High-throughput Flow-cytometry Measurement of Cellular Mechanotype Based on Rupture and Delivery of DNA Tension Probes into Cells

Our lab generally studies molecular mechanisms cells use to convey mechanical stimuli. Recently, we have been developing high-throughput methods to measure the forces cells exert on their surroundings to bring mechanobiology into the omics era. Studies implicate mechanical forces in how T cells are activated by the correct antigen and how stem cells differentiate into specialized cell types.

Recently, targeting processes underlying dysregulated cell mechanics has led to the development of mechanotherapeutics as an avenue to treat disease. DNA duplex and hairpin sensors measure extracellular forces, while genetically encoded sensors allow intracellular measurements. Nearly all require high-end microscopes.

Traction force microscopy can measure the force the cells exert on their surroundings, but it, too, involves high-resolution imaging. We address three challenges in the field in this work, one, using flow cytometry for force measurement instead of imaging, two, employing HOH tag technology to attach ligands and antibodies to DNA probes, and three, utilizing standard cell culture plates without special surface treatments. How changes of cell generated forces can be used as a prognostic marker for disease state, and are these forces contributing to, or are they a secondary effect of said disease state?

To begin, assemble duplexes in a 1.1 to one molar ratio of anchor strand with quencher to ligand strand with fluorophore. Prepare the duplexes at a final concentration of 40 micromolar ligand strands in the solution. For annealing the assembled duplexes, incubate them at 98 degrees Celsius for five minutes.

Then, let the mixture cool at room temperature for one hour away from light. React the annealed duplexes with HUH using the concentration of the least abundant strand from the annealing step, which in this case, is the ligand strand. Mix the duplexes with HUH in a one to two molar ratio at a final concentration of 10 micromolar duplex and 20 micromolar HUH.

Incubate the reaction at 37 degrees Celsius for 30 minutes in a thermal cycler or a light-protected heat block. After incubation, remove the reaction from heat and use a benchtop mini centrifuge to spin down the now-assembled tension gauge tethers, or TGTs. When the TGTs adhere adequately, wash the wells with desired cells three times with PBS.

Follow this with two washes using serum-free media and leave the media in the wells after the final wash. Add 200 microliters of PBS to the peripheral wells to mitigate edge effects. Transfer the plate to a 37 degrees Celsius incubator to prewarm the plate.

Then, retrieve the cells of interest at approximately 80%confluency. Aspirate the media, then wash the cells with PBS and incubate the cells with trypsin at 37 degrees Celsius until the cells dissociate, which takes approximately five to 10 minutes. Once the cells are dissociated, add the original volume of complete cell media used to maintain the cells.

Resuspend the cells using a serological pipette and transfer the cell suspension into a 15 milliliter conical tube. Centrifuge the cells at 300 G for four minutes. Aspirate the media and resuspend the pellet in PBS.

After repeating the centrifugation, resuspend the cells in five milliliters of serum-free media and centrifuge again. Aspirate and resuspend the cells in approximately two milliliters of serum-free media. Determine the density of the resuspended cells to adjust the concentration as desired.

Optionally, if testing drug effects, add the appropriate concentration of the drug to the cells and return the cells to the incubator for pre-incubation time before proceeding. Now, retrieve the original plate from the incubator and remove the residual media left from the wash in the experimental wells. Wash the wells once with the desired media and pipette 100 microliters of the resuspended cells into the desired wells.

Transfer the plated cells to the incubator for 90 minutes. Following incubation, remove approximately 125 to 170 microliters of volume from each well. If required, wash the cells gently with 200 microliters of warmed PBS.

Then, incubate the cells in each well with 50 microliters of trypsin until the cells begin to dissociate, which takes approximately five to 10 minutes. Next, add 160 microliters of freshly prepared flow buffer to each well. Gently pipette up and down to dissociate the cells.

Transfer 196 microliters of this cell suspension into PCR tubes, add four microliters of propidium iodide, and gently mix. Finally, place the tubes in an ice bucket with a lid blocking any light and measure fluorescence intensity using a flow cytometer. Flow cytometry gating confirmed successful live single cell isolation, ensuring accurate mechanotype analysis.

A negative control gating, set at the 99th percentile of Cy5-A fluorescence, provided a baseline for the comparison of experimental conditions. CHO-K1 cells treated with Y27632 showed a significant decrease in the percentage of cells exceeding the negative control gate on shearing TGTs, while unzipping TGTs exhibited only a modest and statistically insignificant decrease. Unzipping TGTs consistently resulted in a higher percentage of cells exceeding the negative control compared to shearing TGTs, confirming their lower force requirement.