Imaging Molecular Adhesion in Cell Rolling by Adhesion Footprint Assay

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

This protocol helps establish the connection between molecular force and cell rolling behavior besides allowing to spatially map and quantifying rolling adhesion at the molecular level. This technique allows to study individual adhesion events during the actual cell rolling allowing us to measure the physiological relevant molecular adhesion force directly. Surface preparation using the appropriate concentration and incubation time in each step is crucial to achieve high-quality surface functionalization.

The quality of bioconjugations and proper conditions are also crucial. Visual demonstration is critical to help researchers replicate this protocol. In particular, the steps such as flow chamber assembly and post-processing images will benefit greatly from a visual demonstration.

Begin by thinly spreading a small amount of epoxy on both sides of the double-sided tape with a razor blade. Using a laser, cut the epoxy-coated tape to create four channels. Create the flow chip by sandwiching the epoxy tape between a four-hole slide and PEG coverslip.

Using a pipette tip, apply gentle pressure along the length of the channels to create a good seal, then cure the epoxy for a minimum of one hour. Align the chip so that the opening of each channel is positioned at the centers of the adapter. Then place two transparent acrylic spacers on top of the chip.

Apply firm pressure in the middle of the block and screw at the ends of each spacer. Screw the inlets into the threaded holes on the other side of the bracket and monitor the sealing condition through the transparent acrylic block. Flow 200 microliters of wash buffer into the chamber to check for leakage.

If bubbles form in the channel, aggressively push an additional 200 microliters of wash buffer to remove the bubbles. Add 40 microliters of BSA to the flow chamber to prevent nonspecific binding and incubate for 10 minutes in the humidity chamber. After incubation, add 40 microliters at Tween 20 to the flow chamber and again incubate for 10 minutes to further reduce the nonspecific binding.

Then wash the channel with 200 microliters of wash buffer to remove all passivation agents. For chamber surface functionalization, add 40 microliters of Streptavidin to the flow chamber and incubate for 20 minutes, then wash the chamber with 200 microliters of wash buffer. Now, add 40 microliters of hybridized protein G-TGT to the flow chamber and incubate for 20 minutes.

After washing with wash buffer, add 40 microliters of protein G-TGT top strand and incubate for 20 minutes to complete any unhybridized TGT bottom strand on the surface and then wash with wash buffer. Finally, add 40 microliters of P-selectin-Fc to the flow chamber and incubate for 60 minutes before washing with wash buffer. Fill a five milliliter glass syringe with the rolling buffer and tap the sides of the syringe to dislodge and push the bubbles out as they float towards the tip.

After inserting a sterile needle into a 200 millimeter piece of polyethylene tubing, connect the needle to the glass syringe. Fix the syringe onto the syringe pump and tilt the syringe pump such that the plunger side is elevated to prevent air bubbles from entering the channel. Insert the end of the tube into the flow chamber inlet.

Insert one end of another 200 millimeter piece of the polyethylene tubing into the outlet and submerge the other end in a waste beaker. Take one to two milliliters of the cell suspension sample and centrifuge to pellet the cells. Remove the medium and gently resuspend the cells in 500 microliters of the rolling buffer.

Carefully disconnect the tubing from the inlets and the outlet and pipette 40 microliters of the cell suspension into the flow chamber. Reconnect the tubing as described previously ensuring bubbles are not introduced into the flow channel. Begin cell rolling experiment by starting the syringe pump at desired flow rates.

Observe cell rolling using a dark field microscope with a 10X objective. Once the experiment is completed, remove the cells from the channel by infusing the rolling buffer at 100 milliliters per hour until the surface is cell free. For imaging the local tracks by DNA-PAINT, add 40 microliters of DNA-PAINT imager strand prepared in DNA-PAINT buffer to the channel.

Perform total internal reflection fluorescence microscopy using the conditions mentioned in the text manuscript. Localize and render the super resolution images. For imaging the long tracks by permanent labeling, add the permanent imager strand and incubate for 120 seconds in T50M5 buffer.

Then wash the channel by infusing 200 microliters of wash buffer. Record an image with the excitation laser off to obtain background camera noise. Image a large area in a grid pattern by TIRF microscopy.

Program the microscope to scan over the area of 400 by 50 images and split the raw data into individual TIP files each containing a maximum of 10, 000 images using the ImageJ program. Flatten all the images using the illumination profile and use the MIST plugin to stitch the images. The result for the protein G ssDNA bioconjugation characterization showed a nearly one is to one ratio of protein G to ssDNA where the protein G Maleimide ssDNA and imidazole elution buffer spectra from the orthogonal basis for fitting to the bioconjugation product spectrum.

Native PAGE was used to confirm the bioconjugation showing the bright green bands coinciding with the monomeric protein G band indicating successful conjugation and good yield. The TIRF illumination profile introduced from a single mode fiber is generally brighter in the middle of the field of view and dimmer around the edges. To compensate for the uneven illumination and flatten the images for quantitative analysis, the illumination profile was determined by averaging thousands of individual frames.

Flattened images were produced by subtracting the camera noise from both raw and illumination profiles and then normalizing by the illumination profile. Raw stitching showed clear periodic patterns corresponding to the uncorrected images, while the same field of view stitched from flattened images produced a flat background. A ramp up flow profile was used to determine the range of shear stress resulting in cell rolling and yield fluorescence tracks under which a typical single-cell adhesion footprint could be seen.

Suboptimal and optimal results of fluorescent tracks with insufficient contrast, excessive track density, optimal track density and contrast, and diffraction limited and DNA-PAINT imaging are shown. incubation time of greater than 60 minutes ensure surface functionalization and proper chamber construction prevents the passage of bubbles that damage functionalized surface. This procedure is applicable for quantitative analysis of molecular force involved in rolling adhesion and allows researchers to understand the rolling behavior of the different cell types.

This technique allows researchers to explore new questions regarding fast adhesion events leading to further advancement of single molecule and mechanobiology research.