Exposure to stress can cause protein aggregation and result in significant alterations in the phenotypic behavior of bacteria. The procedure described in this video allows the extraction and visualization of protein aggregates after stress treatment. In contrast to other published procedures, this protocol requires lower cell numbers and abstains from complicated physical disruption processes, as well as time-consuming washing and incubation steps.
This protocol can be used to study protein aggregation in a wide variety of gram-negative and gram-positive bacteria. You can also analyze the effect of gene deletions or compare the efficacy of proteotoxic compounds. Begin by inoculating a single colony of E.coli strain MG1655 and uropathogenic E.coli strain CFT073, each into five milliliters of lysogeny broth, or LB, medium.
Incubate the two broth cultures at 37 degrees Celsius and 300 rpm for 14 to 16 hours. Dilute each strain to an optical density at 600 nanometers, or OD600, of 0.1 into a 500-milliliter flask containing 70 milliliters of MOPS-g medium. Incubate the flasks at 37 degrees and 300 rpm until the mid-log phase is reached.
After incubation, transfer 20 milliliters of each culture into three pre-warmed 125-milliliter flasks, and incubate them at 37 degrees Celsius and 300 rpm for two minutes. Prepare a two-milligram-per-milliliter antimicrobial compound solution in MOPS-g medium. Add this solution to each culture to reach the desired concentrations, and add the required volume of MOPS-g medium for the untreated control.
Determine the OD600 of each culture after 45 minutes of stress treatment. For each sample, aliquot four milliliters of culture with an OD600 of one into 15-milliliter centrifuge tubes. Resuspend the cell pellets in 50 microliters of ice-cold lysis buffer.
Incubate the samples for 30 minutes on ice. Add 360 microliters of ice-cold buffer A to the samples, and gently mix by pipetting. Transfer the samples into two-milliliter microcentrifuge tubes with around 200 microliters of 0.5-millimeter glass beads.
Incubate the tubes for 30 minutes at eight degrees Celsius in a ThermoMixer with shaking at 1400 rpm. Incubate the tubes on ice for five minutes without shaking to settle the glass beads. Then transfer 200 microliters of the cell lysate into 1.7-milliliter microcentrifuge tubes.
Centrifuge the cell lysate for 20 minutes at 16, 000 times g and four degrees Celsius. Collect the supernatant, which will be used as the soluble protein sample later. Use a pipette to resuspend the pellet in 200 microliters of ice-cold buffer A.Centrifuge of the tubes at 16, 000 times g and four degrees Celsius for 20 minutes.
Then carefully remove the supernatant altogether. Use a pipette to carefully resuspend the pellet in 200 microliters of ice-cold buffer B.Centrifuge the tubes again, and carefully remove the supernatant. Then repeat the wash with buffer A.Resuspend the pellet in 100 microliters of 1x reducing SDS sample buffer, and boil it for five minutes in a ThermoMixer at 95 degrees Celsius.
Mix one volume of 100%TCA with four volumes of the soluble protein sample collected earlier, and incubate for 10 minutes at four degrees Celsius for protein precipitation. Centrifuge the sample at 21, 000 times g and four degrees Celsius for five minutes to obtain the precipitate, and remove the supernatant. Use 200 microliters of ice-cold acetone to wash the pellet, to remove cellular debris.
Centrifuge the sample again to obtain the precipitate, and remove the supernatant. To remove the remaining acetone from the pellets, keep the microcentrifuge tubes in the ThermoMixer at 37 degrees Celsius with their lids open. Then add 100 microliters of 1x reducing SDS buffer to completely dissolve the pellet, and boil the sample at 95 degrees Celsius for five minutes.
Immediately load the sample on an SDS polyacrylamide gel for separation, or store the sample at minus 20 degrees Celsius to proceed later. Prepare two separating gels as described in the text manuscript. Pour the gel between the glass plates using a one-milliliter pipette, ensuring that the upper two centimeters are free of the mixture.
Add 70%ethanol on top of the separating gel, creating an even interface between the two layers. Once the separating gels have polymerized, prepare the stacking gels using instructions in the text manuscript. Then remove the ethanol from the separating gels, and add the stacking gel solution.
Carefully insert a comb with the desired number of pockets without introducing air bubbles, and allow the gel to polymerize for 20 to 30 minutes. Load four microliters of each sample, as well as the protein ladder, into separate wells. Then run the gel in tris-glycine running buffer at 144 volts for 45 minutes at room temperature.
Use pre-warmed Fairbanks solution A to stain the gels on a rocker for 30 minutes and pre-warmed Fairbanks solution D to de-color the gels on a rocker until the desired background is achieved. An increase was observed in the amount of protein aggregate formed when cells were treated with 175 micrograms per milliliter and 200 micrograms per milliliter of the antimicrobial, as compared to untreated cells. The increase in the insoluble protein fraction was more pronounced in the more sensitive MG1655 strain.
A decrease was observed in the amount of soluble proteins when cells were treated with 175 micrograms per milliliter and 200 micrograms per milliliter of the antimicrobial, as compared to untreated cells. When attempting this protocol, keep in mind that normalization to the optical density at 600 nanometers is important for comparison of the extent of protein aggregation in the samples.
