth to which the beads are localized during gel polymerization.
This technique improves spatial resolution of beads in PA gel by increasing the number of distinguishable particles within the field of view. Localizing the beads to a layer near the top of the gel does not significantly affect the composite stiffness of the gel. Isostress composite theory is utilized to compute the composite elastic modulus, Ec, of only the 1 &#956;m-thick layer of gel containing beads as
<img alt="Equation 1" fo:content-width="1.7in" src="/files/ftp_upload/51873/51873equation1.jpg" width="262" />
where Ef is the elastic modulus of the polystyrene beads (3 GPa), Em is the elastic modulus of the gel matrix (10 kPa), and vf and vm are the volume fractions of the beads and gel matrix, respectively. Because the beads comprise only 0.03% of the volume, Ec = 10.0025 kPa is not significantly different than the elastic modulus of an isotropic gel of stiffness 10 kPa. An established method for fabricating PA gels reduces the polymerization time to 30 sec, thus minimizing the effect of gravity on the distribution of beads during polymerization15. Our method relies on the longer (~30 min) polymerization time for the beads to polymerize into the gel such that they do not protrude the gel surface. Because localizing the beads to a single layer does not significantly affect the composite gel stiffness, and the longer polymerization time aids in the transfer of beads into the gel matrix, we do not incorporate any measure to reduce polymerization time in our technique.
Our technique demonstrates how to localize the beads to a known depth inside PA gel, which improves the measurement of substrate deformation, and thus the calculation of cell traction forces. A theoretical approach to quantify the cellular traction forces has been previously reported16,17. This approach treats the gel substrate as a semi-infinite solid in elastic half space. The Boussinesq solution is used to solve the inverse problem of calculating traction forces from a displacement field. We employ a simplified version of the Boussinesq integral to quantify the error in displacement measurement due to beads distal from the gel surface18. The equation for the displacement in a given direction based on an applied shear stress, &#964;, over a given radius, R, is given by
<img alt="Equation 2" fo:content-width="3.7in" src="/files/ftp_upload/51873/51873equation2.jpg" width="558" />
where ux is the displacement in the x direction,&#160;&#956; is the shear modulus, and z is the depth of the beads from the gel surface. For various locations of beads throughout the depth of the gel, corresponding displacements are shown in Table 2.
For the case shown here, the radius is estimated to be an average focal adhesion size of 1 &#956;m. If the beads are assumed to be coplanar with the cells (z = 0), the displacement is given by 0.75&#160;&#964;/&#956;. If the beads are only 1 &#956;m below the gel surface, the displacement is given by 0.24 &#964;/&#956;. This represents greater than a threefold reduction in displacement from a bead coplanar with the surface to a location 1 &#956;m below. This introduces error in the calculation of surface traction forces based on the motion of embedded beads whose precise locations within the gel are unknown. The variation in traction defined by &#964;/&#956;, as a function of the depth of the beads in the gel for both the proposed and traditional fabrication methods is significant. Figure 9 shows the distribution of forces (given by the ratio &#964;/&#956;) based on Boussinesq theory that result from computation utilizing the distribution of bead depths, z, within the optical depth of field when beads are uniformly distributed in the gel and localized near the surface. The most accurate measurement of cell force would result if beads were on the same plane as the cells, where z = 0 and &#964;/&#956; = 0.0133. When compared to this case,&#160;&#964;/&#956; increases by 118%, 80%, and 50% for 1 kPa, 10 kPa, and 40 kPa gels, respectively, when beads are localized near the surface using the technique we propose. When beads are uniformly distributed throughout the gel depth,&#160;&#964;/&#956; increases an additional 60%, 80%, and 150% beyond the increase induced by localizing beads near the surface, for 1 kPa, 10 kPa, and 40 kPa gels, respectively. This represents a total of nearly 200% error in traction calculations when the beads are distributed throughout the gel depth. Thus, localizing the beads to a known depth inside the gel improves accuracy in displacement measurements and cell traction forces.
The technique described here presents a new way to prepare PA gel substrates for traction force microscopy applications such that the beads are localized near the gel surface. Confining the beads to a known depth within PA gel provides new information about the actual displacement of beads upon cell force application, creating a more accurate picture of how the cells are deforming their underlying substrate.

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><tbody><tr><td colspan="4">&lt;strong&gt;Reagents&lt;/strong&gt;</td></tr><tr><td>97% 3-Aminopropyl-trimethoxysliane (APTES)</td><td>Sigma-Aldrich</td><td>281778</td><td></td></tr><tr><td>70% Glutaraldehyde</td><td>Polysciences, Inc.</td><td>111-30-8</td><td></td></tr><tr><td>1 M HEPES buffer solution</td><td>Sigma-Aldrich</td><td>83264</td><td></td></tr><tr><td>40% Acrylamide</td><td>Sigma-Aldrich</td><td>A4058</td><td></td></tr><tr><td>2% Bisacrylamide</td><td>Sigma-Aldrich</td><td>M1533</td><td></td></tr><tr><td>0.1 &amp;micro;m Fluorescent microspheres (mCherry)</td><td>Invitrogen</td><td>F8801</td><td></td></tr><tr><td>Fibronectin, Human, 1 mg</td><td>BD Biosciences</td><td>354008</td><td></td></tr><tr><td>Ammonium persulfate</td><td>Bio-RAD</td><td>161-0700</td><td></td></tr><tr><td>Tetramethylethylenediamine (TEMED)</td><td>Bio-RAD</td><td>161-0801</td><td></td></tr><tr><td>Poly-D-lysine</td><td>Millipore</td><td>A-003-E</td><td></td></tr><tr><td>1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC)&amp;nbsp;</td><td>Thermo Scientific</td><td>22980</td><td></td></tr><tr><td>N-hydroxysulfosuccinimide (NHS)</td><td>Thermo Scientific</td><td>24500</td><td></td></tr><tr><td>0.05% Trypsin-EDTA (1X)</td><td>Life Technologies</td><td>25300-054</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich</td><td>S9888</td><td></td></tr><tr><td>Acrylic acid&amp;nbsp;</td><td>Sigma-Aldrich</td><td>147230</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G7757</td><td></td></tr><tr><td>2-(N-morpholino)ethanesulfonic acid (MES)</td><td>Sigma-Aldrich</td><td>M3671</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;Materials&lt;/strong&gt;</td></tr><tr><td>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><td></td></tr><tr><td>Glass cover slips, 12 mm diam.</td><td>Ted Pella, Inc.</td><td>26023</td><td></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.

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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 ...

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Nanyang Technological University

Research

JoVE Journal

Bioengineering

13.7K Views.  University of Illinois at Urbana-Champaign.  Traditionele technieken voor het vervaardigen polyacrylamide (PA) gels die fluorescente probes omvatten waartussen een gel tussen een hechtend oppervlak en een glasplaatje. Hier laten we zien dat deze dia bekleden met poly-D-lysine (PDL) en fluorescerende probes lokaliseert de probes binnen 1,6 urn uit het gel oppervlak. 

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

Fluorescence Biomembrane Force Probe: Concurrent Quantitation of Receptor-ligand Kinetics and Binding-induced Intracellular Signaling on a Single Cell

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions are likely affected by the mechanical environment in which binding and signaling take place. A recent study demonstrated that mechanical force can regulate antigen recognition by and triggering of the T-cell receptor (TCR). This was made possible by a new technology we developed and termed fluorescence biomembrane force probe (fBFP), which combines single-molecule force spectroscopy with fluorescence microscopy. Using an ultra-soft human red blood cell as the sensitive force sensor, a high-speed camera and real-time imaging tracking techniques, the fBFP is of ~1 pN (10-12 N), ~3 nm and ~0.5 msec in force, spatial and temporal resolution. With the fBFP, one can precisely measure single receptor-ligand binding kinetics under force regulation and simultaneously image binding-triggered intracellular calcium signaling on a single live cell. This new technology can be used to study other membrane receptor-ligand interaction and signaling in other cells under mechanical regulation.

We describe a technique for concurrently measuring force-regulated single receptor-ligand binding kinetics and real-time imaging of calcium signaling in a single T lymphocyte.

Membrane receptor-ligand interactions mediate many cellular functions. Binding kinetics and downstream signaling triggered by these molecular interactions ...

Use of Dual Optical Tweezers and Microfluidics for Single-Molecule Studies

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the averaging that takes place in bulk-phase studies. To achieve visualization, dual optical tweezers, where one trap is fixed and the other is mobile, are focused into one channel of a multi-stream microfluidic chamber positioned on the stage of an inverted fluorescence microscope. The optical tweezers trap single molecules of fluorescently labeled DNA and fluid flow through the chamber and past the trapped beads, stretches the DNA to B-form (under minimal force, i.e., 0 pN) with the nucleic acid being observed as a white string against a black background. DNA molecules are moved from one stream to the next, by translating the stage perpendicular to the flow to enable the initiation of reactions in a controlled manner. To achieve success, microfluidic devices with optically clear channels are mated to glass syringes held in place in a syringe pump. Optimal results use connectors permanently bonded to the flow cell, tubing that is mechanically rigid and chemically resistant, and which is connected to switching valves that eliminate bubbles that prohibit laminar flow.

Visual, single-molecule biochemistry studied through microfluidic chambers is greatly facilitated using glass barrel, gas-tight syringes, stable connections of tubing to flow cells, and elimination of bubbles by placing switching valves between the syringes and tubing. The protocol describes dual optical traps that enable visualization of DNA transactions and intermolecular interactions.

Visual biochemistry is a powerful technique for observing the stochastic properties of single enzymes or enzyme complexes that are obscured in the ...

Biochemistry

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are therefore poised to be useful in the field of mechanobiology. Since the method commonly relies on image-based tracking of magnetic beads, the speed limit in recording and analyzing images, as well as the thermal fluctuations of the beads, has long hampered its application in observing small and fast structural changes in target molecules. This article describes detailed methods for the construction and operation of a high-resolution MT setup that can resolve nanoscale, millisecond dynamics of biomolecules and their complexes. As application examples, experiments with DNA hairpins and SNARE complexes (membrane-fusion machinery) are demonstrated, focusing on how their transient states and transitions can be detected in the presence of piconewton-scale forces. We expect that high-speed MTs will continue to enable high-precision nanomechanical measurements on molecules that sense, transmit, and generate forces in cells, and thereby deepen our molecular-level understanding of mechanobiology.

Here, we describe a high-speed magnetic tweezer setup that performs nanomechanical measurements on force-sensitive biomolecules at the maximum rate of 1.2 kHz. We introduce its application to DNA hairpins and SNARE complexes as model systems, but it will be also applicable to other molecules involved in mechanobiological events.

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are ...

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 ...

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

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

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

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

Analyses of Actin Dynamics, Clutch Coupling and Traction Force for Growth Cone Advance

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes depend on the advances of growth cones that located at the tips of neurites. Axonal growth cones generate driving forces by sensing their local microenvironment and modulating cytoskeletal dynamics and actin-adhesion coupling (clutch coupling). Decades of research have led to the identification of guidance molecules, their receptors, and downstream signaling cascades for regulating neuronal migration and axonal guidance; however, the molecular machineries required for generating forces to drive growth cone advance and navigation are just beginning to be elucidated. At the leading edge of neuronal growth cones, actin filaments undergo retrograde flow, which is powered by actin polymerization and actomyosin contraction. A clutch coupling between F-actin retrograde flow and adhesive substrate generates traction forces for growth cone advance. The present study describes a detailed protocol for monitoring F-actin retrograde flow by single speckle imaging. Importantly, when combined with an F-actin marker Lifeact, this technique can quantify 1) the F-actin polymerization rate and 2) the clutch coupling efficiency between F-actin retrograde flow and the adhesive substrate. Both are critical variables for generating forces for growth cone advance and navigation. In addition, the present study describes a detailed protocol of traction force microscopy, which can quantify 3) traction force generated by growth cones. Thus, by coupling the analyses of&#160;single speckle imaging and traction force microscopy, investigators can monitor the molecular mechanics underlying growth cone advance and navigation.

To advance, growth cones must exert traction forces against the external environment. The generation of traction forces is dependent on actin dynamics and clutch coupling. The present study describes methods for analyzing actin dynamics, clutch coupling and traction forces for growth cone advance.

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes ...

Neuroscience

Imaging Molecular Adhesion in Cell Rolling by Adhesion Footprint Assay

Rolling adhesion, facilitated by selectin-mediated interactions, is a highly dynamic, passive motility in recruiting leukocytes to the site of inflammation. This phenomenon occurs in postcapillary venules, where blood flow pushes leukocytes in a rolling motion on the endothelial cells. Stable rolling requires a delicate balance between adhesion bond formation and their mechanically-driven dissociation, allowing the cell to remain attached to the surface while rolling in the direction of flow. Unlike other adhesion processes occurring in relatively static environments, rolling adhesion is highly dynamic as the rolling cells travel over thousands of microns at tens of microns per second. Consequently, conventional mechanobiology methods such as traction force microscopy are unsuitable for measuring the individual adhesion events and the associated molecular forces due to the short timescale and high sensitivity required. Here, we describe our latest implementation of the adhesion footprint assay to image the P-selectin: PSGL-1 interactions in rolling adhesion at the molecular level. This method utilizes irreversible DNA-based tension gauge tethers to produce a permanent history of molecular adhesion events in the form of fluorescence tracks. These tracks can be imaged in two ways: (1) stitching together thousands of diffraction-limited images to produce a large field of view, enabling the extraction of adhesion footprint of each rolling cell over thousands of microns in length, (2) performing DNA-PAINT to reconstruct super-resolution images of the fluorescence tracks within a small field of view. In this study, the adhesion footprint assay was used to study HL-60 cells rolling at different shear stresses. In doing so, we were able to image the spatial distribution of the P-selectin: PSGL-1 interaction and gain insight into their molecular forces through fluorescence intensity. Thus, this method provides the groundwork for the quantitative investigation of the various cell-surface interactions involved in rolling adhesion at the molecular level.

This protocol presents the experimental procedures to perform the adhesion footprint assay to image the adhesion events during fast cell rolling adhesion.

Rolling adhesion, facilitated by selectin-mediated interactions, is a highly dynamic, passive motility in recruiting leukocytes to the site of ...

Biology

Directly Measuring Forces Within Reconstituted Active Microtubule Bundles

Microtubule networks are employed in cells to accomplish a wide range of tasks, ranging from acting as tracks for vesicle transport to working as specialized arrays during mitosis to regulate chromosome segregation. Proteins that interact with microtubules include motors such as kinesins and dynein, which can generate active forces and directional motion, as well as non-motor proteins that crosslink filaments into higher-order networks or regulate filament dynamics. To date, biophysical studies of microtubule-associated proteins have overwhelmingly focused on the role of single motor proteins needed for vesicle transport, and significant progress has been made in elucidating the force-generating properties and mechanochemical regulation of kinesins and dyneins. However, for processes in which microtubules act both as cargo and track, such as during filament sliding within the mitotic spindle, much less is understood about the biophysical regulation of ensembles of the crosslinking proteins involved. Here, we detail our methodology for directly probing force generation and response within crosslinked microtubule minimal networks reconstituted from purified microtubules and mitotic proteins. Microtubule pairs are crosslinked by proteins of interest, one microtubule is immobilized to a microscope coverslip, and the second microtubule is manipulated by an optical trap. Simultaneous total internal reflection fluorescence microscopy allows for multichannel visualization of all the components of this microtubule network as the filaments slide apart to generate force. We also demonstrate how these techniques can be used to probe pushing forces exerted by kinesin-5 ensembles and how viscous braking forces arise between sliding microtubule pairs crosslinked by the mitotic MAP PRC1. These assays provide insights into the mechanisms of spindle assembly and function and can be more broadly adapted to study dense microtubule network mechanics in diverse contexts, such as the axon and dendrites of neurons and polar epithelial cells.

Here, we present a protocol for reconstituting microtubule bundles in vitro and directly quantifying the forces exerted within them using simultaneous optical trapping and total internal reflection fluorescence microscopy. This assay allows for nanoscale-level measurement of the forces and displacements generated by protein ensembles within active microtubule networks.

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

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