βバレル外膜タンパク質の構造決意を目指して - 構築物から結晶へ

The overall goal of this method is to obtain milligram quantities of outer membrane beta-barrel proteins that will crystallize and defract. This method can help answer key questions in the membrane protein folding and transport field, such as how beta-barrel proteins are inserted in cell membranes. The main advantage of this technique is that it is robust for beta-barrel proteins.

Generally, individuals new to this method will struggle, because choice of detergents for purification, crystallization, and defraction is challenging. Jeremy Guerin, a postdoc from Susan Buchanan's laboratory, will be demonstrating the procedure, in addition to myself and colleagues in the lab. Begin this procedure with construction of a T7 expression vector containing the codon optimized target outer membrane protein, or OMP gene, for in vivo expression to the membrane as described in the text protocol.

Transform the construct into an expression strain of bacteria for expression by pipetting one microliter of plasmid into 50 microliters of BL21(DE3)chemically competent cells, and mix gently by pipetting up and down. Incubate on ice for 30 minutes. Following incubation, heat pulse at 42 degrees Celsius for 30 seconds using a water bath, and then place back on ice for one minute.

Add one milliliter of pre-warmed SOC media and shake at 37 degrees Celsius for one hour at 1000 RPM using a bench-top shaker incubator. Following incubation, plate 100 microliters of cells onto LB Agar plates containing an appropriate antibiotic and incubate overnight at 37 degrees Celsius, inverted. Next, perform small-scale expression tests by inoculating five milliliters of LB containing antibiotic culture with a single colony.

Repeat for five to 10 colonies. Grow at 37 degrees Celsius with shaking to an optical density at 600 nanometers of approximately zero point six. Induce expression of the target OMP by adding five microliters of one molar IPTG to each culture tube and allow to grow an additional one to two hours.

To compare expression levels for all the colonies, centrifuge one milliliter of each culture for one minute at 15, 000 times G, using a microcentrifuge. Remove the supernatant and resuspend the cells in 200 microliters of 1X SDS-PAGE loading buffer. Heat at 100 degrees Celsius for five minutes, and then centrifuge again at 15, 000 times G for five minutes.

Analyze the samples using SDS-PAGE by pipetting 20 microliters into each well of a ten per cent gel. Run the gel for 35 minutes at a constant 200 volts. Finally, harvest the cells by centrifugation at 6000 times G for ten minutes.

Resuspend the cells in lysis buffer at a ratio of five milliliters per gram of cell paste. Lyse the cells using a French press or cell homogenizer. Then, spin the lysed cells at 15, 000 times G for 30 minutes at four degrees Celsius to remove unlysed cells and cell debris.

Transfer the supernatant to a clean tube and centrifuge again at high speed for one hour at four degrees Celsius. The resulting pellet is the membrane fraction which contains the protein of interest. Using a Dounce homogenizer, resuspend the membrane fraction.

First, transfer the membranes and then add solubilization buffer at 2X concentration without detergent. Pour the resuspended membranes into a graduated cylinder and add water until a final concentration of 1X solubilization buffer is reached. Transfer the sample to a beaker and add detergent slowly to a final concentration of approximately 10 times the critical micelle concentration.

Then, stir for 0.5 to 16 hours at four degrees Celsius, depending on how easily the target protein is extracted from the membranes. Finally, centrifuge the solubilized sample at 300, 000 times G for one hour at four degrees Celsius. Prepare a one to five milliliter IMAC column or use a pre-packed column.

Perform subsequent chromatography steps at four degrees Celsius. Flush with water to remove any traces of preservatives. If using an automated purification system, install the column according to manufacturer's instructions.

Prepare 500 milliliters of IMAC buffer A without imidizole and 250 millilliters of IMAC buffer B with one molar imidizole. Equilibrate the IMAC column using 10 column volumes of IMAC buffer A.Add imidizole to the protein sample to a final concentration of 25 millimolar and mix. Then, load the sample onto the equilibrated IMAC column at two milliliters per minute.

Collect the flow-through. Next, wash the IMAC column with five column volumes each of increasing concentrations of imidizole, using buffer B.Collect the washes in two milliliter fractions. Elute the sample with a final concentration of 250 millimolar for five column volumes and collect the eluted sample in two milliliter fractions.

Analyze the flow-through, the wash fractions, and the elution fractions using SDS-PAGE analysis based on their absorbance at 280 nanometers. Pool the fractions containing the target beta-barrel OMP as verified by SDS-PAGE analysis. Then, remove the 6X histadine tag by adding TEV protease to the pooled sample and incubating overnight at four degrees Celsius with gentle rocking.

Load the sample protease solution onto an IMAC column again to separate the target beta-barrel OMP from the cleaved tags and any uncleaved sample. The flow-through will contain the cleaved sample lacking the tag. To prepare for crystallization, load the sample onto a gel filtration column into a buffer containing the detergent to be used for crystallization.

Collect one milliliter fractions and analyze using SDS-PAGE analysis based on their absorbance at 280 nanometers. Pool those fractions containing the target protein, and then concentrate to approximately 10 milligrams per milliliter. Prior to preparing samples, assemble the gel apparatus.

Use pre-cooled gel running buffer or run the gel in the cold room. Filter the sample using a 0.22 micron centrifugal filter to remove particulates and precipitation. Then, pipette 0.25 microliters of the sample into two 1.5 milliliter micro centrifuge tubes.

Label one as boiled"and the other as RT"Next, add 9.75 microliters of the sample buffer to each tube and mix by pipetting. To both samples, also add 10 microliters of 2X SDS loading buffer and mix gently by pipetting. Boil the boiled"sample at 95 degrees Celsius for five minutes, while leaving the RT"sample at room temperature.

Then, spin the boiled"sample briefly. Next, load 20 microliters of both samples onto the pre-assembled native gel. After running the gel for 60 minutes at a constant 150 volts remove the gel and soak in staining solution to visualize the results.

Proceed with crystallization trials in detergent, bi-cells, and lipidic cubic phase, using previously published Jove methods. Crystallization heads are visualized under a UV microscope to verify that the crystals are, indeed, protein and not solved, lipid or detergent. Lead crystals are screened using a home source or at the synchrotron to see if the crystals defract.

After data collection of crystals that defract well, process the data using processing software, such as HKL2000. Transfer resulting files to a software suite for structure determination, such as Phoenix Suit, and perform molecular replacement for initial phasing and structure determination. The crystal structure of YIUR was determined at 2.6 Angstrom resolution using this method.

Once mastered, this technique can be done in two weeks, if it is performed properly. After watching this video, you should have a good understanding of how to express and purify bacterial outer membrane proteins for structural studies.