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Mapping protein–protein interactions by mass spectrometry
Mapping protein–protein interactions by mass spectrometry Julian Vasilescu and Daniel Figeys Mass spectrometry is currently at the forefront of technologies for mapping protein–protein interactions, as it is a highly sensitive technique that enables the rapid identification of proteins from a variety of biological samples. When used in combination with affinity purification and/or chemical crosslinking, whole or targeted protein interaction networks can be elucidated. Several methods have recently been introduced that display increased specificity and a reduced occurrence of false-positives. In the future, information gained from human protein interaction studies could lead to the discovery of novel pathway associations and therapeutic targets. Addresses The Ottawa Institute of Systems Biology, 451 Smyth Road, Ottawa, ON K1H 8M5, Canada Corresponding author: Figeys, Daniel ([email protected])
Current Opinion in Biotechnology 2006, 17:394–399 This review comes from a themed issue on Protein technologies Edited by Deb K Chatterjee and Joshua LaBaer Available online 5th July 2006 0958-1669/$ – see front matter # 2006 Elsevier Ltd. All rights reserved. DOI 10.1016/j.copbio.2006.06.008
Introduction Mass spectrometry (MS) is a powerful technique for identifying protein–protein interactions and for elucidating components of multiprotein complexes. Although MS is highly sensitive and relatively tolerant to heterogeneities and contaminants, the identification of protein complexes remains a significant challenge. This is because many interactions are transient in nature and components that form these unstable complexes can be lost during sample preparation. The purification of native protein complexes is also difficult, because many are expressed at low levels and might not possess a unique physical characteristic that permits their isolation or enrichment from a cell lysate. Affinity purification strategies employing epitope tags and mild buffer conditions have proven to be effective for both large-scale and targeted protein interaction mapping efforts. Another effective tool that has been used to aid the purification of protein complexes employs chemical cross-linkers that effectively ‘freeze’ protein– protein interactions in their in vivo state and stabilizes complexes for subsequent purification. Quantitative Current Opinion in Biotechnology 2006, 17:394–399
approaches using chemical cross-linking, isotope labeling and affinity purification, also now offer the possibility to distinguish bona fide interaction partners from background contaminants. The aim of this short review is to provide an overview of developments in protein interaction mapping by MS that have occurred in the past two years. In the following sections we provide a summary of results from recent large-scale interaction mapping efforts in yeast and human cells. We also describe MS-based methods that employ novel affinity purification strategies and affinity tags as well as in vivo and in vitro chemical cross-linking approaches.
Large-scale protein interaction mapping studies Since the landmark studies of yeast protein complexes by Gavin et al. [1] and Ho et al. [2] were published in 2002, three large-scale protein interaction mapping efforts have been reported [3–5]. All three employed tandem affinity purification coupled to MS (TAP-MS) — a strategy used for identifying a tagged protein and its interaction partners that has gained in popularity in recent years because of its high reproducibility and selectivity [6,7]. Krogan et al. [3] utilized the TAP-MS strategy to process 4562 tagged proteins in Saccharomyces cerevisiae. Matrixassisted laser desorption ionization time-of-flight (MALDI-TOF) MS and liquid chromatography tandem MS (LC-MS/MS) analysis of affinity-purified samples resulted in the identification of 4087 proteins (corresponding to 72% of the predicted yeast proteome) and enabled the generation of a core dataset comprising 7123 protein–protein interactions. A clustering algorithm was then used to organize interacting proteins into 547 protein complexes. Taken together, these results represent the most comprehensive analysis of yeast protein–protein interactions to date. Using a similar TAP-MS strategy, Gavin et al. [4] performed a total of 1993 successful purifications, resulting in 2760 confident protein identifications. The reproducibility of this approach was measured by performing 139 purifications in duplicate and, on average, 69% of proteins were found to be common to both, thus giving an approximation of false-positive rates. Iterative clustering and selection was used to organize the interaction dataset into 491 complexes that differentially combine with additional attachment proteins or protein modules. Of the 491 protein complexes reported, 257 were deemed to be novel. www.sciencedirect.com
Mapping protein–protein interactions Vasilescu and Figeys 395
To map the protein interaction network around the human tumor necrosis factor-a/nuclear factor kB (TNF-a/NF-kB) signal transduction pathway, Bouwmeester et al. [5] TAP-tagged 32 known or postulated components of this pathway and expressed them in embryonic kidney cells by retrovirus-mediated gene transfer. A total of 237 successful TAP purifications were performed, with at least four purifications per tagged component. LC-MS/MS analysis enabled the identification of 680 unique proteins and 171 protein–protein interactions, representing 70% of known interactions in the TNF-a/NF-kB signaling pathway. Functional validation was performed on 28 proteins using RNA interference together with an NF-kB luciferase reporter assay to monitor the phenotypic effect. On the basis of loss-offunction data and results obtained from co-immunoprecipitation experiments, 10 new functional modulators of the TNF-a/NF-kB signaling pathway were identified.
This novel approach, consisting of targeted interaction mapping combined with loss-of-function analysis, is applicable to other disease-related signaling pathways.
Novel affinity purification strategies for mapping protein interactions Although the TAP-MS strategy has been widely used for mapping yeast protein–protein interactions, its applicability to higher eukaryotes has been limited. Unlike yeast, which can be grown in large quantities in flasks, mammalian cells often grow in monolayer cultures and cannot be harvested as easily. To overcome this problem, Drakas et al. [8] modified the original TAP tag (Figure 1a) by inserting a biotinylation tag at the N terminus of the target protein (Figure 1b). Use of this modified tag in combination with avidin beads takes advantage of the high-affinity streptavidin–biotin interaction and increases the overall yield of purified protein complexes. Drakas
Figure 1
Novel affinity tags for mapping protein interactions by MS. (a) Original C-terminal TAP tag [6]. CBP represents the calmodulin-binding peptide, TEV represents a cleavage site for the tobacco etch virus and ProtA represents two immunoglobulin G-binding domains of Staphylococcus aureus Protein A. (b) Modified mammalian TAP tag [8]. BT represents a 17-amino acid sequence that is biotinylated by Escherichia coli biotin holoenzyme synthase and binds to streptavidin. (c) The Strep-II tag is an 8 amino acid peptide that binds a derivative of streptavidin, whereas the Strep-III tag is a sequential arrangement of the Strep-II tag separated by a linker [9]. (d) C-terminal histine-biotin (HB) tag [10]. RGSH6 represents a hexahistidinecontaining sequence that binds to Ni2+ chelate resins. BIO represents a 75-amino acid sequence derived from Propionibacterium shermanii transcarboxylase that is biotinylated and binds to streptavidin. (e) Fusion protein [11]. THIO represents a modified thioredoxin gene, V5 is a 14-amino acid sequence, and 6 His is a hexahistidine sequence. www.sciencedirect.com
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and colleagues demonstrated the usefulness of this tag by identifying numerous insulin receptor substrate-1 interaction partners in mouse embryo fibroblasts. As an alternative to the TAP-MS strategy, Junttila et al. [9] developed a pair of epitope tags that also take advantage of the high binding affinity between streptavidin and biotin. The Strep-tag II is an eight amino acid peptide that strongly binds a derivative of streptavidin called Strep-Tactin, while the Strep-tag III is a sequential arrangement of the Strep-tag II sequence separated by a linker region (Figure 1c). The usefulness of these tags was demonstrated by tagging a component of the protein phosphatase 2A (PP2A) holoenzyme complex and expressing it in a human fibrosarcoma cell line. Lysates expressing the tagged protein were then passed through a Strep-Tactin column and bound proteins were specifically eluted using a derivative of biotin called desthiobiotin. Both known and novel interacting proteins of the PP2A complex were subsequently identified by LC-MS/MS. In another development related to epitope tags, Tagwerker et al. [10] developed a histine–biotin (HB) tag for a two-step purification protocol that can be performed under fully denaturing conditions, such as 8 M urea or 6 M guanidinium chloride. The HB tag consists of a hexahistidine sequence which binds to Ni2+ chelate resins and a biotinylation tag that binds to streptavidin resins (Figure 1d). Tagwerker et al. used the HB tag to perform a targeted ubiquitin profiling experiment and identified several known and candidate ubiquitinated proteins in yeast. The HB tag was also used to enable the identification of interaction partners of a HB-tagged ubiquitin ligase, known as Skp1, using in vivo crosslinking and LC-MS/MS analysis. The stringent purification conditions employed in this method are particularly useful for the analysis of post-translational modifications, because it permits their maximal preservation. A method based on single-step affinity purification has been developed by Markillie et al. [11] that circumvents the need to perform sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) and in-gel digestion before MS analysis. This gel-free method consists of expressing a fusion protein in bacteria that contains the V5 and hexahistidine epitope tags and thioredoxin at the C and N termini, respectively (Figure 1e). Cell extracts containing the fusion protein are incubated with Ni2+ MagneHis beads, which bind to the hexahistidine sequence, or an affi-gel matrix that covalently binds to amino groups. Bound proteins are then eluted from the beads using a denaturing solution (e.g. 8 M urea or 40% acetonitrile) and are subjected to in-solution tryptic digestion and LC-MS/MS analysis. The applicability of this approach was demonstrated by tagging a component of the degradosome complex, known as polynucleotide Current Opinion in Biotechnology 2006, 17:394–399
phosphorylase, and identifying specific interaction partners that included RNase E and RNA helicase. McCraken et al. [12] described an affinity purification method for protein complexes that also avoids SDS– PAGE separation. Cell extracts are incubated with antibody-coupled sepharose beads that bind the target protein. The target protein and its interaction partners are eluted with urea, digested with trypsin in solution and analyzed by LC-MS/MS. This method was used for the characterization of complexes containing the splicing co-activator protein SRm160 from HeLa nuclear cell lysates. Proteins involved in pre-mRNA processing and chromatin regulation were found to associate with SRm160. Affinity purification strategies combined with isotope labeling strategies such as SILAC (stable isotope labeling of amino acids in cell culture) have been described for the analysis of protein interactions. In a recent study, Foster et al. [13] utilized such an approach to identify proteins that interact with the insulin-regulated glucose transporter, GLUT4, in an insulin-dependent manner. Myctagged GLUT4 was stably expressed in basal or insulin-stimulated myoblast cells and grown in medium containing normal leucine or deuterated leucine. Proteins that co-purified with GLUT4 were analyzed by LC-MS/ MS and among the group of proteins quantified, 36 displayed significant insulin-dependent changes in their interaction with GLUT4. Co-immunoprecipitation and fluorescence microscopy experiments were used to validate a number of these interactions.
Chemical cross-linking approaches for studying protein interactions Affinity purification strategies coupled directly to MS analysis are unable to characterize all protein complexes because many protein–protein interactions are transient, or low affinity, and are dependent on the specific cellular environment in which they occur. Therefore, in vivo chemical cross-linking can be used to stabilize these interactions through covalent-bond formation before affinity purification (Figure 2). The most commonly used cross-linker for this purpose is formaldehyde, which is a small, water-soluble, cell-membrane-permeable molecule that efficiently produces reversible protein–protein cross-links [14,15]. Guerrero et al. [16] recently described a method for the study of protein complexes in yeast using formaldehyde cross-linking in combination with SILAC, affinity purification and LC-MS/MS analysis. Using this method, Guerrero et al. identified the full composition of the 26S proteasome complex, including two known ubiquitin receptors, Rad23 and Dsk2. Quantitative MS analysis was also used to identify 64 potential proteasome-interacting proteins, 42 of which were novel interactions, and enabled www.sciencedirect.com
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Figure 2
Formaldehyde cross-linking coupled to affinity purification and MS analysis has also been adapted to mammalian cells. Vasilescu and colleagues [17] studied the interaction partners of a Myc-tagged Ras GTPase, known as MRas, in a murine cell line that stably expressed this protein. Cells were treated with formaldehyde and tagged complexes were purified by immunoaffinity chromatography and the cross-linked complexes dissociated. After SDS–PAGE separation, interacting proteins were identified by LC-MS/MS. Co-immunoprecipitation experiments were also used to confirm a number of these interactions, including one involving the RasGAP-related protein IQGAP-1. A method that combines transcardiac perfusion and formaldehyde cross-linking for the study of protein complexes in living tissues has also been reported [18]. Using mouse brains as the target tissue, this method enabled the purification and identification of more than 20 membrane proteins associated with the cellular prion protein PrPC. To validate their protocol, the authors used high salt and detergent buffers during the purification of a complex known as g-secretase. Despite the stringent washing conditions, LC-MS/MS analysis was able to confirm the presence of a number of its known components, including aph-1, presenilin-1, and nicastrin. In addition to aiding the purification of native protein complexes, chemical cross-linking can be used to detect specific interaction sites and to obtain low-resolution structural information by MS. A study by Kang et al. [19] used chemical cross-linking and the introduction of site-specific mutations to identify inter- and intrasubunit interactions within the bacteriophage p22 procapsid. Three lysine-reactive cross-linkers were used to determine cross-linking between two coat protein subunits at the same residue (Lys183) and between subunits at different residues (Lys175–Lys183). A mutation was introduced at Lys183 and the integrity of the capsid monitored; predicted secondary structure elements were then generated from the data obtained.
In vivo chemical cross-linking. (a) Affinity purification coupled to MS analysis. (b) In vivo cross-linking of a ‘bait’ protein to interacting ‘prey’ proteins before affinity purification prevents the loss of transient and/or weakly binding interaction partners for MS analysis. More stringent washing conditions can be employed to remove contaminants.
co-purifying background proteins to be distinguished. This method, termed QTAX (quantitative analysis of tandem affinity-purified in vivo cross-linked protein complexes), is a powerful integrated approach that could be used for the study of other yeast protein complexes. www.sciencedirect.com
Chemical cross-linking and MS analysis have also been used to study interactions between the protein component of the signal recognition particle (FfH) and its receptor (FtsY) [20]. FfH–FtsY complexes were purified from bacteria and treated with two homo-bifunctional cross-linkers. LC-MS/MS analysis enabled the identification of nine intermolecular cross-linked peptides as well as specific residues involved in cross-linking. This information was used in conjunction with a computational modeling approach that combines geometric restraints optimization with macromolecular docking to determine a structure of the FfH–FtsY complex. The resulting model was found to be in agreement with a later identified crystal structure. Current Opinion in Biotechnology 2006, 17:394–399
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It should be noted that the analysis of cross-linked peptides by MS is not always feasible when dealing with very large protein complexes. Dynamic range issues become a factor when a small number of cross-linked peptides are present among hundreds to thousands of unmodified peptides. Several strategies have been employed to overcome this challenge, including the use of isotope-labeled cross-linkers, cross-linkers that create characteristic ions upon fragmentation, and exact mass measurement of cross-linked peptides using high mass accuracy instruments, such as FTICR (fourier transform ion cyclotron resonance) mass spectrometers. Several recent examples of these strategies have been reported [21–23].
Conclusions Mapping protein–protein interactions by MS remains a significant challenge that requires the use of affinity purification strategies or other complementary techniques such as chemical cross-linking. The development of highly specific epitope tags have made it feasible to pursue large-scale interaction mapping efforts in human cells that parallel those already performed in yeast. However, such studies are likely to target proteins with known disease associations and of known functional classes, rather than systematically tagging and expressing all known and predicted gene products. The information gained from large-scale human protein interaction studies will ultimately lead to the discovery of novel pathway associations and therapeutic targets.
References and recommended reading Papers of particular interest, published within the annual period of review, have been highlighted as: of special interest of outstanding interest 1.
Gavin AC, Bosche M, Krause R, Grandi P, Marzioch M, Bauer A, Schultz J, Rick JM, Michon AM, Cruciat CM et al.: Functional organization of the yeast proteome by systematic analysis of protein complexes. Nature 2002, 415:141-147.
2.
Ho Y, Gruhler A, Heilbut A, Bader GD, Moore L, Adams SL, Millar A, Taylor P, Bennett K, Boutilier K et al.: Systematic identification of protein complexes in Saccharomyces cerevisiae by mass spectrometry. Nature 2002, 415:180-183.
3.
Krogan NJ, Cagney G, Yu H, Zhong G, Guo X, Ignatchenko A, Li J, Pu S, Datta N, Tikuisis AP et al.: Global landscape of protein complexes in the yeast Saccharomyces cerevisiae. Nature 2006, in press, doi:10.1038/nature04670. TAP-MS was used to process 4562 yeast proteins. A core dataset comprising 7123 protein–protein interactions was produced involving 2708 proteins. A clustering algorithm was then used to organize the interactions into 547 protein complexes.
4.
Gavin AC, Aloy P, Grandi P, Krause R, Boesche et al.: Proteome survey reveals modularity of the yeast cell machinery. Nature 2006, in press, doi:10.1038/nature04532. TAP-MS was used to perform a genome wide screen for complexes in yeast. Cellular proteins were found to be partitioned into 491 complexes, of which 257 are novel.
5.
Bouwmeester T, Bauch A, Ruffner H, Angrand PO, Bergamini G, Croughton K, Cruciat C, Eberhard D, Gagneur J, Ghidelli S et al.: A physical and functional map of the human TNF-a/NF-kB signal transduction pathway. Nat Cell Biol 2004, 6:97-105. TAP-MS was used to map the protein interaction network around the human TNF-a/NF-kB pathway. Some 221 protein–protein interactions
Current Opinion in Biotechnology 2006, 17:394–399
were identified, including 80 unknown interactors and 10 new functional modulators. 6.
Puig O, Caspary F, Rigaut G, Rutz B, Bouveret E: The tandem affinity purification (TAP) method: a general procedure of protein complex purification. Methods 2001, 24:218-229.
7.
Rigaut G, Shevchenko A, Rutz B, Wilm M, Mann M, Seraphin B: A generic protein purification method for protein complex characterization and proteome exploration. Nat Biotechnol 1999, 17:1030-1032.
8.
Drakas R, Prisco M, Baserga R: A modified tandem affinity purification tag technique for the purification of protein complexes in mammalian cells. Proteomics 2005, 5:132-137.
9.
Junttila MR, Saarinen S, Schmidt T, Kast J, Westermarck J: Single-step strep-tag purification for the isolation and identification of protein complexes from mammalian cells. Proteomics 2005, 5:1199-1203.
10. Tagwerker C, Flick K, Cui M, Guerrero C, Dou Y, Auer B, Baldi P, Huang L, Kaiser P: A tandem affinity tag for two-step purification under fully denaturing conditions: application in ubiquitin profiling and protein complex identification combined with in vivo cross-linking. Mol Cell Proteomics 2006, 5:737-748. A new histidine-biotin tag is described for two-step purification under denaturing conditions. This method was used to study ubiquitination in yeast and for the analysis of an in vivo cross-linked protein complex containing Skp1. 11. Markillie LM, Lin CT, Adkins JN, Auberry DL, Hill EA, Hooker BS, Moore PA, Moore RJ, Shi L, Wiley HS, Kery V: Simple protein complex purification and identification method for highthroughput mapping of protein interaction networks. J Proteome Res 2005, 4:268-274. 12. McCracken S, Longman D, Marcon E, Moens P, Downey M, Nickerson JA, Jessberger R, Wilde A, Caceres JF, Emili A, Blencowe BJ: Proteomic analysis of SRm160-containing complexes reveals a conserved association with Cohesin. J Biol Chem 2005, 280:42227-422236. 13. Foster LJ, Rudich A, Talior I, Patel N, Huang X, Furtado LM, Bilan PJ, Mann M, Klip A: Insulin-dependent interactions of proteins with GLUT4 revealed through stable isotope labeling by amino acids in cell culture. J Proteome Res 2006, 5:64-75. 14. Orlando V, Strutt H, Paro R: Analysis of chromatin structure by in vivo formaldehyde cross-linking. Methods 1997, 11:1187-1198. 15. Either M, Lambert JP, Vasilescu J, Figeys D: Analysis of protein interactions networks using mass spectrometry compatible techniques. Anal Chim Acta 2006, 564:10-18. 16. Guerrero C, Tagwerker C, Kaiser P, Huang L: An integrated mass spectrometry-based proteomic approach: quantitative analysis of tandem affinity-purified in vivo cross-linked protein complexes (QTAX) to decipher the 26S proteasomeinteracting network. Mol Cell Proteomics 2006, 5:366-378. Affinity purification after in vivo formaldehyde cross-linking was combined with SILAC and MS analysis to identify and quantify specific interactions. The full composition of the yeast 26S proteasome complex was identified using this method. 17. Vasilescu J, Guo X, Kast J: Identification of protein–protein interactions using in vivo cross-linking and mass spectrometry. Proteomics 2004, 4:3845-3854. 18. Schmitt-Ulms G, Hansen K, Liu J, Cowdrey C, Yang J, DeArmond SJ, Cohen FE, Prusiner SB, Baldwin MA: Timecontrolled transcardiac perfusion cross-linking for the study of protein interactions in complex tissues. Nat Biotechnol 2004, 22:724-731. 19. Kang S, Hawkridge AM, Johnson KL, Muddiman DC, Prevelige PE: Identification of subunit–subunit interactions in bacteriophage p22 procapsids by chemical cross-linking and mass spectrometry. J Proteome Res 2005, 5:370-377. 20. Chu F, Shan SO, Moustakas DT, Alber F, Egea PF, Stroud RM, Walter P, Burlingame AL: Unraveling the interface of signal recognition particle and its receptor by using chemical www.sciencedirect.com
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cross-linking and tandem mass spectrometry. Proc Natl Acad Sci USA 2004, 101:16454-16459. 21. Kalkhof S, Ihling C, Mechtler K, Sinz A: Chemical cross-linking and high-performance fourier transform ion cyclotron resonance mass spectrometry for protein interaction analysis: application to a calmodulin/target peptide complex. Anal Chem 2005, 77:495-503.
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22. Petrotchenko EV, Olkhovik VK, Borchers CH: Isotopically coded cleavable cross-linker for studying protein-protein interactions and protein complexes. Mol Cell Proteomics 2005, 4:1167-1179. 23. Tang X, Munske GR, Siems WF, Bruce JE: Mass spectrometry identifiable cross-linking strategy for studying protein–protein interactions. Anal Chem 2005, 77:311-318.
Current Opinion in Biotechnology 2006, 17:394–399
Current Opinion in Biotechnology 2006, 17:394–399 This review comes from a themed issue on Protein technologies Edited by Deb K Chatterjee and Joshua LaBaer Available online 5th July 2006 0958-1669/$ – see front matter # 2006 Elsevier Ltd. All rights reserved. DOI 10.1016/j.copbio.2006.06.008
Introduction Mass spectrometry (MS) is a powerful technique for identifying protein–protein interactions and for elucidating components of multiprotein complexes. Although MS is highly sensitive and relatively tolerant to heterogeneities and contaminants, the identification of protein complexes remains a significant challenge. This is because many interactions are transient in nature and components that form these unstable complexes can be lost during sample preparation. The purification of native protein complexes is also difficult, because many are expressed at low levels and might not possess a unique physical characteristic that permits their isolation or enrichment from a cell lysate. Affinity purification strategies employing epitope tags and mild buffer conditions have proven to be effective for both large-scale and targeted protein interaction mapping efforts. Another effective tool that has been used to aid the purification of protein complexes employs chemical cross-linkers that effectively ‘freeze’ protein– protein interactions in their in vivo state and stabilizes complexes for subsequent purification. Quantitative Current Opinion in Biotechnology 2006, 17:394–399
approaches using chemical cross-linking, isotope labeling and affinity purification, also now offer the possibility to distinguish bona fide interaction partners from background contaminants. The aim of this short review is to provide an overview of developments in protein interaction mapping by MS that have occurred in the past two years. In the following sections we provide a summary of results from recent large-scale interaction mapping efforts in yeast and human cells. We also describe MS-based methods that employ novel affinity purification strategies and affinity tags as well as in vivo and in vitro chemical cross-linking approaches.
Large-scale protein interaction mapping studies Since the landmark studies of yeast protein complexes by Gavin et al. [1] and Ho et al. [2] were published in 2002, three large-scale protein interaction mapping efforts have been reported [3–5]. All three employed tandem affinity purification coupled to MS (TAP-MS) — a strategy used for identifying a tagged protein and its interaction partners that has gained in popularity in recent years because of its high reproducibility and selectivity [6,7]. Krogan et al. [3] utilized the TAP-MS strategy to process 4562 tagged proteins in Saccharomyces cerevisiae. Matrixassisted laser desorption ionization time-of-flight (MALDI-TOF) MS and liquid chromatography tandem MS (LC-MS/MS) analysis of affinity-purified samples resulted in the identification of 4087 proteins (corresponding to 72% of the predicted yeast proteome) and enabled the generation of a core dataset comprising 7123 protein–protein interactions. A clustering algorithm was then used to organize interacting proteins into 547 protein complexes. Taken together, these results represent the most comprehensive analysis of yeast protein–protein interactions to date. Using a similar TAP-MS strategy, Gavin et al. [4] performed a total of 1993 successful purifications, resulting in 2760 confident protein identifications. The reproducibility of this approach was measured by performing 139 purifications in duplicate and, on average, 69% of proteins were found to be common to both, thus giving an approximation of false-positive rates. Iterative clustering and selection was used to organize the interaction dataset into 491 complexes that differentially combine with additional attachment proteins or protein modules. Of the 491 protein complexes reported, 257 were deemed to be novel. www.sciencedirect.com
Mapping protein–protein interactions Vasilescu and Figeys 395
To map the protein interaction network around the human tumor necrosis factor-a/nuclear factor kB (TNF-a/NF-kB) signal transduction pathway, Bouwmeester et al. [5] TAP-tagged 32 known or postulated components of this pathway and expressed them in embryonic kidney cells by retrovirus-mediated gene transfer. A total of 237 successful TAP purifications were performed, with at least four purifications per tagged component. LC-MS/MS analysis enabled the identification of 680 unique proteins and 171 protein–protein interactions, representing 70% of known interactions in the TNF-a/NF-kB signaling pathway. Functional validation was performed on 28 proteins using RNA interference together with an NF-kB luciferase reporter assay to monitor the phenotypic effect. On the basis of loss-offunction data and results obtained from co-immunoprecipitation experiments, 10 new functional modulators of the TNF-a/NF-kB signaling pathway were identified.
This novel approach, consisting of targeted interaction mapping combined with loss-of-function analysis, is applicable to other disease-related signaling pathways.
Novel affinity purification strategies for mapping protein interactions Although the TAP-MS strategy has been widely used for mapping yeast protein–protein interactions, its applicability to higher eukaryotes has been limited. Unlike yeast, which can be grown in large quantities in flasks, mammalian cells often grow in monolayer cultures and cannot be harvested as easily. To overcome this problem, Drakas et al. [8] modified the original TAP tag (Figure 1a) by inserting a biotinylation tag at the N terminus of the target protein (Figure 1b). Use of this modified tag in combination with avidin beads takes advantage of the high-affinity streptavidin–biotin interaction and increases the overall yield of purified protein complexes. Drakas
Figure 1
Novel affinity tags for mapping protein interactions by MS. (a) Original C-terminal TAP tag [6]. CBP represents the calmodulin-binding peptide, TEV represents a cleavage site for the tobacco etch virus and ProtA represents two immunoglobulin G-binding domains of Staphylococcus aureus Protein A. (b) Modified mammalian TAP tag [8]. BT represents a 17-amino acid sequence that is biotinylated by Escherichia coli biotin holoenzyme synthase and binds to streptavidin. (c) The Strep-II tag is an 8 amino acid peptide that binds a derivative of streptavidin, whereas the Strep-III tag is a sequential arrangement of the Strep-II tag separated by a linker [9]. (d) C-terminal histine-biotin (HB) tag [10]. RGSH6 represents a hexahistidinecontaining sequence that binds to Ni2+ chelate resins. BIO represents a 75-amino acid sequence derived from Propionibacterium shermanii transcarboxylase that is biotinylated and binds to streptavidin. (e) Fusion protein [11]. THIO represents a modified thioredoxin gene, V5 is a 14-amino acid sequence, and 6 His is a hexahistidine sequence. www.sciencedirect.com
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and colleagues demonstrated the usefulness of this tag by identifying numerous insulin receptor substrate-1 interaction partners in mouse embryo fibroblasts. As an alternative to the TAP-MS strategy, Junttila et al. [9] developed a pair of epitope tags that also take advantage of the high binding affinity between streptavidin and biotin. The Strep-tag II is an eight amino acid peptide that strongly binds a derivative of streptavidin called Strep-Tactin, while the Strep-tag III is a sequential arrangement of the Strep-tag II sequence separated by a linker region (Figure 1c). The usefulness of these tags was demonstrated by tagging a component of the protein phosphatase 2A (PP2A) holoenzyme complex and expressing it in a human fibrosarcoma cell line. Lysates expressing the tagged protein were then passed through a Strep-Tactin column and bound proteins were specifically eluted using a derivative of biotin called desthiobiotin. Both known and novel interacting proteins of the PP2A complex were subsequently identified by LC-MS/MS. In another development related to epitope tags, Tagwerker et al. [10] developed a histine–biotin (HB) tag for a two-step purification protocol that can be performed under fully denaturing conditions, such as 8 M urea or 6 M guanidinium chloride. The HB tag consists of a hexahistidine sequence which binds to Ni2+ chelate resins and a biotinylation tag that binds to streptavidin resins (Figure 1d). Tagwerker et al. used the HB tag to perform a targeted ubiquitin profiling experiment and identified several known and candidate ubiquitinated proteins in yeast. The HB tag was also used to enable the identification of interaction partners of a HB-tagged ubiquitin ligase, known as Skp1, using in vivo crosslinking and LC-MS/MS analysis. The stringent purification conditions employed in this method are particularly useful for the analysis of post-translational modifications, because it permits their maximal preservation. A method based on single-step affinity purification has been developed by Markillie et al. [11] that circumvents the need to perform sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) and in-gel digestion before MS analysis. This gel-free method consists of expressing a fusion protein in bacteria that contains the V5 and hexahistidine epitope tags and thioredoxin at the C and N termini, respectively (Figure 1e). Cell extracts containing the fusion protein are incubated with Ni2+ MagneHis beads, which bind to the hexahistidine sequence, or an affi-gel matrix that covalently binds to amino groups. Bound proteins are then eluted from the beads using a denaturing solution (e.g. 8 M urea or 40% acetonitrile) and are subjected to in-solution tryptic digestion and LC-MS/MS analysis. The applicability of this approach was demonstrated by tagging a component of the degradosome complex, known as polynucleotide Current Opinion in Biotechnology 2006, 17:394–399
phosphorylase, and identifying specific interaction partners that included RNase E and RNA helicase. McCraken et al. [12] described an affinity purification method for protein complexes that also avoids SDS– PAGE separation. Cell extracts are incubated with antibody-coupled sepharose beads that bind the target protein. The target protein and its interaction partners are eluted with urea, digested with trypsin in solution and analyzed by LC-MS/MS. This method was used for the characterization of complexes containing the splicing co-activator protein SRm160 from HeLa nuclear cell lysates. Proteins involved in pre-mRNA processing and chromatin regulation were found to associate with SRm160. Affinity purification strategies combined with isotope labeling strategies such as SILAC (stable isotope labeling of amino acids in cell culture) have been described for the analysis of protein interactions. In a recent study, Foster et al. [13] utilized such an approach to identify proteins that interact with the insulin-regulated glucose transporter, GLUT4, in an insulin-dependent manner. Myctagged GLUT4 was stably expressed in basal or insulin-stimulated myoblast cells and grown in medium containing normal leucine or deuterated leucine. Proteins that co-purified with GLUT4 were analyzed by LC-MS/ MS and among the group of proteins quantified, 36 displayed significant insulin-dependent changes in their interaction with GLUT4. Co-immunoprecipitation and fluorescence microscopy experiments were used to validate a number of these interactions.
Chemical cross-linking approaches for studying protein interactions Affinity purification strategies coupled directly to MS analysis are unable to characterize all protein complexes because many protein–protein interactions are transient, or low affinity, and are dependent on the specific cellular environment in which they occur. Therefore, in vivo chemical cross-linking can be used to stabilize these interactions through covalent-bond formation before affinity purification (Figure 2). The most commonly used cross-linker for this purpose is formaldehyde, which is a small, water-soluble, cell-membrane-permeable molecule that efficiently produces reversible protein–protein cross-links [14,15]. Guerrero et al. [16] recently described a method for the study of protein complexes in yeast using formaldehyde cross-linking in combination with SILAC, affinity purification and LC-MS/MS analysis. Using this method, Guerrero et al. identified the full composition of the 26S proteasome complex, including two known ubiquitin receptors, Rad23 and Dsk2. Quantitative MS analysis was also used to identify 64 potential proteasome-interacting proteins, 42 of which were novel interactions, and enabled www.sciencedirect.com
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Figure 2
Formaldehyde cross-linking coupled to affinity purification and MS analysis has also been adapted to mammalian cells. Vasilescu and colleagues [17] studied the interaction partners of a Myc-tagged Ras GTPase, known as MRas, in a murine cell line that stably expressed this protein. Cells were treated with formaldehyde and tagged complexes were purified by immunoaffinity chromatography and the cross-linked complexes dissociated. After SDS–PAGE separation, interacting proteins were identified by LC-MS/MS. Co-immunoprecipitation experiments were also used to confirm a number of these interactions, including one involving the RasGAP-related protein IQGAP-1. A method that combines transcardiac perfusion and formaldehyde cross-linking for the study of protein complexes in living tissues has also been reported [18]. Using mouse brains as the target tissue, this method enabled the purification and identification of more than 20 membrane proteins associated with the cellular prion protein PrPC. To validate their protocol, the authors used high salt and detergent buffers during the purification of a complex known as g-secretase. Despite the stringent washing conditions, LC-MS/MS analysis was able to confirm the presence of a number of its known components, including aph-1, presenilin-1, and nicastrin. In addition to aiding the purification of native protein complexes, chemical cross-linking can be used to detect specific interaction sites and to obtain low-resolution structural information by MS. A study by Kang et al. [19] used chemical cross-linking and the introduction of site-specific mutations to identify inter- and intrasubunit interactions within the bacteriophage p22 procapsid. Three lysine-reactive cross-linkers were used to determine cross-linking between two coat protein subunits at the same residue (Lys183) and between subunits at different residues (Lys175–Lys183). A mutation was introduced at Lys183 and the integrity of the capsid monitored; predicted secondary structure elements were then generated from the data obtained.
In vivo chemical cross-linking. (a) Affinity purification coupled to MS analysis. (b) In vivo cross-linking of a ‘bait’ protein to interacting ‘prey’ proteins before affinity purification prevents the loss of transient and/or weakly binding interaction partners for MS analysis. More stringent washing conditions can be employed to remove contaminants.
co-purifying background proteins to be distinguished. This method, termed QTAX (quantitative analysis of tandem affinity-purified in vivo cross-linked protein complexes), is a powerful integrated approach that could be used for the study of other yeast protein complexes. www.sciencedirect.com
Chemical cross-linking and MS analysis have also been used to study interactions between the protein component of the signal recognition particle (FfH) and its receptor (FtsY) [20]. FfH–FtsY complexes were purified from bacteria and treated with two homo-bifunctional cross-linkers. LC-MS/MS analysis enabled the identification of nine intermolecular cross-linked peptides as well as specific residues involved in cross-linking. This information was used in conjunction with a computational modeling approach that combines geometric restraints optimization with macromolecular docking to determine a structure of the FfH–FtsY complex. The resulting model was found to be in agreement with a later identified crystal structure. Current Opinion in Biotechnology 2006, 17:394–399
398 Protein technologies
It should be noted that the analysis of cross-linked peptides by MS is not always feasible when dealing with very large protein complexes. Dynamic range issues become a factor when a small number of cross-linked peptides are present among hundreds to thousands of unmodified peptides. Several strategies have been employed to overcome this challenge, including the use of isotope-labeled cross-linkers, cross-linkers that create characteristic ions upon fragmentation, and exact mass measurement of cross-linked peptides using high mass accuracy instruments, such as FTICR (fourier transform ion cyclotron resonance) mass spectrometers. Several recent examples of these strategies have been reported [21–23].
Conclusions Mapping protein–protein interactions by MS remains a significant challenge that requires the use of affinity purification strategies or other complementary techniques such as chemical cross-linking. The development of highly specific epitope tags have made it feasible to pursue large-scale interaction mapping efforts in human cells that parallel those already performed in yeast. However, such studies are likely to target proteins with known disease associations and of known functional classes, rather than systematically tagging and expressing all known and predicted gene products. The information gained from large-scale human protein interaction studies will ultimately lead to the discovery of novel pathway associations and therapeutic targets.
References and recommended reading Papers of particular interest, published within the annual period of review, have been highlighted as: of special interest of outstanding interest 1.
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