I-motif-stapled and spacer-dependent multiple DNA nanostructures

Abstract

Construction of stimuli-responsive biomaterials as one of the most fascinating research fields plays an important role in various applications including bioanalysis and drug delivery. Functional nucleic acids (e.g. G-quadruplex, i-motif and DNAzyme) have been widely adopted to fabricate stimuli-responsive biomaterials, attributed to its remarkable and editable functionality. Herein, we employed i-motifs to staple DNA building blocks and obtained multiple pH-switchable DNA structures. Through increasing the length of T-spacers between duplex segments and i-motif sequences, DNA building blocks assembled into primary superstructures from multimers, dimers to monomers, which were characterized elaborately by various techniques, electrophoresis, atomic force microscopy and circular dichroism spectroscopy. It was postulated that the spacer effect on the morphology of final DNA assemblies was derived from conformational constraint among adjacent DNA architectures. Our study not only establishes one new DNA-based method for preparing stimuli-responsive biomaterials, but also provides a significant clue that the DNA linkers (spacers) between different nucleic acid structures are capable of influencing the conformation and morphology of the final products.

---

Introduction

Nucleic acids as an attractive biomaterial have been widely studied and introduced into nanotechnology.¹ Owing to their editable properties, high stability and fidelity, DNAs have being utilized as building blocks to fabricate nano/micro-structures which have been applied in various fields, such as bioanalysis,² electronics,³ drug delivery,⁴ logic operation⁵ and so on. Based on Watson–Crick base pairings, many approaches have been reported,^6–9 e.g. tile-based assembling¹⁰ and DNA origami.⁹ Consequently, one-dimensional (1D) structures (e.g. ribbon) and much more complicated (2D, 3D) constructs (e.g. triangle, cubes etc.) have been continuously acquired.^11–14

Functional DNA motifs (e.g. G-quadruplex, i-motif, DNAzyme and triplex) based on non-Watson–Crick interactions have been positioned into DNA structures and biomaterials,^15–18 on the basis of their remarkable functionalities.^19,20 For example, in our previous study,²¹ 1D DNA nanostructures were kinetically produced and functionalized with G-quadruplex DNAs which acted as signal carriers of fluorescence and colorimetry. In addition, various superstructures, such as G-wires,²² fray wires²³ and so on, were built up, and exhibited input (e.g. metal ion)-responsive behavior.²⁴ I-motif DNAs as one of the functional DNA structures, also have attracted wide interest of researchers,²⁵ due to its unique pH sensitivity derived from deprotonation of cytosine (C) and formation of C·CH⁺ hydrogen bonds.²⁶ By virtue of i-motifs, pH-responsive DNA nanomachines,²⁷ logic circuits²⁸ and hydrogels^29,30 were constructed, assembling and disassembling of nanoparticles were realized^31,32 and pH-stimulated drug release systems were designed for cancer therapy,^33–35 etc.³⁶ For example, Famulok et al. functionalized double-stranded DNA nanocircles with inter- and intramolecular i-motif DNAs and realized reversible assembling of defined DNA architectures which can undergo structural changes, such as dimerization of DNA nanocircles.³⁷ However, these studies have not proved the significance of DNA spacers in DNA-based biomaterials. The spacers between different DNA building blocks may affect the conformation and morphology of final structures. In addition, the further development of i-motif-based nanostructures is still required.

In this study, by stapling duplex units with dimeric i-motif structures at acidic environment, multiple pH-switchable DNA nanostructures were obtained, which combined structural advantages of both duplexes and i-motifs (Fig. 1A). Fig. 1B displays sequences of the building block consisting of C1Sn and C2Sn. We found that the base number (n) of spacers between duplex units and i-motif staples correlated with morphology of final assembled nanostructures. Obvious conformational constraint was engendered if there were no spacers between duplex units and i-motif structures, leading to formation of assembled multimers. As the length of spacer increased, distinct and small assemblies (dimer and monomer) were produced. Although some i-motif-based DNA nanostructures have been reported, the researchers did not pay attention to the significance of spacers between different building blocks, which was investigated, herein, for the first time.

Fig. 1 (A) Building blocks cosslinked by intermolecular i-motif DNAs as staples. Primary assemblies were produced from multimer, dimer to monomer by gradually increasing the length (n: T base number) of spacers between duplex units and i-motif structures. (B) The sequences of the building block consisting of two fuel strands, C1Sn and C2Sn.

Results and discussion

As mentioned above, i-motifs have been extensively applied in the DNA nanotechnology.²⁵ It is well-known that C-rich DNA sequences can fold into intramolecular or intermolecular i-motif structures.²⁶ Some DNA sequences containing two C-tracts yield stable bimolecular i-motifs, such as C5TC5, C6TC6 and C6T3C6.³⁷ In our work, C6TC6 (CCCCCCTCCCCCC) which can fold into a bimolecular i-motif with high stability, was selected and linked with one random duplex unit via a thymine (T)-spacer (in blue) (Fig. 1). When the solutions were adjusted to be acidic (pH = 5), the building blocks of different spacer length (T base number, n) were crosslinked by i-motif structures and self-assemble into distinct superstructures, multimer, dimer or monomer. We demonstrated that long DNA chains were obtained if there were no spacers (n = 0) between duplexes and i-motif DNAs. The primary assembled products became dimers and monomers gradually, as T number increased to 2–5.

Native PAGE was performed to confirm the pH-responsive assembling and disassembling of DNA nanostructures. The mixture of C1S0 and C2S0 in the weak basic solution (pH = 8.0) was annealed firstly, then the pH of the building block solution was adjusted to 5.0 to trigger the self-assembling. As shown in Fig. 2A, many bands corresponding to multimers were distinguished obviously (lane 3). DNA polymer chains were obtained which were comparable to duplexes with 500 base pairings (bp). After subjecting the system pH to 8.0, single band corresponding to single building block was observed (lane 6), indicating the dissociation of the DNA mulitmers at the weak basic environment. Once the solution pH was changed to 5.0 again, shifted bands attributed to multimers recovered (lane 9). These data illustrated that cyclic transition between single building blocks and assembled multimers was realized by changing the system pH between 8.0 and 5.0. This pH-controlled reversible assembling and dissociation of DNA multimers are promising for fabrication of reusable nanowire in the future.

Fig. 2 (A) Native PAGE analysis of switchable production of multimers at acidic environment. Lane 1, 4 and 7 represent C1S0 only. Lane 2, 5 and 8 represent C2S0 only. Lane 3, 6 and 9 represent products from mixture of C1S0 and C2S0. Lane 10 represents 50 bp DNA ladder. (B) AFM images of produced multimers from C1S0 and C2S0 mixture. The scale bar is 200 nm. (C) Frequency distribution of height of multimer products.

The assembled multimer products of C1S0/C2S0 system were characterized by atomic force microscopy (AFM). As revealed in Fig. 2B and S1,† one-dimensional DNA polymers with different length were observed clearly. The height of the products was statistically analyzed. The result in Fig. 2C exhibits that the average height is 2.1 nm, which represents single i-motif-duplex hybrid structure.³⁸ Also length histogram (Fig. S2†) shows the average length of DNA polymer chains is 78 nm and these multimer structures are consistent with those in gel electrophoresis assays which exhibit bands around 250 bp (Fig. 2A) based on the nm-to-bp value of 0.334.³⁹ And a few chains with length of around 167 nm correspond to the smearing bands near 500 bp. Circular dichroism spectra (Fig. 3) were also collected to validate the presence of i-motif structures which glued building blocks together. In contrast to the spectrum of C1S0/C2S0 system at pH 8.0, after adjusting the system pH to 5.0, an obviously enhanced positive signal near 290 nm was obtained, as a result of i-motif DNA structure generation. Therefore, the DNA multimers were prepared by i-motif structures as staples. In addition, it should be noted that the appropriate experimental procedures are critical for preparation of DNA multimers. C1S0 and C2S0 were annealed respectively at pH 5.0 and then mixed. Finally, only single band attributed to dimer products was observed clearly (lane 3 in Fig. S3†). Therefore, the efficient way to yield the multimers (lane 3 in Fig. 2A) is to anneal the C1S0 and C2S0 together at pH 8.0 and then adjust its pH to 5.0.

Fig. 3 CD spectra of mixtures containing C1S0 and C2S0 at pH 5.0 (a) and pH 8.0 (b), for demonstrating of i-motif formation.

One-dimensional DNA polymer chains (multimers) were produced using fuel strands without T-spacers (n = 0). To systematically investigate whether the T-spacer could affect the modality of final products, DNA fuel strands (C1Sn/C2Sn) with T-spacers of different length were designed and native PAGE was implemented (Fig. 4). In the presence of one T-spacer, the bands corresponding to multimer DNAs were also generated but decreased drastically (lane 3) as compared to those without spacers (lane 3 in Fig. 2A). With increasing the number of T base to 2 and 3, the dimer products were obtained mostly (indicated by red circles in lane 6 and 9), and massive monomers became dominant if the number of T base reached 5 (indicated by a blue circle in lane 12). If the spacer length was up to 10 T bases, only single band emerged corresponding to monomer products (lane 6 in Fig. S3†), disclosing the self-assembling of an i-motif in single building block. Additionally, small DNA nanostructures with height of ca. 2.0 nm were observed, but no DNA chains were found in the AFM images of assembled products from C1S2/C2S2 (Fig. S4A and B†) and C1S10/C2S10 (Fig. S4C and D†), which were consistent with the PAGE results and suggested the spacer-dependent effect. The formation of i-motif structures at acidic environment were also confirmed for systems, C1S2/C2S2 and C1S5/C2S5, by CD (Fig. S5†). These results revealed that the assembled nanostructures of distinct modalities were obtained by tuning the length of T-spacers between i-motifs and duplex units, probably owing to generation and weakening of conformation constraint between adjacent duplex units which were crosslinked by i-motifs.

Fig. 4 Native PAGE confirming that formation of primary assemblies (lanes 3, 6, 9 and 12) from multimer/dimer to monomer, at pH 5.0, by prolonging the T-spacers between duplexes and i-motifs. The spacer length (T number, n) is corresponding to 1 (lanes 1–3), 2 (lanes 4–6), 3 (lanes 7–9) and 5 (lanes 10–12) T bases, respectively. Lane 13 represents 50 bp DNA ladder.

Conclusions

Stimuli-responsive materials, especially DNA-based biomaterials, have been widely prepared for molecular machines, logic operations, and drug delivery and so on. Because of their remarkable functionalities, functional DNA motifs have contributed to the field so much and are inevitable to be continuously utilized for much more complicated stimuli-responsive DNA architectures²⁸ and biomaterials.¹⁸ In this study, we utilized one significant functional nucleic acid, i-motif DNA structures as staples, and fabricated pH-responsive multiple DNA nanostructures. It was found that the DNA spacers between duplexes and i-motif structures were critical for morphology of final assembled products. For the first time, the spacer effect were systematically investigated by analyzing the assembled products via native PAGE. In the absence of spacers and ascribed to the presence of conformation constraint, the DNA building blocks were crosslinked by i-motifs, resulting in production of multimers. As the spacer was prolonged and conformational constraint was weakened, small assemblies (DNA dimer and monomer) were obtained. The multiple pH-stimulated DNA nanostructures may hopefully be applied into distinct-scale molecular devices.⁴⁰ Moreover, the finding that the DNA linkers (spacers) between different nucleic acid architectures influence the conformation and morphology of final products, is very instructive for constructing other DNA nanostructures in the future.

Experimental

Materials

Ultrapage-purified oligonucleotides (Table S1†) and tris(hydroxymethyl)aminomethane (Tris) were obtained from Sangon Biotechnology Co., Ltd (Shanghai, China). 3-Aminopropyltriethoxysilane (APTES) and ethidium bromide (EB) were purchased from Sigma-Aldrich (USA). Acrylamide, N,N′-methylenebisacrylamide and ammonium persulfate ((NH₄)₂SO₄) were obtained from Beijing Dingguochangsheng Biotechnology Co. Ltd (Beijing, China). Magnesium acetate (Mg(OAc)₂) and tetramethylethylenediamine (TEMED) were purchased from Beijing Chemical Works (Beijing, China) and AMRESCO Inc. (USA), respectively. 50 bp DNA ladder was purchased from New England Biolabs (Beijing), Ltd. The 6× loading buffer (pH = 8.0 or 5.0) was prepared containing 25 mM Tris, 10 mM Mg(OAc)₂ and 36% glycerol. TA buffer (pH = 8.0 or 5.0) containing 25 mM Tris and 10 mM Mg(OAc)₂ was used for sample preparation and electrophoresis. Double distilled water was used throughout.

Native polyacrylamide gel electrophoresis (native PAGE)

All the assembled products were prepared as follows: the mixtures of fuel strands C1Sn (2 μM) and C2Sn (2 μM) in TA buffer (pH = 8.0) were first heated for 3 min and cooled down slowly for 10 hours. Then the solution pHs were adjusted to 5.0 using 500 mM HCl. After incubation overnight, the final assembled products were obtained and subjected to electrophoresis. HCl (500 mM) and NaOH (500 mM) solutions were used to switch the mixture pHs between 8.0 and 5.0. The samples (C1Sn only, C2Sn only) were also prepared as controls. Polyacrylamide gels (6%, w/v) were prepared with TA buffer. 10 μL of each sample with 2 μL of 6× loading buffer (pH = 8.0 or 5.0) was loaded into the gel. The gels were run at 60 V for 90 min and photographed under UV light on a fluorescence imaging system (Vilber Lourmat, Marne laVallee, France).

Atomic force microscopy (AFM)

The assembled products from C1S0 and C2S0 were prepared according to procedures as mentioned above, and then diluted ten times with TA buffer (pH = 5.0). Freshly cleaved micas were immersed in 1% APTES solution for 5 min, then rinsed with double distilled water and dried in a desiccator. After 6 hours, 120 μL of the diluted samples were added onto the pretreated micas. 5 min later, the mica surfaces were rinsed gently and dried. The AFM images were collected on a MultiMode® 8 Scanning Probe Microscope (Veeco Instruments Inc., USA) using tapping mode at room temperature and analyzed with NanoScope® Version 8.1 software. The height data was acquired by analyzing the cross-sections of DNA chains and the length data of DNA chains were measured by drawing short lines along the DNA contour. These data were processed into the histograms of frequency distributions by frequency count in OriginPro 9.1.

Circular dichroism (CD)

The CD measurements from 210 to 350 nm were performed on a JASCO J-820 spectropolarimeter (Tokyo, Japan). The data pitch was 0.1 nm, response time was 0.5 s, scan speed was 200 nm min⁻¹, and bandwidth was 1 nm. 700 μL of each sample containing fuel strands (2 μM) was prepared according to aforementioned procedures.

Acknowledgements

This study was supported by the National Natural Science Foundation of China (No. 21190040, 91227114 and 11174105) and the National Science Foundation.

Notes and references

1. E. Stulz, G. Clever, M. Shionoya and C. Mao, Chem. Soc. Rev., 2011, 40, 5633.
2. C. Lin, Y. Liu and H. Yan, Nano Lett., 2007, 7, 507.
3. D. Liu, S. H. Park, J. H. Reif and T. H. LaBean, Proc. Natl. Acad. Sci. U. S. A., 2004, 101, 717.
4. B. P. Timko, T. Dvir and D. S. Kohane, Adv. Mater., 2010, 22, 4925.
5. Y. Benenson, Nat. Rev. Genet., 2012, 13, 455.
6. X. He, L. Dong, W. Wang, N. Lin and Y. Mi, Chem. Commun., 2013, 49, 2906.
7. Y. Ma, H. Zheng, C. Wang, Q. Yan, J. Chao, C. Fan and S.-J. Xiao, J. Am. Chem. Soc., 2013, 135, 2959.
8. D. Niu, H. Jiang, R. Sha, J. W. Canary and N. C. Seeman, Nucleic Acids Res., 2015, 43, 7201.
9. B. Saccà and C. M. Niemeyer, Angew. Chem., Int. Ed., 2011, 51, 58.
10. C. Lin, Y. Liu, S. Rinker and H. Yan, ChemPhysChem, 2006, 7, 1641.
11. C. S. Nadrian and S. L. Philip, Rep. Prog. Phys., 2005, 68, 237.
12. J. J. Lu, S. Wang, G. Li, W. Wang, Q. Pu and S. Liu, Anal. Chem., 2012, 84, 7001.
13. N. Y. Wong, H. Xing, L. H. Tan and Y. Lu, J. Am. Chem. Soc., 2013, 135, 2931.
14. F. A. Aldaye, A. L. Palmer and H. F. Sleiman, Science, 2008, 321, 1795.
15. D. P. N. Gonçalves, T. L. Schmidt, M. B. Koeppel and A. Heckel, Small, 2010, 6, 1347.
16. Y. Yang, C. Zhou, T. Zhang, E. Cheng, Z. Yang and D. Liu, Small, 2012, 8, 552.
17. F. Wang, J. Elbaz, R. Orbach, N. Magen and I. Willner, J. Am. Chem. Soc., 2011, 133, 17149.
18. J. Ren, Y. Hu, C.-H. Lu, W. Guo, M. A. Aleman-Garcia, F. Ricci and I. Willner, Chem. Sci., 2015, 6, 4190.
19. M. Wang, Z. Mao, T.-S. Kang, C.-Y. Wong, J.-L. Mergny, C.-H. Leung and D.-L. Ma, Chem. Sci., 2016, 7, 2516.
20. H. A. Day, P. Pavlou and Z. A. E. Waller, Bioorg. Med. Chem., 2014, 22, 4407.
21. J. Ren, J. Wang, L. Han, E. Wang and J. Wang, Chem. Commun., 2011, 47, 10563.
22. T. C. Marsh and E. Henderson, Biochemistry, 1994, 33, 10718.
23. E. Protozanova and R. B. Macgregor, Biochemistry, 1996, 35, 16638.
24. R. P. Fahlman, M. Hsing, C. S. Sporer-Tuhten and D. Sen, Nano Lett., 2003, 3, 1073.
25. Y. Dong, Z. Yang and D. Liu, Acc. Chem. Res., 2014, 47, 1853.
26. M. Guéron and J.-L. Leroy, Curr. Opin. Struct. Biol., 2000, 10, 326.
27. D. Liu and S. Balasubramanian, Angew. Chem., Int. Ed., 2003, 42, 5734.
28. T. Li, F. Lohmann and M. Famulok, Nat. Commun., 2014, 5, 4940.
29. E. Cheng, Y. Xing, P. Chen, Y. Yang, Y. Sun, D. Zhou, L. Xu, Q. Fan and D. Liu, Angew. Chem., Int. Ed., 2009, 48, 7660.
30. W. Guo, C.-H. Lu, X.-J. Qi, R. Orbach, M. Fadeev, H.-H. Yang and I. Willner, Angew. Chem., Int. Ed., 2014, 53, 10134.
31. W. Wang, H. Liu, D. Liu, Y. Xu, Y. Yang and D. Zhou, Langmuir, 2007, 23, 11956.
32. C. Wang, Z. Huang, Y. Lin, J. Ren and X. Qu, Adv. Mater., 2010, 22, 2792.
33. D. He, X. He, K. Wang, J. Cao and Y. Zhao, Adv. Funct. Mater., 2012, 22, 4704.
34. L. Song, V. H. B. Ho, C. Chen, Z. Yang, D. Liu, R. Chen and D. Zhou, Adv. Healthcare Mater., 2013, 2, 275.
35. W. Zhang, F. Wang, Y. Wang, J. Wang, Y. Yu, S. Guo, R. Chen and D. Zhou, J. Controlled Release, 2016, 232, 9.
36. S. Modi, M. G. Swetha, D. Goswami, G. D. Gupta, S. Mayor and Y. Krishnan, Nat. Nanotechnol., 2009, 4, 325.
37. T. Li and M. Famulok, J. Am. Chem. Soc., 2013, 135, 1593.
38. Y. Mao, D. Liu, S. Wang, S. Luo, W. Wang, Y. Yang, Q. Ouyang and L. Jiang, Nucleic Acids Res., 2007, 35, e33.
39. K. Hori, T. Takahashi and T. Okada, Eur. Biophys. J., 1998, 27, 63.
40. K. Keren, M. Krueger, R. Gilad, G. Ben-Yoseph, U. Sivan and E. Braun, Science, 2002, 297, 72.

---

Footnotes

† Electronic supplementary information (ESI) available: Detailed materials, Table S1 and Fig. S1–S3. See DOI: 10.1039/c6ra15201e

‡ These authors contributed equally.

---