Conformationally Constrained Oligonucleotides for RNA Targeting

To My Family

List of Papers

This thesis is based on the following papers, which are referred to in the text by their Roman numerals.

I Upadhayaya, R.*; Deshpande, S. G.*; Li, Q.*; Kardile, R. A.; Sayyed, A. Y.; Kshirsagar, E. K.; Salunke, R. V.; Dixit, S. S.; Zhou, C.; Földesi, A.; Chattopadhyaya, J. Carba-LNA-5MeC/A/G/T Modified Oligos Show Nucleobase- Specific Modulation of 3′-Exonuclease Activity, Thermody- namic Stability, RNA Selectivity, and RNase H Elicitation: Synthesis and Biochemistry *Co-first authorship with Upadhayaya, R and Deshpande, S. G J. Org. Chem. 2011, 76, 4408-4431.

II Li, Q.; Yuan, F.; Zhou, C.; Plashkevych, O.; Chattopadhyaya, J. Free-Radical Ring Closure to Conformationally Locked α-L- Carba-LNAs and Synthesis of Their Oligos: Nuclease Stability, Target RNA Specificity, and Elicitation of RNase H J. Org. Chem. 2010, 75, 6122-6140.

III Li, Q.; Plashkevych, O.; Upadhayaya, R.; Deshpande, S. G.; Földesi, A.; Chattopadhyaya, J. The Physicochemical Properties of DNA-RNA Duplexes Con- taining Pure 7′R-Me- or 7′S-Me-Carba-LNA Derivatives of A, G, 5-MeC or T in the DNA Strand: Diastereomer Specific Com- parison of The 3′-Exonuclease Stability and RNase H Elicita- tion Submitted 2012.

Reprints were made with permission from the respective publishers.

Contribution Report

The author wishes to clarify his contributions to the research papers presented in the thesis:

Paper I: Designed research with RamShankar Upadhayaya, Sachin Gangad- har Deshpande and Prof. Jyoti Chattopadhyaya (J. C.). Synthesized modified oligonucelotides, performed studies of thermal denaturation, SVPDE, blood serum, and RNase H digestions. Interpreted the data obtained from enzymatic studies and wrote enzymatic part of the manuscript.

Paper II: Designed research with Prof. Jyoti Chattopadhyaya (J. C.). Synthe- sized, purified and characterized all the intermediates for α-L-carba-LNA derivatives. Incorporated them into oligos and performed all the physicochemical and biochemical studies (including thermal denaturation, circular dichroism, SVPDE, blood serum and RNase H digestions) towards modified oligonucleotides. Interpreted the data and wrote the first draft of the manuscript. [Note: Dr. Oleksandr Plashkevych (O. P.) contributed to the solvation free energy calculations and Dr. Chuanzheng Zhou (C. Z.) helped to revise and correct the manuscript]

Paper III: Designed research with Prof. Jyoti Chattopadhyaya (J. C.). Syn- thesized, purified and characterized all the intermediates for 7′-Me-carba- LNA-A, -G, -MeC and -T analogues. Incorporated them into oligos and performed all the physicochemical (Tm measurement and thermodynamic study) and enzymatic studie (SVPDE, blood serum, E. coli RNase H1). Interpreted the data and wrote the first draft of the manuscript. Computational calculations were performed by Dr. Oleksandr Plashkevych (O. P.) in collaboration with Qing Li (Q. L.) and Prof. Jyoti Chattopadhyaya (J. C.).

Contents

1. Introduction...... 11 1.1 General introduction to nucleic acids...... 11 1.2 Components of nucleic acids...... 12 1.3 Structural properties of nucleotides and nucleic acids ...... 13 1.3.1 Nucleotide conformation ...... 13 1.3.2 Structural features of nucleic acids...... 14 1.4 Nucleic acids for therapeutic application ...... 16 1.4.1 Nucleic acid-based therapeutics ...... 16 1.4.1.1 Antisense oligonucleotide...... 16 1.4.1.2 Triple-helix forming oligonucleotide...... 16 1.4.1.3 Ribozyme and DNAzyme ...... 17 1.4.1.4 RNA interference (siRNA and miRNA)...... 17 1.4.1.5 Nucleic acid aptamers ...... 18 1.4.2 Chemical synthesis of oligonucleotides...... 18 1.4.3 Chemical modifications in oligonucleotides ...... 19 1.4.3.1 Backbone modifications...... 20 1.4.3.2 Sugar Moiety modifications...... 20 1.4.3.3 Base modifications...... 21 1.5 Overview of the thesis...... 22 2. Conformationally constrained nucleosides: introduction, synthesis and structural characterization...... 23 2.1 Brief introduction of conformationally constrained nucleos(t)ides....23 2.1.1 Recent advances of intramolecular free-radical cyclization reactions on pentose sugars ...... 25 2.2 Synthesis and structural elucidation of 7'Me-cLNA-A, -G, 5MeC, and T as well as (6'OH,7'Me)-α-L-carba-LNA-T nucleosides (Paper I- III) ...... 26 2.2.1 Synthesis of diastereomerically pure (7'S- or R-Me)-cLNA-A, -G, -5MeC, and -T nucleosides and their phosphoramidites...... 26 2.2.2 Structural evidence of free-radical ring closure products in the synthesis of cLNA-A, -G, -MeC and -T nucleosides ...... 29 2.2.2.1 Confirmation of bicyclic systems in 5-exo cyclization products...... 29 2.2.2.2 Stereochemistry of cLNA nucleosides...... 30

2.2.3 Synthesis of diastereomerically pure (6'-OH,7'-Me)-α-L- carba-LNA-T nucleosides...... 31 2.2.4 NMR characterization of key intermediates involved in the synthesis of α-L-carba-LNA analogues...... 33 2.2.4.1 Confirmation of bicyclic and tetracyclic systems by HMBC and COSY experiments...... 33 2.2.4.2 Stereochemistry of key intermediates in the synthesis of α- L-carba-LNA analogues...... 34 2.2.5 Mechanism of the free-radical cyclization and radical rearrangement involved in the synthesis of α-L-carba-LNA analogues ...... 36 3. Antisense properties of modified AONs containing α-L-carba-LNAs and 7'-Me-carba-LNAs (Paper I-III)...... 38 3.1 Thermo-stabilities of chemically modified AONs toward the RNA and DNA targets...... 38 3.1.1 Binding affinity of AONs containing α-L-carba-LNA and α-L- LNA thymines toward complementary RNA and DNA...... 38 3.1.2 Binding affinity and thermodynamic properties of 7'-Me- cLNA-A, -G, -MeC and -T modified AONs toward complementary RNA and DNA strands...... 40 3.2 Nuclease stabilities of modified AONs...... 46 3.2.1 Nucleolytic stabilities of AONs modified with α-L-carba-LNA derivatives and α-L-LNA ...... 46 3.2.2 Nucleolytic stabilities of AONs containing 7'R- and S-Me- cLNA-A, -G, -MeC and -T nucleosides ...... 47 3.3 RNase H-mediated RNA degradation in modified AON/RNA hybrids...... 50 3.3.1 RNase H elicitation induced by AONs modified with α-L- carba-LNA and α-L-LNA analogues...... 50 3.3.2 RNase H elicitation induced by AONs modified with 7'-Me- cLNA-A, -G, -MeC, -T and LNA-A, -G, -C, -T analogues...... 52 Sammanfattning...... 54 Acknowledgements...... 56 References...... 58

Abbreviations

1D One dimensional 2D Two dimensional A Adenosine Ade Adenine AIBN Azobisisobutyronitrile AMD Age-related macular degeneration AONs Antisense oligonucleotides aza-ENA 2′-N,4′-C-Ethylene bridged nucleic acid BNA Bridged nucleic acid Bu3SnH Tributyltinhydride C Cytidine CEM 2-Cyanoethoxymethyl cENA Carbocyclic 2′-O,4′-C-ethylene bridged nucleic acid cLNA Carbocyclic locked nucleic acid COSY Correlation spectroscopy Cyt Cytosine DEPT Distortionless enhancement by polarization transfer DMTr 4′,4′-Dimethoxytrityl DNA Deoxyribonucleic acid dsRNA Double-stranded RNA ENA 2′-O,4′-C-Ethylene bridged nucleic acid FDA Food and drug administration G Guanosine Gua Guanine HMBC Heteronuclear multiple bond correlation HMQC Heteronuclear multiple quantum coherence HNA Hexitol nucleic acid LNA Locked nucleic acid miRNA Micro-RNA MOE 2′-O-Methoxyethyl mRNA Messenger RNA ncRNA Non-coding RNA NMO N-Methylmorpholine-N-oxide NMR Nuclear magnetic resonance NOE Nuclear Overhauser effect

PAGE Polyacrylamide gel electrophoresis piRNA Piwi-interacting RNA PNA Peptide nucleic acid PTC Phenoxythiocarbonyl RISC RNA-induced silencing complex RNA Ribonucleic acid RNAi RNA interference RP-HPLC Reverse-phase high performance liquid chromatography SELEX Systematic evolution of ligands by exponential enrichment siRNA Small interfering RNA snoRNA Small nucleolar RNA SVPDE Snake venom phosphodiesterase T Thymidine TBAF Tetra-n-butylammonium fluoride tBDMS Tert-butyldimethylsilyl TEM 2-(4-tolylsulfonyl)ethoxymethyl TFOs Triple-helix forming oligonucleotides Thy Thymine TMS-Cl Trimethylsilyl chloride TMSOTf Trimethylsilyl trifluoromethanesulfonate TOM [(triisopropylsilyl)oxy]methyl TPAP Tetra-n-propylammonium perruthenate tRNA Transfer RNA TS Transition state U Uridine Ura Uracil UV Ultraviolet φm Puckering amplitude

1. Introduction

1.1 General introduction to nucleic acids Nucleic acids are the macromolecules responsible for the hereditary information that regulates the protein synthesis in living organisms. The name “nucleic” derives from the fact that they were discovered (by the Swiss bio- chemist Friedrich Miescher, in 1869) within the cell nucleus. In nature, nucleic acids exist as two analogous chemical forms: deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). The main role of DNA is the long-term storage and carrier of genetic information,1,2 whereas RNA has multiple cellular functions. Three kind of RNAs are identified, the largest subgroup (85 to 90%) being ribosomal RNA, rRNA, together with proteins, is the major component of ribosomes, which has been claimed to be a catalyst for peptide bond formation in the ribosome.3,4 Other two forms of RNA are messenger RNA (mRNA) and transfer RNA (tRNA), in which mRNA is transcribed from DNA, and serves to carry information from DNA to the protein synthesis machinery called ribosomes; tRNA is a small RNA and serve as transporters of amino acids in the translation process. As we know, aforementioned highly abundant and functionally important RNAs such as tRNA and rRNA are not translated into proteins therefore called non-coding RNA (ncRNA).5,6 There are also some other types of ncRNAs such as small nucleolar RNA (snoRNAs),7 microRNAs,8-14 siRNAs,15,16 piRNAs17-19 and riboswitch,20-22 which have been discovered in the past decades. Some of ncRNAs are involved in the regulation of gene expression, however, the number and function of ncRNAs encoded within the human genome are still unknown.6 RNA is considered to be able both to store genetic information, like DNA, and to catalyze chemical reactions, like an enzyme protein. The diverse functions of the RNA molecules led to the “RNA World” hypothesis, which proposes that life based on ribonucleic acid (RNA) pre-dates the current world of life based on deoxyribonucleic acid (DNA), RNA and proteins. It speculates that in a primordial age, RNA could not only act as a carrier of genetic information to the progeny, but also could carry out complex catalytic activities for self-replication and other complex biochemical reactions.23-25

11 1.2 Components of nucleic acids Nucleic acids, i. e. DNA and RNA are polymers of nucleotides linked in a chain through phosphodiester bonds (Figure 1). Therefore, nucleotides are the building blocks of all nucleic acids. Nucleotides have a distinctive structure composed of three components covalently bound together: a nitrogen- containing “base” (either a pyrimidine or purine); a 5-carbon pentofuranose sugar (2′-deoxy-D-ribose in DNA or D-ribose in RNA); and a phosphate group. The combination of a heterocyclic aromatic nucleobase and sugar through an N-glycosidic bond in β configuration is called a nucleoside. There are five common bases: adenine (Ade), guanine (Gua), which are linked at N9 to C1′ of the sugar, and cytosine (Cyt), thymine (Thy), uracil (Ura) linked at N1. The purines (adenine and guanine) and pyrimidines (cytosine and thymine) are represented in DNA, whereas RNA contains the pyrimidine uracil instead of thymine (Figure 1). Moreover, the presence of 2′-OH substituent at the ribose of RNA compared to deoxyribose of DNA makes RNA molecules structurally and functionally different. It is known that DNA exists as a double-stranded form in the cell, while most cellular RNA molecules are single stranded, they can form secondary structures such as stem-loop, bulge-loop and hairpin, etc.26 The extra 2′-OH group makes RNA less stable than DNA due to the easy attack of this hydroxyl group to the neighbouring phosphate.27

NH 2 O N N NH 5' A T N N O N O O O c N 5 h 6 NH a Thymin-1-yl 7 1 i 8 2 n O R G N 4 N NH d 5' 9 2 ir O P O 3 e c O NH2 t 4' 2' 1' io O 4 n 3' 5 N 3 O R C 2 6 N1 O O P O O O O nucleotide unit NH O R U N O O P O O O

O R 3' Figure 1. Atomic numbering and primary structure of DNA and RNA. R = OH represents RNA, R = H and U is replaced by T represent DNA.

12 1.3 Structural properties of nucleotides and nucleic acids

1.3.1 Nucleotide conformation As discussed above, the nucleotide is composed of sugar moiety, phosphate group and nucleobase. Therefore, the conformation of a nucleotide is determined by the torsion angles α, β, γ, δ, ε and ζ in the sugar-phosphate backbone; ν0-ν4 in the pentofuranose sugar ring; and χ for the N-glycosidic bond28 (Figure 2-3 in ref. 29). If the precise torsion angle for a conformation is not available, it is often convenient to specifiy it roughly by defining a conformational region such as cis (c) = 0 ± 30°, +gauche (g+) = 60 ± 30°, +anticlinal (a+) = 120 ± 30°, trans (t) = 180 ± 30°, -anticlinal (a-) = 240 ± 30°, -gauche (g-) = 300 ± 30°. Besides this, the range 0 ± 90° is denoted as syn and the range 180 ± 90° is denoted as anti.28,29 The rotational position of the nucleobase relative to the sugar is sterically restricted, and only two conformational states are preferred, syn (-90° ≤ χ ≤ 120°) and anti (90° ≤ χ ≤ 270°).30,31Anti orientation is more favourable compared to syn orientation, because there is no special steric hindrance between sugar and base in anti conformation, while in syn conformation, the bulky part of the base is located over the the sugar, leading to close interactions. It is noteworthy that the torsion angle χ can be tuned by the sugar pucker (C2′- 32,33 34,35 endo versus C3′-endo), base types (purine versus pyrimidine), and chemical modification on the base.36-39 The sugar moiety of a nucleotide is the five-membered furanose ring, which is never planar. Obviously, the planar furanose is energetically unfavorable, because in this arrangement, all torsion angles are 0° and the substituents of carbon atoms are fully eclipsed.29 Therefore, they are puckered in either envelope (E) or twist (T) forms to reduce the energy.40 In order to accurately describe the conformation of ribose and deoxyribose rings in nucleosides and nucleotides, the conception of pseudorotation has been introduced with two important parameters: the pseudorotation phase angle P (tan P = [(ν4+ν1)-(ν3+ν0)]/2ν2(sin36+sin72) and the puckering 41 amplitude φm (φm = ν2/cos P). In nucleotide structural analysis two ranges 42 of pseudorotation phase angles are preferred: C2′-endo at 137° ≤ P ≤ 194° (in the “South” of the circle, or S), and C3′-endo at -1° ≤ P ≤ 34° (“North” , or N, see Figure 3). Generally, the deoxyribofuranosyl sugars in B-type DNA adopt preferentially C2′-endo twist form (S-type conformation), whereas ribofuranosyl sugars in A-type RNA are in C3′-endo twist form (N- type conformation). Pentofuranose conformations in nucleic acids are not static and in solution they are involved in a dynamic two-state North ↔ South equilibrium. Using temperature-dependent NMR Chattopadhyaya et al first suggested42-44 that different stereoelectronic effects (Gauche and Ano- meric Effects) actually control this two-state dynamic North ↔ South equilibrium. Thus the modification of the sugar ring as well as aromatic nature of the nucleobase can drive the sugar ring to adopt predominantly one type of conformation. 45-48

13 1.3.2 Structural features of nucleic acids When describing structural features of nucleic acids, base-base interactions are always the critical considerations. Hydrogen bonding for base pair formation and base stacking are considered as the main forces which stabilize the associations between bases of nucleic acids. Normally, purine and pyrimidine can form base pair through hydrogen bond. There are twenty- eight possible base-pairs arrangements with dyad, pseudodyad, and no sym- metry.29 Amongst them, Watson-Crick,49 Hoogsteen50,51 and Wobble52,53 type base pairing are the most populated ones (Figure 2). At higher salt concentrations, oligomeric base multiplets such as T.A.T and C.G.C+ triplexes54, and G-quadruplexes55,56 can form. Though first considered as an anomaly, of these, triplex structures have been found in potential use as a therapeutic, by inhibiting transcription,57 or as a sequence specific cleavage reagent. 58

H O H N H O H N O N N N O H N Ura Cyt N H N Gua N N Thy N H N Ade N N N Gua O N N N H N N H O H N O H NH2 Wobble G-U Watson-Crick A-T Watson-Crick G-C

H Hoogsteen N N N H N Cyt N H N Gua N N N O Thy N Gua N O H O H N H H H N N N Hoogsteen H O H O O H N H H N H N O N Hoogsteen M+ H H O Cyt N H N Gua N Thy H N Ade N N N N N N H O H N N N N Gua Gua O H N H O N N H T-A-T triplet N C-G-C+ triplet N H N N H G-quadruplexes Figure 2. Some representatives of base pairing types (horizontal base-base interactions).

Except the horizontal base-base interactions (base pair formation), the vertical base stacking is also of importance for the stabilization of nucleic acid helices.59 The base stacking is stabilized by dipoles, π-electron systems, dipole-induced dipole moments (London dispersion59), hydrophobic bond- ing60,61 and van der Waals interactions. In aqueous solution, base stacking dominates while base-base hydrogen bonding is greatly suppressed due to the competition at binding sites with water molecules. Normally, the stacking strength between purine and pyrimidine bases follows the trend: purine- purine > pyrimidine-purine > pyrimidine-pyrimidine,62-64 which explains why poly(A) oligos mainly exist as helical form, and why poly(U) is predominantly in random coil form at room temperature. Generally, there are mainly three kinds of double-helical structures in DNA, i.e., A-, B- and Z-DNA. Amongst them, A- and B-DNA are right- handed, whereas Z-DNA is left-handed. However, the corresponding RNA

14 duplexes only adopt predominantly structurally closely related A- and A′- type conformations, showing the conservatism of RNAs.29 In nature, B-DNA occurs dominantly, which has wide major groove (ca 22 Å) and narrow mi- nor groove (ca 12 Å).65 In contrast, A-type RNA and DNA have shallow, wide minor groove (ca 10 Å), and narrower, deeper major groove (ca 4 Å). As we know, the essential distinction between the A- and B-forms is in the conformation of the ribose sugar (ring puckering), which is C2′-endo for B- form DNA and C3′-endo for A-forms of DNA or RNA. The structural features of DNA-RNA hybrid have also been well characterized. Since it can be recognized by ribonuclease H family, which cleaves the RNA strand of the hybrid, leading to down-regulation of cellular RNAs,66 the structure of DNA-RNA hybrid has been intensively studied in recent years. Recent NMR and X-ray crystallography studies67-70 suggest that DNA-RNA hybrid adopts an intermediate conformation between A and B form, but globally more like of A-form. The ribose sugar in the RNA chains is in the typical C3′-endo conformation while the deoxyribose in DNA chain 67 is in an unexpected O4′-endo conformation. Hence, chemical modification of sugar moiety in DNA strand of DNA-RNA hybrid to C3′-endo conformation could induce the global conformation of DNA/RNA hybrid to A-form like RNA-RNA duplex, resulting in a more stabilized structure. This explains that the AONs modified with 2′,4′-conformationally constrained nucleosides [such as LNA and carba-LNA derivatives that locking the sugar moiety in a perfect N-type (C3′-endo) conformations] shows favorable RNA specificity when targeted to complementary RNAs and DNAs. Figure 3 shows the representative structures of B-DNA and DNA-RNA hybrid.

Major Groove Major Minor Groove Groove

Minor Groove

DNA-DNA duplex DNA-RNA hybrid Figure 3. Structures of B-form DNA duplex71 and DNA-RNA hybrid.69 These are crystal structures by X-ray crystallography obtained from Nucleic Acid Database

15 1.4 Nucleic acids for therapeutic application

1.4.1 Nucleic acid-based therapeutics Oligonucleotides have proved to be valuable tools in the modulation of gene expression in a highly specific manner. This kind of macromolecules can specifically bind to target genes or proteins, resulting in the downregulation or upregulation of the interested target gene products. Thus, they play important roles in target validation, genomic functions and therapeutic applications. The following sections briefly discuss the different types of oligonucleotide-based strategies, viz. antisense approach,72-74 ribozyme,75,76 DNAzyme,77 RNA interference15 and aptamers,78 etc, their discovery and recent development in therapeutics.

1.4.1.1 Antisense oligonucleotide Antisense oligonucleotides, typically 15-20 nucleotides long, can bind to target RNA sequences through Watson-Crick hybridization, resulting in gene expression regulation in living cells.74 Two major mechanisms have been widely suggested, namely physical blockage and RNase H activa- tion.72,73 One notable discovery in antisense chemistry is the addition of a phosphorothioate backbone to the oligonucleotides, leading to a significant increase in nuclease stability without impairing the ability to hybridize with target mRNA.72 Other chemically modified oligonucleotides including peptide nucleic acid (PNA)79,80 and locked nucleic acid (LNA)81-86 have been developed to increase the efficacy, stability and practicability of antisense molecules. So far, only one antisense oligonucleotide, i.e.VitraveneTM (Fomivirsen) has been approved, which is used against cytomegalovirus-induced retinitis by local injection.87 Other antisense oligonucleotides targeting several genes that are important to severe human diseases such as cancer and infectious diseases have been in clinical trials, some of them even in multiple clinical Phase III trials.74,88 Very recently, antisense oligonucleotides have also been designed and applied to suppress miRNA function efficiently89, making the antisense approach continueously attractive.

1.4.1.2 Triple-helix forming oligonucleotide Triplex-forming oligonucleotides (TFOs)54 are designed to artificially regu- late gene expression through targeting with complementary double-stranded DNA (generally uninterrupted homopurine-homopyrimidine sequence). They are of special interest due to their binding to the gene itself rather than to its transcription product mRNA, as in antisense strategy. This antigene strategy can not only be applied to transciptional inhibition and activa- tion,57,90 but also target DNA cleavage and targeted mutagenesis.58,91 How-

16 ever, the triple helix approach is limited by a few issues including target sequence limitation, target DNA accessibility and delivery system.92 There- fore, no triplex-forming oligonucleotides are on clinical trials.

1.4.1.3 Ribozyme and DNAzyme Ribozymes are based upon catalytic RNAs originally discovered in the pro- tozoan tetrahymena.75,76 Since then, different types of naturally occurring ribozymes have been discovered.93,94 Much like antisense oligonucleosides, ribozymes can be targeted to a variety of molecules, and have been studied against viral infections to cancer cells for potential therapeutic applications. The catalytic nature of ribozymes is highly dependent on their structures, limiting the possibility of modifying their chemistry to improve pharma- cokinetics, efficacy and toxicity profile. Moreover, several ribozymes have been evaluated in clinical trials.95 The DNAzyme molecules were discovered by Ronald and Gerald in 1994 using in vitro selection methodology.77 The most well-characterized DNAzyme is the 10-23 subtype, or called 10-23 DNAzyme, comprising a cation-dependent catalytic core of 15 deoxy- ribonucleotides that is used to bind to and cleave the target RNA between an unpaired purine and paired pyrimidine through a de-esterification reaction.96,97 Unlike ribozymes, DNAzymes are much easier to be synthesized and chemically modified, which makes them attractive therapeutic candidates for further study or clinical evaluation.

1.4.1.4 RNA interference (siRNA and miRNA) First discovered in 1998, RNA interference (RNAi) has since become a standard tool for various types of laboratory research.15 The scientists who discovered this phenomenon have won the 2006 Nobel Prize in Medicine. The plausible mechanism of RNAi is suggested as follows: the endogenous short stretches of dsRNAs (normally 21-23-nucleotide RNA fragments, referred as small interfering RNAs; i.e. siRNAs, derived from the “Dicer” endoribonuclease cleavage of long stretches of dsRNA) or synthetic small dsRNAs enter the multi-nuclease containing RNA-induced silencing complex (RISC), followed by unwinding and subsequent binding to complementary target RNAs, and the enzymes known as “Slicer” in RISC are responsible for specific cleavage of target RNAs98. It has been demonstrated that RNA interference is mediated by 21- and 22-nucleotide small RNAs.99-101 Target genes inactivation by synthetic siRNAs has also been found in different types of mammalian cells.102-104 These findings make RNA interference a new tool for gene function study in higher eukaryotes, and eventually may be used as efficient gene-specific therapeutics. There have already been several siRNA molecules evaluated in human clinical trials.105

17 MicroRNAs (abbreviated miRNAs) are naturally occurring post- transcriptional regulators that bind to complementary mRNA transcripts, usually resulting in translational repression or target RNA degradation.9- 12,106,107 Since its first discovery in 1993,8 hundreds of miRNAs have been recognized in plants and animals.14 It has been suggested that miRNA- mediated regulation of gene products may account for various human diseases including cancer.108,109 Thus, naturally occurring endogenous and ex- ogenous miRNAs represent ideal target for therapy.110 Recently, the first miRNA targeting drug advanced by Santaris Pharma A/S has entered Phase II clinical trial, and it is only a matter of time before the first miRNA-related therapeutic reaches the market.111

1.4.1.5 Nucleic acid aptamers Nucleic acid aptamers are short stretches of DNA or RNA that bind to specific target molecules, usually proteins. It is possible to screen libraries of aptamers employing Systematic Evolution of Ligands by Exponential En- richment (SELEX) to determine the best aptamer for a particular target efficiently.78 Unlike the other oligonucleotide-based approaches, the binding affinity of aptamers are not dependent upon the complementarity, but are highly determined by their specific 3-dimensional structures that can form complexes with target proteins and inhibit their activity.112 For this reason, aptamers can be considered “chemical antibodies”. In addition to their distinct target recognition, aptamers offer advantages over antibodies in that they are readily produced by chemical synthesis, possess desirable storage properties, and elicit little or no immunogenicity in therapeutic applications.113,114 The first aptamer-based drug MacugenTM (Pegaptanib) has been approved by the U.S. Food and Drug Administration (FDA) in treatment for age-related macular degeneration (AMD).115 Nowadays, several aptamers are under different phases of clinical trials.116

1.4.2 Chemical synthesis of oligonucleotides Chemical synthesis of oligonucleotides was first achieved by Todd and Michelson in 1955 through phosphotriester approach.117 Soon after, a completely different strategy named as phosphodiester approach was introduced by Khorana and his co-workers.118-120 In this approach, the internucleotide linkages were left completely unprotected during the assembly of the oligonucleotide sequence.121 Another important innovation in phosphorylation methodology is phosphite triester approach, first developed by Letsinger et. al.122 In 1981, Beaucage and Caruthers improved this P(III) approach and introduced nucleoside phosphoramidites to oligonucleotide synthesis.123 Very soon, the phosphoramidite chemistry was extensively used in solid- supported oligonucleotide synthesis due to its high coupling efficiency and yield.124 The occurrence of phosphoramidite approach combining with solid-

18 supported synthesis make the efficient and high-yielding synthesis of DNA and RNA accessible. Hence, gene machine arised in 1980′s.124 Using this machine, also called DNA/RNA synthesizer, the synthetic time of 20 mer deoxyoligos was significantly shortened to 2 hours with average stepwise yield more than 98%. That means one coupling reaction can be completed within 5 minutes, which greatly avoids irreversible side-reactions. For RNA synthesis, the extra 2′-OH group should be protected during the sequence assembly. Several 2′-OH protecting groups such as 2′-O- tBDMS,125-127 2′-O-TOM,128 2′-O-CEM129 and 2′-O-TEM130 and so on, have been developed and widely used in the RNA automation synthesis. All the protecting groups mentioned above can be removed under the conditions which deprotected RNAs are tolerant to. After the completion of chain assembly, the oligomers are cleaved from the solid support by treatment with ammonia, at the same time, the protecting groups on the base and phosphate linkage are also removed. The crude product hence obtained is purified by RP-HPLC or PAGE to give oligonucleotide with high purity.131

1.4.3 Chemical modifications in oligonucleotides Due to increased target affinity and enzymatic stability compared to native counterparts, modified oligonucleotides have been used for a wide range of purposes in fields such as biotechnology,132 molecular biology,133-135 chemical probes in bioanalysis and diagnostic136-138 as well as therapies aforementioned (antigene,139 antisense,66,140 RNAi,141-143 etc). Normally, the modified nucleosides are transformed to phosphoramidites and incorporated into oligonucleotides through solid-phase DNA/RNA synthesis. It is well known that modifications can be introduced in the base, sugar or phosphate moieties, and they are briefly discussed in the following sections.

19 (A) Examples of bacbone modifications (B) Examples of sugar modifications

a. R1 = CH3, R2 = O; methylphosphonate Base O O O Base Base b. R1 = CH2COOH, R2 = O; phosphonoacetate O O R2 O c. R1 = COOH, R2 = O; phosphonoformate O P R - O 1 d. R1 = S , R2 = O; phosphorothioate - O -O P O O e. R1 = BH3 , R2 = O; borane phosphonate O R O - f. R1 = O , R2 = N; (N3' P5') phosphoramidate R = F, OMe, OMOE, etc. LNA HNA Base (C) Examples of base modifications O O NH2 N NH2 H3C C C N H3C N N NH NH Base N HN N O O N N O N NH O N N N 2 O 2,6-diaminopurine 6-phenyllumazine 5-methylcytosine 5-propynyl U NH Base O H2N O n N C NH O NH NH C NH2 NH O N O N N N N O N N O N N g. Peptide nucleic acid (PNA) N 7-(3-aminopropynyl)-7-deazaadenine pyridopyrimidine phenoxazine tetracyclic adenine Figure 4. Modifications on base, sugar and phosphate linkage moiety of nucleotides

1.4.3.1 Backbone modifications Amongst the very first modifications to oligonucleotides were alterations of phosphodiester backbone to prevent the oligonucleotides from enzymatic degradation within the cell. During recent years, a number of different modifications have been applied to the native phosphodiester linkage in the backbone of oligonucleotides.144 One of the changes involves the alterations to the nonbridging oxygen atoms such as phosphorothioate (P=S),145-147 borane 148,149 150,151 phosphonate (P-BH3), methylphosphonate (P-Me), phosphono- 152 153 acetate (P-CH2COOH) and phosphonoformate (P-COOH), etc, (Figure 5A.a-e) while the bridging atoms change is representatively exemplified by N3′→P5′ phosphoramidate154,155 (Figure 4A.f). Other backbone modifications worth mentioning are those peptide nucleic acids (PNAs, see Figure 4A.g), in which the sugar phosphate backbone is entirely replaced by an achiral polyamide.79,80 Since the backbone of PNA contains no charged phosphate groups, the binding between PNA/DNA strands is stronger than between DNA/DNA strands due to the lack of electrostatic repulsion. Oli- gonucleotides modified at phosphate backbone have shown potential therapeutic properties such as increased binding affinity, improved nuclease resistance, and membrane permeability, etc. Therefore, they are continuously attracting research focus in the field of oligonucleotide-based therapeutics.

1.4.3.2 Sugar Moiety modifications A wide variety of sugar modifications (Figure 4B) have been introduced into antisense oligonucleotides to enhance nuclease stability and target binding affinity.156 Amongst these, the modifications at the 2′-position of the ribose

20 sugar have attracted great interests. The oligonucleotides contaning 2′-O- modifications such as 2′-O-methyl (OMe)157-159 and 2′-O-methoxy-ethyl (MOE)159,160 RNAs are considered to have appropriate electronegative substituents that can induce an RNA-like 3′-endo sugar puckering, which results in improved hybridization affinities. The more electronegative the 2′- substituent, the greater the shift to 3′-endo conformation and the higher Tm values.161 Thus, 2′-F modified oligonucleotides can form the most stable hybrids, but similar to 2′-OMe modified ones, they do not show enough resistance to nucleases for in vivo aplications.162 On the other hand, larger groups at the 2′-position improve nuclease resistance, but at the cost of target affinity. Arguably, the series of sugar-modified oligonucleotide mimics to have had most success is the locked nucleic acid (LNA, also called as BNA), which were established independently by Wengel and Imanishi.81,82,85 The ribose ring of an LNA is “locked” by a methylene bridge connecting the 2′- O-atom and the 4′-C atom, therefore typical 3′-endo locked conformation of the sugar puckering is created. Oligonucleotides containing LNAs are preor- ganized in the A-type canonical structure,163 thus bind to complementary RNA strand with higher affinity relative to native oligomers (plus 3-8 °C per modification). Another type of important sugar modification worth mentioning is hexitol nucleic acid (HNA), in which the pentafuranose sugar is replaced by 1,5- anhydrohexitol.164,165 It draws attentions because this pyranose nucleoside upon incorporation into oligonucleotides can stabilize the duplexes with the target DNA or RNA. The plausible reason is that firstly, the six-membered pyranose ring adopts a rigid chair conformation, requiring less entropy change during the duplex forming; secondly, the larger interstrand phosphate distance in HNA leads to less interstrand charge repulsion compared to the native duplexes.166 Both LNA and HNA belong to the family of conformationally constrained nucleosides. Other important conformationally locked members of this family will be described in the next chapter.

1.4.3.3 Base modifications Different base modifications (Figure 4C) in oligonucleotides167,168 have been used to discover potent antisense compounds for therapeutics. The modifications at the nucleobase that have positive influence on stacking interactions, hydrogen bonding and phosphate electrostatic repulsion can normally enhance the thermodynamic stability of nucleic acid duplexes. It was reported that unsaturated 5-position substitution of a pyrimidine nucleoside such as 5- propynyl-pyrimidines169,170 as well as some size-expanded nucleobase171-174 influenced stacking interactions in a beneficial way. Another method to obtain duplex stabilization is to decrease the electrostatic repulsion between phosphates of duplexes. Thus, a positively charged functional group (normally aminoalkyl group) is introduced, and positioned

21 in the major or minor groove that is able to interact with adjacent phosphate groups, resulting in net charge reduction.168 Besides these, modifications at the base-pairing site can also be beneficial for duplex stability when the number of hydrogen bonds between complementary bases is increased. 2,6-Diaminopurine175-177 when introduced into oligomers stabilizes the duplexes formed with both DNA and RNA because of the stronger base pairing without disturbing the local or global conformations of duplexes.

1.5 Overview of the thesis In this thesis we present the chemical synthesis of conformationally 2′, 4′- locked carba-LNA analogues, as well as their physicochemical and biochemical evaluations as antisense drug candidates upon incorporation into antisense oligonucleosides (AONs). The potential of AONs modified with carba-LNAs for antisense-based therapeutic application has been expected in terms of their high RNA binding affinity, improved nuclease resistance and RNase H elicitation capacity. The present work consists of two parts. The first part describes the chemical synthesis of 7′-Me-carba-LNAs and 6′-OH- 7′-Me-α-L-carba-LNAs through a 5-hexenyl type free-radical ring closure reaction. NMR characterization of key cyclization compounds for bicyclic ring formation and stereochemistry of different substitutions have also been discussed in this part. In the second part, the thermodynamic study of carba- LNA modified AONs toward complementary DNAs and RNAs is presented. Besides this, the AON sequences with modifications at different positions were subjected to 3′-exonuclease (SVPDE), human blood serum, E. coli RNase H1 and HeLa cell cytosolic extract to evaluate their antisense properties. The findings and implications from these enzymatic studies are discussed in this part.

22 2. Conformationally constrained nucleosides: introduction, synthesis and structural characterization

2.1 Brief introduction of conformationally constrained nucleos(t)ides Conventional ribo- and deoxyribonucleos(t)ides equilibrate rapidly in solution between two major conformers: the North (N) and the South (S) types. However, when a nucleoside or nucleotide binds to an enzyme or its phar- macological target, only one conformer is present in the active site. In order to query conformational preferences of enzymes involved in metabolism of nucleos(t)ides or to study the interactions of such compounds with their target enzymes, conformation-activity study is normally carried out. For these reasons, conformationally constrained nucleosides178 have attracted consid- erable interests since they are locked in preferred conformations, which can be a useful tool in such kind of evaluations. In 1994, Altmann and Marquez showed that 4′, 5′-methanocarba nucleoside (N-type sugar pucker) and 1′, 5′- methanocarba nucleoside (S-type sugar pucker) had totally different antiviral and antisense activities.179-181 However, the application of conformationally restricted nucleosides is not only limited to the research into bioactive compounds or biological tools, but also involved in RNA or DNA targeting therapeutics such as antisense, antigene and siRNA approaches. In late 1990s, antisense oligonucleotides (AONs) containing locked nucleic acids (LNAs or called as BNAs) showed unprecedented RNA target affinities as well as improved nuclease stabilities.82,83,85 These exciting discoveries have inspired an upsurge in synthesis of sugar moiety locked nucleos(t)ides. In the past twenty years, many sugar locked nucleosides including 1′,2′- locked,182-184 3′,5′-locked,185-188 2′,4′-locked,82,83,85,86,189-214 3′,4′-locked,215-218 2′,3′-locked,219-224 locked hexose nucleosides225 and methanocarba nucleo- sides225-228 have been synthesized, and some of them have been found to have good antisense and/or RNAi properties. The chemical structures of some representatives of sugar-locked nucleosides are shown in Figure 5.

23 (I) Methanocarba Nucleosides (II) 1',2'-Locked Nucleosides

B B B B HO HO O 5' HO HO T 6' 5' O 1' 4' 2' 4' 1' 3' OH 3' 2' OH R OH R O X 1 2 3 4 5 X = O or NH N-type R = H or OH R = H or OH ref. 227 ref. 228 N-type S-type North-East type ref. 225 ref. 181, 226 ref. 182-184

(III) 3',5'-Locked Nucleosides (IV) Locked Hexose Nucleosides OH T T O B O B O N O O O O HO O T OH O O O 8 9 6 7 OH 10 Tc-DNA E-type S-type Bc-DNA ref. 186 ref. 225 ref. 188 ref. 187 ref. 185

(V) 2',4'-Locked Nucleosides B B B B B B O O O O O O O O O O O O

N R O O O O O O O N O O O NH R R 16 12 13 14 15 11 NC LNA 6'-substituted LNA 2'-amino-LNA ENA aza-ENA 2',4'-BNA ref. 82, 83, 85 ref. 193 ref. 196, 199 ref. 192, 205-207 ref. 208 ref. 197 B B O B B B B O O O O O O O O O O O

R R O O O O 1 O 2 O O CH O O O 22 2 17 18 19 20 21 methylene-cLNA PrNA 2',4'-BNACOC parent cLNA 6',7'-substituted cLNA beta-bicyclonucleoside ref. 203 ref. 194, 200, 201, 203 ref. 198 ref. 202 ref. 205 ref. 195 R1 H C B B B B H B R B 2 O O 2 O O O O O O N O

O O O 28 O R1 O O O O O O 27 R2 25 26 23 24 6'7'-substituted methylene-alpha alpha-L-LNA alpha-L-2'-amino-LNA parent cENA 6',8'-substituted cENA -L-cLNA ref. 213 ref. 210 alpha-L-cLNA ref. 189 ref. 190, 191, 194, 200, 214 ref. 211 ref. 212 A O (VI) 3',4'-Locked Nucleosides T T O O B O T T O O O O O O O O O O O 29 O O Me 5'-cLNA-A Me 34 ref. 209 32 33 O 30 O O OR 31 W-type ref. 215 ref. 215 S-type S-type ref. 216 ref. 217 ref. 218 (VII) 2',3'-Locked Nucleosides T T HO T HO HO T T HO T HO HO O T O O HO O O O O O O O O O Si X X Si Ph Si 39 35: X = O 37: X = O O 40 41 42 43 36: X = N 38: X = N ref. 221 ref. 220 ref. 224 ref. 222 S-type N-type ref. 222 ref. 221 ref. 221

Figure 5. Representatives of conformationally constrained nucleos(t)ides. Note: the variations of structures with same entry number can be found in the original references.

24 2.1.1 Recent advances of intramolecular free-radical cyclization reactions on pentose sugars Of all the free-radical reactions that have been used for the construction of carbon-carbon bonds,229 the hex-5-enyl radical cyclization is the most well- known. The development of this reaction was first reported by Lamb et al230 in early 1960s. After that, synthetic organic chemists realized the power of this method in the assembly of complicated polyfunctional molecules, and hence utilized it into varieties of natural product synthesis.231 In the early 1990s, Chattopadhyaya and co-workers in our lab first reported the synthesis of 2′,3′-fused bicyclic nucleosides 35-39 (see Figure 6) through 5-exo type free-radical cyclization using the different radicals generated.221 Subse- quently, different kinds of functionalized bicyclic nucleosides 33, 34, 40-43 containing 3′,4′-β-fused six-membered ring, 2′,3′-fused five-membered ring as well as 2′,3′-cis and trans-fused seven-membered rings (Figure 5-VII) were synthesized through 6-exo, 5-exo and 7-endo type free-radical ring closure by Chattopadhyaya et al. 215,220,222,224 Inspired by the stereo and regiospecificity of the above free-radical cyclization on the pentose ring to construct bicyclic nucleosides as well as unprecedented antisense properties of LNA incorporated oligonucleotides developed by Imanish and Wengel, Chattopadhyaya realized that if the ring closure took place between the tethered olefin on C4′ and the radical center on C2′, which will lead to a 2′,4′-sugar locked bicyclic nucleoside having the same bicyclic scaffords as the LNA and ENA molecules. This idea prompted the synthesis of carba-LNA and carba-ENA derivatives by intramolecular free-radical cyclization on pentose sugar in our group.232 We believe that these new series of conformationally constrained nucleosides could create the flexibility of steering desired biophysical and biological properties for RNA targeting therapeutics. The first carba-ENA and its analogues were synthesized by Nielson’s group using ring closure metathesis strategy.189 AONs containing these cENA modifications led to increased thermal stabilities (Tm values) by 2.5- 4.5°C/modification compared to the native counterpart. Soon after, Chat- topadhyaya et al reported the synthesis of 7′-Me-carba-LNA-T and 8′-Me- carba-ENA-T, respectively, through 5-hexenyl and 6-heptenyl free-radical cyclization.200 It was demonstrated that 7′-Me-cLNA-T and 8′-Me-cLNA-T enhance the Tm of the modified AON/RNA heteroduplexes by 3.5-5 and 1.5 °C/modification, respectively. AONs containing cLNA and cENA modifications also showed remarkably enhanced lifetime in human blood serum, which may potentially produce the highly desired pharmacokinetic properties due to the unique stability they have. Later on, Chattopadhyaya and co- workers further investigated the effects of modifying the electrostatics of the bridging group in variants of 6′,7′-substituted cLNAs and 6′,8′-substituted cENAs. Effects of modifications on target affinity, nuclease resistance and

25 RNase H elicitation suggest that steric substitution on the edge of the minor groove improves the antisense properties of both cLNA and cENA.191,194

2.2 Synthesis and structural elucidation of 7'Me-cLNA- 5Me A, -G, C, and T as well as (6'OH,7'Me)-α-L-carba- LNA-T nucleosides (Paper I-III)

2.2.1 Synthesis of diastereomerically pure (7'S- or R-Me)-cLNA- A, -G, -5MeC, and -T nucleosides and their phosphoramidites Recently, we have reported the synthesis of 7′-Me-carba-LNA thymine (cLNA-T) and 8′-Me-carba-ENA thymine (cENA-T) through 5-exo type free-radical cyclization, and incorporated them into antisense oligonucleotides (AONs) for their physicochemical and biochemical studies.200 These molecules showed excellent biological properties, and thus prompted us to synthesize the related carba-LNA-A, -G, MeC analogues to investigate their antisense properties. In present work, diastereomerically pure 7′S and R methyl-cLNA-A, -G, -MeC and -T nucleosides and their phosphoramidites were synthesized, and the structural integrity and stereochemistry of these molecules were further proved by 1D and 2D NMR experiments. Detailed synthetic procedures and discussions are described in the following sections. The bicyclic systems of 7′-Me-carba-LNA-A, -G, -MeC and -T nucleosides was achieved by a key 5-exo type free-radical cyclization reaction using compounds 1a-d as the radical precursors (synthesis of 1a-d was described in ref. 202 and Paper I). Thus, the free-radical cyclization of compounds 1a, 1b, 1c and 1d using Bu3SnH and AIBN in refluxed anhydrous toluene led to two different diastereomeric isomers, i.e., 5-exo-cyclization products 7′R-Me-cLNAs 2-5a (38-50% yields) and 7′S-Me-cLNAs 2-5b (4- 7% yields) as well as one 6-endo-cyclization product parent cENAs 2-5c (4- 5% yields) (Scheme 1). The separation of each pure isomer was done by preparative HPLC, and the NMR characterization and structural elucidation of compounds 2-5a/b and 2-5c will be discussed in section 2.2.2. Above all, free-radical ring closure of precursors 1a-d, with various base moieties (A, G, MeC and T) can create both exo- and endo-cyclization products, in an approximate ratio of 10: 1. The results are well consistent with the previous theoretical study of alkenyl radical ring closure by Beckwith and Schiesser et al, where they pointed out that cyclization of 5-hexenyl-1- radical is a kinetically controlled process and generally prefers the exo cyclization products (exo/endo > 98/2).233

26 Scheme 1

NHCOPh OCO-NPh2 5 ' B -B 5 ' B1-B4 BnO B1-B4 1 4 BnO BnO O N O O N N N Bu3SnH, AlBN 4 ' 1' 4 ' 1' 3 ' 3 ' N 2 ' 2 ' N N N NHCOMe Toluene, reflux 6' 6' CH3 7' 8' BnO O O 8' Ph OBn 7' OBn Bz DPC/Ac B1 (A ) B2 (G ) S 2a: B1 (Major : 7'R, 38%) 2c: B1, 4% 1a: B1 O 2b: B1 (Minor : 7'S, 4%) 3c: B2, 4% NHCOPh 1b: B2 Me 4c: B3, 5% Me NH 1c: B3 3a: B (Major : 7'R, 48%) N 2 5c: B4, 5% 1d: B4 3b: B (Minor : 7'S, 4%) N 2 N O O 4a: B3 (Major : 7'R, 47%) 4b: B3 (Minor : 7'S, 7%) Me Bz B (T) B3 ( C ) 4

5a: B4 (Major : 7'R, 50%) 5b: B4 (Minor : 7'S, 6%) Scheme 1. Free-radical ring closure to 7′-Me-cLNAs and parent cENAs. Eight diastereomerically pure 7′S/R-Me-cLNA-A, -G, -MeC and -T nucleosides (2a/b, 3a/b, 4a/b, 5a/b) were isolated by preparative HPLC after free-radical cyclization. To convert all these intermediates to the corresponding phosphoramidites for solid-supported DNA synthesis, different synthetic strategies were employed.

Scheme 2

DMTrO T BnO T HO T O O (a) O (b)

CH3 CH3 CH3 OH OBn OH

6a (7'R) 7a (7'R), 80% in two steps 5a (7'R) 7b (7'S), 75% in two steps 5b (7'S) 6b (7'S)

Reagents and conditions: (a) 20% Pd(OH)2/C, ammonium formate, methanol, reflux, 8 h; (b) DMTrCl, dry pyridine, rt, overnight.

Scheme 2. Synthesis of 5′-O-DMTr-7′S or R-Me-cLNA-T nucleosides.

First, compounds 5a and 5b were debenzylated using 20% Pd(OH)2/C and ammonium formate, followed by selective protection of 5′-OH with DMTr chloride, giving 7a and 7b as precursors of phosphoramidites in 80% and 75% yields, respectively (Scheme 2).

Scheme 10

Me Bz Me C MeCBz BnO MeCBz BnO MeC HO C HO DMTrO O O O O O (d) (a) (b) (c) CH CH CH CH 3 CH3 3 3 3 OH OBn OBn OH OH

4a (7'R) 8a (7'R) 9a (7'R) 10a (7'R), 64% in three steps 11a (7'R), 75% 4b (7'S) 8b (7'S) 9b (7'S) 10b (7'S), 59% in three steps 11b (7'S), 86%

Reagents and conditions: (a) methanolic ammonia, rt, 16 h; (b) 20% Pd(OH)2/C, ammonium formate, methanol, reflux, 8 h; (c) Bz2O, dry pyridine, rt, overnight; (d) DMTrCl, dry pyridine, rt, overnight.

Scheme 3. Synthesis of 5′-O-DMTr-7′S or R-Me-cLNA-MeC nucleosides.

27 To deprotect N4-benzoyl group of cytosine, 4a and 4b were treated with methanolic ammonia at rt overnight to furnish compounds 8a and 8b, which were subjected to 20% Pd(OH)2/C and ammonium formate debenzylation to give 9a and 9b, followed by selective protection of amino group with benzoic anhydride to afford 10a and 10b in 64% and 59% yields, respectively. Thus, selectively protecting 5′-OH of 10a and 10b with DMTr chloride gives 11a and 11b in 75% and 86% yields (Scheme 3).

Scheme 4

HO HO ABz DMTrO ABz BnO Bz BnO A O A O O O A O (a) (b) (c) (d) CH CH CH CH3 52% in 3 80% 3 CH3 3 OH OH OH OBn OBn three steps 14a (7'R) 15a (7'R) 2a (7'R) 12a (7'R) 13a (7'R) 2b (7'S) 12b (7'S) 13b (7'S) Bz HO Bz DMTrO ABz DMTrO A (c) O A O O (d) (e) 50% in CH3 three steps CH3 81% CH3 85% OTMS OTMS OH 14b (7'S) 15b (7'S) 16b (7'S)

Reagents and conditions: (a) methanolic ammonia, rt, 16 h; (b) 20% Pd(OH)2/C, ammonium formate, methanol, reflux, 8 h; (c) i. TMSCl, dry pyridine; ii. BzCl, iii. aq NH3; (d) DMTrCl, dry pyridine, rt, overnight; (e) 1M TBAF, THF, rt, overnight. Scheme 4. Synthesis of 5′-O-DMTr-7′S or R-Me-cLNA-A nucleosides. Similarly, compounds 13a/b were formed from 2a/b using the same procedure as 4a/b to 9a/b. Subsequently, protection of N6-benzoyl group in 9a/b was achieved using Jone’s transient protection method.234 Therefore, compounds 9a/b was treated with TMS-Cl in dry pyridine followed by in situ drop-wise addition of benzoyl chloride, then treated with aqueous ammonia to give 14a and 14b in 52% and 50% yields respectively over three steps. It should be noted that TMS group of 3′-OH in 14b was not cleaved by aqueous ammonia, which is due to the steric hindrance effect caused by the neighbouring 7′S methyl group. Treatment of 14a and 14b with DMTr- Cl gives 15a and 15b in 80% and 81% yields. Compound 15b was then subjected to treatment with 1 M TBAF solution for cleavage of 3′-O-TMS group, giving compound 16b in 85% yield (Scheme 4).

Scheme 5

N2-Ac DMTrO N2-Ac BnO N2-Ac -O6-DPC BnO GN2-Ac HO G O G O G O O (a) (b) (c) CH CH CH3 CH3 3 3 OH OH OBn OBn 19a (7'R), 80% 17a (7'R), 60% 18a (7'R), 60% 3a (7'R) 18b (7'S), 80% 19b (7'S), 67% 3b (7'S) 17b (7'S), 83%

Reagents and conditions: (a) acetic acid, 55°C; (b) 20% Pd(OH)2/C, formic acid, methanol, reflux, 2.5 h; (c) DMTrCl, dry pyridine, rt, overnight. Scheme 5. Synthesis of 5′-O-DMTr-7′S or R-Me-cLNA-G nucleosides. The O6-diphenylcarbamoyl groups in 3a/b were cleaved by treatment with acetic acid under moderately warm conditions, followed by rt stirring, to

28 furnish 17a and 17b in good yield (60% for 17a, 83% for 17b). In the following step, the obtained compounds 17a/b were debenzylated in 235 Pd(OH)2/formic acid conditions instead of the classical Pd(OH)2/ ammonium formate conditions to avoid N2-acetyl group cleavage, giving 18a and 18b in 60% and 80% yields. Subsequently, selective protection of 5′-OH in 18a and 18b with DMTr-Cl in dry pyridine affords 19a and 19b in 80% and 67% yields (Scheme 5).

Scheme 6 DMTrO B O

DMTrO B O (a) CH3 O CH3 N P OH O CN

7a (7'R), B = T 7b (7'S), B = T 20a (7'R), B = T, 70% 11a (7'R), B = MeCBz 20b (7'S), B = T, 77% 11b (7'S), B = MeCBz 21a (7'R), B = MeCBz, 69% 15a (7'R), B = ABz 21b (7'S), B = MeCBz, 88% 16b (7'S), B = ABz 22a (7'R), B = ABz, 81% 19a (7'R), B = GN2-Ac 22b (7'S), B = ABz, 81% 19b (7'S), B = GN2-Ac 23a (7'R), B = GN2-Ac, 77% 23b (7'S), B = GN2-Ac, 56% Reagents and conditions: (a) 2-cyanoethyl-N, N- diisopropylphosphoramidochloridite, DIPEA, dry DCM, rt, 2 h.

Scheme 6. Synthesis of 7′S or R-Me-cLNA-A, -G, -MeC, -T phosphoramidites In the end, 5′-O-DMTr precursors 7a/b, 11a/b, 15a, 16b, 19a/b were subjected to phosphitylation using 2-cyanoethyl N, N-diisopropylphosphor- aminochloridite under standard conditions191,194,200,201,203,211 to furnish 20a/b, 21a/b, 22a/b, 23a/b as a diastereomeric mixture in 56-88% yields (Scheme 13). Thus, eight distereomerically pure 7′-Me-cLNA-A, -G, -MeC, -T phosphoramidites were obtained using the synthetic way described above in de- cent total yields.

2.2.2 Structural evidence of free-radical ring closure products in the synthesis of cLNA-A, -G, -MeC and -T nucleosides

2.2.2.1 Confirmation of bicyclic systems in 5-exo cyclization products The formation of C2′-C7′ bond in cLNA nucleosides 2a/b, 3a/b and 4a/b has been confirmed unequivocally: first, by the presence of correlation between H2′ and H7′ in COSY spectra (Paper I, Figure 5), and second, by the 3 observable JHC HMBC correlation between H1′ and C7′, H2′ with C6′ and 2 JHC HMBC correlation between H7′ with C2′ (Paper I, Figure 6). These proton-proton connectivities in the COSY and proton-carbon connectivities in HMBC corroborated that the oxa-bicyclo[2.2.1]heptane ring systems in

29 cLNA-A, -G and -MeC nucleosides have indeed been formed during the radical cyclization. For 6-endo cyclization product 2c, the ring closure by formation of C2′-C8′ bond has been unequivocally confirmed by the following NMR observations: 3 (1) JHH correlation between H8′ and H2′ in COSY spectrum (Paper I, Figure 3 8A); (2) JHC HMBC correlations between H1′ and C8′ (Paper I, Figure 8C); (3) DEPT (Paper I.SI, Figure SII.98) and HMQC (Paper I, Figure 8B) spectra showed both C7′ and C8′ are secondary carbons, each of them having two protons attached. Similar NMR experiments have also been carried out to prove that the oxa-bicyclo[3.2.1]octane ring system has been formed for compounds 3c (Paper I, Figure 8D-F) and 4c (Paper I, Figure 8G-J).

2.2.2.2 Stereochemistry of cLNA nucleosides The configuration of C1′ and C7′ centers in cLNA nucleosides 2a/b, 3a/b, 4a/b and 5a/b has been determined by 1D NOE experiments (see Paper I. SI Figure SII 65-66, 86-87, SIII 48-51, 68-69, SIV 72-73, 91-92 and Paper III.SI Figure SI 13-14, 27-28). For 2a, irradiation of H3′ led to 3 % of NOE enhancement for H8, suggesting that the 9-adeninyl moiety is in β configuration and in an anti conformation across the glycoside bond. The fact that the NOE enhancement of 2.9 % for H1′ upon irradiation on 7′-CH3 has been observed indicates that the methyl group on C7′ is in close proximity of H1′ (dH1′, 7′Me ≈ 2.1Å) in 2a, thereby confirming the R configuration at C7′ center (Figure 12). On the other hand, in the minor product 2b, irradiation of H1′ led to 5.5 % NOE enhancement for H7′ (dH1′, H7′ ≈ 2.1Å) but none for 7′-Me (dH1′, 7′Me ≈ 3.8Å). Hence, the stereochemistry at C7′ has been assigned to S configuration in compound 2b. In the similar way, the C7′ of the major cyclization products 3a, 4a and 5a have been assigned to R configuration, and the minor products 3b, 4b and 5b to C7′-S configuration (Figure 6).

30 H8 6.5% H8 1.9% H8 H8 3.1% 2.8% H3′ H3′ H3′ H3′ H1′ H1′ 4.7% H1′ H1′ H2′ H2 ′ H2′ H2′ 2.9% 7.3% 1.8% 1.9% 5.5% 6.3% 0.7% 7′-CH 7′-CH3 H7′ 3 H7′

3a (7′R) 3b (7′S) 2a (7′R) 2b (7′S)

H6 H6 4.6% 4.0% H6 4.0% H6 4.7% H3′

H3′ H3′ H1′ H3′ H1′ H1′ H1′ 5.3% H2′ 6.2% 2.1% 1.9% 4.3% 4.6% H7′ 7′-CH H2′ 3 H7′

7′-CH3

4a (7′R) 4b (7′S) 5a (7′R) 5b (7′S) Figure 6. NOE contacts to fix the stereochemistry in 7′S/R-Me-cLNA-A, -G, - MeC and -T nucleosides

2.2.3 Synthesis of diastereomerically pure (6'-OH,7'-Me)-α-L- carba-LNA-T nucleosides One of the most important conformationally constrained nucleosides α-L- LNA, a diastereomer of LNA, was developed by Wengel et al in recent 213,236-238 years. The AONs containing α-L-LNAs have shown comparable Tm values with that of LNA counterparts when targeted to the complementary RNAs. Moreover, α-L-LNA modified AONs were found more nucleolyti- cally stable than LNA modified counterpart.213 In vivo and in vitro experiments also showed that α-L-LNA modified AONs can knockdown the specific genes with higher efficiency than that of LNA modified counterpart.239,240 These favorable physicochemical and biochemical features of α-L- LNA inspired us to synthesize α-L-carba-LNA analogues, in which a methylene group replaces the 2′-oxygen of α-L-LNA, yielding a 2′,4′- carbocyclic locked ring. In present investigation, we succeeded in the synthesis of 6′,7′-substituted α-L-carba-LNA-T nucleosides and their phosphoramidites through a key 5-exo free-radical ring closure reaction which is used extensively for the construction of new carbocyclic rings. The synthetic route and structural elucidation of these novel molecules are presented as follows. The synthetic route to 6′,7′-substituted α-L-carba-LNA-T nucleosides and their phosphoramidites 34a-c is shown in Scheme 7-9. The synthesis started from the known compound 24,241 which was subjected to six steps to furnish the free-radical precursors 25a and 25b in 21% and 18% total yield (see Scheme 1 in Paper II).

31 Scheme 7 Bu3SnH, AlBN, CH HO 7' 8' HO 6'R 7'S 3 Thy dry toluene, reflux, 6h Thy 6'R O 1' O 4' 4' 1' 3' 2' 57% BnO 5' S BnO 3' 2' OBn O 5' HO 6 OBn O 1 O 4 α 26a (α-L) O 6 steps 25a ( -L) BnO 3 2 5 O H3C OBn H 5S O (β 8' 4 24 -L) 7' Bu SnH, AlBN, HO 6'S 6S HO 7'S CH HO 3 7'R 8' 3 Thy dry toluene, NH 6'S 6'S O O N Thy 1' reflux, 6h 2 + O 4' 4' 1' 4' 1' 3' 2' O BnO 5' S BnO 3' 2' BnO 3' 2' OBn O 5' OBn 5'OBn O 25b (α-L) 26c (α-L): 14% 26b (α-L): 43% Scheme 7. Free-radical ring closure reactions to 26a, 26b and 26c. The free-radical cyclization was carried out in refluxing anhydrous toluene with Bu3SnH, using AIBN as the initiator. Cyclization of 25a (6′-R) took place with high stereoselectivity to give α-L-carba-LNA nucleoside 26a (6′R-OH, 7′S-CH3) in good yield (57%, Scheme 7). Under an identical con- dition, cyclization of 25b (6′-S), on the other hand, gave α-L-carba-LNA nucleoside 26b (6′S-OH, 7′S-CH3) in 43% yield along with an unexpected product (6,7'-methylene-bridged)-α-L-carba-LNA-T 26c in a 14% yield (Scheme 7). The mechanism of the free-radical cyclization reaction for the formation of 26a, 26b, and 26c will be discussed in section 2.2.5. In an effort to remove the 6′S-OH in compound 26b by free-radical deoxygenation strategy, 26b was treated with CS2 and MeI along with NaH as a base, giving the precursor 27 in 64% yield. Subsequently, compound 27 was subjected to the standard Barton-McCombie deoxygenation242 in the presence of Bu3SnH and AIBN to furnish 7′S-CH3-α-L-carba-LNA nucleoside 28 in 42% yield as well as an unexpected rearrangement product bicyclo[2.2.1]-2′,6′-methylene-bridged hexopyranosyl nucleoside 29 in 7% yield (Scheme 8).

Scheme 8 CH3 7' S 6' 7'R H3C Bu SnH, AlBN, MeS 3 O Thy O CH3 dry toluene, NaH,CS , 6' 7' 4' 1' 6' O 2 Thy reflux, 2h + 4'R MeI, 0°C, 4h BnO 2'3' Thy O 5' 2' 26b 4' OBn BnO 3'R 64% 1' 5' 1' BnO 3' 2' 28 (α-L): 42% OBn 5'OBn 29 (α-L): 7% 27 (α-L) Scheme 8. Synthesis of 28 and 29 by free-radical deoxygenation and rearrangement. In order to convert 26a/b to the corresponding phosphoramidites for solid-supported DNA synthesis, the 6′-OH group of 26a and 26b must be protected (Scheme 9). Therefore, compounds 26a and 26b were treated with p-toluoylchloride to give 30a and 30b in 73% and 71% yield, respectively. After that, compounds 30a, 30b and 28 were subjected to debenzylation using 20% Pd(OH)2/C and ammonium formate followed by selectively protecting 5′-OH with DMTr group, giving 31a, 31b and 31c, respectively

32 (Scheme 9). For the purpose of inverting the configuration of 3′-OH, compounds 31a-c were oxidized with Dess-Martin periodinane243,244 followed by the reduction with sodium borohydride in ethanol to give 33a-c in 57-71% yield. Phosphi-tylation of 33a-c with 2-cyanoethyl-N,N-diisopropyl phos- phoramidochloridite gave phosphoramidites 34a-c, respectively, as a diastereomeric mixtures in 63-71% yield (Scheme 9).

Scheme 9 i) 20% Pd(OH)2/C, HCOONH4, MeOH, HO R reflux R 7'S CH3 4-toluoyl chloride, 6' 7' CH3 6' 7' CH3 6' ii) DMTrCl, dry py. Dess-Martin O Thy dry pyridine O Thy overnight O Thy periodinane 4' 1' CH2Cl2, rt, 3h 3' 2' DMTrO BnO 5' BnO OBn OBn OH 26a (α-L): 6'R 30a (α-L): R = OTol, 6'R, 73% 31a (α-L): R = OTol, 6'R, 75% 26b (α-L): 6'S 30b (α-L): R = OTol, 6'S, 71% 31b (α-L): R = OTol, 6'S, 59% 28 (α-L): R = H 31c (α-L): R = H, 70%

R 6' CH3 R 6' 7' CH3 7' O Thy O Thy O R 6' 7' 2-cyanoethyl N,N-diisopropyl NaBH4, EtOH CH3 DMTrO 3' phosphoramindochloridite, DMTrO rt, 1h O Thy P CN O OH DIPEA, CH2Cl2, rt, 3 h N O Intermediates DMTrO 3' 32a (α-L): R = OTol, 6'R 32b (α-L): R = OTol, 6'S 33a (α-L-ribo): R = OTol, 6'R, 57% in two steps 34a (α-L-ribo): R = OTol, 6'R, 71% 32c (α-L): R = H. 33b (α-L-ribo): R = OTol, 6'S, 66% in two steps 34b (α-L-ribo): R = OTol, 6'S, 63% 33c (α-L-ribo): R = H, 71% in two steps 34c (α-L-ribo): R = H, 65% Scheme 9. Synthesis of 6′,7′-substituted α-L-carba-LNA-T phosphoramidites 34a-c. The unexpected product 26c obtained during free-radical cyclization of 25b was also transformed to corresponding phosphoramidite using a similar strategy as shown in Scheme 9. The detailed synthetic procedures and yields can be found in Paper II.

2.2.4 NMR characterization of key intermediates involved in the synthesis of α-L-carba-LNA analogues

2.2.4.1 Confirmation of bicyclic and tetracyclic systems by HMBC and COSY experiments The formation of bicyclic systems in compounds 26a, 26b, 29 and the tetracyclic system in compound 26c was confirmed by HMBC and COSY experiments. 3 For compounds 26a and 26b, the observed JHC HMBC correlation between H2′ and carbon of 7′-methyl group, as well as correlation between H2′ and C6′ (Paper II, Figure SI.28) suggested that the fused carbocyclic ring between C4′ and C2′ has indeed been formed in compound 26a and 26b. 3 For compound 26c, the JHH correlation between H2′ and H7′ together 3 with the observation of JHC HMBC correlation of H2′ with C6′ (Paper II, Figure SI.144) gave solid evidence for the cis-fused nature of the 2′,4′- 3 carbocyclic ring system. The observed JHH correlation between H8′ and H7′ 3 3 3 as well as H8′′ and H7′ ( J8′, 7′ = 3.5 Hz, J8′′, 7′ = 4.0 Hz), JHH correlations 3 between H8′ and H6 as well as H8′′ and H6 of the thymine ( J8′, 6 = 4.0 Hz, 3 2 J8′′, 6 = 12.0 Hz), as well as JHC HMBC correlations between H8′, H8′′ and

33 C7′ along with C6 (Paper II, Figure SI.144 and 145) strongly verified that the 6,7′-methylene bridge was actually formed in compound 26c. Further- 3 3 more, the JHH correlation between H6 and H5 of thymine moiety ( J5, 6 = 3 3 10.5 Hz), as well as JHH correlation between H5 and 5-methyl ( J5, 5-CH3 = 7.0 Hz) were also observed in 2D COSY and 1D proton decoupling spectra, suggesting the aromatic property of thymine base was perturbed during the formation of methylene-bridge between C6 and C7′. The unusual product 29 has the same mass as compound 28, both of which were formed from free-radical deoxygenation of compound 27 (Scheme 8). Two NMR observations support that compound 29 was obtained by scission of C4′-O4′ bond and formation of a new O4′-C6′ bond: (i) 3 For compound 29, the JHC HMBC correlation between H1′ and C6′ (Paper II, Figure SI.177) was observed; (ii) The DEPT135 experiments showed that for compound 29, C4′ and C6′ are tertiary carbons (Paper II, Figure SI.180), but for compound 28, C6′ is a secondary carbon and C4′ existes as a quarter- nary carbon (Paper II, Figure SI.179). These observations unequivocally suggested that free-radical deoxygenation of compound 27 actually led to the formation of a rearranged product 29 with hexopyranosyl sugar moiety (putative mechanism for this rearrangement will be discussed in section 2.2.5) along with a deoxygenated compound 28 with furanosyl sugar moiety.

2.2.4.2 Stereochemistry of key intermediates in the synthesis of α-L- carba-LNA analogues The orientation of substituents in the carbocyclic moiety of compound 26a and 26b was determined by 1D NOE experiments (Figure 8) as well as from the vicinal coupling constants evaluation. For compound 26a, irradiation of H6 of thymine led to NOE enhancement for H7′ (2.6%) (dH6-H7′ ≈ 2.5 Å) and for 6′-OH (0.9%) (dH6-6′-OH ≈ 3.2 Å), but none for 7′-CH3 and H6′ (dH6-7′-CH3 ≈ 4.2 Å, dH6-H6′ ≈ 4.6 Å), whereas irradiation of 7′-CH3 led to NOE enhancement for H6′ (1.7%) (dH6′-7′-CH3 ≈ 2.5 Å) and H3′ (1.7%) (dH3′-7′-CH3 ≈ 2.4 Å), but none for 6′-OH (d7′-CH3-6′-OH ≈ 3.9 Å), strongly suggesting that C6′ is in 6′R configuration, and C7′ is in 7′S configuration. In addition, the trans disposition of H6′ and H7′ was also in agreement with the small coupling con- 3 stant for J6′, 7′ (3.6 Hz, hence dihedral angle H6′-C6′-C7′-H7′ ≈ 232° accord- ing to Karplus equation). As for compound 26b, irradiation of H6 of thymine group led to NOE enhancement for H7′ (1.4%) (dH6-H7′ ≈ 2.8 Å) and H6′ (2.7%) (dH6-H7′ ≈ 2.5 Å), but none for 7′-CH3 and 6′-OH (dH6-7′-CH3 ≈ 4.3 Å, dH6-6′-OH ≈ 4.4 Å), whereas irradiation of 7′-CH3 leads to NOE enhancement for H3′ (1.6%) (dH3′-7′-CH3 ≈ 2.4 Å), but none for H6′ (dH6′-7′-CH3 ≈ 3.8 Å), suggesting that C6′ is in 6′S configuration, and C7′ is in 7′S configuration. 3 Furthermore, the large coupling constant for J6′, 7′ (8.5 Hz) corresponding to the dihedral angle of H6′-C6′-C7′-H7′ ≈ 26° also suggested a cis disposition of H6′ and H7′ in 26b (Figure 7).

34 2.7%

0.9% 6'-OH H6' H6 H6

H7' 1.4% 2.6% H7' H6' 7'-Me 1.4% 1.7% 1.6% H2' 7'-Me H3'

H3' 1.7% 26b 26a

H6 4.9% H5

H7' 2.4% H6' H8' 5.2% H8'' 7'-Me H6'

1.2% H1' H7' 1.4% H4' 1.1% H2' 3.8% 1.2% H3' H5'

26c/35 29

Figure 7. Key NOE contacts of carbocyclic compound 26a, 26b, 26c/35 and 29; R = OH for 26c, R = H for 35 (see compound 19 in Paper II).

The stereochemistry of compound 29 can be also determined by 1D NOE experiment (Figure 8). It was found that selective irradiation of H4′ led to distinct NOE enhancement for H3′ (3.8%) (dH3′-H4′ ≈ 2.3 Å), and irradiation of 7′-CH3 led to NOE enhancement for H3′ (1.2%) (dH3′-7′-CH3 ≈ 2.4 Å) and H4′ (1.4%) (dH4′-7′-CH3 ≈ 2.4 Å) respectively, which suggested that H3′ and H4′ are cis oriented and they are on the same face as that of 7′-CH3. The observation that irradiation of H1′ led to NOE enhancement for H5′, 5′′ (1.1%) (dH1′-H5′,5′′ ≈ 2.8 Å) but none for H3′ (dH1′-H3′ ≈ 3.7 Å) and H4′ (dH1′-H4′ ≈ 4.2 Å) suggested that H5′, 5′′ are located on the face close to H1′. Hence, both C3′ and C4′ in compound 29 are in R configuration (Figure 7). For compound 26c, in which four new chiral centers have been formed during a single free-radical cyclization step, the configuration of every chiral center was also well determined by 1D NOE experiments (see Figure 7). Selective irradiation of H6 in thymine moiety led to strong NOE enhancement for H6′ (4.9%) (dH6-H6′ ≈ 2.3 Å) and H8′ (2.4%) (dH6-H8′ ≈ 2.4 Å), but none for 6′-OH and H7′ (dH6-6′OH ≈ 4.1 Å, dH6-H7′ ≈ 3.8 Å), unequivocally suggesting that C6′ is in 6S configuration, C7′ is in 7′S configuration and C6 is in 6S configuration respectively. The trans disposition of H6′ and H7′ was 3 also consistent with the coupling constant ( J6′, 7′ = 2.0 Hz, with dihedral angle H6′-C6′-C7′-H7′ ≈ 116°). In addition, selective irradiation of H8′′ of 35 (having the same carbon skeleton as 26c, see compound 19 in Paper II) led to strong NOE enhancement for H5 (5.2%) (dH5-H8′′ ≈ 2.3 Å) and for H2′

35 (1.2%) (dH2′-H8′′ ≈ 2.8 Å), but none for 5-CH3 group (d5-CH3-H8′′ ≈ 3.8 Å), suggesting that C5 is in 5S configuration.

2.2.5 Mechanism of the free-radical cyclization and radical rearrangement involved in the synthesis of α-L-carba-LNA analogues

After treatment of 25a with Bu3SnH and AIBN, the C2′ radical is supposed to be generated (see TS1 in Scheme 5A in Paper II), which should be capa- ble to attack the C=C double bond from both the “top” and “bottom” faces by the 5-exo cyclization pathway, resulting into two plausible intermediates TS2 and TS3 (Scheme 5A in Paper II). The optimized structures have shown • that in TS2 state, the thymine moiety, developing 7′-CH2 radical and 6′-OH are all occupying the axial positions. The steric hindrance between them makes TS2 much more unstable than TS3 because in TS3 state the thymine • moiety, developing 7′-CH2 radical and 6′-OH are occupying the axial, equa- torial and axial positions, respectively (Scheme 5A in Paper II). This comparison might explain why the exclusive formation of cyclic product 26a (6′R, 7′S) has been observed. Similarly, cyclization of 25b can proceed through intermediates TS5 and TS6 (see Scheme 5B in Paper II). In TS5, the cis orientation of 6′-OH (eq) • and 7′-CH2 radical (eq) is unfavored because of steric hindrance but the • orientation between thymine moiety (ax) and 6′-OH (eq) as well as 7′-CH2 radical (eq) are favored. On the other hand, in TS6, the trans orientation of • 6′-OH (eq) and 7′-CH2 radical (ax) is favored but 1,3-diaxial dispostion of • thymine (ax) and 7′-CH2 radical (ax) is unfavored. Taking together, cyclization of 25b through both TS5 and TS6 are possible to give product 26b and 26c with TS5 predominating since the products 26b and 26c were obtained in ratio 3:1. The formation of minor product 26c starting from TS6 can be easily un- • derstood as follows: First, in TS6, the primary -CH2 is not stable and will be intramolecularly trapped by the double bond of thymine moiety before quenched by Bu3SnH to give intermediate TS7. Then, the newly formed radical center at C5 of thymine moiety was reduced by Bu3SnH from less sterically hindered face, giving chiral C5 in S-configuration (Scheme 5B in Paper II). Hence, tetracyclic nucleoside 26c was obtained through a radical cyclization process. In our previous study, a radical deoxygenation of C6′-OH of β-D-carba- LNA led to an unusual 2′,6′-methylene bridged hexopyranosyl nucleoside which was obtained through a radical rearrangement of the generated 6′C• to 4′C• radicals.245 The formation of compound 29 during radical deoxygenation of the 6′-OH of α-L-carba-LNA nucleoside 27 was supposed to have gone through a similar mechanism. Thus after treatment of compound 27

36 • with Bu3SnH and AIBN in refluxing toluene, 6′C was putatively formed

(Scheme 10), which can be reduced by Bu3SnH directly to give product 28. On the other hand, the 6′C• could also lead to scission of the C4′-O4′ bond to give a new C6′=C4′ double bond. As a result, the 4′C• radical was transformed to 4′O• (TS9). Then the 4′O• radical attacked the C6′=C4′ again, resulting in formation of O4′―C6′ bond and the rearrangement of 4′O• radi- • cal to 4′C , which reacted with Bu3SnH by the less hindered face to furnish hexopyranosyl nucleoside 29.

Scheme 10

S H MeS O CH3 CH CH 7' 3 6' 7' 3 6' O Thy O Thy O Thy 4' 4' 1' 1' 2'3' BnO BnO 3' 2' BnO 5' OBn 5' OBn TS 8 OBn 28 : 42% 27

7' CH H3C 3 CH 6' H 6' 7' 3 O Thy 6' O 4' O Thy 4' BnO Thy 4'5' 1' 2' BnO 3' OBn BnO 3' 2' 5' 1' OBn OBn TS 9 TS 10 29 : 7%

Scheme 10. Mechanism of formation of 29 by free-radical rearrangement.

37 3. Antisense properties of modified AONs containing α-L-carba-LNAs and 7'-Me-carba- LNAs (Paper I-III)

3.1 Thermo-stabilities of chemically modified AONs toward the RNA and DNA targets The synthesized phosphoramidites of α-L-carba-LNAs and 7′-Me-cLNA-A, - G, -MeC and -T nucleosides were introduced into various sequences of antisense oligonucleotides using solid-supported DNA/RNA synthesizer (In present studies, 15 mer and 20 mer sequences were used). Measurement of thermal stabilities (Tm values) of the AON/RNA and AON/DNA duplexes was performed by means of temperature dependent UV spectroscopy. Hence, the thermostabilities of AONs containing different α-L-carba-LNA and carba-LNA modifications toward RNAs and DNAs are briefly discussed in the following sections.

3.1.1 Binding affinity of AONs containing α-L-carba-LNA and α-L-LNA thymines toward complementary RNA and DNA

The Tm values of duplexes formed by AON 2-21 with the complementary RNA or DNA were measured and compared with that of the native counterpart (Table 1). It was found as in previous reports213,236-238 that one α-L-LNA (type IV) modification resulted in roughly 4.4 °C increase in Tm for the AON/RNA hybrid. On the contrary, 7′R-Me-α-L-carba-LNA (type III), 6′S- OH-7′S-Me-α-L-carba-LNA (type I) and 6′R-OH-7′S-Me-α-L-carba-LNA (type II) led to Tm decrease by around 3 °C per modification. This observation indicates that 6′-OH in type I and II α-L-carba-LNAs exert no obvious effect on Tm for the AON/RNA hybrid regardless of their chiralities. Proba- bly, the drop in Tm caused by type I, II and III modifications could be on one hand due to the steric clash of the hydrophobic 7′R-methyl group. Addition- ally, this methyl group pointing toward vicinal 3′-phosphate (d(7′S-methyl)-3′P ≈ 4.1 Å) might impair the AON/RNA thermal stability by perturbing the hy- dration pattern246-251 and other stereoelectronic interactions in the major groove of the DNA/RNA duplex.

38 Table 1. Thermal denaturation of duplexes of native, α-L-carba-LNA derivatives and α-L-LNA modified AONs with complementary RNA or DNAa RNA With With selectiv- Modified LNA RNA DNA Entry AON-Sequence ity structures b b c ΔTm ΔTm ΔΔTm

AON1 Native 5'-d (CTT CAT TTT TTC TTC) Ref.* Ref.* -

AON2 HO CH3 5'-d (CTT CAT TTT TTC TTC) -2.8 -2.6 -0.2 6'S 7'S AON3 O T 5'-d (CTT CAT TTT TTC TTC) -3.0 -5.6 +2.6 AON4 O 5'-d (CTT CAT TTT TTC TTC) -2.2 -5.7 +3.5 O AON5 I 5'-d (CTT CAT TTT TTC TTC) -3.6 -4.1 +0.5

AON6 HO CH3 5'-d (CTT CAT TTT TTC TTC) -2.6 -2.9 +0.3 6'R 7'S 5'-d (CTT CAT TTT TTC TTC) AON7 O T -2.8 -4.5 +1.7 AON8 O 5'-d (CTT CAT TTT TTC TTC) -2.5 -7.0 +4.5 O AON9 II 5'-d (CTT CAT TTT TTC TTC) -3.7 -4.6 +0.9

AON10 CH3 5'-d (CTT CAT TTT TTC TTC) -3.0 -4.0 +1.0 AON11 7'R 5'-d (CTT CAT TTT TTC TTC) -3.0 -6.2 +3.2 O T AON12 O 5'-d (CTT CAT TTT TTC TTC) -2.3 -7.2 +4.9 AON13 O III 5'-d (CTT CAT TTT TTC TTC) -3.6 -5.0 +1.4

AON14 O 5'-d (CTT CAT TTT TTC TTC) +3.4 +1.3 +2.1 O T AON15 O 5'-d (CTT CAT TTT TTC TTC) +5.0 +1.8 +3.2 AON16 O 5'-d (CTT CAT TTT TTC TTC) +5.5 +0.9 +4.6 IV AON17 5'-d (CTT CAT TTT TTC TTC) +3.8 +2.2 +1.6

H3C AON18 5S 5'-d (CTT CAT TTT TTC TTC) -7.6 -6.9 -0.7 H O AON19 6S 5'-d (CTT CAT TTT TTC TTC) -12.6 -16.3 +3.7 7'S N NH AON20 O 5'-d (CTT CAT TTT TTC TTC) -13.8 -18.2 +4.4 O O O AON21 V 5'-d (CTT CAT TTT TTC TTC) -13.9 -16.2 +2.3 a A = native adeninyl, C = cytosinyl, T = thyminyl, ‘T’ indicates α-L-carba-LNA or α-L-LNA modified thymidime monomer with specified structure. * ‘Ref’ indicates reference duplex for Tm comparison. Tm values measured as the maximum of the first derivative of the melting curve (A260nm vs temperature) in medium salt buffer (60 mM Tris-HCl at pH 7.5, 60 mM KCl, 0.8 mM MgCl2) with temperature range from 20 to 65 ˚C using 1 μM concentrations of the two complementary strands. The value b of Tm given is the average of two or three independent measurements. ΔTm values were obtained by comparing the Tm values of AONs 2-21 with that of native AON 1.

39 c The RNA selectivity ΔΔTm was calculated by this equation: ΔΔTm = ΔTm (aver) of AON/RNA - ΔTm (aver) of AON/DNA.

Notably, the 6′,7′-substituted β-D-carba-LNA derivatives were found in 194,200,203 our previous studies lead to increase in Tm by 2-4°C depending on different substitutions at 6′ and 7′ positions. This should be compared to the observation made in the present study showing a Tm drop of 2-3 °C for 6′, 7′- substituted α-L-carba-LNA. This significant difference in thermal stabilities of β-D-carba-LNA vis-à-vis α-L-carba-LNA modified duplexes hints that the substitutions on the carbocyclic ring of α-L-carba-LNA located in major groove of DNA/RNA duplex have significantly destabilized the duplex, while the modifications of the carbocyclic ring of β-D-carba-LNAs located in the minor groove lead to stabilization of the duplexes. It was also found that all the α-L-LNA and α-L-carba-LNA derivatives are RNA selective since ΔΔTm (ΔΔTm = ΔTm (aver) of AON/RNA - ΔTm (aver) of AON/DNA) values were found in the range of 1.6-2.8°C (Table 1). Incorporation of the hyper-constrained 6,7′-methylene bridged-α-L-carba- LNA thymidine (type V) into 15mer oligonucleotides led to dramatic decrease in thermal affinity toward both complementary RNA and DNA (Tm dropped 7 - 14 °C with RNA, and dropped 7-18 °C with DNA, see AON 18- 21 in Table 1). It was speculated that the loss of aromaticity in nucleobase resulted in the increased steric bulk of nucleobase moiety (from planar to tetrahedral geometry at C5 and C6 position), thereby destabilizing the duplex owing to perturbation of the base stacking of modified nucleic acid with neighboring base pair to some extent due to energetically unfavorable in- trastrand interaction. Alternatively, as a consequence of constrained glycosidic torsion angle of type V, the nucleobase participating in hydrogen bonding might not be disposed optimally for efficient Watson-Crick base pairing, therefore also impairing the duplexes stability to some extent.

3.1.2 Binding affinity and thermodynamic properties of 7'-Me- cLNA-A, -G, -MeC and -T modified AONs toward complementary RNA and DNA strands Eight diatereomerically pure 7′S- and R-Me-cLNA-A, -G, -MeC, and -T as well as LNA-A, -G, -C, -T were incorporated into a 20 mer AON sequence as a single modification at different positions. The thermal denaturation study of cLNA and LNA modified AONs duplexed with complementary RNA and DNA was carried out. Hence, the sequence of AONs and Tm values of these duplexes are listed in Table 2. From the Tm measurement, we found that AONs modified with cLNA-A, - G, -MeC and -T analogues exhibited much higher RNA affiniies (plus 1-4 °C/mod in Tms) than that of native DNA strand, which is also comparable

40 with that of LNA modified AON/RNA duplexes (Table 2). In addition, we also found that the Tm enhancement in AON/RNA duplexes modified with different cLNA and LNA analogues is highly position-dependent. It showed in Figure 8 that AONs containing 7-Me-cLNA-As at 3-position (counting from 3′-end, the same as follows) have the lowest Tm increase toward RNA (around 0.7°C), while AONs containing cLNAs with different base moieties Me (A, G, C and T) at position 4, 5, 6 and 7 have around 2°C Tm enhancement. On the other hand, AONs with single modifications at 8-, 9-, 10-, 11-, 13-, 15- and 16-positions showed a 3-4°C Tm enhancement toward RNA. For AONs with LNA modifications, the position-dependent Tm variations were also observed (Figure 8) in that AONs with LNA modifications at 3-, 4-, 5-, 6-, 7- and 9-positions have slightly lower Tm enhancement toward RNA (plus 1-3°C/mod) as compared to that of AONs with modifications at 8-, 10-, 11-, 13-, 15- and 16-positions (plus 3-5°C/mod). Upon incorporation into the 20 mer AONs, the S- and R-configured methyl group in cLNAs (minor vs major isomer) imposed a differential impact on Tm values toward complementary RNA (Figure 8). However, Tm variations caused by reverse stereochemical orientations of methyl substitution in cLNAs are position-dependent and marginal.

Table 2. Tm Values of Duplexes Formed by Modified AONs with Comple- mentary RNA and DNAa

ΔT AON-Sequence* T of T of m Modified LNA m m RNA Entry (Containing modifications in the AON/ AON/ structures selectiv- minor groove) RNA DNA ityb

Native 5'-d (TCC CGC CTG TGA CAT GCA TT) 73.3ºC 70.6ºC +2.7ºC

A AON22 O 5'-d (TCC CGC CTG TGA CAT GCAR TT) 74.0ºC 70.9ºC +3.1ºC O

AON23 5'-d (TCC CGC CTG TGA CART GCA TT) 75.0ºC 70.5ºC +4.5ºC 7'R

O CH3 AON24 5'-d (TCC CGC CTG TGAR CAT GCA TT) 76.1ºC 71.5ºC +4.6ºC Major isomer

A AON25 O 5'-d (TCC CGC CTG TGA CAT GCAS TT) 73.9ºC 70.6ºC +3.3ºC O

AON26 5'-d (TCC CGC CTG TGA CAST GCA TT) 74.7ºC 69.9ºC +4.8ºC 7'S

O CH3 AON27 5'-d (TCC CGC CTG TGAS CAT GCA TT) 75.9ºC 71.6ºC +4.3ºC Minor isomer

41 G AON28 O 5'-d (TCC CGC CTG TGA CAT GRCA TT) 74.5ºC 70.8ºC +3.7ºC O

AON29 5'-d (TCC CGC CTG TGRA CAT GCA TT) 76.8ºC 70.1ºC +6.7ºC 7'R

O CH3 AON30 5'-d (TCC CGRC CTG TGA CAT GCA TT) 76.9ºC 70.4ºC +6.5ºC M ajor isom er

G AON31 O 5'-d (TCC CGC CTG TGA CAT GSCA TT) 75.2ºC 70.9ºC +4.3ºC O

AON32 5'-d (TCC CGC CTG TGSA CAT GCA TT) 76.7ºC 70.4ºC +6.3ºC 7'S

O CH3 AON33 5'-d (TCC CGSC CTG TGA CAT GCA TT) 77.1ºC 70.6ºC +6.5ºC Minor isomer

Me C 5'-d (TCC CGC CTG TGA CAT GMeCRA AON34 O 75.5ºC 71.4ºC +4.1ºC O TT) 5'-d (TCC CGC CTG TGA MeCRAT GCA 76.7ºC 71.8ºC +4.9ºC AON35 TT) 7'R O CH 5'-d (TCC CGMeCR CTG TGA CAT GCA AON36 3 77.0ºC 71.8ºC 5.2ºC Major isomer TT)

MeC 5'-d (TCC CGC CTG TGA CAT GMeCSA AON37 O 75.8ºC 71.7ºC +4.1ºC O TT) 5'-d (TCC CGC CTG TGA MeCSAT GCA 76.7ºC 72.1ºC +4.6ºC AON38 TT) 7'S O Me S CH3 5'-d (TCC CG C CTG TGA CAT GCA AON39 76.7ºC 72.2ºC +4.5ºC Minor isomer TT) T AON40 O 5'-d (TCC CGC CTG TGA CATR GCA TT) 75.9ºC 70.5ºC +5.4ºC O

AON41 5'-d (TCC CGC CTG TRGA CAT GCA TT) 76.6ºC 70.8ºC +5.8ºC 7'R

O CH3 AON42 5'-d (TCC CGC CTRG TGA CAT GCA TT) 77.2ºC 71.9ºC +5.3ºC M ajor isom er

T AON43 O 5'-d (TCC CGC CTG TGA CATS GCA TT) 75.2ºC 70.2ºC +5.0ºC O

AON44 5'-d (TCC CGC CTG TSGA CAT GCA TT) 75.9ºC 70.7ºC +5.2ºC 7'S

O CH3 AON45 5'-d (TCC CGC CTSG TGA CAT GCA TT) 76.7ºC 70.1ºC +4.9ºC Minor isomer

42 A AON46 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 74.7ºC 71.6ºC +3.1ºC O

AON47 5'-d (TCC CGC CTG TGA CAT GCA TT) 75.6ºC 71.5ºC +4.1ºC

O AON48 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 75.9ºC 72.5ºC +3.4ºC

G AON49 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 75.9ºC 72.1ºC +3.8ºC O

AON50 5'-d (TCC CGC CTG TGA CAT GCA TT) 77.1ºC 71.4ºC +5.7ºC

O AON51 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 77.2ºC 71.4ºC +5.8ºC

C AON52 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 76.5ºC 73.6ºC +2.9ºC O

AON53 5'-d (TCC CGC CTG TGA CAT GCA TT) 77.7ºC 74.1ºC +3.6ºC

O AON54 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 77.7ºC 73.8ºC +3.9ºC

T AON55 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 76.6ºC 72.7ºC +3.9ºC O

AON56 5'-d (TCC CGC CTG TGA CAT GCA TT) 76.9ºC 72.6ºC +4.3ºC

O AON57 O 5'-d (TCC CGC CTG TGA CAT GCA TT) 77.9ºC 73.2ºC +4.7ºC

aMolecular weights of all antisense sequences are confirmed by MALDI-TOF mass spectrum (see Table.1 and Figure SII.13-48 in Supporting Information Part II of Paper III). A = adeninyl, G = guaninyl, C = cytosinyl, T = thyminyl. AR = 7'R-Me- cLNA-A, GR = 7'R-Me-cLNA-G, MeCR = 7'R-Me-cLNA-MeC, TR = 7'R-Me-cLNA-T; AS = 7'S-Me-cLNA-A, GS = 7'S-Me-cLNA-G, MeCS = 7'S-Me-cLNA-MeC, TS = 7'S-

Me-cLNA-T; A = LNA-A, G = LNA-G, C = LNA-C, T = LNA-T. Tm Values meas-

ured at the maximum of the first derivative of the melting curve (A260nm Vs temperature) in medium salt buffer (60 mM tris-HCl at pH 7.5, 60 mM KCl, 0.8 mM

MgCl2) with temperature 60°C – 90°C using 1μM concentrations of two comple-

mentary strands. The values of Tm given are averages of three independent measurements (the error of the three consecutive measurements is within + 0.2 °C). d RNA-selectivity: ΔTm = (Tm of AON/RNA) - (Tm of AON/DNA).

43 5.0 C LNA-T Me LNA-C LNA-C C C 4.5 C Me Me Me * -Me-cLNA-T R -Me-cLNA-G 7' S -Me-cLNA- 4.0 LNA-G R -Me-cLNA-G 7' LNA-G R 7' -Me-cLNA-G 7' -Me-cLNA- R LNA-T -Me-cLNA- -Me-cLNA-T -Me-cLNA- -Me-cLNA-G S S R S S 7' -Me-cLNA-T C 7' 7' 7' 7' R 7'

3.5 Me 7' LNA-T LNA-C C Me -Me-cLNA-A

3.0 R -Me-cLNA-T 7' -Me-cLNA-T -Me-cLNA-A S R S -Me-cLNA- 7' LNA-A 7' S 7' LNA-G 7'

2.5 -Me-cLNA- R LNA-A 7' -Me-cLNA-G -Me-cLNA-T S S 7' 7' 2.0 -Me-cLNA-A R 7' -Me-cLNA-A S -Me-cLNA-G 7' 1.5 LNA-A R 7' of AON-RNA duplex (°C) -Me-cLNA-A m 1.0 R -Me-cLNA-A

Τ S 7' 7' Δ 0.5

0.0 pos3 pos4 pos5 pos6 pos7 pos8 pos9 native pos10 pos11 pos13 pos15 pos16

Me Figure 8. Barplot of Tm variations of 7′-Me-cLNA-A, -G, - C, -T and LNA-A, -G, -C, -T modified AON/RNA duplexes compared with that of native counterpart. *ΔTm of AON/RNA was obtained by comparing the Tm of AON/RNA with that of the native AON /RNA.

We have elucidated the experimental thermodynamic parameters of different cLNA modified AON/RNA and AON/DNA duplexes based on the concentration dependent Tm measurement. Through comparison of -ΔG° values obtained, we found that nearly all the cLNA modified AON/RNAs are more RNA affinitive than the native counterpart (Figure 9). Additionally, we also found that both of enthalpy and entropy changes with strong com- pensation resulted in the free-energy changes (-ΔG°) of the duplexes, and therefore well explained the argument that the origin of greater duplex- stability of cLNA modified AONs compared to the native counterpart is either enthalpy or entropy driven, but not both at the same time.252 It shows in Table 2 that ΔΔTm (ΔΔTm = Tm of AON/RNA - Tm of AON/DNA) values of all cLNA modified AONs are from 3.1 to 6.7°C, much higher than that of native AON with ΔΔTm value of 2.7°C. On the other hand, we found that most of the ΔG°RNA selectivity values [ΔG°RNA selectivity = (-ΔG° of AON/RNA duplexes) - (-ΔG° of AON/DNA duplexes)] derived between cLNA modified AON/RNA and cLNA modified AON/DNA duplexes are much higher than that of the native DNA/RNA with native DNA/DNA duplex besides that the AONs 2, 6, 10, 14, 18 have similar or slightly lower ΔG°RNA selectivity values (Figure 10). Therefore, we can arrive at the conclusion that cLNA modified AONs are more RNA selective than the native counterpart not only in terms of Tm analysis but also thermodynamic properties.

44 C Me -Me-cLNA-

17.5 S 7' -Me-cLNA-T R C 7'

15.0 Me

12.5 -Me-cLNA-G S 7' -Me-cLNA-G -Me-cLNA- R R -Me-cLNA-G C 7' 7' 10.0 R Me C 7' C -Me-cLNA-T -Me-cLNA-A Me -Me-cLNA-A R Me kJ/mol) kJ/mol) -Me-cLNA-T R S 7' S ( 7' 7'

7.5 7' * ° C -Me-cLNA-G -Me-cLNA-T -Me-cLNA-T -Me-cLNA- S S R G Me R 7' 7' -Me-cLNA- 7'

5.0 7' -Me-cLNA- S R -Me-cLNA-T 7' S -Me-cLNA-A 7' ΔΔ -Me-cLNA-A -Me-cLNA-A 7'R-Me-cLNA-G R 7' S S 7' 7' 2.5 7' -Me-cLNA-A -Me-cLNA- S R 7' 7' 0.0 7'S-Me-cLNA-G

-2.5

-5.0 RNA affinity affinity RNA duplexes of AON-RNA -7.5

-10.0 pos3 pos4 pos5 pos6 pos7 pos8 pos9 native pos10 pos11 pos13 pos15 pos16 Figure 9. Barplot of free energy difference (RNA affinity) of cLNA modified AON/RNA duplexes compared with that of native duplex. *ΔΔG° = (-ΔG° of cLNA modified AON/RNA duplexes) - (-ΔG° of native DNA/RNA duplex). Therefore ΔΔG° of native DNA/RNA duplex is calculated to zero.

35 C Me C -Me-cLNA-G Me R 30 7' -Me-cLNA-T -Me-cLNA- -Me-cLNA-G R *(kJ/mol) S R 7' 7' 7' C -Me-cLNA- -Me-cLNA-T S R Me -Me-cLNA-G 7' 7' -Me-cLNA-A S C

25 -Me-cLNA-T R 7' -Me-cLNA-T S Me S 7' 7' 7' -Me-cLNA-T R -Me-cLNA- 7' S

20 7' -Me-cLNA- RNA selectivity R -Me-cLNA-A S 7' ° -Me-cLNA-G 7' R 7' G -Me-cLNA-G R Δ 15 7' -Me-cLNA-A S C 7' -Me-cLNA-T

10 Me S -Me-cLNA-A C R 7' Me 7' -Me-cLNA-G S 7' Native -Me-cLNA-A -Me-cLNA-A R S 7' -Me-cLNA-

5 7' S 7' -Me-cLNA- R 7' 0 RNA selectivity selectivity RNA pos3 pos4 pos5 pos6 pos7 pos8 pos9 native pos10 pos11 pos13 pos15 pos16

Figure 10. Barplot of RNA selectivity ΔG°RNA selectivity represented by cLNA modified AONs 22-45. *ΔG°RNA selectivity = (-ΔG° of AON/RNA duplexes) - (-ΔG° of AON/DNA duplexes)

45 3.2 Nuclease stabilities of modified AONs It is known that 3′-exonuclease activity is predominantly responsible for enzymatic degradation of AON in serum-containing medium or in various eukaryotic cell lines, and modifications located at 3′-terminus can significantly contribute to the nuclease resistance of an oligonucleotide.253,254 Thus, AONs containing different cLNA modifications near to the 3′-end were designed and treated with phosphodiesterase I from Crotalus adamant- eus venum (SVPDE) as well as human blood serum that contains mainly 3′- exonuclease. A comparison of digestion rates and implications from the studies of various chemically modified AONs toward SVPDE and blood serum incubations are briefly discussed in the following sections.

3.2.1 Nucleolytic stabilities of AONs modified with α-L-carba- LNA derivatives and α-L-LNA The stabilities of AONs 2, 6, 10, 14, 18 (see Figure 4 in Paper II) toward SVPDE incubation decreased in following order: 6,7′-methylene bridged α- L-carba-LNA-T (type V) modified AON 18 (k = 0.0040 ± 0.0012 min-1) > 7′R-methyl-α-L-carba-LNA (type III) modified AON 10 (k = 0.0068 ± 0.0009 min-1) ≈ 6′R-hydroxyl-7′S-methyl-α-L-carba-LNA (type II) modified AON 6 (k = 0.0076 ± 0.0018 min-1) > 6′S-hydroxyl-7′S-methyl-α-L-carba- LNA (type I) modified AON 2 (k = 0.0125 ± 0.0007 min-1) > α-L-LNA (type IV) modified AON 14 (k = 0.0777 ± 0.0072 min-1) > β-D-LNA (type VI) incorporated AON (k = 0.5331 ± 0.1800 min-1). Hence, all α-L-carba-LNA analogues modified AONs were found to be 3′-exonucleolytically more stable than parent α-L-LNA and β-D-LNA modified counterparts. We have also found that 7′R-methyl-α-L-carba-LNA (type III) modified AON 10 is about 10 times more stable than α-L-LNA (type IV) modified AON 14, which suggests that the replacement of the 2′-O- with hydrophobic methylmethylene function in α-L-LNA can render significantly positive effects on the nuclease resistance. C6′-OH substitution on the α-L-carba-LNA can modulate the stability of modified AONs to different extent depending on the stereochemical orientation of the hydroxyl group (Paper II). 6′R-hydroxyl substitution led to virtu- ally no effect since 6′R-hydroxyl-7′S-methyl-α-L-carba-LNA (type II) modified AON 6 have shown very similar overall stability with the 7′R-methyl-α- L-carba-LNA (type III) modified AON 10. On the other hand, 6′S-hydroxyl substitution [OH group is pointing at the vicinal 3′-phosphate] in 6′S- hydroxyl-7′S-methyl-α-L-carba-LNA (type I) upon incorporation into modified AON 2 led to significantly less 3′-exonucleolytically stable duplex than the duplex incorporating the 7′R-methyl-α-L-carba-LNA (type III) modification (AON 10). This finding is well in agreement with the previous nuclease studies of 6′R-OH-β-D-carba-LNAs, which upon incorporated into AONs

46 showed remarkably reduced 3′-exonucleolytic stability. The reason that is probably the C6′-OH points at the vicinal 3′-phosphate can assist in the departure of 3′-oxyanion during SVPDE mediated 3′O-P bond scission.255 Notably, 6,7′-methylene bridged α-L-carba-LNA-T (type V) modified AON 18 showed better 3′-exonucleolytic resistance than type I, II, III modified AONs. It can be concluded that the extra six-membered ring of 6,7′- methylene bridged α-L-carba-LNA-T (type V), which lies above the pentose sugar, gave slightly higher nuclease resistance compared to other α-L-carba- LNA derivatives (type I, II and III). Blood serum study of AONs 2, 6, 10, 14, 18 was also performed. Due to the presence of alkaline phosphatase in blood serum that gradually removes the 5′-end 32P label, it was impossible to obtain accurate degradation rate for each AON by quantifying the gel picture. Visual comparison of the gel pictures showed that the order of relative stabilities of AONs in human blood serum was similar to that observed upon treatment by 3′-exonucleases.

3.2.2 Nucleolytic stabilities of AONs containing 7'R- and S-Me- cLNA-A, -G, -MeC and -T nucleosides

By comparison of pseudofirst-order reaction rates and half-life times (t1/2) obtained, the stability of modified AONs containing 7′R- and S-Me-cLNA modifications (see Figure 11 & 12) under treatment with SVPDE decreased in the following order: 7′S-Me-cLNA-MeC modified AON 37 (k = 0.0027 ± -1 Me 0.0002 h , t1/2 = 258 ± 19.0 h) > 7′R-Me-cLNA- C modified AON 34 (k = -1 0.0059 ± 0.0006 h , t1/2 = 119 ± 13.0 h ) > 7′S-Me-cLNA-T modified AON -1 43 (k = 0.0985 ± 0.0046 h , t1/2 = 7.10 ± 0.40 h) > 7′R-Me-cLNA-T modified -1 AON 40 (k = 0.1107 ± 0.0084 h , t1/2 = 6.30 ± 0.50 h) > 7′S-Me-cLNA-A -1 modified AON 25 (k = 0.1650 ± 0.0055 h , t1/2 = 4.20 ± 0.10 h) > 7′R-Me- -1 cLNA-A modified AON 22 (k = 0.3954 ± 0.0448 h , t1/2 = 1.80 ± 0.20 h) > -1 7′S-Me-cLNA-G modified AON 31 (k = 0.5246 ± 0.0235 h , t1/2 = 1.35 ± -1 0.05 h) > 7′R-Me-cLNA-G modified AON 28 (k = 0.7354 ± 0.0300 h , t1/2 = 0.95 ± 0.04 h). Therefore, it is evident that all the 7′-Me-cLNAs modified AONs are much more stable toward SVPDE than the native counterpart, which was degraded completely within several minutes. It was found that 7′S-Me-cLNA-MeC modified AON 37 and 7′R-Me- cLNA-MeC modified AON 34 exhibited unprecedented 3′-exonucleolytic resistance compared to other cLNAs modified AONs. Remarkably, the most Me -1 resistant 7′S-Me-cLNA- C modified AON 37 (k = 0.0027 ± 0.0002 h , t1/2 = 258 ± 19.0 h) is around 35 and 41-fold more stable than 7′S-Me-cLNA-T -1 modified AON 43 (k = 0.0985 ± 0.0046 h , t1/2 = 7.10 ± 0.40 h) and 7′R-Me- -1 cLNA-T modified AON 40 (k = 0.1107 ± 0.0084 h , t1/2 = 6.30 ± 0.50 h), which have been reported to show greatly improved nuclease resistance ca- pability compared to LNA modified oligos.200 Likewise, 7′S-Me-cLNA-T

47 modified AON 43 is around 1.7 and 4-fold more stable than 7′S-Me-cLNA- A modified AON 25 and 7′R-Me-cLNA-A modified AON 22, and around 5.3 and 7.5-fold more stable than 7′S-Me-cLNA-G modified AON 31 and 7′R-Me-cLNA-G modified AON 28. These results suggest that the AONs containing cLNAs with 5MeC base moiety are much more resistant to 3′- exonuclease, relative to AONs having cLNAs with T, A and G base moieties. Furthermore, the orientation of C7′-methyl in cLNA-A, -G, -MeC and -T can influence the stability of modified AONs to some extent. By comparing degradation kinetics of 7′S-Me-cLNA-MeC modified AON 37 (k = 0.0027 ± -1 Me 0.0002 h , t1/2 = 258 ± 19.0 h) with 7′R-Me-cLNA- C modified AON 34 (k -1 = 0.0059 ± 0.0006 h , t1/2 = 119 ± 13.0 h), 7′S-Me-cLNA-T modified AON -1 43 (k = 0.0985 ± 0.0046 h , t1/2 = 7.10 ± 0.40 h) with 7′R-Me-cLNA-T modi- -1 fied AON 40 (k = 0.1107 ± 0.0084 h , t1/2 = 6.30 ± 0.50 h), 7′S-Me-cLNA-A -1 modified AON 25 (k = 0.1650 ± 0.0055 h , t1/2 = 4.20 ± 0.10 h) with 7′R- -1 Me-cLNA-A modified AON 22 (k = 0.3954 ± 0.0448 h , t1/2 = 1.80 ± 0.20 -1 h), 7′S-Me-cLNA-G modified AON 31 (k = 0.5246 ± 0.0235 h , t1/2 = 1.35 ± 0.05 h) with 7′R-Me-cLNA-G modified AON 28 (k = 0.7354 ± 0.0300 h-1, t1/2 = 0.95 ± 0.04 h), we can arrive at a conclusion that the 7′-methyl group, when it points towards the vicinal 3′-phosphate (C7′-S), has more positive effect on the enzymatic stability of the vicinal phosphate than when it points away from the phosphate (C7′-R). In addition, it should be noted that the extent of the effect by orientation of C7′ methyl group is much less evident compared to that caused by four various base moieties in cLNAs. This can be obviously proved by the degradation kinetics of modified AONs (Figure 11 & 12) that nucleolytic resistance of cLNA modified AONs is decreased in the following order: 7′S- and R-Me-cLNA-MeC > 7′S- and R-Me-cLNA-T > 7′S- and R-Me-cLNA-A > 7′S- and R-Me-cLNA-G, regardless of the relative stereochemical orientation of 7′-methyl subsitution.

48 100 AON37 (k = 0.0027 ± 0.0002/hr) 5' O B O 4' 1' 3' 80 AON34 (k = 0.0059 ± 0.0006/hr) 2'

6' CH3 8' O 7' 60 AON22: B = A, 7'R (major isomer) AON25: B = A, 7'S (minor isomer) AON28: B = G, 7'R (major isomer) 40 AON43 (k = 0.0985 ± 0.0046/hr) AON31: B = G, 7'S (minor isomer) AON34: B = MeC, 7'R (major isomer) AON40 (k = 0.1107 ± 0.0084/hr) Me %AON remaining %AON AON22 (k = 0.3954 ± 0.0448/hr) AON37: B = C, 7'S (minor isomer) 20 AON40: B = T, 7'R (major isomer) AON25 (k = 0.1650 ± 0.0055/hr) AON43: B = T, 7'S (minor isomer) AON31 (k = 0.5246 ± 0.0235/hr)

0 Native AON28 (k = 0.7354 ± 0.0300/hr)

0 5 10 15 20 25 Time (hr) Figure 11. SVPDE digestion curves of native AON and selected cLNA-A, -G, -MeC, -T modified AONs. The pseudofirst-order rates shown here were obtained by fitting the curve to the single-exponential decay function. Digestion conditions: AON 3 µM (5′-end 32P labeled with specific activity 80 000 cpm) in 100 mM Tris-HCl (pH 8.0) and 15 mM MgCl2, 21 °C, total reaction volume 30 µL, SVPDE concentration (6.7

ng/µL). )

r Me

o 7'S-Me-cLNA- C (minor)

(

e M - 5'-end

A O )

N 258 h r

) B

L o

c O P O

n 12

- i

j O 250 e

a O

)

(

200 - n ' T

i O

)

- A

r B

)

(

r or

119 h

150 N n O P O

L 13

aj i

A O

m c

m O

(

100 (

R or S

M - L

G O CH S

- 3

' A - B

10 R

7 L

N O P14 O

L M

c 7.1 h O

c L

8 -

- c

S O

e 6.3 h

M M

6 - -

M O

R 4.2 h -

S cleavage site

' '

(h) of SVPDE of digestion (h) B

7 7 4 '

7 O P15 O 1/2 1.8 h O t

1.35 h 2 0.95 h O

0 3'-end OH -G C A-A A N NA-T A-Me cL -cL - N e -cL 7-Me 7-Me-cLN 7-M 7-Me

Figure 12. Bar plot of half-life time (t1/2) of SVPDE digestion of selected 7′R- and S-Me-cLNA-A, -G, -MeC, -T modified AONs.

49 AONs 22, 25, 28, 31, 34, 37, 40 and 43 were also investigated for stability in human blood serum. By visually comparing the gel pictures, the relative stabilities of all modified AONs in human blood serum are as follows: cLNA-MeC modified AON 34 and 37 > cLNA-T modified AON 40 and 43 > cLNA-A modified AON 22 and 25 > cLNA-G modified AON 28 and 31 > native AON. It appears that the relative stability of AONs in human blood serum is similar to that observed upon treatment of 3′-exonuclease (SVPDE).

3.3 RNase H-mediated RNA degradation in modified AON/RNA hybrids In the antisense approach, RNase H mediated RNA degradation is one of the most important antisense mechanisms that can be exploited once an antisense drug binds to its target RNA. In present study, α-L-carba-LNA and 7′-Me-cLNA containing AONs were pre-annealed with target RNA, followed by treatment with Escherichia coli RNase H1 or HeLa cell cytosolic extracts for RNA degradation investigations. The cleavage patterns and rates of the modified AON/RNA hybrids are discussed and compared to that of native DNA/RNA counterpart.

3.3.1 RNase H elicitation induced by AONs modified with α-L- carba-LNA and α-L-LNA analogues The RNase H recruitment study of synthesized α-L-carba-LNA derivatives modified AONs (containing a single type I, II, III or V modification) as well as the parent α-L-LNA modified AON (type IV) was carried out by treatment with Escherichia coli RNase H1. It was demonstrated that all the modified AONs 2-21/RNA hybrids are good substrates for RNase H. For type I, II, III and IV modified AON/RNA duplexes (Table 1), the cleavage patterns were found to be very similar and independent of the nature of the modification, but dependent on the site of modification. The cleavage activity of RNase H was suppressed within a stretch of 5 base pairs that starts from the modification site towards the 3′-end in the RNA strand. In addition, the original preferred A8 cleavage site also shifted to the edges of suppressed region if A8 is included within this 5 bp suppressed area (Paper II, Figure 5A). Hence, the observed RNase H cleavage patterns for type I, II, III and IV modified AON/RNA duplexes were found to be very closely similar to the previous studies of β−D-carba-LNA,194,200,203 β−D-carba-ENA191,194,200 and β−D-aza-ENA208,256 modified AON/RNA hybrids. It is noteworthy that the novel tetracyclic 6,7'-methylene bridged-α-L- carba-LNA (type V) modified AON 19/RNA and AON 20/RNA hybrids

50 showed different cleavage patterns compared to the type I, II, III and IV modified AON/RNA duplexes at the same position: a stretch of 3 bp region was observed for RNase H cleavage suppression in AON 19/RNA duplex, whereas a stretch of 5 bp suppressed region was also observed in AON 20/RNA duplex but with different preferred cleavage site and suppressed stretch region (see Paper II, Figure 5B). All the α-L-carba-LNA modified AON/RNA duplexes were degraded by RNase H with comparable cleavage rates as digestion of native AON/RNA hybrid (Figure 13). Remarkably, the cleavage rates of type III (7′R-methyl- α-L-carba-LNA) and IV (α-L-LNA) modified AON/RNA duplex were even two-times higher than cleavage of the native counterpart. It is previously reported that β-D-carba-LNA modified AON/RNA duplexes generally showed less RNase H digestion efficiency than the native counterpart. Therefore, we could arrive at the conclusion that α-L-carba-LNA modifications led to much less effect on RNase H elicitation than β-D-carba-LNA modifications.

0.025

Modification at 3-position from 3'-end Modification at 6-position from 3'-end Modification at 8-position from 3'-end Modification at 10-position from 3'-end 0.020 ) -1

0.015

0.010 Observed rates k (minObserved

0.005

0.000 Native I II III IV V Modification Type

H3C H O HO CH CH CH 5S 7'S 3 HO 3 7'R 3 O 7'S 6S 6'S 6'R 7'S NH Thy O N O O O Thy O Thy O Thy O O O O O O O O O O Type I Type II Type III Type IV Type V Figure 13. Bar plots of the observed cleavage rates of the RNase H1 mediated degradation of RNA in AON1-21/RNA hybrid duplexes.

51 3.3.2 RNase H elicitation induced by AONs modified with 7'- Me-cLNA-A, -G, -MeC, -T and LNA-A, -G, -C, -T analogues The RNase H recruitment study of cLNA-A, -G, -MeC, -T modified AON22- 45/RNA as well as LNA-A, -G, -C and -T modified AON46-57/RNA heteroduplexes (see Table 2 for AON entries) was carried out using native DNA sequence as a comparison. All the duplexes formed by AONs 22-57 with complementary RNA were also found to be excellent substrate for E. coli. RNase H1. As previously reported,191,194,200,201,203,211 the cleavage pattern was not affected by the structural nature of the modification, but depended upon the site of the modification (Paper III, Figure 8). It was shown that RNase H1 cleaved the central part of the RNA strand (between A6 and A13), with a preference on A13 and A6 for native DNA/RNA duplexes. However, for the AON strands modified with 7′-Me-cLNA and LNA analogues, the cleavage activity of RNase H1 was suppressed within a stretch of 5-6 nt region. Moreover, if one of the preferred cleavage sites A6 or A13 were included within this 5-6 nt region, the major cleavage sites were therefore suppressed (Paper III, Figure 8). -Me-cLNA-T) -Me-cLNA-T) R S C) Me -Me-cLNA-T)

0.070 R AON42 (7' AON45 (7' AON51 (LNA-G) 0.065 AON57 (LNA-T) -Me-cLNA- -Me-cLNA-G) AON41 (7' AON41 S S -Me-cLNA-G) ) 0.060 R C) -1 0.055 Me AON48 (LNA-A) AON39 (7' AON33 (7' AON30 (7' -Me-cLNA-T) 0.050 -Me-cLNA-G) S R -Me-cLNA- -Me-cLNA-G) S 0.045 R AON54 (LNA-C) AON44 (7' AON29 (7' 0.040 AON56 (LNA-T) AON32 (7' AON36 (7' C) 0.035 C) Me AON50 (LNA-G) Me -Me-cLNA-A) C) S C)

0.030 Me -Me-cLNA-A) Me R

0.025 -Me-cLNA- -Me-cLNA- S S AON27 (7' -Me-cLNA-T) -Me-cLNA- R AON24 (7' -Me-cLNA-

0.020 -Me-cLNA-A) R R R -Me-cLNA-A) -Me-cLNA-A) -Me-cLNA-G) S -Me-cLNA-G) -Me-cLNA-A) -Me-cLNA-T) R S S R S AON37 (7'

0.015 e (7' AON38

a AON40 (7' AON53 (LNA-C) AON35 (7' AON35

N Observed rates k (min Observed AON34 (7' AON22 (7'

0.010 AON52 (LNA-C) AON49 (LNA-G) AON49 AON46 (LNA-A) AON25 (7' AON55 (LNA-T) AON23 (7' AON47 (LNA-A) AON31 (7' AON26 (7' AON26 AON28 (7' AON28 0.005 AON43 (7' 0.000 pos3 pos4 pos6 pos7 pos8 pos9 Pos5 native pos10 pos11 pos13 pos15 pos16

Figure 14. Bar plots of the observed cleavage rates of the RNase H1 promoted degradation of RNA in native and 7′-Me-cLNA-A, -G, -MeC, -T modified AON22- 57/RNA hybrid duplexes.

52 Through comparison of pseudo first-order digestion rates of each AON/RNA hybrid, we found that the cleavage rates of duplexes formed by AONs single modified at 3-, 4-, 5-, 6-, 7- and 8-positions with RNA were similar or slightly lower than that of native AON/RNA counterpart, whereas the cleavage rates of duplexes formed by AONs single modified at 9-, 10-, 11-, 13-, 15- and 16-positions with RNA were from 2-fold to 8-fold higher than that of native AON/RNA counterpart (Figure 14). This result suggests that the RNA cleavage rates in duplexes formed by AON 22-57 (viz. cLNA and LNA modified oligos) with complementary RNAs were correlated with the modification site of each AON strand. Above all, it can be concluded that the cleavage patterns and rates of cLNA or LNA modified AON/RNA hybrids by RNase H1 are quite position-dependent, regardless of the structural nature of incorporated modifications.

53 Sammanfattning

Syntetiska oligonukleotider har i stor utsträckning tillämpats på nukleinsyrabaserade terapier genom att rikta och hämma cellulär RNA genom olika strategier (såsom RNase H mekanism, translationell blockering eller RNA- interferens). Olika sorter av kemiskt modifierade oligonukleotider har syntetiserats som resultat av detta och dessa har undersökts i syfte att förbättra cellulär stabilitet, “target” affinitet, farmakokinetik och leverans av de po- tentiella terapeutiska medlen. Utav dessa så har strukturellt begränsade oligonukleotider många fördelar på grund av deras gynnsamma egenskaper gentemot RNA affinitet, förbättrad nukleasstabilitet samt felfri RNase H rekryterings kapacitet jämfört med “native” motparter. De senaste åren har en av de viktigaste strukturellt begränsade nukleosider s.k. LNA (“Låst” nukleinsyra) syntetiserats av Imanish och Wengel et al. Vid intergrering i antisensoligonukleotider (AON), visades oöverträffade RNA riktade egenskaper (plus 5°C/mod i Tm jämfört med “native” motsvarighet). Därefter syntetiserades en rad 6 ', 7'-substituerade karba-LNA och ENA tymin analoger med fri radikal cykliseringsstategi av våra kollegor. AONs innehållande cLNA/cENA analoger uppvisade liknande RNA affinitet i jämförelse med LNA innehållande AON, men med mycket större nuk- leolytisk resistans än LNA motsvarigheten. Den gynnsamma funktionen hos 7'-Me-cLNA tymin uppmuntrade oss att syntetisera diastereomeriskt rena 7'S och R-Me-cLNA-A,-G, MeC analoger, jämföra deras RNA specificitet och nukleasstabilitet samt RNas H elicitation beträffande effekter orsakade av olika nukleobaser men även orienteringen av 7'-metyl-gruppen. Olika nukleobaser samt S och R konfiguration hos 7'-metyl i cLNA uppvisade inga up- penbara effekter på Tm värden, men med betydande inflytande på nukleolytiskt motstånd. En LNA diastereomer, omnämnd som α-L-LNA syntetiserades och rap- porterades visa sånär RNA affinitet som den hos LNA motsvarigheten. Ef- tersom AON innehållande α-L-LNA var nukleolytiskt stabilare än LNA motsvarigheten, lyckades vi syntetisera en serie 6', 7'-substituerade α-L- karba-LNA analoger, i syfte att utvärdera deras antisense egenskaper vid intergrening i antisensoligonukleotider. Även AON modifierade med α-L- karba-LNA uppvisade lägre Tm värden (minus 2-3 °C / mod) i jämförelse

54 med “native” motsvarighet, dessa visade dock en bättre nukleasstabilitet jämfört med “native” och α-L-LNA modifierade motsvarigheter. AON modifierade med cLNA-A-G,-MeC,-T och α-L-karba-LNA analoger med varierande positioner var “pre-annealed” med komplementär RNA följt av behandling med E. coli-RNas H 1 eller HeLa-cell cytosoliskt extrakt (rik- ligt med enzymen RNas H). Det visade att alla modifierade AON / RNA- duplexen var utmärkta substrat för RNas H. RNA klyvningsmönster samt hastighet av E. coli RNas H1 klyvning befanns vara beroende av modifie- rings positionen i AON sekvenserna, bortsett ifrån ändringarnas natur. Med hjälp av omfattande fysikalisk-kemiska och biokemiska studier av cLNA och α-L-karba-LNA modifierade AON föreslog vi att våra karba- LNA derivat kan användas som gynnsama RNA riktade medel inte bara terapeutiskt utan även för diagnostiska tillämpningar. Sammanfattningsvis så har strukturellt begränsade oligonukleotider varit ett lovande verktyg till att bredda utsräckningen av nukleinsyrabaserade läkemdel gentemot genterapi.

55 Acknowledgements

This work has been carried out at the Bioorganic Chemistry Program (BOC), Department of Cell and Molecular Biology (ICM), Biomedical Center (BMC), Uppsala University (August 2007-September 2012). Hereby, I wish to appreciate all the people who provided me invaluable suggestion and encouragement. In particular, I would like to express my sincere gratitude to the following individuals: My Ph.D supervisor Professor Jyoti Chattopadhyaya for accepting and supporting me as a Ph.D student in his luxurious equipped laboratory, for introducing me into the field of nucleotide and nucleic acid chemistry, for his thoughtful and creative ideas in scientific research as well as for his un- selfishly sharing rich life experience with me. My co-supervisor Dr. Oleksandr Plashkevych for his good example of se- rious scientific attitude, for helpful advices on scientific research, for fruitful collaborations, for solving so many computer-related problems as well as for sharing happy time on push-up exercises. Docent Andras Földesi for kind and valuable suggestion on scientific research, for his assistance in repairing and maintenance of DNA/RNA synthesizer and NMR instruments, also for his kindness on my thesis correction. All the former and present colleagues in our group: Dr. Chuanzheng zhou for his patient teaching in the synthesis and purification of oligonucleotides, as well as 32P labeling and enzymatic studies, and also for the helpfulness and good collaboration in scientific research; Jaana Evander and Anders Eriksson for their kind assistances with administrational work; Dr. Yi Liu, Dr. Jiangfeng Xu, Sayeh Erfan, Mansoureh Karimiahmadabadi, Fengfeng Yuan, Dr. RamShankar Upadhayaya, Dr. Sachin Gangadhar Deshpande, Dr. Shailesh Satish Dixit, Dr. Naresh Badgujar, Dr. Christelle Dupouy, Dr. Mounir Andaloussi for being good collaborators, encouraging and growing up each other, and for keeping a harmonious atmosphere in the laboratory. Professor Åke Engström for teaching me handle MALTI-TOF machine, also for providing valuable suggestions about mass spectra of oligonucleotides. My Master Supervisor Professor Serge van Calenbergh in Ghent Univer- sity, Belgium, for giving me chance to study in your group, and for leading me into medicinal chemistry of nucleosides and nucleotides.

56 All people in ICM for kind assistance in 32P labeling and phosphorimag- ing. All friends in China as well as in Sweden for sharing happy time together, and good friendship. Special thanks to my beloved Jie Li, for your patient waiting for me in remote China, and for your encouragement when I encountered the difficul- ties, and for supporting all the time. Great thanks to my parents, for you raising me up in a happy and healthy family, for your hard work to create a superior environment and strict re- quirement in my whole youthhood durations, for you encouraging me leave faraway from you and pursue a Ph.D degrees overseas, also for your endless love all the time. In the end, I would like to contribute this thesis to my passing-away grandfather and grandmother for your love and care from child. I have to say that the completion of this thesis is a result of your blessings in the paradise.

57 References

(1) Avery, O. T.; MacLeod, C. M.; McCarty, M. J. Exp. Med. 1944, 79, 137-158. (2) Hershey, A. D.; Chase, M. J. Gen. Physiol. 1952, 36, 39-56. (3) Weinger, J. S.; Parnell, K. M.; Dorner, S.; Green, R.; Strobel, S. A. Nat. Struct. Mol. Biol. 2004, 11, 1101-1106. (4) Trobro, S.; Aqvist, J. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 12395-12400. (5) Eddy, S. R. Nat. Rev. Genet. 2001, 2, 919-929. (6) Huttenhofer, A.; Schattner, P.; Polacek, N. Trends Genet. 2005, 21, 289-297. (7) Bachellerie, J. P.; Cavaille, J.; Huttenhofer, A. Biochimie 2002, 84, 775-790. (8) Lee, R. C.; Feinbaum, R. L.; Ambros, V. Cell 1993, 75, 843-854. (9) Lagos-Quintana, M.; Rauhut, R.; Lendeckel, W.; Tuschl, T. Science 2001, 294, 853-858. (10) Lau, N. C.; Lim., L. P.; Weinstein, E. G.; Bartel, D. P. Science 2001, 294, 858- 862. (11) Lee, R. C.; Ambros, V. Science 2001, 294, 862-864. (12) Bartel, D. P. Cell 2004, 116, 281-297. (13) He, L.; Hannon, G. J. Nat. Rev. Genet. 2004, 5, 522-531. (14) Bushati, N.; Cohen, S. M. Annu. Rev. Cell Dev. Biol. 2007, 23 , 175-205. (15) Fire, A.; Xu, S. Q.; Montgomery, M. K.; Kostas, S. A.; Driver, S. E.; Mello, C. C. Nature 1998, 391, 806-811. (16) Dorsett, Y.; Tuschl, T. Nature Rev. Drug Discov. 2004, 3, 318-329. (17) Houwing, S.; Kamminga, L. M.; Berezikov, E.; Cronembold, D.; Girard, A.; Van den elst, H.; Filippov, D. V.; Blaser, H.; Raz, E.; Moens, C. B.; Plasterk, R. H.; Hannon, G. J.; Draper, B. W.; Ketting, R. F. Cell 2007, 129, 69-82. (18) Seto, A. G. K.; RELau, E.; Lau, N. C. Mol. Cell 2007, 26, 603-609. (19) Siomi, M. C.; Sato, K.; Pezic, D.; Aravin, A. A. Nat. Rev. Mol. Cell. Biol. 2011, 12, 246-258. (20) Nudler, E.; Mironov, A. S. Trends. Biochem. Sci. 2004, 29, 11-17. (21) Vitreschak, A. G.; Rodionov, D. A.; Mironov, A. A.; Gelfand, M. S. Trends Genet. 2004, 20, 44-50. (22) Tucker, B. J.; Breaker, R. R. Curr. Opin. Struct. Biol. 2005, 15, 342-348. (23) Crick, F. H. J. Mol. Biol. 1968, 38, 367-379. (24) Orgel, L. E. J. Mol. Biol. 1968, 38, 381-393. (25) Gilbert, W. Nature 1986, 319, 618-618. (26) Chow, C. S.; Bogdan, F. M. Chem. Rev. 1997, 97, 1489-1513. (27) Oivanen, M.; Kuusela, S.; Lonnberg, H. Chem. Rev. 1998, 98, 961-990. (28) IUPAC-IUB Eur. J. Biochem. 1983, 131, 9-15. (29) Saenger, W. Principles of nucleic acid structure 1984. (30) Donohue, J.; Trueblood, K. N. J. Mol. Biol. 1960, 2, 363-371.

58 (31) Haschemeyer, A. E. V.; Rich, A. J. Mol. Biol. 1967, 27, 369-384. (32) Viswamitra, M. A.; Reddy, B. S.; Lin, G. H.-Y.; Sundaralingam, M. J. Am. Chem. Soc. 1971, 93, 4565-4572. (33) Dickerson, R. E.; Drew., H. R. Proc. Natl. Acad. Sci. U. S. A. 1982, 78, 7318- 7322. (34) Guschlbauer, W.; Fric, I. H., A. Eur. J. Biochem. 1972, 31, 1-13. (35) Son, T. D.; Guschlbauer, W.; Guéron, M. J. Am. Chem. Soc. 1972, 94, 7903- 7911. (36) Tavale, S. S.; Sobell, H. M. J. Mol. Biol. 1970, 48, 109-123. (37) Miles, D. W.; Townsend, L. B.; Robins, M. J.; Robins, R. K.; Inskeep, W. H.; Eyring, H. J. Am. Chem. Soc. 1971, 93, 1600-1608. (38) Suck, D.; Saenger, W. J. Am. Chem. Soc. 1972, 94, 6520-6526. (39) Yasuniwa, M.; Tokuoka, R.; Ogawa, K.; Yamagata, Y.; Fujii, S.; Tomita, K.-I.; Limn, W.; Ikehara, M. Biochim. Biophys. Acta. 1979, 561, 240-247. (40) Hall, L. D. Chem. Ind. (London) 1963, 950-951. (41) Altona, C.; Sundaralingam, M. J. Am. Chem. Soc. 1972, 94, 8205-8212. (42) Thibaudeau, C.; Acharya, P.; Chattopadhyaya, J. Stereoelectronic effects in nucleosides and nucleotides and their structural implications 2005, Second ed., Uppsala University Press. (43) Plavec, J.; Tong, W. M.; Chattopadhyaya, J. J. Am. Chem. Soc. 1993, 115, 9734- 9746. (44) Plavec, J.; Thibaudeau, C.; Chattopadhyaya, J. Pure Appl. Chem. 1996, 68, 2137-2144. (45) Thibaudeau, C.; Plavec, J.; Chattopadhyaya, J. J. Am. Chem. Soc. 1994, 116, 8033-8037. (46) Luyten, I.; Thibaudeau, C.; Sandstrom, A.; Chattopadhyaya, J. Tetrahedron 1997, 53, 6433-6464. (47) Plavec, J.; Roselt, P.; Foldesi, A.; Chattopadhyaya, J. Magn. Reson. Chem. 1998, 36, 732-740. (48) Acharya, P.; Trifonova, A.; Thibaudeau, C.; Foldesi, A.; Chattopadhyaya, J. Angew. Chem. Int. Ed. Engl. 1999, 38, 3645-3650. (49) Watson, J. D.; Crick, F. H. C. Nature 1953, 171, 737-738. (50) Hoogsteen, K. Acta Crystallogr. 1959, 171, 737-738. (51) Hoogsteen, K. Acta Crystallogr. 1963, 16, 907-916. (52) Abrescia, N. G. A.; Thompson, A.; Huynh-Dinh, T.; Subirana, J. A. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 2806-2811. (53) Polonius, F. A.; Müller, J. Angew. Chem. Int. Ed. Engl. 2007, 46, 5602-5604. (54) Rich, A. Gene 1993, 135, 99-109. (55) Simonsson, T. Biological Chemistry 2001, 382, 621-628. (56) Burge, S.; Parkinson, G. N.; Hazel, P.; Todd, A. K.; Neidle, S. Nucleic Acids Res. 2006, 34, 5402-5415. (57) Grigoriev, M.; Praseuth, D.; Guieysse, A. L.; Robin, P.; Thuong, N. T.; Helene, C.; Harel-Bellan, A. Proc. Natl. Acad. Sci. U. S. A. 1993, 90, 3501-3505. (58) Moser, H. E.; Dervan, P. B. Science 1987, 238, 645-650. (59) DeVoe, H.; Tinoco. Jr, I. J. Mol. Biol. 1962, 4, 500-517. (60) Herskovits, T. T. Arch. Biochem. Biophys. 1962, 97, 474-484. (61) Herskovits, T. T. Biochemistry 1963, 2, 335-340.

59 (62) Solie, T. N.; Schellman, J. A. J. Mol. Biol. 1968, 33, 61-77. (63) Gratzer, W. Eur. J. Biochem. 1969, 10, 184-187. (64) Davies, D. B. Jerus. Symp. Quant. Chem. Biochem. 1978, 11, 71-85. (65) Wing, R.; Drew, H.; Takano, T.; Broka, C.; Tanaka, S.; Itakura, K.; Dickerson, R. E. Nature 1980, 287, 755-758. (66) Crooke, S. T. Annu. Rev. Med 2004, 55, 61-95. (67) Fedoroff, O. Y.; Salazar, M.; Reid, B. R. J. Mol. Biol. 1993, 233, 509-523. (68) Horton, N. C.; Finzel, B. C. J. Mol. Biol. 1996, 264, 521-533. (69) Xiong, Y.; Sundaralingam, M. Nucleic Acids Res. 2000, 28, 2171-2176. (70) Nowotny, M.; Gaidamakov, S. A.; Crouch, R. J.; Yang, W. Cell 2005, 121, 1005-1016. (71) Rozenberg, H.; Rabinovich, D.; Frolow, F.; Hegde, R. S.; Shakked, Z. Proc. Natl. Acad. Sci. U. S. A 1998, 95, 15194-15199. (72) Kurreck, J. Eur. J. Biochem. 2003, 270, 1628-1644. (73) Rayburn, E. R. Expert Opin. Emerg. Drugs 2006, 11, 337-352. (74) Rayburn, E. R.; Zhang, R. Drug Discov Today 2008, 13, 513-521. (75) Cech, T. R.; Zaug, A. J.; Grabowski, P. J. Cell 1981, 27, 487-496. (76) Zaug, A. J.; Cech, T. R. Science 1986, 231, 470-475. (77) Breaker, R. R.; Joyce, G. F. Chem Biol 1994, 1, 223-229. (78) Djordjevic, M. Biomol. Eng 2007, 24, 179-189. (79) Nielson, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O. Science 1991, 254, 1497- 1500. (80) Egholm, M.; Buchardt, O.; Christensen, L.; Behrens, C.; Freier, S. M.; Driver, D. A.; Berg, R. H.; Kim, S. K.; Norden, B.; Neilsen, P. E. Nature 1993, 365, 556-568. (81) Obika, S.; Nanbu, D.; Hari, Y.; Morio, K.-I.; Y., I.; Ishida, T.; Imanishi, T. Te- trahedron Lett. 1997, 38, 8735-8738. (82) Koshkin, A. A.; Nielsen, P.; Meldgaard, M.; Rajwanshi, V. K.; Singh, S. K.; Wengel, J. J. Am. Chem. Soc. 1998, 120, 13252-13253. (83) Obika, S.; Nanbu, D.; Hari, Y.; Andoh, J.-I.; Morio, K.-I.; Doi, T.; Imanishi, T. Tetrahedron Lett. 1998, 39, 5401-5404. (84) Singh, S. K.; Wengel, J. Chem. Commun. 1998, 1247-1248. (85) Koshkin, A. A.; Singh, S. K.; Nielsen, P.; Rajwanshi, V. K.; Kumar, R.; Meldgaard, M.; Olsen, C. E.; Wengel, J. Tetrahedron 1998, 54, 3607-3630. (86) Singh, S. K.; Nielsen, P.; Koshkin, A. A.; Wengel, J. Chem. Commun. 1998, 455-456. (87) Crooke, S. T. Antisense Nucleic Acid Drug Dev 1998, 8, 133-134. (88) Pan, W. H.; Clawson, G. A. J. Cell. Biochem. 2006, 98, 14-35. (89) Esau, C. C. Methods 2008, 44, 55-60. (90) Kuznetsova, S.; Ait-Si-Ali, S.; Nagibneva, I.; Troalen, F.; Le Villain, J.-P.; Harel-Bellan, A.; Svinarchuk, F. Nucleic Acids Res. 1999, 27, 3995-4000. (91) Wang, G.; Seidman, M. M.; Glazer, P. M. Science 1996, 271, 802-805. (92) Maher, L. J. Cancer Invest 1996, 14, 66-82. (93) Unlenbeck, O. C. Nature 1987, 328, 596-600. (94) Walter, N. G.; Burke, J. M. Curr. Opin. Chem. Biol. 1998, 2, 24-30. (95) Schubert, S.; Kurreck, J. Curr. Drug Targets 2004, 5, 667-681. (96) Santoro, S. W.; Joyce, G. F. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 4262-4266.

60 (97) Dass, C. R.; Choong, P. F. M.; Khachigian, L. M. Mol Cancer Ther 2008, 7, 243-251. (98) Bernstein, E.; Caudy, A. A.; Hammond, S. M.; Hannon, G. J. Nature 2001, 409, 363-366. (99) Tuschl, T.; Zamore, P. D.; Lehmann, R.; Bartel, D. P.; Sharp, P. A. Genes Dev. 1999, 13, 3191-3197. (100) Zamore, P. D.; Tuschl, T.; Sharp, P. A.; Bartel, D. P. Cell 2000, 101, 25-33. (101) Elbashir, S. M.; Lendeckel, W.; Tuschl, T. Genes Dev. 2001, 15, 188-200. (102) Elbashir, S. M.; Harborth, J.; Lendeckel, W.; Yalcin, A.; Weber, K.; Tuschl, T. Nature 2001, 411, 494-498. (103) Shi, Y. Trends Genet. 2003, 19, 9-12. (104) Zou, G.-M.; Wu, W.; Chen, J.; Rowley, J. D. Biol. Cell 2003, 95, 365-371. (105) de Fougerolles, A.; Vornlocher, H.-P.; Maraganore, J.; Lieberman, J. Nat. Rev. Drug Discov 2007, 6, 443-453. (106) Ambros, V. Cell 2001, 107, 823-826. (107) Bartel, D. P. Cell 2009, 136, 215-233. (108) Georges, M.; Coppieters, W.; Charlier, C. Curr. Opin. Genet. Dev 2007, 17, 166-176. (109) Jeyaseelan, K.; Herath, W. B.; Armugam, A. Expert. Opin. Ther. Targets 2007, 11, 1119-1129. (110) Esau, C. C.; Monia, B. P. Adv. Drug Deliv. Rev 2007, 59, 101-114. (111) http://mirnablog.com/first-microrna-targeted-drug-reaches-phase-ii-clinical- trial/. (112) Ireson, C. R.; Kelland, L. R. Mol. Cancer Ther 2006, 5, 2957-2962. (113) Wilson, C.; Keefe, A. D. Curr. Opin. Chem. Biol. 2006, 10, 607-614. (114) Que-Gewirth, N. S.; Sullenger, B. A. Gene Ther 2007, 14, 283-291. (115) Gragoudas, E. S.; Adamis, A. P.; Cunningham, E. T.; Feinsod, M.; Guyer, D. R. N. Engl. J. Med. 2004, 351, 2805-2816. (116) Ni, X.; Castanares, M.; Mukherjee, A.; Lupold, S. E. Curr Med Chem. 2011, 18, 4206-4214. (117) Michelson, A. M.; Todd, A. R. J. Chem. Soc. 1955, 2362-2368. (118) Khorana, H. G.; Tener, G. M.; Moffatt, J. G.; Pol, E. H. Chem. Ind. (London) 1956, 1523-1523. (119) Khorana, H. G.; Razzell, W. E.; Gilham, P. T.; Tener, G. M.; Pol, E. H. J. Am. Chem. Soc. 1957, 79, 1002-1003. (120) Gilham, P. T.; Khorana, H. G. J. Am. Chem. Soc. 1958, 80, 6212-6222. (121) Reese, C. B. Org. Biomol. Chem. 2005, 3, 3851-3868. (122) Letsinger, R. L.; Lunsford, W. B. J. Am. Chem. Soc. 1976, 98, 3655-3661. (123) Beaucage, S. L.; Caruthers, M. H. Tetrahedron Lett. 1981, 22, 1859-1862. (124) Caruthers, M. H. Science 1985, 230, 281-285. (125) Ogilvie, K. K.; Sadana, K. L.; Thompson, E. A.; Quilliam, M. A.; Westmore, J. B. Tetrahedron Lett. 1974, 33, 2861-2863. (126) Ogilvie, K. K.; Theriault, N.; Sadana, K. L. J. Am. Chem. Soc. 1977, 99, 7741- 7743. (127) Usman, N.; Ogilvie, K. K.; Jiang, M. Y.; Cedergren, R. J. J. Am. Chem. Soc. 1987, 109, 7845-7854.

61 (128) Pitsch, S.; Weiss, P. A.; Jenny, L.; Stutz, A.; Wu, X. Helv. Chim. Acta 2001, 84, 3773-3795. (129) Ohgi, T.; Masutomi, Y.; Ishiyama, K.; Kitagawa, H.; Shiba, Y.; Yano, J. Org. Lett. 2005, 7, 3477-3480. (130) Zhou, C.; Honcharenko, D.; Chattopadhyaya, J. Org. Biomol. Chem. 2007, 5, 333-343. (131) Beaucage, S. L.; Caruthers., M. H. Current Protocols in Nucleic Acid Chemistry 2000, John Wiley & Sons, Inc. (132) Jiang, J.; Sheng, J.; Carrasco, N.; Huang, Z. Nucleic Acids Res. 2007, 35, 477- 485. (133) Harris, M. E.; Pace, N. R. Cell 1995, 1, 210-218. (134) Scott, W. G.; Finch, J. T.; Klug, A. Cell 1995, 81, 991-1002. (135) Chaulk, S. G.; MacMillan, A. M. Nucleic Acids Res. 1998, 26, 3173-3178. (136) Ahlborn, C.; Siegmund, K.; Richert, C. J. Am. Chem. Soc. 2007, 129, 15218- 15232. (137) Zhang, N.; Appella, D. H. J. Am. Chem. Soc. 2007, 129, 8424-8425. (138) Socher, E.; Bethge, L.; Knoll, A.; Jungnick, N.; Herrmann, A.; Seitz, O. Angew. Chem. Int. Ed. Engl. 2008, 47, 9555-9559. (139) Buchini, S.; Leumann, C. J. Curr. Opin. Chem. Biol. 2003, 7, 717-726. (140) Uhlmann, E.; Peyman, A. Chem. Rev. 1990, 90, 543-584. (141) Braasch, D. A.; Jensen, S.; Liu, Y. H.; Kaur, K.; Arar, K.; White, M. A.; Corey, D. R. Biochemistry 2003, 42, 7967-7975. (142) Manoharan, M. Curr. Opin. Chem. Biol. 2004, 8, 570-579. (143) Bumcrot, D.; Manoharan, M.; Koteliansky, V.; Sah, D. W. Y. Nature Chem. Biol. 2006, 2, 711-719. (144) Micklefield, J. Curr. Med. Chem. 2001, 8, 1157-1179. (145) Eckstein, F. Ann. Rev. Biochem. 1985, 54, 367-402. (146) Campbell, J. M.; Bacon, T. A.; Wickstrom, E. Biochem. Biophys. Methods 1990, 20, 259-267. (147) Levin, A. A. Biochim. Biophys. Acta. 1999, 1489, 69-84. (148) Sood, A.; Shaw, B. R.; Spielvogel, B. F. J. Am. Chem. Soc. 1990, 112, 9000- 9001. (149) McCuen, H. B.; Noé, M. S.; Sierzchala, A. B.; Higson, A. P.; Caruthers, M. H. J. Am. Chem. Soc. 2006, 128, 8138-8139. (150) Miller, P. S.; Agris, C. H.; Murakami, A.; Reddy, P. M.; Spitz, S. A.; Ts′o, P. O. P. Nucleic Acids Res. 1983, 11, 6225-6242. (151) Agrawal, S.; Goodchild, J. Tetrahedron Lett. 1987, 28, 3539-3542. (152) Sheehan, D.; Lunstad, B.; Yamada, C. M.; Stell, B. G.; Caruthers, M. H.; Dellinger, D. J. Nucleic Acids Res. 2003, 31, 4109-4118. (153) Yamada, C. M.; Dellinger, D. J.; Caruthers, M. H. J. Am. Chem. Soc. 2006, 128, 5251-5261. (154) Gryaznov, S. M.; Chen, J.-K. J. Am. Chem. Soc. 19946, 3143-3144., 11 (155) Gryaznov, S. M.; Lloyd, D. H.; Chen, J.-K.; Schultz, R. G.; Dedionisio, L. A.; Ratmeyer, L.; Wilson, W. D. Proc. Natl. Acad. Sci. U. S. A. 1995, 92, 5798- 5802. (156) Prakash, T. P. Chem Biodivers. 2011, 8, 1616-1641.

62 (157) Inoue, H.; Hayase, Y.; Imura, A.; Iwai, S.; Miura, K.; Ohtsuka, E. Nucleic Acids Res. 1987, 15, 6131-6148. (158) Lamond, A. I.; Sproat, B. S. FEBS Lett. 1993, 325, 123-127. (159) Martin, P. Helv. Chim. Acta 1995, 78, 486-504. (160) Altmann, K.-H.; Dean, N. M.; Fabbro, D.; Freier, S. M.; Geiger, T.; Haner, R.; Husken, D.; Martin, P.; Monia, B. P. B.; Miller, M.; Natt, F.; Nicklind, P.; Phil- lipsd, J.; Pieles, U.; Sasmor, H.; Moser, H. E. Chimia 1996, 50, 168-176. (161) Gallo, M.; Montserrat, J. M.; Iribarren, A. M. Braz. J Med. Biol. Res. 2003, 36, 143-151. (162) Kawasaki, A. M.; Casper, M. D.; Freier, S. M.; Lesnik, E. A.; Zounes, M. C.; Cummins, L. L.; Gonzalez, C.; Cook, P. D. J. Med. Chem. 1993, 36, 831-841. (163) Petersen, M.; Bondensgaard, K.; Wengel, J.; Jacobsen, J. P. J. Am. Chem. Soc. 2002, 124, 5974-5982. (164) Verheggen, I.; Van Aerschot, A.; Van Meervelt, L.; Rozenski, J.; Wiebe, L.; Snoeck, R.; Andrei, G.; Balzarini, J.; Claes, P.; Clercq, E. D.; Herdewijn, P. J. Med. Chem. 1995, 38, 826-835. (165) Hendrix, C.; Rosemeyer, H.; Verheggen, I.; Seela, F.; Aerschot, A. V.; Seela, F.; Herdewijn, P. Chem. Eur. J. 1997, 1, 110-120. (166) Hunziker, J.; Roth, H.-J.; Böhringer, M.; Giger, A.; Didedrichsen, U.; Göbel, M.; Krishnan, R.; Jaun, B.; Leumann, C.; Eschenmoser, A. Helv. Chim. Acta 1993, 76, 259-352. (167) Luyten, I.; Herdewijn, P. Eur. J. Med. Chem. 1998, 33, 515-578. (168) Herdewijn, P. Antisense Nucleic Acid Drug Dev 2000, 10, 297-310. (169) Froehler, B. C.; Wadwani, S.; Terhorst, T. J.; Gerrard, S. R. Tetrahedron Lett. 1992, 33, 5307-5310. (170) Wagner, R. W.; Matteucci, M. D.; Lewis, J. G.; Gutierrez, A. J.; Moulds, C.; Froehler, B. C. Science 1993, 260, 1510-1513. (171) Inoue, H.; Imura, A.; Ohtsuka, E. Nucleic Acids Res. 1985, 13, 7119-7128. (172) Lin, K.-Y.; Jones, R. J.; Matteucci, M. D. J. Am. Chem. Soc. 1995, 117, 3873- 3874. (173) Matteucci, M. D.; von Krosigk, U. Tetrahedron Lett. 1996, 37, 5057-5060. (174) Buhr, C. A.; Matteucci, M. D.; Froehler, B. C. Tetrahedron Lett. 1999, 40, 8969-8970. (175) Chollet, A.; Kawashima, E. Nucleic Acids Res. 1988, 16, 305-317. (176) Chazin, W. J.; Rance, M.; Chollet, A.; Leupin, W. Nucleic Acids Res. 1991, 19, 5507-5513. (177) Gryaznov, S. M.; Schultz, R. G. Tetrahedron Lett. 1994, 35, 2489-2492. (178) Zhou, C.; Chattopadhyaya, J. Curr. Opin. Drug. Discovery Dev. 2009, 12, 876- 898. (179) Altmann, K. H.; Imwinkelried, R.; Kesselring, R.; Rihs, G. Tetrahedron Lett. 1994, 35, 7625-7628. (180) Marquez, V. E.; Siddiqui, M. A.; Ezzitouni, A.; Russ, P.; Wang, J. Y.; Wagner, R. W.; Matteucci, M. D. J. Med. Chem. 1996, 39, 3739-3747. (181) Altmann, K. H.; Kesselring, R.; Francotte, E.; Rihs, G. Tetrahedron Lett. 1994, 35, 2331-2334. (182) Bogucka, M.; Naus, P.; Pathmasiri, W.; Barman, J.; Chattopadhyaya, J. Org. Biomol. Chem. 2005, 3, 4362-4372.

63 (183) Honcharenko, D.; Varghese, O. P.; Plashkevych, O.; Barman, J.; Chattopadhyaya, J. J. Org. Chem. 2006, 71, 299-314. (184) Pradeepkumar, P. I.; Cheruku, P.; Plashkevych, O.; Acharya, P.; Gohil, S.; Chattopadhyaya, J. J. Am. Chem. Soc. 2004, 126, 11484-11499. (185) Obika, S.; Sekiguchi, M.; Somjing, R.; Imanishi, T. Angew. Chem. Int. Ed. Engl. 2005, 44, 1944-1947. (186) Stauffiger, A.; Leumann, C. Nucleosides, Nucleotides Nucleic Acids 2007, 26, 615-619. (187) Steffens, R.; Leumann, C. Helv. Chim. Acta 1997, 80, 2426-2439. (188) Tarkoy, M.; Bolli, M.; Schweizer, B.; Leumann, C. Helv. Chim. Acta 1993, 76, 481-510. (189) Albaek, N.; Petersen, M.; Nielsen, P. J. Org. Chem. 2006, 71, 7731-7740. (190) Enderlin, G.; Nielsen, P. J. Org. Chem. 2008, 73, 6891-6894. (191) Liu, Y.; Xu, J.; Karimiahmadabadi, M.; Zhou, C.; Chattopadhyaya, J. J. Org. Chem. 2010, 75, 7112-7128. (192) Morita, K.; Hasegawa, C.; Kaneko, M.; Tsutsumi, S.; Sone, J.; Ishikawa, T.; Imanishi, T.; Koizumi, M. Bioorg. Med. Chem. Lett. 2001, 12, 73-76. (193) Seth, P. P.; Vasquez, G.; Allerson, C. A.; Berdeja, A.; Gaus, H.; Kinberger, G. A.; Prakash, T. P.; Migawa, M. T.; Bhat, B.; Swayze, E. E. J. Org. Chem. 2010, 75, 1569-1581. (194) Zhou, C.; Liu, Y.; Andaloussi, M.; Badgujar, N.; Plashkevych, O.; Chattopadhy- aya, J. J. Org. Chem. 2009, 74, 118-134. (195) Hari, Y.; Obika, S.; Ohnishi, R.; Eguchi, K.; Osaki, T.; Ohishi, H.; Imanishi, T. Bioorg. Med. Chem. 2006, 14, 1029-1038. (196) Prakash, T. P.; Siwkowski, A.; Allerson, C. R.; Migawa, M. T.; Lee, S.; Gaus, H. J.; Black, C.; Seth, P. P.; Swayze, E. E.; Bhat, B. J. Med. Chem. 2010, 53, 1636-1650. (197) Rahman, S. M. A.; Seki, S.; Obika, S.; Yoshikawa, H.; Miyashita, K.; Imanishi, T. J. Am. Chem. Soc. 2008, 130, 4886-4896. (198) Seth, P. P.; Allerson, C. R.; Berdeja, A.; Siwkowski, A.; Pallan, P. S.; Gaus, H.; Prakash, T. P.; Watt, A. T.; Egli, M.; Swayze, E. E. J. Am. Chem. Soc. 2010, 132, 14942-14950. (199) Singh, S. K.; Kumar, R.; Wengel, J. J. Org. Chem. 1998, 63, 10035-10039. (200) Srivastava, P.; Barman, J.; Pathmasiri, W.; Plashkevych, O.; Wenska, M.; Chat- topadhyaya, J. J. Am. Chem. Soc. 2007, 129, 8362-8379. (201) Upadayaya, R.; Deshpande, S. G.; Li, Q.; Kardile, R. A.; Sayyed, A. Y.; Kshir- sagar, E. K.; Salunke, R. V.; Dixit, S. S.; Zhou, C.; Földesi, A.; Chattopadhyaya, J. J. Org. Chem. 2011, 76, 4408-4431. (202) Wang, G.; Giradet, J. L.; Gunic, E. Tetrahedron 1999, 55, 7707-7724. (203) Xu, J. F.; Liu, Y.; Dupouy, C.; Chattopadhyaya, J. J. Org. Chem. 2009, 74, 6534-6554. (204) Obika, S.; Nanbu., D.; Hari, Y.; Andoh, J.-I.; Morio, K.-I.; Doi, T.; Imanishi, T. Tetrahedron Lett. 1998, 39, 5401-5404. (205) Morita, K.; Takagi, M.; Hasegawa, C.; Kaneko, M.; Tsutsumi, S.; Sone, J.; Ishi- kawa, T.; Imanishi, T.; Koizumi, M. Bioorg. Med. Chem. 2003, 11, 2211-2226. (206) Morita, K.; Yamate, K.; Kurakata, S.; Abe, K.; Imanishi, T.; Koizumi, M. Nu- cleic Acids Res. Sup. 2002, 2, 99-100.

64 (207) Morita, K.; Yamate, K.; Kurakata, S.-I.; Watanabe, K.; Imanishi, T.; Koizumi, M. Nucleosides, Nucleotides Nucleic Acids 2003, 22, 1619-1621. (208) Varghese, O. P.; Barman, J.; Pathmasiri, W.; Plashkevych, O.; Honcharenko, D.; Chattopadhyaya, J. J. Am. Chem. Soc. 2006, 128, 15173-15187. (209) Kim, H. S.; Jacobson, K. A. Org. Lett. 2003, 5, 1665-1668. (210) Kumar, T. S.; Madsen, A. S.; Wengel, J.; Hrdlicka, P. J. J. Org. Chem. 2006, 71, 4188-4201. (211) Li, Q.; Yuan, F. F.; Zhou, C. Z.; Plashkevych, O.; Chattopadhyaya, J. J. Org. Chem. 2010, 75, 6122-6140. (212) Seth, P. P.; Allerson, C. R.; Berdeja, A.; Swayze, E. E. Bioorg. Med. Chem. Lett. 2011, 21, 588-591. (213) Sørensen, M. D.; Kværnø, L.; Bryld, T.; Håkansson, A. E.; Verbeure, B.; Gau- bert, G.; Herdewijn, P.; Wengel, J. J. Am. Chem. Soc. 2002, 124, 2164-2176. (214) Zhou, C.; Plashkevych, O.; Chattopadhyaya, J. Org. Biomol. Chem. 2008, 6, 4627-4633. (215) Xi, Z.; Agback, P.; Sandstrom, A.; Chattopadhyaya, J. Tetrahedron 1991, 47, 9675-9690. (216) Madsen, A. S.; Hrdlicka, P. J.; Kumar, T. S.; Wengel, J. Carbohydr. Res. 2006, 341, 1398-1407. (217) Obika, S.; Morio, K.; Hari, Y.; Imanishi, T. Bioorg. Med. Chem. Lett. 1999, 9, 515-518. (218) Obika, S.; Sekiguchi, M.; Osaki, T.; Shibata, N.; Masaki, M.; Hari, Y.; Imanishi, T. Tetrahedron Lett. 2002, 43, 4365-4368. (219) Koole, L. H.; Wu, J. C.; Neidle, S.; Chattopadhyaya, J. J. Am. Chem. Soc. 2199, 114, 2687-2696. (220) Velazquez, S.; Jimeno, M. L.; Huss, S.; Balzarini, J.; Camarasa, M. J. J. Org. Chem. 1994, 59, 7661-7670. (221) Wu, J. C.; Xi, Z.; Gioeli, C.; Chattopadhyaya, J. Tetrahedron 1991, 47, 2237- 2254. (222) Xi, Z.; Agback, P.; Plavec, J.; Sandstrom, A.; Chattopadhyaya, J. Tetrahedron 1992, 48, 349-370. (223) Xi, Z.; Glemarec, C.; Chattopadhyaya, J. Tetrahedron 1993, 49, 7525-7546. (224) Xi, Z.; Rong, J. H.; Chattopadhyaya, J. Tetrahedron 1994, 50, 5255-5272. (225) Osaki, T.; Obika, S.; Harada, Y.; Mitsuoka, Y.; Sugaya, K.; Sekiguchi, M.; Roongjang, S.; Imanishi, T. Tetrahedron 2007, 63, 8977-8986. (226) Jeong, L. S.; Marquez, V. E.; Yuan, C. S.; Borchardt, R. T. Heterocycles 1995, 41, 2651-2656. (227) Rodriguez, J. B.; Marquez, V. E.; Nicklaus, M. C.; Mitsuya, H.; Barchi, J. J. J. Med. Chem. 1994, 37, 3389-3399. (228) Shin, K. J.; Moon, H. R.; George, C.; Marquez, V. E. J. Org. Chem. 2000, 65, 2172-2178. (229) Giese, B. Radicals in Organic Synthesis: Formation of Carbon-Carbon Bonds 1986, Pergamon Press: New York. (230) Lamb, R. C.; Ayers, P. W.; Toney, M. K. J. Am. Chem. Soc. 1963, 85, 3483- 3486. (231) Hart, D. J. Science 1984, 223, 883-887. (232) Zhou, C.; Chattopadhyaya, J. Chem. Rev.2 201, 112, 3808-3832.

65 (233) Beckwith, A. L. J.; Schiesser, C. H. Tetrahedron 1985, 41, 3925-3941. (234) Ti, G. S.; Gaffney, B. L.; Jones, R. A. J. Am. Chem. Soc. 1982, 104, 1316-1316. (235) Koshkin, A. A.; Fensholdt, J.; Pfundheller, H. M.; Lomholt, C. J. Org. Chem. 2001, 66, 8504-8504. (236) Hakansson, A. E.; Koshkin, A. A.; Sørensen, M. D.; Wengel, J. J. Org. Chem. 2000, 65, 5161-5166. (237) Rajwanshi, V. K.; Hakansson, A. E.; Dahl, B. M.; Wengel, J. Chem. Commun. 1999, 1395-1396. (238) Rajwanshi, V. K.; Hakansson, A. E.; Kumar, R.; Wengel, J. Chem. Commun. 1999, 2073-2074. (239) Fluiter, K.; Frieden, M.; Vreijling, J.; Rosenbohm, C.; De Wissel, M. B.; Christensen, S. M.; Koch, T.; Ørum, H.; Baas, F. ChemBioChem 2005, 6, 1104- 1109. (240) Frieden, M.; Christensen, S. M.; Mikkelsen, N. D.; Rosenbohm, C.; Thrue, C. A.; Westergaard, M.; Hansen, H. F.; Ørum, H.; Koch, T. Nucleic Acids Res. 2003, 31, 6363-6372. (241) Nacro, K.; Lee, J.; Barchi, J. J.; Lewin, N. E.; Blumberg, P. M.; Marquez, V. E. Tetrahedron 2002, 58, 5335-5345. (242) Barton, D. H. R.; McCombie, S. W. J. Chem. Soc., Perkin Trans. 1 1975, 16, 1574-1585. (243) Dess, D. B.; Martin, J. C. J. Org. Chem. 1983, 48, 4155-4156. (244) Dess, D. B.; Martin, J. C. J. Am. Chem. Soc. 1991, 113, 7277-7287. (245) Zhou, C.; Plashkevych, O.; Liu, Y.; Badgujar, N.; Chattopadhyaya, J. Hetero- cycles 2009, 78, 1715-1728. (246) Auffinger, P.; Westhof, E. Angew. Chem. Int. Ed. 2001, 40 , 4648-4650. (247) Ball, P. Chem. Rev. 2008, 108, 74-108. (248) Egli, M.; Minasov, G.; Teplova, M.; Kumar, R.; Wengel, J. Chem. Commun. 2001, 651-652. (249) Schneider, B.; Patel, K.; Berman, H. M. Biophys. J. 1998, 75, 2422-2434. (250) Teplova, M.; Minasov, G.; Teresshko, V.; Inamati, G. B.; Cook, P. D.; Manoha- ran, M.; Egli, M. Nat. Struct. Biol. 1999, 6, 535-539. (251) Tereshko, V.; Gryaznov, S.; Egli, M. J. Am. Chem. Soc. 1998, 120, 269-283. (252) McTigue, P. M.; Peterson, R. J.; Kahn, J. D. Biochemistry 2004, 43, 5388-5405. (253) Maier, M. A.; Leeds, J. M.; Balow, G.; Springer, R. H.; Bharadwaj, R.; Manoha- ran, M. Biochemistry 2002, 41, 1323-1327. (254) Shaw, J. P.; Kent, K.; Bird, J.; Fishback, J.; Froehler, B. Nucleic Acids Res. 1991, 19, 747-750. (255) Zhou, C.; Chattopadhyaya, J. J. Org. Chem. 2010, 75, 2341-2349. (256) Honcharenko, D.; Barman, J.; Varghese, O. P.; Chattopadhyaya, J. Biochemistry 2007, 46, 5635-5646. (257) Cerritelli, S. M.; Crouch, R. J. FEBS J. 2009, 276, 1494-1505. (258) Figiel, M.; Chon, H.; Cerritelli, S. M.; Cybulska, M.; Crouch, R. J.; Nowotny, M. J. Biol. Chem. 2011, 286, 10540-10550. (259) Pileur, F.; Toulme, J. J.; Cazenave, C. Nucleic Acids Res. 2000, 28, 3674-3683.