ライブセル光親和性標識によって、ネイティブの細胞環境における小分子結合タンパク質の同定

The overall goal of this procedure, is to identify the binding proteins of small molecules by using a photoaffinity probe, that can bind to its target in live cells, allowing the subsequent isolation and identification of the target. This method can be used to elucidate the molecular mechanism of action of drugs or other small molecules. The main advantage of this technique, is that binding and covalent labeling of the target proteins occur within the native cellular environment, removing the risk of disrupting native protein structure and binding conditions upon cell lysis.

Begin this procedure, by preparing sterile six centimeter cell culture dishes, for the number of samples desired. Typically, one dish of cells is used per treatment condition. Three dishes of cells will be prepared for this demonstration, and negative control with DMSO only, labeled D, probe treatment only, labeled P, and probe plus competitor, labeled C.To each culture dish, add 3.5 million HEK 293 T cells in four milliliters of culture medium.

Incubate the cells overnight. Prior to treating the cells, pre-aliquot the needed drugs into 1.5 milliliter microcentrifuge tubes. For the competitor pre-treatment, add four microliters of DMSO to each of two tubes, and four microliters of 10 millimolar competitor to one tube.

For the probe treatment, add 16 microliters of DMSO to one tube, and 16 microliters of 15 micromolar photoaffinity probe to each of two tubes. Bring the cell culture dishes into the cell culture hood for the addition of the pre-aliquoted drugs. Aspirate one milliliter of culture medium from the competition treatment dish, transfer it to the microcentrifuge tube that contains the pre-aliquoted competitor, and resuspend the drug in the medium.

Gently add the medium and drug mixture dropwise back to the dish. In the same manner, add DMSO to both the negative control dish and the probe dish. Return the dishes to the incubator for 30 minutes.

After 30 minutes, bring the dishes back to the culture hood and dim the lights. Add the pre-aliquoted DMSO to the negative control dish, and probe to the probe and competition dishes. Return the dishes to the incubator for one hour.

After one hour, place the dishes on ice. Wash the cells in each dish gently with five milliliters of ice cold PBS to remove excess probe. Re-cover the cells with four milliliters of ice cold PBS.

Place a dish of cells, centered three centimeters under the UV lamp on top of an ice pack, to minimize heating from the lamp, and irradiate for three minutes. Remove the dish and place it on ice. In this manner, irradiate all the samples.

After irradiation of all samples, aspirate the PBS from the cells, and add 200 microliters of ice cold PBS with protease inhibitors to each dish. Detach the cells from the dish using a rubber scraper, and transfer to pre-labeled microcentrifuge tubes on ice. Add SDS to each sample to a final concentration of 0.4%Lyse the cells by sonicating the suspension for 10 pulses, incubating on ice for one minute, and then sonicating for another 10 pulses.

Boil the samples on a heat block set to 95 degrees Celsius for five minutes to complete cell lysis and denature all the proteins. After measuring the protein concentration in each sample, as described in the text protocol, normalize the protein concentration to 2.5 milligrams per milliliter, by adding PBS PH 8.5 plus 0.4%SDS as needed. To begin this procedure, transfer 40 microliters of each cell lysate, prepared in the previous segment to a new microcentrifuge tube.

Add the following reagents in this order, 0.2 microliters of flour-azide, 0.58 microliters of TCEP, and 3.38 microliters of TBTA. Vortex to mix. Add 1.14 microliters of copper sulfate pentahydrate to start the reaction.

Vortex briefly and incubate at room temperature for 30 minutes in the dark. Add 50 microliters of 2X SDS sample buffer to quench the reaction. Run the samples on an SDS-PAGE gel, and when the dye front has reached the end of the gel, continue running the gel for additional five minutes to ensure all of the excess unreacted flour-azide has completely exited the gel.

After washing away all excess flour-azide, place the gel onto a glass plate and scan the gel using a fluorescent gel scanner, according to the manufacturer's instructions. For the attachment of a biotin tag, use the maximum amount of cell lysates after protein normalization, such that all samples are of the same volume. Preclear the lysates by adding each sample to a new tube, containing 50 microliters of pre-washed high capacity streptavidin agarose beads.

Incubate for one hour at four degrees Celsius with rotation. Pellet the beads by centrifugation. Remove the supernatant to a new microcentrifuge tube on ice, and discard the beads.

Per 500 microliters of lysate, add the following reagents, 1.38 microliters of biotin-azide, 5.5 microliters of TCEP, and 32.5 microliters of TBTA. Vortex to mix. Add 11 microliters of copper sulfate pentahydrate per 500 microliters of lysate and vortex briefly.

Incubate at room temperature for 30 minutes. Add four sample volumes of acetone, chilled to 20 degrees Celsius, vortex the samples, and incubate overnight at 80 degrees Celsius, to completely precipitate the proteins and remove unreacted biotin-azide. On the following day, centrifuge the samples at 17, 000 x g for 15 minutes at four degrees Celsius to pellet the precipitated proteins.

Aspirate the supernatant completely, add 150 microliters of PBS PH 7.4 and 1%SDS to each sample, and resoluablize the proteins by sonication. Add 600 microliters of PBS to each sample, to dilute the concentration of SDS to 0.2%then add the sample to a new tube containing 30 microliters of pre-washed high capacity streptavidin agarose beads. Incubate for one hour at four degrees Celsius with rotation.

After one hour, pellet the beads by centrifugation at 1000 x g for three minutes. Aspirate and discard the supernatant, containing unbound proteins. Add one milliliter of wash buffer to the beads, and incubate for five minutes at room temperature with rotation.

Centrifuge, discard the supernatant, and wash with wash buffer again. After the final wash, aspirate the wash buffer completely from the beads, and add 30 microliters of 2X SDS sample buffer. Incubate for five minutes in a 95 degrees Celsius heatblock, to release the proteins from the beads.

Centrifuge the beads at 13000 x g at room temperature for one minute. Carefully pipette the sample buffer, containing proteins off of the beads, for SDS-PAGE. The visualization of proteins after labeling with the fluorescent tag, is illustrated by this fluorescent scanned gel.

The major specific binding protein band, at approximately 35 kilodaltons, is only present in the probe lane, and is competed away by excess parent compound. The same 35 kilodalton band was visualized by silver staining after biotin pulldown, and subsequently identified by mass spectrometry as the membrane protein VDAC1. The identity of the protein was further validated, using a specific antibody for VDAC1.

The signal is present in the probe lane, and decreased in the competition lane. Including the input fraction, ensures that the antibody works and the protein of interest can be detected in the lysate. The slight increase in molecular weight of the pulldown samples, is due to the covelantly attached probe.

The consequences of not performing certain steps in the protocol correctly are illustrated. Incomplete removal of excess flour-azide, results in large black smears at the bottom of the fluorescent scanned gel, while insufficient preclearing of the lysates and or washing of the beads, results in a silver stained gel with very high background staining. From start to finish, the pulldown experiment takes two to three days to complete.

After the target protein has been isolated and identified by mass spectrometry or western blotting, further target specific validation experiments should be performed to assess the relevance of the target to the drug-induced phenotype.