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Application of Mu in vitro transposition for high-precision mapping of protein–protein interfaces on a yeast two-hybrid platform
Methods 49 (2009) 255–262
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Methods journal homepage: www.elsevier.com/locate/ymeth
Application of Mu in vitro transposition for high-precision mapping of protein–protein interfaces on a yeast two-hybrid platform Maria Pajunen a,b, Eini Poussu a, Hilkka Turakainen a, Harri Savilahti a,b,* a b
Program in Cellular Biotechnology, Institute of Biotechnology, Viikki Biocenter, University of Helsinki, Finland Division of Genetics and Physiology, Department of Biology, University of Turku, Finland
a r t i c l e
i n f o
Article history: Accepted 20 April 2009 Available online 3 May 2009 Keywords: Protein–protein interaction Protein interface Yeast two-hybrid analysis Scanning mutagenesis Transposon techniques
a b s t r a c t High-precision mapping of regions involved in protein–protein interfaces of interacting protein partners is an essential component on a path to understand various cellular functions. Transposon-based systems, particularly those involving in vitro reactions, offer exhaustive insertion mutant libraries and highthroughput platforms for many types of genetic analyses. We present here a genetic strategy to accurately map interacting protein regions at amino acid precision that is based on transposition-assisted construction, sampling, and analysis of a comprehensive insertion mutant library. The methodology integrates random pentapeptide mutagenesis of proteins, yeast two-hybrid screening, and high-resolution genetic footprinting. This straightforward strategy is general, and it provides a rapid and easy means to identify critical contact regions in proteins without the requirement of prior structural knowledge. Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction 1.1. Methods to study protein–protein interactions A detailed description of networks of protein–protein interactions is needed for the comprehensive understanding of cell function [1]. The initial identification of interacting protein partners can be readily accomplished using currently available methodologies such as the yeast two-hybrid system [2], tandem affinity purification of protein complexes [3], and computational predictions [4]. However, mapping of the interacting protein interfaces of identified protein partners poses a major challenge, as fully streamlined molecular techniques are lacking. The current means to identify regions involved in protein–protein interactions include mutational analyses (e.g. deletion series and alanine scanning), protein footprinting with proteases [5] or hydroxyl-radicals [6,7], chemical cross-linking combined with mass spectrometry [8], hydrogen–deuterium exchange experiments [9], and structural studies by NMR or X-ray crystallography. Some of the methods rely on time- and labor-consuming production of individual protein mutant variants, and certain methods lack optimal resolution. In addition, several of the methods require highly specialized instrumentation and technical skills. Thus, any methodology that would streamline the process of mapping protein–protein interfaces would be highly beneficial. * Corresponding author. Address: Division of Genetics and Physiology, Department of Biology, University of Turku, Vesilinnantie 5, 20014 Turku, Finland. Fax: +358 2 333 6680. E-mail address: harri.savilahti@utu.fi (H. Savilahti). 1046-2023/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2009.04.014
We present a system for comprehensive mapping of protein– protein interfaces at high resolution without the requirement to construct or isolate mutants separately and without the need for protein structural information. The system is feasible for the study of any interacting protein partner that is amenable to yeast twohybrid screening, and the analysis can be performed in any standard molecular biology laboratory. 1.2. Transposition-based genetic footprinting techniques Genetic footprinting strategies take advantage of large-scale random transposon mutagenesis protocols coupled with an appropriate genetic selection of a gene function to identify genes or gene regions essential for that function [10,11]. On a single-protein scale, genetic footprinting is applicable for the analysis of functional protein regions [12–14]. Genetic footprinting protocols involve the initial generation of a large library of transposon insertion mutations and subsequent negative selection for a certain function en masse; mutants with insertions in essential regions for this particular function are excluded from the selected pool. The mutant pools collected before and after selection are analyzed by PCR. Several transposons have been used in various footprinting studies, such as Mu (e.g. [14,15]) and Ty1 [10], but also a retroviral integrase has been utilized for genetic footprinting (e.g. [11]). In addition to selection, screening methods can also be used as a sampling system for genetic footprinting analyses. Yeast two-hybrid screening provides a straightforward means to identify interacting protein partners. We show that it provides also a platform for the identification of critical contact regions between proteins.
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1.3. Mu for mutagenesis Transposable elements are indispensable tools in modern genetics, and their ability to insert essentially randomly into DNA enables the generation of exhaustive insertion mutant libraries
[16]. One of the most versatile DNA transposition tools is the in vitro reaction derived from bacteriophage Mu transposition [17,18]. This system requires only a simple reaction buffer and three purified macromolecular components: transposon DNA, MuA transposase, and target DNA (typically a gene of interest
Fig. 1. Experimental outline. (A) Flowchart of the high-precision detection of protein–protein interfaces. (B) Construction of the 15-bp insertion mutant library. (C) Yeast twohybrid screen. The yeast strain EGY48 carrying the lacZ marker gene-containing plasmid (p8op-lacZ) is transformed with the plasmid (pGildaB-BAIT) expressing the bait fusion protein and subsequently with the mutant library encoding PREY variants cloned in a modified pB42AD plasmid (pMPH11). Plasmid-containing clones are identified on X-gal-containing selection plates. Clones from white, pale blue, and blue colonies are collected as streaks and grown to form pools of no, weak, or strong protein–protein interaction, respectively. (D) PCR-based genetic footprinting strategy. The prey gene region is initially amplified from the insertion mutant library using vector-specific primers. Secondary amplification is done using a biotinylated vector-specific primer and a radioactively labeled prey gene-specific primer. Following pull-down with streptavidin beads, the PCR products are digested with NotI (recognition site within the 15-bp insertion), and the soluble fraction is analyzed by denaturing PAGE and autoradiography.
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cloned in an appropriate plasmid). The reaction is highly efficient with relatively low target site selectivity [18,19]. These characteristics make the Mu in vitro reaction ideal for the generation of comprehensive mutant DNA libraries usable in a variety of molecular biology applications [15,20–26]. One of the applications is a strategy that can be used to generate essentially randomly distributed five amino-acid insertions in proteins [21,25]. It is also possible to distribute one amino acid deletions and substitutions [27,28], as well as entire protein domains [29]. 1.4. Identifying interacting protein regions at amino acid precision The mapping strategy presented combines (i) the generation of a pentapeptide insertion mutant library for the protein of interest, (ii) en masse screening for altered protein–protein association on a yeast two-hybrid platform, and (iii) parallel analysis of mutant pools using a PCR-based genetic footprinting technique. Overall, the strategy provides a convenient general means to accurately map interacting regions in protein partners. The fully optimized system is applicable to any protein-encoding gene, given that the behavior of the analyzed proteins is compatible with yeast two-hybrid analysis. Besides the yeast platform, the methodology should be readily modifiable to function on bacterial two-hybrid platforms [30] as well.
2. Experimental approach The strategy to fine-map a protein region involved in a specific protein–protein interaction is based on a robust DNA transposition-based en masse insertion mutagenesis system, visual yeast two-hybrid screen, and high-resolution genetic footprinting technique (Fig. 1A). First, a pentapeptide insertion mutant library is generated for the protein partner to be analyzed (Fig. 1B). Then, yeast two-hybrid screen is used to distinguish between those protein variants that are able to interact with the partner protein and those with altered or lost interaction due to a five amino-acid insertion (Fig. 1C). Next, phenotypically different clone classes are pooled and positions of the insertions in each pool are located using a PCR-based footprinting strategy (Fig. 1D). Finally, the critical insertion sites can be mapped to the protein’s primary and, if available, tertiary structure. 2.1. Pentapetide mutagenesis Pentapeptide mutagenesis strategy (Fig. 1B) can be used to generate essentially randomly distributed five amino-acid insertions in proteins with high accuracy and 100% efficiency [25]. In the final protein-encoding plasmid mutant library all of the member clones contain only one insertion. In the current protein interface detection application, the gene of interest is initially cloned into any suitable plasmid to initiate the mutant library construction. In the subsequent steps, the library is finalized as a mutant pool in an expression plasmid compatible with the yeast two-hybrid system. For the purpose, we have modified the commercial pB42AD plasmid vector of the MATCHMAKER kit to include a kanamycin resistance cassette (npt), which can be used to counterselect in Escherichia coli against the other two ampicillin resistance-encoding (bla) plasmids present in the yeast two-hybrid strain (Fig. 1C). This is a favorable feature, as the selected mutant pools will be shuttled from Saccharomyces cerevisiae into E. coli. The gene of interest should be cloned in the initial vector between unique restriction sites such that, following manipulation by transposons, it can be transferred into the final yeast two-hybrid expression plasmid in a correct orientation for protein expression. Importantly, as one of the critical modification steps will be done using
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NotI digestion and religation, both the final vector and the gene of interest must not contain NotI sites. In the final mutant library, each plasmid clone will contain a unique NotI site within the 15-bp insertion, and this site will be exploited later in the genetic footprinting protocol. During the mutagenesis, plasmid clones will be plated three times and sampled twice. Taking into account the initial number of clones and Poisson distribution during sampling, the number of independently generated mutants, present in the final pool, can be estimated. We typically aim at a minimum of 10-fold insertion-per-nucleotide coverage within the target gene to guarantee a high degree of saturation mutagenesis. For example, to successfully cover a gene of 1000 bp in length, a minimum of 10,000 independent clones would be generated. 2.2. Yeast two-hybrid platform In standard yeast two-hybrid protocols, cells are initially selected on the basis of their plasmid content on glucose-containing plates, yielding well-growing yeast colonies. These colonies are subsequently replica-plated onto a two-hybrid screening medium where growth conditions induce the expression of interacting protein partners. In most systems, galactose serves as an inducer via activation of GAL1 promoters, resulting in blue colonies on X-gal indicator plates upon interaction. A simpler protocol used for the analysis presented here utilizes conditions that allow relatively good cell growth and also induce sufficient protein expression, thereby permitting color screening directly on the original transformation plates without prior replica-plating. The plating medium is SD, -Ura, -His, -Trp supplemented with 100 mM sodium phosphate (pH 7.0), 0.6% glucose, 1.4% galactose, and 200 lg/ml X-gal. This protocol yields both deep blue and clean white colonies, with interacting and non-interacting protein pairs, respectively. Colonies with intermediate color (pale blue) will be generated by protein mutants with altered but not totally destroyed binding capacity. The system generates a low-frequency background of false positive and false negative data, as indicated by the appearance of 1–2% of colonies with a switched color phenotype following the reintroduction of plasmids from ‘‘blue” or ‘‘white” pools [14]. However, this low background is tolerable, and it does not interfere with the data analysis. Following screening in yeast, different color phenotype mutants are sampled as pools. Plasmid DNA is isolated from the pools and then introduced into E. coli. At this stage, it is possible to enumerate the total number of clones in a given pool. Subsequent propagation of the pools and isolation of plasmids from E. coli ensures good quality template DNA for PCR-based genetic footprinting analysis. 2.3. Genetic footprinting Analysis of mutants as pools is arguably the most effective means to exploit the potential of an insertion mutant library, as a large number of mutants can be analyzed simultaneously. Initially the entire gene of interest is amplified using a vector-specific primer pair and DNA from the selected mutant library as a template. The amplified segment is then used as a template to amplify shorter gene segments using primer pairs consisting of a radioactively labeled and a biotinylated primer (Fig. 1D). Following pull-down with streptavidin-coated beads and subsequent NotI digestion, reaction products are analyzed by denaturing polyacrylamide gel electrophoresis and autoradiography (Fig. 2). The maximum number of clones present in each analyzed plasmid pool is not very critical. Nevertheless, a moderate number of pooled clones (e.g. 100–600) appear to be optimally suited for the analysis, as the distribution of radioactive label in these cases is restricted to fewer and thus more intensely labeled reaction products, generating easily interpretable visual data (Fig. 3).
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Fig. 2. Genetic footprinting with radioactively labeled primers. On the top is shown the overall PCR strategy. Each radioactively labeled primer (asterisk-labeled arrows specified with A, B, C, D, etc.) forms a pair with biotin-labeled oligonucleotide (Bio-labeled arrow). With several primer pairs the entire prey-expressing gene region can be analyzed. In this example, a total of 174 white (W) and 35 pale blue (PB) colonies were picked to form pools of no protein–protein interaction and weak interaction, respectively. Six independent pools of blue colonies (100 clones each, labeled 1–6) were also collected to represent strong interaction. PCR, denaturing PAGE, and autoradiography were used to analyze the pools. The wild-type prey clone (wt) served as a negative control. The size marker lanes (M) contain a representative selection of individually sequenced insertion variants [14] with a known insertion site (indicated as a nucleotide position above each band), allowing accurate mapping of insertion sites in the footprinting data. Sequencing reactions (C and G) assist in the estimation of band spacing in different parts of the autoradiograph.
3. Materials 3.1. Reagents Escherichia coli strain DH10B (Invitrogen) [31]. This strain yields high-efficiency electrocompetent cells with the protocol used [32]. It also yields excellent quality plasmid DNA for the plasmid library construction. Bacto-agar, Bacto-tryptone, Bacto-yeast extract, and Bacto-yeast nitrogen base (Difco).
Ampicillin, kanamycin, and chloramphenicol (Sigma). Plasmid pEGFP-C1 (Clontech) or similar cloning vector. Note! The selectable marker must not be the same as that present in the transposon used. MATCHMAKER LexA two-hybrid system (Clontech). Plasmid pMPH11 [14]. This is a modified version of pB42AD of MATCHMAKER. Importantly, the plasmid includes a modified kanamycin resistance cassette (XhoI site removed) that interrupts the ampicillin resistance-conferring (bla) gene. As importantly, the plasmid does not contain any NotI sites.
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Fig. 3. An example of data analysis. Location of five amino-acid insertions within the human JFC1 protein (shown schematically on the top) in relation to observed interaction with human Rab8A protein as determined by sequencing or genetic footprinting. The schematic structure of the JFC1 protein indicates the location of the SHD and C2 domains. The interaction interface is located within the SHD1 domain.
Mutation Generation System, MGSTM (Finnzymes), including the cat-Mu(NotI) transposon [25]. Plasmid DNA isolation kits (Qiagen). Qiaquick Gel Extraction kit (Qiagen). PCR Clean-up Nucleospin Extract II kit (Macherey–Nagel, Germany). Seakem LE and Seaplaque GTG agaroses (Lonza). Appropriate restriction endonucleases for cloning steps (New England Biolabs). NotI for library construction and footprinting (New England Biolabs). T4 DNA ligase, T4 polynucleotide kinase, and Vent DNA Polymerase (New England Biolabs). High quality oligonucleotide primers (see Table 1; all our primers have been purchased from the Howard Hughes Medical Institute/Keck Oligonucleotide Synthesis Faculty at Yale University). 10 mM dNTP mix (Finnzymes). DyNAzyme II DNA Polymerase (Finnzymes). Sequenase 2.0 sequencing kit (USB) for the generation of sequencing ladders as size markers. Table 1 Oligonucleotide primers.# Primer
Sequence (50 ? 30 )
Description
HSP508
CCGCCGATCCAGCCTGACTG
HSP509
GTGTCAACAACGTATCTACCAACG
Bio-HSP488
Biotin-GACAAGCCGACAACCTTGATTG
Non-radioactive PCR, pB42AD forward Non-radioactive PCR, pB42AD reverse pB42AD reverse
[c-33P]ATP (1000–3000 Ci/mmol) (GE Healthcare). Streptavidin-coated magnetic beads (Roche). Micro Bio-spin 30 columns (Bio-Rad). 40% acrylamide/bis solution 19:1 (Bio-Rad). Amberlite deionizing resin, IRN-150L (GE Healthcare).
3.2. Equipment Petri dishes (Sterilin). Large 24.5 cm 24.5 cm plates (Corning 431111). 5-mm glass beads for spreading of yeast cultures on plates (approx. 10 beads per a large plate). 0.5-mm glass beads for plasmid isolation from yeast cells. 37 °C shaker (New Brunswick Scientific). 37 °C incubator (Termaks). Centrifuges (Heraeus and Sorvall). DNA electrophoresis equipment (Bio-Rad). Electroporator (Bio-Rad Genepulser II). Thermocyclers, Robocycler Gradient40 (Stratagene) and MiniCyclerTM (MJ Research). Magnetic particle separator (Roche). Test-tube-rotator (Snijders 34528). Sequencing gel apparatus (Life Technologies). Gel dryer (Savant SpeedGel). Phosphorimager (Fuji). Whatman 1 and Whatman 3MM paper. Saran wrap.
3.3. Reagent setup
#
In addition to the primers shown, the strategy employs several gene-specific forward primers that are 50 -labeled with [c-33P]ATP prior to PCR (see Fig. 2). The interval between the binding sites of labeled primers should be approx. 200 bp. This arrangement generates overlapping data, and thus the majority of the gene regions are analyzed twice, yielding high quality data.
Luria Broth (LB) medium, 1 l. 10 g Bacto-tryptone, 5 g Bacto-yeast extract and 5 g NaCl. Adjust pH to 7.0 with 5 M NaOH. Autoclave. Store at RT for up to 1 month.
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LB agar plates. Add 15 g Bacto-agar to 1 liter of LB medium before autoclaving. Cool down to 55 °C, supplement with suitable antibiotics and dispense approx. 20 ml per Petri dish. These plates can be stored at 4 °C for 1 month. Ampicillin (Ap) solution. Dissolve ampicillin in distilled water at 100 mg/ml and pass the solution through a 0.22-lm pore filter. Can be stored at 20 °C for 1 year. Ampicillin is used at the final concentration of 100 lg/ml. Chloramphenicol (Cm) solution. Dissolve chloramphenicol in absolute ethanol at 20 mg/ml. The solution can be stored at 20 °C for 1 year. Chloramphenicol is used at the final concentration of 10 lg/ml. Kanamycin (Km) solution. Dissolve kanamycin in distilled water at 25 mg/ml and pass the solution through a 0.22-lm pore filter. The solution can be stored at 20 °C for 1 year. Kanamycin is used at the final concentration of 25 lg/ml. X-gal. Dissolve at 20 mg/ml in dimethylformamide (DMF). Store in glass or polypropylene bottles at 20 °C in the dark. P1 buffer . 50 mM Tris–HCl, pH 8.0, 10 mM EDTA, 100 lg/ml RNase A. TE buffer. 10 mM Tris–HCl, pH 7.5, 0.5 mM EDTA. TEN buffer. 10 mM Tris–HCl, pH 7.5, 0.5 mM EDTA, 50 mM NaCl. TEN100 buffer . 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 100 mM NaCl. TEN1000 buffer. 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 1 M NaCl. 2 binding buffer . 10 mM Tris–HCl, pH 7.5, 2 mM EDTA, 200 mM NaCl. 2 formamide loading dye . 95% deionized formamide, 10 mM EDTA, 0.1% bromo-phenol blue, 0.1% xylene cyanol. 7 M urea, 6% polyacrylamide solution, 200 ml. Dissolve 84 g urea and 30 ml 40% acrylamide in 50 ml distilled water at 37 °C bath, deionize with Amberlite resin (5 g) for 1–2 h, and pass the solution through two Whatman 1 papers. Add 20 ml 10 TBE and adjust volume to 200 ml with distilled water. Store at 4 °C for up to 1 month.
4. Methodology 4.1. Plasmid constructions (1) Standard DNA techniques are performed as previously described [33]. (2) Insert the wild-type prey gene into any appropriate cloning vector to generate an initial target plasmid for pentapeptide mutagenesis. Because a gel isolation procedure will be used during the library construction, choose a vector plasmid whose size is different than that of the total length of the prey gene plus transposon (cat-Mu(NotI) transposon is 1254 bp in length). In addition, the cloning sites in the vector need to be compatible with those of pMPH11 (EcoRI, NheI, StuI, NcoI, SfiI, XmaI, BglII, and XhoI), the final expression plasmid for the two-hybrid analysis. Plasmid pMPH11 [14] is a modified version of pB42AD of the MATCHMAKER kit, and it is available upon request. The modifications (elimination of a critical NotI site and an insertion of the npt gene) have been described in detail previously [14]. The prey gene must not contain any NotI sites, as this restriction endonuclease is used to remove the transposon bulk during the library construction. If it does, remove the site(s) using any of the currently available site-directed mutagenesis protocols. (3) Clone the wild-type prey gene into pMPH11 using the same cloning strategy that you chose to use for the mutant library. This construct will be used as a positive control in the yeast two-hybrid analysis and as a negative control in genetic footprinting. (4) Clone the bait gene into pGildaB provided by the MATCHMAKER kit.
4.2. Generation of the pentapeptide insertion mutant library (5) The pentapeptide insertion mutant library is generated using the Mutation Generation SystemTM (Finnzymes) essentially as specified by the supplier. This mutagenesis system exploits the MuA transposase-catalyzed in vitro transposition reaction [18] and generates 5-aa insertions in proteins [21,25]. (6) Prepare five standard in vitro transposition reactions using cat-Mu(NotI) [25] as a donor DNA. Note! MuA transposase should be added last. Component Target plasmid (100 ng/ll) M1-CamR transposon (100 ng/ll) 5 reaction buffer for MuA transposase Nuclease-free water MuA transposase (220 ng/ll)
Amount (ll) 3 1 4 11 1
Final 300 ng 100 ng 1 Final volume 20 ll 220 ng
(7) Incubate at 30 °C for 3 h, and then inactivate MuA transposase by incubating at 75 °C for 10 min. (8) Pool the reactions and extract with phenol and subsequently with chloroform. Ethanol precipitate the DNA and resuspend it in water (25 ll). (9) Electroporate several aliquots (2 ll) of DNA into DH10B electrocompetent cells (50 ll) prepared as previously described [32]. (10) Select transposon-containing plasmid clones by plating the bacteria on LB agar plates supplemented with chloramphenicol (Cm, 10 lg/ml) and kanamycin (Km, 25 lg/ml). To avoid background growth, do not plate more than approx. 1000 CFU per plate. (11) Pool an appropriate number of clones by scraping transformant colonies from the plates and grow them in LB–Cm–Km medium at 37 °C for 3 h. In general, we use the length of the gene of interest (in base pairs) and multiply that by 10 to form an estimate for a minimum final pool size. For example: if the gene of interest is 1700 bp, we would need at least 17,000 independently generated clones in the final pool (see step 25 for considerations about sampling). (12) Isolate plasmid DNA from the pool and digest the DNA with appropriate restriction enzymes to release the DNA of interest (the target gene hit by the transposon) from the cloning vector. (13) Separate the resulting fragments by preparative electrophoresis using 0.8% Seaplaque GTG agarose gel in TAE buffer [33]. (14) Isolate from the gel the DNA fragment (pool) that corresponds to transposon insertions into the PREY-encoding DNA segment. We have used a standard electroelution protocol for the purpose, but commercial gel isolation kits may substitute. (15) Ligate the isolated DNA fragment into pMPH11 digested with the same enzymes that were used in step 12. (16) Electroporate the ligation mixture into DH10B electrocompetent cells (50 ll) as specified in step 9. (17) Select transposon-containing plasmid clones by plating the bacteria on LB–Cm–Km plates as specified in step 10. (18) Pool and grow the transformants as specified in step 11. (19) Prepare plasmid DNA from the pool. (20) To eliminate the transposon core sequence from the plasmid pool, digest the plasmid pool with NotI and separate the linearized fragment using preparative electrophoresis on a 1.7% Seaplaque GTG agarose gel.
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(21) Isolate the plasmid backbone as specified in step 14 and recircularize it by ligation at low DNA concentration (approx. 1 ng/ll). (22) Electroporate ligated plasmids into DH10B cells as specified in step 9. (23) Select the transformants on LB–Km plates to generate the final library. Note! Chloramphenicol selection is not used at this stage. (24) Pool and grow transformants in LB–Km medium as specified in step 11. (25) Prepare plasmid DNA from the pool. The insertion library is now ready for your specific application. Note that during the library construction, the original plasmid pool is sampled several times. As the number of clone members in each sampling is known, Poisson distribution can be used to estimate the number of independently generated mutants in the final library. 4.3. Yeast two-hybrid system (26) The yeast two-hybrid protein interaction screen is performed using the MATCHMAKER LexA Two-Hybrid System (Clontech) according to the manufacturer’s specifications. The standard yeast media used are described in the Yeast Protocols Handbook (Clontech). (27) Transform pGildaB-BAIT plasmid into S. cerevisiae reporter strain EGY48 (MATa, his3, trp1, ura3, LexAop(x6)-LEU2) harboring the plasmid p8op-lacZ. Plate on SD-Ura-His agar. (28) Introduce the final pMPH11-PREY insertion mutant library into the above reporter strain harboring the above two plasmids. Plate on SD, -Ura, -His, -Trp agar supplemented with 100 mM sodium phosphate (pH 7.0), 0.6% glucose, 1.4% galactose, 0.2 mg/ml X-gal. We have used large (24.5 cm 24.5 cm) plates. (29) Streak the colonies on Petri dishes (medium from step 28) and grow them to verify their color phenotype. Individual insertion mutants can be isolated and sequenced at this point. These mutants can be used as convenient size markers in the footprinting step. (30) Pool colonies on the basis of their color phenotype and grow them in SD, -Ura, -His, -Trp medium at 30 °C for 3 h. We prefer using pool sizes less than 200 member clones, and recommend generating several ‘‘blue” pools in parallel if necessary. A pool size with about 600 member clones approaches current detection limit [14]. (31) Prepare plasmid DNA from the pools using the Qiagen Plasmid Spin mini kit. Vortex the cells in P1 buffer with glass beads for 10–30 min prior to DNA isolation. (32) Introduce the mutant plasmid pools into E. coli DH10B by electroporation as specified in step 9 using LB–Km plates for selection. Compared to the original pool size, at least 10-fold excess colonies should be obtained after electroporation. For example, a pool of 100 member clones in yeast should be plated as approx. 1000 colonies (or more) in E. coli. (33) Scrape the transformant colonies from the plates and prepare plasmid DNA from each pool. DNA preparations are now ready for PCR-based genetic footprinting. 4.4. PCR (34) Prepare the non-radioactive PCR in a total volume of 50 ll. Component Plasmid DNA (50 ng/ll) 10 mM dNTP mix Primer HSP508 (5 pmol/ll) Primer HSP509 (5 pmol/ll)
Amount (ll) 1 1 5 5
Final 50 ng 200 lM 0.25 lM 0.25 lM
100 mM MgSO4 (contributes 2 mM) 10 ThermoPol reaction buffer Vent DNA polymerase (2 U/ll) Nuclease-free water
1
4 mM
5 0.5 31.5
1 1U Final volume 50 ll
(35) Amplify in the thermocycler (Robocycler) using the following parameters: 5 min at 95 °C followed by 25 cycles of 1 min at 95 °C, 1 min at 60 °C, and 2 min at 72 °C, and finally 5 min at 72 °C. (36) Purify PCR products electrophoretically using the Qiaquick Gel Extraction kit. (37) Label the prey-specific primers radioactively; 50 -labeling with [c-33P]ATP and T4 polynucleotide kinase is performed as previously described [33]. (38) Prepare the radioactive label-containing PCR in a total volume of 50 ll. Using a PCR protocol with one biotinylated (nested) primer increases specificity and decreases background radioactivity in the final gel analysis. Component Non-radioactive PCR product (50 ng/ll) 10 mM dNTP mix Primer Bio-HSP488 (5 pmol/ll) Labeled prey-specific primer (5 pmol/ll) 10 Optimized DyNAzyme Buffer DyNAzyme II DNA polymerase (2 U/ll) Nuclease-free water
Amount (ll)
Final
1
50 ng
1 5 5
200 lM 0.25 lM 0.25 lM
5 0.5
1 1U
32.5
Final volume 50 ll
(39) Amplify in the thermocycler (MiniCycler) using otherwise same parameters as specified in step 35 except that the extension time at 72 °C can be variable (typically from 45 s to 120 s) depending on the length of the desired PCR product. (40) Purify the radioactively labeled PCR product using the PCR Clean-up Nucleospin Extract II kit and elute into 30 ll of 10 mM Tris–HCl, pH 8.5. 4.5. Genetic footprinting (41) Pre-wash streptavidin beads four times with TEN100 and once with 2 binding buffer. Mix the beads with 2 binding buffer using the ratio of 13 volume beads and 17 volume buffer. Use this suspension in step 42. We have done all the washing steps using a magnetic particle separator. Alternatively, washing steps can be done by centrifugation. (42) Add 30 ll of radioactively labeled PCR product to the prewashed streptavidin bead suspension (30 ll) and incubate at room temperature for at least 1 h to allow adsorption. To avoid sedimenting of the beads, use a test-tube-rotator and/or mix occasionally. (43) Wash the beads three times with 0.5 ml of TEN1000 and twice with 1 restriction enzyme buffer 3 (New England Biolabs). (44) Add 50 ll of 1 restriction enzyme buffer 3 containing NotI (25 U) and incubate at least for 4 h at 37 °C. Mix occasionally. (45) Remove the beads using the magnetic particle separator and subsequently purify the supernatant by centrifugation through a Micro Bio-spin 30 column (equilibrated with TEN buffer) at room temperature in a tabletop microcentrifuge (3000 rpm, 4 min).
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(46) Ethanol precipitate the DNA and resuspend it in 3 ll of TE buffer. Add 3 ll of 2 formamide loading dye for gel analysis. (47) Analyze samples by 7 M urea, 6% polyacrylamide gel electrophoresis as previously described [34]. Appropriate sequencing reactions are used as size markers in the analysis. Individually sequenced insertion variants (see step 29), each with a known insertion site, can be used to accurately calibrate the data. (48) Dry gel at 80 °C onto Whatman 3MM paper. (49) Visualize the bands by autoradiography. We have used a Fuji BAS 1500 phosphorimager with BAS-Reader 2.9 software (Raytest). (50) Analyze the data to localize the interacting interface.
[4] [5] [6] [7] [8] [9] [10] [11] [12]
5. Anticipated results
[13] [14]
Functional analysis of a protein interface involved in an interaction with another protein can be accomplished by subjecting the selected protein mutant pools to comparative parallel analysis using a PCR-based strategy. Following gel analysis and autoradiography, this type of genetic footprinting assay generates a visual read-out where reciprocal band patterns can be seen between the insertion mutant pools representing unaltered versus altered protein–protein interaction. In yeast two-hybrid screen, the percentage of white colonies should be indicative of the relative proportion of the interacting interface in the protein. For preliminary data, mutant clones can be analyzed individually by sequencing.
[15]
Acknowledgment This study has been supported by the Academy of Finland and the National Technology Agency of Finland (TEKES). References [1] M.E. Cusick, N. Klitgord, M. Vidal, D.E. Hill, Hum. Mol. Genet. 14 (2005) R171– R181. [2] S. Fields, O. Song, Nature 340 (1989) 245–246. [3] A.-C. Gavin, M. Bösche, R. Krause, P. Grandi, M. Marzioch, A. Bauer, J. Schultz, J.M. Rick, A.-M. Michon, C.-M. Cruciat, M. Remor, C. Höfert, M. Schelder, M. Brajenovic, H. Ruffner, A. Merino, K. Klein, M. Hudak, D. Dickson, T. Rudi, V. Gnau, A. Bauch, S. Bastuck, B. Huhse, C. Leutwein, M.-A. Heurtier, R.R. Copley, A.
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Methods journal homepage: www.elsevier.com/locate/ymeth
Application of Mu in vitro transposition for high-precision mapping of protein–protein interfaces on a yeast two-hybrid platform Maria Pajunen a,b, Eini Poussu a, Hilkka Turakainen a, Harri Savilahti a,b,* a b
Program in Cellular Biotechnology, Institute of Biotechnology, Viikki Biocenter, University of Helsinki, Finland Division of Genetics and Physiology, Department of Biology, University of Turku, Finland
a r t i c l e
i n f o
Article history: Accepted 20 April 2009 Available online 3 May 2009 Keywords: Protein–protein interaction Protein interface Yeast two-hybrid analysis Scanning mutagenesis Transposon techniques
a b s t r a c t High-precision mapping of regions involved in protein–protein interfaces of interacting protein partners is an essential component on a path to understand various cellular functions. Transposon-based systems, particularly those involving in vitro reactions, offer exhaustive insertion mutant libraries and highthroughput platforms for many types of genetic analyses. We present here a genetic strategy to accurately map interacting protein regions at amino acid precision that is based on transposition-assisted construction, sampling, and analysis of a comprehensive insertion mutant library. The methodology integrates random pentapeptide mutagenesis of proteins, yeast two-hybrid screening, and high-resolution genetic footprinting. This straightforward strategy is general, and it provides a rapid and easy means to identify critical contact regions in proteins without the requirement of prior structural knowledge. Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction 1.1. Methods to study protein–protein interactions A detailed description of networks of protein–protein interactions is needed for the comprehensive understanding of cell function [1]. The initial identification of interacting protein partners can be readily accomplished using currently available methodologies such as the yeast two-hybrid system [2], tandem affinity purification of protein complexes [3], and computational predictions [4]. However, mapping of the interacting protein interfaces of identified protein partners poses a major challenge, as fully streamlined molecular techniques are lacking. The current means to identify regions involved in protein–protein interactions include mutational analyses (e.g. deletion series and alanine scanning), protein footprinting with proteases [5] or hydroxyl-radicals [6,7], chemical cross-linking combined with mass spectrometry [8], hydrogen–deuterium exchange experiments [9], and structural studies by NMR or X-ray crystallography. Some of the methods rely on time- and labor-consuming production of individual protein mutant variants, and certain methods lack optimal resolution. In addition, several of the methods require highly specialized instrumentation and technical skills. Thus, any methodology that would streamline the process of mapping protein–protein interfaces would be highly beneficial. * Corresponding author. Address: Division of Genetics and Physiology, Department of Biology, University of Turku, Vesilinnantie 5, 20014 Turku, Finland. Fax: +358 2 333 6680. E-mail address: harri.savilahti@utu.fi (H. Savilahti). 1046-2023/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2009.04.014
We present a system for comprehensive mapping of protein– protein interfaces at high resolution without the requirement to construct or isolate mutants separately and without the need for protein structural information. The system is feasible for the study of any interacting protein partner that is amenable to yeast twohybrid screening, and the analysis can be performed in any standard molecular biology laboratory. 1.2. Transposition-based genetic footprinting techniques Genetic footprinting strategies take advantage of large-scale random transposon mutagenesis protocols coupled with an appropriate genetic selection of a gene function to identify genes or gene regions essential for that function [10,11]. On a single-protein scale, genetic footprinting is applicable for the analysis of functional protein regions [12–14]. Genetic footprinting protocols involve the initial generation of a large library of transposon insertion mutations and subsequent negative selection for a certain function en masse; mutants with insertions in essential regions for this particular function are excluded from the selected pool. The mutant pools collected before and after selection are analyzed by PCR. Several transposons have been used in various footprinting studies, such as Mu (e.g. [14,15]) and Ty1 [10], but also a retroviral integrase has been utilized for genetic footprinting (e.g. [11]). In addition to selection, screening methods can also be used as a sampling system for genetic footprinting analyses. Yeast two-hybrid screening provides a straightforward means to identify interacting protein partners. We show that it provides also a platform for the identification of critical contact regions between proteins.
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1.3. Mu for mutagenesis Transposable elements are indispensable tools in modern genetics, and their ability to insert essentially randomly into DNA enables the generation of exhaustive insertion mutant libraries
[16]. One of the most versatile DNA transposition tools is the in vitro reaction derived from bacteriophage Mu transposition [17,18]. This system requires only a simple reaction buffer and three purified macromolecular components: transposon DNA, MuA transposase, and target DNA (typically a gene of interest
Fig. 1. Experimental outline. (A) Flowchart of the high-precision detection of protein–protein interfaces. (B) Construction of the 15-bp insertion mutant library. (C) Yeast twohybrid screen. The yeast strain EGY48 carrying the lacZ marker gene-containing plasmid (p8op-lacZ) is transformed with the plasmid (pGildaB-BAIT) expressing the bait fusion protein and subsequently with the mutant library encoding PREY variants cloned in a modified pB42AD plasmid (pMPH11). Plasmid-containing clones are identified on X-gal-containing selection plates. Clones from white, pale blue, and blue colonies are collected as streaks and grown to form pools of no, weak, or strong protein–protein interaction, respectively. (D) PCR-based genetic footprinting strategy. The prey gene region is initially amplified from the insertion mutant library using vector-specific primers. Secondary amplification is done using a biotinylated vector-specific primer and a radioactively labeled prey gene-specific primer. Following pull-down with streptavidin beads, the PCR products are digested with NotI (recognition site within the 15-bp insertion), and the soluble fraction is analyzed by denaturing PAGE and autoradiography.
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cloned in an appropriate plasmid). The reaction is highly efficient with relatively low target site selectivity [18,19]. These characteristics make the Mu in vitro reaction ideal for the generation of comprehensive mutant DNA libraries usable in a variety of molecular biology applications [15,20–26]. One of the applications is a strategy that can be used to generate essentially randomly distributed five amino-acid insertions in proteins [21,25]. It is also possible to distribute one amino acid deletions and substitutions [27,28], as well as entire protein domains [29]. 1.4. Identifying interacting protein regions at amino acid precision The mapping strategy presented combines (i) the generation of a pentapeptide insertion mutant library for the protein of interest, (ii) en masse screening for altered protein–protein association on a yeast two-hybrid platform, and (iii) parallel analysis of mutant pools using a PCR-based genetic footprinting technique. Overall, the strategy provides a convenient general means to accurately map interacting regions in protein partners. The fully optimized system is applicable to any protein-encoding gene, given that the behavior of the analyzed proteins is compatible with yeast two-hybrid analysis. Besides the yeast platform, the methodology should be readily modifiable to function on bacterial two-hybrid platforms [30] as well.
2. Experimental approach The strategy to fine-map a protein region involved in a specific protein–protein interaction is based on a robust DNA transposition-based en masse insertion mutagenesis system, visual yeast two-hybrid screen, and high-resolution genetic footprinting technique (Fig. 1A). First, a pentapeptide insertion mutant library is generated for the protein partner to be analyzed (Fig. 1B). Then, yeast two-hybrid screen is used to distinguish between those protein variants that are able to interact with the partner protein and those with altered or lost interaction due to a five amino-acid insertion (Fig. 1C). Next, phenotypically different clone classes are pooled and positions of the insertions in each pool are located using a PCR-based footprinting strategy (Fig. 1D). Finally, the critical insertion sites can be mapped to the protein’s primary and, if available, tertiary structure. 2.1. Pentapetide mutagenesis Pentapeptide mutagenesis strategy (Fig. 1B) can be used to generate essentially randomly distributed five amino-acid insertions in proteins with high accuracy and 100% efficiency [25]. In the final protein-encoding plasmid mutant library all of the member clones contain only one insertion. In the current protein interface detection application, the gene of interest is initially cloned into any suitable plasmid to initiate the mutant library construction. In the subsequent steps, the library is finalized as a mutant pool in an expression plasmid compatible with the yeast two-hybrid system. For the purpose, we have modified the commercial pB42AD plasmid vector of the MATCHMAKER kit to include a kanamycin resistance cassette (npt), which can be used to counterselect in Escherichia coli against the other two ampicillin resistance-encoding (bla) plasmids present in the yeast two-hybrid strain (Fig. 1C). This is a favorable feature, as the selected mutant pools will be shuttled from Saccharomyces cerevisiae into E. coli. The gene of interest should be cloned in the initial vector between unique restriction sites such that, following manipulation by transposons, it can be transferred into the final yeast two-hybrid expression plasmid in a correct orientation for protein expression. Importantly, as one of the critical modification steps will be done using
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NotI digestion and religation, both the final vector and the gene of interest must not contain NotI sites. In the final mutant library, each plasmid clone will contain a unique NotI site within the 15-bp insertion, and this site will be exploited later in the genetic footprinting protocol. During the mutagenesis, plasmid clones will be plated three times and sampled twice. Taking into account the initial number of clones and Poisson distribution during sampling, the number of independently generated mutants, present in the final pool, can be estimated. We typically aim at a minimum of 10-fold insertion-per-nucleotide coverage within the target gene to guarantee a high degree of saturation mutagenesis. For example, to successfully cover a gene of 1000 bp in length, a minimum of 10,000 independent clones would be generated. 2.2. Yeast two-hybrid platform In standard yeast two-hybrid protocols, cells are initially selected on the basis of their plasmid content on glucose-containing plates, yielding well-growing yeast colonies. These colonies are subsequently replica-plated onto a two-hybrid screening medium where growth conditions induce the expression of interacting protein partners. In most systems, galactose serves as an inducer via activation of GAL1 promoters, resulting in blue colonies on X-gal indicator plates upon interaction. A simpler protocol used for the analysis presented here utilizes conditions that allow relatively good cell growth and also induce sufficient protein expression, thereby permitting color screening directly on the original transformation plates without prior replica-plating. The plating medium is SD, -Ura, -His, -Trp supplemented with 100 mM sodium phosphate (pH 7.0), 0.6% glucose, 1.4% galactose, and 200 lg/ml X-gal. This protocol yields both deep blue and clean white colonies, with interacting and non-interacting protein pairs, respectively. Colonies with intermediate color (pale blue) will be generated by protein mutants with altered but not totally destroyed binding capacity. The system generates a low-frequency background of false positive and false negative data, as indicated by the appearance of 1–2% of colonies with a switched color phenotype following the reintroduction of plasmids from ‘‘blue” or ‘‘white” pools [14]. However, this low background is tolerable, and it does not interfere with the data analysis. Following screening in yeast, different color phenotype mutants are sampled as pools. Plasmid DNA is isolated from the pools and then introduced into E. coli. At this stage, it is possible to enumerate the total number of clones in a given pool. Subsequent propagation of the pools and isolation of plasmids from E. coli ensures good quality template DNA for PCR-based genetic footprinting analysis. 2.3. Genetic footprinting Analysis of mutants as pools is arguably the most effective means to exploit the potential of an insertion mutant library, as a large number of mutants can be analyzed simultaneously. Initially the entire gene of interest is amplified using a vector-specific primer pair and DNA from the selected mutant library as a template. The amplified segment is then used as a template to amplify shorter gene segments using primer pairs consisting of a radioactively labeled and a biotinylated primer (Fig. 1D). Following pull-down with streptavidin-coated beads and subsequent NotI digestion, reaction products are analyzed by denaturing polyacrylamide gel electrophoresis and autoradiography (Fig. 2). The maximum number of clones present in each analyzed plasmid pool is not very critical. Nevertheless, a moderate number of pooled clones (e.g. 100–600) appear to be optimally suited for the analysis, as the distribution of radioactive label in these cases is restricted to fewer and thus more intensely labeled reaction products, generating easily interpretable visual data (Fig. 3).
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Fig. 2. Genetic footprinting with radioactively labeled primers. On the top is shown the overall PCR strategy. Each radioactively labeled primer (asterisk-labeled arrows specified with A, B, C, D, etc.) forms a pair with biotin-labeled oligonucleotide (Bio-labeled arrow). With several primer pairs the entire prey-expressing gene region can be analyzed. In this example, a total of 174 white (W) and 35 pale blue (PB) colonies were picked to form pools of no protein–protein interaction and weak interaction, respectively. Six independent pools of blue colonies (100 clones each, labeled 1–6) were also collected to represent strong interaction. PCR, denaturing PAGE, and autoradiography were used to analyze the pools. The wild-type prey clone (wt) served as a negative control. The size marker lanes (M) contain a representative selection of individually sequenced insertion variants [14] with a known insertion site (indicated as a nucleotide position above each band), allowing accurate mapping of insertion sites in the footprinting data. Sequencing reactions (C and G) assist in the estimation of band spacing in different parts of the autoradiograph.
3. Materials 3.1. Reagents Escherichia coli strain DH10B (Invitrogen) [31]. This strain yields high-efficiency electrocompetent cells with the protocol used [32]. It also yields excellent quality plasmid DNA for the plasmid library construction. Bacto-agar, Bacto-tryptone, Bacto-yeast extract, and Bacto-yeast nitrogen base (Difco).
Ampicillin, kanamycin, and chloramphenicol (Sigma). Plasmid pEGFP-C1 (Clontech) or similar cloning vector. Note! The selectable marker must not be the same as that present in the transposon used. MATCHMAKER LexA two-hybrid system (Clontech). Plasmid pMPH11 [14]. This is a modified version of pB42AD of MATCHMAKER. Importantly, the plasmid includes a modified kanamycin resistance cassette (XhoI site removed) that interrupts the ampicillin resistance-conferring (bla) gene. As importantly, the plasmid does not contain any NotI sites.
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Fig. 3. An example of data analysis. Location of five amino-acid insertions within the human JFC1 protein (shown schematically on the top) in relation to observed interaction with human Rab8A protein as determined by sequencing or genetic footprinting. The schematic structure of the JFC1 protein indicates the location of the SHD and C2 domains. The interaction interface is located within the SHD1 domain.
Mutation Generation System, MGSTM (Finnzymes), including the cat-Mu(NotI) transposon [25]. Plasmid DNA isolation kits (Qiagen). Qiaquick Gel Extraction kit (Qiagen). PCR Clean-up Nucleospin Extract II kit (Macherey–Nagel, Germany). Seakem LE and Seaplaque GTG agaroses (Lonza). Appropriate restriction endonucleases for cloning steps (New England Biolabs). NotI for library construction and footprinting (New England Biolabs). T4 DNA ligase, T4 polynucleotide kinase, and Vent DNA Polymerase (New England Biolabs). High quality oligonucleotide primers (see Table 1; all our primers have been purchased from the Howard Hughes Medical Institute/Keck Oligonucleotide Synthesis Faculty at Yale University). 10 mM dNTP mix (Finnzymes). DyNAzyme II DNA Polymerase (Finnzymes). Sequenase 2.0 sequencing kit (USB) for the generation of sequencing ladders as size markers. Table 1 Oligonucleotide primers.# Primer
Sequence (50 ? 30 )
Description
HSP508
CCGCCGATCCAGCCTGACTG
HSP509
GTGTCAACAACGTATCTACCAACG
Bio-HSP488
Biotin-GACAAGCCGACAACCTTGATTG
Non-radioactive PCR, pB42AD forward Non-radioactive PCR, pB42AD reverse pB42AD reverse
[c-33P]ATP (1000–3000 Ci/mmol) (GE Healthcare). Streptavidin-coated magnetic beads (Roche). Micro Bio-spin 30 columns (Bio-Rad). 40% acrylamide/bis solution 19:1 (Bio-Rad). Amberlite deionizing resin, IRN-150L (GE Healthcare).
3.2. Equipment Petri dishes (Sterilin). Large 24.5 cm 24.5 cm plates (Corning 431111). 5-mm glass beads for spreading of yeast cultures on plates (approx. 10 beads per a large plate). 0.5-mm glass beads for plasmid isolation from yeast cells. 37 °C shaker (New Brunswick Scientific). 37 °C incubator (Termaks). Centrifuges (Heraeus and Sorvall). DNA electrophoresis equipment (Bio-Rad). Electroporator (Bio-Rad Genepulser II). Thermocyclers, Robocycler Gradient40 (Stratagene) and MiniCyclerTM (MJ Research). Magnetic particle separator (Roche). Test-tube-rotator (Snijders 34528). Sequencing gel apparatus (Life Technologies). Gel dryer (Savant SpeedGel). Phosphorimager (Fuji). Whatman 1 and Whatman 3MM paper. Saran wrap.
3.3. Reagent setup
#
In addition to the primers shown, the strategy employs several gene-specific forward primers that are 50 -labeled with [c-33P]ATP prior to PCR (see Fig. 2). The interval between the binding sites of labeled primers should be approx. 200 bp. This arrangement generates overlapping data, and thus the majority of the gene regions are analyzed twice, yielding high quality data.
Luria Broth (LB) medium, 1 l. 10 g Bacto-tryptone, 5 g Bacto-yeast extract and 5 g NaCl. Adjust pH to 7.0 with 5 M NaOH. Autoclave. Store at RT for up to 1 month.
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LB agar plates. Add 15 g Bacto-agar to 1 liter of LB medium before autoclaving. Cool down to 55 °C, supplement with suitable antibiotics and dispense approx. 20 ml per Petri dish. These plates can be stored at 4 °C for 1 month. Ampicillin (Ap) solution. Dissolve ampicillin in distilled water at 100 mg/ml and pass the solution through a 0.22-lm pore filter. Can be stored at 20 °C for 1 year. Ampicillin is used at the final concentration of 100 lg/ml. Chloramphenicol (Cm) solution. Dissolve chloramphenicol in absolute ethanol at 20 mg/ml. The solution can be stored at 20 °C for 1 year. Chloramphenicol is used at the final concentration of 10 lg/ml. Kanamycin (Km) solution. Dissolve kanamycin in distilled water at 25 mg/ml and pass the solution through a 0.22-lm pore filter. The solution can be stored at 20 °C for 1 year. Kanamycin is used at the final concentration of 25 lg/ml. X-gal. Dissolve at 20 mg/ml in dimethylformamide (DMF). Store in glass or polypropylene bottles at 20 °C in the dark. P1 buffer . 50 mM Tris–HCl, pH 8.0, 10 mM EDTA, 100 lg/ml RNase A. TE buffer. 10 mM Tris–HCl, pH 7.5, 0.5 mM EDTA. TEN buffer. 10 mM Tris–HCl, pH 7.5, 0.5 mM EDTA, 50 mM NaCl. TEN100 buffer . 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 100 mM NaCl. TEN1000 buffer. 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 1 M NaCl. 2 binding buffer . 10 mM Tris–HCl, pH 7.5, 2 mM EDTA, 200 mM NaCl. 2 formamide loading dye . 95% deionized formamide, 10 mM EDTA, 0.1% bromo-phenol blue, 0.1% xylene cyanol. 7 M urea, 6% polyacrylamide solution, 200 ml. Dissolve 84 g urea and 30 ml 40% acrylamide in 50 ml distilled water at 37 °C bath, deionize with Amberlite resin (5 g) for 1–2 h, and pass the solution through two Whatman 1 papers. Add 20 ml 10 TBE and adjust volume to 200 ml with distilled water. Store at 4 °C for up to 1 month.
4. Methodology 4.1. Plasmid constructions (1) Standard DNA techniques are performed as previously described [33]. (2) Insert the wild-type prey gene into any appropriate cloning vector to generate an initial target plasmid for pentapeptide mutagenesis. Because a gel isolation procedure will be used during the library construction, choose a vector plasmid whose size is different than that of the total length of the prey gene plus transposon (cat-Mu(NotI) transposon is 1254 bp in length). In addition, the cloning sites in the vector need to be compatible with those of pMPH11 (EcoRI, NheI, StuI, NcoI, SfiI, XmaI, BglII, and XhoI), the final expression plasmid for the two-hybrid analysis. Plasmid pMPH11 [14] is a modified version of pB42AD of the MATCHMAKER kit, and it is available upon request. The modifications (elimination of a critical NotI site and an insertion of the npt gene) have been described in detail previously [14]. The prey gene must not contain any NotI sites, as this restriction endonuclease is used to remove the transposon bulk during the library construction. If it does, remove the site(s) using any of the currently available site-directed mutagenesis protocols. (3) Clone the wild-type prey gene into pMPH11 using the same cloning strategy that you chose to use for the mutant library. This construct will be used as a positive control in the yeast two-hybrid analysis and as a negative control in genetic footprinting. (4) Clone the bait gene into pGildaB provided by the MATCHMAKER kit.
4.2. Generation of the pentapeptide insertion mutant library (5) The pentapeptide insertion mutant library is generated using the Mutation Generation SystemTM (Finnzymes) essentially as specified by the supplier. This mutagenesis system exploits the MuA transposase-catalyzed in vitro transposition reaction [18] and generates 5-aa insertions in proteins [21,25]. (6) Prepare five standard in vitro transposition reactions using cat-Mu(NotI) [25] as a donor DNA. Note! MuA transposase should be added last. Component Target plasmid (100 ng/ll) M1-CamR transposon (100 ng/ll) 5 reaction buffer for MuA transposase Nuclease-free water MuA transposase (220 ng/ll)
Amount (ll) 3 1 4 11 1
Final 300 ng 100 ng 1 Final volume 20 ll 220 ng
(7) Incubate at 30 °C for 3 h, and then inactivate MuA transposase by incubating at 75 °C for 10 min. (8) Pool the reactions and extract with phenol and subsequently with chloroform. Ethanol precipitate the DNA and resuspend it in water (25 ll). (9) Electroporate several aliquots (2 ll) of DNA into DH10B electrocompetent cells (50 ll) prepared as previously described [32]. (10) Select transposon-containing plasmid clones by plating the bacteria on LB agar plates supplemented with chloramphenicol (Cm, 10 lg/ml) and kanamycin (Km, 25 lg/ml). To avoid background growth, do not plate more than approx. 1000 CFU per plate. (11) Pool an appropriate number of clones by scraping transformant colonies from the plates and grow them in LB–Cm–Km medium at 37 °C for 3 h. In general, we use the length of the gene of interest (in base pairs) and multiply that by 10 to form an estimate for a minimum final pool size. For example: if the gene of interest is 1700 bp, we would need at least 17,000 independently generated clones in the final pool (see step 25 for considerations about sampling). (12) Isolate plasmid DNA from the pool and digest the DNA with appropriate restriction enzymes to release the DNA of interest (the target gene hit by the transposon) from the cloning vector. (13) Separate the resulting fragments by preparative electrophoresis using 0.8% Seaplaque GTG agarose gel in TAE buffer [33]. (14) Isolate from the gel the DNA fragment (pool) that corresponds to transposon insertions into the PREY-encoding DNA segment. We have used a standard electroelution protocol for the purpose, but commercial gel isolation kits may substitute. (15) Ligate the isolated DNA fragment into pMPH11 digested with the same enzymes that were used in step 12. (16) Electroporate the ligation mixture into DH10B electrocompetent cells (50 ll) as specified in step 9. (17) Select transposon-containing plasmid clones by plating the bacteria on LB–Cm–Km plates as specified in step 10. (18) Pool and grow the transformants as specified in step 11. (19) Prepare plasmid DNA from the pool. (20) To eliminate the transposon core sequence from the plasmid pool, digest the plasmid pool with NotI and separate the linearized fragment using preparative electrophoresis on a 1.7% Seaplaque GTG agarose gel.
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(21) Isolate the plasmid backbone as specified in step 14 and recircularize it by ligation at low DNA concentration (approx. 1 ng/ll). (22) Electroporate ligated plasmids into DH10B cells as specified in step 9. (23) Select the transformants on LB–Km plates to generate the final library. Note! Chloramphenicol selection is not used at this stage. (24) Pool and grow transformants in LB–Km medium as specified in step 11. (25) Prepare plasmid DNA from the pool. The insertion library is now ready for your specific application. Note that during the library construction, the original plasmid pool is sampled several times. As the number of clone members in each sampling is known, Poisson distribution can be used to estimate the number of independently generated mutants in the final library. 4.3. Yeast two-hybrid system (26) The yeast two-hybrid protein interaction screen is performed using the MATCHMAKER LexA Two-Hybrid System (Clontech) according to the manufacturer’s specifications. The standard yeast media used are described in the Yeast Protocols Handbook (Clontech). (27) Transform pGildaB-BAIT plasmid into S. cerevisiae reporter strain EGY48 (MATa, his3, trp1, ura3, LexAop(x6)-LEU2) harboring the plasmid p8op-lacZ. Plate on SD-Ura-His agar. (28) Introduce the final pMPH11-PREY insertion mutant library into the above reporter strain harboring the above two plasmids. Plate on SD, -Ura, -His, -Trp agar supplemented with 100 mM sodium phosphate (pH 7.0), 0.6% glucose, 1.4% galactose, 0.2 mg/ml X-gal. We have used large (24.5 cm 24.5 cm) plates. (29) Streak the colonies on Petri dishes (medium from step 28) and grow them to verify their color phenotype. Individual insertion mutants can be isolated and sequenced at this point. These mutants can be used as convenient size markers in the footprinting step. (30) Pool colonies on the basis of their color phenotype and grow them in SD, -Ura, -His, -Trp medium at 30 °C for 3 h. We prefer using pool sizes less than 200 member clones, and recommend generating several ‘‘blue” pools in parallel if necessary. A pool size with about 600 member clones approaches current detection limit [14]. (31) Prepare plasmid DNA from the pools using the Qiagen Plasmid Spin mini kit. Vortex the cells in P1 buffer with glass beads for 10–30 min prior to DNA isolation. (32) Introduce the mutant plasmid pools into E. coli DH10B by electroporation as specified in step 9 using LB–Km plates for selection. Compared to the original pool size, at least 10-fold excess colonies should be obtained after electroporation. For example, a pool of 100 member clones in yeast should be plated as approx. 1000 colonies (or more) in E. coli. (33) Scrape the transformant colonies from the plates and prepare plasmid DNA from each pool. DNA preparations are now ready for PCR-based genetic footprinting. 4.4. PCR (34) Prepare the non-radioactive PCR in a total volume of 50 ll. Component Plasmid DNA (50 ng/ll) 10 mM dNTP mix Primer HSP508 (5 pmol/ll) Primer HSP509 (5 pmol/ll)
Amount (ll) 1 1 5 5
Final 50 ng 200 lM 0.25 lM 0.25 lM
100 mM MgSO4 (contributes 2 mM) 10 ThermoPol reaction buffer Vent DNA polymerase (2 U/ll) Nuclease-free water
1
4 mM
5 0.5 31.5
1 1U Final volume 50 ll
(35) Amplify in the thermocycler (Robocycler) using the following parameters: 5 min at 95 °C followed by 25 cycles of 1 min at 95 °C, 1 min at 60 °C, and 2 min at 72 °C, and finally 5 min at 72 °C. (36) Purify PCR products electrophoretically using the Qiaquick Gel Extraction kit. (37) Label the prey-specific primers radioactively; 50 -labeling with [c-33P]ATP and T4 polynucleotide kinase is performed as previously described [33]. (38) Prepare the radioactive label-containing PCR in a total volume of 50 ll. Using a PCR protocol with one biotinylated (nested) primer increases specificity and decreases background radioactivity in the final gel analysis. Component Non-radioactive PCR product (50 ng/ll) 10 mM dNTP mix Primer Bio-HSP488 (5 pmol/ll) Labeled prey-specific primer (5 pmol/ll) 10 Optimized DyNAzyme Buffer DyNAzyme II DNA polymerase (2 U/ll) Nuclease-free water
Amount (ll)
Final
1
50 ng
1 5 5
200 lM 0.25 lM 0.25 lM
5 0.5
1 1U
32.5
Final volume 50 ll
(39) Amplify in the thermocycler (MiniCycler) using otherwise same parameters as specified in step 35 except that the extension time at 72 °C can be variable (typically from 45 s to 120 s) depending on the length of the desired PCR product. (40) Purify the radioactively labeled PCR product using the PCR Clean-up Nucleospin Extract II kit and elute into 30 ll of 10 mM Tris–HCl, pH 8.5. 4.5. Genetic footprinting (41) Pre-wash streptavidin beads four times with TEN100 and once with 2 binding buffer. Mix the beads with 2 binding buffer using the ratio of 13 volume beads and 17 volume buffer. Use this suspension in step 42. We have done all the washing steps using a magnetic particle separator. Alternatively, washing steps can be done by centrifugation. (42) Add 30 ll of radioactively labeled PCR product to the prewashed streptavidin bead suspension (30 ll) and incubate at room temperature for at least 1 h to allow adsorption. To avoid sedimenting of the beads, use a test-tube-rotator and/or mix occasionally. (43) Wash the beads three times with 0.5 ml of TEN1000 and twice with 1 restriction enzyme buffer 3 (New England Biolabs). (44) Add 50 ll of 1 restriction enzyme buffer 3 containing NotI (25 U) and incubate at least for 4 h at 37 °C. Mix occasionally. (45) Remove the beads using the magnetic particle separator and subsequently purify the supernatant by centrifugation through a Micro Bio-spin 30 column (equilibrated with TEN buffer) at room temperature in a tabletop microcentrifuge (3000 rpm, 4 min).
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(46) Ethanol precipitate the DNA and resuspend it in 3 ll of TE buffer. Add 3 ll of 2 formamide loading dye for gel analysis. (47) Analyze samples by 7 M urea, 6% polyacrylamide gel electrophoresis as previously described [34]. Appropriate sequencing reactions are used as size markers in the analysis. Individually sequenced insertion variants (see step 29), each with a known insertion site, can be used to accurately calibrate the data. (48) Dry gel at 80 °C onto Whatman 3MM paper. (49) Visualize the bands by autoradiography. We have used a Fuji BAS 1500 phosphorimager with BAS-Reader 2.9 software (Raytest). (50) Analyze the data to localize the interacting interface.
[4] [5] [6] [7] [8] [9] [10] [11] [12]
5. Anticipated results
[13] [14]
Functional analysis of a protein interface involved in an interaction with another protein can be accomplished by subjecting the selected protein mutant pools to comparative parallel analysis using a PCR-based strategy. Following gel analysis and autoradiography, this type of genetic footprinting assay generates a visual read-out where reciprocal band patterns can be seen between the insertion mutant pools representing unaltered versus altered protein–protein interaction. In yeast two-hybrid screen, the percentage of white colonies should be indicative of the relative proportion of the interacting interface in the protein. For preliminary data, mutant clones can be analyzed individually by sequencing.
[15]
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