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Construction of a Structurally Defined Double-Stranded DNA Catenane Thorsten L. Schmidt† and Alexander Heckel* Cluster of Excellence Macromolecular Complexes, Goethe-University Frankfurt, Max-von-Laue-Strasse 9, 60438 Frankfurt/M, Germany
bS Supporting Information ABSTRACT: Topologically interlocked structures like catenanes and rotaxanes are promising components for the construction of molecular machines and motors. Herein we describe the construction of double-stranded DNA catenanes for DNA nanotechnology. For this, C-shaped DNA minicircle fragments were equipped with sequence-specific DNA-binding polyamides and their respective binding site. Formation of catenanes is achieved by self-assembly of two of these fragments and subsequent addition of a ring-closing oligonucleotide. KEYWORDS: DNA nanotechnology, catenanes, topology, topologically interlocked structures, self-assembly
C
atenanes and rotaxanes are multicomponent structures, built up from topologically interlocked molecules. In rotaxanes, a macrocycle is threaded on the axle of a dumbbell-like structure and is held in place by bulky stoppers. In catenanes (from Latin catena), two or more macrocycles are interlocked like links of a chain. In macromolecular chemistry, catenanes and rotaxanes were realized synthetically decades ago1,2 and are now being investigated as components for molecular switches3 or molecular motors.4 DNA nanotechnology is a much younger field that uses DNA as a structural material for building nanometer-scale architectures.5 DNA, in contrast to other biopolymers like RNA or polypeptides, is not only programmable, but also structurally and functionally well behaved and is arguably the most versatile material for bottom-up construction of artificial objects in the nanometer-scale. Moreover, DNA can also be the basis for molecular motors,6 sensors, and logic devices7 and many other devices with a broad range of functions.8,9 Topologically interesting structures like Borromean rings,10 catenanes, and knots11,12 have already been built using DNA. However, generally single-stranded (ss) DNA was used to produce the topological macrocycle units. Examples of catenated structures from ssDNA include Seeman’s famous cube,13 the truncated octahedron,14 or catenated scaffolds for enzymes15,16 in which the catenated topology of individual strands is ensured by the hybridization of two ss rings over one or more helical turns into double-stranded (ds) DNA. So the macrocycles are not only catenated but also wound around each other at least once (the linking number Lk is 2 or higher). The double-stranded regions of such catenanes do not allow the macrocycles to move or rotate freely and single-stranded regions have a persistence length of only few nanometers. Those catenanes are therefore not suitable as mobile components for molecular machines. Exceptions to this ss design approach are the first dsDNA rotaxane that we r 2011 American Chemical Society
recently presented17 and very large Borromean rings and catenanes from DNA origami.18 Catenanes from dsDNA macrocycles generally have a linking number of 1 and no DNADNA interactions. They are also known to exist in biological contexts such as in kinetoplast DNA, a part of the mitochondrial DNA of certain protozoa, which is an extremely complex polycatenane of thousands of mini- and macrocircles.19 Topoisomerase enzymes that release torsional stress created during DNA replication can catalyze the interconversion of topological isomers of DNA,20 for example, knots and catenanes.21 In this case, the catenane or knot formation is reversible and can be controlled by polycationic additives.22 Another route to catenanes and knots is the recombination of supercoiled plasmids, such as the integration system of coliphage λ.23 All of the above examples produce catenanes from plasmids in the kilo base pair (kbp) or micrometer range, thus at least 1 order of magnitude longer than the persistence length of dsDNA (∼150 bp/50 nm). Therefore, also these catenanes are not suitable components for potential nanoarchitectures, both because they are too large and their shapes are not defined. The aim of this study was to construct catenanes made from small ds minicircles which are more than 1 order of magnitude smaller than plasmids. In previous studies, we established reliable syntheses of minicircles with a length of 168 bp or a diameter of 18 nm.2428 At this size in the vicinity of the persistence length, the shape remains close to an ideal circle and supercoiling was never observed. Therefore, a synthesis route with recombination enzymes that requires supercoiling prior to transient strand breaks cannot be applied here. To the best of our knowledge, topoisomerases have never been used with minicircles this small Received: January 26, 2011 Revised: March 12, 2011 Published: March 16, 2011 1739
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Figure 1. (a) Three different representations of the Dervan polyamide used in this study (1). The top left pictogram shows filled circles for imidazole residues and open circles for pyrrole residues. The turn is drawn as line, β-alanine as diamond, and N,N-dimethylaminopropylamine as half circle with a “þ”. In the next level of abstraction (top middle part), the polyamide is shown only as arrow in NC direction. The double-stranded binding site to which this polyamide binds is indicated in green. W (for weak) stands for A or T. (b) An oligonucleotide with an internal amino modification (2) is coupled to the isothiocyanate group of the polyamide (1) yielding the DNA-PA hybrid molecule as a thiourea derivative (3).
and we elected against using an enzymatic approach for catenating minicircles. Instead, we designed a system whereby open minicircle fragments preorganize in a manner that leads to catenanes upon closure of the rings. Design. We used Dervan-type polyamides (PA) as an orthogonal construction element to direct self-organization of open minicircle fragments. These PA are a class of heteroaromatic polymers that can sequence-selectively bind to the minor groove of DNA.29 They were already used to control gene activity29 or to align proteins on a DNA scaffold.30 In previous studies, we have connected two Dervan PA with a linker to a heterodimeric “strut” with the ability to hold two dsDNA objects together.27 Later, we improved the concept by connecting a PA covalently to an oligonucleotide and incorporating this hybrid molecule in minicircles. These covalently modified minicircles yielded complexes with an increased stability.28 In the current study, we applied covalently attached polyamides again (Figure 1). Instead of forming side-on homodimeric complexes, the PAs direct organization of C-shaped minicircle fragments that lack a single oligonucleotide for ring closure (Figure 2). These C-shaped fragments (7) are intrinsically curved due to A-tract sequences and were equipped with a polyamide anchor and the binding site. The orientation of the PA attachment was designed to face down and the (unique) PA binding site was facing up. This fragment should be able to form a homodimeric complex. The palindromic binding site allows two binding orientations but the point of attachment is hardly moved in space in both orientations. The length of the PA-DNA-linker and the orientations of the PA attachment site and the PA binding site were designed so that both potential PA-DNA bonds should be formed only when one fragment first crosses under and then over the other fragment (8). This interlocked complex can be
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Figure 2. (a) Schematic representation of the catenane synthesis. The polyamide-oligonucleotide hybrid (3) with oligonucleotide shown as orange strand and polyamide as green arrow was hybridized with three other oligonucleotides (46) to a C-shaped minicircle fragment (7). This fragment carried both the polyamide (PA) and the polyamide binding site. Two of these fragments could now form a homodimeric complex (8). To demonstrate the topology, one of the identical fragments is drawn in blue. Upon addition of oligonucleotide (9), the minicircles are completed and a catenane of double-stranded minicircles is formed (10). (b) A model of the central region of the catenane. The PA modification is printed in the same color as the minicircle to which it is covalently attached.
assumed to be rather flexible, but the two interactions act cooperatively. Moreover, neither the distance between the PA modification and the binding site, nor the remaining double stranded ends of the “C” exceed a third of the persistence length of DNA. Therefore we believed that the ends of the interlocked C-shapes (8) point away from another, as illustrated in Figure 2. Upon addition of the ring closure oligonucleotide to this complex, the fragments should close and form a pair of topologically interlocked minicircles, a catenane. The nicks present in the catenane could then be ligated for higher mechanical stability and subsequent analyses under denaturing conditions. Experimental Section. The electrophilic isothiocyanate modification of the polyamide (1) was obtained from the corresponding amine with CS2 and tosylchloride as described before.28 Then, a commercially available oligonucleotide with an internal amino modification (2) was coupled under slightly basic conditions to the isothiocyanate (1) yielding the PA-DNA hybrid molecule (3). This hybrid was then mixed with equimolar amounts of three other oligonucleotides (46), heated to 95 °C and then slowly cooled down to form the C-shaped fragment (7). After further incubation at 4 °C, the homodimeric complex was assumed to have hybridized. Isolation or further analysis of this intermediate was not performed. Instead, the ring-closing oligonucleotide (9) was added to form the catenane (10). Finally, the nicks of the minicircles were enzymatically ligated and the catenanes were isolated from unincorporated oligonucleotides, monomeric minicircles, and undefined aggregates by anion exchange chromatography. For details on the synthetic and analytical methods, we refer to the Supporting Information. The catenanes were deposited on mica positivized with polyornithine and imaged in air by atomic force microscopy (AFM). The AFM image (Figure 3) predominantly shows pairs of 1740
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Figure 3. (a) AFM image of the catenated minicircles (10). (b) As a control (11), an AFM image of side-on homodimeric complexes.28 (c) Denaturing PAGE-gel of the catenane (10) and the control complex (11). Whereas catenanes stay intact even under denaturing conditions, the control complex is quantitatively separated to individual minicircles. The components of the marker lane (M) have a much higher electrophoretic mobility than the minicircles or catenanes of the same length. The size of the marker fragments in nucleotides is indicated.
minicircles, but also some unconnected rings and ring fragments. We speculate that these may have been ripped apart during the workup or adsorption to the surface due to an incomplete ligation of one or more of the 10 nicks of the catenane. The reaction could potentially be treated with BAL-31 exonuclease after ligation28 to further increase the purity of completely ligated catenanes. However, this increase in purity would be expected to result in lower yield, as the exonuclease also has some endonuclease activity at nicks and was thus omitted here. As a control structure we imaged noninterlocked homodimeric complexes (11).28 These complexes are very similar and consist of minicircles of the same length (168 bp), carry one PA binding site and are modified with exactly the same PA that was used in this study. The difference to the catenane-minicircle is only a shorter distance between the PA modification and the PA binding site and that they are presented to the outer side of the tori. As a consequence, these minicircles form side on complexes (Figure 3b). However, the topology of the supposedly catenated dimers cannot be determined by AFM alone. Theoretically, it could be possible that pairs of rings lie on top of one-another without being catenated, only held in place by the PA-DNA interactions. To definitively prove the catenated topology, we performed denaturing polyacrylamide gel electrophoresis (PAGE) on both
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catenanes and the control complex (Figure 3c). Under denaturing conditions, hydrogen bonds are broken between base pairs and also between the PA and DNA. The comparison of the two lanes reveals a far lower electrophoretic mobility of the catenane than the control complex. Whereas the minicircles of the catenane are interlocked and cannot be separated, the control complex falls apart into unconnected minicircle monomers. Discussion and conclusion. The construction principle shown here was not our first attempt to construct catenanes. In our first attempt, we wanted to preorganize two C-shaped fragments with one homodimeric PA-strut.27 This attempt was not successful, presumably because of three reasons. First, the alignment of dsDNA with only one PA-strut is much weaker than with two cooperatively acting covalent PA. Second, the interlocked topology of two C-shaped fragments cannot be enforced effectively with only one connection instead of the two connections of the second design. And third, the ring closure efficiencies were not as high as for later designs. Taking into account the yields of every single step of the catenane synthesis, (1) the formation of the C-shaped fragment, (2) the complex formation of two fragments, (3) the formation of the interlocked topology of the fragments, and (4) the ring closure of both of the rings have to be high as every error decreases the overall yield, the failure of the first design becomes understandable in retrospect. In the second design presented here, high-performance liquid chromatography (HPLC) determined that the catenane was the main reaction product (Supporting Information). This design is therefore highly robust and may be applicable in future projects, such as the construction of molecular motors or other nanoscale machinery. For these applications, it may be necessary to have a free rotation of the catenated macrocycles. Here, the rotation of the catenane is presumably locked at low temperatures. At temperatures well above room temperature, the PA-DNA-bonds are dissociated and the minicircles may rotate freely. Earlier studies31 also imply a pH-dependency of the PA-DNA interaction. In our study here, the assembly of the control complex (11) was fastest at pH 5.4, slowed down at pH 7.4, and did not take place effectively any more at a pH of 8.2. Furthermore, heat denaturation of the control complex (11) and rapid cooling to room temperature also resulted in separate minicircles in the AFM analysis (see Supporting Information). However, incubation of this sample for 12 h at 4 °C resulted again in a nearly quantitative formation of the expected twin ring assemblies. Therefore, we believe that also the catenanes have a high degree of freedom to rotate in a buffer slightly more basic than pH 7.4, upon addition of denaturing agents and/or at elevated temperature and that this process is reversible. Another way to induce the free rotation of the minicircles would be to use a photolabile linker for the covalent attachment of the polyamide to the minicircles. This linker could be cleaved by irradiation with soft UV light after the ring closing step, leading to freely rotatable minicircles under all conditions. However, we chose not to use this approach for this study to keep the linker length as short as possible and hence the preorganization in a very defined way. Interestingly, the minicircles themselves are [2]-catenanes from two ss minicircles that are interlocked 16 times (Lk = 16). Therefore, the resulting catenane is actually a [2]-catenane of [2]-catenanes. The catenane can be synthesized as two different topoisomers; in Figure 2, the orange minicircle passes before the blue minicircle. Moving both the attachment site for the PA and the PA binding site 5 bases (half a helical turn) would result in a complex where in the lower junction the blue strand is 1741
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Nano Letters on top of the orange strand. However, these two isomers would become practically identical under conditions where the minicircles can rotate freely. In summary, we have shown a method of building catenanes from small double-stranded DNA minicircles. A C-shaped minicircle fragment was equipped with sequence-specific DNA-binding polyamides and the respective binding site. These fragments selfassembled into homodimeric complexes with interlocked C-shapes, in which the ends point to opposite directions. The addition of a ring-closing oligonucleotide produced catenanes as the main product. The structures were imaged by high-resolution AFM and the interlocked topology was proven through the structural integrity of the catenane under denaturing conditions. The catenanes presented here are at least 1 order of magnitude smaller than naturally occurring dsDNA catenanes. The size of the minicircles was chosen to be in the vicinity of the persistence length of DNA and the rings are thus rather rigid and show no supercoiling. Resembling stylized wedding rings, these catenanes are not only aesthetically appealing but could also be suitable components for the construction of molecular motors or machines made from DNA. The design principle of this study adds another recognition layer on top of the WatsonCrick design rules, that is, without the need of ssDNA templates. It can still be applied once all WatsonCrick base pairs have been formed or where the use of single-stranded regions is not possible (e.g., DNA plasmids or other dsDNA structures).
’ ASSOCIATED CONTENT
bS
Supporting Information. DNA sequences, the synthesis and workup protocol for the DNA-PA hybrid molecule, analytical data, synthesis and hybridization protocols of the catenanes, anion exchange HPLC data and AFM protocols, as well as the result of control experiments on the reversibility of PA-DNA binding. This material is available free of charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION Corresponding Author
*E-mail:
[email protected]. Present Addresses †
Wyss Institute for Biologically Inspired Engineering, Harvard University, 3 Blackfan Circle, 02115 Boston, MA.
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(2) Sauvage, J.; Dietrich-Buchecker, C. Molecular catenanes, rotaxanes and knots: a journey through the world of molecular topology; WileyVCH: New York, 1999. (3) Green, J. E.; Wook Choi, J.; Boukai, A.; Bunimovich, Y.; Johnston-Halperin, E.; DeIonno, E.; Luo, Y.; Sheriff, B. A.; Xu, K.; Shik Shin, Y.; Tseng, H.; Stoddart, J. F.; Heath, J. R. Nature 2007, 445, 414–417. (4) Leigh, D. A.; Wong, J. K. Y.; Dehez, F.; Zerbetto, F. Nature 2003, 424, 174–179. (5) Seeman, N. C. Annu. Rev. Biochem. 2010, 79, 65–87. (6) Omabegho, T.; Sha, R.; Seeman, N. C. Science 2009, 324, 67–71. (7) Willner, I.; Shlyahovsky, B.; Zayats, M.; Willner, B. Chem. Soc. Rev. 2008, 37, 1153–1165. (8) Liedl, T.; Sobey, T. L.; Simmel, F. C. Nano Today 2007, 2, 36–41. (9) Bath, J.; Turberfield, A. J. Nat. Nanotechnol. 2007, 2, 275–284. (10) Mao, C.; Sun, W.; Seeman, N. C. Nature 1997, 386, 137–138. (11) Seemand, N. C.; Chen, J.; Du, S.; Mueller, J.; Zhang, Y.; Fu, T.; Wang, Y.; Wang, H.; Zhang, S. New J. Chem. 1993, 17, 739–755. (12) Wang, H.; Du, S. M.; Seeman, N. C. J. Biomol. Struct. Dyn. 1993, 10, 853–863. (13) Chen, J.; Seeman, N. C. Nature 1991, 350, 631–633. (14) Zhang, Y.; Seeman, N. C. J. Am. Chem. Soc. 1994, 116, 1661–1669. (15) Weizmann, Y.; Braunschweig, A. B.; Wilner, O. I.; Cheglakov, Z.; Willner, I. Proc. Nat. Acad. Sci. U.S.A. 2008, 105, 5289. (16) Wilner, O. I.; Weizmann, Y.; Gill, R.; Lioubashevski, O.; Freeman, R.; Willner, I. Nat. Nanotechnol. 2009, 4, 249–254. (17) Ackermann, D.; Schmidt, T. L.; Hannam, J. S.; Purohit, C. S.; Heckel, A.; Famulok, M. Nat. Nanotechnol. 2010, 5, 436–442. (18) Han, D.; Pal, S.; Liu, Y.; Yan, H. Nat. Nanotechnol. 2010, 5, 712–717. (19) Shapiro, T. A.; Englund, P. T. Annu. Rev. Microbiol. 1995, 49 117–143. (20) Gellert, M. Annu. Rev. Biochem. 1981, 50, 879–910. (21) Krasnow, M. A.; Stasiak, A.; Spengler, S. J.; Dean, F.; Koller, T.; Cozzarelli, N. R. Nature 1983, 304, 559–560. (22) Tse, Y.; Wang, J. C. Cell 1980, 22, 269–276. (23) Spengler, S. J.; Stasiak, A.; Cozzarelli, N. R. Cell 1985, 42, 325–334. (24) Ackermann, D.; Rasched, G.; Verma, S.; Schmidt, T. L.; Heckel, A.; Famulok, M. Chem. Commun. 2010, 46, 4154–4156. (25) Gonc-alves, D. P. N.; Schmidt, T. L.; Koeppel, M. B.; Heckel, A. Small 2010, 6, 1347–1352. (26) Rasched, G.; Ackermann, D.; Schmidt, T. L.; Broekmann, P.; Heckel, A.; Famulok, M. Angew. Chem., Int. Ed. 2008, 47, 967–970. (27) Schmidt, T. L.; Nandi, C. K.; Rasched, G.; Parui, P. P.; Brutschy, B.; Famulok, M.; Heckel, A. Angew. Chem., Int. Ed. 2007, 46, 4382–4384. (28) Schmidt, T. L.; Heckel, A. Small 2009, 5, 1517–1520. (29) Dervan, P. Curr. Opin. Struct. Biol. 2003, 13, 284–299. (30) Cohen, J. D.; Sadowski, J. P.; Dervan, P. B. J. Am. Chem. Soc. 2008, 130, 402–403. (31) Nandi, C.; Parui, P.; Schmidt, T. L.; Heckel, A.; Brutschy, B. Anal. Bioanal. Chem. 2008, 390, 1595–1603.
’ ACKNOWLEDGMENT T.L.S. dedicates this work to his wife and colleague Dr. Diana P. Gonc-alves Schmidt on the occasion of their wedding. We would like to thank Professor A. Terfort for access to the AFM. Dr. Steve D. Perrault and Dr. Diana P. Gonc- alves Schmidt are gratefully acknowledged for helpful discussions and help in the preparation of the manuscript. This work was funded by grants to A.H. from the Deutsche Forschungsgemeinschaft (Cluster of Excellence Macromolecular Complexes EXC 115 and HE 4597/3-1). ’ REFERENCES (1) Schalley, C. A.; Beizai, K.; V€ogtle, F. Acc. Chem. Res. 2001, 34, 465–476. 1742
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