De novo design of functional oligonucleotides with acyclic scaffolds.

Personal Account

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De Novo Design of Functional Oligonucleotides with Acyclic Scaffolds Hiroyuki Asanuma,*[a] Hiromu Kashida,[a] and Yukiko Kamiya[a,b] Department of Molecular Design and Engineering, Graduate School of Engineering, Nagoya University, Furo-cho, Chikusa-ku, Nagoya 464-8603 (Japan), Tel.: (+81) 52-789-2488, Fax: (+81) 52-789-2528, E-mail: [email protected] [b] EcoTopia Science Institute, Nagoya University, Furo-cho, Chikusa-ku, Nagoya 464-8603 (Japan)

[a]

Received: April 29, 2014 Published online: August 29, 2014

ABSTRACT: In this account, we demonstrate a new methodology for the de novo design of functional oligonucleotides with the acyclic scaffolds threoninol and serinol. Four functional motifs—wedge, interstrand-wedge, dimer, and cluster—have been prepared from natural DNA or RNA and functional base surrogates prepared from d-threoninol. The following applications of these motifs are described: (1) photoregulation of formation and dissociation of a DNA duplex modified with azobenzene, (2) sequence-specific detection of DNA using a fluorescent probe, (3) formation of fluorophore assemblies that mimic quantum dots, (4) improved strand selectivity of siRNA modified with a base surrogate, and (5) in vivo tracing of the RNAi pathway. Finally, we introduce artificial nucleic acids (XNAs) prepared from d-threoninol and serinol functionalized with each of the four nucleobases, which have unique properties compared with other acyclic XNAs. Functional oligonucleotides designed from acyclic scaffolds will be powerful tools for both DNA nanotechnology and biotechnology. DOI 10.1002/tcr.201402040 Keywords: fluorescent probes, helical structures, nucleic acids, oligonucleotides, photochemistry

Introduction Naturally occurring DNA is a programmable supramolecule as well as a carrier of genetic information. Beginning in the early 1980s,[1] Seeman’s group developed DNA nanoarchitectures by assembling pieces of synthetic DNA as building blocks. This led to the DNA origami technology, originally developed by Rothemund,[2] and DNA is now regarded as an excellent and easily accessible nanomaterial that can be used to construct various nanomachines and higher-ordered nanoarchitectures as well as biological tools. DNA nanoarchitectures and tools are limited when only the four passive natural nucleotides are used. Chemical modification of DNA or RNA can overcome this

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limitation. Conventionally, modifications are made to nucleotides that are incorporated into oligonucleotides via phosphoramidite chemistry or postsynthetic modification.[3] The 2′-position of the nucleotides and 5-position of uracil are suitable for incorporation of functional moieties because these modifications should not induce significant changes to the nucleic acid duplex. Incorporation of multiple functional residues is complicated using these modification sites, however, as four modified phosphoramidite monomers are necessary to incorporate functional molecules at any position of the sequence. In addition, modification is rather laborious due to

© 2014 The Chemical Society of Japan and Wiley-VCH, Weinheim

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the need to protect hydroxyl and amino groups. Furthermore, even when the structural difference between the native and modified nucleotides is small, these modified duplexes are not necessarily stable, especially when multiple functional molecules are incorporated.[4] To enable multiple incorporations of functional moieties without significant destabilization of the DNA duplex, we have designed scaffolds based on acyclic diols that are readily incorporated within an oligonucleotide. For the last decade, we have prepared base surrogates using d-threoninol (Figure 1a). These analogues extend the “genetic alphabet” and, unlike modification of the nucleotide unit, are simple to prepare. With these surrogates, various functional motifs can easily be designed by the conjugation with natural nucleotides. Furthermore, the surrogate prompted us to design new artificial nucleic acids, acyclic threoninol nucleic acid

Hiroyuki Asanuma is a Professor at the Graduate School of Engineering, Nagoya University. He received his Ph.D. (1989) from the University of Tokyo. In 1989, he joined Fuji Photo Film Co. Ltd. as a researcher, working on the spectral sensitization of silver halide emulsion and the development of lithium ion secondary batteries. He moved to the University of Tokyo in 1995 as Assistant Professor and began investigation of polymer receptors. He was promoted to Associate Professor in 2000 and started work on photoresponsive DNA. In 2005, he moved to Nagoya University as a Full Professor. Currently, his research interests are focused on the development of functional artificial oligonucleotides with acyclic scaffolds and new XNAs and their applications: (1) photoregulation of DNA and RNA functions, (2) fluorescent probes for practical applications, (3) nucleic acid based drugs, and (4) assembled functional molecules.

(aTNA) and serinol nucleic acid (SNA), which form unexpectedly stable homo-duplexes. In this account, we first explain the design of four functional motifs with d-threoninol and then focus on their applications. Unique stereochemical and hybridization properties of aTNA and SNA are also described.

d-Threoninol as a Scaffold of a Functional Molecule To Design Four Artificial Motifs Threoninol (2-amino-1,3-butanediol, see Figure 1a), obtained by the reduction of threonine methyl ester,[5] is a convenient acyclic scaffold that can tether the various functional molecules listed in Figure 1b to an amino group, and which is further converted to a phosphoramidite monomer.[6] d-Threoninol is

Yukiko Kamiya is an Associate Professor at the EcoTopia Science Institute at Nagoya University. In 2008 she received her Ph.D. from Nagoya City University. From 2008 to 2011 she worked at the Institute for Molecular Science at the National Institutes of Natural Sciences as a postdoctoral researcher. In 2012, she joined Professor Hiroyuki Asanuma’s group at the Graduate School of Engineering at Nagoya University as an Assistant Professor. She is currently an Associate Professor in the EcoTopia Science Institute. Her current interests are in the development of molecular tools for application in biological systems, such as gene regulation including RNAi.

Hiromu Kashida is an Associate Professor at the Graduate School of Engineering, Nagoya University. He received his Ph.D. in 2006 from the University of Tokyo. He became an Assistant Professor at Nagoya University in 2007 and was promoted to Associate Professor in 2011. His research interests include the design of artificial nucleic acids and their applications as probes and nanomaterials. He received the PCCP Prize from the Royal Society of Chemistry in 2011 and the Chemical Society of Japan Award for Young Chemists in 2014.

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De Novo Design of Functional Oligonucleotides

Fig. 1. Design of functional motifs by the combination of base surrogates synthesized via d-threoninol with natural nucleotides. (a) Synthesis of a base surrogate from threonine methyl ester. (b) Typical functional molecules tethered on threoninol. (c) Schematic of duplex containing natural nucleotides and a base surrogate.

superior to the l form as a scaffold due to its clockwise winding property.[7–9] A duplex that contains this base surrogate is shown in Figure 1c; the functional moiety on d-threoninol is additionally inserted to form a single-bulge-like structure. NMR analysis revealed that the functional group on the scaffold intercalates between the adjacent base pairs.[7] This sequence design allows multiple introductions of functional molecules at any position in the duplex without sacrificing base pairing. We have characterized four functional motifs with this base surrogate. Wedge motif (Figure 2a). In the wedge motif, functional groups linked to the d-threoninol scaffold (X) are multiply inserted into one strand, whereas the complementary strand is composed of only native nucleotides (N). To avoid significant destabilization of the duplex due to asymmetrical sequence design, at least two natural nucleotides should be inserted between the surrogates as in 5′-(NNX)n-NN-3′.[9,10] A planar functional molecule with a molecular size of around 1.1 nm, similar to that of a Watson–Crick base pair,[11] is desirable to stabilize the wedge motif.[9] Interstrand-wedge (I-W) motif (Figure 2b). In the I-W motif, base surrogates are introduced into both strands of the duplex as in 5′-(NNX)n-NN-3′ and 3′-(NXN)n+1-5′, resulting in alternating natural base pairs and functional molecules. Unlike the wedge motif, the I-W motif is symmetric and is remarkably stable due to the stacking interactions between natural bases and the intercalated functional molecules.[12,13]

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Fig. 2. Four functional motifs designed with functionalized d-threoninol incorporated on one or both strands of a duplex. Ovals represent base surrogates and interlocking shapes represent natural nucleotides.



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Fig. 3. (a) Schematic illustration of reversible photoregulation of hybridization induced by trans–cis isomerization of azobenzenes. (b) Modification of azobenzene results in more effective photoregulation.

NMR analysis proved that canonical Watson–Crick base pairs form, each separated by the surrogate in the I-W motif.[13] Dimer motif (Figure 2c). In this motif, base surrogates are regarded as pseudonucleobases; surrogates on opposite strands form a non-natural pair (dimer).[14] There are two possible arrangements of the dimer. NMR analysis revealed that surrogates are adjacent to the 5′ natural base pair in a DNA duplex but to the 3′ side in an RNA duplex (Figure 2c).[15,16] The surrogate dimers can be aligned alternately with natural base pairs by hybridizing 5′-(NX)n-N-3′ with 3′-(NX)n-N-5′. Since stacking interactions between the surrogates stabilize the duplex, the melting temperature (Tm) of a duplex containing this motif is much higher than that of the duplex without the surrogates.[14] Cluster motif (Figure 2d). The cluster motif can be regarded as an extension of the dimer motif in which surrogates are consecutively multiplied, allowing for zipper-like clusters of functional molecules by combining 5′-Nm-Xn-No-3′ and 3′-Nm-Xn-No-5′.[15–17] As with the dimer motif, the cluster motif stabilizes a duplex due to stacking interactions among the surrogates; the Tm increases almost linearly with the number of surrogates incorporated.[17] It should be noted that canonical Watson–Crick base pairs are formed in all of these motifs, so that introduction of a mismatched pair significantly lowers the Tm.[9,12–14] In dimer and cluster motifs, the base surrogates form stable pairs but surrogates do not pair with natural nucleobases on (deoxy)ribose scaffolds.[17] In this sense, the surrogates prepared from


d-threoninol expand the “genetic alphabet”. In the following sections, we demonstrate several applications attained with these functional motifs.

Reversible Photoregulation of the Formation and Dissociation of Azobenzene-Containing Duplexes Artificial control of biofunctions by external stimuli is one of the current hot topics in chemical biology.[18] Among external stimuli, light has several advantages: (1) unlike molecular stimuli, light does not contaminate the microenvironment of the reaction system; (2) spatiotemporal control of the reaction is possible; and (3) irradiation wavelengths are tunable by suitable molecular design.[19,20] To provide biomacromolecules with photoresponsiveness, the macromolecule can be tethered covalently to a photoswitchable molecule such as a caged compound.[21,22] For reversible regulation, azobenzene derivatives are widely utilized due to their chemical stability, availability, and large structural and polarity changes induced by UV (300 nm < λ < 400 nm) and visible light (λ > 400 nm).[23] Over the last decade, our group has tethered azobenzenes via d-threoninol for the photoregulation of DNA-based nanomachines.[24] Our strategy is based on the reversible formation and dissociation of the duplex induced by light irradiation, as schematically illustrated in Figure 3a. Since most DNA-based nanoarchitectures and bioreactions involve spontaneous hybridization, photocontrol of this supramolecular

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property allows various applications. Photoregulation of hybridization is based on the stabilization of the duplex by intercalated planar trans-azobenzene, whereas trans-to-cis isomerization by UV light irradiation destabilizes the duplex due to the steric hindrance of non-planar cis-azobenzene.[6] Modification of azobenzene at the ortho position of the distal benzene ring (Figure 3b) improves the photoregulatory efficiency.[25–27] For most DNA-based devices, duplexes of about 20 base pairs are used.[10] Hence, with wedge, interstrand-wedge, and dimer motifs, formation and dissociation of DNA duplexes modified with azobenzene could be photoregulated almost completely.[10,12–14] Photoresponsive DNA has recently been reviewed in detail in another account.[24]

Design of Functional Fluorescent Probes for Sequence-Specific Detection of DNA ISMB probe. Sequence-specific recognition of DNA and RNA is of increasing importance in disease diagnosis and in assessment of disease risks and drug responses of individuals.[28] One of the typical fluorescent probe designs is a molecular beacon (MB) originally developed by Tyagi et al.[29] In an MB, fluorophore and quencher dyes are attached at the termini of a stem-loop structure.[29,30] Recently, we proposed a new MB design that we call an in-stem molecular beacon (ISMB), which employs a dimer motif within the stem region to pair fluorophore and quencher surrogates.[31,32] ISMB enables multiple introductions of fluorophore–quencher pairs into the stem without significant destabilization. Accordingly, the signal-to-background (S/B) ratio is remarkably improved relative to an MB of the typical design.[31–33] Furthermore, we can easily design a functional ISMB that can discriminate between wild-type and mutant sequences such as single-nucleotide polymorphisms (SNPs) and insertion/deletion polymorphisms (indels). One to four base pair indels are more frequent than larger indels, and some are related to genetic diseases.[34] An ISMB probe design that can detect deletion mutants is schematically illustrated in Figure 4a.[31,35] Two fluorophores (F and L) are introduced on either side of the target sequence.[35] When the probe hybridizes with the fully matched wild-type sequence (Wild in Figure 4b), the two fluorophores are intercalated between base pairs to form a stable wedge motif, and the target base pairs interrupt the direct contact between F and L. Excitation of F affords monomeric fluorescence from L due to fluorescence resonance energy transfer (FRET) between F and L. However, hybridization with a three-base deletion mutant (Del) results in a bulge structure in which the two perylene derivatives are in close proximity. In this structure, monomer emission is quenched, and exciplex emission with a longer wavelength appears. In order to minimize background emission in the absence of the target, the ISMB (FL-MB) was designed with doubled quenchers opposite the fluorophore.[36]

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A demonstration of the discriminatory ability of this design is shown in Figure 4b. The target was the cystic fibrosis transmembrane conductance regulator (CFTR) gene; a specific three-base deletion is responsible for many cases of cystic fibrosis. In the absence of target, the FL-MB does not emit fluorescence at all (Figure 4b). In the presence of fully matched Wild, monomeric green fluorescence is observed. In contrast, orange-colored exciplex emission is observed in the presence of the RNA with the deletion. Notably, with this system, these polymorphisms can be distinguished with the naked eye. An ideal fluorescent probe satisfies the following prerequisites: (1) no emission in the absence of target, (2) bright emission from the fluorophore in the presence of target DNA or RNA with sufficient sequence specificity, and (3) rapid response to the substrate. An ISMB associated with a chaperone polymer satisfies these prerequisites.[33] Stemless linear probe. We have also designed stemless linear probes that meet the criteria for ideal probes.[37] By taking advantage of the wedge motif, we designed the probe depicted in Figure 5a.[38] Fluorophores on d-threoninols are multiply inserted into the probe. In the single-stranded state, the flexible probe does not emit fluorescence due to self-quenching among the weakly interacting fluorophores. Upon hybridization with the target, intercalation of each dye between base pairs results in strong fluorescence emission as the dyes are separated. In this design we use perylene as the fluorophore because perylenes (unlike pyrenes) efficiently self-quench. Self-quenching in the 5′-(NNX)n-3′ design is more efficient than in the 5′-(NNNX)n-3′ design. Too many fluorophores in the probe may retard hybridization with the target but too few results in low signal. To facilitate self-quenching, a butylene linker was introduced between the amide bond and perylene (E4 in Figure 5a). The single-stranded linear probe, 5E4-2, does not emit light due to the self-quenching of the five incorporated perylenes, whereas in the presence of the target b3a2 significant emission is observed (Figure 5b). The S/B ratio is as high as 180 with this design. Note that in this design no quencher is needed. Our linear probe was able to discriminate fully matched from mismatched target.[38]

Fluorophore Assemblies that Mimic Quantum Dots Quantum dots (QDs) are size-controlled inorganic nanoclusters that have been used as fluorescent tags for various biomacromolecules. QDs are especially useful in singlemolecule imaging due to their brightness, photostability, and large Stokes shift.[39–42] We have constructed organized fluorophore assemblies that mimic inorganic quantum dots using wedge, I-W, dimer, and cluster motifs. Here, we selected pyrene (P) and perylene (E) as excimeremitting and self-quenching fluorophores, respectively. First,



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Fig. 4. (a) Schematic illustration of the design of a functional ISMB that can discriminate between the wild-type sequence and a three-base deletion mutant. (b) Sequences of FL-MB and substrates (red characters in FL-MB denote the stem region, and blue ones in Wild are missing in Del). Fluorescence emission spectra of FL-MB alone (black), FL-MB + Wild (green), and FL + Del (orange) are shown. The inset shows magnified spectra. The photograph demonstrates the color differences. Excitation was at 445 nm. Solution conditions were 0.2 μM FL-MB, 0.4 μM target (Wild or Del), 100 mM NaCl, 10 mM phosphate buffer (pH 7.0), 20°C.

we designed the oligonucleotides shown schematically in Figure 6a (left panel) with six pyrenes.[43] In the I-W motif, each pyrene is shielded by the adjacent base pairs. Since pyrene is severely quenched by the nucleobases, monomeric fluorescence (380 and 400 nm emission) from the pyrenes in the I-W motif was insignificant (Figure 6b, red line). In contrast, excimer emission at around 480 nm was observed in motifs with clustered pyrenes. Maximum emission was observed with the cluster motif involving six pyrenes (Figure 6b, black line). When we used perylene instead of pyrene, emission properties were entirely reversed: in the I-W motif maximum monomeric emission was observed, whereas almost no emission was observed from the cluster motif due to the severe self-quenching.[43] Hetero-assembly of pyrene with perylene afforded different emission properties and depended on the motif. For


example, in the I-W motif, in which each pyrene and perylene are separated by intervening base pairs, we observed strong blue-colored emission from perylene due to the facilitated FRET between the pyrene and perylene (Figure 6c, red line).[43] Note that the excitation wavelength was at 345 nm, where pyrene has an absorption maximum. The apparent Stokes shift is 115 nm. Furthermore, this hetero-assembly could be excited with light ranging from 300 to 470 nm. Use of dimer and trimer motifs shifted the emission band to longer wavelengths due to exciplex formation (Figure 6c, compare blue and green lines with red). In the cluster motif, monomer emission completely disappeared leaving an exciplex emission at around 550 nm (Figure 6c, black line). Color variation depending on the motif was observable with the naked eye, as shown in the inset of Figure 6c. Thus, use of the various motifs allowed us to mimic most of the properties of inorganic QDs, including:

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Fig. 5. (a) Schematic illustration of a stemless linear probe (upper panel) and typical sequence design using perylene as a fluorophore (lower panel). (b) Emission spectra of linear probe (5E4-2) in the presence of b3a2 (solid line) and in its absence (dotted line). Solution conditions were 1.0 μM 5E4-2, 2.0 μM b3a2, 100 mM NaCl, 10 mM phosphate buffer (pH 7.0), 20°C. Adapted from reference [38] with permission from the Royal Society of Chemistry.

(1) Extinction coefficient increased by incorporating multiple fluorophores into the DNA duplex. (2) Photochemical stability by use of fused aromatic fluorophores. (3) Large apparent Stokes shift achieved by FRET from pyrene to perylene and exciplex formation. (4) Emission wavelength tunable by controlling the assembly of pyrene and perylene in the duplex. Oligonucleotides containing these motifs can be efficiently ligated to native DNA using T4 ligase, demonstrating potential for application as fluorescent tags for various biomolecules.[43]

RNA Modification with Base Surrogates for Improving Strand Selectivity of siRNA and In Vivo Analysis of the RNAi Pathway MicroRNAs (miRNAs) and short interfering RNAs (siRNAs)[44] are attracting attention due to their importance in regulation of gene expression. siRNAs are now used routinely to inhibit specific genes in living systems due to their powerful silencing ability and are being evaluated clinically.[45] Unmodified siRNAs have several drawbacks that must be overcome before widespread clinical use becomes possible. These include susceptibility to nuclease digestion, insufficient strand selectivity leading to off-target effects, induction of the immune response, and difficulties in delivery to target cells. Chemical modification is a powerful and promising method to solve these problems. We have applied our base surrogates to improve strand selectivity.

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The RNA interference (RNAi) pathway involves processing of a precursor into a double-stranded RNA of approximately 20 base pairs and formation of a mature RNA-induced silencing complex (RISC) that contains only the antisense (guide) strand of the siRNA.[46] Misincorporation of sense (passenger) strand leads to silencing of non-target genes. Wedge and dimer motifs have been used to design functional siRNAs with improved strand selectivity and RNAi activity (Figure 7a).[47,48] In order to analyze strand selectivity and RNAi activity simultaneously, we constructed two reporter plasmids that enabled us to evaluate on- and off-target RNAi activity. The target contained a region of the mPIASy gene and the off-target reporter contained a sequence complementary to that present in the target gene. As shown in Figure 7b, in this case, the unmodified siRNA more efficiently suppressed off-target expression than target expression, indicating that the sense strand was predominantly misincorporated into RISC. Introduction of azobenzene (AZ), methyl red (MR), or thiazole orange (TO) using the d-threoninol linker into the sense strand near the 5′ terminus remarkably suppressed off-target activity and raised on-target activity (S5/NA and S7/NA in Figure 7b). Furthermore, incorporation of a dimer motif at near the 5′ terminus of the sense strand (S5MR/A5TO and S7MR/A7TO in Figure 7c) also significantly improved strand selectivity, although RNAi activation was not as great as with the wedge motif. Selective incorporation of the antisense strand into RISC may be closely correlated with the recognition of siRNA by the protein components of RISC. In the crystal structure of the complex between human Argonaute 2



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Fig. 6. (a) Illustration of the sequence design of fluorophore assemblies. (b) Fluorescence emission spectra of duplexes containing six pyrenes. The fluorescence emission spectra at 20°C of the I-W (red line), dimer (blue line), trimer (green line), and cluster (black line) motifs are shown. Magnified spectra at 360–430 nm and photographs of the I-W and cluster motifs are depicted in the inset. (c) Fluorescence emission spectra of hetero-duplexes. The fluorescence emission spectra at 20°C of hetero I-W (red line), dimer (blue line), trimer (green line), and cluster (black line) motifs are shown. Magnified spectra at 450–600 nm and photographs of all the motifs are depicted in the inset. The excitation wavelength was 345 nm. Conditions are 100 mM NaCl, 10 mM phosphate buffer (pH 7.0), 1.0 μM DNA. Adapted with permission from reference [43]. Copyright (2010) American Chemical Society.

(hAGO2) and RNA, the first ten residues from the 5′ terminus are contacted by the protein.[49] The fifth and seventh residues are particularly important because 2′-OH groups of these residues form hydrogen bonds with amino acid residues of AGO2. Since modified siRNAs such as S5/NA and S7/NA do not have 2′-OH groups but instead contain the non-ribose threoninol at the fifth and seventh residues, AGO2 cannot interact with the 5′ region of the sense strand in these modified siRNA duplexes. Accordingly, the unmodified antisense strand is selectively loaded onto RISC.


This selective uptake of the antisense strand enables selective labeling of mature RISC and allowed us to trace the fate of the complex.[48] Functional siRNA S5MR/A5TO itself does not emit fluorescence because the fluorophore is quenched by stacking with MR (Figure 8a). Processing of 5′-terminal residues of the siRNA by Dicer did not alter fluorescence (Figure 8a, upperright panel). Upon formation of mature RISC the sense strand is separated from the passenger strand containing the quencher, and strong emission is recovered (Figure 8a, lower-right panel). Using this system, we were able to follow the fate of the siRNA

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Fig. 7. (a) Sequence design of functional siRNAs applying wedge and dimer motifs. (b) Suppression of reporter gene (firefly luciferase) expression and expression of an off-target gene by siRNAs incorporating the wedge motif. (c) Selectivity and activity of siRNAs incorporating the dimer motif. Adapted with permission from references [47] and [48]. Copyright (2011) Wiley-VCH Verlag GmbH & Co. KGaA and copyright (2013) the Royal Society of Chemistry.

after transfection by confocal laser fluorescence microscopy (Figure 8b). At the early stage after transfection of S5MR/ A5TO, fluorescence was hardly observed in living cells. This delay reflects the uptake of the RNA–lipofectamine complex into cells, the translocation to the cytosol, and subsequent assembly with RNAi-related protein. Fluorescence intensity was almost saturated at 16 h, suggesting that mature RISC was formed. In order to confirm that fluorescence originated from TO in RISC, we conducted immunofluorescent analyses. We labeled AGO2 and GW182, two proteins involved in RNAi, and found that fluorescence of Alexa 568 from the secondary antibody used to visualize either AGO2 or GW182 completely merged with that from TO. Hence, selective monitoring of mature RISC was achieved with a functional siRNA containing a fluorophore and a quencher in the dimer motif.

New Artificial Nucleic Acids: aTNA and SNA Acyclic threoninol nucleic acid (aTNA). Certain functional molecules appended to d-threoninols are highly compatible with

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natural DNA and RNA duplex formation and do not alter sequence specificity or duplex stability. In 2005, a very interesting paper was published by Meggers et al.[50] indicating that the glycol nucleic acid (GNA; see Figure 9) synthesized via conventional phosphoramidite chemistry from an acyclic propylene glycol forms a very stable homo-duplex.[50,51] These reports prompted us to synthesize a new artificial nucleic acid from acyclic d-threoninol (Figure 9).[52] We found that the aTNA/aTNA duplex forms a surprisingly stable antiparallel homo-duplex (Table 1). The Tm of one 8-mer aTNA/aTNA duplex is 62.7°C, whereas those of DNA/DNA and RNA/ RNA duplexes of the same base sequence are only 29.0 and 38.9°C, respectively. The aTNA homo-duplex is even more stable than PNA (53.6°C) or GNA homo-duplexes (71.0°C; aTNA of the same sequence, 78.1°C).[53] To the best of our knowledge, aTNA is the most stable homo-duplex composed of canonical Watson–Crick base pairs yet synthesized. GNA forms a remarkably stable homo-duplex because it is preorganized in the single-stranded state.[50] Unlike GNA or DNA, single-stranded aTNA does not have a circular



THE CHEMICAL RECORD

Fig. 8. (a) Schematic illustration of the selective visualization of mature RISC using S5MR/A5TO with the thiazole orange (fluorophore) and methyl red (quencher) pair. (b) Time-lapse analysis of S5MR/A5TO in living HeLa cells. Fluorescence images were captured at the indicated time points after transfection by confocal laser fluorescence microscopy. Adapted with permission from reference [48]. Copyright (2013) the Royal Society of Chemistry.

Fig. 9. Chemical structures of DNA, RNA, GNA and aTNA.

dichroism (CD) spectrum, suggesting that single-stranded aTNA is not structured. Thus, pre-organization is not necessarily a prerequisite for stable duplex formation. aTNA does not form a stable hybrid with native DNA or with RNA. Serinol nucleic acid (SNA). When designing artificial nucleic acids (XNAs) that can recognize native DNA or RNA,


one might think that the scaffold should be rigid to reduce entropy loss. In fact, most of the XNAs that hybridize with DNA or RNA possess rigid cyclic scaffolds;[54] the exception is PNA. Bridged (locked) nucleic acid (B(L)NA) appears to minimize entropy loss due to binding to RNA by pre-adopting an A-form conformation.[55] We took the opposite strategy for design of an

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Table 1. Sequences and melting temperatures (Tm) of homo- and hetero-duplexes. Sequences

Tm / °C[a]

D8a: 5′-GCATCAGT-3′ D8b: 3′-CGTAGTCA-5′ R8a: 5′-GCAUCAGU-3′ R8b: 3′-CGUAGUCA-5′ R8a: 5′-GCAUCAGU-3′ D8b: 3′-CGTAGTCA-5′ D8a: 5′-GCATCAGT-3′ R8b: 3′-CGUAGUCA-5′ S8a: (S)-GCATCAGT-(R) D8b: 3′-CGTAGTCA-5′ S8a: (S)-GCATCAGT-(R) R8b: 3′-CGUAGUCA-5′ T8a: 1′-GCATCAGT-3′ D8b: 3′-CGTAGTCA-5′ T8a: 1′-GCATCAGT-3′ R8b: 3′-CGUAGUCA-5′ T8a: 1′-GCATCAGT-3′ T8b: 3′-CGTAGTCA-1′ T8c: 1′-CGTAGTCA-3′ T8d: 3′-GCATCAGT-1′ S8a: (S)-GCATCAGT-(R) S8b: (R)-CGTAGTCA-(S) S8c: (S)-CGTAGTCA-(R) S8d: (R)-GCATCAGT-(S) S8e: (S)-GCATTACG-(R) S8f: (R)-CGTAATGC-(S) P8a: (NH2)-GCATCAGT-(CONH2) P8b: (CONH2)-CGTAGTCA-(NH2)

29.0

Duplexes 8-mer DNA/DNA RNA/RNA DNA/RNA DNA/RNA DNA/SNA RNA/SNA DNA/aTNA RNA/aTNA aTNA/aTNA aTNA/aTNA SNA/SNA SNA/SNA SNA/SNA PNA/PNA 15-mer DNA/DNA RNA/RNA SNA/SNA aTNA/aTNA GNA/GNA SNA/RNA

D15a: 5′-CACATTATTGTTGTA-3′ D15b: 3′-GTGTAATAACAACAT-5′ R15a: 5′-CACAUUAUUGUUGUA-3′ R15b: 3′-GUGUAAUAACAACAU-5′ S15a: (S)-CACATTATTGTTGTA-(R) S15b: (R)-GTGTAATAACAACAT-(S) T15a: 1′-CACATTATTGTTGTA-3′ T15b: 3′-GTGTAATAACAACAT-1′ G15a: 3′-CACATTATTGTTGTA-2′ G15b: 2′-GTGTAATAACAACAT-3′ S15a: (S)-CACATTATTGTTGTA-(R) R15b: 3′-GUGUAAUAACAACAU-5′

38.9 25.7 27.3 23.5 35.0 — — 62.7 58.1 51.1 51.2 47.0 53.6

47.0 49.3 68.7 78.1 71.0[b] 47.8

[a] Conditions: 2.0 μM oligonucleotide, 100 μM NaCl, 10 mM phosphate buffer (pH 7.0). [b]From reference [51]. Adapted with permission from reference [53]. Copyright (2013) Wiley-VCH Verlag GmbH & Co. KGaA.

XNA: we engineered the flexibility necessary to accommodate DNA or RNA by sacrificing entropy loss. We used “serinol” as a new scaffold by removing the methyl group on the main chain of threoninol.[56] This design affords the serinol nucleic acid

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oligomer with a very unique stereochemical property: unlike threoninol, serinol is not only flexible but achiral due to the lack of the methyl group (Figure 10a). Hence, the chirality or helicity of the SNA oligomer synthesized from corresponding optically pure phosphoramidite monomers depends only on the sequence. As shown in Figure 10b, the mirror image of a symmetrical sequence such as T→T is identical to the original oligomer (Figure 10b, lower panel), indicating that an SNA oligomer with symmetrical sequence is achiral. In contrast, an asymmetrical SNA oligomer such as A→T is chiral (Figure 10b, upper panel). However, interestingly, its enantiomer is identical to the oligomer of reversed sequence, T→A. More specifically, chirality of the oligomer can be exactly inverted by reversing the sequence of SNA monomers (i.e., two enantiomers can be synthesized from the same chiral monomers but not from their enantiomeric monomers). These stereochemical properties of SNA oligomers as illustrated in Figure 11a are reflected in their melting behaviors and CD spectra. The Tm of the chiral duplex S8a/S8b is 51.1°C (Table 1), and its CD spectrum is characterized by a positive-to-negative Cotton effect (Figure 11b, solid line), indicating that the S8a/S8b duplex is a right-handed helix. In contrast, S8c/S8d, with the reversed sequence relative to S8a/S8b (i.e., it is the enantiomer of S8a/S8b) has an inverse CD signal (Figure 11b, broken line) and thus forms a lefthanded helix. As expected, the Tm of S8c/S8d was the same as that of S8a/S8b (51.2°C). The symmetrical S8e/S8f duplex (Tm 47.0°C) has no induced CD (Figure 11b, dotted line). As described above, aTNA does not hybridize with DNA or RNA. However, removal of the methyl group from threoninol allowed hybridization with both DNA and RNA! The Tm of the SNA/RNA (S8a/R8b) hetero-duplex was 35.0°C, which was remarkably higher than that of the corresponding DNA/RNA duplex (D8a/R8b, 27.3°C). The Tm of the fifteen-base-pair S15a/R15b hetero-duplex (47.8°C) was almost the same as the corresponding RNA homo-duplex (R15a/R15b, 49.3°C), demonstrating the potential of the use of SNAs in antisense strategies.[53] SNA also hybridizes with DNA, although the Tm of the hybrid is slightly lower than that of the corresponding DNA/DNA duplex (Table 1). To the best of our knowledge, SNA is the first XNA prepared from an acyclic scaffold that contains a phosphodiester linkage that hybridizes with both DNA and RNA with reasonable stability. Why are aTNA and SNA duplexes so stable? The Tm of the SNA homo-duplex (S8a/S8b, 51.1°C) is lower than that of the corresponding aTNA duplex (T8a/T8b, 62.7°C), but still far higher than DNA and RNA duplexes (29.0 and 38.9°C, respectively). This fact unambiguously demonstrates that preorganization is not critical to the stabilities of aTNA and SNA duplexes. The stabilities of homoduplexes of the same sequences are in the order of aTNA > PNA ≈ GNA ≥ SNA >> RNA > DNA.[53] The stability of the PNA duplex is likely due to the lack of negative



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Fig. 10. (a) Chemical structure of SNA. S and R termini were named based on the chirality of the terminal residue. The SNA monomer used on the DNA synthesizer is also shown. (b) The mirror image of SNA. An asymmetrical sequence ((S)-AT-(R)) is identical to SNA with the reverse sequence ((S)-TA-(R)). SNA with a symmetrical sequence ((S)-TT-(R)) is identical to its mirror image. Adapted with permission from reference [56]. Copyright (2011) Wiley-VCH Verlag GmbH & Co. KGaA.

Fig. 11. (a) Schematic illustration of the relationship between the SNA duplexes, and (b) their CD spectra. Note that the S8c/S8d duplex is an enantiomer of S8a/S8b, and the S8e/S8f duplex with symmetrical sequence is achiral. Conditions: 4.0 μM oligonucleotide, 100 μM NaCl, 10 mM phosphate buffer (pH 7.0). Adapted with permission from reference [56]. Copyright (2011) Wiley-VCH Verlag GmbH & Co. KGaA.

charge on the main chain.[57] However, the Tm of the PNA duplex is similar to that of the SNA duplex and far lower than the aTNA duplex, suggesting that the neutral backbone is not the only reason for the stability. We hypothesize that PNA, GNA, aTNA, and SNA homo-duplexes are highly stable because they are composed of acyclic scaffolds. Flexibility of acyclic scaffolds may allow the chains to adopt conformations optimal for duplex formation; however, too much flexibility significantly reduces the stability of a duplex due to the excessive entropy loss.[58] We believe that the cyclic structures of the ribose rings of DNA and RNA destabilize these duplexes due to


the steric hindrance relative to the duplexes formed from acyclic scaffolds. Sequence specificity may be one of the reasons why ribose is the natural scaffold: the Tm reduction caused by a mismatch in an XNA hybrid is less than that caused by a mismatch in a DNA or RNA duplex.[52,53,56] Thus, although not as stable, cyclic backbones enable mismatch recognition.

Summary and Outlook Using d-threoninol as a scaffold for the tethering of functional moieties such as dyes, we have shown that the functional motifs

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depicted in Figure 2 can be applied to: (1) photoregulation of DNA functions, (2) design of functional fluorescent probes, (3) creation of fluorophore assemblies that mimic inorganic quantum dots, (4) preparation of siRNAs with improved strand selectivity, and (5) fluorescent labeling of RISC. Acyclic scaffolds have also been utilized by other groups to functionalize nucleic acids.[59–61] With d-threoninol, there is no limitation on the available functional molecules. Even quaternized molecules that are difficult to tether on ribose due to their instability during the DNA synthesis are easily incorporated.[62] Acyclic threoninol was used as a scaffold for nucleobases to afford the artificial nucleic acid termed aTNA, which forms the most stable homo-duplex yet characterized. SNA, created from a scaffold that lacks the methyl group present on the main chain of aTNA, recognizes both DNA and RNA with reasonable affinity. To date, most functionalized oligonucleotides have been developed based on the ribose scaffold. The results summarized here definitively and clearly show that cyclic ribose is not the only scaffold that can tether nucleobases and functional molecules. The ease of synthesis of functionalized d-threoninol and serinol monomers is a great merit for practical applications. Functional base surrogates can be synthesized from d-threonine with low cost and can be introduced at any position and in any number in an otherwise natural oligonucleotide chain using an automated DNA synthesizer. SNA is also easily synthesized from inexpensive l-serine. In the future, we expect that SNA will be used rather than PNA when hybrids with RNA or DNA are desired, because SNA oligomers are free from several of the drawbacks of PNA, such as non-specific binding to proteins and low water solubility (aggregation) due to its hydrophobic nature.[63,64] We expect that SNA phosphoramidite monomers will be coupled more efficiently than conventional DNA monomers because trivalent phosphorus is bound to not a secondary but a primary hydroxyl group. Thus, very long SNA may be obtained with good yield. Facile conjugation of SNA with DNA and RNA via conventional phosphoramidite chemistry will allow antisense and siRNA modifications. Functional nucleobases can be easily introduced into SNA. Threoninol and serinol open up a new “acyclic world” for the creation of various functional artificial oligonucleotides for practical applications. Our studies of acyclic scaffolds raise several questions. Since a stable cluster motif can be prepared, complementary hydrogen bonds appear unnecessary for stable duplex formation. Häner et al. also reported a new supramolecular assembly stabilized strictly by stacking interactions.[65] Furthermore, ribose-like rigid ring structures are not necessary for formation of double-helical structures containing the natural bases. Flexible acyclic scaffolds functionalized with the natural bases form more stable duplexes than DNA or RNA. Why has Nature selected (deoxy)ribose as a scaffold? Glycerol and threoninol are much simpler and synthetically less costly than ribose. One

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possible answer is that acyclic scaffolds do not as effectively discriminate mismatches as do DNA and RNA, but this is likely not the complete answer. The Eschenmoser group has begun pioneering work on the chemical etiology of nucleic acid structure with a sugar backbone.[54] We anticipate that as this research proceeds several mysteries of double-helical structure will be clarified.

Acknowledgements This work was supported by a Grant-in-Aid for Scientific Research (A) (no. 25248037), a Grant-in-Aid for Scientific Research for Young Scientists (B) (no. 24750173), and Grantsin-Aid for Scientific Research on Innovative Areas “Molecular Robotics” (no. 24104005) and “Dynamical Ordering & Integrated Functions” (no. 26102518) from the Ministry of Education, Culture, Sports, Science and Technology, Japan. Support from the Naito Foundation Natural Science Scholarship (for H.K.) and the Canon Foundation (for H.A.) is also acknowledged.

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De novo design of functional proteins: Toward artificial hydrogenases.

de novo computational enzyme design.

Nanomedicine: de novo design of nanodrugs.

De novo design of sequences for nucleic acid structural engineering.

Accurate de novo design of hyperstable constrained peptides.

De novo design - hop(p)ing against hope.

Advances in multiparameter optimization methods for de novo drug design.

Protein folding and de novo protein design for biotechnological applications.

Multidimensional de novo design reveals 5-HT2B receptor-selective ligands.

Multi-objective molecular de novo design by adaptive fragment prioritization.

High-fidelity de novo synthesis of pathways using microchip-synthesized oligonucleotides and general molecular biology equipment.

Computer design of bioactive molecules: a method for receptor-based de novo ligand design.

De Novo Design of an Allosteric Metalloprotein Assembly with Strained Disulfide Bonds.

Computational de novo design of a self-assembling peptide with predefined structure.

De novo design of multitarget ligands with an iterative fragment-growing strategy.

De novo design of protein homo-oligomers with modular hydrogen-bond network-mediated specificity.

De novo status epilepticus with isolated aphasia.

De novo phasing with optimized XFEL data.

De novo AVM formation.

De novo selection of oncogenes.

De novo design of transmembrane helix-helix interactions and measurement of stability in a biological membrane.

Rational design of inducible CRISPR guide RNAs for de novo assembly of transcriptional programs.

De Novo Design of Skin-Penetrating Peptides for Enhanced Transdermal Delivery of Peptide Drugs.

De novo design and experimental characterization of ultrashort self-associating peptides.