Biology
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Purification of Native Complexes for Structural Study Using a Tandem Affinity Tag Method
Chapters
Summary July 27th, 2016
The Tandem Affinity Purification (TAP) method has been used extensively to isolate native complexes from cellular extract, primarily eukaryotic, for proteomics. Here, we present a TAP method protocol optimized for purification of native complexes for structural studies.
Transcript
The overall goal of this protocol is to purify a native macromolecular complex of suitable quality for biophysical studies. This protocol serves not only as a way to purify a complex, but as a detailed guide to assessing the structural quality of the sample during purification. The main advantage of this protocol is that it allows one to purify a native assembly under near physiological buffer conditions simply and rapidly, so as to ensure compositional homogeneity.
After centrifugation of S.cerevisiae cells, and decanting the supernatant according to the text protocol, add two milliliters of chilled lysis buffer, and swirl and/or use a 10 milliliter serelogical pipette to suspend the pellet. Transfer the cell suspension into an empty 50 milliliter conical polypropylene centrifuge tube kept on ice. Then use an additional two milliliters of chilled lysis buffer to wash each centrifuge bottle and add the solution to the 50 milliliter tube.
After determining the wet weight of the cell pellet, create a liquid nitrogen bath in and around a 50 milliliter conical polypropylene centrifuge tube. Next, draw suspended cells into a five milliliter syringe and connect a 16 gauge needle to the syringe. Then, freeze the cell suspension by passing it through the syringe and needle into the bath at a rate of approximately one milliliter per minute to generate droplets.
To lyse the yeast cells, begin by using liquid nitrogen to pre-chill a coffee grinder. Then add up to 40 grams of yeast cells and grind for 25 seconds, repeating the grinding eight or nine times. It is recommended not to purify for more than 10 liters of cell culture at one time.
This minimizes purification time and ensures that structural integrity of the complex is maintained. After every two rounds of grinding, add a shallow layer of liquid nitrogen to the grinder and allow it to evaporate. To keep the cells from clumping, prevent the cells from thawing by using a liquid nitrogen chilled spatula to stir the cells.
Following lysis, the cells should appear as a fine white powder. After checking cell lysis and preparing lysis buffer with PMSF, DTT and protease inhibitors according to the text protocol, use a liquid nitrogen chilled spatula to incrementally scoop the frozen lysed cell powder into the prepared 50 milliliter tubes of the buffer. With each added increment, gently rotate the 50 milliliter tubes at room temperature so as to thaw and dissolve the cells and to prevent bubbles from forming.
As the cells dissolve, add more cell powder. After all the cells have been added, continue thawing and dissolving the cells until there are no frozen cell clumps observed in the tubes. This should take approximately 50 minutes.
Next, centrifuge the cell lysate at 25, 000 times g and four degrees Celsius for 20 minutes. Once cells are lysed and following centrifugation, you may observe a small dark layer in the insoluble pellet. Transfer the supernatant to polycarbonate ultracentrifuge tubes and add 10 microliters of 200 millimolar PMSF to each full tube.
Centrifuge the samples at 100, 000 times g and four degrees Celsius for one hour. At the end of the spin, four layers should be visible. A hard clear pellet containing mostly ribosomal complexes;a soft lipid rich pellet;a large, clear, yellow-tinged layer containing most of the cell's soluble proteins and complexes;and a scaly top layer consisting of lipids.
In a cold room at four degrees Celsius, use a one milliliter pipette to remove as much of the top scaly lipid layer as possible and discard. Use a 10 milliliter serological pipette to recover most of the clear yellow layer. Then, using a one milliliter pipette to avoid disturbing the bottom two layers, recover the last few milliliters of this layer.
From 40 grams of wet cell pellet, approximately 48 milliliters of the middle layer is typically recovered. After preparing IgG sepharose according to the text protocol, combine the IGG resin with the middle phase supernatant and two mini protease inhibitor tablets per 40 grams of cells. Then place on a rotator at four degrees Celsius for two hours.
This is the IgG batch solution. Prepare two 10 milliliter polyprep columns by cutting the ends of the columns to produce a flat opening. Pour the 24 milliliters of suspend IgG resin slurry or batch solution into each column, and allow to sediment by gravity.
This corresponds to 200 microliters of packed IgG resin. If sedimentation takes longer than 30 minutes, the middle phase may have been contaminated by lipids. When sedimentation is complete, use four successive 10 milliliter volumes of IgG D150 buffer to wash the packed column.
To carry out tobacco etch virus or TEV protease cleavage on the column, seal the bottom of the column and add one milliliter of IgG D150 buffer and 100 microliters of TEV protease. Seal the top of the column and using a thermal mixer at 750 RPM and 18 degrees Celsius, mix for 20 minutes. Re-suspend the resin and mix again for 20 minutes.
Then re-suspend and repeat the 20 minute mixing a third time. Return the column to four degrees Celsius and use gravity to elute the protein complex. Then, add 200 microliters of IgG D150 to elute the dead volume.
After preparing calmodulin affinity resin according to the text protocol, add the calmodulin binding buffer and sample to the resin, and use a tube rotator to incubate at four degrees Celsius for one hour. Prepare a two milliliter polyprep column per 100 microliters of calmodulin slurry by cutting the end of the column to create a flat opening. The protocol details specific buffer and residence volumes chosen so as to keep the complex concentrated and to minimize its resin residence time.
If cultural volume is changed, it is recommended to change the gravity columns'resin and buffer volumes to account for the change. Pack the column with no more than 100 microliters of calmodulin slurry and by gravity, use 5 milliliters of calmodulin binding buffer to wash the resin three times. Using 200 microliters of calmodulin elution buffer, elute the protein three times.
Then, seal the bottom of the column and add 20 microliters of elution buffer and incubate for 2.5 minutes before releasing the seal and allowing the protein to elute by gravity. Seal the bottom of the column again and add 200 microliters of elution buffer before incubating for five minutes. Then, unseal the bottom of the column and elute the solution.
For the sixth and final elution, seal the bottom of the column and incubate with elution buffer for 10 minutes. Then, unseal and elute. Carry out post-purification analysis and store samples according to the text protocol.
As shown here, an initial TAP purification of the complex following a published protocol yielded a complex that migrated as three bands on a silver-stained gel. Multiple rounds of optimization of the TAP method gives us a complex that migrated as a single band on a native gel, indicative of a more homogeneous assembly. Consistent with the native gel result, western blotting using a TAP antibody against the complex, purified according to the published protocol, detected proteolysis of the TAP tagged protein, SNU-71.
Modifications of the TAP method significantly reduced the amount of proteolysis, as nearly all of the 17 proteins in the U1 snRNP complex were resolved by SDS-PAGE and positively identified by multi mass spectrometry. Negative stain electron microscopy shows monodispersed particles from a successful purification after the first and second steps of TAP. In contrast, no particles of the correct size are observed when the purification is unsuccessful.
Final complex quality was assessed on a lawn of purified particles, shown here, that appear monodispersed in the negative stain EM raw image. In addition, class averages of the particles reveal distinct features indicating that the sample is possibly suitable for structural study. Once mastered, this protocol can be completed within 10 to 12 hours of thawing the frozen lysed cell powder.
While following this protocol, it's important to ensure that all solutions are free of proteases and nucleases, as well as to work rapidly to prevent aggregation or dissociation of the complex. After watching this video, you should understand how to purify a native complex of suitable quality for structural study, as well as how to assess that this has been achieved.
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