Purification of Native Complexes for Structural Study Using a Tandem Affinity Tag Method

Summary

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.

Abstract

Affinity purification approaches have been successful in isolating native complexes for proteomic characterization. Structural heterogeneity and a degree of compositional heterogeneity of a complex do not usually impede progress in conducting such studies. In contrast, a complex intended for structural characterization should be purified in a state that is both compositionally and structurally homogeneous as well as at a higher concentration than required for proteomics. Recently, there have been significant advances in the application of electron microscopy for structure determination of large macromolecular complexes. This has heightened interest in approaches to purify native complexes of sufficient quality and quantity for structural determination by electron microscopy. The Tandem Affinity Purification (TAP) method has been optimized to extract and purify an 18-subunit, ~ 0.8 MDa ribonucleoprotein assembly from budding yeast (Saccharomyces cerevisiae) suitable for negative stain and electron cryo microscopy. Herein is detailed the modifications made to the TAP method, the rationale for making these changes, and the approaches taken to assay for a compositionally and structurally homogeneous complex.

Introduction

Many major cellular processes are carried out by large protein and protein-RNA complexes¹. A significant bottleneck to conducting biophysical and structural studies of such complexes is obtaining them of a suitable quality (i.e., homogeneity) and at an appropriate concentration. Isolating a complex from a native source has many advantages, including retaining relevant post-transcriptional and/or translational modifications of subunits and insuring proper complex assembly. However, large cellular complexes are often present in a cell at a low copy number and the purification must be highly efficient and occur under near physiological conditions to ensure complex integrity is maintained. Purifying a complex from a eukaryotic source is particularly challenging and can be financially prohibitive. Thus, strategies or methods that are efficient and yield a homogeneous complex are highly desired.

A strategy that has been successful in purifying native complexes from eukaryotic cells for their initial characterization is the Tandem Affinity Purification (TAP) method^2,3. The TAP method was initially devised to purify a native protein from the budding yeast (S. cerevisiae) in complex with interacting factor(s)². The TAP method utilized two tags, each tag fused in tandem to the same protein-coding gene sequence. The tags were selected so as to balance a need for tight and selective binding to an affinity resin with a desire to maintain near physiological solution conditions. This balance serves to preserve stable interaction(s) of the tagged protein with interacting factor(s) for post-purification characterization. The genomically incorporated TAP tag was placed at the end (C-terminal) of a protein-coding gene and consisted of a sequence coding for a Calmodulin Binding Peptide (CBP) followed by Protein A - an addition of just over 20 kDa to the tagged protein. CBP is short, 26 amino acids, and recognized by the ~ 17 kDa protein Calmodulin (CaM) in the presence of calcium with a K_D on the order of 10^-9 M ⁴. The Protein A tag is larger, consisting of two repeats of 58 residues with a short linker between the repeats. Each 58 amino acid repeat is recognized by immunoglobulin G (IgG) with a K_D ~ 10^-8 M ⁵. Between these two tags was incorporated a recognition site for TEV protease, an endopeptidase from the Tobacco Etch Virus^6,7. As illustrated in Figure 1, in the first affinity step of the method the TAP tagged protein is bound to an IgG resin via the Protein A. The tagged protein is eluted by on column cleavage upon the addition of TEV protease, site-specifically cleaving between the two tags. This is a necessary step as the interaction of IgG and Protein A is very strong and can only be adequately perturbed under denaturing solution conditions. Lacking a Protein A tag, the protein is bound to CaM resin in the presence of calcium and eluted from this resin with addition of the metal ion chelator EGTA (ethylene glycol tetraacetic acid) (Figure 1).

Soon after the introduction of the TAP method, it was used in a large-scale study to generate a 'map' of complex interactions in S. cerevisiae⁸. Importantly, as a consequence of this effort an entire yeast-TAP Tagged Open Reading Frame (ORF) library as well as individual TAP tagged ORFs⁹ are available from a commercial source. Thus, one can obtain any yeast strain with a tagged protein for any yeast complex. The TAP method also spurred modifications or variations of the TAP tag, including: its use for purification of complexes from other eukaryotic as well as bacterial cells^10,11; the design of a "split tag", wherein the Protein A and CBP are placed on different proteins¹²; and the tags changed, so as to accommodate the need of the investigator, such as sensitivity of the complex to Ca²⁺ or EGTA¹³.

Recent advances in both instrumentation and methodology have led to significant advances in the application of electron microscopy (EM) for structure determination, that have led to high, near atomic resolution images of macromolecular complexes¹⁴. The resolution obtainable of a complex by EM, however, remains contingent upon the quality of the complex under study. This study has utilized the TAP tag approach to purify from S. cerevisiae the U1 snRNP, an 18-subunit (~ 0.8 MDa) low copy number ribonucleoprotein complex that is part of the spliceosome^15,16. A number of steps have been taken to purify this complex such that it is homogeneous and of an adequate concentration. Potential problems encountered at various stages of the purification are described and strategies taken to overcome challenges highlighted. By carefully assessing and optimizing steps in the purification, the U1 snRNP purified is of a quality and at a quantity suitable for negative stain and electron cryo microscopy (cryo-EM) studies. An optimized TAP method protocol for purification of native complexes for structural studies is described herein.

Protocol

Note: The following protocol was devised for purification of a complex from 4 L of cell culture, approximately 40 g wet weight of cells. Once prepared, all buffers should be stored at 4 °C and used within a month of their preparation. Reducing agent and protease inhibitors are only added to buffers just prior to use.

1. Preparation of Whole Cell Extract for Tandem Affinity Purification 

1. Growth of S. cerevisiae Cells
   1. Streak the desired TAP tagged S. cerevisiae strain from storage (-80 °C) onto a yeast peptone dextrose (YPD) plate. Incubate for 3 - 5 days (30 °C), until yeast cells appear white and fluffy.
   2. Inoculate a streak of approximately 2 mm² of the fresh cells from the plate into 10 ml YPD media in a 50 ml conical polypropylene centrifuge tube. Shake the tube at 180 revolutions per min (rpm) overnight (30m°C).
   3. Inoculate 2 ml of the overnight culture per liter of YPD media in a 2 L Erlenmeyer or conical flask. Shake at 180 rpm (30 °C). Monitor cell growth at an optical density of 600 nm (OD₆₀₀) until late log phase, OD₆₀₀ of 1.8 to 2, typically 20 - 24 hr.
2. Harvesting S. cerevisiae Cells
   1. Harvest cells by centrifugation at 5,400 x g for 15 min (4 °C).
   2. Quickly decant the supernatant and keep cells at 4 °C or on ice.
   3. Suspend each liter of cell pellet with 2 ml chilled Lysis Buffer (10 mM Tris-HCl, pH 8.0; 300 mM NaCl; 10 mM KCl; 0.2 mM EDTA, pH 8.0; 5 mM imidazole, pH 8.0; 10% glycerol; 0.1% v/v NP-40). Suspend cells by swirling and/or repeat pipetting using a 10 ml serological pipet.
      Note: The importance of NP-40 as it pertains to complex quality and post-purification analysis is discussed below.
   4. Transfer the cell suspension into an empty 50 ml conical polypropylene centrifuge tube kept on ice. Wash each centrifuge bottle with an additional 2 ml of chilled Lysis Buffer and add to the cell suspension in the 50 ml tube.
   5. Determine the wet weight of cell pellet by weighing the tube and subtracting the weight of the empty tube as well as the added Lysis Buffer.
      Note: A liter of cell culture having an OD₆₀₀ of 1.8 is approximately 10 g.
   6. Create a liquid nitrogen bath in a 50 ml conical polypropylene centrifuge tube. Draw suspended cells into a 5 ml syringe. Connect a 16 G needle to the syringe. Freeze the cell suspension by passing it through the syringe/needle to generate cell "droplets." Rate of freezing should be approximately 1 min/ml.
   7. Store frozen cell droplets (-80 °C) or proceed to lysis.
3. Lysing Cells and Lysate Clarification
   1. Lyse the yeast cells using a coffee grinder. Lyse cells eight to nine times with 25 sec grinding bursts, while shaking the coffee grinder. Prior to use, pre-chill the coffee grinder with liquid nitrogen. Every two rounds of grinding, add a shallow layer of liquid nitrogen to the grinder and allow it to evaporate.
   2. Stir with a liquid nitrogen chilled spatula as needed to keep cells from clumping. Take caution to prevent thawing of cells, which can result in cell clumping. The cells should appear as a fine white powder following lysis. A maximum cell pellet from 4 L of yeast culture (~ 40 g) can be ground at a time.
      Note: Observations regarding method of lysis are discussed below.
   3. Assess the degree of cell lysis using a hemocytometer or alternative cell counting method. To provide an adequate analysis, take the following samples: (1) before lysis; (2) after four rounds of lysis; and (3) following lysis. To check these samples, take a small amount of cells with a liquid nitrogen chilled spatula and place in a 1.5 ml tube.
   4. Once cells are thawed, pipette 5 µl of cells and mix with 495 µl water. If 40 - 60% lysis of cells is observed, proceed to next step in the protocol.
      Note: Observations regarding effect of long periods of lysis are discussed below.
   5. Once adequate lysis is observed, store lysed cells (-80 °C) or proceed to next step.
   6. Prepare two 50 ml conical polypropylene centrifuge tubes each with 15 ml Lysis Buffer, including 1 mM phenylmethanesulfonyl fluoride (PMSF), 1 mM dithiothreitol (DTT), and a commercially available cocktail of protease inhibitors.
      Note: If significant proteolysis is observed, consider use of a protease deficient strain.
   7. Use a liquid nitrogen chilled spatula to scoop the frozen lysed cell powder into the prepared 50 ml tubes. Add the cell powder/solid incrementally to the Lysis Buffer, containing both a reductant and protease inhibitor(s).
   8. Rotate the 50 ml tubes gently at RT so as to thaw/dissolve the cells, but also to prevent bubbles from forming. As the cell powder/solid dissolves, add additional cell solid/powder.
   9. Fill each of the two 50 ml tubes until each has approximately 20 g of cells or all cells added. Continue thawing/dissolving until there are no frozen cell clumps observed in the 50 ml tubes. This should take approximately 50 min.
   10. Centrifuge the suspended cell lysate for 20 min at 25,000 x g (4 °C). Well-lysed cells may exhibit a small amount of black precipitate.
   11. Transfer the 50 ml of supernatant to polycarbonate ultra-centrifuge tubes (26.3 ml tubes used) and add 10 µl 200 mM PMSF to each full tube.
   12. Centrifuge for 1 hr at 100,000 x g (4°C). Four layers should be visible, from bottom to top: (1) a hard clear pellet, containing ribosomal complexes; (2) a soft lipid-rich pellet; (3) a large clear yellow-tinged layer containing most of the cell's soluble proteins and complexes; and (4) a 'scaly' top layer consisting of lipids.
   13. Perform all subsequent steps in a cold room (4 °C). Remove as much of the top 'scaly' lipid layer as possible using a 1 ml pipet and discard. Use a 10 ml serological pipet to recover most of the yellow, clear layer.
   14. Recover the last few ml of this layer using a 1 ml pipette to avoid disturbing the bottom two layers. From 40 g wet cell pellet, approximately 48 ml of the middle layer is typically recovered and 8 ml of lipid layer removed/discarded.
       Note: Column purification flow is greatly hindered by the presence of lipids, which also may reduce the quality of the complex, discussed below.

2. Column Purification step 1: IgG Chromatography

1. Resin Equilibration and Binding
   1. Prepare a 300 µl IgG Sepharose slurry for 40 g of cell pellet. Cut the end of a 1 ml pipet tip to facilitate pipetting the slurry. Wash/equilibrate the IgG resin three times, each time with 5 ml IgGD150 Buffer (10 mM Tris-HCl, pH 8.0; 150 mM NaCl; 150 mM KCl; 1 mM MgCl₂; 5 mM imidazole, pH 8; 0.1% v/v NP-40; 1 mM DTT; a protease inhibitor tablet per 50 ml volume).
      1. Spin to pellet at 160 x g (4°C) between the washes and remove supernatant.
   2. Rotate the IgG resin with the middle phase supernatant and two mini protease inhibitor tablets per 40 g of cells, using an appropriate rotator, for 2 hr (4 °C). This is the IgG batch solution.
   3. Prepare two 10 ml poly-prep columns for use by cutting the end of the column to produce a flat cut. To expedite the packing and washing of the column by gravity flow, load a maximum of 200 µl of packed IgG resin or ~ 25 ml of the IgG batch solution per 10 ml column.
   4. Pour the IgG batch solution into the column and allow to sediment by gravity. If sedimentation takes longer than 30 min, there most likely was lipid contamination when recovering the middle phase from the centrifuge tubes.
   5. Wash the packed column with four successive 10 ml volumes of IgGD150 Buffer.
2. On Column TEV Cleavage
   1. Seal the bottom of column and add 1 ml of IgGD150 Buffer and 100 µl of TEV protease (0.6 mg/ml stock, mutant variant S219V of TEV produced in-house). Seal the top of the column and mix for 20 min (18 °C) using a thermal mixer, shaking at 750 rpm. Resuspend resin and mix again for 20 min, repeat once more for a total of 1 hr incubation.
      Note: It is critical to keep time of incubation to a minimum.
   2. Return the column to 4 °C and elute the protein/complex by gravity. Elute the dead-volume with an additional 200 μl of IgGD150 Buffer.

3. Column Purification Step 2: Calmodulin Affinity Chromatography

1. Resin Equilibration and Binding
   1. Prepare 200 µl of Calmodulin affinity resin per 40 g of cell pellet. Wash the resin three times, each time with 5 ml Calmodulin Binding Buffer (10 mM Tris-HCl, pH 8.0; 150 mM NaCl; 10 mM KCl; 1 mM MgCl₂; 5 mM imidazole, pH 8; 2 mM CaCl₂; 0.1% v/v NP-40; 10 mM β-mercaptoethanol), centrifuging at 160 x g (4 °C) to pellet for each wash.
   2. To each 1 ml IgG eluate, add 3 volumes of Calmodulin Binding Buffer. To keep the calcium level constant, add CaCl₂ and β-mercaptoethanol to account for the lack of these constituents in the IgG elution. Final concentrations should be 2 mM CaCl₂ and 10 mM β-mercaptoethanol.
   3. Add the sample to the Calmodulin resin and bind for 1 hr (4 °C) using a tube rotator.
2. Packing and Eluting from Calmodulin Resin
   1. Prepare a 2 ml poly-prep column per 100 µl Calmodulin slurry by cutting the end of the column to create a flat opening. Load no more than 100 µl Calmodulin slurry per column to minimize the amount of resin the sample comes in contact with during elution.
   2. Pack the column by gravity and wash the resin in the column three times with 5 ml Calmodulin Binding Buffer.
   3. Elute protein with addition of 200 µl Calmodulin Elution Buffer (10 mM Tris-HCl, pH 8.0; 150 mM NaCl; 10 mM KCl; 1 mM MgCl₂; 5 mM imidazole, pH 8.0; 4 mM EGTA, pH 8.0; 0.08% v/v NP-40; 10 mM β-mercaptoethanol). Repeat six times the addition of 200 µl of Calmodulin Elution Buffer.
   4. For elutions 1 to 3, apply the volume to the column and collect the eluate immediately. For elution 4, seal the bottom of the column and incubate with elution buffer for 2.5 min and then unseal and allow to elute by gravity. For elution 5, incubate for 5 min and then unseal and elute. For elution 6, incubate for 10 min and then unseal and elute.
      Note: The integrity of the complex may vary between elutions/fractions (see representative results section).
   5. Dialyze elutions 2 to 6 individually overnight using an appropriate micro dialysis method. Dialyze the fractions against Dialysis Buffer (10 mM Tris-HCl, pH 8.0; 150 mM NaCl; 10 mM KCl; 1 mM MgCl₂; 5 mM imidazole, pH 8; 10 mM β-mercaptoethanol).
      Note: NP-40 does not pass appreciably, if at all, through the dialysis membrane.

4. Post-Purification Analyses to Assess Complex Quality

1. Native Gel and Silver Staining
   1. Prepare a 4 - 16% precast native gel for analysis of the complex as per the instructions from the manufacturer. Use an appropriate molecular weight protein standard and load 15 µl of each of the dialyzed Calmodulin elution/fractions onto the gel.
   2. Electrophorese the gel at 150 V for approximately 3 hr (4 °C).
   3. Silver stain the gel to assess the quality of each of the six elutions. Alternatively, use equally sensitive commercial fluorescent stain(s). At this stage, the concentration of the complex is most likely below the level of detection for Coomassie blue staining.
2. Additional Gel Analysis Methods
   1. To detect protein by Western blotting, resolve the complex by SDS or native PAGE and then probe the gel for the Calmodulin Binding Peptide (CBP) epitope using a TAP tag antibody as per the instructions from the manufacturer.
   2. Using a specific fluorescent stain(s), confirm the presence and integrity of protein and/or nucleic acid in a TAP purified complex. Take care that no spectral overlap is detected between these stains.
3. Negative Stain Electron Microscopy Visualization
   1. Use continuous carbon grids glow-discharged at -15 mA for 30 sec, within 30 min of complex application.
   2. Apply 3 µl of TAP purified complex (typically at a concentration of 1 nM) to grid. Incubate for a minute. Blot from the side of the grid to remove excess buffer.
   3. Wash twice with 20 µl drops of deionized water. Side blot between washes.
   4. Apply grid to a 50 µl negative stain drop and incubate for a min. Staining with uranyl acetate (2%) or uranyl formate (0.75%) is recommended.
      Caution: Uranyl salts and solutions are weakly radioactive and contain heavy metal. These should be handled with care and all material that comes in contact with these solutions should be disposed of following institutional recommended waste guidelines.
   5. Apply grid to a fresh 50 µl negative stain drop, without blotting. Incubate for a min, then side blot, and finally air dry and visualize.
4. Cryo Electron Microscopy (cryo-EM) Visualization
   1. Perform cryo-EM with TAP purified complex, though optimization may be necessary, according to manufacturer's protocol.
      Note: Presence of detergent at high concentrations may impede progress and is discussed below.

5. Complex storage

1. Preparing Complex for Storage
   1. Concentrate the complex, if needed, using a centrifugal filter having an appropriate molecular weight cut-off for the complex. Spin at 14,000 x g for 1 min increments until desired concentration is reached.
      Note: NP-40 concentration will increase in concentration during concentrating, as it does not pass through the filter nor dialysis membrane.
   2. Flash freeze the complex in 30 µl aliquots in liquid nitrogen or dry ice cooled ethanol bath and then store at -80 °C.
2. Optional Post-Purification Removal of Detergent

Note: The presence of a detergent such as NP-40 and at a concentration above that of its critical micelle concentration may hinder or inhibit progress for some applications. The consequences of its removal from the protocol as well as replacement with other additives or co-solvents are discussed below. If no NP-40 is desired, an option is to use a commercial application for removal, for example Bio-beads.

1. Pre-equilibrate commercially available detergent absorbing beads in Dialysis Buffer. Mix five beads per 30 μl of complex (usually at 10 nM concentration).
2. Incubate equilibrated beads from Step 5.2.1 with TAP purified complex for 15 min on ice. Use complex within a few hours following the removal of detergent.

Results

A modified TAP method was used to purify from S. cerevisiae the U1 snRNP, an 18-subunit ribonucleoprotein complex. An initial TAP purification of the complex following the published protocol^2,3 yielded a complex that appeared heterogeneous, migrating as three bands on a silver stained native polyacrylamide gel (Figure 2A). Multiple rounds of optimization of the TAP method, yielded a complex that migrated as primarily a single band on a native gel indic...

Discussion

The TAP method utilizes two tags that balance a need for tight and selective binding to an affinity resin with a desire to maintain near physiological solution conditions. This balance serves to preserve stable interaction(s) of the tagged protein with interacting factor(s) for post-purification characterization. In addition, individual TAP tagged ORFs are available from a commercial source, so that one can obtain any yeast strain with a tagged protein for any yeast complex. Preserving the integrity of a complex and the ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
S. cerevisiae TAP tagged strain	Open Biosystems	YSC1177	This is the primary yeast strain used to develop the TAP protocol. Its background is S288C: ATCC 201388: MATa, his3Δ1, leu2Δ0, met15Δ0, ura3Δ0, SNU71::TAP::HIS3MX6
Coffee grinder	Mr. Coffee	IDS77	Used for cell lysis
Hemocytometer, Bright Line	Hausser Scientific	3120	Used to assess cell lysis
JA 9.100 centrifuge rotor	Beckman Coulter, Inc.		Used to harvest the yeast cells
JA 20 fixed-angle centrifuge rotor	Beckman Coulter, Inc.		Used to clear the cell extract of non-soluble cellular material
Ti 60 fixed-angle centrifuge rotor	Beckman Coulter, Inc.		Used to further clear the soluble cell extract
Thermomixer	Eppendorf	R5355	Temperature controlled shaker
Novex gel system	Thermo Fisher Scientific
IgG resin	GE Healthcare	17-0969-01	Sepharose 6 fast flow
Calmodulin resin	Agilent Technologies, Inc.	214303	Affinity resin
Protease inhibitor cocktail, mini tablets	Sigma Aldrich	589297	Mini cOmplete ultra EDTA-free tablets
Protease inhibitor cocktail, large tablets	Sigma Aldrich	5892953	cOmplete ultra EDTA-free tablets
Phenylmethanesulfonyl fluoride (PMSF)			Dissolved in isopropanol
2 ml Bio-spin column	Bio-Rad Laboratories, Inc.	7326008	Used to pack and wash the Calmodulin resin
10 ml poly-prep column	Bio-Rad Laboratories, Inc.	7311550	Used to pack and wash the IgG resin
Precast native PAGE Bis-Tris gels	Life Technologies	BN1002	Novex NativePAGE Bis-Tris 4 - 16% precast polyacrylamide gels
NativeMark protein standard	Thermo Fisher Scientific	LC0725	Unstained protein standard used for native PAGE. Load 7.5 μl for a silver stained gel and 5 μl for a SYPRO Ruby stained gel
Precast SDS PAGE Bis-Tris gels	Life Technologies	NP0321	Novex Nu-PAGE Bis-Tris 4 - 12% precast polyacrylamide gels
PageRuler protein standard	Thermo Fisher Scientific	26614	Unstained protein standard used for Western blotting
SDS running buffer	Life Technologies	NP0001	1x NuPAGE MOPS SDS Buffer
TAP antibody	Thermo Fisher Scientific	CAB1001	Primary antibody against CBP tag
Secondary antibody	Thermo Fisher Scientific	31341	Goat anti-rabbit alkaline phosphatase conjugated
BCIP/NBT	Thermo Fisher Scientific	34042	5-bromo-4-chloro-3-indolyl-phosphate/nitro blue tetrazolium
Dialaysis units	Thermo Fisher Scientific	88401	Slide-A-Lyzer mini dialysis units
Centrifugal filter units, 100kDa MWCO	EMD Millipore	UFC5100008	Amicon Ultra-0.5 Centrifugal Filter Unit with Ultracel-100 membrane
Detergent absorbing beads	Bio-Rad Laboratories, Inc.	1523920	Bio-bead SM-2 absorbants
SYBR Green II	Thermo Fisher Scientific	S-7564	Flourescent dye for nucleic acid staining, when detecting with SYPRO Ruby present, use excitation wavelength of 488 nm and emission wavelength of 532 nm
SYPRO Ruby	Molecular Probes	S-12000	Flourescent dye for protein staining, when detecting with SYBR Green II present, use excitation wavelength of 457 nm and emission wavelength of 670 nm
Copper grids	Electron Microscopy Sciences	G400-CP