Temperature-Programmed Assembly of DNA: Au Nanoparticle

NANO LETTERS

Temperature-Programmed Assembly of DNA:Au Nanoparticle Bioconjugates

2006 Vol. 6, No. 1 16-23

Lisa M. Dillenback, Glenn P. Goodrich,† and Christine D. Keating* Department of Chemistry, The PennsylVania State UniVersity, UniVersity Park, PennsylVania 16802 Received May 12, 2005; Revised Manuscript Received November 21, 2005

ABSTRACT Temperature has been used to control the order of assembly events in a solution containing three types of particles to be linked by two different sets of complementary DNA. At higher temperatures, only the duplexes having higher thermal stability were able to form. By starting at a high temperature and then cooling the sample, these more stable sequences hybridized first, followed by the less stable sequences at lower temperatures. Because of the use of thiolated DNA on Au particles, some loss and exchange of the DNA strands occurred at elevated temperatures. However, since cooperativity favors the “correct” assemblies, Au−S bond lability did not appreciably impact the order of the assembly process. Temperature programming combines the selectivity of DNA-directed assembly with the ability to control the order in which several complementary strands hybridize in a common solution and could contribute to the synthesis of more complex nanostructured materials.

Nanoparticles are promising building blocks for the bottomup assembly of high-density electronics and other functional materials because of their small size and tunable optical, electronic, and physical properties.1 A variety of methods have been developed for nanoparticle assembly, including those based on electrostatics, hydrophobic interactions, surface patterning, Langmuir-Blodgett methods, and magnetic or electric fields.2-16 These approaches can provide simple, high-yield routes to nanoparticle structures but are usually nonspecific and therefore unable to selectively build structures from multiple different particles. Biorecognition-directed assembly has been employed using proteins17-22 and DNA22-41 to exert more control over the specificity of the interparticle attractions and the resulting structures. DNA is particularly appealing because it can be synthesized readily in a desired length and sequence with functional groups for conjugation or detection.29,30 The sequence selectivity of DNA duplex formation provides a route to synthesis of complex assemblies by incorporating multiple different sequence pairs as selective “glues”. Alivisatos and co-workers demonstrated the synthesis of discrete few-nanosphere structures (dimers, trimers, etc.).24,27,39 Mirkin and co-workers have used nanospheres functionalized with many DNA strands to assemble the spheres into larger aggregates in solution, or monolayers on planar supports.23,29 Highly selective assembly can be achieved, enabling sensitive and specific bioanalysis.23,29,34,42 When constructing on the macroscale, it is important to not only control the placement of different objects with respect to each other but also the * Corresponding author. E-mail: [email protected]. † Current address: Nanospectra Biosciences, Houston, TX. 10.1021/nl0508873 CCC: $33.50 Published on Web 12/10/2005

© 2006 American Chemical Society

order in which the objects are assembled (e.g., foundation before walls). In contrast, on the microscale, although biomolecules such as DNA offer selectivity in the placement of building blocks, reported methods exhibit little control over the timing of nanoparticle assembly processes. In this report, we take advantage of the temperaturedependent melting properties of DNA to control the order in which nanoparticles assemble. It is well-known that the melting transition temperature (Tm) of a DNA duplex, that is, the temperature at which one-half of a DNA population is in duplex form and one-half is single-stranded, depends strongly on the sequence length of the DNA duplex.43 By immobilizing DNA of various lengths onto the nanoparticles, the sequence of assembly events can be dictated simply by controlling the hybridization temperature (Scheme 1). All particles for assembly are present in solution at an initial elevated temperature. Upon lowering the solution temperature in discreet steps, the DNA:Au bioconjugates with longer DNA strands begin to assemble. As the temperature is lowered further, DNA:Au bioconjugates with successively shorter DNA sequences join the growing aggregate. The same DNA:Au bioconjugates would form a different assembly if they reacted only at the lowest temperature, where all the conjugates could assemble at the same time; temperature-programmed assembly (TPA) provides a means of controlling the temporal sequence of assembly events. We present a simple proof-of-principle demonstration of the TPA process in which the order of assembly for 12-nm-diameter Au nanospheres is controlled. Three different batches of DNA:Au bioconjugates (A9: Au, B18:Au, and C9/18:Au) were prepared by derivatizing the

Scheme 1.

Temperature-Programmed Assembly of Nanoparticles

surface of 12-nm-diameter Au spheres with thiolated DNA oligonucleotides (Table 1) as described previously.42,44,45 C9/18:Au conjugates were modified with a mixture of C9 and C18. Note that none of the DNA on any of the particle surfaces was directly complementary to sequences on the other particles. The nanoparticles were assembled by the addition of the linking oligonucleotides, A9′C9′ and B18′C18′. Half of the linker sequence is complementary to one of the sequences on particle A or B (Scheme 1), and the other half is complementary to one of the sequences on the particles with mixed DNA (C). The melting transitions for these DNA: Au conjugate aggregates are 53 °C for the shorter duplex and 69 °C for the longer duplex in the high salt buffer (0.3 M NaCl, 100 mM phosphate, pH 7.0) used for these experiments. Therefore, when the hybridization is carried out at 65 °C, only the longer sequences can hybridize, forming aggregates composed of only B18:Au and C9/18:Au conjugates. When the hybridization temperature is lowered to 25

Figure 1. Absorbance spectra of solutions containing conjugates and linking oligonucleotides taken after overnight hybridizations at various temperatures. The control sample contains all of the conjugate types, but no linking DNA, at 25 °C.

°C, aggregates that contain each of the three conjugates should form. Figure 1 shows the absorbance spectra for samples after the hybridization is complete. The intensity of the peak at 520 nm, which is the characteristic plasmon absorbance for the isolated 12-nm Au particles, decreases as the nanoparticles assemble (note that because the aggregates have sedimented, their redshifted absorbance is not observed). The absorbance for the 65 °C system shows approximately onethird the number of particles as in the starting solution. This correlates well with the aggregates being formed from only two of the conjugates (B18:Au and C9/18:Au), leaving the A9: Au conjugates in solution. When the hybridization was carried out at 25 °C, the resulting absorbance at 520 nm dropped significantly, suggesting that nearly all of the particles participated in aggregation.

Table 1. DNA Sequences Used in This Worka name

sequence 5′ to 3′

A9 B18 C9 C18(f)

GTC AAT CTC TTT TTT C3H6-SH ACT CAG TGT GTG CCT TCT TTT TTT C3H6-SH HS-C6H12 AAA AAA AAG TCC TGT HS-C6H12 AAA AAA TTC CTG AGA TGT GTC GGT

A9′C9′ B18′C18′ A9′ B18′ C9′ C18′ N9

GAG ATT GAC ACA GGA CTT AGA AGG CAC ACA CTG AGT ACC GAC ACA TCT CAG GAA GAG ATT GAC AGA AGG CAC ACA CTG AGT ACA GGA CTT ACC GAC ACA TCT CAG GAA ATT AGA CAT

F10 D9 D9′ D21 D21′

6-FAM TAC AGG ACT T C3H6-SH HS-C6H12 (AAA)6 ATT GCC TGC HS-C6H12 (AAA)7 CGA GGC AAT HS-C6H12 (AAA)5 TTA AGA CGA GGC AAT CAT GCA HS-C6H12 (AAA)4 TGC ATG ATT GCC TCG TCT TAA

description on A particles; links to C9 via A9′C9′ (Tm ) 53 °C) on B particles; links to C18 via B18′C18′ (Tm ) 69 °C) on C particles; links to A9 via A9′C9′ (Tm ) 53 °C) on C particles; links to B18 via B18′C18′ (Tm ) 69 °C) (in the fluorescent version of this sequence, 6-FAM is attached on the 3′ end) links A9 and C9 (Tm ) 53 °C) links B18 and C18 (Tm ) 69 °C) complement to A9 complement to B18 complement to C9 complement to C18 not complementary to any sequence in the three-strand TPA system fluorescent, thiolated oligonucleotide complement to D9′ (Tm ) 42 °C) complement to D9 (Tm ) 42 °C) complement to D21′ (Tm ) 84 °C) complement to D21 (Tm ) 84 °C)

a PolyA or polyT spacers were incorporated at the thiolated end of each sequence to reduce steric constraints on hybridization. For this reason, the subscripts refer to the number of bases to be involved in the hybridization process, not the total length of the sequence.

Nano Lett., Vol. 6, No. 1, 2006

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Figure 3. Photograph of an agarose gel after particle separation. Aliquots from the 25 °C hybridization and the 65 °C hybridizations were loaded onto the gel along with particle standards. (A) B18: Au; (B) A9:Au; (C) C9/18:Au; (D) supernatant from 25 °C hybridization; (E) mixture of all three conjugates with no linker; (F) supernatant from 65 °C hybridization. Because the concentration of the particles in the supernatant from the 65 °C sample is lower than that of the control samples, lane F is shown again, on the far right, with increased contrast.

Figure 2. Thermal denaturation (top) and 1st derivative plot (bottom) for the 25 °C hybridization sample containing A9:Au, B18: Au, and C9/18:Au conjugates with oligonucleotide linkers A9′C9′ and B18′C18′.

After hybridization, the aggregates were melted while monitoring the absorbance at 260 nm, which is characteristic for single-stranded DNA (Figure 2). As the temperature was raised, there was an increase in the absorbance at 260 nm, and two distinct melt transitions corresponding to those of the individual duplexes were observed. A greater increase in absorbance is observed for the higher-temperature transition. This is not surprising because, when the shorter duplex melts, only A9:Au conjugates and A9′C9′ linker strands are released from the aggregate, whereas, in the second step, all of the remaining DNA (B18:Au and C9/18:Au conjugates and B18′C18′ linker) are released into the solution. Importantly, as evident in the first derivative plot, both melt transitions are sharp and well-defined. Mirkin, Schatz, and co-workers have reported that cooperativity in the melting of DNA:Au bioconjugate aggregates results in very sharp melt transitions.46 In our experiments, this enables the complete dissociation of the shorter duplexes before the longer duplexes begin to melt. Indeed, Figure 2 shows a nearly 10 °C window between the two melts, suggesting the possibility of incorporating more than two duplexes into a single assembly. It has been shown previously that DNA:Au bioconjugates can be separated by gel electrophoresis based on the length of the oligonucleotide immobilized on the outside of the particle.42,47-49 To identify which conjugates are excluded from the aggregates during hybridization, aliquots were taken from the solutions before thermal denaturation and loaded onto an agarose gel. Figure 3 shows a photograph of the resulting gel. Controls are shown in lanes A, B, and C, which contain conjugates B18:Au, A9:Au, and C9/18:Au, respectively. A mixture of the three types of conjugates with no linker is shown in lane E as a fourth control. Lane D contains the supernatant from the 25 °C hybridization. Because all of the 18

conjugates are hybridized, none remain in the solution, and no particles are observed in this lane of the gel. In lane F, which contains the supernatant from the 65 °C hybridization, the primary band correlates best to A9:Au conjugates. This is consistent with the expectation that conjugates with the shorter DNA (A9:Au) were not incorporated into the aggregate formed at 65 °C. It should be noted that the C9/18:Au conjugates (lane C) run significantly slower than A9:Au and B18:Au, despite having a mixture of the longer and shorter DNA. Because intermediate migration would be expected for a mixture of the two lengths, assuming the total number of DNA strands per particle was unchanged, we hypothesized that the surface coverage of the DNA on the particles was different between the bioconjugate types. The number of DNA strands bound to the Au particles in each conjugate type was determined by fluorescence as described elsewhere.42,50 Conjugate C9/18: Au had a surface coverage of 90 ( 7 oligonucleotides per particle, whereas A9:Au and B18:Au had 57 ( 7 oligonucleotides per particle. Although the conjugates were all prepared under the same conditions, the oligonucleotides on C9/18:Au conjugates were attached via a 5′ thiol moiety with a 6-carbon alkyl chain spacer between the sulfur and the DNA sequence. Conjugates A9:Au and B18:Au were prepared using DNA with the thiol on the 3′ end; these molecules generally have only a three-carbon spacer. The lower coverage for the 3′ oligonucleotides may result from steric hindrance during attachment to the particles. The increased surface coverage for C9/18:Au explains the reduced electrophoretic mobility of these conjugates as compared to A9:Au and B18:Au. It has been shown that at lower surface coverages of oligonucleotide, the DNA can coil, thus reducing the apparent particle radius. At high surface coverages, the DNA extends further out from the Au nanosphere surface, resulting in a larger apparent radius and slower migration.51 Figure 4 compares the electrophoretic mobility of conjugates with either 3′- or 5′-thiol-modified DNA with the same strand lengths. All of the 3′ conjugates migrated more quickly than the 5′ conjugates, and the C9/18: Au conjugates have intermediate migration with respect to C9:Au and C18:Au. The gel in Figure 3 also shows that the conjugates from the 65 °C sample (lane F) run slightly faster than the Nano Lett., Vol. 6, No. 1, 2006

Figure 5. Photograph of an agarose gel after particle separation. Mixtures of conjugates with long DNA and conjugates with short DNA were separated after incubation at 25 °C or 65 °C (two lanes of each are shown). Figure 4. Photograph of an agarose gel illustrating the difference in electrophoretic mobility of DNA:Au bioconjugates prepared with 3′ and 5′ thiolated DNA. The conjugates run in each lane are noted on the photograph. Note that sequences A9 and B18 have a 3′ thiol, whereas sequences C9 and C18 have a 5′ thiol.

unheated A9:Au conjugates (lanes B and E). Given the kinetic lability of Au-S bonds at elevated temperature,42 a reasonable explanation for this behavior is DNA loss from the conjugates when heated at 65 °C overnight. To investigate this possibility, conjugates with a fluorescent sequence (C18f) were prepared. Each batch was split into two samples, one left at room temperature and the other heated overnight. The conjugates were then washed by centrifugation and the amount of DNA remaining on the particles in each sample was quantified by fluorescence.42,50 The sample that had been heated at 65 °C overnight had only ∼64% as many oligonucleotides per particle as compared to the sample stored at room temperature, a loss of 36%. Kinetics experiments in which absorbance at 260 and 540 nm was followed as a function of time indicated that the hybridization at 65 °C is complete after 5.5 h (Supporting Information Figure S1). Therefore, the above experiment was repeated with samples heated for this shorter hybridization time. In this case, the retention of oligonucleotides on the particles was improved to 76%, a loss of 24%. The substantial loss of DNA from the particles during heating at 65 °C introduces the possibility for exchange of thiolated sequences between particles in the TPA experiments. Such exchange could be detrimental to assembly control. To investigate this possibility, equimolar amounts (1.3 picomoles each) of conjugates Au:D21 and Au:A9 were mixed together in two samples. One was incubated at 65 °C overnight, and the other was left at room temperature. These samples were then run on an agarose gel for comparison (Figure 5). The large difference in DNA sequence length results in good separation. As expected based on the experiment described above, the heated sample runs faster because of the DNA lost from those particles. However, for both samples, two distinct bands are visible. Although a slight smear between the bands is apparent in both samples (indicating a range of DNA surface coverages for particles within a batch), the spreading of the 65 °C sample is no Nano Lett., Vol. 6, No. 1, 2006

greater than that of the 25 °C sample, indicating that there is not a great deal of exchange of oligonucleotides between the conjugates. Although the gel electrophoresis results indicate that exchange of DNA between the bioconjugates is limited, the possibility of exchange occurring is not entirely precluded. Therefore, we used fluorescent oligonucleotides to quantitatively evaluate the extent of DNA exchange between particles at 65 °C. Several batches of A9:Au conjugates were prepared, and each batch was split into two samples. The concentration of each sample was equivalent to that of the particles remaining in solution after the 65 °C hybridization. Free thiolated oligonucleotides of sequence F10 were added to both samples in an amount equivalent to the DNA that would have been lost from the other two types of particles in the TPA sample at 65 °C. One of the conjugate samples was heated to 65 °C for 6 h, while the other was left at room temperature. Excess DNA was then washed from the samples, and the amount of F10 adsorbed to the particles from solution was quantified by fluorescence.42,50 The heated samples showed 8 ( 1 fluorescent oligonucleotides per particle (14% of total DNA strands), while the samples left at room temperature showed 2 ( 1 fluorescent oligonucleotides per particle (3.5% of total DNA strands).52 Thus, although 24% of the oligonucleotides dissociate from the conjugates with heat (as shown above), only 14% exchange of oligonucleotides is observed. This difference may result from electrostatic repulsions and/or steric hindrance to reassociation of the thiolated DNA to the bioconjugates, which already host a large number of DNA strands. It is also likely that some of the free DNA adheres to the tube wall. In addition, thiolated oligonucleotides may react with each other in solution to form disulfides, which would experience greater steric and electrostatic repulsions to particle adsorption. In the actual TPA system, with the other two types of particles in solution, some of the oligonucleotides will exchange onto particles of the same type as that from which they dissociated. Although some interparticle exchange of oligonucleotides has occurred, gel electrophoresis data (Figure 3) suggest that the exchange had little effect on the assembly process; that is, the conjugates remaining in solution after the 65 °C 19

hybridization step appear to be primarily those with the lower-melting DNA sequences. Thus, it appears that “correct” assembly is driven by cooperativity46 because of the much larger numbers of nonexchanged sequences over exchanged sequences on any given particle type. We note that the gold-thiol attachment, although quite common in DNA:nanoparticle bioconjugation, is not required for the TPA process. Exchange could be eliminated by using a more stable nanoparticle-DNA attachment, such as silane-based chemistries on SiO2.53 Because DNA loss and exchange introduce complexity to the system that may not be observable via gel electrophoresis or thermal denaturation, and because the gel in Figure 3 does not preclude the presence of the similar mobility B18:Au conjugates in the supernatant of the 65 °C hybridization sample, we sought an independent means of verifying the TPA process. After the 65 °C hybridization, we compared the ability of DNA sequences to interfere with further assembly at 25 °C. Six identical TPA samples (prepared as for the experiments in Figures 2 and 3 above) were hybridized at 65 °C for 6 h, at which point, each sample contained an aggregate at the bottom of the tube, as well as some bioconjugates in solution. If TPA is proceeding as designed, then the conjugates in solution should be primarily A9:Au (with perhaps a small number of C9/18:Au) and addition of A9′ should interfere with further assembly. Because DNA:Au assembly is highly cooperative,46 complete inhibition by a complementary strand is unlikely. Nonetheless, inhibition can be expected only when sequences complementary to those on the particles in solution are added, thus providing a means of differentiating which particles are present in solution. We compared the effect of added A9′ to that of added C9′, C18′, and B18′, as well as a sequence noncomplementary to any of the TPA strands (N9), and buffer alone. Samples were then allowed to hybridize at room temperature overnight. Figure 6 shows photographs of the samples before and after the overnight hybridization. The only sample that does not fully aggregate overnight is the sample with the complement to the shorter TPA sequence (A9′). All other samples with complementary sequences appear the same as the samples with buffer and noncomplementary DNA. This is a strong indication that the conjugates with the shorter DNA (A9:Au) did remain in solution, whereas those with the longer DNA (B18:Au and C9/18:Au) were assembled, during the 65 °C hybridization. Absorbance spectra were taken at each step in this experiment (Figure 7, top). In the sample containing A9′ there is an approximately 2-fold loss in absorbance at 520 nm, whereas the loss in absorbance for all other samples is nearly complete. The decrease in absorbance in this sample results from competition of A9′-A9:Au hybridization with the normal progression of the TPA process. Because the aggregate containing B18:Au and C9/18:Au conjugates is still present in each sample, the hybridization with A9′ must compete with the highly cooperative binding interactions between the A9:Au conjugates in solution and complementary sites on the aggregate (A9:Au binds C9/18:Au via the 20

Figure 6. Photograph of TPA samples after 65 °C hybridization, (a) before and (b) after hybridization with complementary sequences. Buffer, A9′, B18′, C9′, C18′, and N9, were added to samples 1-6, respectively.

A9′C9′ linker, which is also still in solution). Even though a 10-fold excess of A9′ was added over the A9′C9′ linker, cooperativity due to the high density of binding sites on the particles, which favors assembly with the aggregate, is strong enough to have a significant effect. The addition of the other complementary sequences results in low absorbances similar to the buffer and noncomplementary samples, verifying that B18′, C9′, and C18′ do not prevent completion of the TPA process at 25 °C.54 Only the addition of the complement to the A9:Au conjugates prevents the completion of the assembly process. In the absence of the aggregate, complete assembly cannot occur (Figure 7, bottom). As in the previous experiment, the result for A9′ addition differed substantially from the other samples. In this case, no aggregation occurred, which we attribute to blocking of available hybridization sites on the predominantly A9:Au conjugates in solution. Note that essentially no loss of particles occurs in this sample: the slight decrease in absorbance is due to dilution upon addition of A9′. A much smaller, but still substantial decrease in absorbance due to bioconjugate assembly was also observed for all of the other samples.55 The fact that TPA could continue at all in the absence of an aggregate suggests that A9:Au is not the only type of particle remaining in solution. We envision two possible explanations. First, there may be a small population of C9/18: Au conjugates that were not incorporated into the initial aggregate during the 65 °C hybridization (a large population is precluded by both gel electrophoresis and the data in Figure 7, top). However, large numbers of A9:Au conjugates can be cross-linked with only a few C9/18:Au conjugates. When left at room temperature with no competing interaction, these would assemble with the A9:Au conjugates via the Nano Lett., Vol. 6, No. 1, 2006

Scheme 2.

Figure 7. Absorbance spectra of TPA samples after 65 °C hybridization followed by incubation at 25 °C with buffer or oligonucleotides in the presence (top panel) and absence (bottom panel) of aggregates formed during 65 °C hybridization. (a) All six samples after 65 °C hybridization (in the bottom panel, these samples have not yet been split up). (b) Sample after hybridization with complementary sequence A9′. (c) Sample after hybridization with complementary sequence C9′. (d) Samples after hybridization with buffer, B18′, C18′, and N9′, respectively. Both insets show the original TPA mixture that corresponds to each sample set before hybridization at 65 °C. The same volume of buffer or DNA was added to each of the b, c, and d samples.

A9′C9′ linker. Because only a small minority of the particles left in solution could be the C9/18:Au conjugates, there would not be enough to hybridize to all of the A9:Au conjugates, so only a partial decrease in absorbance is observed. This behavior could also result from DNA exchange between the various types of conjugates in the initial mixture. Because there is a low density of exchanged sequences on the particles in solution, cooperativity is greatly diminished, if not eliminated, for these “incorrect” interactions. The gel in Figure 3 suggests that the number of C9/18:Au conjugates remaining is solution after the 65 °C hybridization is much less than the number of A9:Au conjugates. Even considering exchange of oligonucleotides between particles, in the actual TPA system, the surface density of exchanged sequences as compared with the “correct” initial sequences is quite small, such that cooperativity favors correct assembly. Even if some C9/18:Au conjugates are still in solution at 65 °C, the interaction between B18:Au and C9/18:Au conjugates still occurs first, and the sequence in time of the assembly events is thus controlled. Although three-strand hybridization systems such as those described above are well documented in the literature,56 Nano Lett., Vol. 6, No. 1, 2006

Three-Strand and Two-Strand DNA Hybridization Systems

directly complementary two-strand systems are also possible and are more straightforward for assembly purposes. Temperature-programmed experiments were repeated with directly complementary oligonucleotides (Scheme 2). To demonstrate that the controlling factor in this assembly is temperature, rather than DNA sequence, we took the hybridizing portions of the shorter sequences, D9 and D9′ (Tm ) 42 °C), from the longer sequences, D21 and D21′(Tm ) 84 °C), respectively (Table 1). PolyA spacers were added between the hybridizing sequence and the thiol so that the spacing between the particles after hybridization would be approximately the same for both duplexes. It should be noted that cross hybridization (eg. D9 with D21′) is possible. However, because the stability of a D9:D21′ pair is the same as that for a D9:D9′ pair, they can form only at lower temperatures. The results for the two-strand TPA experiments were very similar to the three-strand system described above. Gel electrophoresis indicates that, for samples incubated at a temperature between the Tms for the two duplexes, the D9: Au conjugates remain in solution (data not shown). At 25 °C, all of the conjugates participate in aggregation. Figure 8 shows two distinct melt transitions during thermal denaturation of samples incubated at 25 °C. The separation between the two melting transitions leaves room for additional duplexes in one assembly sample, which would provide greater control over the order of assembly events. The feasibility of temperature-programmed assembly has been demonstrated for nanoparticle assembly in solution, using both three-strand and two-strand hybridization systems. We have also shown that TPA can be performed in conjunction with or in the absence of sequence differences, allowing for a wide variation in assembly design. Despite the instability of gold-thiol attachment chemistry at elevated temperatures, cooperativity effects resulted in the desired order of nanoparticle assembly. Alternative attachment chemistries such as silane-based strategies on SiO2 particles, SiO2-coated metal particles, or glass surfaces53 would eliminate DNA strand exchange between nanoparticles and improve the robustness of the approach. Although unnecessary for the experiments described here, inert oligonucleotide attachment chemistries would be desirable for assembly of larger numbers of different building blocks. Alternatively, thiol exchange could be reduced by using lower salt buffers to lower the Tms. TPA could prove to be an important tool in bottom-up assembly strategies of more complex nanostructures. In these proof-of-principle experiments, the aggregates formed via 21

Figure 8. Thermal denaturation (top) and 1st derivative plot (bottom) for the 25 °C hybridization of D9:Au, D21:Au, and D9′/21′:Au.

temperature-programmed assembly are not functionally distinct; however, by incorporating more elaborate particle combinations into the assembly strategy, functionally distinct nanostructures will be possible. Acknowledgment. This work was supported by the NSF (MCB-0074845), DARPA/ONR (contract no. ONR-N0001498-1-00846), NSF-NIRT (CCR-0303976), and Pennsylvania State University. C.D.K. also acknowledges support from a Beckman Foundation Young Investigator Award, a Sloan Fellowship, and a Camille Dreyfus Teacher-Scholar Award. Supporting Information Available: Experimental details and kinetic data for the hybridization of the three-particle TPA system. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) (a) Metal Nanoparticles: Synthesis, Characterization, and Applications; Feldheim, D. L., Foss, C. A., Jr., Eds; Marcel Dekker: New York, 2002. (b) Nanobiotechnology: Concepts, Applications, and PerspectiVes; Niemeyer, C. M., Mirkin, C. A., Eds.; Wiley-VCH: Weinheim, Germany, 2004. (2) Ginger, D. S.; Zhang, H.; Mirkin, C. A. Angew. Chem., Int. Ed. 2004, 43, 30-45. (3) Yin, Y.; Lu, Y.; Gates, B.; Xia, Y. J. Am. Chem. Soc. 2001, 123, 8718-8729. (4) Cui, Y.; Bjork, M. T.; Liddle, J. A.; Sonnichsen, C.; Boussert, B.; Alivisatos, A. P. Nano Lett. 2004, 4, 1093-1098. (5) (a) Kovtyukhova, N. I.; Mallouk, T. E. Chem.sEur. J. 2002, 8, 4354-4363. (b) Kovtyukhova, N. I.; Martin, B. R.; Mbindyo, J. K. N.; Smith, P. A.; Razavi, B.; Mayer, T. S.; Mallouk, T. E. J. Phys. Chem. B 2001, 105, 8762-8769. (6) Huang, Y.; Duan, X.; Cui, Y.; Lauhon, L. J.; Kim, K.; Lieber, C. M. Science 2001, 294, 1313-1317. (7) Wu, Y.; Yan, H.; Huang, M.; Messer, B.; Song, J. H.; Yang, P. Chem.sEur. J. 2002, 8, 1260-1268. (8) Xia, Y.; Yang, P.; Sun, Y.; Wu, Y.; Mayers, B.; Gates, B.; Yin, Y.; Kim, F.; Yan, H. AdV. Mater. 2003, 15, 353-389. 22

(9) Genov, D. A.; Sarychev, A. K.; Shalaev, V. M.; Wei, A. Nano Lett. 2004, 4, 153-158. (10) Wang, W.; Gu, B.; Liang, L.; Hamilton, W. J. Phys. Chem. B 2003, 107, 3400-3404. (11) (a) Kim, F.; Kwan, S.; Akana, J.; Yang, P. J. Am. Chem. Soc. 2001, 123, 4360. (b) Kim, B.; Tripp, S. L.; Wei, A. J. Am. Chem. Soc. 2001, 123, 7955-7956. (12) (a) Whang, D.; Jin, S.; Lieber, C. M. Nano Lett. 2003, 3, 951-954. (b) Whang, D.; Jin, S.; Wu, Y.; Lieber, C. M. Nano Lett. 2003, 3, 1255-1259. (13) Ryan, D.; Nagle, L.; Rensmo, H.; Fitzmaurice, D. J. Phys. Chem. B 2002, 106, 5371-5377. (14) Smith, P. A.; Nordquist, C. D.; Jackson, T. N.; Mayer, T. S.; Martin, B. R.; Mbindyo, J.; Mallouk, T. E. Appl. Phys. Lett. 2000, 77, 13991401. (15) Hermanson, K. D.; Lumsdon, S. O.; Williams, J. P.; Kaler, E. W.; Velev, O. D. Science 2001, 294, 1082-1086. (16) Chung, S.-W.; Ginger, D. S.; Morales, M. W.; Zhang, Z.; Chandrasekhar, V.; Ratner, M. A.; Mirkin, C. A. Small 2005, 1, 64-69. (17) (a) Niemeyer, C. M. Angew. Chem., Int. Ed. 2001, 40, 4128-4158. (b) Ryadnov, M. G.; Ceyhan, B.; Niemeyer, C. M.; Woolfson, D. N. J. Am. Chem. Soc. 2003, 125, 9388-9394. (18) Mann, S.; Shenton, W.; Li, M.; Connolly, S.; Fitzmaurice, D. AdV. Mater. 2000, 12, 147-150. (19) Stevens, M. M.; Flynn, N. T.; Wang, C.; Tirrell, D. A.; Langer, R. AdV. Mater. 2004, 16, 915-918. (20) Caswell, K. K.; Wilson, J. N.; Bunz, U. H. F.; Murphy, C. J. J. Am. Chem. Soc. 2003, 125, 13914-13915. (21) Wang, Y.; Tang, Z.; Tan, S.; Kotov, N. A. Nano Lett. 2005, 5, 243248. (22) Wang, J. Small 2005, 1, 1036-1043. (23) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. Nature 1996, 382, 607-609. (24) Alivisatos, A. P.; Johnsson, K. P.; Peng, X.; Wilson, T. E.; Loweth, C. J.; Bruchez, M. P.; Schultz, P. G. Nature 1996, 382, 609-611. (25) Mucic, R. C.; Storhoff, J. J.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1998, 120, 12674-12675. (26) Le, J. D.; Pinto, Y.; Seeman, N. C.; Musier-Forsyth, K.; Taton, T. A.; Kiehl, R. A. Nano. Lett. 2004, 4, 2343-2347. (27) (a) Loweth, C. L.; Caldwell, W. B.; Peng, X.; Alivisatos, A. P.; Schultz, P. G. Angew. Chem., Int. Ed. 1999, 38, 1808-1812. (b) Zanchet, D.; Micheel, C. M.; Parak, W. J.; Gerion, D.; Williams, S. C.; Alivisatos, A. P. J. Phys. Chem. B 2002, 106, 11758-11763. (28) (a) Mbindyo, J. K. N.; Reiss, B. D.; Martin, B. R.; Keating, C. D.; Natan, M. J.; Mallouk, T. E. AdV. Mater. 2001, 13, 249-254. (b) Goodrich, G. P.; Helfrich, M. R. Overberg, J. J.; Keating, C. D. Langmuir 2004, 20, 10246-10251. (29) (a) Mirkin, C. A. Inorg. Chem. 2000, 39, 2258-2272. (b) Storhoff, J. J.; Mirkin, C. A. Chem. ReV. 1999, 99, 1849-1862. (30) Seeman, N. C. Acc. Chem. Res. 1997, 30, 357-363. (31) (a) Niemeyer, C. M.; Ceyhan, B.; Hazarike, P. Angew. Chem., Int. Ed. 2003, 42, 5766-5770. (b) Niemeyer, C. M.; Ceyhan, B.; Noyong, M.; Simon, U. Biochem. Biophys. Res. Commun. 2003, 311, 995999. (32) (a) Hamad-Schifferli, K.; Schwartz, J. J.; Santos, A. T.; Zhang, S.; Jacobson, J. M. Nature 2002, 415, 152-155. (b) Keren, K.; Berman, R. S.; Buchstab, E.; Sivan, U.; Braun, E. Science 2003, 302, 13801382. (33) Ongaro, A.; Griffin, F.; Nagle, L.; Iacopino, D.; Eritja, R.; Fitzmaurice, D. AdV. Mater. 2004, 16, 1799-1803 (34) (a) Reichert, J.; Csaki, A.; Kohler, J. M.; Fritzsche, W. Anal. Chem. 2000, 72, 6025-6029. (b) He, L.; Musick, M. D.; Nicewarner, S. R.; Salinas, F. G.; Benkovic, S. J.; Natan, M. J.; Keating, C. D. J. Am. Chem. Soc. 2000, 122, 9071-9077. (c) Taton, T. A.; Mirkin, C. A.; Letsinger, R. L. Science 2000, 289, 1757-1760. (d) Cao, Y. C.; Jin, R.; Mirkin, C. A. Science 2002, 297, 1536-1540. (35) Li, M.; Mann, S. J. Mater. Chem. 2004, 14, 2260-2263. (36) Deng, Z.; Mao, C. Nano. Lett. 2003, 3, 1545-1548. (37) Kannan, B.; Kulkarni, R. P.; Majumdar, A. Nano Lett. 2004, 4, 15211524. (38) Li, Z.; Mirkin, C. A. J. Am. Chem. Soc. 2005, 127, 11568-11596. (39) Claridge, S. A.; Goh, S. L.; Frechet, J. M. J.; Williams, S. C.; Micheel, C. M.; Alivisatos, A. P. Chem. Mater. 2005, 17, 1628-1635. (40) Sadasivan, S.; Dujardin, E.; Li, M.; Johnson, C. J.; Mann, S. Small 2005, 1, 103-106. (41) Deng, Z.; Tian, Y.; Lee, S.-H.; Ribbe, A. E.; Mao, C. Angew. Chem., Int. Ed. 2005, 44, 3582-3585.

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(42) Pena, S. R. N.; Raina, S.; Goodrich, G. P.; Fedoroff, N. V.; Keating, C. D. J. Am. Chem. Soc. 2002, 124, 7314-7323. (43) Bloomfield, V. A.; Crothers, D. M.; Tinoco, I., Jr. Nucleic Acids: Structures, Properties and Functions; University Science Books: Sausalito, CA, 2000. (44) Elghanian, R.; Storhoff, J. J.; Mucic, R. C.; Letsinger, R. L.; Mirkin, C. A. Science 1997, 277, 1078-1081. (45) Mucic, R. C.; Storhoff, J. J.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1998, 120, 12674-12675. (46) Jin, R.; Wu, G.; Li, Z.; Mirkin, C. A.; Schatz, G. A. J. Am. Chem. Soc. 2003, 125, 1643-1654. (47) Zanchet, D.; Micheel, C. M.; Parak, W. J.; Gerion, D.; Williams, S. C.; Alivisatos, A. P. J. Phys. Chem. B 2002, 106, 11758-11763. (48) Zanchet, D.; Micheel, C. M.; Parak, W. J.; Gerion, D.; Alivisatos, A. P. Nano Lett. 2001, 1, 32-35. (49) Sandstro¨m, P.; Åkerman, B. Langmuir 2004, 20, 4182-4186. (50) Demers, L. M.; Mirkin, C. A.; Mucic, R. C.; Reynolds, R. A., III; Letsinger, R. L.; Elghanian, R.; Viswanadham, G. Anal. Chem. 2000, 72, 5535-5541. (51) Parak, W. J.; Pellegrino, T.; Micheel, C. M.; Gerion, D.; Williams, S. C.; Alivisatos, A. P. Nano Lett. 2003, 3, 33-36. (52) Note that two fluorescent strands bound in the room-temperature control experiment may be the result of exchange or may simply be due to nonspecific adsorption. In the latter case, actual exchange would be lower (10%). (53) Andreadis, J. D.; Chrisey, L. A. Nucleic Acids Res. 2000, 28, e5ie5viii. (54) One curious result of this experiment is that the addition of C9′ does not prevent further assembly of the particles. This sequence should compete with the A9′C9′ linker for hybridization to the C9/18:Au conjugates just as A9′ competes with the same linker for hybridization

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to the A9:Au conjugates. However, addition of C9′ gave the same result as noncomplementary DNA or buffer alone (Figure 7, top). We believe this to be due to the greater accessibility of the A9:Au conjugates in solution relative to the aggregate. This is further evidenced by the repetition of this experiment with the samples shaking during the overnight hybridization with the complementary sequences. In this case, all samples, including that with A9′ added, showed complete assembly of the particles in solution to the aggregate. (55) When C9′ is added, again the result is similar to the samples with buffer and noncomplementary DNA, although it should be noted that the aggregation in this sample is slightly less than in the others. Again, one might imagine that the addition of C9′ would have the same result as the addition of A9′. However, when C9′ is added, the A9′C9′ linker hybridizes with the A9:Au conjugates, resulting in the majority of particles in solution with a high density of preorganized pieces for assembly and only a small minority of particles with the less favorable hybridization to the complementary sequence C9′. Alternatively, when A9′ is added, the A9′C9′ linker hybridizes with only half of the DNA on the C9/18:Au conjugates, resulting in a small minority of particles with a lower density of preorganized pieces for assembly and the majority of particles already fully hybridized with the complementary sequence A9′. It has been shown previously that a decrease in surface density of the oligonucleotide to be assembled results in a decreased stability and cooperativity in the resulting structure.45 (56) Rosi, N. L.; Mirkin, C. M. Chem. ReV. 2005, 105, 1547-1562.

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