The sequencing of complete genomes of several organisms provides an exceptional opportunity to analyze thedifferent functions governed by their genes. Insightsinto these complex biological systems can be gained byanalysis of gene regulatory networks and by determining the identity, modification, and expression levels ofencoded proteins as well as by defining interactionsexisting among proteins (proteomic analyses). Large-scale two-hybrid screening has been used for this latter purpose (1–3). However, false-positive and false-negative results, the lack of information about stoichiometry,and the limited set of conditions testable make it desirable to use additional strategies to easily detect protein interactions.
Biochemical purification of proteins in combinationwith mass spectrometry allows identification of interacting partners. This strategy is becoming an importanttool to define relations existing among gene products(4, 5). Currently, ,100 fmol of a protein can be detectedand identified by mass spectrometry, allowing rapidcharacterization of any protein present in a complexmixture, provided that the target complex is sufficientlypurified in reasonable quantity. Identification of proteins by mass spectrometry is currently facilitated forseveral organisms by the availability of complete genomic sequences. The current limiting step in protein complex characterization appears therefore to be proteinpurification rather than protein identification. Eachprotein has unique properties, which can be exploitedfor its purification (6). This makes it, however, impossi-ble to design a general purification strategy valid forall cases. A generic purification protocol is thereforedesirable to allow routine and possibly automated protein complex purification for proteome analysis. Thefusion of tags, peptides, or protein domains to proteintargets appeared best suited toward this goal. Aftercomparative testing of several tags, we have recentlydeveloped a new tag, the tandem affinity purification(TAP) tag, and we have optimized a procedure, the TAPmethod, for the native purification of protein complexes(7). This strategy allows for fast purification with highyield of protein complexes under standard conditions.Ultimately, the purified complex can be used for protein identification, functional, orstructuralstudies. Furthermore, variations on the original strategy, including the use of C- or N-terminal tags, the use of a split tag, and/or the use of a subtraction step can easily be developed. These various aspects are described below starting with a presentation of the basic TAP method from gene tagging to protein analysis. We also present several applications of the method and discuss different variations from the original protocol and potential problems. General guidelines useful for various organisms are given; however, as the TAP method was developed with yeast, emphasis is given to applications in this organism. Detailed protocols and latest developments can also be found on our web site(http://www.emblheidelberg.de/ExternalInfo/seraphin/TAP.html).
METHODS
1.Over view of the TAP Method and the TAP Tag
The TAP method involves the fusion of the TAP tag (seebelow) to the target protein and the introduction of the construct into the host cell or organism. For optimal results, it is preferable to maintain expression of the fusion protein at, or close to, its natural level. Indeed, over expression of the protein often induces its association with nonnatural partners (heat shock proteins, proteasome; Ref. (8)). Cell extracts are prepared and the fusion protein as well asassociated partners is recovered by two specific affinity purification/elution steps. The material recovered can be analyzed in several ways. For protein complex characterization, proteins are concentrated, and eventually fractionated on a denaturing gel, before identification by mass spectrometry. (Alternatively, Edman degradation or Western blot may be used.) Because the various TAP purification steps are performed in a gentle native manner, purified complexes may also be tested for their activities or used in structural analysis.
The TAP tag consists of two IgG binding domains of Staphylococcus aureus protein A (ProtA) and a calmodulin binding peptide (CBP) separated by a TEV protease cleavage site. Originally, a C-terminal TAP tag was described (7) (Fig. 1A). We have now also generated an N-terminal TAP tag (Fig. 1A, seebelow). Note that the relative order of the modules of the TAP tag are inversed in the two tags because the ProtA module needs to be located at the extreme N or C terminus of the fusion protein. Both affinity tags have been selected for highly efficient recovery of proteins present at low concentration. ProtA binds tightly to an IgG matrix, requiring the use of the TEV protease to elute material under native conditions (Fig. 1B). The eluate of this first affinity purification step is then incubated with calmodulin coated beads in the presence of calcium. After washing, which removes contaminants and the TEV protease remaining after the first affinity selection, the bound material is released under mild conditions with EGTA (Fig. 1B). Optimized conditions have been developed for the generic use of the TAP strategy (see below). The TAP tag is, however, very tolerant to buffer conditions and changes can easily be implemented to optimize recovery of specific complexes.
2.Tagging the Target Protein with the TAP Tag
The choice of the strategy for fusing the TAP tag to the target protein depends on the methods available to introduce recombinant nucleic acids into the corresponding cell or organism. One should also keep in mind that strong over expression of the target protein is not preferable except if one is interested in producing large amounts of this protein by itself. Indeed, protein overexpression may often lead to the formation of nonspecific and/or nonnatural protein interactions with host proteins (8). This should be avoided if one wants to identify the structure, composition, and/or activity of acomplex. The TAP tag has been specifically designed to allow recovery of proteins expressed at their low natural levels. Usually, standard DNA cloning procedures can be used to introduce the N- or C- terminal TAP tag in-frame with the coding region of the protein of interest in an appropriate expression vector. For this purpose, unique
restriction sites present upstream and downstream of the N- and C-terminal TAP cassettes are available (Fig. 2A). The recombinant vector can then be transiently or stably introduced into recipient cells ororganisms. Optimally, the tagged construct should be used to replace the endogenous wild-type gene. However, depending on the organism analyzed, this is not always possible and often time consuming (e.g., construction of transgenic mice).
The high efficiency of homologous recombination in yeast bypasses the need to construct a plasmid to fuse the TAP tag to the protein of interest. Polymerase chain reaction (PCR) fragments can indeed be used to integrate the TAP tag directly in the genome (9,10). We routinely prefer to use the C- terminal TAP tag for this purpose as this maintains expression of the target protein under the control of its natural promoter. However, some proteins undergo loss of function when a peptide is added to its C-terminus. While from our experience this is not very frequent (about 5% of fusions), it is worthwhile to introduce the TAP tag into both haploid and diploid cells in parallel to test this possibility. For cases where problems are encountered with the C-terminal TAP tag, we have designed a strategy that allows genomic fusion of an N-terminal TAP tag to the protein of interest while maintaining its expression under control of the endogenous promoter (see variations of the TAP method below).
The two plasmids constructed in our laboratory to introduce the C-terminal TAP tag into the yeast genome differ by the presence of either a URA3 or a TRP1 marker from Kluyveromyces lactis adjacent to the TAPc assette(Fig. 2A, pBS1479 and pBS1539, respectively). Primers containing a region of similarity to the yeast genome (40-50 nt long) and a constant priming regio (Fig. 2B) are synthesized. Primer A hybridizes at the 5' end of the CBP coding sequence and primer B in the vector downstream of the selection marker. Primer A should be carefully designed such that the last C-terminal residue of the target protein gets fused in-frame to the TAP tag. These primers are used to amplify by PCR the TAP tag from plasmid pBS1479 or pBS1539. The PCR product is extracted with phenol/chloroform/isoamylalcohol, precipitated, and used to transform haploid and diploid yeast cells (11,12). Correct integration of the cassette is verified by PCR and/or Southern blot (13,14). To check for expression of the tagged protein, Western blot is used. Briefly, the cellular pellets corresponding to 1.5 ml of cellculture are vortexed 3×30s with30 μl siliconized glass beads and 100 μl of SDS-PAGE loading buffer. Samples are boiled, vortexed once more, and loaded directly on an SDS-Cpolyacrylamid gel. Western blots are developed with a peroxidase-antiperoxidase complex (PAP, Sigma P-2026) that detects ProtA. However, one should remember that this strategy might not be sufficiently sensitive if the target protein is expressed at a very low level.
3. Extract preparation
Various extraction procedures can be used to prepare extracts from cells or organisms expressing the target protein fused to the TAP tag. The choice of the appropriate extract preparation procedure will depend on the target protein and on prior experience in the field that can be found in the literature. Cell fractionation and/or tissue dissection can facilitate purification by providing a preenrichment step or can be used to assay specifically protein complex composition in various tissues or cell compartments. In general, however, it is advisable to check, by detecting the ProtA moiety of the TAP tag by Western blot, whether extraction is efficient and if the TAP tag is not degraded under these specific conditions.
For yeast, we recommend the following standard procedure that has been extensively used in our laboratory. However, this method is unlikely to be optimal for all proteins and alternative protein extraction methods may be used (see variations in the purification protocol below). Extracts are routinely prepared from 2 liters of yeast cells grown to late log phase (OD600～2–3). Cell pellets are washed once with water and pelleted againin a 50-ml polypropylene tube (Falcon). The packed cell volume (PCV) is measured and the tube is frozen with liquid nitrogen. Frozen cell pellets may be stored at
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