Duplex Pinching: A Structural Switch Suitable for Contractile DNA

Such a “pinching” transition should provide a valuable “contractile” feature to the design of new, ... a DNA duplex,21,24 the pinched duplex t...
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Duplex Pinching: A Structural Switch Suitable for Contractile DNA Nanoconstructions

2003 Vol. 3, No. 8 1073-1078

Richard P. Fahlman,*,† Michael Hsing, Caroline S. Sporer-Tuhten, and Dipankar Sen Department of Molecular Biology & Biochemistry, Simon Fraser UniVersity, Burnaby, British Columbia V5A 1S6, Canada Received April 28, 2003; Revised Manuscript Received June 6, 2003

ABSTRACT A versatile role for guanine−guanine mismatches within DNA double helices such as the formation of specific, interhelical synaptic events has previously been reported. Here, we demonstrate that certain categories of guanine−guanine mismatches within double helices enable a different structural/conformational transitionsa reversible, intramolecular “pinching” of the duplex, initiated and reversed by the binding and dissociation of certain specific cations. Such a “pinching” transition should provide a valuable “contractile” feature to the design of new, DNA-based molecular motors and nanoconstructions.

Within the emergent field of nanotechnology, the nucleic acid polymer, DNA, has found surprising utility. Even prior to the widespread use of the term “nanotechnology”, DNA had been used in the assembly of novel geometric objects such as cubes.1 The underlying property of DNA that was exploited in nanostructure assembly was the highly specific and nucleotide sequence-dependent formation of duplex (double-helical) DNA via Watson-Crick base pairing. The rules of Watson-Crick pairing permit highly predictable associations of single-stranded sequences to form double helices. This property has enabled the “programmed” assembly of other materials. Nanoscale objects such as colloidal gold2-5 or quantum dots6 derivatized with single-stranded DNA have been spatially organized using DNA duplex assembly. Duplex DNA has also been successfully used to template the formation of silver7 and platinum wires.8 In combination with sequence specificity, the ability to generate stable branched structures (three-way and four-way immobile junctions) has led to the construction of a variety of structures created solely from DNA.9-11 In addition to the double helix, other structural/conformational motifs of DNA have also shown promise for nucleic acid-based nanostructure construction. Examples include guanine quadruplexes, which are made up guanine base quartets (G-quartets). G-quadruplexes have been used in the formation of G-wires,12 frayed wires,13,14 and synapsable * Corresponding author. E-mail: [email protected]. Tel: 847491-6762. † Current address: Department of Biochemistry, Molecular Biology & Cell Biology, Northwestern University, Evanston, Illinois 10.1021/nl034267i CCC: $25.00 Published on Web 06/28/2003

© 2003 American Chemical Society

DNA.15 Additionally, known RNA tertiary contacts have also been successfully used in forming RNA-based arrays.16,17 The ability to create objects on the nanometer scale is only one potential role of DNA in nanotechnology applications. DNA is also capable of controlled conformational changes, which, when harnessed, can be used to generate molecular motors and other mechanical devices. Conformational changes induced by the binding of small molecules or other oligonucleotides have been harnessed for the construction of allosteric nucleic acid enzymes18 as well as electrical sensors.19 Conformational changes have also been harnessed for the construction of DNA-based motors. One such device utilizes the salt-dependent B-DNA to Z-DNA transformation to generate rotational motion.20 Other examples have used the input of “fuel” DNA oligomers to induce structural changes relevant to the functioning of such motors.21-24 In this paper, we report the use of G-quartets to enable an unusual “pinching” conformational change in doublestranded DNA, which can be reversibly regulated in aqueous solution by the presence or absence of certain specific cations (i.e., K+ or Sr2+). Although G-quartets have been used to generate other conformational changes in DNA, for instance, the transition from a single-stranded intramolecular quadruplex to either an unstructured single-stranded sequence25 or a DNA duplex,21,24 the pinched duplex that we describe always exists within a double-stranded DNA context and therefore should prove particularly useful for the construction of “contractile” DNA nanostructures. A key feature of guanine base quartets (Figure 1A) that makes them attractive for use in regulated structure formation

Figure 1. (A) Guanine quartet. The association of the four participating guanine bases is stabilized by a total of eight hydrogen bonds and by the site- and cation-size-specific binding of ions such as K+ or Sr2+ in the internal cavity formed by two successive guanine quartets. The guanine bases are linked via their N-9 positions to deoxyribosephosphate backbones, which are depicted by ovoid shapes. (B) Synapsable duplexes. Certain duplexes possessing internal guanine-guanine (G‚G) mismatches are able to dimerize to form a synapsed duplex dimer via the formation of intermolecular guanine quartets. (C) Pinched duplex, where certain duplexes possessing G‚G mismatches along a stretch are able to fold back upon themselves to form intramolecular guanine quartets.

is their requirement for certain specific cation species (i.e., K+ or Sr2+) in order to form. A G-quartet consists of four guanine bases associated into a planar structure, stabilized by a total of eight hydrogen bonds (Figure 1A). The stacking of two adjacent G-quartets results in a cation binding cavity, which, when occupied by a cation of the correct size, dramatically increases the stability of the quadruplex.26,27 Typically, most single-stranded DNA sequences containing stretches of guanosine nucleotides are able to form these structures, but association is not highly specific, per se, other than the observation that the formation of certain conformers is favored by certain cation species.28 Previously, we demonstrated that duplex DNA incorporating guanineguanine mismatches are able to dimerize with similar mismatch-containing duplexes via the formation of interduplex G-quartets in the presence of potassium15 (Figure 1B). We have also demonstrated that by “patterning” G‚G mismatches by introducing thymine-thymine (T‚T) mismatches within them the G‚G domains can be made selfselective for dimerization such that a given domain will 1074

dimerize with another identical domain and will not dimerize easily with duplexes possessing other G‚G mismatch patterns.29 In our earlier work,29,30 we had also shown preliminary evidence that certain G‚G domains dimerized with a lower efficiency than expected, likely owing to the preferential formation of intramolecular G-quartets, resulting in what we termed a pinched duplex (Figure 1C). The formation of such a structure might be enabled if the mismatch domain was long enough to allow folding back on itself. For a more detailed description of G-quartets and their properties, one is referred to various review articles.26,27 We wished to investigate further the requirements for duplex-duplex synapsis. For this study, we designed a series of synapsable duplexes, each containing a total of six G‚G mismatches with T‚T mismatches flanking either side and acting as buffers between the G‚G domain and the two Watson-Crick base-paired stems. The four duplex constructs differed in that the G‚G mismatches were disrupted by the insertion of zero to three A‚T base pairs (Figure 2A). The resulting duplexes were designated as TA-0, TA-1, TA-2, and TA-3. Constituent single strands within these duplexes are individually identified, for instance, as T-3 for the upper strand and A-3 for the lower strand in the case of the TA-3 duplex (Figure 2A). The four kinds of duplexes mentioned above were constructed in two sizes such that they were either 53 base pairs long (TA-0, TA-1, TA-2, and TA-3) or 43 base pairs long (TA-0-s and TA-2-s). Long and short duplexes were utilized together such that a cross-dimerized species formed by a long and a short duplex (for instance, TA-1‚TA-2-s) could be distinguished electrophoretically by possessing intermediate mobility from the homosynapsed dimers (TA-1)2 and (TA-2-s)2). The complete sequences of individual duplexes used are listed in Supporting Information. Duplexes were assembled in buffer A (10 mM Tris-Cl, pH 7.9, 100 mM tetramethylammonium chloride and 0.1 mM EDTA) with one of the two constituent single strands carrying a 32P label at its 5′ end. (The kinasing and purification procedures were as previously described.29) When individual duplexes were incubated overnight at 37° C in buffer A containing 1 M LiCl followed by analysis on nondenaturing PAGE, none of the duplexes were found to form synapsed dimers (data not shown). Following incubations in 1 M KCl, however, slowly migrating synapsed duplex dimers were observed by nondenaturing PAGE (with the exception of the TA-3 duplex, which, under no conditions tested, appeared to form synapsed duplexes (data not shown)). Figure 2B displays the nondenaturing gel electrophoretic results of incubations in 1 M KCl combining the short TA-0-s duplex with, in turn, three longer synapsable duplexes (TA-1, TA-2, and TA-3). Figure 2B also shows analogous experiments using the short TA-2-s duplex instead of TA-0-s. The results indicate that (a) duplexes possessing a TA-0 domain dimerize only with other TA-0 domains; (b) TA-1 and TA-2 duplexes, however, exhibit cross reactivity with each other, as demonstrated by the formation of the hybrid synapsed duplex TA-2-s‚TA-1 (indicated in Figure 2B with an asterisk); and (c) duplexes containing the TA-3 domain do not dimerize either with themselves (vide the lane Nano Lett., Vol. 3, No. 8, 2003

Figure 2. (A) Design of G‚G domains. Domains consisting of six G‚G mismatches are flanked by pairs of thymine-thymine (T‚T) mismatches on each side. In different domains, the stretch of G‚G mismatches is interrupted by 1, 2, or 3 A‚T base pairs (these domains are named TA-1, TA-2, and TA-3, respectively) or left uninterrupted as a stretch of six G‚G contiguous mismatches (named TA-0). All G‚G domains are flanked by double-stranded arms of sufficient length to make up duplexes that are either 43 or 53 base pairs long. In all cases, the upper strand depicted is termed the T strand, and the lower strand, the A strand. (B) Screening for dimerization. Samples were 1 µM in each duplex and 1 M KCl in buffer A (10 mM Tris-Cl, pH 7.9, 100 mM tetramethylammonium chloride and 0.1 mM EDTA) and were incubated at 37 °C for at least 12 h. Samples where then analyzed by nondenaturing gel electrophoresis, as previously described.29 Dried gels were visualized by phosphorimaging. Undimerized duplexes as well as synapsed duplexes are indicated; the lower (faster migrating) electrophoretic band of synpased duplexes correlates to the dimer product of the 43 base pair duplex and of the 53 base pair duplex for the upper electrophoretic band of synapsed duplexes. The hybrid synapsed product that formed between the short TA-2-s duplex and the long TA-1 and TA-2 duplexes is indicated with an asterisk. (C) Chemical probing of the TA-3 duplex when the T-3 strand is 32P-5′-labeled. Samples of the duplex (1 µM) in buffer A were made up to either 1 M LiCl or KCl and were incubated at 37 °C overnight prior to reaction with DMS or KMnO4 or with no chemical probing reagent added (control). All samples, including the control, were treated with hot piperidine (at 90 °C), lyophilized, and then analyzed by denaturing PAGE. (D) Identical to C, but the TA-3 duplex was 32Pend-labeled on the A-3 strand. This duplex was also treated with DEPC to examine the disposition of the adenosine bases within the G‚G domain. Nano Lett., Vol. 3, No. 8, 2003

showing mixed incubation of TA-0-s and TA-3 in Figure 2B) or with duplexes incorporating other dimerization domains. The ability of TA-0 domains to discriminate between “self” and “other” (such as the TA-1 and TA-2 domains) is understandable in terms of thermodynamics, given that the formation of a maximum of six contiguous G-quartets by TA-0 can be achieved only via self-dimerization. The A‚T internal spacers in TA-1 and TA-2 would discourage the alignment of all available G‚G mismatches so that, likely, only a suboptimal number of G-quartets could form in, say, a TA-0‚TA-2 heterodimer. Conversely, the lack of specificity in dimerization between TA-1 and TA-2 is likely the result of the flexibility of the A‚T spacer bases. In TA-1‚TA-2 heterodimers, one set of adenosine and thymine bases may be extruded to allow for the aligning of G‚G mismatches to form the six G-quartets possible between these domains. There is some evidence in the literature that intervening bases separating stacks of G-quartets may be displaced out of the helical structure in this way.31 The inability of TA-3 domains to dimerize under a wide variety of salt conditions tested (data not shown) is consistent with the preferential formation of an alternative, stable, intramolecular structure. Such a potential structure, a “pinched duplex”, is depicted in Figure 1C. To investigate the formation of such a structure in KCl-containing buffers, chemical probing experiments were carried out on this duplex. Either the T-3 or the A-3 component strand of the TA-3 duplex was 5′ labeled with 32P and then subjected to partial covalent modification with dimethyl sulfate (DMS), diethyl pyrocarbonate (DEPC), and potassium permanganate (KMnO4). The reactivity of specific DNA bases with these chemicals (guanine with DMS, adenine with DEPC, and thymine with KMnO4) depends on the conformation and hydrogen-bonding status of the base concerned. Nucleobases modified by these reagents lead to the lability of the intact DNA strands at their sites when heated in aqueous base. Figure 2C shows that when the TA-3 duplex (with a 32P label on its T-3 strand) was incubated in a LiCl-containing buffer all guanine bases in the duplex were reactive to DMS (as observed by the cleavage of the DNA strand at every single guanine residue). In the presence of KCl, only the six guanines in the G‚G domain became resistant to methylation (as demonstrated by a significant loss of DNA cleavage at these base sites), whereas the guanines in the duplex regions remained reactive to DMS. Such potassium-dependent methylation protection is a signature of G-quartet formation.32 Other probing experiments, where the duplex was reacted with KMnO4, showed that in lithium-containing buffer (the concentration of added KMnO4 was insignificant compared to the lithium concentration) only the thymines within the G‚G domain were reactive to KMnO4, consistent with the expectation of greater conformational mobility of these thymines relative to that of thymines base-paired within Watson-Crick duplex regions.33 In the presence of KCl, however, hyperreactivity was observed at the two 5′-most thymines of the A‚T “spacer” element within the G‚G domain. The thymines separating the G‚G domain from the 1075

Watson-Crick domain in the 5′ direction also exhibited increased reactivity. Such observed hyperreactivity to KMnO4 in potassium-containing buffers suggests that the duplex has undergone a structural transition that enables a significantly greater solvent accessibility to the abovementioned thymines. Such a scenario is consistent with the model of a “pinched duplex” shown in Figure 1C. The most compelling evidence for the formation of a pinched duplex, however, comes from chemical probing of the TA-3 duplex, where the constituent A-3 strand has a 32P label on its 5′ end. Figure 2D shows the results of these experiments. The reagent DEPC is known preferentially to target adenines that are present in loops, bulges, and singlestranded regions of DNA;34 adenines base-paired in WatsonCrick duplexes are generally unreactive toward DEPC.34 When the TA-3 duplex was treated with DEPC in lithium solution, only a small difference in the DEPC reactivity of the spacer adenines and the adenines in the flanking duplex regions could be observed. However, when TA-3 was treated with DEPC in the presence of potassium, the two most 5′ adenines in the A‚T spacer showed a strong hyperreactivity, suggesting a large structural change commensurate with the greatly enhanced access of solvent to these adenines. A simple model for such hyperreactive adenines would have them jutting out into the solution, as illustrated in the pinched duplex shown in Figure 1C. The involvement of the G‚G domain guanines of TA-3 in G-quartet formation in potassium solution, yet not forming the synapsed dimers, and the gross structural changes observed on the thymines and adenines within and flanking the G‚G domain cumulatively indicate the formation of an intramolecular pinched duplex, such as that depicted in Figure 1C. What is particularly notable about the TA-3 G‚ G domain is that it readily forms the pinched duplex while remaining completely incapable of forming the intermolecularly synapsed duplex dimers. This singular property of the TA-3 G‚G domain suggests that it could find utility in the construction of nucleic acid superstructures in which a pinching conformational change could be readily induced without concern for the G‚G domains also dimerizing, in a competing process, to give undesired aggregated products. To determine whether the pinching of the TA-3 G‚G domain could be easily reversed, we studied the properties of the TA-3 duplex in buffered solutions containing 5 mM strontium chloride rather than 1 M potassium chloride. The divalent strontium cation (unlike magnesium or calcium) strongly promotes G-quartet formation and does so at concentrations much lower than those required for potassium.35,36 A further advantage that Sr2+ offers is that the free hydrated ion can be cleanly sequestered by a strong chelator such as EDTA. The TA-3 duplex, with its A-3 strand 5′-labeled with 32P, was first probed, in separate experiments, with DMS and DEPC to define whether the pinched duplex formed in Mg2+or Sr2+-containing buffer solutions. Figure 3A shows that the characteristic signatures for the formation of a TA-3 pinched duplex are present when the duplex is incubated in 5 mM SrCl2 for 30 min at 37 °C. The characteristic DMS 1076

Figure 3. (A) Reversibility of formation of the pinched duplex formed by TA-3. Duplexes containing a 32P label on the 5′ end of the A-3 strand were probed with either DMS or DEPC as described in Figure 2 and were examined by denaturing PAGE. The control lane shows the background cleavage produced by hot piperidine treatment. Lanes 1 and 5 are of samples incubated for 30 min at 37 °C in the presence of 5 mM SrCl2 in buffer A. Lanes 2 and 6 are of samples identical to those in 1 and 5, but following incubation with SrCl2, 12-µL samples were diluted with 3 µL of 100 mM EDTA and incubated for an additional hour prior to chemical probing. Lanes 3 and 7 show samples incubated in 5 mM MgCl2 instead of SrCl2. Lanes 4 and 8 show samples preincubated in MgCl2 prior to incubation in excess EDTA, as with 2 and 6 above. (B) Schematic illustrating the reversibility of pinched duplex formation. Control over the formation of a pinched duplex is simply achieved by the addition of a G-quartet-promoting cation such as Sr2+. The process can be reversed by the removal of the cation from solution by using a chelator such as EDTA. The angle between the flanking duplexes is proposed only because there is currently no data determining the overall extent of DNA bending.

methylation protection observed with the guanines in the G‚ G domain (lane 1) as well as the DEPC hyperreactivity of the adenines in the A‚T spacer region (lane 5) are observed. Subtle differences between these data and those obtained in 1 M potassium were noted; however, this strontium protection pattern persisted even when the sample was incubated for more than 18 h in strontium under the conditions described above (data not shown), and again, no formation of the synapsed dimer of TA-3 was observed. Following incubation in 5 mM Sr2+, the TA-3 sample was treated with excess EDTA, and both characteristic probing patterns of the pinched duplex disappeared (lanes 2 and 6). Interestingly, when exactly analogous experiments were carried out with Mg2+ replacing Sr2+, no differences in the chemical probing patterns were observed either before (lanes 3 and 7) or after (lanes 4 and 8) EDTA treatment, and there was also no indication of the formation of a pinched duplex. These results demonstrate the ability of the TA-3 duplex to undergo a structural cycle from an open linear conformation to a pinched duplex and back by the addition and removal of specific cation species such as K+ and Sr2+, as depicted in Figure 3B. The actual degree of distortion or bending of the linear axis of the duplex DNA is currently undetermined Nano Lett., Vol. 3, No. 8, 2003

Figure 4. Schematics for the potential incorporation of G‚G domains into larger DNA assemblies to serve as a device for inducing regulated structural changes in (A) a contractile fiber and (B) a contractile sheet. These examples have been chosen to highlight the potential of contraction that can be achieved when the units of assembly either can (as in four-way junctions) or cannot (as in double crossovers) be deformed. In both kinds of architecture, the upper “extended” forms result from the incorporated G‚G domains existing in an extended conformation. Upon the addition of Sr2+, the G‚G mismatches form pinched duplexes as a result of intramolecular guanine quartet formation. Formation of the pinched duplexes results in an overall contraction of the structures. Upon the removal of Sr2+, the structure reverts to the original extended form.

and may depend significantly on the presence or absence of the flanking T‚T mismatches. The ability to induce a defined and “contractile” conformational change within a DNA double-stranded region by the addition or removal of specific cations may prove useful with larger DNA architectures. It should be possible to incorporate multiple G‚G domains into a single DNA double helix as well as into different DNA nanostructures to generate contractile DNA fibers or sheets. Architectures simultaneously incorporating immobile fourway junctions and G‚G mismatch domains such as TA-3, as depicted in Figure 4, could allow for the creation of such devices. For these applications, the G‚G domains may require the removal of the T‚T mismatches flanking the G‚G domains because the dynamic mobility of these flanking T‚T mismatch regions could significantly reduce the global level of duplex bending. A contractile fiber, as shown schematically in Figure 4A, would, in the absence of the cations that promote G-quartet formation, be expected to adopt a long, extended structure. The design of such a construct, based on DNA four-way (Holliday) junctions, is shown in Figure 4A. (The structure and properties of immobile four-way junctions have been reviewed in detail by Lilley.37) Upon the addition of a cation such as Sr2+, the repeating G‚G domains within this construct would be expected to pinch, resulting in a global contraction of the structure. Contraction would require overcoming the natural preference of four-way junctions to stack in a certain way, but the energetic barrier for changes to the preferred stacking-mode switch is small.38 We would anticipate that the formation of highly stable intramolecular G-quartets by the G‚G domains should easily drive the system into the contracted state depicted in Figure 4A. G‚G domains could also be used to bridge conformationally rigid structures such as DNA double crossovers. Double Nano Lett., Vol. 3, No. 8, 2003

crossovers are closely spaced pairs of four-way junctions that are topologically more constrained and hence structurally less dynamic than single four-way junctions (reviewed by Seeman39). Double crossovers have been used with great success as building blocks for the construction of DNA 2D arrays.10,11,40 Interspersing G‚G domains between adjacent double crossover “tiles” would enable the construction of reversibly contractile DNA sheets (shown schematically in Figure 4B). In summary, we have demonstrated the ability of a new class of G‚G mismatch domains to undergo a reversible, “contractile” structural change induced by specific G-quartet stabilizing cations such as K+ or Sr2+. The inherent ability of G‚G mismatch domains to be incorporated within doublehelical DNA frameworks should allow for their facile integration into DNA nanostructures and should lead to new and contractile variants of such intricate DNA structures. Acknowledgment. This work was supported by the Natural Sciences and Engineering Research Council of Canada (NSERC). D.S. is a Senior Scholar of the Michael Smith Foundation for Health Research. R.P.F. was an NSERC postgraduate scholar during this work and is currently a Canadian Institute of Health Research (CIHR) postdoctoral fellow. Supporting Information Available: A listing of the complete DNA sequences used in the assembly of the different duplexes. Detailed procedures used for chemically probing the duplexes with DMS, DEPC, and KMnO4. This material is available free of charge via the Internet at http:// pubs.acs.org. References (1) Chen, J.; Seeman, N. C. Nature 1991, 350, 631-633. (2) Mirkin, C.A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. Nature 1996, 382, 607-609. (3) Alivisatos, A. P.; Johnsson, K. P.; Peng, X.; Wilson, T. E.; Loweth, C. J.; Bruchez, M. P., Jr.; Schultz, P. G. Nature 1996, 382, 609611. (4) Mucic, R. C.; Storhoff, J. J.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1998, 120, 12674-12675. (5) Loweth. C. J.; Caldwell. W. B.; Peng, X.; Alivisatos, A. P.; Schultz, P. G. Angew. Chem., Int. Ed. 1999, 38, 1808-1812. (6) Mitchell, G. P.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1999, 121, 8122-8123. (7) Braun, E.; Eichen, Y.; Sivan, U.; Ben-Yoseph, G. Nature 1998, 391, 775-778. (8) Mertig, M.; Ciacchi, L. C.; Seidel, R.; Pompe, W.; De Vita, A. Nano Lett. 2002, 2, 841-844. (9) Zhang, Y.; Seeman, N. C. J. Am. Chem. Soc. 1994, 116, 16611669. (10) Winfree, E.; Liu, F.; Wenzler, L. A.; Seeman, N. C. Nature 1998, 394, 539-544. (11) Mao, C.; Sun, W.; Seeman, N. C. J. Am. Chem. Soc. 1999, 121, 5437-5443. (12) Marsh, T. C.; Henderson, E. Biochemistry 1994, 33, 10718-10724. (13) Sen, D.; Gilbert, W. Biochemistry 1992, 31, 65-70. (14) Protozanova, E.; Macgregor R. B., Jr. Biochemistry 1996, 35, 1663816645. (15) Venczel, E. A.; Sen, D. J. Mol. Biol. 1996, 257, 219-224. (16) Jaeger, L.; Leontis, N. B. Angew. Chem., Int. Ed. 2000, 39, 25212524. (17) Jaeger, L.; Westhof, E.; Leontis, N. B. Nucleic Acids Res. 2001, 29, 455-463. (18) Soukup, G. A.; Breaker, R. R. Trends Biotechnol. 1999, 17, 469476. 1077

(19) Fahlman, R. P.; Sen, D. J. Am. Chem. Soc. 2002, 124, 4610-4616. (20) Mao, C.; Sun, W.; Shen, Z.; Seeman, N. C. Nature 1999, 397, 144146. (21) Li, J. J.; Tan, W. Nano Lett. 2002, 2, 315-318. (22) Yan, H.; Zhang, X. Shen, Z.; Seeman, N. C. Nature 2002, 415, 6265. (23) Yurke, B.; Turberfield, A. J.; Mills, A. P., Jr.; Simmel, F. C.; Neumann, J. L. Nature 2000, 406, 605-608. (24) Alberti, P.; Mergny, J. L. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 1569-1573. (25) Ueyama, H.; Takagi, M.; Takenaka, S. J. Am. Chem. Soc. 2002, 124, 14286-14287. (26) Wellinger, R.; Sen, D. Eur. J. Cancer 1997, 33, 735-749. (27) Simonsson, T. Biol. Chem. 2001, 382, 621-628. (28) Sen, D.; Gilbert, W. Nature 1990, 344, 410-414. (29) Fahlman, R. P.; Sen, D. J. Am. Chem. Soc. 1999, 121, 11079-11085.

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Nano Lett., Vol. 3, No. 8, 2003