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
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Abstract |
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Keywords: cellular targeting/drug delivery/polyomavirus VP1/protein design/WW domain
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Introduction |
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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, 1994). 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., 1985
; Schmidt et al., 2000
; Stubenrauch et al., 2000
). It is a pentameric protein which forms virus-like particles in vitro consisting of 72 pentamers (Salunke et al., 1986
; Salunke et al., 1989
). 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., 2000
). The crystal structures of the capsid and of truncated pentameric VP1 have been reported (Stehle et al., 1994
; Stehle and Harrison, 1996
, 1997
). 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., 1995
; Braun et al., 1999
).
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, 1994; Sudol et al., 1995
). 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., 1999
). Until now, WW domains were found in several proteins of different species where they contribute to signal transduction processes or to proteinprotein interactions in general. In analogy to SH3 domains, WW domains bind proline-rich peptide sequences which differ in their consensus sequences (Pawson and Scott, 1997
). WW domains can either be subdivided into four classes according to their binding specificity (Bedford et al., 2000
) or into three subclasses according to sequence similarity (Macias et al., 2000
). 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., 1996
) and the FBP28 and YJQ8 WW domains (Macias et al., 2000
). 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., 1996
). 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., 1997
).
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Materials and methods |
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The structure of VP1-WW150 was modeled using the program Modeller 4 (Sali and Blundell, 1993) based on the published structures of VP1 at 1.9 and 3.65 Å (Stehle and Harrison, 1996
, 1997
). The inserted FBP11 WW domain was modeled using the NMR structure of the YAP WW domain (Macias et al., 1996
). Figures were generated by MolScript (Kraulis, 1991
) and Raster3D (Merritt and Bacon, 1997
).
Cloning and vector construction
The first FBP11 WW domain was genetically fused into the VP1 sequence as shown in Figure 2 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., 2000
).
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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., 1997; Schmidt et al., 2000
).
In vitro assembly and size-exclusion chromatography
Particles were assembled in vitro according to Salunke et al. (Salunke et al., 1986, 1989
). 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., 2000
).
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., 1992) 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) coppercarbon-grids according to standard protocols.
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Results |
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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, 1996). 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 1
.
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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 2). 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., 1997; Schmidt et al., 2000
) 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 3). The spectra of VP1 and VP1-WW292 had similar shapes, whereas the VP1-WW150 spectrum had a significantly increased negative ellipticity difference (
) 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 3
). 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., 2000
), 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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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., 1997). 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 I
), in good agreement with the value of 21 nM published earlier (Bedford et al., 1997
).
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However, VP1-WW150 showed a high affinity for the ligand (Figure 4). Using the same conditions as for GST-WW, the equilibrium dissociation constant KD of VP1-WW150 was increased to 7.7 ± 5 nM (Table I
). VP1-WW150 exhibited a high affinity for the proline-rich ligand in the range of KD = 415 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 I
). Association and dissociation reactions (represented by the association/dissociation rate constants ka and kd in Table I
) of VP1-WW150 were approximately 10 times faster than the respective values of GST-WW.
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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., 2000). 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., 2000). Analysis of the assembly of the VP1-WW proteins revealed that VP1-WW150 that still remained in solution assembled to only 15% (Figure 5
). 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 5
). 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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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 6). 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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Discussion |
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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., 1999). 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 I). 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 6
). 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, 1997
).
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., 2000). 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., 1999
).
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.
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Notes |
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4 To whom correspondence should be addressed. E-mail: ulis{at}ichr.uwa.edu.au
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Acknowledgments |
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Received June 30, 2000; revised June 11, 2001; accepted July 10, 2001.