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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#916;1, leu2&#916;0, met15&#916;0, ura3&#916;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&#160;</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&#160;</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 &#956;L for a silver stained gel and 5 &#956;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&#160;</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&#160;</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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JoVE Journal

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

Purification and Visualization of Influenza A Viral Ribonucleoprotein Complexes

The influenza A viral genome consists of eight negative-sense, single stranded RNA molecules, individually packed with multiple copies of the influenza A nucleoprotein (NP) into viral ribonulceoprotein particles (vRNPs). The influenza vRNPs are enclosed within the viral envelope. During cell entry, however, these vRNP complexes are released into the cytoplasm, where they gain access to the host nuclear transport machinery. In order to study the nuclear import of influenza vRNPs and the replication of the influenza genome, it is useful to work with isolated vRNPs so that other components of the virus do not interfere with these processes. Here, we describe a procedure to purify these vRNPs from the influenza A virus. The procedure starts with the disruption of the influenza A virion with detergents in order to release the vRNP complexes from the enveloped virion. The vRNPs are then separated from the other components of the influenza A virion on a 33-70% discontinuous glycerol gradient by velocity sedimentation. The fractions obtained from the glycerol gradient are then analyzed on via SDS-PAGE after staining with Coomassie blue. The peak fractions containing NP are then pooled together and concentrated by centrifugation. After concentration, the integrity of the vRNPs is verified by visualization of the vRNPs by transmission electron microscopy after negative staining. The glycerol gradient purification is a modification of that from Kemler et al. (1994)1, and the negative staining has been performed by Wu et al. (2007).2

The genome of the influenza A virus consists of eight separate complexes of RNA and proteins, termed viral ribonucleoprotein complexes (vRNPs). This paper describes the glycerol gradient purification and transmission electron microscopy visualization of influenza A vRNPs.

The influenza A viral genome consists of eight negative-sense, single stranded RNA molecules, individually packed with multiple copies of the influenza A ...

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

The Drosophila peripheral nervous system (PNS) is a powerful model for investigating the complex processes of neuronal development and dendrite morphogenesis at the functional and molecular levels. To aid in these analyses, we have developed a strategy for the isolation of a subclass of PNS neurons called dendritic arborization (da) neurons that have been widely used for studying dendrite morphogenesis1,2. These neurons are very difficult to isolate as a pure population, due in part to their extremely low occurrence and their difficult-to-reach location below the tough chitinous larval cuticle. Our newly developed method overcomes these challenges, and is based on a fast and specific cell enrichment using antibody-coated magnetic beads. For our magnetic bead sorting studies, we have used age-matched third instar larvae expressing a mouse CD8 tagged GFP fusion protein (UAS-mCD8-GFP)3 under the control of either the class IV dendritic arborization (da) neuron-specific pickpocket (ppk)-GAL4 driver4 or the control of the pan-da neuron-specific GAL421-7 driver5. Although this protocol has been optimized for isolating PNS cells which are attached to the inner wall of the larval cuticle, by varying a few parameters, the same protocol could be used to isolate many different cell types attached to the cuticle at larval or pupal stages of development (e.g. epithelia, muscle, oenocytes etc.), or other cell types from larval organs depending upon the GAL4-specific driver expression pattern. The RNA isolated by this method is of high quality and can be readily used for downstream genomic analyses such as microarray gene expression profiling studies. This approach offers a powerful new tool to perform studies on isolated Drosophila dendritic arborization (da) neurons thereby providing novel insights into the molecular mechanisms underlying dendrite morphogenesis.

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

In this video-article we present a method for the isolation and purification of Drosophila peripheral neurons using a fast magnetic bead assisted cell sorting strategy. RNA obtained from the isolated cells can be readily used for downstream applications including microarray analyses.

The Drosophila peripheral nervous system (PNS) is a powerful model for investigating the complex processes of neuronal development and dendrite ...

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an array of dynamic, multi-protein complexes1. To gain a mechanistic understanding of the various cellular processes, it is crucial to determine the structure of such protein complexes, and reveal how their structural organization dictates their function. Many aspects of multi-protein complexes are, however, difficult to characterize, due to their heterogeneous nature, asymmetric structure, and dynamics. Therefore, new approaches are required for the study of the tertiary levels of protein organization.
One of the emerging structural biology tools for analyzing macromolecular complexes is mass spectrometry (MS)2-5. This method yields information on the complex protein composition, subunit stoichiometry, and structural topology. The power of MS derives from its high sensitivity and, as a consequence, low sample requirement, which enables examination of protein complexes expressed at endogenous levels. Another advantage is the speed of analysis, which allows monitoring of reactions in real time. Moreover, the technique can simultaneously measure the characteristics of separate populations co-existing in a mixture. 
Here, we describe a detailed protocol for the application of structural MS to the analysis of large protein assemblies. The procedure begins with the preparation of gold-coated capillaries for nanoflow electrospray ionization (nESI). It then continues with sample preparation, emphasizing the buffer conditions which should be compatible with nESI on the one hand, and enable to maintain complexes intact on the other. We then explain, step-by-step, how to optimize the experimental conditions for high mass measurements and acquire MS and tandem MS spectra. Finally, we chart the data processing and analyses that follow. Rather than attempting to characterize every aspect of protein assemblies, this protocol introduces basic MS procedures, enabling the performance of MS and MS/MS experiments on non-covalent complexes. Overall, our goal is to provide researchers unacquainted with the field of structural MS, with knowledge of the principal experimental tools.

Mass spectrometry has proven to be a valuable tool for analyzing large protein complexes. This method enables insights into the composition, stoichiometry and overall architecture of multi-subunit assemblies. Here, we describe, step-by-step, how to perform a structural mass spectrometry analysis, and characterize macromolecular structures.

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an ...

Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) for Analysis of Multiprotein Complexes from Cellular Lysates

Multiprotein complexes (MPCs) play a crucial role in cell signalling, since most proteins can be found in functional or regulatory complexes with other proteins (Sali, Glaeser et al. 2003). Thus, the study of protein-protein interaction networks requires the detailed characterization of MPCs to gain an integrative understanding of protein function and regulation. For identification and analysis, MPCs must be separated under native conditions. In this video, we describe the analysis of MPCs by blue native polyacrylamide gel electrophoresis (BN-PAGE). BN-PAGE is a technique that allows separation of MPCs in a native conformation with a higher resolution than offered by gel filtration or sucrose density ultracentrifugation, and is therefore useful to determine MPC size, composition, and relative abundance (Sch&auml;gger and von Jagow 1991); (Sch&auml;gger, Cramer et al. 1994). By this method, proteins are separated according to their hydrodynamic size and shape in a polyacrylamide matrix. Here, we demonstrate the analysis of MPCs of total cellular lysates, pointing out that lysate dialysis is the crucial step to make BN-PAGE applicable to these biological samples. Using a combination of first dimension BN- and second dimension SDS-PAGE, we show that MPCs separated by BN-PAGE can be further subdivided into their individual constituents by SDS-PAGE. Visualization of the MPC components upon gel separation is performed by standard immunoblotting. As an example for MPC analysis by BN-PAGE, we chose the well-characterized eukaryotic 19S, 20S, and 26S proteasomes.

Blue Native Polyacrylamide Gel Electrophoresis BN-PAGE for Analysis of Multiprotein Complexes from Cellular Lysates

In this video, we describe the characterization of multiprotein complexes (MPCs) by blue native polyacrylamide gel electrophoresis (BN-PAGE). In a first dimension, dialyzed cellular lysates are separated by BN-PAGE to identify individual MPCs. In a second dimension SDS-PAGE, MPCs of interest are further subdivided to analyze their constituents by immunoblotting.

Multiprotein complexes (MPCs) play a crucial role in cell signalling, since most proteins can be found in functional or regulatory complexes with other ...

Orthogonal Protein Purification Facilitated by a Small Bispecific Affinity Tag

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a demand for general methods that do not require target protein specific optimization1 . To achieve this, purification tags that genetically can be fused to the gene of interest are commonly used2 . The most widely used affinity handle is the hexa-histidine tag, which is suitable for purification under both native and denaturing conditions3 . The metabolic burden for producing the tag is low, but it does not provide as high specificity as competing affinity chromatography based strategies1,2.
Here, a bispecific purification tag with two different binding sites on a 46 amino acid, small protein domain has been developed. The albumin-binding domain is derived from Streptococcal protein G and has a strong inherent affinity to human serum albumin (HSA). Eleven surface-exposed amino acids, not involved in albumin-binding4 , were genetically randomized to produce a combinatorial library. The protein library with the novel randomly arranged binding surface (Figure 1) was expressed on phage particles to facilitate selection of binders by phage display technology. Through several rounds of biopanning against a dimeric Z-domain derived from Staphylococcal protein A5, a small, bispecific molecule with affinity for both HSA and the novel target was identified6 .
The novel protein domain, referred to as ABDz1, was evaluated as a purification tag for a selection of target proteins with different molecular weight, solubility and isoelectric point. Three target proteins were expressed in Escherishia coli with the novel tag fused to their N-termini and thereafter affinity purified. Initial purification on either a column with immobilized HSA or Z-domain resulted in relatively pure products. Two-step affinity purification with the bispecific tag resulted in substantial improvement of protein purity. Chromatographic media with the Z-domain immobilized, for example MabSelect SuRe, are readily available for purification of antibodies and HSA can easily be chemically coupled to media to provide the second matrix.
This method is especially advantageous when there is a high demand on purity of the recovered target protein. The bifunctionality of the tag allows two different chromatographic steps to be used while the metabolic burden on the expression host is limited due to the small size of the tag. It provides a competitive alternative to so called combinatorial tagging where multiple tags are used in combination1,7.

A novel and highly efficient two-step affinity chromatography protocol has been developed and is described in detail. The method is based on a small purification tag with two inherent affinities and is applicable to a wide range of target proteins with different properties.

Due to the high costs associated with purification of recombinant proteins the protocols need to be rationalized. For high-throughput efforts there is a ...

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 ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

GST-His purification: A Two-step Affinity Purification Protocol Yielding Full-length Purified Proteins

Key assays in enzymology for the biochemical characterization of proteins in vitro necessitate high concentrations of the purified protein of interest. Protein purification protocols should combine efficiency, simplicity and cost effectiveness1. Here, we describe the GST-His method as a new small-scale affinity purification system for recombinant proteins, based on a N-terminal Glutathione Sepharose Tag (GST)2,3 and a C-terminal 10xHis tag4, which are both fused to the protein of interest. The latter construct is used to generate baculoviruses, for infection of Sf9 infected cells for protein expression5. GST is a rather long tag (29 kDa) which serves to ensure purification efficiency. However, it might influence physiological properties of the protein. Hence, it is subsequently cleaved off the protein using the PreScission enzyme6. In order to ensure maximum purity and to remove the cleaved GST, we added a second affinity purification step based on the comparatively small His-Tag. Importantly, our technique is based on two different tags flanking the two ends of the protein, which is an efficient tool to remove degraded proteins and, therefore, enriches full-length proteins. The method presented here does not require an expensive instrumental setup, such as FPLC. Additionally, we incorporated MgCl2 and ATP washes to remove heat shock protein impurities and nuclease treatment to abolish contaminating nucleic acids. In summary, the combination of two different tags flanking the N- and the C-terminal and the capability to cleave off one of the tags, guaranties the recovery of a highly purified and full-length protein of interest.

In the present protocol, we demonstrate a highly efficient and cost-effective small-scale protein purification method, which allows purification of recombinant proteins by uniquely combining a cleavable GST-tag and a small His-tag.

Key assays in enzymology for the biochemical characterization of proteins in vitro necessitate high concentrations of the purified protein of interest. ...

Protein Purification-free Method of Binding Affinity Determination by Microscale Thermophoresis

Quantitative characterization of protein interactions is essential in practically any field of life sciences, particularly drug discovery. Most of currently available methods of KD determination require access to purified protein of interest, generation of which can be time-consuming and expensive. We have developed a protocol that allows for determination of binding affinity by microscale thermophoresis (MST) without purification of the target protein from cell lysates. The method involves overexpression of the GFP-fused protein and cell lysis in non-denaturing conditions. Application of the method to STAT3-GFP transiently expressed in HEK293 cells allowed to determine for the first time the affinity of the well-studied transcription factor to oligonucleotides with different sequences. The protocol is straightforward and can have a variety of application for studying interactions of proteins with small molecules, peptides, DNA, RNA, and proteins.

Microscale thermophoresis (MST) can be widely used for determination of binding affinity without purification of the target protein from cell lysates. The protocol involves overexpression of the GFP-fused protein, cell lysis in non-denaturing conditions, and detection of MST signal in the presence of varying concentrations of the ligand.

Quantitative characterization of protein  interactions is essential in practically any field of life sciences,  particularly drug discovery. Most of ...

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

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Chapters in this video

Purification and Visualization of Influenza A Viral Ribonucleoprotein Complexes

Isolation and Purification of Drosophila Peripheral Neurons by Magnetic Bead Sorting

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) for Analysis of Multiprotein Complexes from Cellular Lysates

Orthogonal Protein Purification Facilitated by a Small Bispecific Affinity Tag

Identification of Protein Interacting Partners Using Tandem Affinity Purification

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

High-throughput Purification of Affinity-tagged Recombinant Proteins

GST-His purification: A Two-step Affinity Purification Protocol Yielding Full-length Purified Proteins

Protein Purification-free Method of Binding Affinity Determination by Microscale Thermophoresis