Traction Microscopy Integrated with Microfluidics for Chemotactic Collective Migration

Collective cell migration is often guided by the gradient of signaling molecules. In this video, we demonstrate how to quantify cellular forces during collective migration guided by biochemical gradient. So, we integrate the microfluidic chip with traction microscopy, then we use the microfluidic chip to generate biochemical gradient, and we use the traction microscopy to measure cellular force.

Dr.Hwanseok Jang, a postgrad in my laboratory, will demonstrate the procedure. Prepare the PDMS solution by mixing the base elastomer and curing agent at a ratio of ten to one. Place 15 milliliters of the base elastomer in a 50 milliliter conical tube, and add 1.5 milliliters of curing agent.

Prepare two of these tubes. Vortex the PDMS mixture for five minutes before centrifuging at 196 time g for one minute to remove bubbles. To fabricate the PDMS stencil, pour approximately one milliliter of the PDMS mixture on the wafer while avoiding SU-8-patterned regions so that the PDMS touches the side of the SU-8 pillars but not the top of the SU-8 pattern.

Place the wafer on a flat surface for over 30 minutes at room temperature, then cure the PDMS in a dry oven at 80 degrees Celsius for over two hours. Carefully peel off the PDMS from SU-8 mold, trim the thin PDMS membrane using a 14 millimeter hollow punch. Remove the dust on the surface of the PDMS pieces using sticky tape before autoclaving the PDMS stencils.

To fabricate PDMS microchannel, pour approximately 30 milliliters of PDMS mixture over the SU-8 mold. Degas for 30 minutes in a vacuum chamber, and then cure the PDMS in a dry oven at 80 degrees Celsius for over two hours. Carefully peel off the PDMS from SU-8 mold, and cut the PDMS to a size of 24 millimeters by 24 millimeters.

In each PDMS block, create one outlet and three inlets using a one millimeter biopsy punch. Manufacture and silanize glass slides as described in the text protocol. Cover the edged surface of the bottom glass with 100 microliters of the bind-silane solution.

After leaving the glass at room temperature for one hour, rinse the glass three times with deionized water. Then let the glass dry at ambient air temperature or by blowing compressed air. Prepare the polyacrylamide gel solution as described in the text protocol.

Then transfer for 10 microliters of mixed gel solution onto the rectangular microwell and place a circular cover slip on top. Fix the assembly of custom glass, gel solution, and cover slip on the cover of a six-well plate with sticky tape and flip it. Then centrifuge for 10 minutes at 96 times g to bring fluorescent particles to the top layer of the polyacrylamide gel.

Remove the assembly from the centrifuge and place it on a flat surface with the cover slip facing down. After 30 minutes, flip the assembly and place it in a 35 millimeter Petri dish. Fill the dish with two milliliters of deionized water.

Using forceps, gently remove the cover slip by sliding it to one side. To code collagen on the polyacrylamide gel, dissolve one milligram per milliliter sulfo-SANPAH in a warm 50 millimolar HEPES buffer. Drop 200 microliters of the solution onto the gel surface and activate by UV light for 10 minutes.

Following UV activation, rinse the gel two times with 0.1 molar HEPES buffer, and then once with PBS. Code the polyacrylamide gel with collagen solution at four degrees Celsius overnight. On the following day, wash the gel three times with PBS.

Immerse the autoclaved PDMS stencil in F-127 solution. Keep it in a 37-degree Celsius incubator for one hour. Wash the PDMS stencil with PBS three times, and remove liquid from both the PDMS stencil and polyacrylamide gel.

Then place the PDMS stencil on the polyacrylamide gel, and add PBS to the stencil. Remove bubbles in the holes of the PDMS stencil by pipetting gently. After removing bubbles, clear the PBS from the surface of the PDMS stencil.

Now add 200 microliters of the cell solution onto PDMS stencil and place the gel in an incubator for one hour so that the cells attach to the polyacrylamide gel. Following incubation, gently wash off the cell solution with cell culture media and add more cell culture media. Remove the PDMS stencil, check the formation of cell islands under a microscope.

Next, treat the surface of the PDMS microchannel with oxygen plasma for 30 seconds. After removing any fluid on the polyacrylamide gel-filled bottom glass, place the PDMS microchannel on top of the bottom glass, and put the assembly on the custom glass holder. Fill the microchannel with cell culture medium.

To prepare the inlet tubing of the integrated microfluidic system, connect a trimmed needle and a 30-centimeter mini volume line with a three-way stopcock. Prepare three of these arrangements. For outlet tubing, connect a trimmed needle and a 75-centimeter mini volume line with a three-way stopcock.

Fill the tubing lines with the medium that has been preheated for one hour. Prepare reservoirs by removing plungers from syringes and connecting inlet tubing lines. Plug the needle connectors of each tubing line into the three inlets and one outlet of the microfluidic device.

Fill each reservoir with three milliliters of fresh medium or conditioned medium. For the gradient test, fill the left inlet reservoir with 20 nanograms per milliliter hepatocyte growth factor, or HGF, in the cell culture medium. For visualization of the concentration gradient, add 200 micrograms per milliliter of fluorescent dye to the left inlet reservoir.

Connect the outlet tubing line to a syringe pump. Place the integrated microfluidic system on the stage of a conventional epifluorescent microscope. Take images every 10 minutes for up to 24 hours using an automated microscope housed in an incubator.

At each time point, take a set of images using a 4X objective lens in three different channels, including a phase image to visualize cell migration, a green fluorescent image to visualize fluorescent beads embedded in the gel, and a red fluorescent image to visualize the concentration gradient of a chemical. After taking time-lapse images, infuse 0.25%Trypsin-EDTA solution into microchannels to detach cells from the polyacrylamide gel. Upon completely removing cells from the gel, take a green fluorescent image to be used as a reference image for traction microscopy.

Proceed to data analysis as described in the text protocol. To measure contractile force and intercellular stress, traction microscopy and monolayer stress microscopy were combined with the microfluidic channel. The maps were plotted in radio coordination with outward traction using warm color and inward traction using cold color.

At time zero, all islands showed similar traction distributions with strong inward traction on the edge and fluctuation within the island. After 10 hours of applying the HGF gradient, while the degree of island expansion was different in each column, the traction distributions were largely similar to time zero. The average traction did not change for 10 hours.

When calculating the monolayer stress, however, each column showed different trends. Where HGF concentration was low, the average tension within islands was maintained around 200 pascals throughout a 10-hour period. Where HGF concentration was high, the average tension within islands gradually decreased from 230 pascals to 100 pascals over 10 hours, as previously shown.

Where the HGF concentration was high on the left half and low on the right half of the island, the average tension was maintained around 150 pascals. Filling the microfluidic channel must be gentle. It is important to remove bubbles trapped in the channel while simultaneously not disrupting micro parent cell islands.

By using this integrated system, one can investigate it how cell collective respond mechanically to various chemical gradients, such as chemo attractants or growth factors. This platform will help us further explore the mechanics of collective cell migration under chemical gradients, which is the key to understand regeneration, cancer metastasis, and development.