The authors are grateful for the support and advice of Nikolaus Grigorieff. We thank Anna Loveland, Axel Brilot, Chen Xu, and Mike Rigney for helpful discussions and EM guidance. This work was funded by the National Science Foundation, Award No. 1157892. The Brandeis EM facility is supported by National Institutes of Health grant P01 GM62580.

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 &#176;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&#160;
<ol>
	<li>Growth of S. cerevisiae Cells
	<ol>
		<li>Streak the desired TAP tagged S. cerevisiae strain from storage (-80 &#176;C) onto a yeast peptone dextrose (YPD) plate. Incubate for 3 - 5 days (30 &#176;C), until yeast cells appear white and fluffy.</li>
		<li>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&#176;C).</li>
		<li>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 &#176;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.</li>
	</ol>
	</li>
	<li>Harvesting S. cerevisiae Cells
	<ol>
		<li>Harvest cells by centrifugation at 5,400 x g for 15 min (4 &#176;C).</li>
		<li>Quickly decant the supernatant and keep cells at 4 &#176;C or on ice.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#34;droplets.&#34; Rate of freezing should be approximately 1 min/ml.</li>
		<li>Store frozen cell droplets (-80 &#176;C) or proceed to lysis.</li>
	</ol>
	</li>
	<li>Lysing Cells and Lysate Clarification
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Once cells are thawed, pipette 5 &#181;l of cells and mix with 495 &#181;l 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.</li>
		<li>Once adequate lysis is observed, store lysed cells (-80 &#176;C) or proceed to next step.</li>
		<li>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.</li>
		<li>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).</li>
		<li>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.</li>
		<li>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.</li>
		<li>Centrifuge the suspended cell lysate for 20 min at 25,000 x g (4 &#176;C). Well-lysed cells may exhibit a small amount of black precipitate.</li>
		<li>Transfer the 50 ml of supernatant to polycarbonate ultra-centrifuge tubes (26.3 ml tubes used) and add 10 &#181;l 200 mM PMSF to each full tube.</li>
		<li>Centrifuge for 1 hr at 100,000 x g (4&#176;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&#39;s soluble proteins and complexes; and (4) a &#39;scaly&#39; top layer consisting of lipids.</li>
		<li>Perform all subsequent steps in a cold room (4 &#176;C). Remove as much of the top &#39;scaly&#39; 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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
2. Column Purification step 1: IgG Chromatography
<ol>
	<li>Resin Equilibration and Binding
	<ol>
		<li>Prepare a 300 &#181;l 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).
		<ol>
			<li>Spin to pellet at 160 x g (4&#176;C) between the washes and remove supernatant.</li>
		</ol>
		</li>
		<li>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 &#176;C). This is the IgG batch solution.</li>
		<li>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 &#181;l of packed IgG resin or ~ 25 ml of the IgG batch solution per 10 ml column.</li>
		<li>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.</li>
		<li>Wash the packed column with four successive 10 ml volumes of IgGD150 Buffer.</li>
	</ol>
	</li>
	<li>On Column TEV Cleavage 
	<ol>
		<li>Seal the bottom of column and add 1 ml of IgGD150 Buffer and 100 &#181;l 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 &#176;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.</li>
		<li>Return the column to 4 &#176;C and elute the protein/complex by gravity. Elute the dead-volume with an additional 200 &#956;l of IgGD150 Buffer.</li>
	</ol>
	</li>
</ol>
3. Column Purification Step 2: Calmodulin Affinity Chromatography
<ol>
	<li>Resin Equilibration and Binding 
	<ol>
		<li>Prepare 200 &#181;l 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 &#946;-mercaptoethanol), centrifuging at 160 x g (4 &#176;C) to pellet for each wash.</li>
		<li>To each 1 ml IgG eluate, add 3 volumes of Calmodulin Binding Buffer. To keep the calcium level constant, add CaCl2 and &#946;-mercaptoethanol to account for the lack of these constituents in the IgG elution. Final concentrations should be 2 mM CaCl2 and 10 mM &#946;-mercaptoethanol.</li>
		<li>Add the sample to the Calmodulin resin and bind for 1 hr (4 &#176;C) using a tube rotator.</li>
	</ol>
	</li>
	<li>Packing and Eluting from Calmodulin Resin
	<ol>
		<li>Prepare a 2 ml poly-prep column per 100 &#181;l Calmodulin slurry by cutting the end of the column to create a flat opening. Load no more than 100 &#181;l Calmodulin slurry per column to minimize the amount of resin the sample comes in contact with during elution.</li>
		<li>Pack the column by gravity and wash the resin in the column three times with 5 ml Calmodulin Binding Buffer.</li>
		<li>Elute protein with addition of 200 &#181;l 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 &#946;-mercaptoethanol). Repeat six times the addition of 200 &#181;l of Calmodulin Elution Buffer.</li>
		<li>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).</li>
		<li>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 &#946;-mercaptoethanol). 
		Note: NP-40 does not pass appreciably, if at all, through the dialysis membrane.</li>
	</ol>
	</li>
</ol>
4. Post-Purification Analyses to Assess Complex Quality
<ol>
	<li>Native Gel and Silver Staining
	<ol>
		<li>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 &#181;l of each of the dialyzed Calmodulin elution/fractions onto the gel.</li>
		<li>Electrophorese the gel at 150 V for approximately 3 hr (4 &#176;C).</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Additional Gel Analysis Methods
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Negative Stain Electron Microscopy Visualization
	<ol>
		<li>Use continuous carbon grids glow-discharged at -15 mA for 30 sec, within 30 min of complex application.</li>
		<li>Apply 3 &#181;l 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.</li>
		<li>Wash twice with 20 &#181;l drops of deionized water. Side blot between washes.</li>
		<li>Apply grid to a 50 &#181;l 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.</li>
		<li>Apply grid to a fresh 50 &#181;l negative stain drop, without blotting. Incubate for a min, then side blot, and finally air dry and visualize.</li>
	</ol>
	</li>
	<li>Cryo Electron Microscopy (cryo-EM) Visualization
	<ol>
		<li>Perform cryo-EM with TAP purified complex, though optimization may be necessary, according to manufacturer&#39;s protocol. 
		Note: Presence of detergent at high concentrations may impede progress and is discussed below.</li>
	</ol>
	</li>
</ol>
5. Complex storage
<ol>
	<li>Preparing Complex for Storage
	<ol>
		<li>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.</li>
		<li>Flash freeze the complex in 30 &#181;l aliquots in liquid nitrogen or dry ice cooled ethanol bath and then store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Optional Post-Purification Removal of Detergent</li>
</ol>
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.
<ol style="margin-left: 40px;">
	<li>Pre-equilibrate commercially available detergent absorbing beads in Dialysis Buffer. Mix five beads per 30 &#956;l of complex (usually at 10 nM concentration).</li>
	<li>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.</li>
</ol>

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The authors have nothing to disclose.

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 availability of a resource to test use of different tagged protein subunits in a complex for purification are two advantages for utilizing the TAP method for purifying a native complex. However, the original TAP method protocol was not devised to ensure that the complex purified is compositionally and structurally homogeneous. The TAP method has therefore been modified, as detailed above, to achieve this goal.
Various steps in TAP method were assessed to establish the optimal approach and conditions to purify a compositional and structural homogeneous complex. An early critical step is cell lysis. This step requires a balance between one&#39;s desire for a high yield and thus maximum lysis, while insuring that the complex obtained is of high quality, i.e., homogeneous. Different approaches to lysing yeast cells were explored, including using a bead-beater, high sheer fluid processor, and ball mill. Use of the less expensive coffee grinder was less efficient than the alternative methods (40 - 60% lysis), but the complex isolated and purified appeared monodisperse while several of these approaches appeared to either perturb particle integrity or cause a degree of aggregation. In the end, a significant number of cells were not lysed so as to ensure high quality complex was obtained. While 40 - 60% lysis is ideal for the yeast U1 snRNP, optimization is suggested when applying this protocol to a new biological complex.
It was critical to identify the steps in the method that could hasten purification to ensure that complex integrity was not compromised by cellular proteases, subunit dissociation as a result of dilution, and extended incubation at 18 &#176;C during TEV protease cleavage. One simple step to achieve time efficient purification was to ensure that lipids did not carry over from the early lysate clarification, since lipids impede flow during gravity purification. Other steps included identifying an optimal balance of the amount resin and column size/dimensions to achieve efficient purification and a maximum flow rate. A particularly critical step was to reduce time of on column cleavage by TEV protease. The TEV protease used was purified in house to ensure it was protease/nuclease free and at a high concentration17. Significant amounts of TEV protease were used to obtain complete cleavage in under an hour.
In initial TAP purifications proteolysis was observed. Proteolysis during purification can be limited by working efficiently, adding a battery of inhibitors, and including in the purification high salt washes. A broad range of protease inhibitors supplied over the first half of the purification greatly improved the stability of the TAP tagged protein and complex. In addition, proteolysis was minimized by increasing the monovalent concentration of 300 to 500 mM during the cell lysis and IgG steps of purification. Even with the above modifications to the TAP method, successive eluted fractions off of the CBP affinity resin varied in their degree of compositional homogeneity. Thus, selective pooling of fractions from the second, CBP chromatography step, is warranted.
An aspect of TAP method that received particular focus was the importance of the non-ionic detergent NP-40 to complex purification and at a concentration above its CMC. The presence of NP-40 at a concentration of 0.08% or higher had an overall beneficial effect on complex integrity and the final yield. The apparent beneficial effect(s) of NP-40 may be due in part to reducing non-specific interactions between resin and the complex. While NP-40 appears to have a beneficial effect during the purification, it could be removed post-purification without causing complex aggregation.
Finally, critical to optimizing the TAP method for structural study were several complementary assays used to assess the integrity and homogeneity of the complex. These included light scattering, SDS and native PAGE probed using the TAP antibody by Western blot and/or stained with fluorescent dyes, as well as negative stain electron microscopy. The changes made to the TAP method detailed herein, yielded a homogeneous complex at sufficient quantities for electron microscopy. Additional changes may need to be made to the TAP method for other complexes, in particular a determination of the importance of NP-40.
In summary, the TAP purification method can be used to successfully purify macromolecular complexes from a eukaryotic source of suitable quality for structural study. We have established a suitable approach for purification of the S. cerevisiae U1 snRNP and detail the steps taken as well as the rationale for many of these changes. For other complexes, we suggest that the investigator similarly explore the steps in the protocol, which may yield adequate amounts and quality of complex. The steps that should be examined include cell lysis, salt and additive type and concentrations including detergent, and sensitivity of the complex to calcium and metal chelator EGTA. Further, it is critical for long-term studies that the complex can be frozen and thawed without perturbing structure. Importantly, one should have several alternative ways to assay the structure of the complex during and following purification. Native and SDS PAGE, negative stain EM, and light scattering have served us well in our investigation of how best to purify a homogeneous complex using the TAP method.

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) method2,3. 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 KD 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 KD ~ 10-8 M 5. Between these two tags was incorporated a recognition site for TEV protease, an endopeptidase from the Tobacco Etch Virus6,7. 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 &#39;map&#39; 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 cells10,11; the design of a &#34;split tag&#34;, 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 spliceosome15,16. 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.

<table><tbody><tr><td>S. cerevisiae TAP tagged strain</td><td>Open Biosystems</td><td>YSC1177</td><td>This is the primary yeast strain used to develop the TAP protocol. Its background is S288C: ATCC 201388: MATa, his3&amp;Delta;1, leu2&amp;Delta;0, met15&amp;Delta;0, ura3&amp;Delta;0, SNU71::TAP::HIS3MX6</td></tr><tr><td>Coffee grinder</td><td>Mr. Coffee</td><td>IDS77</td><td>Used for cell lysis</td></tr><tr><td>Hemocytometer, Bright Line</td><td>Hausser Scientific</td><td>3120</td><td>Used to assess cell lysis</td></tr><tr><td>JA 9.100 centrifuge rotor&amp;nbsp;</td><td>Beckman Coulter, Inc.</td><td></td><td>Used to harvest the yeast cells</td></tr><tr><td>JA 20 fixed-angle centrifuge rotor</td><td>Beckman Coulter, Inc.</td><td></td><td>Used to clear the cell extract of non-soluble cellular material</td></tr><tr><td>Ti 60 fixed-angle centrifuge rotor</td><td>Beckman Coulter, Inc.</td><td></td><td>Used to further clear the soluble cell extract</td></tr><tr><td>Thermomixer</td><td>Eppendorf</td><td>R5355</td><td>Temperature controlled shaker</td></tr><tr><td>Novex gel system</td><td>Thermo Fisher Scientific&amp;nbsp;</td><td></td><td></td></tr><tr><td>IgG resin</td><td>GE Healthcare</td><td>17-0969-01</td><td>Sepharose 6 fast flow</td></tr><tr><td>Calmodulin resin</td><td>Agilent Technologies, Inc.</td><td>214303</td><td>Affinity resin</td></tr><tr><td>Protease inhibitor cocktail, mini tablets</td><td>Sigma Aldrich</td><td>589297</td><td>Mini cOmplete ultra EDTA-free tablets</td></tr><tr><td>Protease inhibitor cocktail, large tablets</td><td>Sigma Aldrich</td><td>5892953</td><td>cOmplete ultra EDTA-free tablets</td></tr><tr><td>Phenylmethanesulfonyl fluoride (PMSF)</td><td></td><td></td><td>Dissolved in isopropanol</td></tr><tr><td>2 ml Bio-spin column</td><td>Bio-Rad Laboratories, Inc.</td><td>7326008</td><td>Used to pack and wash the Calmodulin resin</td></tr><tr><td>10 ml poly-prep column</td><td>Bio-Rad Laboratories, Inc.</td><td>7311550</td><td>Used to pack and wash the IgG resin</td></tr><tr><td>Precast native PAGE Bis-Tris gels</td><td>Life Technologies</td><td>BN1002</td><td>Novex NativePAGE Bis-Tris 4 - 16% precast polyacrylamide gels</td></tr><tr><td>NativeMark protein standard</td><td>Thermo Fisher Scientific</td><td>LC0725</td><td>Unstained protein standard used for native PAGE. Load 7.5 &amp;mu;l for a silver stained gel and 5 &amp;mu;l for a SYPRO Ruby stained gel</td></tr><tr><td>Precast SDS PAGE Bis-Tris gels</td><td>Life Technologies</td><td>NP0321</td><td>Novex Nu-PAGE Bis-Tris 4 - 12% precast polyacrylamide gels&amp;nbsp;</td></tr><tr><td>PageRuler protein standard</td><td>Thermo Fisher Scientific</td><td>26614</td><td>Unstained protein standard used for Western blotting</td></tr><tr><td>SDS running buffer</td><td>Life Technologies</td><td>NP0001</td><td>1x NuPAGE MOPS SDS Buffer</td></tr><tr><td>TAP antibody</td><td>Thermo Fisher Scientific</td><td>CAB1001</td><td>Primary antibody against CBP tag</td></tr><tr><td>Secondary antibody</td><td>Thermo Fisher Scientific</td><td>31341</td><td>Goat anti-rabbit alkaline phosphatase conjugated</td></tr><tr><td>BCIP/NBT</td><td>Thermo Fisher Scientific</td><td>34042</td><td>5-bromo-4-chloro-3-indolyl-phosphate/nitro blue tetrazolium&amp;nbsp;</td></tr><tr><td>Dialaysis units</td><td>Thermo Fisher Scientific</td><td>88401</td><td>Slide-A-Lyzer mini dialysis units</td></tr><tr><td>Centrifugal filter units, 100kDa MWCO</td><td>EMD Millipore</td><td>UFC5100008</td><td>Amicon Ultra-0.5 Centrifugal Filter Unit with Ultracel-100 membrane</td></tr><tr><td>Detergent absorbing beads</td><td>Bio-Rad Laboratories, Inc.</td><td>1523920</td><td>Bio-bead SM-2 absorbants</td></tr><tr><td>SYBR Green II</td><td>Thermo Fisher Scientific</td><td>S-7564</td><td>Flourescent dye for nucleic acid staining, when detecting with SYPRO Ruby present, use excitation wavelength of 488 nm and emission wavelength of 532 nm</td></tr><tr><td>SYPRO Ruby</td><td>Molecular Probes</td><td>S-12000</td><td>Flourescent dye for protein staining, when detecting with SYBR Green II present, use excitation wavelength of 457 nm and emission wavelength of 670 nm</td></tr><tr><td>Copper grids</td><td>Electron Microscopy Sciences</td><td>G400-CP</td><td></td></tr></tbody></table>

purification of native complexes for structural study using a tandem affinity tag method

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.

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 protocol2,3 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 indicative of a more homogeneous assembly (Figure 2A). Using fluorescent dyes to stain for both protein and nucleic acid, it was established that the purified complex had both biopolymers present (Figure 2B).
The purified complex was also analyzed by SDS PAGE and Western blotting using a TAP tag antibody (Figure 3A). Consistent with the native gel result, the complex purified according to the published protocol exhibited proteolysis of the TAP tagged protein (Snu71). The amount of proteolysis, however, was significantly reduced with modifications of the TAP method. Nearly all of the seventeen proteins in the U1 snRNP complex were resolved by SDS PAGE and positively identified by MALDI mass spectrometry (Figure 3B). As well as native and SDS PAGE, complex quality was assessed at different stages of the purification by negative stain electron microscopy (Figure 4). The monodispersed particles observed in Figure 4A and B but absent from 4C are examples of good sample for structural study. This analysis method provided critical insight as to the impact of changes to the TAP method.
The TAP method calls for the inclusion of the non-ionic detergent NP-40 and at a concentration above that of its critical micelle concentration (CMC). The robustness of the methods for assessing the quality of the complex described in the protocol are well demonstrated by an experiment where the zwitterionic surfactant CHAPS, glycerol or no co-solvent were substituted for NP-40 in all buffers (Figure 5). By native PAGE and negative stain EM, the U1 snRNP appears most homogeneous and stable in NP-40 with decreasing stability from CHAPS to glycerol to no co-solvent added. It might be that the yeast U1 snRNP has a hydrophobic face and the presence of NP-40 prevents aggregation by coating this surface. Unfortunately, NP-40 cannot be removed by dialysis and concentrating the complex will then concentrate the NP-40. While the higher NP-40 did not appear to perturb the complex, it did negatively impact the freezing of the particle in vitreous ice for cryo-EM and large micelle-like aggregates were observed. Detergent absorbing beads were used successfully to remove the majority of the NP-40 from the U1 snRNP sample without appearing to impact the complex structurally.
Following the purification of the complex with the modifications detailed in the protocol, a complex suitable for both negative stain and cryo-EM was obtained, as evidenced by images of the lawn of particles and distinctive representative class averages (Figure 6).
<img alt="Figure 1" src="/files/ftp_upload/54389/54389fig1.jpg" /> 
Figure 1. Overview of Tandem Affinity Purification (TAP) Method. (A) Schematic of the two step TAP method. Complex of interest (grey) includes a TAP tagged subunit (black; U1 snRNP subunit Snu71) genomically fused with a sequence comprised of a Calmodulin Binding Peptide (CBP, green), a recognition site for the site-specific protease TEV (Blue), and two IgG binding regions of Protein A (Pr A, Red). In the 1st step, cell lysate is incubated with an IgG resin and the complex is retained via its interaction with the Protein A sequence. The complex is released from the resin by TEV mediated cleavage. In the 2nd step, the complex is incubated with a Calmodulin resin, a Ca2+ dependent reaction. The complex is released via addition of the Ca2+ chelator EGTA. (B) Western blot following a TAP tagged protein through the purification steps illustrated in (A). Samples were taken of cell lysate, after the 1st step, and after the 2nd step of the TAP method. The reactive protein observed in cell lysate migrates faster after the 1st step, following TEV protease cleavage and thus loss of the Protein A sequence. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54389/54389fig2.jpg" /> 
Figure 2. Optimization of the TAP Method. (A) Silver stained native gel of TAP purified complexes. Native PAGE of: protein standard (lane 1); complex purified according to original TAP method3 (lane 2), three bands observed (arrowheads); three consecutive elutions from the same TAP purification following alterations to buffer, two bands observed (arrowheads) (lanes 3 - 5); further changes to the TAP method, yielding primarily a single complex band migrating at ~ 800 kDa (lane 6). (B) Analysis of composition of a purified complex by staining a single gel lane with both protein and RNA reactive stains. Native PAGE of: protein standard (lane 1) stained with a protein fluorescent stain; complex imaged with protein fluorescent stain (lane 2), two bands observed; same lane in lane 2, now imaged with a nucleic acid fluorescent stain (lane 3), two bands observed; and fluorescent signal overlay of stained complex in lanes 2 and 3 (lane 4). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54389/54389fig3.jpg" /> 
Figure 3. Optimization of TAP Method. (A) Western blot analysis of improvement in complex quality, following TAP tagged subunit Snu71 probed with &#945;-TAP antibody. Several proteolyzed bands are visible initially (lane 1) but improvements reduce the amount of lower bands considerably (lanes 2 and 3). (B) A representative SDS gel stained with a fluorescent protein gel stain. Gel resolves 12 of the 17 protein subunits of the complex. Identity of protein in the bands was established by mass spectrometry. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/54389/54389fig4.jpg" /> 
Figure 4. Sample Quality at Stages in the TAP Method Assessed by Negative Stain Electron Microscopy. Assessment by negative stain electron microscopy of complex after: (A) the 1st step of TAP; (B) the 2nd step of TAP; (C) complex destabilized. Scale bar, 125 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" src="/files/ftp_upload/54389/54389fig5.jpg" /> 
Figure 5. Impact on Sample Quality of Additives Present in TAP Method Buffers. Lanes 1 - 4 are the first four elutions of the final or 2nd step of TAP purification of complex purified in the presence of: (A) the non-ionic detergent nonyl phenoxypolyethoxylethanol (NP-40), analyzed by native PAGE (top) and negative stain EM (bottom); (B) the zwitterionic detergent 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate (CHAPS), analyzed by native PAGE (top) and negative stain EM (bottom); (C) glycerol, analyzed by native PAGE (top) and negative stain EM (bottom); and (D) no additive, analyzed by native PAGE (top) and negative stain EM (bottom). Scale bar, 50 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" src="/files/ftp_upload/54389/54389fig6.jpg" /> 
Figure 6. Visualization of TAP Method Purified Complex. (A) Top, a representative image of a negative stain electron microscopy (EM) micrograph. Observed is a lawn of similarly shaped particles, two are boxed, purified by the optimized TAP method. Bottom, a selection of class averages taken of the particles that highlight particle distinguishing features. Scale bar, 50 nm. (B) Top, a representative image of a cryo-EM micrograph. Observed are regions of higher contrast representative of particles, see two such regions boxed. Bottom, class averages of selected particles frozen in vitreous ice. Scale bar, 50 nm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54389/54389fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Purification of Native Complexes for Structural Study Using a Tandem Affinity Tag Method at JoVE.com

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

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.

Affinity purification approaches have been successful in isolating native complexes for proteomic characterization. Structural heterogeneity and a degree ...

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Nanyang Technological University

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Biology

9.5K Views.  Brandeis University. 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. 

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

Protein Complex Affinity Capture from Cryomilled Mammalian Cells

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this technique is also frequently referred to as immunoprecipitation. Affinity capture can be applied in a bench-scale and in a high-throughput context. When coupled with protein mass spectrometry, affinity capture has proven to be a workhorse of interactome analysis. Although there are potentially many ways to execute the numerous steps involved, the following protocols implement our favored methods. Two features are distinctive: the use of cryomilled cell powder to produce cell extracts, and antibody-coupled paramagnetic beads as the affinity medium. In many cases, we have obtained superior results to those obtained with more conventional affinity capture practices. Cryomilling avoids numerous problems associated with other forms of cell breakage. It provides efficient breakage of the material, while avoiding denaturation issues associated with heating or foaming. It retains the native protein concentration up to the point of extraction, mitigating macromolecular dissociation. It reduces the time extracted proteins spend in solution, limiting deleterious enzymatic activities, and it may reduce the non-specific adsorption of proteins by the affinity medium. Micron-scale magnetic affinity media have become more commonplace over the last several years, increasingly replacing the traditional agarose- and Sepharose-based media. Primary benefits of magnetic media include typically lower non-specific protein adsorption; no size exclusion limit because protein complex binding occurs on the bead surface rather than within pores; and ease of manipulation and handling using magnets.

Here we describe protocols to disrupt mammalian cells by solid-state milling at a cryogenic temperature, produce a cell extract from the resulting cell powder, and isolate protein complexes of interest by affinity capture upon antibody-coupled micron-scale paramagnetic beads.

Affinity capture is an effective technique for isolating endogenous protein complexes for further study. When used in conjunction with an antibody, this ...

Biochemistry

Tandem Affinity Purification of Protein Complexes from Eukaryotic Cells

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo identification of novel subunits and post-translational modifications. Bacterial systems allow for the expression and purification of a wide variety of single polypeptides and protein complexes. However, this system does not enable the purification of protein subunits that contain post-translational modifications (e.g., phosphorylation and acetylation), and the identification of novel regulatory subunits that are only present/expressed in the eukaryotic system. Here, we provide a detailed description of a novel, robust, and efficient tandem affinity purification (TAP) method using STREP- and FLAG-tagged proteins that facilitates the purification of protein complexes with transiently or stably expressed epitope-tagged proteins from eukaryotic cells. This protocol can be applied to characterize protein complex functionality, to discover post-translational modifications on complex subunits, and to identify novel regulatory complex components by mass spectrometry. Notably, this TAP method can be applied to study protein complexes formed by eukaryotic or pathogenic (viral and bacterial) components, thus yielding a wide array of downstream experimental opportunities. We propose that researchers working with protein complexes could utilize this approach in many different ways.

We describe here a novel, robust, and efficient tandem affinity purification (TAP) method for the expression, isolation, and characterization of protein complexes from eukaryotic cells. This protocol could be utilized for the biochemical characterization of discrete complexes as well as the identification of novel interactors and post-translational modifications that regulate their function.

The purification of active protein-protein and protein-nucleic acid complexes is crucial for the characterization of enzymatic activities and de novo ...

Resolving Affinity Purified Protein Complexes by Blue Native PAGE and Protein Correlation Profiling

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological processes. Affinity purification coupled to mass spectrometry (AP-MS) has become the method of choice for identifying protein-protein interactions. However, conventional AP-MS studies provide information on protein interactions, but the organizational information is lost. To address this issue, we developed a strategy to unravel the distinct functional assemblies a protein might be involved in, by resolving affinity-purified protein complexes prior to their characterization by mass spectrometry. Protein complexes isolated through affinity purification of a bait protein using an epitope tag and competitive elution are separated through blue native electrophoresis. Comparison of protein migration profiles through correlation profiling using quantitative mass spectrometry allows assignment of interacting proteins to distinct molecular entities. This method is able to resolve protein complexes of close molecular weights that might not be resolved by traditional chromatographic techniques such as gel filtration. With little more work than conventional AP-geLC-MS/MS, we demonstrate this strategy may in many cases be adequate for obtaining protein complex topological information concomitantly to identifying protein interactions.

Here we present protocols for affinity purification of protein complexes and their separation by blue native PAGE, followed by protein correlation profiling using label free quantitative mass spectrometry. This method is useful to resolve interactomes into distinct protein complexes.

Most proteins act in association with others; hence, it is crucial to characterize these functional units in order to fully understand biological ...

Affinity Purification of Chloroplast Translocon Protein Complexes Using the TAP Tag

Chloroplast biogenesis requires the import of thousands of nucleus-encoded proteins into the plastid. The import of these proteins depends on the translocon at the outer (TOC) and inner (TIC) chloroplast membranes. The TOC and TIC complexes are multimeric and probably contain yet unknown components. One of the main goals in the field is to establish the complete inventory of TOC and TIC components. For the isolation of TOC-TIC complexes and the identification of new components, the preprotein receptor TOC159 has been modified N-terminally by the addition of the tandem affinity purification (TAP) tag resulting in TAP-TOC159. The TAP-tag is designed for two sequential affinity purification steps (hence "tandem affinity"). The TAP-tag used in these studies consists of a N-terminal IgG-binding domain derived from Staphylococcus aureus Protein A (ProtA) followed by a calmodulin-binding peptide (CBP). Between these two affinity tags, a tobacco etch virus (TEV) protease cleavage site has been included. Therefore, TEV protease can be used for gentle elution of TOC159-containing complexes after binding to IgG beads. In the protocol presented here, the second Calmodulin-affinity purification step was omitted. The purification protocol starts with the preparation and solubilization of total cellular membranes. After the detergent-treatment, the solubilized membrane proteins are incubated with IgG beads for the immunoisolation of TAP-TOC159-containing complexes. Upon binding and extensive washing, TAP-TOC159 containing complexes are cleaved and released from the IgG beads using the TEV protease whereby the S. aureus IgG-binding domain is removed. Western blotting of the isolated TOC159-containing complexes can be used to confirm the presence of known or suspected TOC and TIC proteins. More importantly, the TOC159-containing complexes have been used successfully to identify new components of the TOC and TIC complexes by mass spectrometry. The protocol that we present potentially allows the efficient isolation of any membrane-bound protein complex to be used for the identification of yet unknown components by mass spectrometry.

We here present a proven and tested protocol for the purification of chloroplast protein import complexes (TOC-TIC complex) using the TAP-tag. The one-step affinity-isolation protocol can potentially be applied to any protein and be used to identify new interaction partners by mass spectrometry.

Chloroplast biogenesis requires the import of thousands of nucleus-encoded proteins into the plastid. The import of these proteins depends on the ...

Optimizing FPLC for Efficient Purification of His-Tagged FEN1 Protein

Affinity Purification of a 6X-His-Tagged Protein using a Fast Protein Liquid Chromatography System

Functional characterization of proteins requires them to be expressed and purified in substantial amounts with high purity to perform biochemical assays. The Fast Protein Liquid Chromatography (FPLC) system allows high-resolution separation of complex protein mixtures. By adjusting various parameters in FPLC, such as selecting the appropriate purification matrix, regulating the protein sample's temperature, and managing the sample's flow rate onto the matrix and the elution rate, it is possible to ensure the protein's stability and functionality. In this protocol, we will demonstrate the versatility of the FPLC system to purify 6X-His-tagged flap endonuclease 1 (FEN1) protein, produced in bacterial cultures. To improve protein purification efficiency, we will focus on multiple considerations, including proper column packing and preparation, sample injection using a sample loop, flow rate of sample application to the column, and sample elution parameters. Finally, the chromatogram will be analyzed to identify fractions containing high yields of protein and considerations for proper recombinant protein long-term storage. Optimizing protein purification methods is crucial for improving the precision and reliability of protein analysis.

Author Spotlight: Optimizing Affinity Chromatography for His-Tagged FEN1 Protein

This article provides a procedure for the affinity purification of a human recombinant protein, flap endonuclease 1 (FEN1), which has been labeled with a 6X-histidine tag. The protocol involves the utilization of two distinct immobilized metal ion columns for the purification of the tagged protein.

Functional characterization of proteins requires them to be expressed and purified in substantial amounts with high purity to perform biochemical assays. ...

Identification of Protein Interacting Partners Using Tandem Affinity Purification

A critical and often limiting step in understanding the function of host and viral proteins is the identification of interacting cellular or viral protein partners. There are many approaches that allow the identification of interacting partners, including the yeast two hybrid system, as well as pull down assays using recombinant proteins and immunoprecipitation of endogenous proteins followed by mass spectrometry identification1. Recent studies have highlighted the utility of double-affinity tag mediated purification, coupled with two specific elution steps in the identification of interacting proteins. This approach, termed Tandem Affinity Purification (TAP), was initially used in yeast2,3 but more recently has been adapted to use in mammalian cells4-8.
As proof-of-concept we have established a tandem affinity purification (TAP) method using the well-characterized eukaryotic translation initiation factor eIF4E9,10.The cellular translation factor eIF4E is a critical component of the cellular eIF4F complex involved in cap-dependent translation initiation10. The TAP tag used in the current study is composed of two Protein G units and a streptavidin binding peptide separated by a Tobacco Etch Virus (TEV) protease cleavage sequence. The TAP tag used in the current study is composed of two Protein G units and a streptavidin binding peptide separated by a Tobacco Etch Virus (TEV) protease cleavage sequence8. To forgo the need for the generation of clonal cell lines, we developed a rapid system that relies on the expression of the TAP-tagged bait protein from an episomally maintained plasmid based on pMEP4 (Invitrogen). Expression of tagged murine eIF4E from this plasmid was controlled using the cadmium chloride inducible metallothionein promoter.
Lysis of the expressing cells and subsequent affinity purification via binding to rabbit IgG agarose, TEV protease cleavage, binding to streptavidin linked agarose and subsequent biotin elution identified numerous proteins apparently specific to the eIF4E pull-down (when compared to control cell lines expressing the TAP tag alone). The identities of the proteins were obtained by excision of the bands from 1D SDS-PAGE and subsequent tandem mass spectrometry. The identified components included the known eIF4E binding proteins eIF4G and 4EBP-1. In addition, other components of the eIF4F complex, of which eIF4E is a component were identified, namely eIF4A and Poly-A binding protein. The ability to identify not only known direct binding partners as well as secondary interacting proteins, further highlights the utility of this approach in the characterization of proteins of unknown function.

Tandem affinity purification is a robust approach for the identification of protein binding partners. As proof of concept, this methodology was applied to the well-characterized translation initiation factor eIF4E to co-precipitate the host cell factors involved in translation initiation. This method is easily adapted to any cellular or viral protein.

A critical and often limiting step in understanding the function of host and viral proteins is the identification of interacting cellular or viral protein ...

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the identification of the subunit composition of multi-protein complexes can provide insight into gene function and enhance understanding of biological systems1, 2. Physical interactions can be mapped with high confidence vialarge-scale isolation and characterization of endogenous protein complexes under near-physiological conditions based on affinity purification of chromosomally-tagged proteins in combination with mass spectrometry (APMS). This approach has been successfully applied in evolutionarily diverse organisms, including yeast, flies, worms, mammalian cells, and bacteria1-6. In particular, we have generated a carboxy-terminal Sequential Peptide Affinity (SPA) dual tagging system for affinity-purifying native protein complexes from cultured gram-negative Escherichia coli, using genetically-tractable host laboratory strains that are well-suited for genome-wide investigations of the fundamental biology and conserved processes of prokaryotes1, 2, 7. Our SPA-tagging system is analogous to the tandem affinity purification method developed originally for yeast8, 9, and consists of a calmodulin binding peptide (CBP) followed by the cleavage site for the highly specific tobacco etch virus (TEV) protease and three copies of the FLAG epitope (3X FLAG), allowing for two consecutive rounds of affinity enrichment. After cassette amplification, sequence-specific linear PCR products encoding the SPA-tag and a selectable marker are integrated and expressed in frame as carboxy-terminal fusions in a DY330 background that is induced to transiently express a highly efficient heterologous bacteriophage lambda recombination system10. Subsequent dual-step purification using calmodulin and anti-FLAG affinity beads enables the highly selective and efficient recovery of even low abundance protein complexes from large-scale cultures. Tandem mass spectrometry is then used to identify the stably co-purifying proteins with high sensitivity (low nanogram detection limits).
Here, we describe detailed step-by-step procedures we commonly use for systematic protein tagging, purification and mass spectrometry-based analysis of soluble protein complexes from E. coli, which can be scaled up and potentially tailored to other bacterial species, including certain opportunistic pathogens that are amenable to recombineering. The resulting physical interactions can often reveal interesting unexpected components and connections suggesting novel mechanistic links. Integration of the PPI data with alternate molecular association data such as genetic (gene-gene) interactions and genomic-context (GC) predictions can facilitate elucidation of the global molecular organization of multi-protein complexes within biological pathways. The networks generated for E. coli can be used to gain insight into the functional architecture of orthologous gene products in other microbes for which functional annotations are currently lacking.

Identification of Protein Complexes in Escherichia coli using Sequential Peptide Affinity Purification in Combination with Tandem Mass Spectrometry

Affinity purification of tagged proteins in combination with mass spectrometry (APMS) is a powerful method for the systematic mapping of protein interaction networks and for investigating the mechanistic basis of biological processes. Here, we describe an optimized sequential peptide affinity (SPA) APMS procedure developed for the bacterium Escherichia coli that can be used to isolate and characterize stable multi-protein complexes to near homogeneity even starting from low copy numbers per cell.

Since most cellular processes are mediated by macromolecular assemblies, the systematic identification of protein-protein interactions (PPI) and the ...

The MultiBac Protein Complex Production Platform at the EMBL

Proteomics research revealed the impressive complexity of eukaryotic proteomes in unprecedented detail. It is now a commonly accepted notion that proteins in cells mostly exist not as isolated entities but exert their biological activity in association with many other proteins, in humans ten or more, forming assembly lines in the cell for most if not all vital functions.1,2 Knowledge of the function and architecture of these multiprotein assemblies requires their provision in superior quality and sufficient quantity for detailed analysis. The paucity of many protein complexes in cells, in particular in eukaryotes, prohibits their extraction from native sources, and necessitates recombinant production. The baculovirus expression vector system (BEVS) has proven to be particularly useful for producing eukaryotic proteins, the activity of which often relies on post-translational processing that other commonly used expression systems often cannot support.3 BEVS use a recombinant baculovirus into which the gene of interest was inserted to infect insect cell cultures which in turn produce the protein of choice. MultiBac is a BEVS that has been particularly tailored for the production of eukaryotic protein complexes that contain many subunits.4 A vital prerequisite for efficient production of proteins and their complexes are robust protocols for all steps involved in an expression experiment that ideally can be implemented as standard operating procedures (SOPs) and followed also by non-specialist users with comparative ease. The MultiBac platform at the European Molecular Biology Laboratory (EMBL) uses SOPs for all steps involved in a multiprotein complex expression experiment, starting from insertion of the genes into an engineered baculoviral genome optimized for heterologous protein production properties to small-scale analysis of the protein specimens produced.5-8 The platform is installed in an open-access mode at EMBL Grenoble and has supported many scientists from academia and industry to accelerate protein complex research projects.

Protein complexes catalyze key cellular functions. Detailed functional and structural characterization of many essential complexes requires recombinant production. MultiBac is a baculovirus/insect cell system particularly tailored for expressing eukaryotic proteins and their complexes. MultiBac was implemented as an open-access platform, and standard operating procedures developed to maximize its utility.

Proteomics  research revealed the impressive complexity of eukaryotic proteomes in  unprecedented detail. It is now a commonly accepted notion that ...

One-step Purification of Twin-Strep-tagged Proteins and Their Complexes on Strep-Tactin Resin Cross-linked With Bis(sulfosuccinimidyl) Suberate (BS3)

Affinity purification of Strep-tagged fusion proteins on resins carrying an engineered streptavidin (Strep-Tactin) has become a widely used method for isolation of protein complexes under physiological conditions. Fusion proteins containing two copies of Strep-tag II, designated twin-Strep-tag or SIII-tag, have the advantage of higher affinity for Strep-Tactin compared to those containing only a single Strep-tag, thus allowing more efficient protein purification. However, this advantage is offset by the fact that elution of twin-Strep-tagged proteins with biotin may be incomplete, leading to low protein recovery. The recovery can be dramatically improved by using denaturing elution with sodium dodecyl sulfate (SDS), but this leads to sample contamination with Strep-Tactin released from the resin, making the assay incompatible with downstream proteomic analysis. To overcome this limitation, we have developed a method whereby resin-coupled tetramer of Strep-Tactin is first stabilized by covalent cross-linking with Bis(sulfosuccinimidyl) suberate (BS3) and the resulting cross-linked resin is then used to purify target protein complexes in a single batch purification step. Efficient elution with SDS ensures good protein recovery, while the absence of contaminating Strep-Tactin allows downstream protein analysis by mass spectrometry. As a proof of concept, we describe here a protocol for purification of SIII-tagged viral protein VPg-Pro from nuclei of virus-infected N. benthamiana plants using the Strep-Tactin polymethacrylate resin cross-linked with BS3. The same protocol can be used to purify any twin-Strep-tagged protein of interest and characterize its physiological binding partners.

One-step Purification of Twin-Strep-tagged Proteins and Their Complexes on Strep-Tactin Resin Cross-linked With Bissulfosuccinimidyl Suberate BS3

A method is described for efficient purification of twin-Strep-tagged fusion proteins and their specific complexes on modified streptavidin (Strep-Tactin) resin covalently cross-linked with Bis(sulfosuccinimidyl) suberate (BS3). The method has the advantages of fast speed, good target protein recovery and high purity, and is compatible with subsequent analysis by mass spectrometry.

Affinity purification of Strep-tagged fusion proteins on resins carrying an engineered streptavidin (Strep-Tactin) has become a widely used method for ...

Combining Chemical Cross-linking and Mass Spectrometry of Intact Protein Complexes to Study the Architecture of Multi-subunit Protein Assemblies

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is therefore fundamental for the understanding of cellular processes. However, many protein complexes are dynamic assemblies and are not accessible by conventional structural techniques. Mass spectrometry contributes to the structural investigation of these assemblies, and particularly the combination of various mass spectrometric techniques delivers valuable insights into their structural arrangement.
In this article, we describe the application and combination of two complementary mass spectrometric techniques, namely chemical cross-linking coupled with mass spectrometry and native mass spectrometry. Chemical cross-linking involves the covalent linkage of amino acids in close proximity by using chemical reagents. After digestion with proteases, cross-linked di-peptides are identified by mass spectrometry and protein interactions sites are uncovered. Native mass spectrometry on the other hand is the analysis of intact protein assemblies in the gas phase of a mass spectrometer. It reveals protein stoichiometries as well as protein and ligand interactions. Both techniques therefore deliver complementary information on the structure of protein-ligand assemblies and their combination proved powerful in previous studies.

The architecture of protein complexes is essential for their function. Combining various mass spectrometric techniques proved powerful to study their assembly. We provide protocols for chemical cross-linking and native mass spectrometry and show how these complementary techniques help to elucidate the architecture of multi-subunit protein assemblies.

Proteins interact with their ligands to form active and dynamic assemblies which carry out various cellular functions. Elucidating these interactions is ...

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

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