Name: identification of protein interacting partners using tandem affinity purification
Uploaded: 2012-02-25T14:00:00
Duration: 10 min 2 s
Description: Watch this Scientific Journal Video about Identification of Protein Interacting Partners Using Tandem Affinity Purification at JoVE.com

This research was funded by a Wellcome Trust Senior Fellowship awarded to Dr. Ian Goodfellow.

1. Generation of cell lines: pMEP4 transfection / expression
<ol>
	<li>Transfect cells with the pMEP4 expression vector and select with 100 &mu;g of hygromycin B /ml (Roche) until all mock transfected cells have been killed off. The pMEP4 plasmid is maintained episomally so there is no need to isolate specific clones.</li>
	<li>Cells containing the pMEP4 vector can be induced by treatment with 10 &mu;M CdCl2 for 16 hours. Expression and inducibility should be confirmed on a small scale before amplification of the cell lines. The TAP-tagged protein can be readily detected as the protein G domains bind to antibodies from almost all species. For large-scale purifications typically 10 confluent 175 cm2 flasks of cells are required. This equates to approximately 2 X108 expressing cells.</li>
</ol>
2. Cell lysate preparation
<ol>
	<li>Scrape cells into PBS and combine into a single 50 ml tube. Spin 1200X g for 5 minutes (this should result in 2-3 ml of packed cells to start with, which reduces to about 1.5 ml after washing). </li>
	<li>Wash the cells three times in ice-cold PBS (50 mls each time).</li>
	<li>Lyse cells in 5 ml lysis buffer (50 mM Tris-HCl (pH 7.5), 125 mM NaCl, 5 % Glycerol, 0.2 % NP-40, 1.5 mM MgCl2, 25 mM NaF, 1 mM Na3VO4 and protease inhibitor). Note: NaF, Na3VO4 and protease inhibitors should be added fresh. Pipette up and down 10 times before leaving for 5 minutes on ice. Syringe up and down 5-10 times using a blunt needle before leaving again on ice for 5 minutes on ice. Repeat syringing and leave for a further 5 minutes on ice.</li>
	<li>Freeze-thaw the lysate twice (liquid N2 / or dry ice and ethanol). Do not allow sample to reach greater than 4&deg;C. Note: you can store this sample at -80&deg;C until you have time to process.</li>
	<li>Aliquot sample into 1.5 ml tubes and remove unlysed cells and debris by centrifugation (10 minutes at 4&deg;C, 16,000X g). </li>
	<li>Recover supernatant, combine in a 15 ml falcon tube and (optionally) pass through a 0.45 &mu;m filter before taking a 20 &mu;l sample for protein yield quantification. Remove a further 50 &mu;l aliquot for subsequent analysis (Sample 1).</li>
</ol>
3. Binding to Rabbit IgG-agarose
(Note: All spins should be carried out at 1200X g in a refrigerated centrifuge at 4&deg;C for 1 minute, unless stated otherwise)
<ol>
	<li>Gently resuspend the rabbit IgG-agarose solution (Sigma-Aldrich) by swirling bottle. Take 380 &mu;l of IgG agarose (using a cut 1ml pipette tip) and remove shipping/preservative solution by centrifugation for 1 minute. Wash the agarose three times in chilled lysis buffer (4&deg;C) using centrifugation to clarify. The final yield of agarose should be about 250 &mu;l of packed beads.</li>
	<li>Add the cleared cell lysate from 2.6 (in a 15 ml falcon tube) to the washed rabbit-IgG agarose and incubate for 3 hours (or overnight) at 4&deg;C using a rotating mixer. </li>
	<li>Spin down the agarose beads for 5 minutes at 4&deg;C and remove the supernatant for subsequent analysis (Sample 2). Note: The TAP tagged protein should now be associated with the beads.</li>
</ol>
4. TEV protease cleavage
(Note: All spins should be carried out at 1200X g in a refrigerated centrifuge at 4oC for 1 minute, unless stated otherwise)
<ol>
	<li>Wash the rabbit IgG agarose beads three times in chilled lysis buffer (4&deg;C) using centrifugation to clarify (note: the lysis buffer should not contain protease inhibitors). Care should be taken not to remove or lose any beads during the washing steps.</li>
	<li>Wash the beads a further two times with TEV-protease cleavage buffer (10 mM Tris-HCl (pH 7.5), 100 mM NaCl, and 0.2 % NP-40) using centrifugation to clarify. After the last wash carefully remove all liquid from the beads.</li>
	<li>Prepare a TEV protease cleavage mix; for each sample include 467.5 &mu;l H2O, 25 &mu;l 20x TEV Buffer (Invitrogen), 5 &mu;l 0.1M DTT and 2.5 &mu;l (25 U) TEV Protease (Invitrogen). Add 500 &mu;l of this TEV cleavage mix to each sample of packed beads and transfer the entire mix to a 1.7 ml pre-lubricated tube (Costar). Prior to incubation remove a 30 &mu;l aliquot for subsequent analysis (Sample 3).</li>
	<li>Incubate the TEV protease reaction overnight at 4&deg;C using a rotating mixer. Note that shorter incubations times may be used depending on the nature of the bait protein and the accessibility of the TEV protease cleavage site but the minimal incubation time should be empirically determined.</li>
</ol>
5. Binding to Ultralink Immobilized Streptavidin Plus beads
(Note: All spins should be carried out at 1200X g in a refrigerated centrifuge at 4&deg;C for 1 minute, unless stated otherwise)
<ol>
	<li>Centrifuge the TEV cleavage reaction containing the rabbit IgG agarose beads for 5 minutes at 1200X g (4&deg;C). Remove a 20 &mu;l aliquot of the clarified supernatant for analysis (Sample 4)</li>
	<li>Transfer the remaining supernatant (roughly 480 &mu;l) to a fresh 1.5 ml pre-lubricated tube (Costar) and leave this on ice. Do not throw away this supernatant &#x2013; this contains your TEV cleaved bait protein and any associated binding proteins.</li>
	<li>Add 500 &mu;l of chilled lysis buffer (Section 2.3) to the remaining rabbit IgG agarose beads and resuspend. Centrifuge this mixture to clarify the beads before removing the supernatant and combining it with that from step 5.2. Leave this tube on ice (it should now contain ~980 &mu;l of sample).</li>
	<li>Retain the rabbit IgG agarose beads for subsequent analysis (Sample 5). Note: if you run this sample on an SDS-PAGE gel (or western blot) it will give a lot of background due to the presence of the IgG heavy and light chains.</li>
	<li>Meanwhile gently resuspend the Ultralink Immobilized Streptavidin Plus agarose beads (Pierce) by swirling the bottle. Using a 200 &mu;l pipette tip with the end cut, aliquot 70 &mu;l of streptavidin beads per sample into pre-lubricated 1.5 ml tubes and remove shipping/preservative solution by centrifugation. Note: these beads are very small and so "duck-billed" or flat/narrow-ended tips can be used to minimize bead loss during washing.</li>
	<li>Wash the streptavidin beads three times in chilled lysis buffer (4&deg;C) using centrifugation to clarify. This should leave approximately 45-50 &mu;l of packed beads per tube. </li>
	<li>Transfer the TEV-protease cleavage supernatant (i.e. the 980 &mu;l of sample from step 5.3) to the tube containing the washed streptavidin beads and incubate for 3 hours (or overnight) at 4&deg;C using a rotating mixer.</li>
	<li>Spin down the streptavidin beads for 5 minutes at 4&deg;C and remove the supernatant for subsequent analysis (Sample 6). Wash the streptavidin beads three times in chilled lysis buffer (4&deg;C) using centrifugation to clarify. After the final wash, carefully remove all remaining wash buffer.</li>
</ol>
6. Biotin elution of the streptavidin binding peptide and bait protein
(Note: All spins should be carried out at 1200X g in a refrigerated centrifuge at 4&deg;C for 1 minute, unless stated otherwise)
<ol>
	<li>Elute tagged proteins from the streptavidin beads by adding 500 &mu;l of PBS:Biotin (1 mM D-biotin) and incubating at 4&deg;C for 3 hours (or overnight) using a rotating mixer.</li>
	<li>Spin down the streptavidin beads for 5 minutes at 4&deg;C and remove the supernatant to a 1.5 ml pre-lubricated tube (Note: this is your final sample/ elution (Sample 7). </li>
	<li>Add another 500 &mu;l Biotin:PBS to the remaining streptavidin beads and incubate at 4&deg;C for 2-3 hours (or overnight) using a rotating mixer.</li>
	<li>Repeat step 6.2 and combine the two elutions (Sample 7). This final elution can be stored at -80&deg;C.</li>
	<li>To assess the efficiency of the biotin elution, freeze the remaining streptavidin beads and boil later in SDS sample buffer (Sample 8) (recommendations: LaneMarker 5x reducing sample buffer from Fisher). </li>
</ol>
7. Protein concentration
<ol>
	<li>The final eluate (Sample 7) needs to be concentrated prior to analysis because of the low protein concentration and relatively high volume. This can achieved using low molecular weight (&lt;5 KDa) Vivaspin spin columns (Vivascience). Aim to concentrate the volume down to less than 100 &mu;l. This final sample can be boiled and stored in protein sample buffer (recommendations: LaneMarker 5x reducing sample buffer from Fisher).</li>
</ol>
8. Analysis
<ol>
	<li>Samples 1-8 can be analyzed by western blot to determine the efficiency of pull down. Note: Most secondary antibodies will cross-react with the Protein G domains of the fusion protein but this is removed after TEV protease cleavage.</li>
	<li>Sample 7 can be analyzed by 1D or 2D SDS-PAGE. Staining can be performed using silver or Coomassie (recommendations: SilverQuest Silver staining and Colloidal Blue Coomassie staining kits from Invitrogen). Protein identification can be performed using mass spectrometry.</li>
</ol>
9. Representative Results
An example of 1D SDS-PAGE analysis of the final elution (Sample 7) from this protocol to identify binding partners of TAP tagged eIF4E is provided in Figure 2. This representative gel reflects the complex and abundant nature of eIF4E interactions with other proteins in the cell. Comparison with the negative control also shown in Figure 2, generated from a cell line expressing only the TAP tag, illustrates the specificity of this eIF4E baited pull-down. In this instance 15% of the concentrated final elution (Sample 7) was analyzed by 1D SDS-PAGE using commercially available pre-cast gradient gels before being stained using the Silverquest silver staining kit from Invitrogen. 
Typically 50-85% of the remaining concentrated final elution (Sample 7) is analyzed with Colloidal Coomassie stain (Invitrogen). Samples (gel slices/bands) from the entire lane were then extracted and analysed by mass-spectrometry.
Proteins identified in the eIF4E pull-down were filtered against the binding partners from the negative control to identify non-specific binding partners. The final proteins identified using this process of TAP tagging eIF4E can be seen in Figure 2, which is representative of the normal proteins identified using this technique. In addition, a number of the eIF3 subunits were also identified (data not shown). 
<img src="/files/ftp_upload/3643/3643fig1.jpg" alt="Figure 1" /> 
Figure 1. Schematic of the tandem affinity purification procedure. The six step tandem affinity purification (TAP) protocol involves cell line generation, cell lysis, Rabbit IgG agarose immuno-precipitation, TEV protease cleavage, streptavidin bead affinity purification and finally biotin elution.
<img src="/files/ftp_upload/3643/3643fig2.jpg" alt="Figure 2" /> 
Figure 2. Tandem affinity purification of the murine eIF4E protein. Interacting partners of the N terminally TAP tagged eIF4E were purified from eukaryotic HEK293 cells using the attached protocol. A 20% fraction of the final elution (Sample 7) was analysed by SDS-PAGE on a pre-cast 4-12% gradient gel (TAP-eIF4E lane). An equivalent analysis was performed for the tag alone (TAP lane). Proteins were identified using silver staining. Proteins subsequently identified by mass spectrometry of the same sample are highlighted on this gel. 
Abbreviations: eIF4G; eukaryotic translation initiation factor 4 gamma, PABP; polyA binding protein, eIF4A: eukaryotic translation initiation factor 4 alpha, SBP-eIF4e; the remaining TAP bait protein containing the streptavidin binding peptide fused to the eukaryotic translation initiation factor 4E, 4EBPs; eukaryotic translation initiation factor 4E binding proteins, eIF4eNiF1l eukaryotic translation initiation factor 4E nuclear import factor 1, SBP; the remaining streptavidin binding peptide from the TEV unfused TAP peptide.

<ol>
	<li>Williamson, M. P., Sutcliffe, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein-protein+interactions.">Protein-protein interactions.</a> Biochem. Soc. Trans. 38 (4), 875-878 (2010).</li><li>Rigaut, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+generic+protein+purification+method+for+protein+complex+characterization+and+proteome+exploration.">A generic protein purification method for protein complex characterization and proteome exploration.</a> Nat. Biotechnol. 17 (10), 1030-1032 (1999).</li><li>Blackwell, C., Brown, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+application+of+tandem-affinity+purification+to+Candida+albicans.">The application of tandem-affinity purification to Candida albicans.</a> Methods Mol. Biol. 499, 133-148 (2009).</li><li>Cochrane, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+complex+formation+between+HIV+Rev+and+the+nucleosome+assembly+protein,+NAP1,+affects+Rev+function.">Stable complex formation between HIV Rev and the nucleosome assembly protein, NAP1, affects Rev function.</a> Virology. 388 (1), 103-111 (2009).</li><li>Fernandez, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+tandem+affinity+purification+of+PSD-95+recovers+core+postsynaptic+complexes+and+schizophrenia+susceptibility+proteins.">Targeted tandem affinity purification of PSD-95 recovers core postsynaptic complexes and schizophrenia susceptibility proteins.</a> Mol. Syst. Biol. 5, 269-269 (2009).</li><li>Gloeckner, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tandem+affinity+purification+of+protein+complexes+from+mammalian+cells+by+the+Strep&#47;FLAG+(SF)-TAP+tag.">Tandem affinity purification of protein complexes from mammalian cells by the Strep&#47;FLAG (SF)-TAP tag.</a> Methods Mol. Biol. 564, 359-372 (2009).</li><li>Holowaty, M. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Protein+profiling+with+Epstein-Barr+nuclear+antigen-1+reveals+an+interaction+with+the+herpesvirus-associated+ubiquitin-specific+protease+HAUSP/USP7.">Protein profiling with Epstein-Barr nuclear antigen-1 reveals an interaction with the herpesvirus-associated ubiquitin-specific protease HAUSP/USP7.</a> J. Biol. Chem. 278 (32), 29987-29994 (2003).</li><li>Burckstummer, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+efficient+tandem+affinity+purification+procedure+for+interaction+proteomics+in+mammalian+cells.">An efficient tandem affinity purification procedure for interaction proteomics in mammalian cells.</a> Nat. Methods. 3 (12), 1013-1019 (2006).</li><li>Goodfellow, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calicivirus+translation+initiation+requires+an+interaction+between+VPg+and+eIF+4+E.">Calicivirus translation initiation requires an interaction between VPg and eIF 4 E.</a> EMBO Rep. 6 (10), 968-972 (2005).</li><li>Goodfellow, I. G., Roberts, L. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Eukaryotic+initiation+factor+4E.">Eukaryotic initiation factor 4E.</a> Int. J. Biochem. Cell Biol. 40 (12), 2675-2680 (2008).</li></ol>

No conflicts of interest declared.

The TAP tagging technique illustrated here demonstrates a highly specific and stringent method for isolating the binding partners of bait proteins in eukaryotic cells. This approach can be applied to both cellular and viral proteins. To our knowledge, this is the first time such a technique has been applied to the translation initiation factor eIF4E. Identification of the known eIF4E binding proteins eIF4G and the 4EBPs using this technique confirms the validity of such an approach. In addition, the identification of the remaining component of the eIF4F complex, namely eIF4A, and PABP confirms that indirect interaction and tertiary complexes remain intact during the purification process. The identification of multiple isoforms of the canonical eIF4e binding proteins was also evident. These are described in more detail in Figure 2. 
With regards to the limitations of the approach, certain care should be taken with regards to the choice of bait protein and whether or not to place the TAG tag at the N- or C-terminus. It might be advisable to undertake a functional assay or examine the localization of the fusion protein by microscopy prior to full scale purification to ensure the tagged derivative is functional. Integral membrane or nuclear proteins may not necessarily be released by the relatively mild detergent conditions described in the lysis step. As with all immunoprecipitations and similar pull-down assays, modifications could be made to the nature and concentrations of the detergent in the lysis buffer to increase the efficiency of lysis. The ionic concentration of the lysis and wash buffers may also be modified to increase or decrease the stringency of the purification. It is also possible to perform the initial purification under standard mild conditions (described above) but subsequently divide the streptavidin beads (section 5.8) into 4-5 aliquots, which can subsequently be washed under increasing ionic conditions. This may enable the user to identify the optimal conditions that remove non-specific interacting proteins. The entire process may also be performed using transient transfection where the interacting partners are known and their presence or absence determined by subsequent western blot only. In this case we would suggest two 100cm2 dishes of cells each transfected with 8 &mu;g of expression plasmid. This modified procedure is of particular use when examining the ability of mutant derivatives of the TAP-fusion protein to interact with known binding partners.
Analyzing samples 1 through 8 on silver stained SDS-PAGE gels (or by western blot if an antibody is available) is an excellent way of troubleshooting, should the technique fail to generate acceptable results. The efficiency of each affinity based precipitation and specific elution can be analyzed using this systematic sampling approach. Samples one to eight should be taken during optimization of the protocol for each new bait.
The TAP tagging technique provides a powerful and robust alternative to other approaches such as GST pull-downs, yeast two hybrid assays etc. The double affinity purification and specific elution steps (TEV cleavage and biotin elution) provide specificity and stringency to be maintained at a high level throughout the purification procedure. Critically this technique can be applied to any protein and therefore represents an excellent method for identifying binding partners for a protein target of interest.

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
		</tr>
		<tr>
			<td>Hygromycin B</td>
			<td>Roche</td>
			<td>10843555001</td>
		</tr>
		<tr>
			<td>Rabbit IgG Agarose</td>
			<td>Sigma</td>
			<td>A2909</td>
		</tr>
		<tr>
			<td>AC-TEV Protease</td>
			<td>Invitrogen</td>
			<td>12575-015</td>
		</tr>
		<tr>
			<td>Protease inhibitor cocktail</td>
			<td>Calbiochem</td>
			<td>539134</td>
		</tr>
		<tr>
			<td>Ultralink Immobilized Streptavidin Plus beads </td>
			<td>Pierce</td>
			<td>53116</td>
		</tr>
		<tr>
			<td>Vivaspin 500 centrifugal concentrator (5KDa)</td>
			<td>Vivaspin</td>
			<td>V50112</td>
		</tr>
		<tr>
			<td>SilverQuest silver staining kit</td>
			<td>Invitrogen</td>
			<td>LC6070</td>
		</tr>
		<tr>
			<td>Novex Colloidal Blue Coomassie staining kit</td>
			<td>Invitrogen</td>
			<td>LC6025</td>
		</tr>
		<tr>
			<td>1.7ml prelubricated tubes</td>
			<td>Costar</td>
			<td>3207</td>
		</tr>
		<tr>
			<td>Microcapillary pipette tips</td>
			<td>VWR</td>
			<td>37001-150</td>
		</tr>
		<tr>
			<td>NuPage 4-12% Bis-Tris gradient gels</td>
			<td>Invitrogen</td>
			<td>NP0322BOX</td>
		</tr>
		<tr>
			<td>CdCl2</td>
			<td>Sigma</td>
			<td>202908</td>
		</tr>
		<tr>
			<td>5x SDS Sample Buffer</td>
			<td>Fisher</td>
			<td>PN39000</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Identification of Protein Interacting Partners Using Tandem Affinity Purification at JoVE.com

Identification of Protein Interacting Partners Using Tandem Affinity Purification

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-protein-interacting-partners-using-tandem-affinity

Research

JoVE Journal

Biology

Staining of Proteins in Gels with Coomassie G-250 without Organic Solvent and Acetic Acid

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents (Methanol, Ethanol or 2-Propanol) and acetic acid are used for fixation, staining and destaining of proteins in a gel after SDS-PAGE. To speed up the procedure, heating the staining solution in the microwave oven for a short time is frequently used. This usually results in evaporation of toxic or hazardous Methanol, Ethanol or 2-Propanol and a strong smell of acetic acid in the lab which should be avoided due to safety considerations. In a protocol originally published in two patent applications by E.M. Wondrak (US2001046709 (A1), US6319720 (B1)), an alternative composition of the staining solution is described in which no organic solvent or acid is used. The CBB is dissolved in bidistilled water (60-80mg of CBB G-250 per liter) and 35 mM HCl is added as the only other compound in the staining solution. The CBB staning of the gel is done after SDS-PAGE and thorough washing of the gel in bidistilled water. By heating the gel during the washing and staining steps, the process can be finished faster and no toxic or hazardous compunds are evaporating. The staining of proteins occurs already within 1 minute after heating the gel in staining solution and is fully developed after 15-30 min with a slightly blue background that is destained completely by prolonged washing of the stained gel in bidistilled water, without affecting the stained protein bands.

A short protocol for protein staining with Coomassie Brilliant Blue (CBB) G-250 in polyacrylamide gels is described without using organic solvents or acetic acid as in the classical staining procedures with CBB.

In classical protein staining protocols using Coomassie Brilliant Blue (CBB), solutions with high contents of toxic and flammable organic solvents ...

Purification of Hsp104, a Protein Disaggregase

Hsp104 is a hexameric AAA+ protein1 from yeast, which couples ATP hydrolysis to protein disaggregation2-10 (Fig. 1). This activity imparts two key selective advantages. First, renaturation of disordered aggregates by Hsp104 empowers yeast survival after various protein-misfolding stresses, including heat shock3,5,11,12. Second, remodeling of cross-beta amyloid fibrils by Hsp104 enables yeast to exploit myriad prions (infectious amyloids) as a reservoir of beneficial and heritable phenotypic variation13-22. Remarkably, Hsp104 directly remodels preamyloid oligomers and amyloid fibrils, including those comprised of the yeast prion proteins Sup35 and Ure223-30. This amyloid-remodeling functionality is a specialized facet of yeast Hsp104. The E. coli orthologue, ClpB, fails to remodel preamyloid oligomers or amyloid fibrils26,31,32.
Hsp104 orthologues are found in all kingdoms of life except, perplexingly, animals. Indeed, whether animal cells possess any enzymatic system that couples protein disaggregation to renaturation (rather than degradation) remains unknown33-35. Thus, we and others have proposed that Hsp104 might be developed as a therapeutic agent for various neurodegenerative diseases connected with the misfolding of specific proteins into toxic preamyloid oligomers and amyloid fibrils4,7,23,36-38. There are no treatments that directly target the aggregated species associated with these diseases. Yet, Hsp104 dissolves toxic oligomers and amyloid fibrils composed of alpha-synuclein, which are connected with Parkinson's Disease23 as well as amyloid forms of PrP39. Importantly, Hsp104 reduces protein aggregation and ameliorates neurodegeneration in rodent models of Parkinson's Disease23 and Huntington's disease38. Ideally, to optimize therapy and minimize side effects, Hsp104 would be engineered and potentiated to selectively remodel specific aggregates central to the disease in question4,7. However, the limited structural and mechanistic understanding of how Hsp104 disaggregates such a diverse repertoire of aggregated structures and unrelated proteins frustrates these endeavors30,40-42.
To understand the structure and mechanism of Hsp104, it is essential to study the pure protein and reconstitute its disaggregase activity with minimal components. Hsp104 is a 102kDa protein with a pI of ~5.3, which hexamerizes in the presence of ADP or ATP, or at high protein concentrations in the absence of nucleotide43-46. Here, we describe an optimized protocol for the purification of highly active, stable Hsp104 from E. coli. The use of E. coli allows simplified large-scale production and our method can be performed quickly and reliably for numerous Hsp104 variants. Our protocol increases Hsp104 purity and simplifies His6-tag removal compared to a previous purification method from E. coli47. Moreover, our protocol is more facile and convenient than two more recent protocols26,48.

Here, we describe a protocol for the purification of highly active Hsp104, a hexameric AAA+ protein from yeast, which couples ATP hydrolysis to protein disaggregation. This scheme exploits a His6-tagged construct for affinity purification from E. coli followed by anion-exchange chromatography, His6-tag removal with TEV protease, and size-exclusion chromatography.

Hsp104 is a hexameric AAA+ protein1 from yeast, which couples ATP hydrolysis to protein disaggregation2-10 (Fig. 1). This activity imparts two key ...

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 and Characterization of Protein Glycosylation using Specific Endo- and Exoglycosidases

Glycosylation, the addition of covalently linked sugars, is a major post-translational modification of proteins that can significantly affect processes such as cell adhesion, molecular trafficking, clearance, and signal transduction1-4. In eukaryotes, the most common glycosylation modifications in the secretory pathway are additions at consensus asparagine residues (N-linked); or at serine or threonine residues (O-linked) (Figure 1). Initiation of N-glycan synthesis is highly conserved in eukaryotes, while the end products can vary greatly among different species, tissues, or proteins. Some glycans remain unmodified ("high mannose N-glycans") or are further processed in the Golgi ("complex N-glycans"). Greater diversity is found for O-glycans, which start with a common N-Acetylgalactosamine (GalNAc) residue in animal cells but differ in lower organisms1.
The detailed analysis of the glycosylation of proteins is a field unto itself and requires extensive resources and expertise to execute properly. However a variety of available enzymes that remove sugars (glycosidases) makes possible to have a general idea of the glycosylation status of a protein in a standard laboratory setting. Here we illustrate the use of glycosidases for the analysis of a model glycoprotein: recombinant human chorionic gonadotropin beta (hCG&beta;), which carries two N-glycans and four O-glycans 5. The technique requires only simple instrumentation and typical consumables, and it can be readily adapted to the analysis of multiple glycoprotein samples.
Several enzymes can be used in parallel to study a glycoprotein. PNGase F is able to remove almost all types of N-linked glycans6,7. For O-glycans, there is no available enzyme that can cleave an intact oligosaccharide from the protein backbone. Instead, O-glycans are trimmed by exoglycosidases to a short core, which is then easily removed by O-Glycosidase. The Protein Deglycosylation Mix contains PNGase F, O-Glycosidase, Neuraminidase (sialidase), &beta;1-4 Galactosidase, and &beta;-N-Acetylglucosaminidase. It is used to simultaneously remove N-glycans and some O-glycans8 . Finally, the Deglycosylation Mix was supplemented with a mixture of other exoglycosidases (&alpha;-N-Acetylgalactosaminidase, &alpha;1-2 Fucosidase, &alpha;1-3,6 Galactosidase, and &beta;1-3 Galactosidase ), which help remove otherwise resistant monosaccharides that could be present in certain O-glycans.
SDS-PAGE/Coomasie blue is used to visualize differences in protein migration before and after glycosidase treatment. In addition, a sugar-specific staining method, ProQ Emerald-300, shows diminished signal as glycans are successively removed. This protocol is designed for the analysis of small amounts of glycoprotein (0.5 to 2 &mu;g), although enzymatic deglycosylation can be scaled up to accommodate larger quantities of protein as needed.

Using specific glycosidases to remove sugars from glycoproteins followed by SDS-PAGE is a valuable method to detect glycan modifications on protein samples and is a good choice for initial glycobiology studies. Changes following deglycosylation can be detected as shifts in gel mobility or by staining with glycan sensitive reagents.

Glycosylation, the addition of covalently linked sugars, is a major post-translational modification of proteins that can significantly affect processes ...

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 ...

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in current research and a plethora of methods for their investigation is available1. Protein-protein interactions often are highly dynamic and may depend on subcellular localization, post-translational modifications and the local protein environment2. Therefore, they should be investigated in their natural environment, for which co-immunoprecipitation approaches are the method of choice3. Co-precipitated interaction partners are identified either by immunoblotting in a targeted approach, or by mass spectrometry (LC-MS/MS) in an untargeted way. The latter strategy often is adversely affected by a large number of false positive discoveries, mainly derived from the high sensitivity of modern mass spectrometers that confidently detect traces of unspecifically precipitating proteins. A recent approach to overcome this problem is based on the idea that reduced amounts of specific interaction partners will co-precipitate with a given target protein whose cellular concentration is reduced by RNAi, while the amounts of unspecifically precipitating proteins should be unaffected. This approach, termed QUICK for QUantitative Immunoprecipitation Combined with Knockdown4, employs Stable Isotope Labeling of Amino acids in Cell culture (SILAC)5 and MS to quantify the amounts of proteins immunoprecipitated from wild-type and knock-down strains. Proteins found in a 1:1 ratio can be considered as contaminants, those enriched in precipitates from the wild type as specific interaction partners of the target protein. Although innovative, QUICK bears some limitations: first, SILAC is cost-intensive and limited to organisms that ideally are auxotrophic for arginine and/or lysine. Moreover, when heavy arginine is fed, arginine-to-proline interconversion results in additional mass shifts for each proline in a peptide and slightly dilutes heavy with light arginine, which makes quantification more tedious and less accurate5,6. Second, QUICK requires that antibodies are titrated such that they do not become saturated with target protein in extracts from knock-down mutants.
Here we introduce a modified QUICK protocol which overcomes the abovementioned limitations of QUICK by replacing SILAC for 15N metabolic labeling and by replacing RNAi-mediated knock-down for affinity modulation of protein-protein interactions. We demonstrate the applicability of this protocol using the unicellular green alga Chlamydomonas reinhardtii as model organism and the chloroplast HSP70B chaperone as target protein7 (Figure 1). HSP70s are known to interact with specific co-chaperones and substrates only in the ADP state8. We exploit this property as a means to verify the specific interaction of HSP70B with its nucleotide exchange factor CGE19.

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

We present a variation of the QUICK (QUantitative Immunoprecipitation Combined with Knockdown) approach that was introduced previously to distinguish between true and false protein-protein interactions. Our approach is based on 15N metabolic labeling, the modulation of affinities of protein-protein interactions by the presence/absence of ATP, immunoprecipitation, and quantitative mass spectrometry.

Protein-protein interactions are fundamental for many biological processes in the cell. Therefore, their characterization plays an important role in ...

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 ...

A Protocol for Phage Display and Affinity Selection Using Recombinant Protein Baits

Using recombinant phage as a scaffold to present various protein portions encoded by a directionally cloned cDNA library to immobilized bait molecules is an efficient means to discover interactions. The technique has largely been used to discover protein-protein interactions but the bait molecule to be challenged need not be restricted to proteins. The protocol presented here has been optimized to allow a modest number of baits to be screened in replicates to maximize the identification of independent clones presenting the same protein. This permits greater confidence that interacting proteins identified are legitimate interactors of the bait molecule. Monitoring the phage titer after each affinity selection round provides information on how the affinity selection is progressing as well as on the efficacy of negative controls. One means of titering the phage, and how and what to prepare in advance to allow this process to progress as efficiently as possible, is presented. Attributes of amplicons retrieved following isolation of independent plaque are highlighted that can be used to ascertain how well the affinity selection has progressed. Trouble shooting techniques to minimize false positives or to bypass persistently recovered phage are explained. Means of reducing viral contamination flare up are discussed.

Phage display is a powerful technique to capture proteins or protein moieties that interact with an immobilized molecule of interest. Once a decision of the type of phage cDNA library to create and screen has been made, the protocol described here permits efficient affinity selection leading to identification of interactors.

Using  recombinant phage as a scaffold to present various protein portions encoded by  a directionally cloned cDNA library to immobilized bait molecules ...