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

Summary

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

Abstract

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

Introduction

Over the past few decades, the importance of the mechanical properties of a cell’s environment has garnered increasing recognition in cell biology. Different tissues and extracellular matrices have different relative stiffnesses and, as cells migrate throughout the body, they navigate these changes, using these mechanical properties to guide them^{1,2,3,4,5,6,7}. Cells use the stiffness of a given tissue to inform their motile behavior during processes such as development, wound healing, and cancer metastasis. However, the molecular mechanisms that allow sensation of and response to these mechanical inputs remain largely unknown^{1,2,3,4,5,6,7}.

In order to study the mechanisms through which cells respond to physical environmental cues, the rigidity or stiffness of the substrate underlying adherent cells must be manipulated. In 2000, Chun-Min Lo, Yu-Li Wang and colleagues developed an assay⁸ whereby an individual cell’s motile response to changing mechanical cues could be directly tested by stretching deformable extracellular matrix (ECM)-coated polyacrylamide hydrogels on which the cells were plated. Cells exhibits a significant preference for migrating towards stiffer substrates, a phenomenon they dubbed “durotaxis.”

Since the original report in 2000, many other techniques have been employed for the study of durotaxis. Steep stiffness gradients have been fabricated by casting gels over rigid features such as polystyrene beads⁹ or stiff polymer posts¹⁰ or by polymerizing the substrate around the edges of a glass coverslips¹¹ to create mechanical ‘step-boundaries’. Alternatively, hydrogels with shallower but fixed stiffness gradients have been fabricated by a variety of methods such as gradients of crosslinker created by microfluidic devices^12,13 or side-by-side hydrogel solution droplets of differing stiffness⁸, or hydrogels with photoreactive crosslinker treated with graded UV light exposure to create a linear stiffness gradient^14,15. These techniques have been used to great effect to investigate durotactic cellular movement en masse over time. However, typically these features are fabricated in advance of cell plating and their properties remain consistent over the course of the experiment, relying on random cell movement for sampling of mechanical gradients. None of these techniques are amenable to observation of rapid changes in cellular behavior in response to acute mechanical stimulus.

In order to observe cellular responses to acute changes in the mechanical environment, single cell durotaxis assays offer several advantages. In these assays, individual cells are given an acute, mechanical stimulus by pulling the underlying substrate away from the cell with a glass micropipette, thereby introducing a directional gradient of cell-matrix tension. Changes in the motile behavior, such as speed or direction of migration, are then observed by live-cell phase contrast microscopy. This approach facilitates direct observation of cause and effect relationships between mechanical stimuli and cell migration, as it allows rapid, iterative manipulation of the direction and magnitude of the tension gradient and assessment of consequent cellular responses in real time. Further, this method can also be used to mechanically stimulate cells expressing fluorescent fusion proteins or biosensors to visualize changes in the amount, activity, or subcellular localization of proteins suspected to be involved in mechanosensing and durotaxis.

This technique has been employed by groups who study durotaxis^8,16 and is described here as it has been adapted by the Howe Laboratory to study the durotactic behavior of SKOV-3 ovarian cancer cells and the molecular mechanisms that underly durotaxis¹⁷. Additionally, a modified method is described for fabrication of hydrogels with a single, even layer of fluorescent microspheres near the cell culture surface; this facilitates visualization and optimization of micropipette-generated strain gradients and may allow assessment of cell contractility by traction force microscopy.

Protocol

1. Fabrication of Deformable Polyacrylamide Hydrogels with Embedded Fluorescent Microspheres

NOTE: Directions describe polymerization of a 25 kPa hydrogel that is 22 μm in diameter and approximately 66 μm thick. Each or all of these parameters can be modified and directions to do so can be found in Table 1 and in the notes¹⁷.

1. Activation of glass-bottom dishes or coverslips
   1. Prepare the bind silane working solution for activation of a glass-bottom imaging dish or a coverslip that fits into a live-cell imaging chamber. Mix 950 μL of 95% ethanol, 50 μL of glacial acetic acid, and 5 μL of bind silane (y-methacryloxypropyltrimethoxysilane).
      NOTE: Using a larger bottom coverslip compared to the top coverslip will give additional room to work when preparing the gel and will facilitate positioning the glass micropipette in later steps. Also, if using a coverslip rather than a glass-bottom imaging dish, clean the coverslip as described in the following section.
   2. Activate the surface of the glass for 20 s with a corona wand and immediately overlay 50 μL of the bind silane working solution. Allow the solution to dry for 10 min.
   3. Rinse two times with 95% ethanol, then two times with isopropanol and then allow the coverslips to airdry for approximately 20 min.
      NOTE: Activated glass can be stored for up to one week in a desiccator.
2. Cleaning top coverslips
   1. Clean 22 mm top coverslips by incubating in 2% HCl at 70 °C for 30 min, then wash in ddH₂O for 10 min two times.
   2. Incubate the coverslips in a solution of 2% cuvette cleaning concentrate in ddH₂O at 50 °C for 30 min, then wash in ddH₂O for 10 min two times.
   3. Incubate the coverslips in ddH₂O at 90 °C for 30 min, then in 70% ethanol at 70 °C for 10 min, and then air dry at 60 °C for a minimum of 2 h.
      NOTE: Cleaned coverslips can be stored indefinitely in a clean desiccator.
3. Fluorescent microsphere/bead deposition onto top coverslips
   1. Sonicate the stock solution of fluorescent microspheres for 1 h in an ultrasonic water bath. Make a working bead solution by diluting bead stock 1:200 in 100% ethanol and sonicate again for 1 h.
   2. 15 min before the bead solution has finished sonicating, thoroughly clean the coverslips by placing them vertically in a ceramic coverslip holder and treating with room-air plasma for 3 min in a tabletop plasma cleaner.
   3. To facilitate handling and prevent sliding of the coverslip during subsequent steps, place a piece of parafilm in a 60 mm Petri dish lid or a similar container. Place the coverslip in the stabilizer and lightly tap down, ensuring good contact between the parafilm and the coverslip.
   4. For a 22 mm coverslip, add 150 μL of the working bead solution to the top of the coverslip. Immediately aspirate the ethanol solution off from the side of the coverslip, leaving the beads on the coverslip. Allow the coverslip to airdry.
      NOTE: The amount of working bead solution added should be ~4 μL/cm² and can be scaled to accommodate any size coverslip.
4. Casting hydrogels with embedded fluorescent beads
   1. Prepare the hydrogel solution of acrylamide and bis-acrylamide. Mix the solution according to Table 1, then add 2.5 μL of 10% APS and 0.5 μL of TEMED. Mix well. Immediately move to the next step.
      NOTE: The hydrogel solution mixture can be altered to vary the Young’s modulus, or stiffness, of the hydrogel by changing the ratio of acrylamide to bis-acrylamide as shown in Table 1. These values have been verified for use in the Howe laboratory using atomic force microscopy but should be confirmed within one’s institution.
   2. Immediately after making the hydrogel solution, add a 25 μL drop to the activated side of the glass-bottom dish or bottom coverslip, then immediately place the bead-coated coverslip onto the solution, bead side down. Contacting the drop with the far side of the coverslip followed by slow lowering helps avoid trapping air bubbles within the hydrogel.
      NOTE: The height of the hydrogel should be well within the working distance of the objective lens to be used in the later experiment. A hydrogel height of 66 μm works well for most systems. The size of the hydrogel can be scaled by adding more or less hydrogel solution depending on the size of the coverslip. To calculate the appropriate volume of hydrogel solution, use the equation for the volume of a cylinder, V = πr²h where r is the coverslip radius and h is the desired hydrogel height. Typically, this calculation predicts with fair accuracy the actual height of the hydrogel, as measured by preparing a gel with bead-coated coverslips on both the top and bottom and using a confocal microscope to measure the distance between the two bead planes. However, it has been observed that the actual height of the hydrogel can deviate from this calculation by ± 20 μm (e.g., depending on the thickness and manufacturer of the top glass coverslip). Direct measurement of gel height using the method described above is recommended.
   3. Allow the gel to polymerize for 30 min, then remove the top coverslip gently with forceps. Adding 50 mM HEPES pH 8.5 to the dish can facilitate removal. Wash for 5 min in 50 mM HEPES pH 8.5 three times.
5. Hydrogel activation and extracellular matrix coating
   1. Activate the hydrogel surface by incubating in 0.4 mM Sulfo-SANPAH (sulfosuccinimidyl 6-(4'-azido-2'-nitrophenylamino) hexanoate) in 50 mM HEPES pH 8.5. Immediately expose to a UV arc lamp in an enclosed area.
      NOTE: Protect Sulfo-SANPAH from light prior to activation. For a 400 W lamp, position the gel 10 cm away from bulb within the light box and illuminate for 100 s. The Sulfo-SANPAH solution will change from bright orange to dark brown.
   2. Wash for 5 min in 50 mM HEPES pH 8.5 three times.
      NOTE: Hydrated gels may be stored at 4 °C for up to one week.
   3. Incubate the activated hydrogel in 20 μg/mL fibronectin in 50 mM HEPES pH 8.5 for 1 h at 37 °C.
   4. Aspirate the fibronectin solution and wash for 5 min in phosphate-buffered saline (PBS) three times. Sterilize the hydrogel and the lid of the dish for 15 min under UV light in a tissue culture hood with a low volume of PBS. Wash once in sterile PBS.
      NOTE: Other types of ECM protein can be used to coat the hydrogel including collagen and laminin.

2. Plating cells

1. Add 3 mL of media containing 21,000 cells to fill a 60 mm dish for a final cell density of ~1000 cells/cm². Adjust the seeding density as needed to prevent crowding and allow free movement of individual cells.
2. Allow the cells to recover at 37 °C for at least 4 h and for up to 18 h before imaging. Prepare for imaging by rinsing with imaging media two times before adding imaging media. Allow the cells to equilibrate for at least 30 min before imaging.
   NOTE: Screen media conditions in advance to determine conditions that will optimize migration in the cell line being used. For SKOV-3 cells, DMEM without phenol red, containing 20 mM HEPES and 12.5 ng/mL epidermal growth factor stimulates the most migration. Optimal conditions for Ref52 cells are Ringer’s Buffer with 10% fetal bovine serum (FBS) and 25 ng/mL platelet-derived growth factor.

3. Preparation of glass micropipette: pipette pulling and forging

1. Pull 100 mm long borosilicate glass micropipettes with a 1.0 mm exterior and 0.58 mm interior diameter in a two-step process to obtain a taper over 2 mm that reduces to ~50 μm in the first millimeter and extends to a long, parallel 10 μm diameter tube in the last millimeter.
2. Load pulled pipet into a microforge. Shape the pipet to have a 15 μm blunted tip that is enclosed at the very end of a 250 μm section bent at a ~35° angle from the rest of the pipet. The approximate diameter at the bend should be around 30 μm to lend strength to the tip.
   NOTE: Taper and tip dimensions can be adjusted to properly apply desired force (see step 5). Pulling micropipettes at 65 °C for the first step for 3 mm, and 60 °C for the second step produces the dimensions described in step 3.1. Results using different pipet pullers may vary.
3. Sterilize the micropipette in 70% ethanol before use.

4. Positioning the micromanipulator and the micropipette

1. Remove the dish lid and load the dish onto the microscope stage and center. Use a 10X or similarly low magnification objective. Cover the media with mineral oil to prevent evaporation of the media.
2. Inserting pulled pipet
   1. Insert the pulled pipet into the micropipette sheath, pointing the hook down toward the dish. The tip of the hook will be the lowest point when lowered to the gel.
   2. Insert sheath into micromanipulator and adjust until the tip of the pipet is centered over the objective lens in both X and Y directions.
   3. Lower the pipet using coarse manipulator until it just touches the surface of the liquid.
3. Using phase contrast or brightfield, focus the microscope on the bead layer at the top of the gel. This will be the reference plane.
4. Ensuring that the objective is in no danger of hitting the sample or stage, bring focus above the gel to find the tip of the micropipette, using small adjustments of the coarse manipulator in the X and Y directions to cast shadows on the focal plane. Only lower the micropipette when certain that the very tip of the pipet is in the field of view.
5. Ensure that the blunted tip of micropipette is pointing down by rotating the pipet in the sheath or rotating the sheath in the micromanipulator until the tip is perpendicular to the focal plane. Repeat steps 4.4 and 4.5 as needed. Focus on the tip of the pipet.
6. Focus back down to the top bead layer of the gel to gauge how far the pipet is from the gel surface. Focus back up to a plane that is part way between the gel and the tip of the pipet. Slowly lower the pipet to reach the intermediate focal plane.
7. Repeat step 4.6 until very faint shadows from the tip of the micropipette can be appreciated when focusing on the hydrogel. Increase to the next highest magnification.
8. Lower the micromanipulator until shadows and refractions of the very tip of the micropipette can be appreciated within the bead layer focal plane.
9. Increase the magnification to that which will be used in the experiment. Lower the pipet until it hovers just above the surface of the hydrogel.

5. Calibrating the micromanipulator and force generation

1. In phase or brightfield, lower the hovering micropipette to touch the surface of the hydrogel. Observe how the pipet looks upon contact with the hydrogel. Continue to lower micropipette in Z until adjustments in X and Y cause pulling and deflection of the hydrogel in those directions. Use the microspheres or nearby cells as fiduciary marks.
   NOTE: If the micromanipulator is attached to the phase condenser arm or the bench and not the sample stage itself, always disengage the gel before moving the stage to avoid breaking the pipet or disturbing cells. If the pipet breaks, go back to step 3 and step 4.
2. Find an area devoid of cells to engage the gel. Pull it in all directions and get comfortable with the way micromanipulation translates to deformation of the gel.
3. Take fluorescent images of the bead field with no manipulation, with the pipet engaging the gel, and with the engaged pipet pulling the gel. Repeat this several times taking good notes regarding the tick marks on the micromanipulator, the way the pipet tip looks in phase or brightfield at each stage of pulling, and the distance the tip moves using that manipulation.
4. Use ImageJ as previously described^16,17 to calculate relative bead displacements and force applied to the beads by comparing the null bead field to the bead field without pipet engagement, the bead field with the gel engaged, and the pulled gel.
5. To fine tune the tensional stimulus, compare force application using differing micropipette tip dimensions, distances from the cell, or distance pulled by the micromanipulator from initial point of touchdown. The effect of the micropipette tip dimension on force application gives great flexibility to the user but also demonstrates the need to generate force maps for new micropipettes, even when the dimensions and shape closely resemble previously calibrated tips.

6. Conducting the durotaxis assay

1. Before performing the experiment, practice engaging the gel near a cell and observe the deformation of the cell when the micromanipulator is repositioned.
2. Monitor a group of cells that have clear polarity and appear to be moving for 30 min to identify cells that are moving in a directed manner.
3. Choose a cell that is moving in a single, clear direction and monitor it at the desired frame rate for an additional 30 min.
4. If determination of forces exerted on the cell or tension exerted by the cell is desired, capture bead field images at each acquisition. If the cell changes its course of direction during monitoring, choose a different cell to monitor as this will make it difficult to determine the effect of stimulation.
5. Engage the hydrogel approximately 50 μm away from the cell. Position the pipet in front of the near side of the leading edge and move the micromanipulator such that the gel is deformed orthogonally to the cell’s direction of travel. Observe the cell over time as it responds to the acute, local gradient of stiffness.
   NOTE: The timing provided here is effective when monitoring SKOV3 or Ref52 fibroblasts, however, the interval and overall time course should be adjusted to suit the cell type and biological event being observed. If pairing with fluorescence microscopy, pause fluorescent acquisition immediately before step 6.5., use phase contrast or brightfield to position micropipette and pull, and restart fluorescent acquisition immediately after.
6. If the pipet slips or if the gradient is otherwise relaxed or released, find a new cell by repeating steps 6.2 and 6.3.

7. Determining durotactic migration response

1. Using ImageJ¹⁹ or another image analysis program, calculate the turn angle by drawing a line between the middle of the leading edge of the cell at 0 min and 30 min post monitor (reflecting the cell’s original trajectory) and another line between the middle of the leading edge just before and 80 min after stimulation and measuring the angle between these two lines.

Representative Results

By preparing micropipettes (Figure 1) and normalizing the force generation of the pulls (Figure 2 and Figure 3) as described above, optimal durotactic conditions have been identified for multiple cell lines. Using this technique, as outlined in Figure 4, both SKOV-3 ovarian cancer cells¹⁷ and Ref52 rat embryonic fibroblasts (Figure 5) move toward increased stiffness in gradients applied by a glass micropipette. In addition to durotaxis, this method can be used to study dynamic signaling events using fluorescent biosensors and markers. For example, the structure of and signaling within focal adhesion structures can be observed upon durotactic stimulation. Vinculin tension sensor (VinTS) is a FRET-based biosensor which localizes to focal adhesions, allowing for fluorescent observation of focal adhesion dynamics and measurement of changes in tension within those structures¹⁹. Ref52 cells transiently expressing VinTS on 125 kPa polyacrylamide gels show the formation of focal adhesions in the direction of stretch over time period of 40 min (Figure 6A). FRET analysis²⁰ reveals that vinculin localized to focal adhesions experiences an immediate change in tension when presented with acute durotactic stimulation (Figure 6B) expanding the utility of this assay to the observation of subcellular signaling events in response to durotactic stimulation.

Figure 1. Diagrams of typical pulled (A) and forged (B) micropipettes. (A) Micropipettes are pulled using a two-step protocol to achieve a taper from 1 mm to 10 μm over 2 mm. (B) Micropipettes are then loaded into the microforge and their tips are bent, enclosed, and shortened so that the last 250 μm of the micropipette is bent at a ~35° angle and tapers from ~30 μm to a rounded tip that measures ~15 μm. Please click here to view a larger version of this figure.

Figure 2. Improved bead field after ethanol coating as compared to traditional method using poly-L-lysine. Representative hydrogel bead fields from poly-L-lysine and ethanol (EtOH) evaporation coverslip coating methods using yellow-green, red, and dark-red fluorescent beads. Scale bar: 25 μm. Please click here to view a larger version of this figure.

Figure 3. Generation of force map for an example durotactic stretch. (A) Position of fluorescent microspheres before and after (pseudo-colored green and red, respectively) deforming the hydrogel with a micropipette (located beyond the right edge of the panel). Scale bar: 25 μm. Displacement vectors (B) and displacement heat map (C) between null and pulled bead fields generated by Traction Force Microscopy plugins in ImageJ highlight the gradient of bead deflection and hydrogel strain. Please click here to view a larger version of this figure.

Figure 4. Schematic of durotaxis assay and determination of deflection angle. (A) A cell is observed for at least 30 min to determine its original trajectory. (B) The micropipette is positioned orthogonally to the cell’s trajectory, 50 μm from the cell edge. The hydrogel is engaged by the micropipette such that moving the micropipette will exert force on the surface of the hydrogel. (C) The micropipette is pulled an additional 20 μm away from the cell, orthogonal to the cell’s trajectory which creates an acute, local gradient of tension (denoted in blue) which increases toward the micropipette. (D) The cell is observed over time as it navigates the applied gradient. (E) In ImageJ or an image analysis program, the original trajectory (dashed line) is marked by a line drawn from the middle of the cell through the center of the leading edge in the first frame. The final trajectory (solid line) is marked by a line drawn after the cell is allowed to navigate the applied tension gradient. The angle between these two lines toward the stimulus is termed “turn angle,” marked here by θ. Please click here to view a larger version of this figure.

Figure 5. Rat Embryonic Fibroblasts move toward regions of increased substrate stiffness in durotaxis. Time course showing durotactic movement of a Ref52 cell 10 min before the pull (panel 1), 1 min before the pull (panel 2), at the time of pull (panel 3), and 1 h after pull (panel 4). Arrow indicates direction of stretch. Scale bar: 50 μm. Please click here to view a larger version of this figure.

Figure 6. Protein localization and activity during durotactic stimulation using fluorescent markers or biosensors. Ref52 cells transiently expressing Vinculin Tension Sensor (VinTS)¹⁹ migrating on 125 kPa polyacrylamide gels are presented with acute durotactic stimulation. (A) After stimulation, new focal adhesions form in the direction of stretch as cells re-orient along the stiffness gradient. For two 10 min periods, starting 20 min before mechanical stimulation and 21 min after stimulation, cell morphology (top) and focal adhesion formation (bottom) were monitored. Red color indicates the first timepoint within the time period and green color indicates the timepoint 10 min later. New focal adhesions formed within the 10 min period are shown in green. Before stimulation, new focal adhesions form in the direction of travel. After stimulation, new focal adhesions form in the direction of stretch. Arrow indicates the direction of stretch. Arrowheads indicate areas with focal adhesions formed over that 10 min period. Scale bar: 25 μm. (B) FRET analysis of VinTS fluorescence indicates a change in tension within focal adhesions proximal to durotactic stretch. Outline of cell membrane before and after stretch highlight deformation of cell upon stimulation. Arrowheads indicate examples of focal adhesions experiencing changes in FRET ratio upon stretch. Scale bar: 10 μm. Please click here to view a larger version of this figure.

Desired hydrogel stiffness	3 kPa	25 kPa	125 kPa
7.5% Acrylamide	100 μL	100 μL	160 μL
0.5% Bis-Acrylamide	10 μL	100 μL	100 μL
ddH2O	287 μL	197 μL	137 μL

Table 1. Acrylamide gel solutions.

Discussion

Demonstrated here is a repeatable, single-cell durotaxis assay that allows assessment of a cell’s ability to alter its migration behavior in response to acute mechanical cues. This technique can also be used in combination with fluorescence microscopy and appropriate fusion proteins or biosensors to examine subcellular signaling and cytoskeletal events within seconds of mechanical stimulation or over a longer timescale during durotactic movement. Understanding a cell’s relationship to its environment involves the study of the impact of both the chemical and mechanical aspects of that environment. Though potentially difficult to master, this durotaxis assay can be widely used to understand the cellular response to changes in its mechanical microenvironment.

Significance with respect to existing methods

As mentioned before, this micropipette-based method of durotactic stimulation is highly manipulable, allowing a high degree of spatiotemporal control over mechanical stimuli, a major advantage over other techniques, such as pre-formed linear or step-gradients of rigidity. The magnitude and direction of the imparted strain gradients can be visualized by tracking the displacement of fluorescent beads embedded in the hydrogel, near the cell culture surface.

Restricting these fiducial markers to a single layer just below the culture surface increases the accuracy of this tracking. Microspheres located below the plane of deflection (imparted either by the micropipette or, for traction force microscopy, by cellular contractility), as would occur with mixing the microspheres evenly throughout the hydrogel, will move less than in-plane microspheres, which can lead to underestimation of applied forces. Also, this modification is easier to perform and more reliable than methods in which beads are overlaid in an extremely thin layer of polyacrylamide cast on top of a pre-formed gel²¹ or brought to the hydrogel surface by gravity-assisted settling²² and produces a more even dispersal of beads across the hydrogel than previously described methods^17,23.

Modification and future applications

The specifics of this assay can be modified to best suit the cell line of interest. For example, a variety of extracellular matrix molecules (e.g., collagen I, collagen IV, laminin) or other adhesive ligands can be used to functionalize the hydrogel. Also, the starting stiffness of the hydrogel can easily be raised or lowered by tuning the ratio of acrylamide to bis-acrylamide (see Table 1). By changing the dimensions of the micropipette tip and the magnitude of the pull, this assay can be optimized to impart a repeatable and effective durotactic stimulus for the cell type in question.

Critical steps and troubleshooting

Only cells following a steady, linear trajectory of migration prior to hydrogel manipulation should be stimulated to ensure that changes in trajectory are due to the mechanical stimulus and not random fluctuation. Care must be taken to fabricate a glass micropipette that engages the hydrogel surface without slipping but does not tear the gel when pulled. It is important to apply a steady, constant stretch to the hydrogel during the course of the experiment to obtain clean results meaning that the user should be practiced at placing and moving the micropipette before encountering a cell. Any unintentional movement of the micropipette that leads to changes in the tension gradient could affect the cell’s ability to durotax. Similarly, manipulation of the gel should be practiced with each new micropipette that is forged as slight changes in pipet shape can cause the pipet/gel interaction to vary.

Failure to position the micropipette within the microscopic field of view before increasing magnification can lead to the accidental breakage of the fragile glass tip. Ensure that the height and X-Y position of the tip is known before lowering the micropipette with the micromanipulator. Always monitor the position of the micropipette to reduce the risk of breakage. It is recommended that the magnification is decreased back down to 10X for each new micropipette loaded into the micromanipulator as slight movements of the micromanipulator and pipet sheath can lead to large apparent changes in the position of the newly loaded micropipette.

Before finding cells to observe, it is important to first test the micropipette to confirm that it will engage the hydrogel as expected and that it is suitable for applying the desired stretch. Finding tip dimensions that suit the experiment and cell type is critical to success in applying durotactic stimulation. The end of the micropipette should be rounded enough so that it does not break through the gel, but not so rounded that it fails to grip it. If the pipet does not pull gel effectively, it may be sliding along the surface. The shape of the micropipette tip may be too rounded to properly engage with the gel surface. The dimensions at the very tip of the micropipette should be adjusted until firm, steady contact can be achieved consistently. In some cases where the micropipette is slipping across the gel surface, it may be necessary to lower the micropipette further into the hydrogel to gain more traction. If the micropipette tears through gel, the tip may be too fine or too sharp. Gel tearing may also indicate too much force is being applied while pulling. The micropipette should be raised slightly to reduce gel deformation and pulling shorter distances.

Often, if the cell or the edge of the cell is pulled out of focus by the micropipette, the tip of the micropipette is engaged too close to the cell or the stretch is too forceful. Move the micropipette further from the cell, only slightly deforming the cell in the X-Y planes. Moving the cell out of focus will not only make cellular events impossible to monitor and cause optical aberrations, but it will cause the cell to experience more stimulation than the 2-dimensional tension gradient intended.

Most importantly, it is critical to record and analyze only responses that have consistent durotactic manipulation with minimal human error. If the lowering of the tip is imprecise or if the micropipette is repositioned, the results of the experiment will be clouded. Since this assay is complex and many steps are prone to error, care must be taken at every step to avoid unintentional changes in the stimulation of cells. Failure at any step can lead to inconsistent stretch application and unreliable results.

Limitations

There are limitations to this technique that should be considered. Most prominently, accurate forging and manipulation of the glass micropipettes can present a steep learning curve for new users. Additionally, the position and magnitude of the hydrogel pull must be optimized for different cell lines. Examining fluorescent bead displacements before and after hydrogel manipulation can help with this aspect of the technique. Also, while the technique allows high spatiotemporal observation of durotactic behavior in individual cells, this makes it a low-throughput assay. It is therefore important to point out that this assay can also be complemented by other techniques with lower manipulability but higher throughput, such as using hydrogels with pre-formed gradients of rigidity, to analyze the durotactic behavior of larger populations of cells at once. In summary, the high degree of spatiotemporal control of mechanical cues afforded by the single-cell durotactic assay make it very useful for parsing the molecular mechanisms contributing to the durotactic behavior of many different cell types under many conditions.

Materials

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