NANO LETTERS
Natural Nanochemical Building Blocks: Icosahedral Virus Particles Organized by Attached Oligonucleotides
2004 Vol. 4, No. 8 1385-1389
Erica Strable, John E. Johnson,* and M. G. Finn* Departments of Chemistry and Molecular Biology, and The Skaggs Institute for Chemical Biology, The Scripps Research Institute, 10550 N. Torrey Pines Rd., La Jolla, California 92037 Received April 27, 2004; Revised Manuscript Received June 11, 2004
ABSTRACT Virus particles, like metal nanoparticles and organic polymer beads, may be induced to assemble together by virtue of duplex base pairing of attached oligonucleotide chains and to disassemble on heating with unusually sharp temperature profiles. Icosahedral virions of the type used here have the added property of being structurally known to atomic resolution, offering unique opportunities to program assembly characteristics at the level of particle structure.
The developing science of nanotechnology requires molecular building blocks of a variety of sizes. Colloidal metal particles, particularly gold, have received much attention as supporting scaffolds, decorated with functional units that allow their use in sensors and devices.1 The strong interaction between n-alkylthiols and the metal surface provides the most popular method for the attachment of molecular groups.2-4 Such bonding is convenient to engineer but is reversible at moderate temperatures and is kinetically unstable with respect to movement of thiols on the surface.4 Thus, groups that are deposited onto a metallic surface cannot be precisely fixed with respect to one another, although average spacings can be arranged by diluting the monolayer of functionalized alkylthiols with analogues lacking the functional group.5 Precise, angstrom-level control of reactivity in nanotechnology requires nanoscale building blocks of known structure at atomic resolution. Biology is a ready source of such materials, and we focus on nonenveloped virus particles as their prototype. We describe here the functional derivatization and directed aggregation of such species by virtue of Watson-Crick duplex base pairing of deoxyribonucleotides, in analogy to the work of Alivisatos6 and Mirkin1,7-11 with gold nanoparticles, and others12 with gold surfaces. As has been previously recognized, the energetics, rates, and structures of duplex DNA interactions are controllable, making them a promising vehicle for interconnecting components in complex molecular- and nanoscale systems.13-15 Cowpea mosaic virus (CPMV), an icosahedral assembly of 60 identical coat protein units which packages its RNA genome, is 30 nm in diameter and can be addressed at selected lysine residues by organic reagents.16,17 The X-ray crystal structure of CPMV is known,18 and mutants have been 10.1021/nl0493850 CCC: $27.50 Published on Web 07/21/2004
© 2004 American Chemical Society
Figure 1. (a) CPMV subunit organization. (b) A single protein asymmetric unit showing the positions of the reactive lysine (green, K38 in the small subunit) of wild-type CPMV and cysteine residue (white, inserted between G98 and K99 of the large subunit) in CPMV-EF-Cys.
constructed which bear reactive cysteines at designed positions. These factors make the system programmable in terms of the number and spacing of attachment points on both the exterior and interior surfaces of the protein shell.19 Purified wild-type and mutant CPMV preparations are available in gram quantities from infected cowpea plants using a simple and convenient isolation procedure that is amenable to scaleup.20-22 The particles are sufficiently stable to variations in temperature (up to 50 °C), pH (from 3 to 10), and solvent (aqueous/organic mixtures) to be used in chemical and material synthesis and manipulation. The repeating unit of the CPMV capsid (the “asymmetric unit”) is composed of two proteins - a single-domain sequence of 24 kD and a two-domain sequence of 42 kD, designated the “small” and “large” subunits, respectively. The structure and subunit organization of the particle is shown in Figure 1. The viruses chosen for this exploratory study were wild-type CPMV (WT-CPMV), which contains
Figure 2. CPMV-oligonucleotide conjugates used in these studies. The designation for each derivatized virus particle incorporates the type of virus, linker, and oligonucleotide used. Viruses labeled 1 and 1Fl contain the complementary oligonucleotide sequence to 2 and 2Fl; each oligonucleotide was attached to the virus at its 5′ end.
Figure 3. Detection of fluorescein-oligonucleotide hybridization to virus-oligonucleotide conjugates by electronic spectroscopy.
a more reactive lysine at position 38 of the small subunit,16,17 and a mutant virus with a Cys-Ala insert installed between G98 and K99 in the EF-loop of the large subunit.19 The mutant is designated here as CPMV-EF-Cys to denote the highly reactive cysteine residue in each asymmetric unit. The positions of these reactive groups are shown in Figure 1. Complementary 20-mer oligonucleotides composed of a repeating GC-rich 3-base pair sequence were attached to CPMV scaffolds either at lysine residues or at genetically engineered cysteines, as shown in Figure 2. The yields of recovered virus ranged from 75% to 90%, and the resulting materials were shown to be composed of intact particles by sucrose gradient sedimentation, the observation of characteristic A260/A280 ratios, and size-exclusion chromatography, as detailed in the Supporting Information. These results confirm that there is little or no disassembly of the virus during the coupling process. The extent of derivatization was measured quantitatively with the use of fluorescein-labeled oligonucleotides, with attachment to lysine residues of wild-type CPMV up to the level of 30 ( 3 oligonucleotides per virion, and 30 ( 3 per virion to cysteine residues of CPMV-EF-Cys. In the following discussion, these values are denoted after the sequence 1386
designation for each virus, as in EF-Cys‚S130. All conjugates were characterized by SDS-PAGE, anion-exchange FPLC, transmission electron microscopy (TEM), and multiangle dynamic light scattering.23 The last of these showed the virus-oligonucleotide structures to have substantially larger hydrodynamic radii than the unlabeled capsids. Selective hybridization of complementary strands to virusoligonucleotide conjugates was demonstrated as shown in Figure 3. Incubation of WT‚N130 with fluorescein oligonucleotide 2Fl, or WT‚N230 + 1Fl, followed by purification of the virus gives products that show absorbance of the fluorescein dye. Quantitative measurement of dye absorbance relative to virus protein showed the formation of 26-28 duplex chains per virus, representing approximately 90% of the covalently attached single strands. Incubation of a virusoligo conjugate with a fluorescein-labeled oligonucleotide of the same, and therefore noncomplementary, sequence (e.g., WT‚N130 + 1Fl, or WT‚N230 + 2Fl) resulted in no dye attachment (Figure 3). The mixing of CPMV particles bearing complementary oligonucleotides should result in their aggregation by virtue of duplex base-pairing, reversible upon the addition of a competing oligonucleotide, as shown in Scheme 1. This Nano Lett., Vol. 4, No. 8, 2004
Figure 4. Fluorescence spectroscopy of virus-oligonucleotide-dye conjugates and their aggregates (excitation wavelength in parentheses) at identical concentrations. (A) WT‚N110/Fl48 (495 nm); (B) WT‚N210/Rd48 (495 nm); (C) WT‚N210/Rd48 (520 nm); (D) WT‚N110, WT‚ N210 (495 and 520 nm, superimposed); (E) WT/Rd57 mixed with WT/Fl58; (F) WT‚N110/Fl48 + WT‚N210/Rd48 (495 nm); (G) aggregate from (F) treated with excess 1 (495 nm). Not shown: WT‚N210/Rd48 + 1 (520 nm; overlays with spectrum C, WT‚N110/Fl45 + WT‚N210 (495 nm; overlays with A), and WT‚N110 + WT‚N210/Rd45 (495 nm; overlays with B). Scheme 1
phenomenon was explored with the observation of light scattering, fluorescence resonance energy transfer (FRET) and TEM, as follows. The mixing of virions bearing oligonucleotide sequences 1 and 2 was accompanied by an immediate and sequencedependent aggregation of the particles, revealed most conveniently by an increase in light scattering throughout the UV-visible spectrum. Maximal scattering was reached within 1 h at virus concentrations of approximately 1 mg/mL (0.18 µM in virions), although aggregation was shown by TEM to continue for much longer than this period (see below). The addition of excess oligonucleotide 1 or 2 (with or without the aminohexyl linker at the 5′ end) resulted in a decrease in light scattering intensity of the mixture back to the original value, consistent with the disruption of duplex-oligonucleotide aggregates by competing soluble oligonucleotides.23 As before, no such spectroscopic changes were observed for any oligonucleotide-decorated virus alone, nor upon their mixing with additional wild-type or the cysteine mutant CPMV. To further explore the solution-phase association of labeled virions, CPMV conjugates WT‚N110 and WT‚N210 were labeled on the capsid with either fluorescein or rhodamine dyes, giving the constructs shown in Figure 4. Quantitation Nano Lett., Vol. 4, No. 8, 2004
of the dye absorbance relative to virus revealed a loading of 48 ( 5 dyes per capsid in each case. The fluorescence properties of each virus sample at long wavelength were those of their respective dyes. Thus, excitation at 495 nm gave rise to an intense fluorescein emission at 514 nm (Figure 4A) and relatively weak rhodamine emisison at 550 nm (Figure 4B); much more intense rhodamine emission was observed upon excitation at 520 nm (Figure 4C). Virusoligonucleotide samples without dye showed no emission in the region of interest (Figure 4D). Mixing equimolar samples of viruses bearing complementary oligonucleotides and complementary dyes gave products with fluorescence spectra showing enhanced rhodamine emission at 550 nm upon excitation at 495 nm (Figure 4F), relative to samples consisting of virus particles lacking the complementary oligonucleotides (Figure 4E), indicative of FRET between fluorescein and rhodamine dyes in the former case. When excess competing oligonucleotide was added, the enhanced emission band at 550 nm disappeared (Figure 4G); added oligonucleotide did not quench rhodamine emission upon excitation at 520 nm, nor did the emission properties of dyelabeled particles change by virtue of aggregation with complementary virions lacking the partner dye (not shown). 1387
Figure 5. Temperature dependence of oligonucleotide-directed aggregation using equimolar mixtures of EF-Cys‚S115 + EF-Cys‚S215. (A) A final virus concentration of 1 mg/mL, 4 °C, 16 h; (B) 1 mg/mL, room temperature, 16 h; (C) 1 mg/mL, 40 °C, 16 h; (D) sample from C heated at 45 °C for 12 h and immediately applied to the TEM grid. (E) Light scattering from a solution of WT‚N130 + WT‚N230 as a function of temperature and time. The arrows mark the times at which the bath temperature was switched to the indicated setting; the time required for the solution to reach thermal equilibrium in the cell was estimated to be about 1 min per degree C. Very similar behavior was observed for EF-Cys‚S115 + EF-Cys‚S215. (F) Calculated base pair overlap vs melting temperature curve for the 1+2 duplexed sequence. The dotted line relates the observed melting temperature of the virus aggregate mediated by the 1+2 interaction with the calculated overlap length of a single 1+2 duplex. (G) Schematic representation of the maximum average overlap between displayed oligonucleotide strands.
Thus, hybridization of complementary oligonucleotide strands brings some fraction of the virus-bound donor (fluorescein) and acceptor (carboxyrhodamine) dyes into close enough proximity (Fo¨rster radius, R0 ) 55 Å) to achieve detectable energy transfer, and added oligonucleotide breaks up this association. TEM analysis of mixtures involving complementary oligonucleotide sequences revealed sequence-specific aggregation of virions into arrays, the form and size of which depended on a variety of variables. The assemblies were broken apart using DNase1, excesses of competing complimentary oligonucleotide, or reducing agents (for cases in which linkers contained a disulfide bond), and resisted RNaseA, high salt concentration, and a noncomplementary sequence.23 Figure 5 shows differences in order, size, and dimensionality of aggregates that were observed as a function of the annealing temperature. Small aggregates approaching two-dimensional hexagonal packing resulted from annealing at 4 °C (Figure 5A). At room temperature, a dramatic increase in the size of the predominantly two-dimensional arrays was observed by electron microscopy (Figure 5B). Increasing the temperature to 35-40 °C generated threedimensional arrays, the precise packing of which cannot be determined by TEM (Figure 5C). Unmodified virus and virus particles with one oligonucleotide sequence remained as discrete individual particles upon incubation at 40 °C (data not shown), indicating that base pairing is essential for aggregation. The repeating nature of the oligonucleotide sequence employed here allows the system to reach a variety of 1388
aggregated states. At lower temperatures and/or shorter reaction times, duplex interactions of less than the maximum length are formed and have insufficient energy or time to break up. The result is less extensive and weaker arrays, as shown in Figure 5A,B. In these cases, deposition and drying on the TEM grid imposes a force on the particles normal to the grid surface. This causes the three-dimensional clusters to collapse to two-dimensional aggregates because the oligonucleotide bridges between the particles have longer single-stranded connections and weaker double-stranded connections. When allowed to anneal at higher temperatures, the population of various duplex interactions is shifted to those of greater overlap until the system reaches its thermodynamic equilibrium and a relatively strong aggregated network. Heating three-dimensional arrays to 45 °C induced rapid dissociation back to individual particles, with little or no loss to capsid decomposition (Figure 5D). This sharp melting behavior was observed for aggregates of particles bearing both 15 and 30 oligonucleotides per virion and suggests that the duplex links between virus particles have an upper melting temperature limit of 45 °C, which corresponds to an overlap of 11 base pairs of the repeating sequence (Figure 5F,G). Such sharp melting behavior has been observed by the Mirkin and Schatz laboratories for arrays of oligonucleotide-derivatized gold nanoparticles and may be a characteristic of systems composed of multivalent components.24 These authors have noted that multiple factors, including DNA density on the particles, particle size, interparticle spacing in the aggregates, and salt concentration, influence Nano Lett., Vol. 4, No. 8, 2004
the melting profile. Close packing of oligonucleotides seems to give rise to decreased melting temperatures,24 and our virus system (with a minimum interoligonucleotide spacing of approximately 5 nm) may be placed in this category. It must be noted that we do not yet have firm evidence that the covalent attachment of oligonucleotides to the CPMV capsid occurs with a random distribution over the particle surface. This work shows that virions can function as organizational scaffolds for DNA-directed assembly just as metal nanoparticles can, but with more precise characterization of the nanoparticle-oligonucleotide building block, including better knowledge of the relative placement of the oligonucleotide chains. Assembly is shown to be dependent on a variety of factors and to exhibit a kinetic profile similar to previous observations of the behavior of gold-oligonucleotide conjugates. More detailed insights, including the dependence of the assembly process on the nature and spacing of oligonucleotides, are the goal of ongoing studies. Acknowledgment. We thank the David and Lucille Packard Foundation Interdisciplinary Science Program, the NIH (EB00432-01), and the La Jolla Interfaces in Science Program - Burroughs Wellcome Fund (graduate fellowships to E.S.) for support of this work. Supporting Information Available: Experimental details and additional data. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Storhoff, J. J.; Elghanian, R.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1998, 120, 1959-1964. (2) Bain, C. D.; Troughton, E. B.; Tao, Y.-T.; Evall, J.; Whitesides, G. M.; Nuzzo, R. G. J. Am. Chem. Soc. 1989, 111, 321-335. (3) Weisbecker, C. S.; Merritt, M. V.; Whitesides, G. M. Langmuir 1996, 12, 3763-3772.
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(4) Templeton, A. C.; Wuelfing, W. P.; Murray, R. W. Acc. Chem. Res. 2000, 33, 27-36. (5) Houseman, B. T.; Mrksich, M. Angew. Chem., Int. Ed. Engl. 1999, 38, 782-785. (6) Loweth, C. J.; Caldwell, W. B.; Peng, X.; Alivisatos, A. P.; Schultz, P. G. Angew. Chem., Int. Ed. Engl. 1999, 38, 1808-1812. (7) Storhoff, J. J.; Mirkin, C. A. Chem. ReV. 1999, 99, 1849-1862. (8) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. Nature 1996, 382, 607-609. (9) Elghanian, R.; Storhoff, J. J.; Mucic, R. C.; Letsinger, R. L.; Mirkin, C. A. Science 1997, 277, 1078-1081. (10) Storhoff, J. J.; Lazarides, A. A.; Mucic, R. C.; Mirkin, C. A.; Letsinger, R. L.; Schatz, G. C. J. Am. Chem. Soc. 2000, 122, 46404650. (11) Taton, T. A.; Mirkin, C. A.; Letsinger, R. L. Science 2000, 289, 1757-1760. (12) Brockman, J. M.; Frutos, A. G.; Corn, R. M. J. Am. Chem. Soc. 1999, 121, 8044-8051. (13) Niemeyer, C. M. Angew. Chem., Int. Ed. Engl. 1997, 36, 585-587. (14) Seeman, N. C. TIBTECH 1999, 17, 437-443. (15) Yan, H.; Zhang, X.; Shen, Z.; Seeman, N. C. Nature 2002, 415, 6265. (16) Wang, Q.; Lin, T.; Tang, L.; Johnson, J. E.; Finn, M. G. Angew. Chem., Int. Ed. 2002, 41, 459-462. (17) Wang, Q.; Kaltgrad, E.; Lin, T.; Johnson, J. E.; Finn, M. G. Chem. Biol. 2002, 9, 805-811. (18) Lin, T.; Chen, Z.; Usha, R.; Stauffacher, C. V.; Dai, J.-B.; Schmidt, T.; Johnson, J. E. Virology 1999, 265, 20-34. (19) Wang, Q.; Lin, T.; Johnson, J. E.; Finn, M. G. Chem. Biol. 2002, 9, 813-819. (20) Lin, T.; Porta, C.; Lomonossoff, G.; Johnson, J. Fold. Des. 1996, 1, 179-187. (21) Goldbach, R.; van Kammen, A. In Molecular Plant Virology; Davies, J., Ed.; CRC Press: Boca Raton, 1985; Vol. 2, pp 83-120. (22) Spall, V. E.; et al. In Engineering Crops for Industrial End Uses; Shewry, P. R., Napier, J. A., Davis, P., Eds.; Portland Press: London, 1998; pp 35-46. (23) See Supporting Information for details. (24) Jin, R.; Wu, G.; Li, Z.; Mirkin, C. A.; Schatz, G. C. J. Am. Chem. Soc. 2003, 125, 1643-1654.
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