Measurement of Force-Sensitive Protein Dynamics in Living Cells Using a Combination of Fluorescent Techniques

This method can answer key questions in the field of mechanobiology, such as how forces are transmitted and sensed within cells as well as between cells and their environment. The main advantage of this technique is it allows access to the molecular scale information regarding how proteins respond to mechanical forces inside the native context of the living cell. Though this method has been applied specifically to studying the focal adhesion protein vinculin, it can also be applied to other mechanically-sensitive proteins, such as talin, cadherins or nesprin.

Visual demonstration of this method is critical, as proper imaging setup is difficult but necessary for the success of the analysis. Begin this procedure with stable expression of the tension sensor construct in the desired cell type, as detailed in the text protocol. To prepare substrates for cell seeding, acquire four 35 millimeter glass-bottomed dishes.

Working in a cell culture hood, in a 15 milliliter conical tube, make four milliliters of 10 micrograms per milliliter fibronectin in sterile phosphate-buffered saline solution, or PBS. Gently invert the tube once to mix and let the solution sit for five minutes in the cell culture hood. Then pipette one milliliter of fibronectin solution onto each glass-bottomed dish.

Leave the fibronectin solution on the dishes for one hour at room temperature or overnight at four degrees Celsius. Then aspirate the fibronectin solution and rinse once with PBS. Leave one milliliter of PBS on the dishes until the cells are ready for seeding.

After counting the cells, seed 30, 000 cells onto each fibronectin-coated glass dish with the appropriate complete media for a final volume of 1.5 milliliters. Allow the cells to spread for four hours following seeding. At two hours of spreading, aspirate the growth media and rinse once with imaging media.

Leave 1.5 milliliters of the imaging media. Warm up the microscope as described in the text protocol before beginning calibration. To calibrate the FRAP laser, first open the laser configuration window.

Set Illumination setting to the appropriate FRAP Illumination settings for laser exposure to the sample. Then set Illumination setting to the illumination settings appropriate for imaging only the acceptor fluorophore. Select the objective to calibrate under Coordinate system setting.

Uncheck Manually click calibration points and check Display images during calibration. Set the Dwell time to 10, 000 microseconds and the Number of pulses to 100. Now place the calibration slide, made of ethidium bromide sealed between a glass slide and a cover slip, into the stage adapter with the cover slip side down.

Use the acceptor illumination settings to focus on the surface of the slide, identifiable as the focal plane with the brightest signal. Small defects in the coating will be visible to aid in focusing. Move the slide to an area with uniform fluorescence across the imaging plane.

Click on Create Setting for the first calibration or Update Setting for subsequent calibrations. The software will initialize the calibration process, automatically bleaching and detecting the position of the bleached point. Ensure a successful calibration by assessing the final image, which will be a three-by-three grid of bleached points that should be evenly distributed and in focus.

Save the calibration image for future reference. Remove the calibration slide and safely store. Calibration should be performed before beginning each experiment but does not need to be performed between samples.

Prepare the microscopy setup for live cell imaging, preferably with a heated stage and objective, as well a carbon dioxide control. Allow to equilibrate for 20 minutes. Now place one of the generated samples of cells expressing the tension sensor into the microscope holder for imaging.

Allow the cells to equilibrate for 10 minutes. To ensure the health of the imaged cells, maintain the temperature at 37 degrees Celsius in the imaging chamber. Use a peristaltic pump to pass humidified 5%carbon dioxide over the samples at 15 milliliters per minute to maintain the pH.

Open the Multi-Dimensional Acquisition, or MDA, tool and set it up with FRET imaging parameters, including the different filter sets. Save this MDA to the Experimental Folder with the name of MDA_FRET_date. Set up another MDA with the FRAP imaging parameters, including the different filter sets, the Timelapse settings and the Journal to pulse the laser after pre-bleach acquisition.

Save this MDA to the Experimental Folder with the name of MDA_FRAP_date. In the toolbar at the top of the screen, select Journal, Start Recording. Open the MDA window, load the MDA_FRET_date state and press Acquire.

Then load the MDA_FRAP_date state and press Acquire. At the end of the acquisition, in the toolbar at the top of the screen, select Journal, Stop Recording. Save this Journal to the Experimental Folder with the name of FRETFRAP_date and add it to a toolbar for easy access.

Close the MDA window. Navigate the sample using the image acquisition under Acquire, Acquire with minimal exposure time and neutral density filter to identify the cells of interest. Set continuous autofocus by navigating to Devices, Focus.

Manually focus on the sample until reaching the correct imaging plane. Click Set Continuous Focus and wait for the system to adjust. Once the system adjusts, click Start Continuous Focusing.

Then find a cell expressing the tension sensor with clear localization to a structure of interest and snap an image. Draw a rectangular region of interest, or ROI, to highlight where to bleach. Store this ROI location.

Now initialize the FRETFRAP_date journal, which will begin the acquisition of FRET images, followed by the initialization of the FRAP timelapse. Repeat these steps until 10 to 15 image sets are acquired. Then analyze the FRET-FRAP data as detailed in the text protocol.

Shown here are representative results for FRET imaging of the vinculin tension sensor, following segmentation and masking to isolate focal adhesions. Spacial variation of vinculin load can be seen across the cell. FRAP imaging and analysis can be affected by focal adhesion stability.

A focal adhesion that is translocating rapidly during the imaging period can be difficult to accurately track. Often this is indicated by a FRAP curve that contains discontinuities and in uninterpretable. For a wild-type vinculin, there is an inverse correlation between protein load and turnover rate, indicating that vinculin is stabilized by force.

Introducing a mutation in vinculin to effect its binding to talin results in a reversal of the correlation, indicating that vinculin that is unable to bind talin is destabilized by force. While attempting this procedure, it’s important to remember to prepare samples properly, ensure that cells are expressing the sensor at endogenous levels and choose imaging parameters carefully. Following this procedure, other methods like immunofluorescence, measuring cellular scale dynamics or challenging cells with various inhibitors can be performed in order to answer additional questions like how the protein of interest interacts with other subcellular components to coordinate response to force.