Biochemistry
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Combining Wet and Dry Lab Techniques to Guide the Crystallization of Large Coiled-coil Containing Proteins
Chapters
Summary January 6th, 2017
We describe a framework incorporating straightforward biochemical and computational analysis to guide the characterization and crystallization of large coiled-coil domains. This framework can be adapted for globular proteins or extended to incorporate a variety of high-throughput techniques.
Transcript
The overall goal of this procedure is to rapidly identify and test possible domain boundaries within large coiled-coil proteins to improve the likelihood of finding an intrinsic fragment that will crystallize. This method can help researchers streamline the process of crystallization for large coiled-coil proteins, which can be notoriously difficult to crystallize. The advantage of this technique is that it integrates both experimental and computational methods to quickly identify and screen likely candidates for crystallization, thereby improving their chances for success.
Demonstrating this technique will be Jenna Zalewski, a graduate student in my lab, and Keith O'Conor, an undergraduate researcher in my lab. To begin, combine output from established web-based tools to generate predictions of possible coiled-coil domain boundaries as described in the text protocol. After generating expression plasmids and testing their expression levels as detailed in the text, begin purification of each strain that successfully expressed Ruby tagged protein.
To lyse the cells, first resuspend each thawed cell pellet in 1.5 milliliters of lysis buffer that has been supplemented with protease inhibitors. Add 30 microliters of lysozyme at a concentration of 10 milligrams per milliliter and incubate at room temperature for 20 minutes, then sonicate, following the instructions for the instrument being used. Centrifuge the sample in a tabletop centrifuge for 30 minutes at 14, 000 times G to pellet insoluble material or unlysed cells.
Save both the supernatant and the pellet for subsequent analysis via SDS-PAGE. To perform nickel affinity purification, first incubate 100 microliters of the nickel NTA beads, prewashed in lysis buffer with the soluble lysate for five to 10 minutes. Invert the suspension several times to mix the beads.
Centrifuge the sample at 800 times G for 30 seconds in a tabletop centrifuge to pellet the beads. Follow this by washing the pellet three to five times with lysis buffer to clean off non-specific contaminants. Finally, elute the Ruby fusion protein from the beads with lysis buffer, supplemented with one molar imidazole.
Use a spectrophotometer set at 280 nanometers to measure the concentration of the Ruby fusion protein, then assess the homogeneity of the sample by loading one to five micrograms of fusion protein onto a native PAGE gel. Run the 10%native PAGE gel at four degrees Celsius for 140 minutes at 175 volts. Image the native PAGE using an imager equipped to visualize fluorescence, observing where the Ruby tagged fusion migrates in this assay.
Perform limited proteolysis to identify stably folded domains by first incubating 95 microliters of Ruby-Shroom SD2 fusion with five microliters of 0.025 Subtilisin A.Sample the reaction at time points of zero, 0.5, two, five, 15, 60 and 120 minutes, removing 10 microliters from the reaction for each time point and visualizing the progress of the reaction via SDS-PAGE using a 15%acrylamide gel. Stain the gel with Coomassie blue to evaluate whether a protease-resistant species can be identified. Integrate the data from the purification efficiency, behavior in native PAGE and limited proteolysis on the partially purified mRuby2-Shroom fusion proteins into a comprehensive assessment of the overall behavior of these proteins in solution.
Perform large scale expressions with two liter cultures of mRuby2-Shroom SD2 with the goal of achieving five to 20 milligrams of completely purified sample. Following cell growth, pellet the cells by centrifugation at 8, 000 times G for 10 minutes. The resulting cell pellet can be stored at minus 80 degrees Celsius until needed.
Next, re-suspend the pellet in approximately eight milliliters of lysis buffer per gram of frozen cell pellet. Lyse the re-suspended cells using a homogenizer. Following cell lysis, pellet the cellular debris via centrifugation at 30, 000 times G for 30 minutes.
Next, batch bind the soluble fraction using 10 milliliters of pre-washed nickel NTA resin. Pour the bead suspension into an appropriate gravity column and allow unbound proteins to drain off, then wash the resin three to five times with 40 milliliters of lysis buffer, followed by a wash with lysis buffer supplemented to one molar sodium chloride. Perform a final wash of the resin with 40 milliliters of lysis buffer supplemented with 80 millimolar imidazole.
Elute the Ruby-Shroom fusion protein with 40 milliliters of lysis buffer containing one molar imidazole. Manually fractionate the eluted protein into four to 10 milliliter fractions. Confirm fractions containing Ruby-Shroom fusion protein using a 10%SDS-PAGE gel.
Add one milligram of TEV protease for every 25 to 50 milligrams of fusion protein to remove the Ruby tag. Next, dialyze the appropriate fractions into TEV cleavage buffer using six to eight kilodalton molecular weight cutoff dialysis tubing. Dialyze the protein overnight at room temperature with slow stirring.
Following dialysis, repeat the nickel affinity purification steps. The Shroom-SD2 domain should flow through into the unbound fraction while the Ruby tag remains bound to the resin. Using a spin concentrator, concentrate the peak fractions to approximately 10 milligrams per milliliter, then perform size exclusion chromatography analyzing peak fractions via a 12%SDS-PAGE gel.
After pooling the peak fractions, dialyze the solution into crystallization buffer, then concentrate the dialyzed protein to 10 milligrams per milliliter prior to crystallization. Screen a small array of 12 conditions to identify an optimal protein concentration for crystallization trials using 24 well sitting drop crystallization trays. Perform crystallization trials using the vapor diffusion method.
First, pipet 500 microliters of each of the 12 conditions into separate wells. On each pedestal, make drops that contain one microliter of well solution and one microliter of protein sample. Quickly seal the tray with clear tape, then move the tray to a suitable temperature controlled environment that is vibration-free.
Examine the drops in the trays over the next three days using a microscope at up to 100X magnification. At the end of the three day period, score each drop as containing no precipitation, light precipitation, heavy precipitation or crystals. A suitable protein concentration should contain no more than six heavy precipitations.
Using the determined protein concentration, screen a broad range of commercially available crystallization screens. The use of a liquid handling or crystallization robot greatly speeds up this process while also minimizing error in the drops. It requires far less protein, allowing the user to screen more conditions with a single sample.
Improve the initial conditions identified from broad screens by systematically varying each of the variables within the crystallization drop using 24 well screen trays, as before. Limited proteolysis is used to identify regions of the protein that might be flexible or otherwise more accessible to protease. Shown here is a degradation pattern for an SD2 domain that is proteolytically sensitive, and another which is quite stable.
Limited proteolysis was performed within the context of the Ruby fusion proteins. In this case, the linker sequence between Ruby and the SD2 domain was rapidly cleaved, as expected, but the resulting Ruby and SD2 fragments remained largely stable. Analysis by native PAGE can quickly reveal good candidates for crystallization.
Four Shroom-Rock complexes with different protein batteries were run on native PAGE gels. The complex containing Rock 834 to 913 had the best behavior and crystallized readily, while the others only crystallized at degradation or failed to crystallize entirely. After watching this video, you should have a good understanding of how to develop a set of potential domain boundaries for your large coiled-coil protein and how to use the Ruby system to quickly and rapidly identify the best candidates for crystallization.
While crystallography remains a time-consuming endeavor, using this technique will focus your search for proteins that will crystallize using straightforwards assays to remove poor candidates, allowing you to spend more time on the candidates with better prospects for success. While attempting this procedure, it's important to remember to include as much information as possible when selecting your possible domain boundaries. The information that you choose to select when utilizing this procedure will vary for each protein.
If possible, other methods, like cell-based or biochemical assays, should be performed to ensure that the protein fragments in question still contain the function of biological interest. Although this method is focused on coiled-coil proteins, the overall framework could easily be adapted for almost any protein. Don't forget that working with certain instrumentation and biological reagents can be hazardous, and precautions, such as using gloves or wearing protective eyewear, should always be followed when performing this procedure.
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