Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins

Here, we characterize protein structure and interaction sites in living cells using a protein footprinting technique termed in-cell fast photochemical oxidation of proteins (IC-FPOP).

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In-cell FPOP coupled with mass spectrometry is a powerful new technique used to study protein protein interactions and protein structural changes in living cells. With in-cell FPOP, you do not need to isolate proteins of interest, but can rather study them in their native environments. This is particularly helpful for membrane proteins.

With this method, you can study changes in structure and protein protein interactions in a variety of diseases, including cancer and genetic disorders, so you can better understand and characterize them. To begin the assembly of the flow system, use a cleavage stone to cut the fused silica tubing to the appropriate sizes. Next, place six small cylindrical magnets in one 500 microliter syringe and fill the syringe, a second 500 microliter syringe, and two five milliliter syringes with sheath buffer.

After removing the air, place the syringes on the syringe pump and tighten the syringe pump stopper so that the cell syringe has roughly 50 microliters left when the motor stalls. Using a lure adaptor, connect a manual valve to each syringe and assemble the silica tubing as illustrated, making sure to thread the cell plus hydrogen peroxide silica line through the cross and insert it directly into the 450 ID silica line. When all of the tubing has been connected, position the flow system next to a laser and use a lighter to burn away the silica coating on the 450 micrometer inner diameter tubing to make a window for laser irradiation.

Place a magnetic stirrer above the cell syringe containing the six magnets and set the syringe pump to 492.4 microliters per minute for a final flow rate of 1083.3 microliters per minute. Flow buffer through the system three times to flush the system, checking each connection for any leaks. Use a convex lens to focus the excimer laser onto the silica tubing.

To test the irradiation window, place a small piece of paper behind the silica tubing and turn on the laser. Then, measure the region burnt from the irradiation and use the irradiation window and flow rate to calculate the needed laser frequency to obtain an exclusion fraction of zero. To collect the cells for the procedure, wash the cells from a 70 to 90%confluent T175 flask culture with an appropriate cell culture grade salt solution.

Resuspend the cells in 10 milliliters of buffer for counting and collect the cells by centrifugation. It is critical to count the cells to ensure that they are enough for downstream analysis, but not too many to cause them to aggregate and clog the flow system. Then resuspend the cells at a two times 10 to the sixth cells per milliliter of sheath buffer concentration and aliquot the cells into one 500 microliter volume per sample.

For in-cell FPOP, fill the two five milliliter syringes with fresh sheath buffer, the 500 microliter syringe containing the magnets with one aliquot of cells, and the final 500 microliter syringe with freshly prepared 200 millimolar hydrogen peroxide. Turn on the magnetic stirrer and spike in 220 microliters of dimethyl sulfoxide to one aliquot of Quench. After gentle mixing, place the Quench behind the flow system to collect the irradiated samples and turn on the laser.

After seven seconds, turn on the flow system. Once the sample finishes flowing, turn off the laser and gently mix the Quench with the collected sample. Next, fill all four of the syringes with fresh buffer and flush the buffer through the flow system.

After the system finishes flushing, repeat the procedure with a new aliquot of cells and solutions without irradiation as the no laser control to account for background oxidation within the cells. While the next sample is running, centrifuge the first sample and resuspend the pellet in 100 microliters of an appropriate cell lysis buffer. Then, transfer the sample to a microcentrifuge tube for flash freezing in liquid nitrogen.

When all of the samples have finished running, disassemble the flow system and discard the used silica tubing. Then, clean all the other connections by sonication for one hour in a 50%water 50%methanol solution. After isolating and digesting the proteins from the cell lysate according to standard protocols, analyze the digested cell lysate according to standard tandem LCMS protocols to localize the FPOP modification protium-wide.

Then, load the data into an appropriate protein analysis software program and analyze the data against a relevant protein database and the relevant digest enzyme. Once the files are finished searching, calculate the extent of FPOP oxidation on the protein of interest. Fluorescence imaging of orthogonal YZ stacked images shows a clear separation of the sheath buffer from the cell solution as it flows past the laser irradiation window.

Three dimensional average heat maps of the sheath buffer or cell solution can be generated to illustrate the minimal mixing of the two solutions. The use of the single cell flow system increases the number of oxidatively modified proteins by 13 fold. To confirm modification of the proteins of interest within the intact cells, fluorescence imaging can be performed following hydrogen peroxide treatment and irradiation.

Using tandem mass spectrometry, these modifications can further be localized to specific amino acids on a protein. The shift observed in this extracted ion chromatogram translates to the change in hydrophobicity caused by the oxidized methionine in the modified peptide. Comparing in-cell labeled actin by in-vitro footprinting reveals similar levels of oxidization, indicating that actin has a similar solvent accessibility for both in-cell and in-vitro studies.

Further, comparison of the extent of the fast photochemical oxidation of the protein of interest modifications to the solvent accessibility, the labeled residues calculated from two actin crystal structures demonstrates that in-cell fast photochemical oxidation of the protein of interest efficiently probes the solvent accessibility of the monomeric protein. There are several additional steps one can take to increase the detection of the lower abundant FPOP modifications, including 2D chromatography, subcellular fractionation, and proteomics multiplexing techniques. With the use of in-cell FPOP, we can now perform proteomyostructural biology to better connect protein structures with the cellular function.

Due to 248 nanometer wavelength required for FPOP, be sure to always wear UV protective goggles and proper clothing to avoid unintended exposure to reflected or scattered radiation.

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Journal (Biochemistry)

Journal (Biochemistry)

Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins

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