A system based on specific protein–RNA interactions for analysis of target protein–protein interactions in vitro: successful selection of membrane-bound Bak–Bcl-xL proteins in vitro

Shinya Y. Sawata1, Eigo Suyama1,2 and Kazunari Taira1,2,3

1Gene Function Research Center, National Institute of Advanced Industrial Science and Technology (AIST), Central 4, 1–1–1 Higashi, Tsukuba Science City 305-8562 and 2Department of Chemistry and Biotechnology, School of Engineering, The University of Tokyo, 7–3–1 Hongo, Tokyo 113-8656, Japan

3 To whom correspondence should be addressed, at the Tokyo address. E-mail: taira{at}chembio.t.u-tokyo.ac.jp


    Abstract
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 Abstract
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 Materials and methods
 Results and discussion
 References
 
Ribosome display systems are very effective and powerful tools for in vitro screening of transcribed mRNAs that encode proteins (or peptides) with specific (known or unknown) functions. We have modified such a system by exploiting the interaction between a tandemly fused MS2 coat-protein (MSp) dimer and the RNA sequence of the corresponding specific binding motif, C-variant (or Cv). We placed the MSp dimer at the N-terminus of a nascent protein and the Cv binding motif was attached to the 5' end of the protein's mRNA. This configuration enhanced the stability of the ribosome–mRNA complex. We demonstrate here that this improved ribosome display system provides an effective method for identifying the gene for a protein that binds to a protein of interest. We visualized the formation of polysome complexes in this advanced polysome display by atomic force microscopy (AFM) and found that the AFM images of polysomes in our system were different from those observed in the case of conventional ribosome display systems. Our results suggest that our technology might usefully complement yeast two-hybrid assays.

Keywords: atomic force microscopy/in vitro selection/MS2 coat protein/ribosome display/RNA–protein interaction


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
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Many biochemical techniques have been used to detect interactions between proteins. Phage display (Smith, 1985Go) and yeast two-hybrid assays (Fields and Song, 1989Go; Chien et al., 1991Go; Bartel and Fields, 1995Go) are commonly used for the identification of the gene for a protein that binds to a protein of interest. However, these assays depend on living cells and their properties and therefore they have some inherent disadvantages, namely (i) limitations with respect to the variety of the genotype library (<107–109), (ii) the waiting period that is required for the culture of host cells and (iii) the impossibility of assays for cell-toxic proteins.

The ribosome display method was established to overcome the problems of phage display and two-hybrid assays (Mattheakis et al., 1994Go; Hanes and Plückthun, 1997Go; He and Taussig, 1997Go; Gersuk et al., 1997Go) and we recently reported a modified ribosome display method (Figure 1) that incorporates an additional protein–RNA interaction (Sawata and Taira, 2003Go), namely, stabilization of the mRNA–ribosome–nascent protein complex using the specific and stable interaction between an RNA-binding protein (mutant coat protein of bacteriophage MS2; mMSp) and an RNA motif (Cv) which binds to the MS2 protein (LeCuyer et al., 1995Go; Peabody, 1997Go; Valegård et al., 1997Go; Convery et al., 1998Go; Johansson et al., 1998Go; Rowsell et al., 1998Go). In this paper, we describe the establishment of a system that utilizes a ‘GST-pulldown’ step and our modified ribosome display system and we demonstrate that our novel system works not only as a system for the detection of protein–protein interactions, but also as a screening system that allows us to identify the gene for a binding partner of a protein of interest without the use of living cells.



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Fig. 1. The predicted scheme of interactions between glutathione–agarose beads, the GST–Bak fusion protein and the mRNA–ribosome–nascent protein complex that included the Cv–mMSp interaction.

 
We tested our novel system for the detection of protein–protein interactions by taking advantage of the established interactions between human Bak and human Bcl-xL proteins (Hammond et al., 2001Go). Bak is a pro-apoptotic member of the Bcl-2 family (Chittenden et al., 1995Go; Farrow et al., 1995Go; Kiefer et al., 1995Go; Suyama et al., 2002Go) and it forms a heterodimeric complex with Bcl-xL (Farrow et al., 1995Go; Sattler et al., 1997Go), which is an anti-apoptotic member of the Bcl-2 family (Boise et al., 1993Go). The dissociation constant (Kd) of the complex between the Bcl-xL and a synthetic peptide that represents the BH3 domain of Bak was estimated by Sattler et al. (1997)Go to be 0.20 µM. We also used the precursor to the human caspase 3 as a negative control in the assay of protein–protein interactions. The precursor to caspase 3 is a pre-activated form of caspase 3, a member of the family of cysteine proteases, that is involved in apoptosis (Thornberry and Lazebnik, 1998Go).

We prepared three mRNAs to investigate whether our system would work as designed: an mRNA encoding the GST–Bak fusion protein for use as ‘bait’; an mRNA encoding Bcl-xL as a selection-positive ‘prey’; and an mRNA encoding the precursor to caspase 3 as a selection-negative ‘false prey’. We prepared these mRNAs by RT–PCR amplification, cloning and transcription. An equimolar mixture of the mRNAs for Bcl-xL and precursor to caspase 3 was translated in a cell-free translation system in vitro.

Since the translated mRNAs lacked a termination codon, they were unlikely to be released efficiently from ribosomes by the release factor (Mattheakis et al., 1994Go; Gersuk et al., 1997Go; Hanes and Plückthun, 1997Go; He and Taussig, 1997Go). In an attempt to improve the efficiency of selection of our system, we introduced an additional interaction between the mRNA and its translated product, using a mutant from the MS2 coat protein (mMSp) and the Cv RNA motif to which mMSp binds with high affinity (LeCuyer et al., 1995Go; Valegård et al., 1997Go; Convery et al., 1998Go; Johansson et al., 1998Go; Rowsell et al., 1998Go; Sawata and Taira, 2003Go). We used a mutant dimeric form of MSp, mMSp, in order to avoid capsid formation by MSp, without any reduction in its affinity for the RNA motif (LeCuyer et al., 1995Go; Peabody, 1997Go). We designed a cDNA that placed mMSp between Bcl-xL (or the precursor to caspase 3) and a linker protein and we placed Cv at the 5' end of the mRNA. This configuration was expected to result in binding of nascent mMSp to the Cv motif, thereby increasing the stability of the mRNA–ribosome–protein complex, as depicted in Figure 1 (Sawata and Taira, 2003Go). In such a system, in which the nascent polypeptide binds to an RNA motif and the ratio of ribosomes to mRNA is >10 (Promega Corporation, personal communication), we predicted that the mRNA–ribosome–nascent protein complex would exist in the form of polysomes. Moreover, we found evidence for the presence of such polysomes by atomic force microscopy (AFM).

Polysome or ribosome display is potentially suitable for investigations of protein–protein interactions that include a membrane protein since the solubility of the protein of interest in an aqueous solution depends not on features of the protein itself but on the entire mRNA–ribosome–protein complex, which is relatively water soluble. We demonstrate here that our advanced ribosome display system can be used to identify protein–protein interactions that include a hydrophobic membrane protein, overcoming the limitations of some in vitro protein-selection systems that are due to limited solubility of the protein of interest. From the effectiveness of our system, as shown here, it seems likely that our system should be a useful complement to the widely used yeast two-hybrid interaction assay and should contribute to efforts aimed at characterizing networks of functional proteins in living cells.


    Materials and methods
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 Abstract
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 Materials and methods
 Results and discussion
 References
 
Preparation of a mutant gene for MSp (mMSp)

A lysate of Saccharomyces cerevisiae L40 ura MS2 strain, obtained from Invitrogen (Carlsbad, CA), included the gene for MS2p. We used this lysate as the source of template DNA for PCR. We subcloned the amplified gene for MSp and used the resultant genome as template for mutagenesis by PCR, which allowed us to introduce the desired mutations into MSp, namely, mutation of valine 75 to glutamic acid, alanine 81 to glycine and arginine 82 to tryptophan. These mutations yielded MSp that was not incorporated into capsids by self-assembly, but they did not eliminate the RNA-binding ability of the protein (LeCuyer et al., 1995Go).

Preparation of the Cv motif

The Cv motif is an RNA motif that binds to MSp and mutant MSp (mMSp) (LeCuyer et al., 1995Go; Valegård et al., 1997Go; Convery et al., 1998Go; Johansson et al., 1998Go; Rowsell et al., 1998Go). We prepared this motif as double-stranded DNA from two chemically synthesized DNA oligomers (sense sequence, 5'-TCG AGA CAT GAG GAT CAC CCA TGT G-3'; and antisense sequence, 5'-TCG ACA CAT GGG TGA TCC TCA TGT C-3'). To clone the Cv motif into the vector, we included SalI sites upstream and downstream, respectively, of the artificial double-stranded DNA.

Construction of plasmids

The original plasmid was prepared by cloning the gene for dihydrofolate reductase (dhfr) from Escherichia coli into the multicloning site of pET30-a(+) (Novagen, Madison, WI,), to yield the plasmid designated pD. This plasmid had EcoRI and SalI sites between the T7 promoter and the Shine–Dalgarno (SD) sequences. Therefore, we were able to insert the Cv motif between the sites of initiation of transcription and translation by cloning the DNA fragment that contained the Cv motif between the EcoRI and SalI sites of this vector. The term Cv in the names of plasmids indicates presence of a Cv motif. We then cloned the gene for mMSp at the PstI site in the plasmid to generate pCvmM2D. The inclusion of mM2 refers to the presence of two sets of gene encoding mMSp in tandem.

We cloned two cDNAs from a lysate of HeLa cells for this study, namely the cDNAs for human Bcl-xL and the human gene for the precursor to caspase 3 by RT–PCR. The reverse primers for RT–PCR had been designed such that they contained no termination codon in the correct frame and, therefore, the cDNAs did not include a termination codon in their respective open reading frames (ORFs). Insertion of these cDNAs into plasmids generated pCvBlmM2D, pCvCamM2D, pBlD and pCaD. The names of plasmids containing Bcl-xL cDNA include Bl and those of plasmids containing the cDNA for the precursor to caspase 3 include Ca. We used the same method to clone the human Bak gene, with the exception that the reverse primer was designed to include the native termination codon of the Bak gene. The plasmid created by insertion of Bak cDNA into the NspV site of pD was designated pBk. Finally, we prepared a gene for the glutathione S-transferase (GST) for construction of the template DNA of the mRNA that encoded the ‘bait’ protein. The gene for GST was prepared in the same way as mentioned for the subcloning of the Bcl-xL gene and the GST cDNA was inserted into the NdeI site of pBk to yield pGBk.

Preparation of template DNA for transcription in vitro

We employed two pairs of DNA oligomers as primers for PCR in order to prepare template DNA for transcription in vitro. The upstream primer in the first pair was designated fP-1 and the downstream primer was designed rP-1. The sequence of fP-1 was 5'-GCG TAG AGG ATC GAG ATC GA-3' and that of rP-1 was 5'-CCG GAT ATA GTT CCT CCT TTC-3'. When the plasmids described above were employed as templates for PCR with this pair of primers, the products of PCR contained a T7 promoter in the upstream region and a termination codon downstream of the longest ORF. The first pair of primers and pGBk were used for preparation of dGBk(+) [see Figure 2 (VII)]. The second pair of primers included fP-1 and rP-2, whose sequence was 5'-GTT ATT GCT CAG CGG TGG CA-3'.



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Fig. 2. The constructs prepared as templates for transcription in vitro. We designed the following template DNAs: dCvBlmM2D(–) (I); dCvCamM2D(–) (II); dBlmM2D(–) (III); dCamM2D(–) (IV); dBlD(–) (V); dCaD(–) (VI); and dGBk(+) (VII). The template DNAs (I)–(VI) did not have a termination codon, whereas template DNA (VII) had a termination codon. The expected lengths of constructs were 2358 bp (I), 2490 bp (II), 2255 bp (III), 2387 bp (IV), 1487 bp (V), 1619 bp (VI) and 1517 bp (VII). An ‘r’ in the name of a construct indicates an mRNA prepared for translation in vitro. Inclusion of ‘(–)’ in the name of a construct means that there was no termination codon within the longest ORF. Inclusion of ‘(+)’ in the name of a construct means that the final codon was a termination codon in the longest ORF. The number on the right of each construct indicates the expected number of amino acids in each respective product of translation.

 
After PCR with this second pair of upstream and downstream primers and the plasmids mentioned above as templates, the products of PCR each contained a T7 promoter in the upstream region but no termination codon within the longest ORF. The second pair of primers and plasmids pCvBlmM2D, pCvCamM2D, pBlD and pCaD were used for the preparation of dCvBlmM2D(–), dCvCamM2D(–), dBlD(–) and dCaD(–), respectively [see Figure 2 (I), (II), (V) and (VI)]. The third pair of primers included rP-2 and fP-2, whose sequence was 5'-TAA TAC GAC TCA CTA ATA GGG GTC GAC AAT AAT TTT GTT TAA CTT-3'. After PCR with this third pair of upstream and downstream primers and the plasmids mentioned above as templates, the products of PCR each contained a T7 promoter in the upstream region but no Cv sequence and no termination codon. The third pair of primers and the plasmids pCvBlmM2D and pCvCamM2D were used for preparation of dBlmM2D(–) and dCamM2D(–), respectively [see Figure 2 (III) and (IV)]. For PCR, the final volume of each reaction mixture, which contained Ex Taq (Takara Shuzo, Kyoto, Japan), was 200 µl. The conditions for PCR were as follows in all cases: denaturation at 94°C for 1 min, annealing at 55°C for 1 min and elongation at 74°C for 1 min. The cycle was repeated 30 times. The products of PCR were purified with a QIAquick PCR purification kit (Qiagen, Hilden, Germany) and concentrations of purified products of PCR were estimated from the absorption at 260 nm of each solution.

Each template DNA was named according to the scheme described above for naming plasmids and a schematic representation of each is shown in Figure 2. Inclusion of ‘d’ in the name of a construct means that it was a product of PCR that had been prepared as template DNA. Inclusion of ‘(–)’ in the name of a construct means that there was no termination codon within the longest ORF in the product of PCR. By contrast, inclusion of ‘(+)’ in the name of a construct means that the final codon was a termination codon in the longest ORF.

Preparation of mRNA for translation and selection in vitro

Transcription in vitro was performed with a RiboMAX Large Scale RNA Production System (Promega, Madison, WI, USA) according to the general protocol supplied with the system. The concentration of template DNA was 1 µg per 50 µl of reaction mixture. The final concentration of the cap analog was adjusted to 0.4 mM. After transcription, deoxyribonuclease I (Takara) was added to the reaction mixture, which then was incubated at 37°C for 1 h. The transcribed RNA was purified with an RNeasy Mini kit (Qiagen) and the concentration of purified RNA was estimated from the absorption at 260 nm of the solution. Then, 0.5 µl of the solution of purified RNA was added to 2 µl of formamide that contained 20 mM EDTA and 40 mg/ml blue dextran. This mixture was heated at 96°C for 2 min and then put on ice for 1 min. After the addition of 0.5 µl of loading buffer [30% (v/v) glycerol, 0.01% (w/v) bromophenol blue and 0.01% (w/v) xylene cyanol FF (Sigma, St Louis, MO)], the sample of denatured RNA was loaded on a 2.0% (w/v) non-denaturing agarose gel prepared with SeaKem GTG agarose (BioWhittaker Molecular Applications, Rockland, ME). Each mRNA was named according to the rules for naming plasmids (see above). An ‘r’ in the name of a construct indicates an mRNA prepared for transcription in vitro. Inclusion of ‘(–)’ in the name of a construct means that there was no termination codon within the longest ORF. By contrast, inclusion of ‘(+)’ in the name of a construct means that the final codon was a termination codon in the longest ORF. For internal labeling with radioactive phosphate, transcription in vitro was performed as described above, with the addition of 1.85 MBq of [{alpha}-32P]CTP (NEN Life Science Products, Boston, MA) to the reaction mixture for transcription in vitro.

Translation in vitro

We used an S30 extract of E.coli (Promega) for translation in vitro of linear templates in the present study. We estimated the approximate molar concentration of each solution of mRNA and we adjusted the working concentration of input mRNA to 0.1 µM when the mRNA was not radiolabeled. Transcripts were translated according to the general protocol from Promega. First we prepared a mixture of 5 µl of a complete amino acid mixture [including 1 mM each amino acid (Promega)], 0.5 µl of a solution of 10 mg/ml rifampicin (Sigma), 20 µl of the enzyme premix without amino acids that was included in the translation kit, 15 µl of the S30 extract from E.coli included in the translation kit and appropriate amounts of the solution of mRNA- and RNase-free water. This mixture was kept on ice until the start of the translation reaction, which was initiated by floating the sample tube in a water-bath that had been adjusted to 37°C. Translation of the mRNA that encoded a prey protein was allowed to proceed at 37°C for 10 min as suggested by protocols in the literature (Hanes and Plückthun, 1997Go). By contrast, independent translation of the mRNA that encoded a bait protein was allowed to proceed at 37°C for 30 min.

Preparation of buffers for selection in vitro

The binding buffer was prepared by mixing solutions as follows: 420 µl of distilled water, 25 µl of 1 M K2HPO4 (pH 9.4), 50 µl of 0.5 M KH2PO4 (pH 5.6), 300 µl of 1 M NaCl, 5 µl of 10% (v/v) polyoxyethylene(20) sorbitan monolaurate (Wako Pure Chemical Industries, Osaka, Japan), 100 µl of 20% (w/v) Block Ace solution (Dainippon Pharmaceutical, Osaka, Japan) and 100 µl of 0.5 M Mg(CH3COO)2. The selection buffer was a mixture of 430 µl of binding buffer and 20 µl of a suspension of glutathione Sepharose 4B beads (Amersham Biosciences, Uppsala, Sweden). The elution buffer was prepared by mixing solutions as follows: 418 µl of distilled water, 25 µl of 1 M K2HPO4 (pH 9.4), 50 µl of 0.5 M KH2PO4 (pH 5.6), 300 µl of 1 M NaCl, 5 µl of 10% (v/v) polyoxyethylene(20) sorbitan monolaurate, 100 µl of 20% (w/v) Block Ace solution, 2 µl of 0.5 M EDTA (pH 8.0) and 100 µl of 0.1 M glutathione.

Selection in vitro

We first prepared equimolar mixtures as models of genotype libraries of prey proteins, namely a mixture of rCvBlmM2D(–) and rCvCamM2D(–), a mixture of rBlmM2D(–) and rCamM2D(–) and a mixture of rBlD(–) and rCaD(–). We also prepared rGBk(+) as an mRNA that encoded a bait protein (i.e. GST–Bak fusion protein). These mRNAs were translated in vitro by the protocol described above. We prepared two microtubes that contained 460 µl of binding buffer and 20 µl of glutathione–agarose beads. The translation in vitro of rGBk(+) was performed in 50 µl and two 20 µl aliquots of the sample after translation were added to respective aliquots of the mixture of binding buffer and glutathione–agarose beads mentioned above. The two resultant mixtures were incubated at room temperature for 10 min with constant gentle mixing by inversion of the sample tubes. This step was designed to allow the binding of the GST tag of the bait protein to glutathione that had been covalently immobilized on agarose beads. This binding step was followed by a washing step.

First, the sample was centrifuged at 2000 g for 1 s and the supernatant was carefully removed. Then, 200 µl of binding buffer were added to the glutathione–agarose beads, which had been pelleted by centrifugation, with gentle mixing. Centrifugation was repeated and the supernatant was removed. While the bait protein was immobilized, the translated products of the mixture of rCvBlmM2D(–) and rCvCamM2D(–), the mixture of rCvBlmM2D(–) and rCvCamM2D(–) and the mixture of rBlD(–) and rCaD(–) were added to 450 µl of binding buffer. Each of these mixtures was then added to the glutathione–agarose beads on which the GST–Bak fusion protein had been immobilized according to the protocol mentioned above. These samples were incubated at room temperature with constant gentle mixing by inversion of the sample tubes. The samples were centrifuged at 2000 g for 1 s and then the supernatants were carefully removed. Then, 200 µl of binding buffer were added and beads and buffer were mixed gently. This washing step was repeated three times in all.

After the third removal of the supernatant, 200 µl of the elution buffer were added to the pelleted glutathione–agarose beads instead of the binding buffer. The mixture was incubated at room temperature for 10 min with repeated gentle inversion of the sample tube, as described above. The glutathione–agarose beads were then pelleted by centrifugation at 2000 g for 1 s and the supernatant, containing the eluted mRNA of interest, was carefully removed to a new microtube. The mRNA in this solution was purified with an RNeasy Mini kit.

The purified mRNA was reverse-transcribed by M-MLV reverse transcriptase (Promega) to yield complementary single-stranded DNA. We designed a single-stranded DNA oligomer as the primer for reverse transcription (RT), which we designed rP-3; its sequence was 5'-CTT GTC GTC GTC GTC GGTA-3'. A mixture of 10 µl of the solution of purified mRNA and 4 µl of 1 µM rP-3 was incubated at 70°C for 5 min and then placed on ice for 1 min. The sample containing mRNA and the annealed RT primer was mixed with 1 µl of PCR Nucleotide Mix (Promega), namely, a premixed solution of 10 mM each dNTP in water and 4 µl of optimized buffer for RT (Promega). Then, 1 µl of the solution of M-MLV reverse transcriptase was added to the sample, which was kept on ice until the reaction was started by floating the tube in a water-bath adjusted to 42°C. The RT reaction was allowed to proceed for 2 h and was followed by PCR. The volume of the reverse-transcribed sample that was subsequently used as the template for PCR was 2 µl. We used the three DNA oligomers as primers for PCR, that is, the forward primer was fP-4 (5'-GGA TAA CAA TTC CCC TCG AGA-3') or fP-5 (5'-GTC GAC AAT AAT TTT GTT TAA CTT-3') and the reverse primer was rP-3 (see above). We performed 15 cycles of PCR when the reverse-transcribed product originated from the initial pool and 23 cycles when the reverse-transcribed product originated from the pool of selected mRNA.

We analyzed the products of RT–PCR by electrophoresis on a 2% (w/v) SeaKem GTG agarose gel. The standard DNA markers used for the identification of products of RT–PCR were 100 Base-Pair Ladders (Amersham Biosciences). The bands of products of RT–PCR on agarose gels were stained with SYBR Gold (Molecular Probes, Eugene, OR). The patterns of the bands after electrophoresis were analyzed quantitatively using Fluor Imager 595 and ImageQuaNT software (Molecular Dynamics, Sunnyvale, CA).

Comparative analysis of the efficiency of the selection in vitro on the basis of the radioactivity of radioisotope-labeled mRNA

To estimate efficiency of our selection system, we used internally radiolabeled mRNA and followed the selection protocol, as described above, but without eluting the selection-positive mRNA. In other words, we measured the radioactivity associated with the pelleted glutathione–agarose beads before elution of mRNA. We confirmed first that the background radioactivity of the pelleted glutathione–agarose beads was close to zero. Hence the radioactivity that remained associated with the glutathione–agarose beads after the screening reflected the amount of mRNA recovered during the screening. This assay allowed us to estimate the efficiency of our system. For this comparative analysis, we attempted to eliminate artifacts due to stoichiometric factors as follows. We chose one mRNA as the standard for the normalization. We then estimated the cytidine content per molecule of each mRNA and calculated the theoretical ratios of cytidine content relative to that of the mRNA chosen as the standard. Then we adjusted the relative radioactivity (c.p.m.) of each input mRNA by reference to the theoretical relative cytidine content, estimated as indicated above. Such normalization allowed us to perform both translation and selection in vitro under conditions wherein the number of input mRNA molecules was identical for each sample, even when the respective sequences and lengths of the mRNAs to be subjected to comparison were significantly different.

AFM analysis of the polysome complex

The preparation of the products of translation in vitro was performed in the same way as the preparation of prey protein that we described above. The amount of each input mRNA was 5 pmol and the working volume of mixture for translation in vitro was 50 µl. Purified mRNA, prepared as the negative control for polysome formation, was prepared by dilution of the original mRNA that was used for the assay of selection in vitro. The amount of each purified mRNA was 50 pmol and the volume of the prepared sample was 50 µl. The samples translated in vitro were diluted 106-fold with PBS buffer (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4) that contained 10 mM MgCl2 and purified mRNA was diluted 103-fold with 10-fold diluted PBS buffer that contained 10 mM MgCl2. The volume of the sample deposited on a freshly cleaved mica surface for AFM was 40 µl. After a 5 min incubation at room temperature, the sample was rinsed with 5 ml of Milli-Q-purified water (Millipore, Bedford, MA) and blown dry with dry nitrogen. The sample was then dried on a vacuum dryer for 15 min at 0.05 MPa and 40°C. The analysis by AFM was performed with a NanoScope IIIa (Digital Instruments, Santa Barbara, CA) in tapping mode at room temperature in air, using silicon cantilevers 119 µm in length with a spring constant of 55 N/m. Typical resonance frequencies of these cantilevers were ~379 kHz. An area of 1000x1000 nm was scanned at <1.5 Hz and then captured images were flattened. AFM was performed at the Research Institute of Biomolecule Metrology (RIBM; http://www.ribm.co.jp/index.htm).

Detailed AFM investigations of ribosomes from E.coli were reported by Vanzi et al. (2003)Go. They estimated that the density of immobilized ribosomes was 0.02 pmol (1.2 x 1010 molecules) of immobilized ribosomes/cm2, which was the average of results derived from 21 repeated measurements, when the concentration of the ribosomes in the sample solution that had been deposited on the mica surface was 10 nM. Thus, one ribosome molecule was observed when an area of 100 x100 nm on the surface of the mica was scanned. In the present experiments, we estimated that the concentration of ribosomes deposited on the mica surface was 1.9 pM; therefore, we expected that two molecules of ribosomes would be observed when an area of 10 000 x 10 000 nm on the surface of the mica was scanned. We scanned an area of 1000 x 1000 nm on the surface of the mica. From the estimations mentioned above, it is reasonable to assume that all of the AFM images shown in Figure 5A and B are meaningful.



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Fig. 5. Representative AFM images of mRNA being translated. (A) The mRNA being translated in vitro was rCvBlmM2D(–), which included the Cv motif and a tandem dimer of the mRNA for mMSp but no termination codon in the longest ORF. (B) The mRNA being translated in vitro was rBlD(–), which did not include a Cv motif, the mRNA for mMSp or a termination codon in the longest ORF. This mRNA was the ancestral type of mRNA used for the first ribosome display system to be developed.

 

    Results and discussion
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 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Design and construction of template DNA and mRNA

The design of each template DNA is shown schematically in Figure 2. As mentioned above, we chose human Bcl-xL as prey as a positive control and the precursor to human caspase 3 as prey as a negative control. When we introduced the Cv–mMSp interaction into the selection system, each prey protein was translated as a fusion protein with a tandem dimer of mMSp and a linker protein (Sawata and Taira, 2003Go). The mMSp dimer was joined to the C-terminus of each prey protein and the linker protein was joined to the C-terminus of the mMSp dimer [Figure 2 (I)–(IV)]. No Cv was included at the 5'-end of mRNAs depicted in Figure 2 (III) and (IV), hence these mRNAs served as negative controls for the assessment of the contribution of the Cv–mMSp interaction. We also prepared the two template DNAs depicted in Figure 2 (V) and (VI), which represented the simplest style of ribosome display. Therefore, to determine the effects of the extra mMSp sequences, only the stop codon was removed and no additional motifs were added to these two templates. In all the prey constructs, the original termination codon of the gene for each respective prey protein was eliminated and there was no termination codon in the longest ORF of the mRNA for each prey protein. We chose dihydrofolate reductase (DHFR) from E.coli as the linker protein and we expected it to function as a spacer between the end of the nascent protein on the large subunit of the ribosome and the C-terminus of the mMSp dimer [Figure 2 (I)–(IV)] or prey protein [Figure 2 (V) and (VI)].

We adopted human Bak as bait, in the form of a GST–Bak fusion protein, namely glutathione S-transferase (GST) linked to the N-terminus of Bak [see Figure 2 (VII)]. The termination codon was maintained at its natural position in the correct ORF of the gene for Bak. The transcribed mRNAs were designated rCvBlmM2D(–) [Figure 2 (I)], rCvCamM2D(–) [Figure 2 (II)], rBlmM2D(–) [Figure 2 (III)], rCamM2D(–) [Figure 2 (IV)], rBlD(–) [Figure 2 (V)], rCaD(–) [Figure 2 (VI)] and rGBk(+) [Figure 2 (VII)], with (+) or without (–) a termination codon. Three kinds of mixture of mRNAs were prepared, as follows: a mixture of rCvBlmM2D(–) and rCvCamM2D(–) encoding Bcl-xL and caspase 3, respectively, to test the specificity of Bcl-xL for binding to Bak; a mixture of rBlmM2D(–) and rCamM2D(–) without the Cv motif to examine the importance of the Cv–mMSp interaction; and a mixture of the simplest ribosome display constructs, rBlD(–) and rCaD(–), without Cv and mMSp. These mRNAs were translated in vitro and mRNA–ribosome–nascent protein complexes displaying Bcl-xL were pulled down using agarose beads with immobilized GST–Bak. GST–Bak was also translated from rGBk(+) in vitro. A schematic hypothetical representation of our strategy is shown in Figure 1. After washing the GST–Bak pulled-down complex, we eluted the mRNA from the complex and analyzed it by RT–PCR and electrophoresis or we determined the radioactivity of radioisotope-labeled mRNA.

Confirmation of in vitro selection by RT–PCR

Results of the application of our selection system are shown in Figure 3. When the templates originated from rCvBlmM2D(–) and rBlD(–), the length of each product of RT–PCR, with primers fP-4 and rP-3 (see Materials and methods for the sequences of primers), was 865 bp in each case. Moreover, when the templates originated from rBlmM2D(–), the length of each product of RT–PCR, with primers fP-5 and rP-3, was 813 bp. By contrast, when the templates originated from rCvCamM2D(–) and rCaD(–), the length of each product of RT–PCR, with primers fP-4 and rP-3, was 997 bp in each case. Finally, when the templates originated from rCamM2D(–), the length of each product of RT–PCR, with primers fP-5 and rP-3, was 945 bp.



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Fig. 3. The results of selection in vitro, performed to confirm that our system functioned as we designed it. The patterns of products of RT–PCR after agarose gel electrophoresis are shown. Lanes 1, 2 and 5 show products of RT–PCR amplified from each mRNA pool after selection in vitro and lanes 3, 4 and 6 show products of RT–PCR amplified from each initial mRNA pool before selection in vitro. Lane 1, the initial mRNA pool was an equimolar mixture of rCvBlmM2D(+) and rCvCamM2D(+); lane 2, the initial mRNA pool was an equimolar mixture of rBlD(–) and rCaD(–); lane 3, the product of RT–PCR amplified from an equimolar mixture of rCvBlmM2D(+) and rCvCamM2D(+) prior to selection in vitro; lane 4, the product of RT–PCR amplified from an equimolar mixture of rBlD(–) and rCaD(–) prior to selection in vitro; lane 5, the initial mRNA pool was an equimolar mixture of rBlmM2D(+) and rCamM2D(+); lane 6, the product of RT–PCR amplified from an equimolar mixture of rBlmM2D(+) and rCamM2D(+) prior to selection in vitro; lane M, markers.

 
Lanes 1 and 5 in Figure 3 show products of RT–PCR derived from a pool of eluted mRNA obtained with (lane 1) and without (lane 5) the Cv–mMSp interaction. Lanes 3 and 6 represent products of RT–PCR derived from an initial pool of mRNA, before selection in vitro, with the Cv–mMSp interaction [lane 3; a mixture of constructs (I) and (II) in Figure 2] and without the Cv [lane 6; a mixture of constructs (III) and (IV) in Figure 2]. Lanes 2 and 4 represent the products of RT–PCR derived from a pool [a mixture of constructs (V) and (VI) in Figure 2] of eluted mRNA (lane 2) and the initial pool of the mRNA (lane 4) that represented the simplest ribosome display, as mentioned above. In all lanes, the lower band represents the product of RT–PCR derived from the mRNA for the selection-positive Bcl-xL and the upper band represents the product derived from the mRNA for the selection-negative precursor to caspase 3. Lanes 3, 4 and 6 show that the ratio of the selection-positive mRNA [i.e. rCvBlmM2D(–), rBlD(–) or rBlmM2D(–)] with Bcl-xL to the selection-negative mRNA [i.e., rCvCamM2D(–), rCaD(–) or rCamM2D(–)] without Bcl-xL but with caspase 3 was 1:1 in each initial pool.

In lane 1 in Figure 3, the intensity of the lower band is significantly higher than the intensity of the upper band. Therefore, we can conclude that the desired rCvBlmM2D(–) was selected and the negative control rCvCamM2D(–) was eliminated during the selection in vitro. By contrast, the intensities of the upper and lower bands in lanes 2 and 5 are obviously low, even though lane 5 indicates that the selection itself was successful when we used the conventional ribosome display system. This result suggests that selection in vitro was more efficient when the mRNA participated in the Cv–mMSp interaction (lane 1). Therefore, the introduction of the Cv–mMSp interaction contributed significantly and positively to a successful selection system in vitro.

Estimation of yields of radioisotope-labeled mRNAs recovered by selection in vitro

In order to assess the contribution of the Cv–mMSp interaction quantitatively, we performed a direct comparison of yields of radioisotope-labeled (RI-labeled) mRNAs recovered by in vitro selection. We radiolabeled the following mRNAs with {alpha}-32P-labeled CTP during transcription in vitro: rCvBlmM2D(–), rCvCamM2D(–), rBlmM2D(–), rCamM2D(–), rBlD(–) and rCaD(–). The results are shown in Figure 4. The yield of RNA (radioactivity remaining on agarose beads with immobilized GST–Bak after each wash during selection in vitro) was plotted for each washing step (Figure 4A and C).



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Fig. 4. Direct comparison of yields of RI-labeled mRNA recovered after selection in vitro. The RI-labeled mRNAs prepared for this comparison were rCvBlmM2D(–), rCvCamM2D(–), rBlD(–), rCaD(–), rBlmM2D(–) and rCamM2D(–). The numerals on the horizontal axis in (A) and (C) represent sequential washing steps during the selection in vitro, and the vertical axis represents the yield of each mRNA (%), which was the percentage of the radioactivity (c.p.m.) remaining on the glutathione–agarose beads after each wash during the selection in vitro. The yield of each mRNA after the third wash is shown in (B) and (D).

 
The yield of rCvBlmM2D(–), indicated by green lines in Figure 4A and C, in the presence of the Cv–mMSp interaction was consistently higher than the yields of the other mRNAs after each washing step. We repeated this assay several times and we detected a greater recovery of rCvBlmM2D(–) than that of the other mRNAs in each assay, even though the yield (%) differed among experiments, ranging from 0.025 to 0.063% after the third washing step. A comparison of the yields of each mRNA after the third washing step is presented in Figure 4B and D. The dominant recovery of rCvBlmM2D(–),represented by green bars, as distinct from that of rCvCamM2D(–) (dark-blue bars) is obvious in Figure 4B and D, reflecting the importance of the Cv–mMSp interaction. By contrast, in the absence of the Cv motif, the dominance, in terms of yields of rBlmM2D(–) (light-blue bar) relative to that of rCamM2D(–) (gray bar) was very low (Figure 4D), despite the fact that selection was performed under conventional ribosome display conditions. Similarly, in the absence of the Cv and mMSp, in a system that corresponded to the simplest ribosome display system, the yields of rBlD(–) (purple bar) and rCaD(–) (brown bar) were almost the same (Figure 4B).

The results described above show that the introduction of the Cv–mMSp interaction was a very effective addition to our in vitro selection system. By contrast, in our previous simpler selection system, in which we included a histidine tag, the introduction of the Cv–mMSp interaction improved the stability of the mRNA–protein complex to only a limited extent (Sawata and Taira, 2003Go). Although we cannot explain the discrepancy between the results obtained with the two systems, the present system that exploited the natural protein–protein (Bak–Bcl-xL) interaction clearly demonstrated the importance of the additional Cv–mMSp interaction in our new in vitro selection system, which might complement selection by the yeast two-hybrid system.

The dominance of rBlmM2D(–) (light-blue bar) relative to rCamM2D(–) (gray bar) that is shown in Figure 4D was reproducible and, therefore, we regard it as meaningful. However, in the absence of mMSp, in a system that represented the simplest ribosome display, the yields of rBlD(–) (purple bar) and rCaD(–) (brown bar) were almost identical (Figure 4B). Although the selections for which results are shown in Figure 4B and D depended on the conventional ribosome display system, we consider it likely that the differences between the results of selection from the mixture of rBlmM2D(–) and rCamM2D(–) (Figure 4D) and the results from the mixture of rBlD(–) and rCaD(–) (Figure 4B) were due to the length of the linker peptide. In the former case (Figure 4D), the mMSp dimer might act as a spacer and keep the Bcl-xL away from the ribosome tunnel. Thus, the positive selection was slightly more powerful than in the latter case (Figure 4B). It should be emphasized, however, that the dominance of rCvBlmM2D(–)shows unambiguously that the Cv–mMSp interaction made a positive contribution to in vitro selection and that its contribution was significantly greater than the effect of the length of the linker peptide. Hence we conclude that the advantage of our system depends mainly on the Cv–mMSp interaction.

The recovered yield should be dependent on the Kd for Bcl-xL and Bak, which was estimated by Sattler et al. (1997)Go to be 0.20 µM, as described in the Introduction. The yield and the selective-enrichment ratio of Bcl-xL cDNA relative to caspase 3 cDNA might be higher if Bak were to be prepared by over-expression in E.coli and the amount of immobilized Bak were to be increased. It should be emphasized, however, that our system worked well even when the protein used as bait was prepared by simple small-scale translation in vitro.

Direct observations of ribosome–mRNA complexes by atomic force microscopy

To examine the way in which the Cv–mMSp interaction contributed to the improvement in yield of our in vitro selection system, we examined the mRNA–ribosome–nascent protein complexes by AFM. We postulated that the Cv–mMSp interaction might circularize the mRNA–ribosome–nascent protein complex as depicted in Figure 1. We hoped that AFM would provide direct images of stable polysomes similar to the AFM images of polysomes extracted from Tetrahymena pyriformis (Yoshida et al., 1997Go).

We obtained AFM images of samples after translation in vitro. As shown in Figure 5, the complex in which the Cv–mMSp interaction occurred (Figure 5A) seemed relatively more compact and complicated than the complex in which the Cv–mMSp interaction was not present (Figure 5B). Each image in Figure 5A and B was enclosed within a square of approximately 250x250 nm. Since Vanzi et al. (2003)Go suggested use of a low concentration in order to avoid overlap of ribosomes, the observation area was limited. The estimated number of images, according to the reference, was about two under the conditions of measurements. The sizes of the complexes seen in Figure 5 were similar to those of the natural polysomes extracted from Tetrahymena pyriformis, as estimated by electron microscopy (Yazaki et al., 2000Go). Therefore, the AFM images in Figure 5 probably represented polysomes.

The AFM images of untranslated mRNAs with and without the Cv–mMSp interaction appeared shrunken and similar (data not shown) and different from the images in Figure 5A and B. The difference between the images in Figure 5A and B was probably due to the nascent protein that was being translated in vitro and not just to the structure of the mRNA. The results are suggestive of a significant contribution by the Cv–mMSp interaction to the stability of ribosomes in our display system.

Conclusion

In this study, we were able to detect the interactions between proteins prepared by small-scale translation in vitro as a result of the introduction of the Cv–mMSp interaction into a ribosome display system. We chose the interaction between human Bcl-xL and human Bak for our analysis of the efficacy of our system. Bcl-xL can be inserted either into synthetic lipid vesicles or into planar lipid bilayers, where it forms ion-conducting channels similar to those formed by bacterial toxins (Minn et al., 1997Go) and it also binds to the mitochondrial protein channel (the voltage-dependent anion channel, VDAC) (Shimizu et al., 1999Go). Similarly, Bak is localized on the outer membrane of mitochondria in healthy cells (Nechushtan et al., 2001Go). Thus, both Bcl-xL and Bak are membrane proteins. In the present analysis, we were able to prepare both membrane proteins as entire proteins by translation in vitro despite the fact that both contain hydrophobic domains.

We are not the first to isolate Bcl-xL-binding proteins in vitro. The selection of the Bcl-xL-binding proteins in vitro has been achieved using cDNA libraries prepared for mRNA display technology (Hammond et al., 2001Go). However, cDNA libraries often do not contain full-length ORFs. Therefore, to our knowledge, this study provides the first demonstration that ribosome display can be applied to selection in vitro that depends on the interaction between intact, full-sized membrane proteins that have been translated in vitro. This system can easily be expanded for high-throughput screening (HTS) and should be a very useful tool for molecular biologists and for the identification of peptide drugs.


    Acknowledgments
 
The authors thank Dr Renu Wadhwa (AIST) and Dr Izumi Yoshizaki (Japan Aerospace Exploration Agency, JAXA) for very helpful comments on the manuscript. This research was supported by grants from the Program for the Promotion of Basic Research Activities for Innovative Biosciences (PROBRAIN) and from the Ministry of Economy, Trade and Industry (METI) of Japan.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results and discussion
 References
 
Bartel,P.L. and Fields,S. (1995) Methods Enzymol., 254, 241–263.[ISI][Medline]

Boise,L.H., Gonzalez-Garcia,M., Postema,C.E., Ding,L., Lindsten,T., Turka,L.A., Mao,X., Nunez,G. and Thompson,C.B. (1993) Cell, 74, 597–608.[ISI][Medline]

Chien,C., Bartel,P.L., Sternglanz,R. and Fields,S. (1991) Proc. Natl Acad. Sci. USA, 88, 9578–9582.[Abstract]

Chittenden,T., Harrington,E.A., O'Connor,R., Flemington,C., Lutz, R.J., Evan,G.I. and Guild,B.C. (1995) Nature, 374, 733–736.[CrossRef][ISI][Medline]

Convery,M.A., Rowsell,S., Stonehouse,N.J., Ellington,A.D., Hirao,I., Murray,J.B., Peabody,D.S., Phillips,S.E.V. and Stockley,P.G. (1998) Nat. Struct. Biol., 5, 133–139.[ISI][Medline]

Farrow,S.N., White,J.H.M., Martinou,I., Raven,T., Pun,K.T., Grinham,C.J., Martinou,J.C. and Brown,R. (1995) Nature, 374, 731–733.[CrossRef][ISI][Medline]

Fields,S. and Song,O. (1989) Nature, 340, 245–246.[CrossRef][ISI][Medline]

Gersuk,G.M., Corey,M.J., Corey,E., Stray,J.E., Kawasaki,G.H. and Vessella,R.L. (1997) Biochem. Biophys. Res. Commun., 232, 578–582.[CrossRef][ISI][Medline]

Hammond,P.W., Alpin,J., Rise,C.E., Wright,M. and Kreider,B.L. (2001) J. Biol. Chem., 276, 20898–20906.[Abstract/Free Full Text]

Hanes,J. and Plückthun,A. (1997) Proc. Natl Acad. Sci. USA, 94, 4937–4942.[Abstract/Free Full Text]

He,M. and Taussig, M.J. (1997) Nucleic Acids Res., 25, 5132–5134.[Abstract/Free Full Text]

Johansson,H.E., Dertinger,D., LeCuyer,K.A., Behlen,L.S., Greef,C.H. and Uhlenbeck,O.C. (1998) Proc. Natl Acad. Sci. USA, 95, 9244–9249.[Abstract/Free Full Text]

Kiefer,M.C., Brauer,M.J., Powers,V.C., Wu,J.J., Umansky,S.R., Tomei,L.D. and Barr,P.J. (1995) Nature, 374, 736–739.[CrossRef][ISI][Medline]

LeCuyer,K.A., Behlen,L.S. and Uhlenbeck,O.C. (1995) Biochemistry, 34, 10600–10606.[ISI][Medline]

Mattheakis,L.C., Bhatt,R.R. and Dower,W.J. (1994) Proc. Natl Acad. Sci. USA, 91, 9022–9026.[Abstract]

Minn,A.J., Vèlez,P., Schendel,S.L., Liang,H., Muchmore,S.W., Fesik,S.W., Fill,M. and Thompson,C.B. (1997) Nature, 385, 353–357.[CrossRef][ISI][Medline]

Nechushtan,A., Smith,C.L., Lamensdorf,I., Yoon,S.H. and Youle,R.J. (2001) J. Cell Biol., 153, 1265–1276.[Abstract/Free Full Text]

Peabody,D.S. (1997) Arch. Biochem. Biophys., 347, 85–92.[CrossRef][ISI][Medline]

Rowsell,S., Stonehouse,N.J., Convery,M.A., Adams,C.J., Ellington,A.D., Hirao,I., Peabody,D.S., Stockley,P.G. and Phillips,S.E.V. (1998) Nat. Struct. Biol., 5, 970–975.[CrossRef][ISI][Medline]

Sattler,M. et al. (1997) Science, 275, 983–986.[Abstract/Free Full Text]

Sawata,S.Y. and Taira,K. (2003) Protein Eng., 16, 1115–1124.[ISI][Medline]

Shimizu,S., Narita,M. and Tsujimoto,Y. (1999) Nature, 399, 483–487.[CrossRef][ISI][Medline]

Smith,G.P. (1985) Science, 228, 1315–1317.[ISI][Medline]

Suyama,E., Kawasaki,H. and Taira K (2002) FEBS Lett., 528, 63–69.[CrossRef][ISI][Medline]

Thornberry,N.A. and Lazebnik,Y. (1998) Science, 281, 1312–1316.[Abstract/Free Full Text]

Valegård,K., Murray,J.B., Stonehouse,N.J., van den Worm,S., Stockley,P.G. and Liljas,L. (1997) J. Mol. Biol., 270, 724–738.[CrossRef][ISI][Medline]

Vanzi,F., Vladimirov,S., Knudsen,C.R., Goldman,Y.E. and Cooperman,B.S. (2003) RNA, 9, 1174–1179.[Abstract/Free Full Text]

Yazaki,K., Yoshida,T., Wakiyama,M. and Miura,K. (2000) J. Electron Microsc., 49, 663–668.[Abstract]

Yoshida,T., Wakiyama,M., Yazaki,K. and Miura,K. (1997) J. Electron Microsc., 46, 503–506.[Abstract]

Received October 26, 2003; revised April 20, 2004; accepted April 29, 2004.

Edited by Haruki Nakamura





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