Patent ID: 12209241

TABLE 1Masses of the different forms of Cp and kinase (SRPKΔ) used,as determined by ESI-MS mass spectrometryExpected mass (Da)Observed mass (Da)SRPKΔ45615.445614.7 ± 1.37PhosphorylatedCp18521995.421995 ± 0.71Cp18521395.321395.6 ± 0.86Cp14916852.316851.7 ± 0.06

TABLE 2Association of Alexa-Fluor-488 labelled PS1 with Cp.FluorescenceTotalPolarisationFluorescenceSample−RNase+RNase−RNase+RNasePS1 oligo7243.57363774102PS1 VLP1301283018733564PS1 + empty52.815.56933670672VLP

Anisotropy was used to determine if 15 nM of Alexa-Fluor-488 labelled RNA PS oligos can bind to, or enter, 125 nM of preformed shells of Cp. The latter were formed by reassembly in the absence of RNA at high concentration(3) (FIG.8c). Fluorescence polarisation values are influenced by the mass of the dye-labelled species(4). The polarisation value for PS1 oligo goes down following addition of RNase, as expected but remains unchanged when incorporated in VLPs assembled in the presence of the oligo. When labelled PS1 is added to the empty Cp VLP its fluorescence emission is unaffected, suggesting that it is not quenched, and it remains RNase sensitive confirming that it does not bind the outside of the protein shell or get internalised.

TABLE 3Sequence changes and corresponding assembly behaviour of PS1variant oligonucleotides, L1-5 and B1. Assembly behaviouris indicated as follows, the first “+” indicatesRNA-Cp binding, the second signifies formation of T =3/T = 4 sized species, and the third indicates RNaseprotection. “−” indicates failure in that assay.AssemblyRNA OligoLoopBulgebehaviourCommentPS1GGGAGGGGG+ + +L1UUUAUUGGG+ − −Loop G's are importantL2GUUAGGGGG+ − −Loop G's are importantL3UGGAUUGGG+ + −Loop G's are importantL4GGGUGGGGG+ + −Loop A is importantL5GGGGGGGGG+ + −Loop A is importantB1GGGAGGAAC+ + −Bulgesequence/structure isimportant

TABLE 4B3 sequence variants.Loop motif (left), full sequence (middle) andfolding free energy values (right) of the B3sequence variants.ΔG offoldingLoop(kcalmotifSequencemol−1)ACAAACAUGCAACAAUGCACAC (SEQ ID NO 10)−2.6AUUUACAUGCAAUUUUGCACAC (SEQ ID NO 11)−2.6UUUAACAUGCAUUUAUGCACAC (SEQ ID NO 12)−2.6GUUUACAUGCAGUUUUGCACAC (SEQ ID NO 13)−2.1UUUGACAUGCAUUUGUGCACAC (SEQ ID NO 14)−2.9AUUGACAUGCAAUUGUGCACAC (SEQ ID NO 15)−2.6GUUAACAUGCAGUUAUGCACAC (SEQ ID NO 16)−3.4GUUGACAUGCAGUUGUGCACAC (SEQ ID NO 17)−2.5AUUAACAUGCAAUUAUGCACAC (SEQ ID NO 18)−2.6

TABLE 5Analysis of suboptimal structures of RNA assembly cassettes. Mfold was used to foldeach cassette with a suboptimality setting of 500. These folds were then assessedby the following criteria: The presence of the correct -A.X.X.A- loop in PSs 1 through5 were verified and shown as a percentage (green = 60+, orange = 40+, red0-39 throughout the table). The nucleotide spacing between each stem loop was measured,compared to the expected value (FIG. 22) and also displayed as a percentage. Wherethese spacings differed, the maximum nucleotide difference is given.% of Correctly presented% of ‘correct’Maximum-A.X.X.A- motifsspacingsspacingCassettePS1PS2PS3PS4PS51-22-33-44-5varianceWT PS1-572080108000+7Synthetic1008710075739697672+7Stable PS1-5Stable PS1-5100879968315725566+7Unstable PS1-5989707327730067+7All PS310099100100100929010090+4PS1-37829N/AN/A00N/AN/AN/APS2-4N/A020N/AN/A00N/A+3PS3-5N/AN/A3446N/AN/A00N/A

TABLE 6Yield and Rhvalues from QELS experiments. Measured Rhvaluestaken from the midpoint of the main peak (20 min, FIG. 25)eluted from the TSKgel G6000PWxl column. Yields of genomicchimera reassemblies were calculated by integrating thearea under the main peak (20 min, FIG. 25) using the peakanalyser function in Origin Pro 9. Yields were then normalisedto the highest value and shown as percentages.RelativeSampleRhvalue/nmYield/%STNV-19.180Unstable PS1-5 + Δ1-127 STNV-18.940Synthetic, Stabilised PS1-5 + Δ1-1279.3100STNV-1

TABLE 7RNA oligonucleotide primersSEQTmPrimer NameSequence 5′-3′ID NO° C.ForwardGACATTAATACGACTCACTATAGGGACATGCA1965.5AUUArevGTGTGCATAATTGCATGTCCCTATAGTGAGTCG2068.2GUUGrevGTGTGCACAACTGCATGTCCCTATAGTGAGTCG2170.0AUUGrevGTGTGCACAATTGCATGTCCCTATAGTGAGTCG2269.5GUUArevGTGTGCATAACTGCATGTCCCTATAGTGAGTCG2369.5UUUArevGTGTGCATAAATGCATGTCCCTATAGTGAGTCG2468.2AUUUrevGTGTGCAAAATTGCATGTCCCTATAGTGAGTCG2568.2GUUUrevGTGTGCAAAACTGCATGTCCCTATAGTGAGTCG2669.5UUUGrevGTGTGCACAAATGCATGTCCCTATAGTGAGTCG2769.5ACCArevGTGTGCATGGTTGCATGTCCCTATAGTGAGTCG2864.3AAAArevGTGTGCATTTTTGCATGTCCCTATAGTGAGTCG2962.0AGGArevGTGTGCATCCTTGCATGTCCCTATAGTGAGTCG3064.0AUGArevGTGTGCATCATTGCATGTCCCTATAGTGAGTCG3162.9AGUArevGTGTGCATACTTGCATGTCCCTATAGTGAGTCG3262.5Unstable 1-5 forwardAGTAATACGACTCACTATAGGGGGGCTGCCCTC3362AAGGACCAGGGCAGAAAAGAGGAAAAGAAUnstable 1-5 templateGGCAGAAAAGAGGAAAAGAAAAGTGACAGAAC3462ACTTATAAGGAAATACACAAGTATAAGGAAAAAAGGAAGCTGCAATAGCGCAAGGAAUnstable 1-5 reverseTTCCTTTCCGAATTTTCGGATTCCTTGCGCTAT3562TGCAGCTTAll PS3 forwardGGGCCCCGCAACAATGCGGGGAAGGAAGGAA3665GGAAGAAAACGTACAAACGTTTTAll PS3 templateAGAAAACGTACAAACGTTTTAAGGAACAACGCA3765ACAATGCGTTGAAGGAAGGAAGGAAGGGGCGTACAAACGCCCCAAGGAATTTTAll PS3 reverseTTCCTTTTTTGCATTGTTGCAAAATTCCTTGGG3865GCGTTTGTACGCStable 1-5 templateGGCAGAAAAGAGGAAAAGAAAAGTGACAGAAC3962ACTTATAAGGAACCACACAAGTGGAAGGAAAAAAGGAAGCTGCAATAGCGCAAGGAASynthetic stable 1-5GGCAGAAAAGAGGAAAAGAAAAGTGACAGAAC4062templateACTTATAAGGAAAAAACGUACAAACGUUUUAAGGAAAAAAGGAAGCTGCAATAGCGCAAGGAAPS1-5 forwardAGTAATACGACTCACTATAGGGAGTAAAGACAG4162GAAACTTTACTGACTAACATGGCAAAACPS1-5 templateACTGACTAACATGGCAAAACAACAGAACAACAG4262GCGAAAATCCGCAACAATGCGTGCAGTGAAGCGCATGATAAATACACPS1-5 reverseTCAGTGCAAACCTTTTATGCTCCAAGTGTGTAT4362TTATCATGCGCTPS1-3 forwardAGTAATACGACTCACTATAGGGAGTAAAGACAG4461GAAACTTTACTGACTAACATGGCAAAACPS1-3 templateACTGACTAACATGGCAAAACAACAGAACAACAG4561GCGAAAATPS1-3 reverseCGCATTGTTGCGGATTTTCGCCTGTTGT4661PS2-4 forwardAGTAATACGACTCACTATAGGGTGGCAAAACAA4758CAGAACAACAGGCGAAAATPS2-4 templateAACAGAACAACAGGCGAAAATCCGCAACAATGC4858GTGCAGTGAAGCGCATGATAAATAPS2-4 reverseCCAAGTGTGTATTTATCATGCGCTTCACTGCAC4958GCATTGTTGCGGPS3-5 forwardAGTAATACGACTCACTATAGGGCCGCAACAATG5061CGPS3-5 templateCCGCAACAATGCGTGCAGTGAAGCGCATGATA5161AATACACPS3-5 reverseTCAGTGCAAACCTTTTATGCTCCAAGTGTGTAT5261TTATCATGCGCT

Materials and Methods

Cloning, Expression and Purification of Proteins Used.

We obtained anE. coliCp-expressing plasmid (a gift of Prof. Nicola Stonehouse), known to produce assembled HBV VLPs containing host RNAs(5). The Cp encoded has the following amino acid sequence differences compared to the current GenBank reference strain (NC_003977.2): A61, E77-FAGAS (single letter amino acid code)-D78 insertion, S92N, F102I, I121L, R156-RD-R157 insertion. Since the wild-type C61 has been implicated in assembly (6), this was restored to the gene before expression in a PET28b plasmid in BL21 (DE3)E. colicells. The inserted FAGAS epitope was also removed. Induction with 1 mM IPTG at 0.6 OD was followed by growth for 20 hrs at 21° C. Cells were lysed using a Soniprep 150 with 5× 30 sec bursts on ice. The lysate was then clarified by spinning at 11,000 g for 1 hr. VLPs were then pelleted by centrifugation at 120,000 g for 14 hr, resuspended in 20 mM Hepes (pH 7.5), 250 mM NaCl, and 5 mM DTT and applied to an XK50 column packed with 25 ml of Capto™ core 700 resin (GE Life Sciences). Fractions containing VLPs were pooled and precipitated with 40% (w/v) ammonium sulphate. The Cp appeared pure on SDS-PAGE and its identity, and that of variants, was confirmed by mass spectrometry (Table 1). Cp lacking the ARD, i.e. Cp149, was produced by mutagenesis (Q5 site-directed mutagenesis kit, NEB) and prepared similarly. Note, the Cp149VLP expressed inE. colilacks significant encapsidated cellular RNA. VLPs were visualised by negative stain transmission electron microscopy (TEM). Full length Cp VLPs were additionally purified by sucrose density gradient before dye-labelling using Alexa Fluor®-488 SDP ester fluorophore (Invitrogen) over 4 hrs at room temperature in 200 mM sodium carbonate buffer (pH 8.3), followed by desalting over a NAP5 column. There were two over-lapping VLP peaks on the gradient and it was impossible to separate them. TEM and smFCS confirm that they are the expected T=3 and T=4 shells, with the latter the predominant form (FIG.7a). The Cp region 140-148 has been shown to be a determinant of morphology, the shorter versions producing more T=3 shells (7). It is possible that the dipeptide insertion adjacent to the linker region at position 157 may alter the properties of the Cp. However, when we removed the RD insertion, yielding Cp183, we found no differences with Cp185, either in RNA binding, ability to form VLPs with PS RNAs or preference for the dominant quasi-conformer shell formed. Since longer Cp was used for SELEX and the high resolution EM work, those are the data shown throughout.

All HBV variants used for assembly assays were dissociated from VLPs into protein dimers as previously described(3), with the exception that dissociation was at pH 9.5, as opposed to 7.5. This was done in the presence of Complete Protease Inhibitor Tablets (Thermofisher Scientific). HBV core dimer concentration was determined by UV absorbance. Fractions with an A260:A280ratio of approximately 0.6 or lower were used in assembly assays. SRPKΔ kinase was expressed and purified from a pRSETb plasmid, as previously described(8).

SELEX Protocol

Purified HBV capsids (˜360 μg) were immobilised onto 6 mg of M270 carboxylic acid Dynabeads® microspheres (Thermo Fisher Scientific) following the manufacturer's protocol. Beads were washed twice with selection buffer (25 mM Hepes, pH 7.5, 250 mM NaCl, 2 mM DTT, EDTA-free complete protease inhibitor) and unreacted N-hydroxysuccinamide blocked with a 15 m 50 mM Tris-HCl pH 7.4 wash. Beads were washed a further three times with selection buffer. Immobilised capsids were dissociated with a 30 minute incubation of 2 M guandinium chloride in 0.5 M LiCl2. Beads were then washed three times with B&W buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 2 M NaCl) and then washed three times with selection buffer. Beads were resuspended in selection buffer so that concentration of beads was 10 mg/mL. Negative selection beads were also prepared in the same manner but with no capsids. Ten rounds of SELEX were performed in vitro using a synthetic, combinatorial N40 2′OH RNA library (˜1024potential sequences) as described previously (9). The amplified DNA of round 10 was then subjected to Next Generation Sequencing on an Illumina MiSeq® sequencing-platform. This yielded ˜1.6M sequence reads, in which one sequence occurs 65,802 times and there are 1149 aptamers with a multiplicity of 100 or higher. The overall frequencies of the four nucleotides in this aptamer pool is A34.30%; C9.09%; G40.97% & U15.64%, and compares with the same data for the unselected naïve library of A26.10%; C22.03%; G24.64% & U27.22%. The highest multiplicity for sequences in the latter pool is 4. These data confirm that selection from the naïve pool occurred, and that the base composition of the selected aptamers is consistent with the RGAG motif identified within the HBV genomes.

PS Identification

PS identification was carried out using the laboratory HBV strain (*NC_003977.1). The aptamer library contained 1,664,890 unique sequences, each 40 nts in length that have been aligned against the genome as follows: Each aptamer sequence was slid along the genome in increments of 1 nt. For each such position of the reference frame, the subset of the aptamer sequence with the best alignment to the genome was identified according to the Bernoulli score B, which benchmarks the probability of a non-contiguous alignment to that of a contiguous alignment of B nucleotides. The Bernoulli scores for all reference frames of a given aptamer sequence in the library were rank-ordered starting from the largest score, and all matches with the genome up to a Bernoulli score of 12 counted. The procedure was then repeated for the other aptamer sequences and corresponding matches added, resulting in the peaks inFIG.2a.

Identification of a Consensus Motif

HBV genome sequences with the following accession numbers were randomly extracted from 750 complete HBV genomes found in GenBank: KCS10648.1; *AF223955.1; AY781181.1; *AB116266.1; AB195943.1; KR014086.1; *KR014072.1; KR014055.1; KR013939.1; KR013921.1; KR013816.1; KR013800.1; EU796069.1; AB540582.1, and the NCBI HBV reference strain (GenBank Seq ID *NC_003977.2) and the laboratory strain (GenBank Seq ID NC_003977.1) were added to the ensemble. Sequences used for the statistical analysis inFIG.2care marked by an asterisk. Bernoulli peaks, which occurred within at most 10 nts of each other in at least 80% of these 16 HBV strain variants, were marked by a green cross inFIG.2ato indicate their conservation. To identify the putative PS recognition motif, we extracted sequences of 60 nts, centred around the peak nucleotide of each Bernoulli peak, from three representative strains (AF223955.1, NC_003977.1, & NC_003977.2) and determined all possible stem-loops of negative free energy via Mfold(10). We carried out a similarity analysis of these stem-loops, comparing both sequence and structure elements, we identified for each peak area that representative that has the highest degree of similarity both with secondary structure elements in the other peak areas in the same genome and stem-loops corresponding to the same peak area in the other strains. This returned a stem-loop for each peak. An alignment of the corresponding loop sequences is shown inFIG.2b.

RNA Dye-Labelling

PS1, PS2 and PS3 (47 nucleotides long) were purchased from Integrated DNA Technologies with a 5′ C6-amino group. To label RNA, 6 μL of RNA (200 μM) was mixed with 1 μL 1 M sodium borate buffer, pH 8 and 3 μL 10 mM Alexa Fluor®-488-SDP fluorophore (Thermo Fisher Scientific) and rolled at room temperature for 4 hours. 10 μL of 2× denaturing loading dye was then added to the RNA, boiled for 5 minutes and loaded onto a pre-warmed denaturing PAGE. RNA was gel extracted, isopropanol precipitated and finally re-suspended in DEPC-H2O and frozen at −80° C. until needed.

Assembly Assays

Assembly reactions were performed by adding HBV Cp in dissociation buffer (50 mM Tris (pH 9.5), 1.5 M GuHCl, 500 mM LiCl and 5 mM DTT) to 15 nM Alexa Fluor®-488 fluorophore labelled RNA in a reassembly buffer containing 20 mM Hepes (pH 7.5), 250 mM NaCl, 5 mM DTT and 0.05% (v/v) Tween®-20 polysorbate emulsifier at 25° C. Successive additions of dimer were performed until assembly was deemed complete by the measured Rhvalue plateauing, but never exceeded 10% of total reaction volume. Each addition of Cp is marked by a vertical dashed grey line in the titration plots and the expected hydrodynamic radii of T=3 and T=4 particles (as determined for dye-labelled particles expressed inE. coli) are marked by an orange horizontal dashed line within figures.

Manual mixing throughout the reactions caused an approximate 1 min delay at the start of FCS data collection. FCS measurements were made using a custom-built FCS setup with 30 sec data accumulation per autocorrelation function (CF). Individual CFs were decomposed into triplet state relaxation and diffusion (characterized by diffusion time, TD) components, and the latter was converted into an apparent hydrodynamic radius, Rh(11). Samples for TEM were taken at the end of each measurement. Plots of Rhover time (thin dashed line) were smoothed (thick solid line) using the FFT filter in Origin Pro-8 with a cutoff percentage of 35%. Plots of Rhdistribution were also fitted using Origin Pro-8 software, to a normal single or multiple peak Gaussian function. Samples taken for negative stain TEM analysis were placed on to a glow discharged carbon coated formvar 300 mesh Cu grid. Grids were stained with 2% uranyl acetate and dried.

Assembled Particle Labelling

Assembly was carried out as in smFCS experiments. In particular, Cp was titrated into reassembly buffer with and without 15 nM unlabelled PS1 to a final concentration of 250 nM. This was allowed to incubate at room temperature for 1 hour, and then buffer exchange was carried out via dialysis to remove guanidinium hydrochloride present. Labelling of protein was then carried out by adding Alexa Fluor®-488 SDP ester fluorophore (1:50 ratio of dye to Cp dimer) and incubating overnight at 4° C. The resulting sample was then measured via smFCS in 30 s bins for 100 min and the Rhdata plotted as above in a hydrodynamic radial distribution plot. A sample was then removed for analysis via TEM. Post labelling, Cp dimer became assembly incompetent, therefore Cp could not be tracked during real time assembly.

Photobleaching

HBV VLPs containing Alexa-488 labelled PS1 were assembled as described in smFCS assembly assays. Under those conditions all RNA is bound to protein as judged from fluorescence quenching and photon counting in the FCS experiments. VLPs were then added to two glow discharge-irradiated Carbon/Formvar 300-mesh grids (Agar Scientific), and one grid stained with 2% (w/v) uranyl acetate and viewed with a Jeol 1400 microscope at 40,000× magnification. The remaining, unstained grid was positioned Formvar side down onto a clean microscope coverslip and mounted onto an inverted TIRF microscope. The laser (Coherent Sapphire, 488 nm, 25 mW) power was adjusted to excite and photobleach the labelled RNA within the time frame of several minutes. Sequential images were taken with an emCCD camera (Andor iXon) with 0.2 sec exposures and em gain of 200. An unexposed field of view was used for each series.

Fluorescent spots were identified in the collected frames using previously described procedures and converted into time traces(12). These were then inspected and classified according to the number of photobleaching steps. Frequencies of traces with a defined number of steps were collated in a histogram. Several bright spots per field of view exhibited continuous intensity decay, presumably representing larger aggregates. These were used to estimate the overall photobleaching rate (0.003 per frame) and formally included in the histogram as representing 10 steps. The histogram without the bin representing continuum events was modelled as a weighted sum of binomial distributions for up to quadruple occupancy and probability of labelling of 0.56 estimated from UV-Vis spectra.

Electron Microscopic Reconstructions

Large Scale VLP Preparation

smFCS experiments were scaled up into 96 well plates. Two 96 well plates (Non-Binding Surface, Corning) were used. PS1 RNA was labelled and gel purified as described earlier and HBV dimer was purified as described above. Each well contained 200 μL of 15 nM PS1 in reassembly buffer. As in smFCS, ten 2 μL injections of 2.5 μM dimer in dissociation buffer were performed. A Perkin-Elmer Envision plate reader was used to carry out the injections and record the anisotropy of the PS1 RNA (FITC excitation and emission filters). VLPs were purified away from free RNA and capsid using a 1.33 g/mL caesium chloride gradient and spun at 113,652×g for 90 hours using an SW40Ti rotor. A single band was observed and fractionated. The band was dialysed into reassembly buffer to remove caesium chloride. The 2 mL fraction of VLP was concentrated to 200 μL using an Amicon 100 kDa MWCO spin concentrator.

CryoEM Specimen Preparation

After recovery of the PS1-containing VLPs and removal of caesium chloride by dialysis, their structures were analysed using single-particle cryo-EM. VLPs were vitrified. 200 mesh EM grids with Quantifoil R 2/1 support film and an additional ˜5 nm continuous carbon film were washed using acetone and glow discharged for 40 s prior to use. CryoEM grids were prepared by placing 3 μl of ˜3.2 mg/ml HepB VLP on the grid, before blotting and plunge freezing using a Leica EM GP freezing device. Chamber conditions were set at 8° C. and 95% relative humidity, with liquid ethane temperature at −175° C. Data was collected on a FEI Titan Krios (eBIC, Diamond Light Source, UK) transmission electron microscope at 300 keV using an electron dose of 27 e−/Å2/s, 2.5 s exposure, yielding a total electron dose of 67.5 e−/Å2. Data was recorded on a 17 Hz FEI Falcon 11 direct electron detector. The dose was fractionated across 33 frames. Final object sampling was 1.34 Å per pixel. A total of 2397 micrographs were recorded using EPU (FEI) automated data collection software.

Single Particle Image Processing

2397 micrographs were motion corrected and averages of each movie were generated using MotionCorr(13), and contrast transfer function (CTF) parameters for each were determined using CTFFIND4(14). Micrographs with unacceptable astigmatism or charging, as determined by examining the output from CTFFIND4, were discarded leaving a total dataset of 1710 micrographs. All particle picking, classification and alignment was performed in RELION 1.3(15).

Approximately 57,000 particles were manually picked and classified using reference-free 2D classification in RELION 1.3. This classification confirmed the initial visual impression that although the VLPs were purified as a single band on a caesium gradient, two sizes of VLPs were present. A selection of resulting 2D class averages were used as templates for automated particle picking. The particle stack generated using auto-picking was subject to 2D classification to separate T=3 and T=4 particles, and to remove particles not corresponding to VLPs. The subsequent particle stacks (5589 for T=3, 42,411 for T=4) were subject to 3D classification, using a sphere with the approximate diameter of the VLP as a starting model. Subsets of the data were reconstructed including data out to the Nyquist frequency using the 3D autorefine option in RELION with 13 symmetry imposed to generate all structures presented in this work. Within the T=4 42,411 particle dataset it was clear that a further subset (10,851 particles) of the data contained a significant asymmetric feature inside the Cp shell where RNA binding would be expected to occur. An asymmetric (C1) reconstruction was performed on a relatively homogenous set of 10,851 such particles, giving the reconstruction at 11.5 Å resolution.

The 3D model of PS1 RNA was made using RNA Composer(16). The cryoEM figures were rendered using USCF Chimera(17).

Purification of Recombinant STNV CPs

Recombinant STNV VLPs were purified fromE. coli(18). STNV charge-change mutant plasmids were created using primers designed using Agilent, and a Quikchange site directed mutagenesis kit (Agilent). CP monomers were purified by disassembly in 50 mM Tris (pH 8.5), 10 mM EDTA, in the presence of Complete Protease Inhibitor Cocktail (Roche, United Kingdom). STNV CP was separated from the mRNA by sequential Q-Sepharose, and SP-Sepharose columns (GE Healthcare, Sweden). STNV CP was washed with 20 column volumes of 50 mM HEPES (pH 7.5) and 25 mM NaCl to remove residual EDTA, and subsequently eluted using a 0.025-2 M NaCl gradient in buffer. CP elutes at 0.8 M NaCl. STNV CP was analysed by SDS-PAGE and its concentration determined by UV absorbance. Fractions with an A260:A280 ratio of 0.6 or lower were used in assembly assays. Mutant CPs that did not form VLPs during overexpression were purified using the same sequential Q-Sepharose and SP-Sepharose columns method.

Preparation of RNA Oligonucleotides

dsDNA transcripts encoding the RNA oligonucleotides used in this study were produced using primers and the KAPA2G system (KAPA biosystems) following the manufacturer's protocol. Transcriptions were carried out using the HiScribe® T7 High yield RNA synthesis kit (NEB). Products were run on a denaturing RNA gel. The Alexa Fluor®-488 fluorophore labelled B3 oligonucleotide used throughout was synthesised and HPLC purified by DNA Technology A/S (Denmark). Other RNA oligonucleotides requiring a 5′ fluorophore were labelled with an amino GMP during transcription and cross linked to an Alexa Fluor®-488 SDP ester fluorophore (Invitrogen) prior to gel purification as described previously (19).

Genomic chimeras were created by purchasing Gene blocks of the Synthetic, stabilised and Unstable+Δ1-127 STNV-1 constructs with a 5′ T7 promoter (Integrated DNA technologies), possessing BamHI and HindIII cleavage sites at either end to create sticky ends after restriction digestion and dephosphorylation using Antarctic phosphatase (NEB). This gene block was then ligated into a PACYC184 plasmid using T4 DNA ligase (NEB). Transcription was carried out as above after linearization using BamHI.

RNA was annealed prior to each experiment by heating to 80° C. for 90 s and cooling slowly to 4° C. in a buffer containing 50 mM NaCl, 10 mM HEPES and 1 mM DTT at pH 7. Genomes were only heated to 65° C.

STNV Reassembly in the Presence of B3 Variants and Sedimentation Velocity Analytical Ultracentrifugation (svAUC) Reassembly reactions were carried out in the presence and absence of B3 variants in a 1:3 RNA:CP ratio at a final CP concentration of 4.5 μM, by dialysis into a buffer containing 50 mM HEPES (pH 7.5) and 2 mM Ca2+. All samples were analysed by TEM and AUC. For AUV, 0.32 mL of each sample was placed in a 1.2 cm path length 2-sector meniscus matching epon centrepiece cell constructed with sapphire windows. The samples were centrifuged at 15,000 rpm in an Optima XL-1 analytical ultracentrifuge at 20° C. in an An50-Ti rotor. Changes in absorbance at 260 nm were detected by absorbance optics with 100 scans taken in approximately 11 hrs 30 min. Data were fitted and analysed using the program Sedfit.

smFCS Data Collection and Analysis

FCS measurements were performed on a custom-built smFCS facility. Excitation laser (Sapphire CW blue laser, 488 nm, Coherent, USA) power was set to 65 μW. The focus position was adjusted to 20 μm from the cover slip inner surface (maintained by piezoelectric feedback loop, Piezosystems Jena, Germany). Immersion oil (refractive index 1.515, type DF, Cargille Laboratories, USA) was used with immersion oil objective (63× magnification, numerical aperture 1.4). The photon count was recorded and analysed by an ALVL5000 multiple tau digital correlator (ALV-GmbH) in single channel mode. FCS data was analysed using non-linear, least-squares fitting with a single component diffusion model autocorrelation function corrected for the triplet state in Matlab® software. Diffusion time was used in the calculation of apparent hydrodynamic radius (Rh) and plotted as a function of assembly time. Rh calculations were based on the measured diffusion time for Alexa Fluor®-488 dye with the estimated Rh of the dye (=˜0.7 nm in assembly buffer).

smFCS Assembly and Competition Assays

Initial measurements of Alexa Fluor®-488 fluorophore labelled RNA oligonucleotides were taken for at least 10 runs of 30 secs (5 min). Purified STNV CP was titrated into labelled RNA. Each titration was measured for a minimum of 10 30 secs runs. In assembly assays this was repeated until full capsid assembly had occurred. At this point RNase A was added to confirm RNA protection. In competition assays, once the sample had formed a capsomer structure (Rh=˜5 nm) the sample was monitored for a further 120 runs of 10 secs (20 min to ensure stability). At this point unlabelled B3 short/B3 variant competitor was added in 100-fold molar excess and measured for 120 runs of 10 secs.

CD Analysis

Transcribed oligonucleotides were diluted to 1.5 μM in 300 μl, in a buffer containing 10 mM MES, 50 mM NaCl and 1 mM DTT at pH 6. Measurements were performed on a Jasco J715 spectropolarimeter, from 200 to 350 nm, with a bandwidth of 2 nm. Each Ca2+ and STNV titration was inverted 5 times and allowed to reach equilibria for 2 min prior to the next measurements. Thermal denaturations were performed using a Peltier temperature control from 10-95° C. in 5° C. steps, and an end scan was performed at 10° C. to check for cleavage. Each measurement was performed in triplicate and averaged. Data was converted to molar ellipticity using the equation: Δε (cm2 mM-1)=θ/(32980 C(mM) L(cm) N(no. of nt)).

Light Scattering Assay of Reassembly with Genomic RNA Variants

Reassemblies were performed with genomic chimeras in a 96 well plate as in the smFCS assays, with 1 nM genome and CP titrated in until a final concentration of 400 nM STNV CP was reached. This was concentrated through a 100 kDa Centricon® concentrator (Millipore) at 10k xg for 5 min and run on a TSKgel G6000PWxl SEC column (Tosoh) with an AKTA PURE® chromatography system (GE Healthcare) connected to a DAWN® multi-angle static light scattering detector, HELEOS® photometer, Optilab® TrEX refractometer for QELS and refractive index measurements. The column flow-rate was 0.4 ml min-1 for 50 min. Peaks were fractionated, A260/280 ratios measured and EM images obtained (FIG.25). The yield of the Unstable PS3 sample, as calculated by integration of light scattering signal, is dramatically lower than that of the wild-type STNV RNA, whilst the stabilised synthetic cassette results in a significantly higher (>20%) yield in VLP compared to the natural sequence. QELS estimates the Rh values similar to those from smFCS, 9.3±0.1 nm, 9.1±0.3 nm and 8.9±0.2 nm for Synthetic, stabilised PS1-5+Δ1-127STNV-1, wild type STNV-1 and Unstable PS1-5+Δ1-127STNV-1, respectively (Table 6). The A260/280 ratios of the assembled VLPs eluting from the gel filtration column are also informative. Both the PS1-5+Δ1-127STNV-1 and the wild-type STNV-1 genome samples have identical values (1.62), whilst the Unstable PS1-5+Δ1-127STNV-1 sample has a higher value (1.89). This is consistent with there being a constant amount of RNA in the first two samples fully enclosed in shells containing the same number of CPs, whilst the final sample has the same RNA content in an incomplete shell.

EXAMPLE 1

The HBV pgRNA Contains Preferred Cp Binding Sites

HBV VLPs assembled from (full-length) Cp subunits expressed inE. coliwere purified as described(3) (FIG.7a& Table 1). They form a mixture of T=3 and predominantly T=4 shells. These were immobilised onto magnetic beads, disassembled by treatment with guanidinium chloride and then washed to remove host RNA, resulting in immobilised Cp dimers(20) with their ARDs accessible. RNA SELEX was carried out using our standard protocols (FIG.7b) and the aptamer pool from the 10th round analysed by NextGen DNA sequencing (Methods).

The RNA sequences that bind Cp in the selected library were aligned to the HBV pre-genome most closely related to the protein used for the SELEX experiments (the laboratory strain, GenBank Seq id NC_003977.1 (21)). Statistically significant matches (a Bernoulli score of 12 or more, Methods) to the pgRNA of this strain (the blue peaks inFIG.2a) were benchmarked against an alignment of the unselected library (grey curve inFIG.2a) to identify peaks that occur with significant frequency. This identifies multiple sites dispersed across the pgRNA having similar sequences/structures to Cp binding aptamers, consistent with our expectation for PS-like sites across the genome. We applied the same procedure to 14 randomly selected HBV strain variants from GenBank, the current NCBI HBV reference strain (GenBank Seq ID NC_003977.2) as well as the laboratory strain (GenBank Seq ID NC_003977.1) and identified all those peaks that are conserved in at least 80% of these strains (marked with green crosses inFIG.2a). These genomic regions are thus likely to encompass PSs. The three peaks with the highest conservation (100%) and peak heights, the latter indicating how many aptamers matched these sites, are labelled PS1, PS2 and PS3 inFIG.2a. For the nine sites with high conservation between strains, we extracted 30 nts 5′ and 3′ to the peak nucleotide in the genomic sequences of three representative strain variants, including the laboratory strain and the reference genome, and considered all their possible secondary structure folds with negative free energy via Mfold (Methods). A similarity analysis of primary and secondary structure revealed the predicted existence of stem-loops sharing a purine-rich loop recognition motif, RGAG (FIG.2b).

We computed the frequency of this motif in stem-loops across the 16 HBV strains analysed. Across all strains, the RGAG motif occurs in stem-loops on average ˜25.4 times (precisely 25 times in the laboratory strain). Compared to 10,000 randomised versions of the pgRNAs, the frequency of occurrence of RGAG in the actual genome is 4.68 standard deviations above the average (FIG.2c), strongly implying a functional role(s).

EXAMPLE 2

pgRNA Oligonucleotides Trigger VLP Formation In Vitro

PS1, 2 & 3 oligonucleotides (FIG.8a), were tested for their ability to bind Cp dimers using single molecule fluorescence correlation spectroscopy (smFCS) (FIGS.3&8b). This technique yields a real time estimate of the hydrodynamic radius (Rh) of dye-labelled species. Importantly, it allows reactions to be followed at low nanomolar concentrations, where we have shown that binding specificity more closely reflects the situation in vivo compared to most in vitro reactions. The latter are typically carried out at higher (e.g. 0.1-0.8 μM) concentrations(20), where the specificity of PS-mediated assembly is reduced or lost. In order to avoid electrostatic effects due to differing oligo lengths, each PS was produced as part of a 47 nt long fragment, each dye-labelled at its 5′ end (Methods(19)). The labelled oligos (˜15 nM) were then titrated with increasing amounts of Cp (5-250 nM Cp dimer) and the Rhvalues tracked over time (FIG.3a). After each addition there was a pause of ˜10 min to allow reactions to equilibrate. The titrations lead to distortions in the data collection and the averaging, which is visible in the plots as noisy signals. After equilibration at 250 nM Cp, RNase was added to each reaction and the Rhvalues monitored for ˜10 min. If these declined steeply, it was assumed that the VLPs produced were incomplete. Negative stain EM images were obtained for the samples before RNase addition, and the sizes of the complexes present at this point were also assessed by calculation of Rhdistribution plots (FIG.3bandFIG.8c, respectively).

Each of the PS fragments stimulates assembly of both T=3 and T=4 complete VLPs with roughly equal efficiency under these conditions (FIGS.3a&b), with the latter being the dominant product, as expected(22). Addition of Cp>250 nM does not increase the Rhvalues obtained, implying that by this stage all the RNAs have been incorporated into VLPs. In order to assess whether these effects are a direct consequence of Cp-PS interaction, we carried out a number of controls. Dye-labelled PS fragments do not bind to preformed VLPs and remain RNase sensitive in their presence (Table 2), implying that the PSs only get internalised in assembling VLPs. To determine if the RNA triggers assembly, we compared assembly efficiency of Cp with and without PS RNA present by adding a protein modifying dye after incubation of Cp alone or completion of a titration of unlabelled PS1. The Rhdistribution plots are shown inFIG.3b. In the absence of RNA, <5% of Cp assembles under these conditions, in contrast to >80% of the Cp for assembly in the presence of RNA. It appears that Cp-PS interaction triggers an increase in the assembly efficiency. This effect varies with the age of the Cp, consistent with oxidation of an assembly-inhibiting disulphide at the dimer interface(6). Comparative statements here are based on the results of both positive and negative control experiments with each batch of Cp.

We then probed the RNA sequence-specificity of these reactions (FIG.9a). Test oligos comprised the epsilon stem-loop, as well as loop and bulge variants of PS1. This included a variant in which the bulge region was fully base-paired. In similar assays to the PS1-3 reactions the Rhvalues for all three RNAs remain sensitive to nuclease action, implying that assembly of closed shells requires a specific RNA sequence/structure. EM images and distribution plots confirm this interpretation. The sequence sensitivity of the assembly reaction is further highlighted by additional PS1 variants (FIGS.9b&c; Table 3). Their effects on assembly confirm the importance of the bulge and/or sequences within it, and the loop RGAG (here a GGAG) motif. A DNA oligonucleotide encompassing the PS1 sequence (FIG.9d) elicits aggregation, showing that faithful assembly is a specific property of the PS in its RNA form, i.e. with an A helical duplex stem, as well as the Cp-recognition motif in the loop.

The C-terminal ARD of the HBV Cp is believed to mediate interactions with the pgRNA, and the 1-149 Cp fragment that lacks the ARD readily assembles in the absence of nucleic acid(23). We therefore assessed the ability of Cp149to respond to PSs in the smFCS assay. No RNA-dependent assembly, or PS binding by Cp149, occurs under these conditions (FIG.10a), although EM images show that the truncated Cp alone readily assembles, confirming that the ARD is essential for the interaction with RNA. The ARD is extensively phosphorylated in vivo, although the responsible cellular kinase remains unknown(24). Lowering the positive charge on the C-terminus of Cp should reduce its ability to bind PS RNAs. We phosphorylated Cp in vitro(8) (Table 1) and tested its properties. EM images show that modified Cp readily assembles but does not bind to PS1 in smFCS assays (FIG.10b).

EXAMPLE 3

HBV NC Assembly is Triggered by Formation of a Sequence-Specific RNA-Core Protein Complex.

The VLPs assembled around PS1 were purified on a larger scale and their structures determined by cryo-EM, yielding icosahedrally-averaged reconstructions of the T=3 and T=4 particles (FIG.4). A significant fraction (˜25%) of the T=4 particles also contained an asymmetric feature located just below the protein shell. An asymmetric reconstruction of these particles was also calculated (FIG.5). The result suggests the asymmetric feature represents a complex between PS1 oligonucleotides and the ARD domains of the overlying Cp subunits.

From the EM map at this resolution it is not possible to determine the number of PS oligonucleotides present in the complex. The A260/280ratio of the purified VLP suggests that the RNA content, assuming T=4 morphology, is ˜5 oligos/particle(25). An additional estimate of this stoichiometry was obtained by studying photobleaching of PS1 VLPs (FIG.4, Methods). VLPs show multiple bleaching steps, confirming that there are multiple oligos within each shell. Given the labelling efficiency of the oligos, the data are consistent with 2-4 oligos/VLP. We built a 3D model of PS1 and manually positioned it within the EM map (FIG.4f, Methods). From the relative volume of the asymmetric density and the size of the PS1 oligo, it appears that at least two copies of the PS are present within the density. We cannot exclude the possibility that other RNA molecules are bound to the protein shell elsewhere, but are not visible due to mobility or an irregular location with respect to the ordered RNA density. The biochemical and structural data are consistent with the asymmetric structure being an assembly initiation complex, where an RNA preferred site(s) has initiated assembly culminating in the formation of the T=4 NC.

The cryo-EM data hint at a further insight into HBV biology. A minority of HBV particles, whether from assembly reactions or wild-type virus infections, assemble with T=3 quasi-symmetry and both types of particles are visible in our cryo-EM data. Using 2D and 3D classification the T=3 (˜11%) and T=4 (89%) particles are readily separable.FIG.4shows 3D reconstructions of the two particles with imposed icosahedral symmetry at 5.6 Å and 4.7 Å resolution, respectively. In addition to the obvious differences in size and number of Cp dimers that the two VLP structures contain, the T=4 and T=3 maps are different in the features visible on their inner surfaces, where the ARDs are located and where RNA binding occurs. As might be expected for icosahedrally-averaged maps of a sub-stoichiometrically occupied VLP, both structures are essentially devoid of density attributable to RNA. The capsid shell of the T=4 structure is visibly thinner than the T=3 equivalent, however, and closer examination of the T=3 map suggests that additional density corresponding to ordered segments of the ARDs is visible (FIG.4), which is absent in the T=4 structure (FIG.4c& d). This difference persists when the T=4 map is Fourier filtered to be at a similar resolution as the T=3 (FIG.11). This is consistent with previous studies that showed that the Cp C-terminal region, including the ARD, plays a role(s) in determining capsid geometry(22, 26).

EXAMPLE 4

Sequence-Specific Recognition of Individual PS Sites

There are multiple consequences of sequence-specific RNA-CP recognition in the STNV system (FIG.12). Titration of CP into oligonucleotides encompassing only PS3 (or B3) initially results in formation of a trimeric capsomer (Rh ˜5 nm), followed by formation of T=1 VLPs (Rh˜11.3 nm) as the CP concentration is raised gradually. Rh distribution plots of the smFCS data at the end of the titration suggest that the VLPs formed are homogeneous, whilst electron microscopy images (EM) and RNase challenge assays suggest that they are composed of complete protein shells. A similar titration with a PS3/B3 variant having a loop sequence of -U.U.U.U-, showed that CP binds such SLs, but the complex formed is unable to assemble to VLPs(19). The natural 127-mer, encompassing PSs1-5, shows more complex behaviour. Addition of low CP concentrations triggers a collapse in its Rhby about 20-30%, mimicking the behaviour seen for the full length genome(27). Subsequent CP additions result in co-operative conversion to T=1 VLPs with the same properties as those formed around PS3 alone. PS sequence variants within this fragment confirm that -A.X.X.A- is a CP recognition motif and its presence is only absolutely required in PS3, however the variants no longer assemble with wild-type co-operativity(19). STNV-1 CP alone shows no tendency to aggregate below 15 μM under these conditions, and therefore everything in the titrations shown here is a consequence of RNA-CP binding.

These results highlight the importance of PS3 recognition by CP for assembly. In order to identify the critical features of that recognition, we produced a series of SLs encompassing variant loop sequences with the PS3 stem (FIG.17& Table 4). The variants have altered nucleotides in the “inner” two positions (-C.C-; -A.A-; -G.G-; -G.U- & -U.G-) compared to the wild-type -C.A- of PS3. “Outer” variants (-A.U.U.A-; -A.U.U.G-; -G.U.U.A-; -G.U.U.G-; -G.U.U.U-; -U.U.U.G-; -U.U.U.A- & -A.U.U.U-), in which both inner nucleotides were altered to uridines, were also tested. Our expectation was that there would be no base specificity at the middle positions while the adenines would be preferred at the first and last positions of the tetraloop. We examined their abilities to support assembly of both the T=1 shell and the trimeric capsomer. Test RNAs and CP were mixed at ˜5 μM concentrations in reassembly buffer and the results assayed by velocity sedimentation analysis and in EM images. Under these conditions, the inner nucleotide variants form T=1 capsids with roughly similar efficiency as PS3, confirming that their identities are not part of the CP recognition motif (FIGS.13A/B &FIG.17). The outer nucleotide variants showed differing behaviour, with only the -A.U.U.U-, -U.U.U.A- and -A.U.U.A- variants having a peak in a similar position to PS3, confirming that the outer adenines are part of the CP recognition motif.

In order to examine their relative importance for CP affinity, we adapted the smFCS assay (FIG.13B). Labelled B3 was titrated with CP to form the trimer, as judged by the Rh value, and then a 100-fold molar excess of each sequence variant was added to compete off the B3. Variants that do not bind with a similar affinity to B3 fail to displace the labelled RNA, whereas B3 and other variants outcompete the labelled species restoring the Rh of CP-free RNA. The results (FIG.13D) show the percentage Rh change following this challenge, revealing a wide variation in the ability of the variant RNAs to compete. All those with guanine substitutions, and the -A.U.U.U- variant, fail to compete. The superior performance of the -U.U.U.A- variant suggests that either the 3′ A is the most important for CP recognition, or that the A-U base pair at the top of the adjacent stem breaks, presenting an -A.U.U.U.A.U- variant of the B3 motif that is still recognised by the CP. Either way, -A.X.X.A- outperforms all variants, suggesting that SLs carrying tetraloop motifs of -A.X.X.A- encompass the best CP recognition motif for assembly into VLPs.

EXAMPLE 5: THE ROLES OF ELECTROSTATICS AND PS COOPERATIVITY IN VLP ASSEMBLY

PS-mediated assembly explains features of viral genome packaging that purely electrostatically driven reactions do not, although there is clearly a beneficial effect of charge neutralisation in supplying some of the free energy to drive encapsidation. We therefore examined the importance of these effects on STNV assembly using a series of charge-change CP variants. Mutations at three positively charged residues R8, R14 and K17 in the N-terminal arm of the CP (FIG.12), were produced with A or D in place of K or R. Since R14 and K17 are adjacent in three-dimensions, their variants were made as the double mutants, i.e. R14 Å/K17 Å and R14D/K17D. The mutated CPs express normally (FIG.18), but both double mutants fail to assemble significantly inE. coliunder these conditions. Since VLPs obtained fromE. colicontain host cell RNA, as well as the recombinant mRNA encoding the viral CP, this outcome suggests that R14 and K17 play important roles in assembly.

All the variant proteins were examined for their abilities to bind RNA oligos encompassing either a single PS (B3) or the 127-mer fragment (FIG.14&FIG.19). Neither double mutant bound either RNA under these conditions. R8A assembles around B3 but requires a much higher (>10 fold) CP concentration to do so, consistent with it having a lowered affinity for the RNA. By 1 μM CP it forms T=1 shells that are resistant to RNase challenge. The R8D variant confirms the importance of the positive charge, failing to form any stable higher order species, even at higher concentrations, with the RNA remaining accessible to RNase digestion. This dependence on favourable electrostatic interaction remains when R8D is titrated against the 127-mer (FIG.14A). However, with this RNA both R8A and wild-type CP show very similar binding curves, including the initial Rh decrease. It appears that the co-operativity arising from CP binding at multiple PS sites overcomes the deleterious effect on intrinsic RNA-CP affinity of reduced electrostatic attraction. If we assume that the altered charge(s) on the N-terminal arm does not significantly alter the unliganded CP conformation, these effects probe the role(s) of electrostatic interactions during RNA sequence-specific triggering of assembly. They imply that charge neutralisation is not an absolute requirement for assembly on longer natural RNA fragments, consistent with the PS-mediated, but not the electrostatic assembly mechanism. Given that the cooperativity of multiple RNA PSs can overcome diminution of electrostatic attraction, as expected for a process in which PSs act collectively, we then examined how many PSs are required to generate cooperative assembly. Given the importance of recognition at PS3 and the effects seen for fragments containing five PSs, three sub-fragments of the 127-mer each containing PS3 were tested (FIG.14B&FIG.20). These are PS1-PS3 (nts 1-76); PS2-PS4 (nts 38-104) and PS3-PS5 (nts 66-127), each potentially able to bind CP at PS3 but differing in the numbers of flanking sites, from two 5′ or 3′ of PS3, to just one on each flank. Only the fragment with the PS3 centrally located assembles T=1 shells that are RNase resistant, although it does not show a collapse and the overall yield is lower than for the 127-mer. The other fragments appear to form non-specific aggregates that eventually spontaneously dissociate.

The interpretation of these results is non-trivial. The effects are clearly not purely electrostatic in nature since the PS2-4 fragment (66 nts) is shorter than PS1-3 (76 nts) and 1 nt shorter than PS3-5. To understand the specificity of the reactions we need to consider the folding propensity of each of the PS-encompassing sites. The secondary structure of the 127-mer shown (FIG.15) was arrived at by constraining its folds to capture the maximum number of SLs with -A.X.X.A- loop motifs present. In this fragment only PS1 and PS3 are predicted to have a favourable folding free energy (Mfold,(10). in isolation. This is consistent with our previous smFCS assays, in which alterations of the CP recognition motifs within each PS and variations in the relative spacing of PS3 with its neighbours resulted in markedly different assembly behaviour(28). We expect these RNA molecules to exist in solution as an ensemble of differing conformations. Interaction with the STNV CP will displace this equilibrium, preferentially selecting a single or few assembly competent conformations in which the PSs are present. The assembly efficiency we see may therefore be directly related to the population of such conformers in the ensemble and thus related to the free energy costs in imposing this conformation. Assessing the extent of a conformational ensemble is difficult, but a sense of the likelihood of alternate structures can be obtained from Mfold by altering the usual default parameters to explore an ensemble of suboptimal folds within 500 percent from the minimum free energy fold (suboptimality=500).

When such structures are examined for the three PS-containing fragments, a possible explanation for their assembly competencies emerges. For PS1-3, the dominant folds encompass PS1 with a minority also containing PS3 (Table 5). In principle, that minor conformer could promote assembly, but the critical spacing between PS1 and PS3 is too large to facilitate the co-operative effects of multiple PSs. A similar analysis of PS2-4 suggests that the dominant secondary structure does not contain any of the PS folds expected for the 127-mer. However, its predicted secondary structure contains two alternative SLs that are almost always present, one of which presents an -A.X.X.A- sequence (FIG.22). Their relative spacing (4 nt) is short enough to see a co-operative effect. The PS3-5 fragment forms two SLs within 10-12 nts of one another, one presenting an -A.X.X.A- motif as PS5. This would suggest an assembly-competent structure. However, in the ensemble of possible structures, this SL is only present in 6% of the potential folds (Table 5), which may account for the assembly behaviour (FIG.14B).

The conformational scrambling behaviour described above for the fragments encompassing three PSs probably reflects events in vivo where it is known that sequences within the 127-mer participate in formation of a translational enhancer with sequences in the 3′ UTR(29). That complex cannot be present in the assembly competent conformer. In order to explore the effects of such secondary structure folding propensity further, we turned to the design of artificial PS-containing sequences.

EXAMPLE 6:ASSEMBLY OF NON-VIRAL SUBSTRATES

In order to investigate the requirements for an efficient assembly substrate, we produced synthetic cassettes mimicking aspects of the wild-type 127 mer (PS1-5) in which most of the natural viral sequence has been replaced (˜77%). Attempts to create these sequences using a simple base substitution scheme, e.g. swapping all As for Us; Cs for Gs, Gs for Cs and Us for As in the regions other than the CP recognition motifs, all resulted in unstable secondary structures. We therefore chose to modify the existing SLs by conversion of base pairs to G-C, inversion of existing G-C pairs, or adding extra base pairs and then checking that they would likely fold into similar secondary structures to those in the wild-type 127-mer. The natural viral sequences connecting these SLs were then replaced with strings of As and Gs until only one fold was most likely (FIG.18&FIG.23). The relative separations of the base-paired stems were kept identical to those in the wild-type 127-mer. As a result of these changes, PSs 1, 2, 4, and 5 have been stabilised compared to the wild-type 127mer, with all SLs having favourable folding propensity (FIG.23).

To assess the importance of the folding propensity of the dominant PS3 site we also created the following synthetic versions: 1) Unstable PS1-5, in which the folding free energy of PS3, the central PS, is positive (0.3 vs −2.6 kcal/mol), i.e. a scenario in which PS3 is unlikely to fold spontaneously; 2) Stable PS1-5, in which the folding free energy of the central PS is more negative (−3.5 vs −2.6 kcal/mol for the 127-mer), i.e. where PS3 is more stable; 3) AII PS3, in which all five PSs mimick PS3, with stems of all PSs extended to the same length (7 bp) and all CP recognition motifs identical to that in wild-type PS3; & 4) Synthetic, stabilised PS1-5, containing the artificial PSs 1, 2, 4 and 5 from Stable PS1-5, and the artificial extended stem-loop for PS3 from the AII PS3 construct. The latter is hyper-stabilised with respect to the PS3 in both the wild-type 127-mer and the Stable PS1-5 (−7.6 vs −2.6 or −3.5 kcal/mol, respectively).

EXAMPLE 7

In order to compare the behaviours of these test variant oligonucleotides we examined their potential secondary structures. Table 5 lists the frequency of occurrence of each PS in ensembles created using the suboptimality feature in Mfold, together with their relative spacings. In addition, we compared their circular dichroism (CD) spectra. CD provides a physical signal(30), the molar ellipticity at 260 nm, that is proportional to the percentage of base-paired residues and/or tertiary structure. The measurements were made in a buffer containing calcium ions since these are required in the reassembly buffer, there being several Ca2+ binding sites within the STNV capsid (38, 39). Titration of the test RNAs up to 2 mM calcium, the concentration in reassembly buffers, results in mild increases (9-17%) in the 260 nm ellipticity, as expected (FIG.24A). The only exception is Unstable PS1-5, which does not respond to the presence of the cation. The molar ellipticity values of all test RNAs in this buffer decline as expected with temperature (FIG.24B). All the RNAs have different CD ellipticities at 260 nm, illustrating the complexity of comparing RNA conformational ensembles. The Unstable PS1-5 sample is much less structured throughout the temperature range. Perhaps surprisingly given the apparent Mfold structures, the wild-type 127-mer has the highest amount of structure at the lower temperatures. At the highest temperature tested all the RNAs except Unstable PS1-5 have roughly similar ellipticity values, implying that they had reached similar levels of denaturation.

All these synthetic variants trigger assembly of T=1 capsids and are able to protect the encapsidated RNA from challenge by nuclease but with very different CP concentration dependences. All but the Unstable PS1-5 show similar initial decreases in Rh to the 127-mer (FIG.15B). The Unstable PS1-5 assembly behaviour resembles that of PS3 alone, suggesting that it has lost co-operativity, and its distribution plot and appearance in EM images (FIG.24) suggests that it has also lost the ability to regulate capsid formation efficiently. In contrast, the importance of the central PS folding propensity is illustrated by the behaviour of Stable PS1-5. Despite the potential issues with a folding ensemble, it shows a similar collapse to the 127-mer and a cooperative assembly to T=1 particles with a similar Rh distribution to the wild-type fragment. It assembles into VLPs at lower CP concentrations than the wild-type 127-mer, i.e. under these conditions it is a better assembly substrate. Remarkably, AII PS3 also assembles more efficiently than wild-type even though it encompasses PSs that are longer than those found in the 127-mer, suggesting that there is some leeway in the PS secondary structure context in which the recognition motif is presented. This is a little surprising given the critical dependency on PS spacing around PS3 observed previously(19). The efficiency of assembly and the folding propensity of the AII PS3 variant notwithstanding, the Synthetic, stabilised PS1-5 is by far the best assembly substrate, assembling to VLPs most efficiently (i.e. it assembles more quickly following 100 nM CP titration point) (FIG.15B).

EXAMPLE 8

These results suggest that it is possible to abstract the critical assembly features from a viral genomic RNA fragment. Given the alterations in the stem lengths and loop sizes in the synthetic fragments it would also appear that there is considerable scope for engineering templates with improved PS folding propensity.

Transfer of Critical Assembly Features to Genomic-Scale RNAs

As a test of whether these experiments have successfully identified essential assembly features we examined how inclusion of this improved RNA “cassette” alters the assembly efficiency of a natural RNA. That RNA must be inherently able to be assembled into the small volume of the STNV virion. The genomic fragment from 128-1239 nts of the STNV-1 RNA is the obvious test fragment. We therefore constructed two genomic chimeras: [Unstable PS1-5+Δ1-127STNV-1], which is 1242 nts long and [Synthetic, stabilised PS1-5+Δ1-127STNV-1], 1248 nts long, and compared their assembly efficiencies in vitro relative to the wild-type STNV-1 RNA (FIG.16A). All three behaved differently in these assays, implying that the sequence and structure of the 5′ 127-mers regulates the assembly pathway of a fragment that is over 10 times its size. The wild-type genome shows the expected initial collapse in Rh (to ˜7.5 nm) followed by a slight rise in Rh consistent with the formation of T=1 particles (FIG.16B). Note, the data inFIG.16Aare for a CP titration compared to the single step addition of a complete complement of CPs described previously (12). In contrast, the Unstable PS1-5 leader sequence results in a larger initial collapse in Rh (to ˜5 nm; 65%) followed by only a small rise that implies VLPs are not made efficiently. Indeed the RNA in this species remains susceptible to RNase degradation, implying that it has not been completely encapsidated. Synthetic, stabilised PS1-5+Δ1-127STNV-1 has roughly the same initial, CP-free Rh, but appears simply to shrink to values consistent with VLP formation, rather than collapsing and recovering. Rh distributions and EM images of the assembly products are consistent with this interpretation. It is noticeable that the Synthetic, stabilised PS1-5+Δ1-127STNV-1 VLP trace is less noisy than the other samples and has a distribution in Rh sizes that more closely matches that of authentic VLPs isolated fromE. coli. Confirmation of the interpretation of these results is provided by quasi-elastic light scattering (QELS) of the products following elution from a gel filtration column (FIG.25).

EXAMPLE 9

We have shown that the dual code inherent in RNA PS-mediated virus assembly, i.e. that genomic RNAs simultaneously encode a genetic message as well as instructions for efficient capsid assembly, are separable. An important question is why do the codes not separate during the course of viral evolution, especially as replication in ssRNA viruses occurs via error-prone processes that lead to creation of a quasi-species of genome variants. There are now three examples of viruses using RNA PS-mediated virus assembly where we have structural information that partially answer this question. In bacteriophage MS2(31), human parechovirus-1(32) and STNV(19), at least one of the PS sites in the genome also encodes amino acid residues forming part of the PS binding site. This intimate embedding of both codes has the consequence of favouring assembly only of progeny RNAs in which PS-mediated assembly persists. Similarly the density of functions encoded within such RNAs is well known. The natural 5′ 127-mer in the STNV genome also forms an essential transcriptional/translational enhancer contact with the 3′ end sequence. Since that structure and assembly are mutually excluding functions, the natural sequence has evolved to balance the propensity that they form such that the viral lifecycle can proceed efficiently.

The focus here is the property of the assembly code liberated from the wild-type viral RNA sequence. Indeed, by sequentially investigating each aspect of the STNV assembly sequence in its natural context we have been able to reproduce its effects in triggering in vitro assembly of STNV CPs using a synthetic non-viral RNA. Additional refinements allow us to produce sequences that are either less or more efficient than the wild-type STNV 127-mer. These results confirm the nature of PS-mediated assembly for STNV. Assembly in vitro initiates within the 127-mer by recognition by CP subunits of the PS3 stem-loop. Higher-order CP binding is dependent on the correct positioning and folding of the neighbouring PSs (PS2 and 4), each presenting a consensus CP recognition motif in the loop. The 127-mer potentially encompasses five PSs that make the initial binding co-operative with respect to protein concentration leading to a collapse in the hydrodynamic radius of the RNA, a necessary precursor to encapsidation. Electrostatic interactions contribute to these protein-RNA contacts but are not the major driving force, which instead is a high-affinity sequence-specific interaction of the stem and loop regions of the PSs with the inner surface of the protein capsid. Despite its minimal sequence content, the -A.U.U.A- sequence is bound with low nanomolar affinity by the CP. Remarkably, grafting the synthetic variant 127-mers onto the remainder of the natural STNV-1 genome results in chimeras whose assembly properties are dominated by the first ˜10% of the RNA.

Previously, Wilson and colleagues showed they could direct assembly of non-viral RNAs into rods of Tobacco Mosaic Virus (TMV) CP by creating RNA chimeras encompassing the TMV assembly initiation site(33, 34). This was successful, with the length of the protein-coated rods formed being determined by the length of the RNA being packaged, as expected from the known assembly mechanism(35). This approach was less successful when applied to spherical ssRNA viruses(36), the highest affinity MS2 PS having positive effects on in vitro encapsidation of short RNAs but being less important on longer ones(37). Note, all these experiments were done at micromolar concentrations where the effects of PS-mediated assembly are lost(31, 37). The results described above suggest an efficient route for encapsidation of bespoke, non-viral RNAs in shells of viral CPs. In vitro assembly may be possible for a large number of CP-RNA combinations, but it differs from in vivo assembly where, in many viruses, there is good evidence suggesting that only nascent genomic transcripts emerging from the viral polymerase complex are packaged into progeny virions. In such reactions, the RNA is very likely to fold kinetically, avoiding some of the issues with RNA conformational ensembles in the in vitro reactions such as those described here.

Viruses and virus-like particles are finding increasing potential medical applications as gene therapy or drug-delivery vectors, as well as acting as non-replicating synthetic vaccines. Viral protein shells are also of interest for nanotechnology applications. The results described here offer an important insight into ways to create such structures with high efficiency and potentially carrying non-viral RNAs with advantageous properties. This will be essential for the production of designer synthetic virions.

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