Temporal and reversible control of a DNAzyme by orthogonal

Subscriber access provided by University of South Dakota

Communication

Temporal and reversible control of a DNAzyme by orthogonal photoswitching Michael W Haydell, Mathias Centola, Volker Adam, Julián Valero, and Michael Famulok J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.8b08738 • Publication Date (Web): 16 Nov 2018 Downloaded from http://pubs.acs.org on November 16, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 6 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of the American Chemical Society

Temporal and reversible control of a DNAzyme by orthogonal photoswitching Michael W. Haydell1, Mathias Centola1, Volker Adam1, Julián Valero1,2*, and Michael Famulok1,2* 1LIMES

Chemical Biology Unit, Universität Bonn, Gerhard-Domagk-Straße 1, 53121 Bonn, Germany, 2Center of Advanced European Studies and Research, Ludwig-Erhard-Allee 2, 53175 Bonn, Germany. Supporting Information Placeholder ABSTRACT: The reversible switching of catalytic

systems capable of performing complex DNA Mathias was here computing operations using the temporal control of two orthogonal photoswitches is described. Two distinct photoresponsive molecules have been separately incorporated into a split horseradish peroxidase mimicking DNAzyme. We show that its catalytic function can be turned on and off reversibly upon irradiation with specific wavelengths of light. The system responded orthogonally not only to a sophisticated selection of irradiation wavelengths but depended also on different durations of irradiation. Furthermore, the DNAzyme exhibits reversible switching and retains this ability throughout multiple switching cycles. We apply the DNAzyme to act as a light-controlled 4:2 multiplexer. Orthogonally photoswitchable DNAzyme-based catalysts as introduced here have potential use for controlling complex logical operations and for future applications in DNA nanodevices.

The rapidly developing field of DNA nanotechnology seeks to build devices that can perform nanoscale operations. DNA origami1 has undergone remarkable progress in the past decade and was used for constructing 2D and 3D structural elements2 or complex dynamic machines3 and logic gates4 that can be triggered by various inputs5. DNA also was used to build mechanically interlocked structures such as catenanes6, rotaxanes7, and daisy-chains8 as precursors for other structures such as molecular motors9 or logic gates for DNA computing6b, 7e, 10. More complex applications of DNA nanomachines require establishing or refining simple and robust methods for controlling inputs and reading outputs. Previous DNA nanodevices mostly relied on strand displacement11 requiring oligodeoxynucleotides (ODNs) as inputs that can pollute the system after several switching cycles, thereby reducing efficiency

and robustness. Photoresponsive molecules incorporated into ODNs have gained increasing attention as optical control devices. Such systems use light as an external stimulus that eliminates the pollution due to addition of external ODNs. Asanuma and coworkers showed that different azobenzene derivatives12 can independently control DNA hybridization; others used light for irreversible uncaging13 or orthogonal molecular switching14.

Figure 1. A photoresponsive DNAzyme. (A) Chemical structures of DM-Azo and AAP. (B) Switching cycle for DM-Azo and AAP HRP-DNAzymes.

ACS Paragon Plus Environment

Journal of the American Chemical Society 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Here we report the combination of two distinct photoswitches that can reversibly and orthogonally control a DNAzyme by applying four different wavelengths. 2’,6’-Dimethylazobenzene (DM-Azo) and N-methyl-arylazopyrazole (AAP) (Figure 1A) effectively control duplex hybridization when incorporated into a DNA sequence15,16. Intercalated DM-Azo and AAP in trans conformation allow DNA hybridization while exposure to UV light sterically hinders hybridization by conversion into cis isomers. DM-Azo and AAP both switch back to trans when irradiated with 450 nm light, but orthogonality is demonstrated because only AAP switches to trans after 590 nm irradiation16. Time-dependent orthogonality is achieved by switching AAP to cis after 2 minutes exposure to 350 nm light, while efficient DM-Azo switching requires >6 minutes of irradiation. All irradiations were performed at 50 ºC.

Figure 2. Photoswitching of CDM-Azo- and CAAP at 1 µM evaluated by UV/Vis spectrometry. (A) Absorbance spectra for DM-Azo and AAP in cis and trans conformations. Values at absorbance maxima for DMAzo at different irradiation parameters (B) and with different irradiation sequences for DM-Azo (C) and AAP (D). Error bars: S.D. (n=3).

Recently an RNA scissor DNAzyme17 was reported to use visible light, via two azo derivatives, to silence genes. Here, we use photoresponsive molecules for guiding the supramolecular assembly of Gquadruplex18 (G4) horseradish peroxidase (HRP) mimicking DNAzymes, which can be applied in DNAbased functional devices, especially for biosensing4c, 19 and DNA computing4b. A split HRP-DNAzyme contains a G4 catalytic center consisting of two separated DNA strands20. It was recently employed as a reporter by using dehybridization to disassemble the G4 structure leading to drastically reduced catalytic activity7f. We use a 3:1 split HRPDNAzyme21 that forms when the two strands composing the G4 (A- and B-ODNs) hybridize to a

DM-Azo- or AAP-modified C-ODN (CDM-Azo, CAAP) when combined in solution with K+. With hemin and H2O2 the DNAzyme oxidizes ABTS [2,2’-azino-bis(3ethylbenzothiazoline-6-sulfonic acid)] to ABTS•+, monitored at 414 nm absorbance22. The photoresponsive DNAzyme can exist in four possible states depending on the irradiation wavelength and duration (Figure 1B). When the Aand B-ODNs are combined in equimolar amounts with the sum of the CDM-Azo and CAAP and subsequently irradiated at 365 nm or 350 nm for >6 minutes, the cis conformations of DM-Azo and AAP prevent hybridization of the A-ODN to CDM-Azo or CAAP. Thus, no G4 structure can form and the catalytic activity is effectively off (Figure 1B, state 1). Upon irradiation at 590 nm, AAP switches to trans and AODN can bind to CAAP (state 2). At this point, the DNAzyme activity is considered “partially on” compared to irradiation at 450 nm that switches both DM-Azo and AAP to trans. With both photoswitches in trans, A-ODN can hybridize to CDMAzo and CAAP (state 3) rendering the fully active DNAzyme. Irradiation at 350 nm for 2 minutes switches AAP to cis while DM-Azo remains in trans (state 4). CDM-Azo and CAAP switching efficiency was characterized by UV/Vis spectrometry. Based on the characteristic UV/Vis spectra for DM-Azo and AAP in their trans and cis conformations (Figure 2A and S1) the absorbance maxima for DM-Azo at 340 nm15 (Figure 2B, C) and for AAP at 350 nm16 (Figure 2D) were monitored after irradiation at different wavelengths. All irradiations were done at each LED’s maximum intensity (30 mA for 350 nm and 700 mA for all others) unless otherwise stated. Figure 2B shows the effect of irradiation at 350 nm on DM-Azo switching. Moieties were switched to trans by irradiating at 450 nm. A large portion of DMAzo remained in trans after 2 minutes of irradiation at 350 nm (Fig. 2B, red dot). Two subsequent 2minute irradiations of 350 nm (Fig. 2B, blue and green dots respectively) resulted in consistently decreasing levels of trans isomers, indicating that 350 nm light can isomerize DM-Azo with temporal control. To verify that the slower kinetics for DM-Azo switching at 350 nm is not due to the reduced LED intensity, the sample was also irradiated with 365 nm light at 30 mA (Fig. 2B, blue triangle). The reduced intensity irradiation at 365 nm successfully switched DM-Azo to cis, as indicated by similar absorbance after further irradiation with 365 nm light at 700 mA. Thus, the specific wavelength, and not the reduced intensity, of the 350 nm LED slows DM-Azo trans-cis switching kinetics. Subsequent 590


Page 2 of 6

Page 3 of 6 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


nm irradiation did not affect DM-Azo but 450 nm irradiation returned it to trans (Figure 2C). The absorbance values measured for AAP indicate that 2 minutes irradiation at 350 nm switches AAP to cis and irradiation with 590 nm light switches it to trans (Figure 2D, red dot and black triangle). At 365 nm and 30 mA intensity, complete switching to cisAAP was observed (blue dot), verified by further irradiation at 365 nm (700 mA). Irradiation with 450 nm light switched AAP to trans and another cycle of irradiations at 365 nm, 590 nm, and 450 nm shows that AAP remained in trans after both 590 nm and 450 nm irradiation steps. The above results demonstrate temporal control since 350 nm light can either switch both DM-Azo and AAP or selectively switch AAP only, depending on the duration of irradiation. This orthogonal behavior of the two Azoderivatives provides greater flexibility to future DNA nano-systems. The activity of the split DNAzyme is a function of hybridization and G-quadruplex formation. Fluorescence was employed to study the hybridization efficiency in isolation from G4 formation. The A- and B-ODNs were modified, substituting the G-triplets for a fluorophore (A-Hex) or quencher (B-BHQ2) respectively. A-Hex fluoresces when DM-Azo or AAP are in cis, preventing its hybridization to CDM-Azo or CAAP. Fluorescence is quenched when DM-Azo or AAP are in trans, allowing A-Hex to hybridize to CDM-Azo or CAAP (Figure S2). These experiments indicate that hybridization can be adequately and orthogonally photocontrolled for both photoswitches (Figure 3,S3). Hybridization efficiency of CDM-Azo is higher than CAAP (likely due to steric hindrance of AAP’s N-methyl group and imperfect π-π stacking) as A-Hex hybridizes to transCDM-Azo with 75% of the maximal normalized fluorescence level while it achieves only 47% when hybridizing to trans-CAAP. We used circular dichroism (CD) to determine the extent of G-quadruplex formation of the DNAzyme with and without K+ when DM-Azo and AAP switch from cis to trans. With K+ we observe a slight CD signal shift from the dsDNA peak at 280 nm (photoswitches in cis) towards the characteristic anti-parallel G-quadruplex peak at 270 nm (photoswitches in trans), indicating G-quadruplex formation (Figure S4A,B). Subtraction of the cis from the trans signal for both DM-Azo and AAP (Figure S4C) confirmed the increase in positive signal at 270 nm for each of the trans isomers. Having shown that A-ODN hybridizes to CAAP less efficiently than to CDM-Azo, we combined them in a 2:1 ratio to investigate the orthogonality of the system

based on catalytic activity of the DM-Azo and AAPmodified DNAzymes, over one cycle (Figure S5) or multiple switching cycles (Figure 4).

Figure 3. Hybridization efficiency of A-Hex to CDM-Azo or CAAP based on fluorescence signal after irradiation, normalized as per SI. Combined sample A- and B-ODNs are 600 nM while CDM-Azo is 200 nM and CAAP is 400 nM. ODNs in separate samples are at 300 nM. Error bars: S.D. (n=3).

The DNAzyme reaction rate was measured as absorbance per second (Abs/sec) at 414 nm to quantify the oxidation of ABTS to ABTS•+ over time. CDM-Azo in absence of CAAP only switched from cis to trans at 450 nm (Figure 4A, blue), and back to cis at 365 nm (purple). CAAP isomerized from cis to trans at 590 nm (Figure 4B, orange) and back to cis at 350 nm (green). Since AAP, but not DM-Azo, switched upon irradiation at either 590 or 350 nm, the results validate the orthogonal behavior of the described photoswitches. We finally combined the A- and BODNs in equimolar amounts with the mixture of CAAP and CDM-Azo in the 2:1 ratio and then irradiated over five cycles as described before (Figure 4C). Irradiation at 365 nm (Figure 4C, purple) switched both DM-Azo and AAP to cis, and neither DNAzyme was active. After irradiation at 590 nm (orange), only AAP switched to trans, and the combined DNAzymes exhibited intermediate activity. Irradiation at 450 nm (blue), switched both DM-Azo and AAP to trans, recovering full DNAzyme activity. Exposure to 350 nm (green) for 2 minutes switched only AAP to cis, while DM-Azo remained in trans, reducing the DNAzyme activity to an intermediate level. This switching behavior was maintained over 5 complete cycles, during which a decrease in maximum reaction rate (blue bars) was observed. However, the relative proportions of AAP to DM-Azo switching remained similar throughout the five cycles. However, the relative proportions of AAP to DM-Azo switching remained similar throughout the five cycles.



Figure 4. Reaction rates of DNAzymes with (A) CDM-Azo only, (B) CAAP only, and (C) CAAP and CDM-Azo 2:1 combination. DA: dark adapted positive control (on-state); AB: negative control (off-state lacking both CDM-Azo and CAAP). A- and BODNs are at 600 nM while CDM-Azo is 200 nM and CAAP is 400 nM. Rates are average of the linear phase slopes and normalized to the highest rate. Error bars: S.D. (n=3).

These results clearly indicate that DM-Azo and AAP can be orthogonally controlled to be in either cis or trans conformations.

Figure 5. DNAzyme 4:2 multiplexer. (A) Logic diagram with DM-Azo and AAP as select inputs and irradiation wavelength as data inputs. (B) Table describing the correlation between absorbance readouts and binary outputs. (C) Truth table for DNAzyme activity experiments.

In the current configuration, the DM-Azo- and AAPphotocontrolled DNAzyme behaves as a 4:2 multiplexer (Figure 5). Complex logic circuits such as multiplexers, demultiplexers23, encoders, and decoders24 are interesting objectives in DNA nanotechnology and molecular computing, since DNA biocompatibility offers potential for using DNA

computing in vivo25. Setting three different thresholds on the absorbance signal converts the output into two binary outputs (Q1 and Q2, Figure 5B): Absorbance below 20% outputs “0” for Q1 and Q2. In the 20%-40% absorbance range Q1 is “0” while Q2 is “1”. Readouts between 40% and 65% correlate with Q1 =”1” and Q2=“0”. Above 65% absorbance set both outputs to “1”. The truth table in Figure 5C describes the logic of the activity experiments (Figure 4C) with DM-Azo and AAP as select inputs and the four irradiation wavelengths as data inputs. In short, when both select inputs are “0,” inputting 590 nm irradiation returns “0” for the output Q1 and “1” for the output Q2. When the AAP select input is “1” and the DM-Azo select input is “0”, inputting 450 nm irradiation returns “1” for both outputs. When both select inputs are “1”, a data input of 350 nm irradiation returns “1” for Q1 and “0” for Q2. Finally, when the DM-Azo select input is “1” and the AAP is “0”, a data input of 365 nm irradiation returns “0” for both outputs (Figure 5C, Figure S6). Adding hemin and ABTS after irradiation precludes one-pot sequential operation, although system optimization may overcome such issues. Furthermore, different nano-architectures employing more complex arrangements of azobenzenes/pyrazoles, as well as better light sources such as lasers, may result in even more capable logic circuits such as full-adder or finite-state devices. In conclusion, we report an unprecedented light- and time-controlled orthogonal and reversible switching of catalytic DNAzymes for complex DNA computing operations. Compared to previously reported systems23-24, our study introduces novel and interesting possibilities for controlling complex DNA computing devices (see Figure S7). Advantages include using light as the sole data input as well as using DNAzyme activity (which provides signal amplification and good sensitivity) to directly monitor output. The system works orthogonally as


Page 4 of 6

Page 5 of 6 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


designed, is robust, and can be switched cleanly over multiple cycles because external addition of ODNs is not required. AAP is thermostable in the cis conformation for 100 hours at room temperature16 and DM-Azo for 25 hours at 60 ºC15, 26. Potentially, our strategy allows developing more complicated DNA architectures such as biosensors, logic gates, and DNA nanomachines. ASSOCIATED CONTENT Supporting Information. Supplementary Methods and Figures. This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Authors

Michael Famulok ([email protected]), Julián Valero ([email protected])

ACKNOWLEDGMENT We thank D. Keppner for technical assistance and the ERC (grant 267173) and Max-Planck Society for support.

REFERENCES 1. Rothemund, P. W. K., Nature 2006, 440, 297-302. 2. (a) Serpell, C. J.; Edwardson, T. G.; Chidchob, P.; Carneiro, K. M.; Sleiman, H. F., J. Am. Chem. Soc. 2014, 136, 15767-74; (b) Douglas, S. M.; Dietz, H.; Liedl, T.; Hogberg, B.; Graf, F.; Shih, W. M., Nature 2009, 459, 414-8. 3. (a) Funke, J. J.; Dietz, H., Nat. Nanotechnol. 2016, 11, 4752; (b) Ketterer, P.; Willner, E. M.; Dietz, H., Science Adv. 2016, 2, e1501209; (c) Kopperger, E.; List, J.; Madhira, S.; Rothfischer, F.; Lamb, D. C.; Simmel, F. C., Science 2018, 359, 296-301; (d) Mariottini, D.; Idili, A.; Vallee-Belisle, A.; Plaxco, K. W.; Ricci, F., Nano Lett. 2017, 17, 3225-3230. 4. (a) Zhang, C.; Yang, J.; Jiang, S.; Liu, Y.; Yan, H., Nano Lett. 2016, 16, 736-41; (b) Guo, Y.; Zhou, L.; Xu, L.; Zhou, X.; Hu, J.; Pei, R., Sci. Rep. 2014, 4, 7315; (c) Willner, I.; Shlyahovsky, B.; Zayats, M.; Willner, B., Chem. Soc. Rev. 2008, 37, 1153-65; (d) Zhu, J.; Zhang, L.; Dong, S.; Wang, E., ACS Nano 2013, 7, 10211-7. 5. (a) Amodio, A.; Del Grosso, E.; Troina, A.; Placidi, E.; Ricci, F., Nano Lett. 2018, 18, 2918-2923; (b) Amodio, A.; Zhao, B.; Porchetta, A.; Idili, A.; Castronovo, M.; Fan, C.; Ricci, F., J. Am. Chem. Soc. 2014, 136, 16469-72; (c) Hollenstein, M.; Hipolito, C.; Lam, C.; Dietrich, D.; Perrin, D. M., Angew. Chem. Int. Ed. 2008, 47, 434650; (d) Thomas, J. M.; Yu, H. Z.; Sen, D., J. Am. Chem. Soc. 2012, 134, 13738-48; (e) Li, W.; Yang, Y.; Yan, H.; Liu, Y., Nano Lett. 2013, 13, 2980-8. 6. (a) Lohmann, F.; Valero, J.; Famulok, M., Chem. Commun. 2014, 50, 6091-3; (b) Li, T.; Lohmann, F.; Famulok, M., Nat.

Commun. 2014, 5, 4940; (c) Lu, C. H.; Cecconello, A.; Qi, X. J.; Wu, N.; Jester, S. S.; Famulok, M.; Matthies, M.; Schmidt, T. L.; Willner, I., Nano Lett. 2015, 15, 7133-7; (d) Sannohe, Y.; Sugiyama, H., Bioorg. Med. Chem. 2012, 20, 2030-4; (e) Schmidt, T. L.; Heckel, A., Nano Lett. 2011, 11, 1739-42. 7. (a) Ackermann, D.; Jester, S. S.; Famulok, M., Angew. Chem. Int. Ed. 2012, 51, 6771-5; (b) Ackermann, D.; Schmidt, T. L.; Hannam, J. S.; Purohit, C. S.; Heckel, A.; Famulok, M., Nat. Nanotechnol. 2010, 5, 436-442; (c) List, J.; Falgenhauer, E.; Kopperger, E.; Pardatscher, G.; Simmel, F. C., Nat. Commun. 2016, 7, 12414; (d) Lohmann, F.; Ackermann, D.; Famulok, M., J. Am. Chem. Soc. 2012, 134, 11884-7; (e) Lohmann, F.; Weigandt, J.; Valero, J.; Famulok, M., Angew. Chem. Int. Ed. 2014, 53, 1037210376; (f) Centola, M.; Valero, J.; Famulok, M., J. Am. Chem. Soc. 2017, 139, 16044-16047. 8. Weigandt, J.; Chung, C.-L.; Jester, S.-S.; Famulok, M., Angew. Chem. Int. Ed 2016, 55, 5512-5516. 9. (a) Valero, J.; Pal, N.; Dhakal, S.; Walter, N. G.; Famulok, M., Nat. Nanotechnol. 2018, 13, 496-503; (b) Lu, C. H.; Cecconello, A.; Elbaz, J.; Credi, A.; Willner, I., Nano Lett. 2013, 13, 2303-8. 10. Lu, C. H.; Cecconello, A.; Willner, I., J. Am. Chem. Soc. 2016, 138, 5172-85. 11. Yurke, B.; Turberfield, A. J.; Mills, A. P.; Simmel, F. C.; Neumann, J. L., Nature 2000, 406, 605-608. 12. (a) Wang, S.; Yue, L.; Li, Z. Y.; Zhang, J.; Tian, H.; Willner, I., Angew. Chem. Int. Ed. 2018, 57, 8105-8109; (b) Zhou, M.; Liang, X.; Mochizuki, T.; Asanuma, H., Angew. Chem. Int. Ed. 2010, 49, 216770; (c) Nishioka, H.; Liang, X.; Kato, T.; Asanuma, H., Angew. Chem. Int. Ed. 2012, 51, 1165-8. 13. Rodrigues-Correia, A.; Weyel, X. M.; Heckel, A., Org. Lett. 2013, 15, 5500-3. 14. Lerch, M. M.; Hansen, M. J.; Velema, W. A.; Szymanski, W.; Feringa, B. L., Nat. Commun. 2016, 7, 12054. 15. Nishioka, H.; Liang, X.; Kashida, H.; Asanuma, H., Chem. Commun.s 2007, (42), 4354-6. 16. Adam, V.; Prusty, D. K.; Centola, M.; Skugor, M.; Hannam, J. S.; Valero, J.; Klockner, B.; Famulok, M., Chemistry 2018, 24, 10621066. 17. Kamiya, Y.; Arimura, Y.; Ooi, H.; Kato, K.; Liang, X. G.; Asanuma, H., Chembiochem 2018, 19, 1305-1311. 18. Travascio, P.; Li, Y.; Sen, D., Chem. Biol. 1998, 5, 505-17. 19. Pelossof, G.; Tel-Vered, R.; Elbaz, J.; Willner, I., Anal. Chem. 2010, 82, 4396-402. 20. Xiao, Y.; Pavlov, V.; Gill, R.; Bourenko, T.; Willner, I., Chembiochem 2004, 5, 374-9. 21. Deng, M.; Zhang, D.; Zhou, Y.; Zhou, X., J. Am. Chem. Soc. 2008, 130, 13095-102. 22. Collins, P. J.; Dobson, A. D. W.; Field, J. A., Appl. Environ. Microbiol. 1998, 64, 2026-31. 23. Orbach, R.; Remacle, F.; Levine, R. D.; Willner, I., Chem. Sci. 2014, 5, 1074-1081. 24. Kang, D.; White, R. J.; Xia, F.; Zuo, X. L.; Vallee-Belisle, A.; Plaxco, K. W., NPG Asia Mater. 2012, 4, e1. 25. Li, H.; Liu, Y.; Dong, S.; Wang, E., NPG Asia Mater. 2015, 7, e166. 26. Nishioka, H.; Liang, X.; Asanuma, H., Chemistry 2010, 16, 2054-62.




Page 6 of 6

Temporal and reversible control of a DNAzyme by orthogonal

Recommend Documents