Binding of external ligands onto an engineered virus capsid

Uli Schmidt1,3,4, Rainer Rudolph1 and Gerald Böhm1,2

1 Institut für Biotechnologie, Martin-Luther-Universität Halle-Wittenberg, Kurt-Mothes-Str. 3, 06120 Halle and 2 ACGT Progenomics AG, Weinbergweg 22, 06120 Halle, Saale, Germany


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
The development of novel delivery systems for therapeutic substances includes targeting of the carriers to a specific site or tissue within the body of the recipient. This can be accomplished by appropriate receptor-binding domains and requires linking of these domains to the carrier. We have used recombinantly expressed polyomavirus-like particles as a model system and inserted the sequence of a WW domain into different surface loops of the viral capsid protein VP1. In one variant, the WW domain maintained its highly selective binding properties of proline-rich ligands and showed an increased affinity but also an accelerated association/dissociation equilibrium compared to the isolated WW domain, thus allowing a short-term coupling of external ligands onto the surface of the virus-like particles.

Keywords: cellular targeting/drug delivery/polyomavirus VP1/protein design/WW domain


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In recent years, there have been intense efforts to generate targetable, injectable vectors for gene and drug delivery based on a variety of viral and non-viral systems (Anderson, 1998Go). It is envisaged that the ideal targeted vector would be administered by intravenous infusion or injection, and then concentrates in the tissues and organs harboring the targeted cells (Peng and Russel, 1999Go). Strategies to achieve a cell-type specific targeting often use direct genetic modification of the coat proteins with receptor-binding domains or sequences, e.g. the insertion of integrin-binding RGD sequences into an adenoviral capsid (Wickham et al., 1997Go; Vigne et al., 1999Go). Another approach is the use of soluble bifunctional crosslinkers that bind both to the vector and to a cell-surface receptor (Douglas et al., 1996Go). A combined approach is to display an immunoglobulin-binding domain of protein A on the vector as a genetic fusion to the coat protein, and then use a monoclonal antibody to crosslink the vector with the targeted cell (Ohno and Meruelo, 1997Go; Ohno et al., 1997Go). Here, we investigate a more general strategy using a genetic fusion of a viral capsid protein with a WW domain, in order to bind any recombinantly produced polyproline-tagged protein which can be, for example, an antibody or a receptor-binding domain.

Surface loops of viral capsid proteins often show a high sequence variability due to the influence of the immune system in viral life-cycles which makes these loops susceptible to the insertion of foreign sequences (Stirk and Thornton, 1994Go). Recombinantly expressed polyomavirus-like particles were used as a model for a viral protein shell. The outer capsid protein VP1 of murine polyomavirus can be expressed from recombinant Escherichia coli (Leavitt et al., 1985Go; Schmidt et al., 2000Go; Stubenrauch et al., 2000Go). It is a pentameric protein which forms virus-like particles in vitro consisting of 72 pentamers (Salunke et al., 1986Go; Salunke et al., 1989Go). The in vitro assembly process was studied in great detail and is thought to involve a pre-existing equilibrium between free capsomeres and capsids which is completely shifted to capsids upon oxidation of a single disulfide bridge (Schmidt et al., 2000Go). The crystal structures of the capsid and of truncated pentameric VP1 have been reported (Stehle et al., 1994Go; Stehle and Harrison, 1996Go, 1997Go). Polyomavirus VP1 gained attention for the in vitro packaging of plasmid DNA and oligonucleotides for the development of non-viral gene transfer vectors (Forstova et al., 1995Go; Braun et al., 1999Go).

In order to specifically bind proline-rich ligands onto the outer surface of VP1 capsids, the sequence of a WW domain was genetically fused into ß-turns of VP1. WW domains are the smallest protein domains known so far and were first discovered in the Yes-kinase associated protein (YAP) of Saccharomyces cerevisiae (Bork and Sudol, 1994Go; Sudol et al., 1995Go). They were named after two conserved tryptophan residues which are essential for the maintenance of the native fold and for ligand binding (Koepf et al., 1999Go). Until now, WW domains were found in several proteins of different species where they contribute to signal transduction processes or to protein–protein interactions in general. In analogy to SH3 domains, WW domains bind proline-rich peptide sequences which differ in their consensus sequences (Pawson and Scott, 1997Go). WW domains can either be subdivided into four classes according to their binding specificity (Bedford et al., 2000Go) or into three subclasses according to sequence similarity (Macias et al., 2000Go). The NMR structures of WW domains representing all three sequence subtypes have been reported, including the YAP WW domain in complex with a proline-rich peptide (Macias et al., 1996Go) and the FBP28 and YJQ8 WW domains (Macias et al., 2000Go). The WW domain contains a three-stranded antiparallel ß-sheet; the proline-rich peptide is mostly bound by hydrophobic interactions. In this study the first of two WW domains of the mouse formin-binding protein 11 (FBP11) was used which binds PPLP-motifs (Chan et al., 1996Go). The affinity of this WW domain to its ligand is the highest reported so far for WW domains. Its equilibrium dissociation constant KD is 21 nM (Bedford et al., 1997Go).


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Protein modeling

The structure of VP1-WW150 was modeled using the program Modeller 4 (Sali and Blundell, 1993Go) based on the published structures of VP1 at 1.9 and 3.65 Å (Stehle and Harrison, 1996Go, 1997Go). The inserted FBP11 WW domain was modeled using the NMR structure of the YAP WW domain (Macias et al., 1996Go). Figures were generated by MolScript (Kraulis, 1991Go) and Raster3D (Merritt and Bacon, 1997Go).

Cloning and vector construction

The first FBP11 WW domain was genetically fused into the VP1 sequence as shown in Figure 2Go using the oligonucleotides FBP11-WWa-5' (5'-ATACTCTTCA GGCAGCGGCT GGACAGAACA TAAATCACCT GATGG-3'), FBP11-WWa-3' (5'-ATACTCTTCT ACCACTACCA TCATCCGGCT TTTCCCAGGT AGACTG-3'), VP1-150-WWaC (5'-ATACTCTTCA GGTAGCGGCG TAAACACAAA AGGAATTTCC ACTCCAG-3'), VP1-150-WWaN (5'-ATACTCTTCA GCCGCTGCCT GTATCTGTCG GTTTGTTGAA CCCATG-3'), VP1-292-WWaC (5'-ATACTCTTCA GGTAGCGGCG TTACAAGAAA CTATGATGTC CATCAC-3'), VP1-292-WWaN (5'-ATACTCTTCA GCCGCTGCCC CAGCCCATTA TATCTACGCT CGAG-3'), VP1-Nde I-5' (5'-TATACATATG GCCCCCAAAA GAAAAAGC-3') and VP1-Sma I-3' (5'-ATATCCCGGG AGGAAATACA GTCTTTGTTT TTCC-3'). The resulting final PCR product was cloned via introduced NdeI and SmaI restriction sites into the plasmid pET21-Int which contained a T7lac-promoter for high level expression in E.coli, and a C-terminal fusion with an intein and a chitin-binding domain for affinity chromatography (Schmidt et al., 2000Go).



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Fig. 2. Cloning of the fusion constructs VP1-WW150 and VP1-WW292. The strategy avoids changes in the original sequences and involves the ligation and reamplification of three separate fragments that were generated by PCR. The recognition sequence of the endonuclease Eam1 104 I is printed in bold letters.

 
Protein expression and purification

All proteins were expressed from recombinant E.coli as C-terminal fusion proteins with modified intein and chitin-binding domains and were purified as described before (Chong et al., 1997Go; Schmidt et al., 2000Go).

In vitro assembly and size-exclusion chromatography

Particles were assembled in vitro according to Salunke et al. (Salunke et al., 1986Go, 1989Go). For the binding onto virus-like particles the proline-rich ligands were added after completion of the assembly process. The capsid assembly was quantitatively analyzed by size-exclusion chromatography using 14 ml TSKgel 5000/6000PWXL columns (Tosoh Biosep, Stuttgart, Germany) as described before (Schmidt et al., 2000Go).

Surface plasmon resonance

In order to determine the affinity of the VP1-WW fusion proteins to polyproline sequences, surface plasmon resonance was measured using a Biacore X (Biacore AB, Uppsala, Sweden) and a CM5 sensorchip which was coated with PPLP peptide (sequence: CSGP6PPLP) following the manufacturer's protocol. The protein concentrations were varied between 5 and 50 nM. For the screening of different buffers and additives the protein solution was diluted in the buffer which was also used for the measurement. Kinetic parameters were calculated with the BIAevaluation software using a simple Langmuir binding model.

Circular dichroism (CD) spectroscopy

Far-UV CD spectra of VP1 variants were measured from 195 to 260 nm in 0.1 mm cuvettes. The proteins (concentration: 0.5 to 1.0 mg/ml) were dialyzed against a buffer containing 10 mM HEPES, 100 mM NaCl, pH 7.2. The secondary structure contents of the proteins were calculated with the program CDNN (Böhm et al., 1992Go) from the buffer-corrected spectra.

Electron microscopy

For electron microscopy studies, an EM 912 instrument (Zeiss) was used with a magnification factor of 63 000. Staining of the specimen was performed with uranyl acetate on bacitracin-incubated (0.1 mg/ml, 1 min) copper–carbon-grids according to standard protocols.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Protein design and model building

Two variants of the fusion protein were designed for the presentation of the WW domain on the surface of polyomavirus-like particles. The first incorporates the WW domain in the DE loop at position 150 in the VP1 sequence and the second in the HI loop at position 292. Both loops are well accessible from the outer surface and are flexible according to the temperature factors determined from crystallographic data (Stehle and Harrison, 1996Go). These positions are also distant from the CD loop and C-terminal sequences which are required for the formation of virus-like particles. Therefore, insertions at these positions should not inhibit the formation of virus-like particles. A sequence alignment did not reveal any conservation in those regions between polyomavirus strains from different species (data not shown). The WW domain insert was flanked by spacers of five amino acids consisting of serine/glycine repeats to allow maximal flexibility and solubility. The overall size of the inserted sequence was 38 amino acids. A model of the VP1-WW150 structure is presented in Figure 1Go.



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Fig. 1. Model structure of VP1-WW150: (a) monomeric subunit and (b) pentameric capsomere. The WW domain is represented in purple and the serine/glycine-linker in blue. The WW domain folds towards the central hole of the capsomere and does not interfere with the pentameric structure.

 
Cloning and protein expression

For the genetic fusion of the WW domain into the VP1 sequence, PCR products with overhangs containing the serine/glycine linker sequences and recognition sites for the type IIs restriction endonuclease Eam1 104 I were generated (Figure 2Go). Eam1 104 I cleaves outside its recognition sequence and therefore allows ligation of fragments without the introduction of suitable restriction sites.

The expression of both proteins as C-terminal fusions with intein and chitin-binding domains allowed a single-step purification (Chong et al., 1997Go; Schmidt et al., 2000Go) and yielded approximately 6 mg of purified protein per liter of culture medium.

CD analysis reveals increased ß-sheet contents

Far-UV CD spectra were recorded and compared with the authentic VP1 in order to verify the native fold of the VP1-WW fusion proteins (Figure 3Go). The spectra of VP1 and VP1-WW292 had similar shapes, whereas the VP1-WW150 spectrum had a significantly increased negative ellipticity difference ({Delta}{varepsilon}) below 207 nm, indicating a higher portion of ß-sheet secondary structure. Secondary structure deconvolution of the spectra revealed an increase of antiparallel ß-sheets in VP1-WW150 and VP1-WW292, which was, however, significantly higher for VP1-WW150. The difference spectra of VP1-WW150/VP1-WW292 minus VP1 should represent the spectrum of the single WW domain (Figure 3Go). The difference CD spectrum of VP1-WW150 had a maximum at 225 nm and an intense negative ellipticity at 198 nm. This spectrum corresponds to a typical WW domain spectrum that is shifted approximately 5 nm towards shorter wavelengths (Macias et al., 2000Go), indicating that the WW domain obtains its native fold in the fusion protein VP1-WW150. In contrast, the difference spectrum of VP1-WW292 showed a much smaller amplitude, suggesting that the WW domain is not or only partially folded in this construct.



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Fig. 3. Circular dichroism analysis. (a) CD spectra of pentameric VP1, VP1-WW150 and VP1-WW292. (b) Deconvolution with the program CDNN (Böhm et al., 1992) indicates an increased secondary structure content of antiparallel ß-sheet in the fusion constructs. (c) Difference spectra of VP1-WW150/VP1-WW292 minus VP1, representing the spectra of the WW domains in the fusion proteins as described in the literature. (d) Thermal denaturation experiments demonstrate that the thermal stability of VP1-WW150 is 6°C lower than that of VP1 (51°C). VP1-WW292 starts to aggregate at 40°C.

 
Ellipticity changes during thermal denaturation yielded a Tm of 51°C for the VP1 capsomer (Figure 3Go). The thermal transition midpoint for VP1-WW150 was lowered by 6°C to 45°C. This value corresponds well to the typical midpoint transition temperature determined for a prototype WW domain (44°C; Macias et al., 2000Go). The denaturation of VP1-WW292 did not have a sigmoidal shape and began to decrease already at 40°C and increased again beyond 60°C, probably due to aggregation followed by sedimentation of the protein that may be caused by large unfolded regions in the protein. The starting aggregation at 40°C was therefore set to be the starting point of thermal denaturation of VP1-WW292.

VP1-WW150 specifically binds proline-rich ligands

For an analysis of the binding properties of the inserted WW domains, a proline-rich peptide with the sequence +H3N-CSGP6PPLP-COOminus; was immobilized via its N-terminal cysteine residue on the surface of a sensorchip for surface plasmon resonance measurements. This peptide contains the PPLP consensus motif that makes it an ideal ligand for binding of the FBP11 WW domain used in this study (Bedford et al., 1997Go). The sensorchip was tested with a linear construct of the WW domain as an N-terminal fusion with glutathione-S-transferase (GST-WW). For this protein, the equilibrium dissociation constant KD was determined to be 18 nM (Table IGo), in good agreement with the value of 21 nM published earlier (Bedford et al., 1997Go).


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Table I. Kinetic parameters of the binding reaction of the WW domain to a proline-rich peptide using a Langmuir binding model
 
The kinetic parameters of the interaction of the WW domains within the VP1 sequence with the peptide were determined with serial dilutions of the fusion proteins under different buffer conditions. VP1 without a WW domain did not interact with the sensorchip surface or with the immobilized peptide (data not shown). The protein VP1-WW292 did not bind to the ligand under any condition tested so far, indicating that the WW domain in this construct did not obtain a native fold or had a dramatically lowered affinity which, therefore, could not be measured using surface plasmon resonance.

However, VP1-WW150 showed a high affinity for the ligand (Figure 4Go). Using the same conditions as for GST-WW, the equilibrium dissociation constant KD of VP1-WW150 was increased to 7.7 ± 5 nM (Table IGo). VP1-WW150 exhibited a high affinity for the proline-rich ligand in the range of KD = 4–15 nM under all buffer conditions tested. In order to mimic a cell culture or physiological system, the measurement was carried out in phosphate buffered saline (PBS) and in Dulbecco's modified Eagle cell culture medium supplemented with 10% fetal calf serum. The serum proteins did not compete with the binding sites on the VP1 surface and their presence did not inhibit the specific binding of VP1-WW150 to its PPLP-ligand, a prerequisite for therapeutic applications. Although the equilibrium constants were similar for the inserted and the linear WW domain, the interaction of the inserted WW domain is accompanied by an accelerated exchange of the ligands (Table IGo). Association and dissociation reactions (represented by the association/dissociation rate constants ka and kd in Table IGo) of VP1-WW150 were approximately 10 times faster than the respective values of GST-WW.



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Fig. 4. Surface plasmon resonance with an immobilized PPLP ligand. Analysis with WW domain concentrations of 20, 15, 10 and 5 nM, respectively, in a buffer containing 20 mM HEPES, 1 mM EDTA, 200 mM NaCl, pH 7.2 of (a) cysteine free VP1-WW150 and (b) VP1-WW150 containing cysteines 19 and 114.

 
Altered in vitro assembly properties of VP1-WW fusion proteins

An essential function of VP1 is its ability to form virus-like particles in vitro. For easier handling, the VP1-WW fusion proteins used so far did not contain cysteine residues and were therefore unable to form an intrapentameric disulfide bridge which is needed for a complete capsid assembly. However, upon addition of Ca2+ ions an equilibrium with 55% reduced capsids and 45% free capsomeres should be reached which is an early step during the assembly process (Schmidt et al., 2000Go). However, removal of EDTA and the addition of Ca2+ resulted in a loss of 90% of the protein due to significant aggregation of VP1-WW150; this indicated a strong decrease of the solubility of the reduced capsid species which resulted in precipitation until the protein concentration fell below a critical limit. This was not due to aggregation via the WW domain in the presence of Ca2+ since the solubility of similar concentrations of the GST-WW protein did not decrease upon addition of CaCl2 (data not shown). Also, in contrast to VP1-WW150, the protein VP1-WW292 did not aggregate and remained in solution.

The in vitro assembly can be quantitatively analyzed by size-exclusion chromatography (Schmidt et al., 2000Go). Analysis of the assembly of the VP1-WW proteins revealed that VP1-WW150 that still remained in solution assembled to only 15% (Figure 5Go). VP1-WW292 did not form capsids at all and represents the first described variant of VP1 which is totally blocked in capsid formation, although the C-terminal domain for the interaction of capsomeres is present (Figure 5Go). These results demonstrate that large inserts, like the 38 amino acids containing the WW domain, into the viral capsid protein had long-distance effects on parts of the protein which are essential for capsid formation.



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Fig. 5. In vitro assembly of VP1-WW fusion proteins. (a) Size-exclusion chromatography (TSKgel 5000PWXL) demonstrates that only VP1-WW150, which contains cysteines 19 and 114, forms quantitatively virus-like particles. Elution volumes of capsids (6–8 ml) and free capsomeres (10–11 ml) are marked. (b) Electron microscopy shows an homogeneous population of disulfide-stabilized VP1-WW150 capsids.

 
An intrapentameric disulfide bridge between cysteines 19 and 114 of neighboring monomers is needed for a complete capsid assembly of the wild-type protein (Stehle et al., 1994Go; Schmidt et al., 2000Go). In order to improve the inefficient cysteine-free in vitro assembly and to test whether oxidized, disulfide-linked VP1-WW150 capsids are also prone to aggregation, cysteines 19 and 114 were reintroduced into VP1-WW150. The disulfide bridge forming variant of VP1-WW150 had no differences in the binding properties of proline-rich ligands (Figure 4Go; Table IGo) and did not aggregate upon initiation of in vitro assembly by dialysis. Size-exclusion chromatography demonstrated that the assembly efficiency of the modified protein was improved to >95%, similar to the wild-type protein. Electron micrographs also showed an homogeneous population of virus-like particles consisting of VP-WW150 (Figure 5Go). This single intrapentameric disulfide bridge could therefore completely restore the assembly properties of the wild-type protein.

VP1-WW150 capsids can bind external ligands in solution

In order to test the ability of polyomavirus-like particles to bind external ligands in solution, as would be necessary for therapeutic applications, completely assembled disulfide stabilized capsids of VP1-WW150 were mixed with an equal molar amount of green fluorescent protein (GFP) that was tagged with a PPLP motif at its C-terminus. The formation of the complex consisting of capsids and GFP was analyzed by size-exclusion chromatography (Figure 6Go). GFP could be detected by its specific absorbance at 490 nm while detecting the protein absorbance at 280 nm for both capsids and GFP in parallel. Absorbance at 490 nm at the elution volume of the capsids represented GFP bound to the particle's surfaces. Control experiments with VP1 wild-type capsids showed no co-elution of GFP with the capsids (data not shown). Integration of the peak areas and calculation of the molar ratio of capsids and GFP, respectively, indicated that 25 ± 5 molecules of GFP were bound to the capsid surface, demonstrating the general applicability of our approach. The binding ratio was probably limited by the rapid dissociation reaction of the inserted WW domain.



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Fig. 6. Coupling of polyproline-tagged GFP onto the surface of virus-like particles. GFP-loaded capsids (elution volume 8–10 ml) are separated from excess GFP and minor impurities by size-exclusion chromatography (TSKgel 6000PWXL).

 

    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In this work polyomavirus-like particles were used as a model for viral protein shells in order to study the influence of insertion mutants with a module for the binding of external ligands onto the particle's surfaces. The binding module must fulfil certain criteria: It should specifically bind peptide ligands with a high or regulatable affinity. Preferably, it should also have a small, compact structure to avoid interferences with other functions of the protein shell. A module which combines many of these properties is the WW domain, the smallest, independently folding protein domain known to date. The highest affinity has been reported for the first FBP11 WW domain to a proline-rich sequence with a PPLP consensus motif (Bedford et al., 1997Go).

Although theoretical considerations and calculations suggested that the WW domain could fold independently in both ß-turns, which we chose for insertion into the VP1 sequence, only one variant, VP1-WW150, could specifically interact with proline-rich ligands. This was expected from CD and thermal denaturation experiments that demonstrated that the WW domain is only folded in VP1-WW150. Probably, the conformation of the DE loop of VP1 is more compatible with the WW domain structure than the alternative HI loop. This might not be a general conclusion since previous experiments demonstrated that insertion of an enzyme, namely dihydrofolate reductase, into the polyomavirus HI loop, yielded pentamers with enzymatic activity; however, these pentamers formed smaller virus-like particles than the VP1-wild-type protein (Gleiter et al., 1999Go). In contrast to these data we did not observe morphological differences between VP1-WW150 and VP1-wild-type capsids; the assembly efficiency of the modified capsomeres was similar to the wild-type protein, provided that an intrapentameric disulfide bridge could be formed during in vitro assembly.

The binding kinetics of the WW domain within the fusion protein has implications for the use of external ligand binding for therapeutic applications. The binding of a PPLP motif by the integrated WW domain (VP1-WW150) compared to a linear construct (GST-WW) resulted in an increased affinity in equilibrium while the ligand exchange was accelerated 10 times (Table IGo). Nevertheless, it could be demonstrated that a complex of polyproline-tagged GFP and virus-like particles can be isolated and that this complex is stable during this procedure (Figure 6Go). However, the coupling efficiency as well as the stability of the complex was limited by the fast dissociation reaction that would possibly reduce the effectiveness of the bound ligand in vivo. This is a general problem of non-covalently linked external ligands and was also discussed for the IgG-binding protein A fusion constructs with viral vector surfaces (Wickham, 1997Go).

In order to extend the stability of the complexes, modified WW domains were designed that allow the formation of a disulfide bridge with the ligand, and the analysis of binding and covalent linking looks very promising (C.Parthier and U.Schmidt, own unpublished results). It has also been proposed that the WW domain fold may constitute a template into which binding sites from unrelated proteins may be introduced in order to mimic proteins for use in drug design because of the high degree of sequence variability allowed by WW domains (Macias et al., 2000Go). Therefore, random mutagenesis of WW domains, selection for a desired binding activity and reintroduction of the altered domain into the protein shell could result in viral particles with a new cellular tropism without the need for external receptor-binding domains or antibody fragments. Alternatively, a two-step therapeutic approach would be conceivable that marks in a first step the target cells with an antibody fragment presenting a proline-rich sequence, thereby directing the WW domain to the target cells in a second step. Similar strategies involving biotin/streptavidin interactions were used for a tissue specific radiation therapy (Paganelli et al., 1999Go).

In summary, our experiments underline the applicability of protein design for the modification and incorporation of new functions into viral capsid proteins. WW domains as modules for the binding of external ligands constitute a flexible tool for future investigations in this direction. These or similar approaches will be useful for individual vector targeting systems and could have a broad range of applications in the drug delivery field.


    Notes
 
3 Present address: WAIMR Cancer Biology Division, Institute for Child Health Research, 100 Roberts Road, Subiaco, WA 6008, Australia Back

4 To whom correspondence should be addressed. E-mail: ulis{at}ichr.uwa.edu.au Back


    Acknowledgments
 
We thank Dr Mark Bedford, Harvard Medical School, USA, for providing the plasmid containing the gene of the FBP11 WW domain and Dr Dieter Neumann, IPB Halle, Germany, for his help with electron microscopy. We also thank Constanze Günther from our laboratory for providing the GFP with the PPLP-tag and for excellent technical assistance. This work was supported by a grant from Land Sachsen-Anhalt.


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Anderson,W.F. (1998) Nature, 392(Suppl.), 25–30.[ISI][Medline]

Bedford,M.T., Chan,D.C. and Leder,P. (1997) EMBO J., 16, 2376–2383.[Abstract/Free Full Text]

Bedford,M.T., Frankel,A., Yaffe,M.B., Clarke,S., Leder,P. and Richard,S. (2000) J. Biol. Chem., 275, 16030–16036.[Abstract/Free Full Text]

Böhm,G., Muhr,R. and Jaenicke,R. (1992) Protein Eng., 5, 191–195.[Abstract]

Bork,P. and Sudol,M. (1994) Trends Biochem. Sci., 19, 531–533.[ISI][Medline]

Braun,H., Boller,K., Löwer,J., Bertling,W.M. and Zimmer,A. (1999) Biotechnol. Appl. Biochem., 29, 31–43.[ISI][Medline]

Chan,D.C., Bedford,M.T. and Leder,P. (1996) EMBO J., 15, 1045–1054.[Abstract]

Chong,S. et al. (1997) Gene, 192, 271–281.[ISI][Medline]

Douglas,J.T., Rogers,B.E., Rosenfeld,M.E., Michael,S.I., Feng,M. and Curiel,D.T. (1996) Nat. Biotechnol., 14, 1574–1578.[ISI][Medline]

Forstova,J., Krauzewicz,N., Sandig,V., Elliot,J., Palkova,Z., Strauss,M. and Griffin,B.E. (1995) Hum. Gene Ther., 6, 297–306.[ISI][Medline]

Gleiter,S., Stubenrauch,K. and Lilie,H. (1999) Protein Sci., 8, 2562–2569.[Abstract]

Koepf,E.K., Petrassi,H.M., Ratnaswamy,G., Sudol,M. and Kelly,J.W. (1999) Biochemistry, 38, 14338–14351.[ISI][Medline]

Kraulis,P. (1991) J. Appl. Crystallogr., 24, 946–950.[ISI]

Leavitt,A.D., Roberts,T.M. and Garcea,R.L. (1985) J. Biol. Chem., 23, 12803–12809.

Macias,M.J., Hyvönen,M., Baraldi,E., Schultz,J., Sudol,M., Saraste,M. and Oschkinat,H. (1996) Nature, 382, 646–649.[ISI][Medline]

Macias,M.J., Gervais,V., Civera,C. and Oschkinat,H. (2000) Nat. Struct. Biol., 7, 375–379.[ISI][Medline]

Merritt,E.A. and Bacon,D.J. (1997) Methods Enzymol., 277, 505–524.[ISI]

Ohno,K. and Meruelo,D. (1997) Biochem. Mol. Med., 62, 123–127.[ISI][Medline]

Ohno,K., Sawai,K., Lijima,Y., Levin,B. and Meruelo,D. (1997) Nat. Biotechnol., 15, 763–767.[ISI][Medline]

Paganelli,G., Grana,C., Chinol,M., Cremonesi,M., De Cicco,C., De Braud,F., Robertson,C., Zurrida,S., Casadio,C., Zoboli,S., Siccardi,A.G. and Veronesi,U. (1999) Eur. J. Nucl. Med., 26, 348–357.[ISI][Medline]

Pawson,T. and Scott,J.D. (1997) Science, 278, 2075–2080.[Abstract/Free Full Text]

Peng,K.W. and Russel,S.J. (1999) Curr. Opin. Biotechnol., 10, 454–457.[ISI][Medline]

Sali,A. and Blundell,T.L. (1993) J. Mol. Biol., 234, 779–815.[ISI][Medline]

Salunke,D.M., Caspar,D.L. and Garcea,R.L. (1986) Cell, 46, 895–904.[ISI][Medline]

Salunke,D.M., Caspar,D.L. and Garcea,R.L. (1989) Biophys. J., 56, 887–900.[Abstract]

Schmidt,U., Rudolph,R. and Böhm,G. (2000) J. Virol., 74, 1658–1662.[Abstract/Free Full Text]

Stehle,T. and Harrison,S.C. (1996) Structure, 4, 183–194.[ISI][Medline]

Stehle,T. and Harrison,S.C. (1997) EMBO J., 16, 5139–5148.[Abstract/Free Full Text]

Stehle,T., Yan,Y., Benjamin,T.L. and Harrison,S.C. (1994) Nature, 369, 160–163.[ISI][Medline]

Stirk,H.J. and Thornton,J.M. (1994) Protein Eng., 7, 47–56.[Abstract]

Stubenrauch,K., Bachman,A., Rudolph,R. and Lilie,H. (2000) J. Chromatogr. B Biomed. Sci. Appl., 737, 77–84.[Medline]

Sudol,M., Bork,P., Einbond,A., Kastury,K., Druck,T., Negrini,M., Huebner,K. and Lehman,D. (1995) J. Biol. Chem., 270, 14733–14741.[Abstract/Free Full Text]

Vigne,E., Mahfouz,I., Dedieu,J.F., Brie,A., Perricaudet,M. and Yeh,P. (1999) J. Virol., 73, 5156–5161.[Abstract/Free Full Text]

Wickham,T.J. (1997) Nat. Biotechnol., 15, 717.[ISI][Medline]

Wickham,T.J., Tzeng,E., Shears II,L.L., Roelvink,P.W., Li,Y., Lee,G.M. Lizonova,A., Brough,D.E. and Kovesdi,I. (1997) J. Virol., 71, 8221–8229.[Abstract]

Received June 30, 2000; revised June 11, 2001; accepted July 10, 2001.