Aspects of Antisense and Antigene Chemistry of Oligonucleotides Tethered

to Intercalators

BY

DIMITRI OSSIPOV

Dissertation for the Degree of Doctor of Philosophy in Bioorganic Chemistry presented at Uppsala University in 2002 Abstract Ossipov, D., 2002. Aspects of Antisense and Antigene Chemistry of Oligonucleotides Tethered to Intercalators. Acta Univ. Ups., Comprehensive Summaries of Uppsala Dissertations from the Faculty of Science and Technology 735. 67 pp. Uppsala. ISBN 91-554-5365-1. Synthetic and physicochemical studies on appropriately functionalized ODN-conjugates have been performed to evaluate their abilities to act as antisense agents against RNA or as intramolecular DNA cross-linking agents. Intercalating aromatic systems [phenazine (Pnz), dipyridophenazine (DPPZ)] and metallointercalators such as Ru2+(phen)2(DPPZ) and Ru2+(tpy)(DPPZ)L [where L = chemically or photochemically labile ligand, phen = phenanthroline, tpy = terpyridine], which are covalently tethered to the oligodeoxynucleotides (ODNs), have been chosen for this purpose. The ODN-conjugates were typically prepared by automated solid phase synthesis using phosphoramidite building blocks, or on solid supports, both functionalized with the chromophore groups. The photosensitive metal complex, Ru2+(tpy)(DPPZ)(CH3CN), has been incorporated by post-synthetic coupling to the amino-linker modified ODNs via an amide bond. The intercalating ability of the tethered chromophores gave enhanced stability of the duplexes and triplexes formed with ODN-conjugates and their complementary targets: DNA, RNA, or double-stranded DNA. The conjugation of DPPZ chromophore to ODN (at 3', 5' or at the middle) led us to incorporate Ru2+(phen)2(DPPZ) through the DPPZ ligand, for the first time. The corresponding (Ru2+-ODN)•DNA duplexes showed dramatic stabilization (∆Tm = 19.4 – 22.0ºC). The CD and DNase I footprinting experiments suggest that the stabilization is owing to metallointercalation by threading of the Ru2+(phen)2 moiety through the ODN•DNA duplex core, thus “stapling” the two helical strands from the minor to major groove. On the other hand, Ru2+(tpy)(DPPZ)(CH3CN)-ODN conjugates represent a new class of oligonucleotides containing the photoactivatible Ru2+ complexes, which can successfully crosslink to the complementary strand. The mechanism of cross-linking upon photoirradiation of [Ru2+(tpy)(DPPZ)(CH3CN)-ODN]•DNA involves in situ conversion to the reactive [Ru2+(tpy)(DPPZ)(H2O)-ODN]•DNA which are subsequently cross-linked through the G residue of the complementary DNA strand. All starting materials and products have been purified by HPLC and/or by PAGE and subsequently charcterized by MALDI-TOF as well as ESI mass spectroscopy. Terminal conjugation of the planar Pnz and DPPZ groups through the flexible linkers were also shown to improve thermal stability of the ODN•RNA hybrid duplexes without alteration of the initial AB-type global helical structure as revealed from CD experiments. As a result, RNase H mediated cleavage of the RNA strand in the intercalator-tethered ODN•RNA duplexes was more efficient compared to the natural counterpart. The RNase H cleavage pattern was also found to be dependent on the chemical nature of the chromophore. It appeared that introduction of a tether at the 3'-end of the ODN can be most easily tolerated by the enzyme regardless of the nature of the appending chromophore. The tethered DPPZ group has also been shown to chelate Cu2+ and Fe3+, like phenanthroline group, followed by the formation of redox-active metal complex which cleaves the complementary DNA strand in a sequence-specific manner. This shows that the choice of appropriate ligand is useful to (i) attain improved intercalation giving Tm enhancement, and (ii) sequence-specifically inactivate target RNA or DNA molecules using multiple modes of chemistry (RNase H mediated cleavage, free-radical, oxidative pathways or photocross-linkage).

Dimitri Ossipov, Department of Bioorganic Chemistry, Biomedical Centre, Box 581, Uppsala University, SE-751 23 Uppsala, Sweden © Dimitri Ossipov 2002 ISSN 1104-232X ISBN 91-554-5365-1 Printed in Sweden by Tryck & Medier, Uppsala 2002

To memory of my father

who should have lived

to see this thesis

THE ORIGINAL PUBLICATIONS

This thesis is based on the following original publications, which are referred to by the Roman numerals. I. Ossipov, D.; Chattopadhyaya, J. Synthesis of 1'-Phenazine-Tethered Psicofuranosyl

Oligonucleotides: The Thermal Stability and Fluorescence Properties of Their Duplexes and Triplexes.

Tetrahedron 1998, 54, 5667-5682. II. Ossipov, D.; Zamaratski, E.; Chattopadhyaya, J. Dipyrido[3,2-a:2',3'-c]phenazine-Tethered Oligo-

DNA: Synthesis and Thermal Stability of Their DNA-DNA and DNA-RNA Duplexes and Triplexes.

Helvetica Chimica Acta 1999, 82, 2186-2200. III. Zamaratski, E.; Ossipov, D.; Pradeepkumar, P.I.; Amirkhanov, N.; Chattopadhyaya, J. The 3'-

Modified Antisense Oligos Promote Faster Hydrolysis of the Target RNA by RNase H than the Natural Counterpart.

Tetrahedron 2001, 57, 593-606. IV. Ossipov, D.; Pradeepkumar, P.I.; Chattopadhyaya, J. Synthesis of [Ru(phen)2dppz]2+-Tethered

Oligo-DNA and Studies on the Metallointercalation Mode into the DNA Duplex. Journal of the American Chemical Society 2001, 123, 3551-3562. V. Ossipov, D.; Gohil, S.; Chattopadhyaya, J. First Synthesis and Reactivity of the DNA-

[Ru(tpy)(dppz)(CH3CN)]2+ Conjugates. Journal of the American Chemical Society 2002 (submitted).

Reprints were made with permission from the publishers.

Table of Contents Abbreviations

1. Introduction 8

1.1 Formation of duplex and triplex structures by oligonucleotide-intercalator conjugates 9 1.2 Antisense oligodeoxynucleotides activating RNase H mediated cleavage of mRNA 10 1.3 Use of metal ions in the antisense and antigene strategies 11

2. Present Work 2.1 Synthesis of the ODNs tethered to phenazine, dipyridophenazine, and Ru2+- dipyridophenazine chromophores 14

2.1.1 Synthesis of Pnz labeled ODNs (Paper I) 14 2.1.2 Synthesis of amino-linkers of different lengths tethered to the glycerol moiety (Paper II) 16 2.1.3 Derivatization of DPPZ containing chromophores for conjugation (Papers II, IV & V) 20 2.1.4 Synthesis of DPPZ and Ru2+(phen)2(DPPZ) labeled ODNs (Papers II & IV) 21 2.1.5 Synthesis of Ru2+(tpy)(DPPZ)(CH3CN) labeled ODNs (Paper V) 23 2.1.6 Characterization of Ru2+(tpy)(DPPZ)(CH3CN) labeled ODNs (Paper V) 26 2.2 Thermal stability of duplexes and triplexes formed by phenazine, dipyridophenazine, and Ru2+-dipyridophenazine conjugated ODNs 27 2.2.1 Thermal stability of conjugated ODN•DNA duplexes (Papers I, II, IV & V) 27 2.2.2 Thermal stability of conjugated ODN•RNA duplexes (Papers II & IV) 30 2.2.3 Thermal stability of conjugated ODN×DNA•DNA triplexes (Papers I, II & IV) 31 2.2.4 Circular dichroism (CD) and DNase I footprinting studies on [Ru(phen)2(DPPZ)]2+- labeled ODN•DNA duplexes performed for the elucidation of the metallointercalation binding mode in the DNA double helixes (Paper IV) 34 2.3 RNase H degradation studies on the AON•RNA duplexes conjugated with various intercalators at AON strand (Paper III) 39 2.3.1 Physicochemical properties of RNA targets and their duplexes formed with AON conjugates 39 2.3.2 RNase H mediated cleavage studies on the conjugated AON•RNA duplexes 42 2.3.3 Nuclease resistance of the 3'-Pnz and 3'-DPPZ-conjugated AONs 46 2.4 Specific chemical modifications of the complementary DNA strand by DPPZ and Ru2+- DPPZ tethered ODNs in ODN•DNA duplexes 46 2.4.1 Sequence-specific cleavage of target DNA by metal-binding DPPZ-group linked to the internucleotide position of the complementary ODN strand (unpublished data) 46 2.4.3 Photochemical generation of Ru2+(tpy)(DPPZ)(H2O)-ODN (Paper V) 48 2.4.4 Photochemical cross-linking of Ru2+(tpy)(DPPZ)(CH3CN)-ODN conjugates to the complementary DNA strand in ODN•DNA duplexes (Paper V) 49 2.4.5 The extension of the idea of the Ru2+ complex activation via photoaquation (Paper V) 55 3. Acknowledgements 58 4. References 59

Abbreviations

AON antisense oligodeoxynucleotide

biq 2,2'-biquinoline

bp base pair

bpy 2,2'-bypyridine

CPG control pore glass

dA 2'-deoxyadenosine

dC 2'-deoxycytidine

dG 2'-deoxyguanosine

dmbpy 6,6'-dimethyl-2,2'-bypyridine

DMF dimethylformamide

DMT 4,4'-dimethoxytrityl

dpp 4,7-diphenyl-1,10-phenanthroline

DPPZ dipyrido[3,2-a;2',3'-c]phenazine

DTT dithiothreitol

dsDNA double-stranded DNA

EDTA ethylenediaminetetraacetic acid

ESI electrospray ionisation

Fmoc 9-fluorenylmethoxycarbonyl

HPLC high perfomance liquid chromatography

MALDI-TOF matrix-assisted laser desorption ionization – time-of-flight

MPE methidiumpropyl-EDTA

mRNA messenger RNA

MS mass spectroscopy

NHS N-hydroxysuccinimidyl

ODN oligodeoxynucleotide

ON oligonucleotide

PAGE polyacrylamide gel electrophoresis

phen 1,10-phenanthroline

phi 9,10-phenanthrenequinone diimine

Pnz phenazine

PO phosphodiester

PS phosphorothioate

pypz 1-(2'-pyridyl)-3,5-dimethylpyrazole

ssDNA single-stranded DNA

T thymidine

tap 1,4,5,8-tatraazaphenanthrene

TBAF n-tetrabutylammonium fluoride

THF tetrahydrofurane

Tm melting temperature

TPDS 1,1,3,3-tetraisopropyldisiloxan-1,3-diyl

tpy 2,2':6',2''-terpyridine

TSU N,N,N',N'-tetramethyl(succinimidyl)uronium tetrafluoroborate

U uridine

UV-vis ultraviolet-visible

• Watson-Crick hydrogen bond

× Hoogsteen hydrogen bond

8

1. Introduction.

The idea to regulate gene expression by the use of synthetic oligonucleotides (ONs), which can

specifically recognize particular base sequence of nucleic acids, was first proposed in 1978 by

Zamecnik and Stephenson.1,2 Later, two main strategies have been considered: in the antisense

strategy,3-7 the ON is targeted to a specific mRNA via Watson-Crick8 base paring (ON•RNA duplex

formation), thereby inhibiting its translation into the corresponding protein;9,10 in the antigene

strategy,11-13 a dsDNA sequence is the target of the ON which is then expected to hybridize with

dsDNA involving Hoogsteen hydrogen bonding14 (ON×DNA•DNA triplex formation) and block

transcription of the specific gene where the target is located (symbols × and • denote here Hoogsteen

and Watson-Crick hydrogen bonding, respectively). Although short ONs are able to bind selectively to

target sequences under physiological conditions, their potential therapeutic use as artificial gene control

agents is possible if they comply with the following requirements:15 (1) ONs must be able to penetrate

the cell membrane to reach its site of action. Unlike low molecular weight agents, natural ONs are

polyanionic and cannot passively diffuse across cell membranes. (2) ONs must be resistant to

intracellular enzymes (endo- and exonucleases) that degrade nucleic acids. This precludes the use of

phosphodiester (PO) linked natural ONs as therapeutics because of their instability against nucleases.

(3) ONs must be able to interact with their cellular targets (such as mRNA or genomic dsDNA) i.e.

target sequences should be accessible for hybridization. As targets are invariably protein bound or

structurally folded, many sites are probably not accessible for interaction with ONs of defined

sequences. (4) ONs should not interact in a non-specific manner with other macromolecules as well as

their interaction within target nucleic acid should be sequence specific. (5) The complex formed

between the ON and its complementary target sequence must be sufficiently stable under physiological

conditions. Especially it concerns the triplex forming oligonucleotides (TFOs) bound to an oligopurine

target sequence in dsDNA, since the formation of dC+×dG•dC triplet is pH dependent with an optimum

stability at non-physiological pH 5.6-6.0, and the T×dA•T triplet is only stable under conditions of high

ionic strength.16,17 (6) Antisense oligodeoxynucleotides (AONs) have been found to inhibit translation

by steric blocking of the ribosomal machinery18-21, as well as by recruitment of an endogenous nuclease

RNase H which degrades the RNA strand in the AON•RNA duplex.22,23 Thus, the ability of an

AON•RNA duplex to be a substrate for the cellular RNase H enzyme is essential because of irreversible

degradation of mRNA.

Whereas the requirement for specificity is satisfactorily met by the natural ONs, sufficient passage

through membranes, adequate stability to nucleases, and improved binding affinity to the target

molecule can be achieved only by modification of the ON. Moreover, modification of natural ON by

9

tethering of chemically reactive groups can be used to induce irreversible sequence specific

modifications and/or cleavage of targeted nucleic acids24 thus permitting permanent inactivation of

unwanted genetic information. Among different chemical modifications of ONs the conjugation is

considered as a result of the coupling of ON to one or more molecules with distinct properties. In this

way the affinity of ON residue to complementary sequence is combined with specific characteristics of

the appending group(s) in the resulting ON conjugate.25 We report here the synthesis of ODN

conjugates, which enhance the binding affinity of the parent natural ODNs toward various targets

(single-stranded DNA or RNA molecules, and DNA duplexes), as well as show improvement of their

properties to elicit RNase H activity for complementary RNA cleavage in the corresponding

heteroduplex. Finally, we show when an appropriate functional group is tethered to a synthetic ODN, it

can specifically modify the target DNA strand in the ODN•DNA duplex.

1.1 Formation of duplex and triplex structures by oligonucleotide-intercalator conjugates.

In order to increase the stability of duplexes and triplexes formed by natural ONs, intercalating agents

have been attached to oligonucleotides without loss of specificity. It has generally been accepted that

planar aromatic chromophores can increase the π-stacking interactions if they are inserted between

adjacent base pairs of a duplex thus stabilizing the helical structure.26,27 Such polyaromatic agents as

phenazinium,28,29 acridine,30-34 ethidium,35 anthraquinone,36-38 fagaronine,39 pyrine,40-42 daunomycin,43

anthracene,44,45 and fluorescein42 were conjugated to a number of ONs at the 3'-, 5'-terminals,

internucleotide position, or through the base residue. Most of these ON conjugates were shown to

significantly increase the thermodinamic stability (expressed with melting temperature Tm) by 10-30°C.

The stabilizing effect of a chromophore was found to be dependent on (i) the type of the tether, (ii) the

linker length, and (iii) its site of attachment within the duplex. Generally, 5'-conjugation had more

noticeable effect on Tm than conjugation at the 3'-terminal of ODN sequence, whereas incorporation of a

potential intercalator at the central position of oligonucleotide chain mostly led to the destabilizing of a

resulting duplex.

Conjugation of intercalating agents was also applied to the triplex forming oligonucleotides

(TFOs) with the expectation of stabilization of resulting triplex provided by insertion of an intercalator

between base pairs at the triplex-duplex junction. This was indeed observed upon triplex formation by

TFOs derivatized with acridine at 5'-46-48 or 3'-ends49. Similar results were obtained with psoralen,50

phenanthroline,48 daunorubicin,51 and naphthalene diimade derivative52 attached to the 5'-end of an

oligonucleotide.53 Triple helix-specific intercalators, benzopyridoindoles and benzopyridoquinoxalines,

were covalently linked54,55 to the 5'-end as well as to the interior of a TFO for the remarkable

improvement of triplex stability under physiological conditions as compared with unmodified and

10

acridine modified counterparts. Despite the attractive idea to efficiently block dsDNA instead of many

copies of mRNA which are continuously produced from one equivalent of DNA, the major limitation of

antigene strategy is: (i) the low accessibility of the DNA strands which are protected with bound

proteins forming complex chromatin structure, and (ii) the requirement of the polypurine sequence of a

dsDNA target recognized by oligopyrimidine TFO. Therefore, several studies have been aimed at

circumventing this limitation. The extension of recognition sequences has been realized by

simultaneous binding of the homopyrimidine TFO to adjacent purine tracts on alternate strands of the

DNA duplex.56-58 The reverse orientation of the pyrimidine oligo-α-nucleotide TFO strand with respect

to that of the corresponding β-oligomer59 was also used to construct another set of molecules which are

able to bind to adjacent homopurine sequences on alternate DNA strands. Varios strategies have been

developed to accommodate a single pyrimidine mismatch in the homopurine tract. The incorporation of

an abasic site into a TFO at a site opposite a single interruption resulted in considerable destabilization

of the triplex.60 The reduction of stability was also observed upon incorporation of base triads

characterized by single hydrogen bond between the third strand base and the pyrimidine interruption

(like dG×T•dA).61 The recognition of base pair inversions within polypurine•polypyrimidine tract was

also achieved by TFO carrying a non-natural nucleoside, N4-(6-amino-2-pyridinyl)-deoxycytidine,62

which simultaneously form two hydrogen bonds with both bases of T•dA interruption. The

incorporation of acridine at an internal site of TFO allowed to stabilize the mismatched triplexes,63,64

thus extending the range of DNA sequences available for triplex formation. The stabilizing effect

brought by acridine intercalation at the base pair inversion site compensated the destabilization

introduced by mismatched base triplet. However, this intercalation did not provide much specificity.

1.2 Antisense oligodeoxynucleotides activating RNase H mediated cleavage of mRNA.

It has been demonstrated that the heteroduplex between natural phosphodiester (PO) AONs and the

target RNA is recognized by an intracellular nuclease RNase H that cleaves only the RNA strand of this

duplex. The RNase H is highly sensitive to structural alterations of the AON, and therefore only limited

number of certain modifications have been shown to support the RNase H-mediated cleavage of RNA

target of an AON•RNA duplex.65,66 On the other hand, it has been found that the inhibition of

translation does not necessarily occur with many antisense oligonucleotides, which do not recruit RNase

H despite their high binding affinity to the target RNA.18,20,67,68 Contrary, naturally occurring AONs

with lower Tm were very effective in inhibition of gene expression through RNase H mechanism. It

appears that irreversible degradation of mRNA by RNase H can amplify the efficacy of AONs

compared to those which are only able to arrest translation by steric blockage of mRNA via the

formation of AON•RNA duplex (RNase H-independent mechanism).69 For instance, 2'-O-alkyl,70 2'-

11

aminoalkyl,71,72 2'-fluoro-modified73 AONs, as well as locked nucleic acids (LNA)74 hybridizes to RNA

resulting in significantly stabilized but not RNase H cleavable AON•RNA duplexes. The same negative

result has been observed for AON phosphodiester backbone modifications as in

methylphosphonates,9,75 phosphoro-N-morpholidates,76 phosphoro-N-butylamidates,76

methylenemethyl-imines,77 and N3'→P5' phosphoramidates.78

To date, only few examples of modified AONs have been shown to support RNase H hydrolysis,

among of which are backbone modified phosphorothioates (PS)15,67 and boranophosphates79 and sugar

modified 2'-deoxy-2'-fluoro-arabinonucleic acids (2'F-ANA).80-82 Unfortunately, such modifications

were accompanied by lowering of AON•RNA duplex stability and, as a consequence, led to the less

RNase H cleavage efficiency compared to the native counterpart.80 To improve AON•RNA duplex

stability without loss of ability to activate RNase H, chimeric oligonucleotides consisting of the middle

part, which supports RNase H hydrolysis (PO or PS-backbone) and flanked with modified sequences to

provide high binding affinity to RNA (2'-O-methyl-ribonucleic acid, locked nucleic acid (LNA), were

synthesized.83-89 The introduction of one or more oxetane modified thymidine blocks into

oligonucleotide allowed to modulate the cleavage pattern of complementary RNA strand by changing

the location of modification.90-92 Recent work has shown that in the gapmer constructs with 2'-O-methyl

nucleosides, one requires at least a minimum stretch of four or five deoxyresidues89. In oxetane92 and

LNA89 modified AONs, one requires at least a stretch of six deoxy nucleotide residues from the

modified site (cleavage site at the sixth phosphate from the 3'-end). Another approach is based on the

terminal conjugation of intercalating agent which can significantly stabilize AON•RNA duplex without

alteration of conformation inherent in the native DNA•RNA hybrid.93 However, only few reports have

been devoted to the investigation of RNase H activity induced by such conjugates. Acridine,94

psoralen,95 and phenanthroline-derivatized95 AONs have been shown to promote specific RNase H

cleavage when bound to their RNA targets.

1.3 Use of metal ions in the antisense and antigene strategies.

Another attractive mechanism by which the potency of ONs might be increased is to synthesize

derivatives that inactivate their nucleic acid targets directly by either irreversible cross-linkage to the

target or by secondary reaction leading ultimately to the disruption of the target molecule. Photocross-

linking96 and cross-linking by an alkylating electrophile97,98 are among the most widely applied

approaches. Cleavage of target strands either hydrolytically or through redox chemistry was mainly

achieved by metal complexes attached to ONs. In the case of cleavage reactions through backbone

hydrolysis, it is relatively easy to hydrolyze RNA target while the estimated half-life of a

phosphodiester bond in DNA is 200 million years at pH 7 and 25°C – unless metal ions come into play

12

when this process can be accelerated enormously.99 A number of cleaving agents,100 Cu2+-terpyridine,101

Lu3+-,Th3+-,Eu3+-iminodiacetate,102 Eu3+-,103 Dy3+-texaphyrin,104,105 Cu2+-N-(2-

mercaptopropionyl)glycine106 complexes, have been covalently attached to ONs and shown to hydrolyze

RNA target. Among the metal ions which cleave DNA at an unprecedented rate are ions of the

lanthanides and combinations thereof, e.g. Ce4+ and Pr3+.107 The introduction of these ions into ONs

represents a major challenge to preparative chemistry. In the case of oxidative cleavage reactions, the

most widely applied pathway is based on the initial generation of hydroxyl radical from molecular

oxygen by its reduction with transition metal ion and subsequent HO•-radical attack on heterocyclic

bases or ribose residues of nucleic acid (Fenton-type chemistry). Fe2+-EDTA-ON conjugates, pioneered

by Dervan, cleaved ssDNA (or RNA)108,109 as well as dsDNA (after triplex formation)110-112 under mild

conditions. The nuclease activity of Cu+-phenanthroline (Cu+(phen)2), introduced by Sigman, was

aimed at ssDNA,113-115 RNA,116 and dsDNA115,117 by attachment of phen ligand at different positions of

the ON. Once delivered to the target strand(s), some metal complexes covalently linked to ONs were

oxidized to the reactive higher oxidation states with co-reactants, such as H2O2 or another peroxo

compounds. The resulting oxo complexes are able to initiate oxidative cleavage of target molecule as it

was shown for metalloporphyrine-11,118,119 and Fe2+-bleomycin-linked120,121 oligonucleotides.

Polypyridyl oxo complexes of ruthenium, electrochemically or chemically obtained from their

aquaruthenium(II) precursors, cleave dsDNA and RNA122-124 through the guanine base oxidation and

H1' abstraction from the ribose moiety, but no attempts have been undertaken to covalently link this

metal species to ODNs because of their high reactivity. Finally, the cleavage of target nucleic acid has

been observed upon photoactivation of some metal complexes tethered to oligonucleotides125-128:

Excited Rh3+(phi)2(bpy)-ODN conjugates underwent the formation of radical cation centered on phi-

ligand which can, depending on the excitation wavelength, either decompose125 or initiate electron

transfer from the remote GG site of the target sequence. High energy activation (313 nm) led to the

direct DNA cleavage near the complex binding site while low energy activation (365 nm) promoted

long-range electron transfer mediated oxidation of distal GG doublet.126,128 Several Ru2+-complexes

with reduction potential in their excited state above 1.1 V (vs. SCE) have been found to promote DNA

strand breaks via an electron transfer.129-132 However, ODN labeled with one of that complexes

(Ru2+(tap)2(dpp)) surprisingly revealed that the dominant process is not a strand break, but the

formation of a cross-linked ODN•DNA duplex.133 The energy transfer from the photoexcited metal

complex to the molecular oxygen has been also proposed to occur thus generating reactive singlet

oxygen (1O2) species which damage the DNA.134

Much less success has been attained in sequence specific cross-linking of nucleic acids by metal

complexes tethered to oligonucleotides. Numerous functional metal complexes, cisplatin and its various

13

platinum analogs,135 as well as RuIICl2(DMSO)4,136 RuII(bpy)2Cl2,137,138 RuIII(tpy)Cl3,137,139

[RuII(NH3)5Cl]Cl,140,141 showed antitumor activity which was generally accepted as a result of their

covalent binding to dsDNA. The profound spatial disturbance and destabilization of the double-helix

caused by metal complex coordination lead to the inhibition of dsDNA replication. Structural studies

performed on model compounds revealed that N7 of a guanine base is the most preferable coordination

site for metal complex binding.135,139,142-145 Unfortunately, these compounds act on dsDNA with low

selectivity, and the problem of toxicity thus becomes an important factor in the use of such agents. The

only reactive metal complexes covalently linked to oligonucleotides to give sequence specific

interstrand cross-linked products were platinum derivatives, currently used in cancer chemotherapy. In

the early studies, the bifunctional platinum complex was directly introduced to ODN containing

exclusively the pyrimidine bases T and dC or one dG residue.146-149 Apart from non-selective

platination at N3 of different cytosine residues, all platinated species formed were sufficiently reactive

to form intrastrand cross-links or could be inactivated by the reactions with other nucleophiles present

in the reaction medium. Thus, the undesirable toxicity cannot be excluded for such ODN conjugates as

a consequence of the presence of the reactive group. Keeping a monofunctionally bound platinum entity

in a reactive state for cross-linking is still a major challenge. Automated insertion of platinated building

blocks during the ODN solid phase synthesis150,151 solved the problem of selective incorporation of Pt2+

to the ODN sequence but not the problem of side reactions. Ideally, functional metal species have to be

protected and back activated after binding to the target molecule. Recently, some efforts have been

made in this direction when [Pt(NH3)2(X)Cl]+-ODN conjugate (X is a 5'-aminolinker of ODN) was

obtained by acidic substitution of cyclohexylmethylthyminate group to Cl on the last stage of ODN

preparation.152,153 However, the possible occurrence of depurination154 in the acid-mediated (HCl, pH

2.3) exchange process limits this scheme to homopyrimidine ODNs only. The activation of protected

Pt2+-ODN conjugates which is exclusively triggered by the duplex formation with target DNA, has been

observed, but the activation was also accompanied by the complete release of the platinum

complex.155,156

14

2. Present Work.

2.1 Synthesis of the ODNs tethered to phenazine, dipyridophenazine, and Ru2+-

dipyridophenazine chromophores.

In the present work, we have covalently linked two sets of chromophore groups to the terminal or

internucleotide positions of ODNs listed in Table 1 and 2. One set consists of simple planar aromatic

labels phenazine (Pnz) and dipyrido[3,2-a;2',3'-c]phenazine (DPPZ), while in the second set the DPPZ

ligand is coordinated to the Ru2+ ion together with other ancillary polypyridyl ligands in octahedral

geometry. Generally, the conjugation sites have been designed in all cases as follows:

ChromophoreLinker Nucleoside residue or non-nucleosidic analog

R2

R2 = 3'-ODN part or H

R1 = 5'-ODN part or H

R1

Two different strategies have been exploited for the coupling of conjugate groups to ODNs: (i) coupling

of chromophore through a linker to a nucleoside residue or non-nucleosidic analog and subsequent

incorporation of the resulting building block into ODN during the solid phase synthesis, and (ii) post-

synthetic modification of amino-linker functionalized ODN with N-hydroxysuccinimidyl (NHS) ester of

the appropriate carboxyl-containing chromophore.

2.1.1 Synthesis of Pnz labeled ODNs (Paper I)

To label ODNs with Pnz, 2-(N-methylamino)ethanol has been chosen as a linker and coupled first to

Pnz at C2 position by nucleophilic substitution in 9-methyl phenazinium methylsulphate 69 (Scheme 1),

which gave us 2-(N-(2-hydroxyethyl)-N-methyl)aminophenazine 70.53,157 Compound 70 was

converted in the usual way158 to the corresponding phosphoramidite 71 and then coupled via tetrazole

activation159 to the 4',6'-TPDS-protected 1-(3'-deoxy-β-D-psicofuranosyl)uracil nucleoside 73 prepared

in six steps from D-fructose.160 Nucleosides 72 and 73 mimics 2'-deoxyribonucleosides but has an

additional hydroxymethyl functionality at 1'-position which can be used for the attachment of conjugate

and linker groups. The coupling product 74, in which Pnz is connected to the nucleoside unit through

the methyl phosphotriester, was deprotected by treatment with TBAF in THF and then 6'-O-DMT-

blocked (74→75→76, Scheme 1). After 75 was phosphitylated or anchored to the

15

Table 1. Oligonucleotides synthesized for ODN•DNA and ODN•RNA duplex study

Sequence Oligo N

5'-TCCAAACAT-3'

5'-TCCATACAT-3'

1

2

5'-PnzTCCAAACAT-3'

5'-PnzUCCAAACAT-3'

5'-TCCAAACAPnzU-3'

5'-TCCAPnzUACAT-3'

5'-PnzUCCAAACAPnzU-3'

5'-TCCAAACAUPnz-3'

3

4

5

6

7

8

DPPZ-Ln-5'-pTCCAAACAT-3'

5'-TCCAAACATp-3'-Ln-DPPZ

5'-TCCAp-Ln(DPPZ)-pAACAT-3'

5'-TCCAp-Ln(DPPZ)-pACAT-3'

9,10,11 (n=1,2,3)

12,13,14 (n=1,2,3)

15,16,17 (n=1,2,3)

18,19,20 (n=1,2,3)

Phos

phod

iest

er

back

bone

(phen)2Ru2+DPPZ-L2-5'-pTCCAAACAT-3'

5'-TCCAAACATp-3'-L2-DPPZ Ru2+(phen)2

5'-TCCAp-Ln[DPPZ Ru2+(phen)2]-pAACAT-3'

21

22

23

Sequ

ence

I

Phos

phor

othi

oate

bac k

bone

PS-5'-TCCAAACAT-3'

PS-5'-PnzTCCAAACAT-3'

PS-5'-TCCAAACAUPnz-3'

PS-DPPZ-L2-5'-pTCCAAACAT-3'

PS-5'-TCCAAACATp-3'-L3-DPPZ

24

25

26

27

28

DNA targets

for sequence I

5'-CATGTTTGGAC-3'

5'-CATGTATGGAC-3'

29

30

RNA targets

for sequence I

5'-r(CAUGUUUGGAC)-3'

5'-r(ACUCAUGUUUGGACUCU)-3'

5'-r(UAACAUGUUUGGACUCU)-3'

31

32

33

5'-CTTACCAATC-3' 34

L2-5'-pCTTACCAATC-3'

5'-CTTACCAATCp-3'-L2

5'-CTTACp-L2-pCAATC-3'

L2-5'-pCTTACCAATCp-3'-L2

35

36

37

38

Sequ

ence

II

Phos

phod

iest

er

back

bone

MeCN(tpy)Ru2+DPPZ-L2-5'-pCTTACCAATC-3'

5'-CTTACCAATCp-3'-L2-DPPZ Ru2+(tpy)NCMe

5'-CTTACp-L2[DPPZ Ru2+(tpy)NCMe]-pCAATC-3'

X-5'-pCTTACCAATCp-3'-X, X=L2-DPPZ Ru2+(tpy)NCMe

39

40

41

42

DNA target

for sequence II 5'-TGATTGGTAAG-3' 43

16

Table 2. Oligonucleotides synthesized for ODN×DNA•DNA triplex study

Sequence Oligo N

5'-TTCT6CT6CT-3'

44

5'-PnzTTCT6CT6CT-3'

5'-PnzUTCT6CT6CT-3'

5'-TTCT6CT6CPnzU-3'

5'-TTCT5PnzUCT6CT-3'

5'-PnzUTCT6CT6CPnzU-3'

45

46

47

48

49

DPPZ-Ln-5'-pTTCT6CT6CT-3'

5'-TTCT6CT6CTp-3'-Ln-DPPZ

5'-TTCT5Tp-Ln(DPPZ)-pCT6CT-3'

5'-TTCT5Tp-Ln(DPPZ)-pT6CT-3'

50,51,52 (n=1,2,3)

53,54,55 (n=1,2,3)

56,57,58 (n=1,2,3)

59,60,61 (n=1,2,3)

Triplex forming

oligos

(TFOs)

(phen)2Ru2+DPPZ-L2-5'-pTTCT6CT6CT-3'

5'-TTCT6CT6CTp-3'-L2-DPPZ Ru2+(phen)2

5'-TTCT5Tp-L2[DPPZ Ru2+(phen)2]-pCT6CT-3'

62

63

64

Duplex targets

for triplex

formation

5'-GCCAAGA6GA6GACGC-3'

3'-CGGTTCT6CT6CTGCG-5'

5'-GCCAAAAAAGA6GA6GACGC-3'

3'-CGGTTTTTTTCT6CT6CTGCG-5'

65

66

67

68

3-aminopropyl-CPG support,161 the resulting 4'-O-phosphoramidite 77 and Pnz-modified support 78

were used as building blocks in solid phase synthesis, permitting Pnz incorporation to the 5'-, 3'- or

internucleotide positions of the oligo chain (ODNs 4 –7, 46 – 49, Tables 1 and 2). Phosphoramidite 71

(R = CH2CH2CN) was also directly used for 5'-Pnz-conjugation as in ODNs 3, 25, 45.

2.1.2 Synthesis of amino-linkers of different lengths tethered to the glycerol moiety (Paper II)

The DPPZ chromophore including its Ru2+ complexes have been attached to the non-nucleosidic

glycerol moiety through the ethylene glycol linkers of different length. The shortest 3-aminopropyl

linker has been connected to the glycerol moiety by the addition of readily available solketal 79 to

acrylonitrile with subsequent reduction of the adduct 80 with NaBH4 in the presence of CoCl2 to give

finally 3-(alkyloxy)propanamine 81 (Scheme 2).162,163 Longer tri- and penta(ethylene glycol) arms 82

17

Abbreviations used in Table 1 and Table 2

N

N

N

Ru

N

N

N

N

N

CH3

O2+

NH

O

ON

OOPO

O-

O

ON

N N

NH

O

ON

OO

P O

-O

OO

N

NNO

CH3

CH3

NH

O

ON

OO

ON

N

O

PO-O

OHN

N

N

N

NO

NN

NN

Ru

2+

N

N

N

NO

HN

OO

O

O

n

HNO

O

O5'

3'

5'

5'

3'

3'

5'

3'

3'

PnzT =

PnzU =

UPnz =

L1 = L2 (n=2)L3 (n=4) =

DPPZ =

(phen)2Ru2+DPPZ =

MeCN(tpy)Ru2+DPPZ =

18

N

NOH

N

N N

CH3

O

N

N N

CH3

PN(iPr)2

OR

NH

O

ON

OHO

OHOH

NH

O

ON

OO

OOH

Si

SiO

NH

O

ON

OO

P O

O

OCH3O

N

NNO

CH3

Si

SiO

NH

O

ON

OR1O

P O

O

OCH3OR2

N

NNO

CH375: R1 = H, R2 = H76: R1 = DMT, R2 = H

Pnz 70 71: R = CH3 or CH2CH2CN

73

7274

NH

O

ON

ODMTO

P O

O

OCH3O

N

NNO

CH377

78

POCH2CH2CN(iPr)2N

NH

O

ON

ODMTO

P O

O

OCH3O

N

NNO

CH3

HN

O

O

S

N

N

69CH3CH3OSO3

-

Scheme 1. Synthesis of building blocks for Pnz incorporation into ODNs 3 – 7, 25, 45 – 49 (Tables 1 and 2).

and 83 were first mono-tosylated and the tosyloxy groups in the resulting compounds 84 and 85 were

then substituted with phthalimido function to afford the protected precursors 86 and 87. They were

treated with allyl bromide in the presence of NaH affording olefins 88 and 89. Subsequently, double

bond oxidation with OsO4 or KMnO4 gave diols 90 and 91, in order to build the sn-glycerol moiety

19

OHO

O

OO

O

CNO

O

O

NH2

O

DMTO

O

DMTO

POCH2CH2CN(iPr)2N

HN

O

O

S

OHO OR

n

OHO

nN

O

O

OO

nN

O

O

OO

N

O

OR2O

R1O

n

OO

NHR2R1O

DMTO

n

79 8180

82: n = 2, R = H83: n = 4, R = H

84: n = 2, R = Ts85: n = 4, R = Ts

86: n = 2

90: n = 2, R1 = R2 = H91: n = 4, R1 = R2 = H

92: n = 2, R1 = DMT, R2 = H93: n = 4, R1 = DMT, R2 = H

94: n = 2, R1 = R2 = H95: n = 4, R1 = R2 = H

96: n = 2, R1 = H, R2 = Fmoc

OO

NHFmoc

2

OO

NHFmoc

297 98

87: n = 488: n = 289: n = 4

Scheme 2. Preparation of linkers for conjugation with dipyridophenazine (DPPZ) containing chromophores (including building blocks 97 and 98 which were directly used for the synthesis of ODNs 35 – 38, Table 1). connected to the linker arms.164 DMT-protection of primary hydroxyl in these diols with subsequent

removal of phthalimido group by refluxing with hydrazine165 gave finally sn-glycerol conjugated

aminolinkers 94 and 95. The amino functionalized compounds 81, 94, and 95 were then used for

condensation with carboxyl derivatives of DPPZ and its Ru2+ complexes. Moreover, 94 was also

utilized for the preparation of amino-linkertethered ODNs 35 – 38 (Table 1) which were subsequently

subjected for the post-synthetic labeling reactions. For this purpose, amino group in 94 was Fmoc-

protected to give compound 96, which was converted into O-(2-cyanoethyl)(N,N-

diisopropyl)phosphoramidite block 97.158 The remaining part of 96 was succinylated and the

corresponding succinate was attached166 to the 3-aminopropyl-CPG support to provide 49 µmol of

bound 96 per gram CPG (98).

20

N

N

N

NO

NN

NN

RuOH

2+

N

N

N

NO

OH

N

N

N

N O

O

99 (phen) 100 101 (DPPZ-COOH)

RuCl3 Ru(phen)2Cl2

102

99(i) 100

2 PF6-

103

H2N

H2NO

OH

N

N

N

Ru

N

N

N

N

COOH

Cl

+

N

N

N

Ru

N

N

N

N

COOH

N

2+

CH3

O

O

N

N

NN

NN

Ru

2+ 2 PF6-

2 Cl-Cl-Ru(tpy)Cl3

terp

yrid

ine

101

104 105

(ii) NH4PF6

Scheme 3. Preparation of dipyridophenazine (DPPZ) containing chromophores for conjugation.

2.1.3 Derivatization of DPPZ containing chromophores for conjugation (Papers II, IV & V)

To be able to attach DPPZ and its Ru2+ complexes to the amino-linker containing blocks 81, 94, 95

(Scheme 2) or to the aminolinker modified ODNs 35 – 38 (Table 1), we prepared dipyrido[3,2- a;2',3'-

c]phenazine-11-carboxylic acid 101 in two steps from 1,10-phenanthroline 99 by its oxidation to 1,10-

phenanthroline-5,6-dione 100 and the following condensation with 3,4-diaminobenzoic acid in ethanol

(Scheme 3).167 Next, we coordinated DPPZ-COOH ligand 101 around Ru2+ center to obtain

tris(polypyridyl) ruthenium complex [Ru(phen)2(dppz-COOH)]2+ 103 as well as bis(polypyridyl)

ruthenium complexes, such as [Ru(tpy)(DPPZ-COOH)Cl]+ 104 and [Ru(tpy)(DPPZ-

COOH)(CH3CN)]2+ 105 containing monodentate ligands (Cl, CH3CN). Common synthetic approaches

to the preparation of complexes of general formulas [Ru(NN)2(NN)']2+ and [Ru(NNN)(NN)X]n+ (NNN

21

and NN denote tri- and bidentate polypyridyl ligands respectively, X is Cl or CH3CN) have been

applied in the synthesis of 103 - 105. Ru(phen)2Cl2167 was coordinated with ligand 100 in refluxing

ethanol-water and the product 102 was condensed with 3,4-diaminobenzoic acid to give finally complex

103.168 In a similar manner, complex 104 was formed starting from Ru(tpy)Cl3169 by coordination with

DPPZ-COOH (101) in refluxing ethanol-water in the presence of LiCl and triethylamine.170 In this

reaction triethylamine functions as a reducing agent to help the dissociation of Cl from Ru(tpy)Cl3,

whereas LiCl is used to prevent any dissociation of Cl from the product. The Cl ligand was then

readily replaced in 104 by CH3CN ligand upon heating under reflux in acetonitrile-water mixture171,172

to afford complex 105.

2.1.4 Synthesis of DPPZ and Ru2+(phen)2DPPZ labeled ODNs (Papers II & IV)

Carboxyl-derivatized DPPZ chromophore 101 (Scheme 3) was subjected for the condensation with the

amines 81, 94, and 95 (Scheme 2) using N,N'-carbonyldiimidazole173 as a condensing agent in pyridine

under heating (80ºC) which resulted in the formation of corresponding amides 106, 111, and 112

(Scheme 4). These compounds were highly contaminated with side products arising from heating of the

reaction mixture but poor solubility of 101 in any organic solvent prevented utilization of other more

soft amide-forming strategies. The DPPZ derivative 106 was then treated with 1M HCl / THF (1:1, v/v)

to remove isopropylidene protection, and the crude product 107 was tritylated to give 108. Finally,

compounds 108, 111, and 112 were converted in the usual manner158 to the corresponding

phosphoramidites 109, 113, and 114 as well as immobilized161 onto 3-aminopropyl-CPG support

(110, 115, 116). The building blocks thus prepared enabled incorporation of the DPPZ chromophore

through the linkers of different lengths at any desired position of an oligo sequence as in ODNs 9 –

20, 27 – 28, and 50 – 61 (Tables 1 and 2) by solid phase synthesis. The present ODN conjugates are the

first examples of oligonucleotides tethered with DPPZ chromophore.

Analogously, hexafluorophosphate salt of Ru2+ complex 103 (Scheme 3) was treated with N,N'-

carbonyldiimidazole174 in pyridine at room temperature and the intermediate acylimidazole was

quenched with amine 94 (Scheme 2) in one pot giving compound 117 (Scheme 4). Next, 2-

cyanoethylchloro-(N,N-diisopropyl)phosphoramidite was reacted with 117 in dry CH3CN175 to afford

metallo-phosphoramidite 118. After drying, 118 was used in the solid phase synthesis of 5'-Ru2+-ODN

conjugates 21, 62, and internucleotide conjugated Ru2+-ODNs 23, 64 (Tables 1 and 2). Compound 117

was also attached161 to the 3-aminopropyl-CPG to assemble ODNs modified with [Ru(phen)2DPPZ]2+-

moiety at the 3'-terminal as in 22 and 63 (Tables 1 and 2). Thus, the novel type of attachment through

the DPPZ moiety of [Ru(phen)2DPPZ]2+ complex has been implemented for its ODN conjugation.

22

OO

OHN DPPZ

O

OHO

ROHN DPPZ

O

OO

DMTOHN DPPZ

O

OO

DMTOHN DPPZ

OPOCH2CH2CN(iPr)2N

HN

O

O

S

N

N

N

NO

OH

101 (DPPZ-COOH)

81

106 107: R = H108: R = DMT

OO

HN

HO

DMTO

n

DPPZ

O

94

95or

110

109

111: n = 2112: n = 4

OO

HN

O

DMTO

n

DPPZ

O113: n = 2114: n = 4

POCH2CH2CN(iPr)2N

OO

HN

O

DMTO

n

DPPZ

O115: n = 2116: n = 4

HN

O

O

SN

N

N

NO

NN

NN

Ru OH

2+ 2 PF6-

103

OO

HN

HO

DMTO

2

DPPZ Ru2+(phen)2

O

94117

OO

HN

O

DMTO

2

DPPZ Ru2+(phen)2

O118

OO

HN

O

DMTO

2

DPPZ Ru2+(phen)2

O119

POCH2CH2CN(iPr)2N

HN

O

O

S

Scheme 4. Synthesis of building blocks for DPPZ and (phen)2Ru2+DPPZ incorporation into ODNs 9 – 23, 27 - 28, and 50 – 64 (Tables 1 and 2).

23

2.1.5 Synthesis of Ru2+(tpy)(DPPZ)(CH3CN) labeled ODNs (Paper V)

At present time, incorporation of metal complexes into ODNs is generally restricted to the linking of

only substitution-inert polypyridyl complexes of Ru2+ 133,134,174-190 and Rh3+.126,127,191,192 One of the

biggest advantages of utilizing such complexes is the fact that their chelation chemistry is less

demanding than that of the functional metal species, and can be compatible both with the synthesis of

ODN building blocks and post-synthetic labeling strategy. Contrary, conjugation of functional metal

complexes requires the use of protection ligand which should be chemically and thermally stable on one

hand, and on the other hand, it should be readily substituted onto a labile ligand under specific

conditions without altering of the ODN part. In the course of our attempts to synthesize

[Ru(tpy)(DPPZ)(H2O)]2+-ODN conjugates, we used [Ru(tpy)(DPPZ)(X)]n+-precursors (X = Cl,

CH3CN) in which X can be thermally (X = Cl)171 or photolytically (X = CH3CN)172,193,194 replaced by

aqua ligand. Due to the thermal dissociation of Cl ligand in [Ru(tpy)(DPPZ-COOH)Cl]+ complex 104

(Scheme 3) occurring in aqueous solutions, its conjugation is not compatible with the conditions

normally applied for the post-synthetic deprotection of the synthesized ODN (conc. aq. NH3, 55ºC) and

subsequent HPLC purification. Replacement of Cl¯ by NH3 during deprotection step as well as by

CH3CN or other coordinating ligands present in HPLC buffers leads to the mixture of conjugates in

which different monodentate ligands are bonded to the Ru2+ center. On the other hand, Meyer at

al.195,196 reported on the incorporation of [Ru(tpy)(bpy')Cl]+ (bpy' = 4-carboxy-4'-methyl-2,2'-

bypyridine) into polymers and peptides indicating that Cl remains unaltered under conditions of

condensation in anhydrous solutions, although no clear spectroscopic evidence was given. This

prompted us to prepare anchored to the solid support building block 123 (Scheme 5A) and use it in

automated solid phase ODN synthesis with subsequent substitution of Cl ligand in ODN-conjugated

Ru2+ complex by thermally stable CH3CN ligand after mild deprotection with conc. aq. NH3 at room

temperature. According to Scheme 5A, the key compound 122 was prepared through either amide bond

formation (as in 104→124→122) or through the coordination of compounds 111 or 120 to Ru(tpy)Cl3

(as in 111→122 or 120→121→122). The amide bond formation has been accomplished by

conversion of a chloride salt of 104 to its NHS-ester 124 upon treatment with TSU in dry DMF.133 The

activated complex 124 was then reacted with amine 94 (Scheme 2) in dry DMF and chloride salt of the

product was metathesized to the hexafluorophosphate salt 122 by treatment with excess ethanolic

NH4PF6. Alternatively, complex 121 was formed by coordination of Ru(tpy)Cl3 with the functionalized

DPPZ ligand 120 in boiling ethanol-water (120 was obtained by deprotecton of 111 with aqueous 80%

acetic acid). The presence of reducing agent triethylamine in the reaction between 120 and Ru(tpy)Cl3

24

N

N

N

Ru

N

N

N

N

COOH

Cl

+

N

N

N

Ru

N

N

N

N

COOH

N

2+

CH3

2 Cl-

Cl-

Ru(tpy)Cl3

104

105

OO

HN

HO

RO

2

DPPZ

O120: R = H111: R = DMT

OO

HN

HO

RO

2

DPPZ Ru2+(tpy)Cl

O

121: R = H122: R = DMT

PF6-

N

N

N

Ru

N

N

N

N

Cl

+ Cl-

124

OO

NO

O

OO

HN

O

DMTO

2

DPPZ Ru2+(tpy)Cl

OPF6

-

HN

O

O

S

(i) 94

123

N

N

N

Ru

N

N

N

N

125

OO

NO

O

N

CH3

2+ 2 Cl-

Oligonucleotides 39 - 42

OO

NH2R2O

R1O

235: R1 = H, R2 = pCTTACCAATC-3'36: R1 = 5'-CTTACCAATCp, R2 = H37: R1 = 5'-CTTACp, R2 = pCAATC-3'38: R1 = H, R2 = pCTTACCAATCp-3'-CH2CH(OH)((OCH2CH2)3NH2)

A

B

(ii) NH4PF6

Scheme 5. (A) Synthesis of solid support for the 3'-incorporation of MeCN(tpy)Ru2+DPPZ moiety (ODN 40, Table 1). (B) Post-synthetic labeling of ODNs 35 – 38 for MeCN(tpy)Ru2+DPPZ incorporation at different terminals (ODNs 39 – 42, Table 1).

25

enabled us to similarly modify DMT-protected analog 111 and prepare compound 122.

Correspondingly, both 121 and 122 were isolated as PF6 salts to ensure their solubility in organic

solvents. Complex 121 was also shown to produce 122 upon treatment with DMT-Cl in dry pyridine at

room temperature. Attachment of 122 to 3-aminopropyl-CPG allowed to prepare [Ru(tpy)(dppz)Cl]+-

modified solid support and use it for automated assembly of 3'-[Ru(tpy)(dppz)Cl]+-ODN conjugate. The

fast deprotecting 5'-O-DMT-N4-isobutyryl-2'-deoxycytidine-3'-O-(β-cyanoethyl-N,N-diisopropyl)- and

5'-O-DMT-N6-phenoxyacetyl-2'-deoxyadenosine-3'-O-(β-cyanoethyl-N,N-diisopropyl)-phosphor-

amidites197,198 utilized in this ODN synthesis enabled to apply mild ammonia deprotection at room

temperature which does not effect the Cl displacement. This was proven by us on a model

[Ru(tpy)(dppz)Cl](PF6) complex. After removal from solid support and lyophilization, the crude

material was incubated in CH3CN-H2O (1:1, v/v) at 55ºC for 17 h to substitute Cl to the

thermally stable CH3CN at the metal center to give 3'-[Ru(tpy)(dppz)(CH3CN)]2+-ODN conjugate 40

(Table 1). Unfortunately, our attempts to prepare metallo-phosphoramidite from compound 122 and use

it for 5'- or internucleotide incorporation failed. This can be explained by the possible displacement of

Cl¯ during interaction between the Ru2+ and phosphite center which accompanies the phosphitilation of

OH group.199,200

Contrary to the [Ru(tpy)(dppz)Cl]+, its Ru2+-CH3CN analog undergoes base hydrolysis201 of the

coordinated CH3CN ligand to acetamide which is released very rapidly from the coordination sphere of

Ru2+, since amides are poor π-acceptor ligands.202 Such decomposition excludes the use of Ru2+-

CH3CN species in the solid phase synthesis, requiring removal of the ODN from solid support and

deprotection with conc. aq. NH3. Hence, the [Ru(tpy)(DPPZ-COOH)(CH3CN)]2+ complex 105 (Scheme

3) was introduced to ODN by amide bond formation between NHS-ester 125 and aminolinker-

functionalized ODNs 35 – 38 (Scheme 5B). For this purpose, complex 105 was activated analogously

as in 104→124 step (Scheme 5A) to give 125 which was incubated with appropriate ODN in 33%

CH3CN / 10 mM sodium tetraborate (pH 8.5) buffer and the product was separated from the excess of

Ru2+ complex by cation exchange column chromatography. This post-synthetic modification has been

proven to be valuable for incorporation of metallating species containing sensitive functionality, as in

ODNs 39 – 42 (Table 1). The methodology described here shows for the first time that mono-

acetonitrile polypyridyl Ru2+ complex tethered to a oligodeoxynucleotide (as in ODNs 39 – 42) can be

constructed by post-synthetic labeling of amino-linker modified ODN, thus opening the way to design

new class of metal-ODN conjugates carrying photochemically reactive metal entity.

26

2.1.6 Characterization of Ru2+(tpy)(DPPZ)(CH3CN) labeled ODNs (Paper V)

The main structural feature of the newly synthesized ODNs 39 – 42 (Table 1) tethered with

monofunctional Ru2+(tpy)(DPPZ)(CH3CN) complex is that they carry photolabile CH3CN ligand in

contrast to other Ru2+- 133,134,174-190 or Rh3+-conjugated126,127,191,192 ODNs prepared till present and

characterized by inertness towards the ligand substitution reactions in the metal coordination sphere. To

show the presence of the sensitive functionality, ODNs 39 – 42 have been carefully characterized by

enzymatic digestion nucleobase analysis, PAGE, as well as by UV-vis and MS-spectroscopy. All

metallated ODNs exhibited the low-energy metal-to-ligand charge transfer (MLCT Ru2+→CH3CN)

bands at 453 nm and DPPZ intraligand transition at 376 nm identically with those for the not tethered

monomer block [Ru(tpy)(DPPZ-CO-L2)(CH3CN)]2+ (L2 is a linker, see page 17).172,194,201 The

substitution of CH3CN by other monodentate ligands leads to the shift of the MLCT band as has been

shown for [Ru(tpy)(DPPZ-CO-L2)X]2+ (X = Cl, py) complexes. The A260/A453 absorption ratio for the

ODN conjugates was found to be higher (9.1) then the corresponding ratio for the not tethered Ru2+-

CH3CN complex (3.5) reflecting the contribution of the DNA bases at 260 nm in the ruthenated oligos.

PAGE analysis revealed that the metal complex-ODN conjugates have less electrophoretic mobility

compared to those of the amino-linker L2 modified counterparts (ODNs 35 – 38, Table 1). This is

owing to the increase of molecular weight (685 dalton per [Ru(tpy)(DPPZ-CO-)(CH3CN)]2+ residue)

and the decrease of the overall negative charge by a factor of 2 in the Ru2+-ODNs. The base

composition as well as the presence of tethered Ru2+ complex was further confirmed by the degradation

of ODN conjugates to nucleosides upon treatment with a mixture of snake venom phosphodiesterase

(SVDP) and alkaline phosphotase (AP).203 The identity of nucleoside peaks in the digest mixtures was

proven by comparison of their retention times with those of the authentic mixture of dC, dT, and dA

nucleosides. The ratio of normalized areas under nucleoside peaks in the digest of mono-Ru2+-labeled

ODNs was found 4.1 : 3 : 2.9 which is close to the nucleoside composition in the native ODN 34 (Table

1). On the other hand, the appearance of two strongly retarded peaks in the HPLC profile of the digest

mixtures was assigned to the Ru2+ species. We suspect that they are the products of the hydrolysis of

coordinated CH3CN ligand under digestion conditions (pH 9.0)201 and subsequent acetamide

substitution onto other entering groups existing in the enzymatic coctail.

The final proof of the identity of [Ru(tpy)(DPPZ)(CH3CN)]2+-ODN conjugates has been obtained

by MALDI-TOF and ESI-mass spectrometry. The development of these soft ionization techniques

enabled us to characterize both natural and synthetic oligonucleotides at the structural level.204-208 The

negative ion mode ESI MS of the mono-metallated ODNs 39 – 41 (calculated MW 3916 in the fully

phosphodiester protonated and 2+ charged state) and bis-metallated ODN 42 (calculated MW 4885 for

the 4+ charged state) showed a number of multiply charged ions, which after deconvolution, revealed

27

the expected ODN anions in the range of mass/charge (m/z) = 3912.5 – 3913.8 for the ODNs 39 – 41

and 4880.9 for the ODN 42 corresponding respectively to [M-3H+] and [M-5H+] anions. From this

data one can conclude that all conjugates have the CH3CN ligand intact in the coordination sphere of

their Ru2+-label. On the other hand, when the ODNs were ionized in the negative mode by laser pulse

(MALDI-TOF MS), the observed mass/charge (m/z) ratio were in the range of 3871.9 – 3872.9 for the

single-modified ODNs 39 – 41 in the 1- charge state. The difference between the m/z values obtained

from MALDI and ESI ionizations for the single Ru2+ modified ODNs 39 – 41 is approximately 41 Da

and expected from the photolabilization of CH3CN ligand occurring during the laser irradiation.

Similarly, laser ionization of the 5',3'-bis-Ru2+- modified ODN 42 led to the loss of two acetonitrile

ligands affording the 1- charged ions with m/z of 4798.4: M[(C186H208N51O74P11102Ru2)4+]-M[2CH3CN

+ 5H+]. Thus, MS-analysis of the Ru2+-ODN conjugates established the presence of the covalently

linked [Ru(tpy)(DPPZ)(CH3CN)]2+ monofunctional complex in the ODN chain. Comparison of the MS

data obtained on the bases of use of the different ionization techniques clearly showed that synthesized

ODN conjugates are photochemically active and undergo the expected CH3CN ligand decomposition

under light irradiation.

2.2 Thermal stability of duplexes and triplexes formed by phenazine, dipyridophenazine,

and Ru2+-dipyridophenazine conjugated ODNs.

2.2.1 Thermal stability of conjugated ODN•DNA duplexes (Papers I, II, IV & V)

Different chromophore groups tethered to natural ODNs are expected to influence the hybridization

properties of the resulting ODN conjugates with the complementary DNA and RNA molecules. The

analysis of melting temperatures of differently conjugated ODN•DNA duplexes shows (Table 3) that

Pnz chromophore tethered at the 1'-position of uridine moiety (PnzU) through the relatively short linker

has the least ability to stabilize a duplex. The stabilization was only achieved when the Pnz-tethered PnzU was attached at the 5'-terminal (∆Tm = 6.1°C) while 3'-conjugation gave slight destabilization (∆Tm

= -0.9°C) and middle-conjugation led to almost complete loss of stability (∆Tm = -12.9°C) of the

duplex. Double Pnz-modification at both 5'- and 3'-termini gave also positive ∆Tm value (2.5°C), albeit

lower than for single 5'-Pnz-conjugation, probably as a result of negative influence of 3'-Pnz-tether.

Interestingly, the same type of Pnz-tethering at 5'-hydroxyl of a 5'-terminal T (PnzT as in the duplex

3•29) allowed to strengthen the stabilization effect (∆Tm = 10.0°C) reflecting better positioning of a

stacker when it is delivered from the 5'-hydroxy terminal of the duplex. Contrary, DPPZ conjugation

allowed us to stabilize ODN•DNA duplexes more effectively. As expected, the stabilization effect of

the DPPZ chromophore depended on the linker length as well as on the position of the linker

attachment to the ODN. As a whole, 5'- and 3'-DPPZ-conjugation lead to comparably equal

28

Table 3. Thermal stability of ODN•DNA duplexes studied#

Native duplexes

5'-TCCAAACAT-3' 5'-TCCATACAT-3' 5'-CTTACCAATC-3'

3'-CAGGTTTGTAC-5' 3'-CAGGTATGTAC-5' 3'-GAATGGTTAGT-5'

1•29,$ Tm = 25.9

2•30,

Tm = 22.4

34•43, Tm = 33.0

Internucleotide 5',3'-bis-

5'- modification

3'- modification modification

modification

PnzT 3•29, 35.9(+10.0)

UPnz 8•29, 31.6(+5.7)

Pnz

- con

juga

tion

PnzU

4•29, 32.0(+6.1)

5•29, 25.0(-0.9)

6•30, 9.5(-12.9)

7•29, 28.4(+2.5)

L1 - linker 9•29, 33.5(+7.6)

12•29, 34.3(+8.4) 15•29, 29.0(+3.1) 18•29, 14.7(-11.2)

L2 - linker 10•29, 36.8(+10.9)

13•29, 33.2(+7.3) 16•29, 30.1(+4.2) 19•29, 14.6(-11.3)

DPP

Z - c

onju

gatio

n

L3 - linker 11•29, 33.2(+7.3)

14•29, 35.6(+9.7) 17•29, 26.4(+0.5) 20•29, not formed

(phen)2Ru2+DPPZ - conjugation

(L2 - linker)

21•29, 47.9(+22.0)

22•29, 38.7(+12.8)

23a•29, 42.3(+16.4) 23b•29, 45.3(+19.4) 23c•29, 39.5(+13.6) 23d•29, 40.2(+14.3)

MeCN(tpy)Ru2+DPPZ -conjugation

(L2 - linker)

39•43, 50.5(+17.5)

40•43, 41.4(+8.4)

41•43, 48.3(+15.3)

42•43 56.6(+23.6)

L2-linker conjugation

35•43, 34.5(+1.5)

36•43, 34.4(+1.4)

37•43, 24.0(-9.0)

38•43 35.5(+2.5)

# Tm(∆Tm) in ºC are shown after numbers of duplex strands (symbol • stands for Watson-Crick hydrogen bonding between duplex strands). ∆Tm = Tm(modified duplex) - Tm(corresponding native duplex) $ For sequence composition of oligos see Table 1 (page 15)

increase of duplex stability with the maximal values observed for 5'-tethering through the 12-atom

linker L2 (∆Tm = 10.9°C) and for 3'-tethering through the 18-atom linker L3 (∆Tm = 9.7°C).

Compared to these values, covalent linkage of DPPZ to the 5'-terminal through the linkers L1 and L3 as

well as 3'-DPPZ-conjugation through the linkers L1 and L2 had slightly less stabilizing effect.

Conversely, internucleotide DPPZ incorporation induced only marginal increase of stability with ∆Tm

varying from 0.5 to 4.2°C depending on the linker length. The relatively positive increase of affinity

of middle-DPPZ tethered ODNs 15 – 17 toward DNA target reflects the possible chromophore

intercalation which compensates the strong destabilizing influence arising from the insertion of a linker

29

between internal dA residues of natural ODN 1. Such linker insertion structurally increases the distance

between the neighboring dAs which should lead to bulge formation in the middle-modified duplexes to

provide Watson-Crick paring of given dAs with complementary nucleobases.34 This strongly disrupts

the helix as is clearly seen from the introduction of a 12-atom linker L2 into the interior position of a

duplex as in 37•43 (∆Tm = -9.0°C). Note that linker L2 externally tethered at 5'-, 3'- or both terminals

does not have a great influence on the hybridization affinity of ODNs 35, 36, 38 as is seen from the

thermal stability of the corresponding duplexes (∆Tm = 1.4 – 2.5°C). In order to exclude the bulge

formation effected by internal insertion of DPPZ unit, we prepared duplexes 18•29, 19•29, and 20•29,

in which the DPPZ-linker units were introduced instead of the central dA nucleotide of the parent

sequence 1, with the expectation that the DPPZ chromophore would place the opposite T base of the

complementary target outside of the helix.34 Experimental results showed that duplexes so formed are

very unstable indicating appearance of a mismatch pair DPPZ-T. In general, more efficient

enhancement of hybridization affinity of DPPZ-ODN conjugates towards DNA compared to the Pnz-

tethered analogs can be explained on the basis of the following considerations: (1) DPPZ itself has more

ability for intercalation because of its more extended planar surface. (2) The use of flexible linkers

attaching the chromophore to the non-nucleosidic glecerol residue of ODN sequence provides the

delivery and optimal orientation for the stacker more effectively.

When coordinated to Ru2+, as in Ru2+(DPPZ)-labeled ODNs, the intercalating ability of DPPZ is

modulated by other ancillary ligands. It was shown earlier that positively charged polypyridyl Ru2+

complexes can bind to dsDNA electrostatically209,210 as well as through hydrophobic interactions in

DNA grooves211 or intercalation by one of the extended ligands.209,212,213 Because in our Ru2+(DPPZ)-

ODN conjugates the DPPZ ligand is sterically hampered by covalent attachment of a linker arm L2 (see

abbreviations on page 17) we first evaluated the ability of complex [Ru(phen)2(DPPZ-CO-L2)]2+ to

intercalate into the native duplex 1•29 by comparison with binding affinity of typical metallointercalator

[Ru(phen)2DPPZ]2+ and groove-binder [Ru(phen)3]2+.214-217 The results of titration of duplex 1•29 with

Ru2+ complexes clearly showed that our linker-tethered Ru2+ complex and [Ru(phen)2DPPZ]2+ behave

similarly, sharply increasing duplex stability upon increasing Ru2+ concentration up to 3 – 4 equiv. per

duplex after which Tm increase becomes slow. This trend is general for intercalation of metal complexes

taking place until all intercalation sites are saturated. After this the stabilization is due to electrostatic

binding, and Tm increases less steeply. On the contrary, growing amount of [Ru(phen)3]2+ does not

practically result in the duplex stabilization.129 Thus, the [Ru(phen)2(DPPZ-CO-L2)]2+ binding should

proceed via intercalation of DPPZ ligand despite its derivatization with a long linker arm.

In conformity with the above data, tethering of the [Ru(phen)2DPPZ]2+ and

[Ru(tpy)(DPPZ)(CH3CN)]2+ complexes to ODN through the linker L2 dramatically enhanced the

30

hybridization affinity of the corresponding conjugates towards target DNA (Table 3). The magnitude of

duplex stabilization for various site-specific [Ru(tpy)(DPPZ)(CH3CN)]2+ incorporations increases as

follows: 3'-Ru2+(∆Tm = 8.4°C) < middle Ru2+(∆Tm = 15.3°C) < 5'-Ru2+(∆Tm = 17.5°C) < 5',3'-bis-

Ru2+(∆Tm = 23.6°C). The same trend was also found for the [Ru(phen)2DPPZ]2+-ODNs with ∆Tm

varying from 12.8°C to 22.0°C. The absolute ligand configuration at the Ru2+ center of the separated

middle-[Ru(phen)2DPPZ]2+-ODN conjugates plays an essential role in the duplex formation. It has been

observed that Λ-Ru2+ tethered complexes (as in duplexes 23a•29 and 23b•29) gave higher stabilization

(∆Tm = 16.4°C and 19.4°C respectively) while ∆-Ru2+ tethered stereoisomers (as in duplexes 23c•29

and 23d•29) yielded relatively poorer stabilization (∆Tm = 13.6°C and 14.3°C respectively). The

stabilization effect gained thus by attachment of metal complexes considerably exceeds those observed

with DPPZ- and Pnz-ODN conjugates irrespective of the attachment site. It is also noteworthy that the

best stabilization achieved before this work was 3°C134 and 8°C181 for 5'-tethered [Ru(phen)2DPPZ]2+

when the phen ligand was used for covalent attachment, while for other types of Ru2+-ODN

conjugates133,182,184,185 no improvement in the stabilization has been observed.

2.2.2 Thermal stability of conjugated ODN•RNA duplexes (Papers II & IV)

The DPPZ- and [Ru(phen)2DPPZ]2+-labeled ODNs, prepared in this work, were also tested (Table 4)

for their ability to act as antisense ONs to form stable duplexes with complementary RNA target.

Generally, all ODN•RNA duplexes, including the native counterpart, were less stable than the

corresponding ODN•DNA analogs as well as the stabilizing effect of intercalator was less pronounced

on ODN•RNA hybrids compared to the ODN•DNA counterparts. Analogously to (DPPZ-ODN)•DNA

duplexes (Table 3), 12-atom linker L2 provided maximal stability for 5'-DPPZ-tethered ODN•RNA

hybrids (∆Tm = 5.4°C for 10•31 in Table 4) while 3'-DPPZ-ODN conjugates showed greater affinity

towards RNA in case of chromophore linking through the 18-atom linker L3 (∆Tm = 7.4°C for 14•31).

The thermal stability of ODN•RNA duplexes internally modified with DPPZ was comparable with that

of the native duplex 1•31, indicating that destabilization induced by the bulge formation is compensated

by the stabilizing impact of DPPZ moiety within ODN•RNA helical structure as it was found for the

middle-DPPZ-modified ODN•DNA duplexes.

The advantage of [Ru(phen)2DPPZ]2+-conjugation for the aim of ODN•RNA hybrid stabilization

was not so remarkable as for ODN•DNA duplexes. Interesting to note that the effect of metal

conjugation at the 3'-terminal of both types of duplexes was almost equal [∆Tm = 7.9°C (Table 4) and

12.8°C (Table 3) for ODN•RNA 22•31 and ODN•DNA 22•29 duplexes respectively]. For the 5'-

31

Table 4. Thermal stability of DNA•RNA duplexes studied#

Native duplex

5'-TCCAAACAT-3'

3'-r(CAGGUUUGUAC)-5'

1•31,$ Tm = 20.6

Internucleotide

5'- modification

3'- modification modification

L1 - linker 9•31, 25.2(+4.6)

12•31, 27.1(+6.5) 15•31, 19.2(-1.5) 18•31, not formed

L2 - linker 10•31, 26.1(+5.4)

13•31, 25.2(+4.5) 16•31, 21.5(+0.9) 19•31, not formed

DPP

Z - c

onju

gatio

n

L3 - linker 11•31, 25.5(+4.9)

14•31, 28.0(+7.4) 17•31, 21.1(+0.4) 20•31, not formed

(phen)2Ru2+DPPZ - conjugation

(L2 - linker)

21•31, 29.7(+9.1)

22•31, 28.5(+7.9)

23a•31, 20.6(+0.0) 23b•31, 21.6(+1.0) 23c•31, 20.1(-0.5) 23d•31, 20.1(-0.5)

# Tm(∆Tm) in ºC are shown after numbers of duplex strands (symbol • stands for Watson-Crick hydrogen bonding between duplex strands). ∆Tm = Tm(modified duplex) - Tm(corresponding native duplex) $ For sequence composition of oligos see Table 1 (page 15)

terminal conjugation the difference in stabilization effect was strikingly higher [∆Tm = 9.1°C (Table 4)

and 22.0°C (Table 3) for ODN•RNA 21•31 and ODN•DNA 21•29 duplexes respectively], much more

than the corresponding difference between (DPPZ-ODN)•RNA and (DPPZ-ODN)•DNA duplexes

(compare ∆Tms for 9•31 - 17•31 with those for 9•29 - 17•29). Finally, the ODN•RNA duplexes carrying

metal complex at the internucleotide position (as in stereoisomeric 23a-d•31, Table 4) were as stable as

the native counterpart 1•31 as opposed to very stabilized (middle-[Ru(phen)2DPPZ]2+-ODN)•DNA

duplexes 23a-d•29 (Table 3). This indicates that interactions of the bulky Ru2+ complex with duplex

nucleobases are sensitive to the duplex conformation, while the planar DPPZ moiety can be more easily

tolerated by both DNA•DNA and DNA•RNA helical structures.

2.2.3 Thermal stability of conjugated ODN×DNA•DNA triplexes (Papers I, II & IV)

All triplex forming ODN 18mer conjugates (TFOs) consisted of pyrimidines T and dC with the

expectation of Hoogsteen base pair formation in a parallel orientation to the complementary purine-rich

strand of a duplex target.11,218 It can be seen (Table 5) that conjugation of the different chromophores to

the native ODN 44 had no or much less stabilization effect compared to the influence of chromophore

conjugation on the duplex formation. Pnz-modified uridine (PnzU) nucleotide when placed at the 5'-end

32

Table 5. Thermal stability of ODN×DNA•DNA triplexes studied#

Native triplexes

5'-TTCT6CT6CT-3' 5'-TTCT6CT6CT-3'

5'-GCCAAGA6GA6GACGC-3' 5'-GCCAAAAAAGA6GA6GACGC-3'

3'-CGGTTCT6CT6CTGCG-5' 3'-CGGTTTTTTTCT6CT6CTGCG-5'

Tm = 13.5 (pH 7.3) Tm = 13.7 (pH 7.3) 44×65•66,$

Tm = 21.5 (pH 6.5) 44×67•68, Tm = 16.8 (pH 6.5)

Tm = 25.6 (pH 6.0)

Tm = 19.3 (pH 6.0) Internucleotide 5',3'-bis-

5'- modification

3'- modification modification

modification

pH 7.3 45×65•66, 17.4(+3.9) pH 6.5 45×65•66, 24.3(+2.8)

PnzT pH 6.0 45×65•66, 30.3(+4.7)

pH 7.3

46×65•66, 17.5(+4.0) 46×67•68, 20.4(+6.7)

47×65•66, not formed

48×65•66, not formed

49×65•66, 18.7(+5.2)

pH 6.5

46×65•66, 23.7(+2.2) 46×67•68, 25.4(+8.6)

47×65•66, 16.6(-4.9)

48×65•66, not formed

49×65•66, 25.2(+3.7)

Pnz

- con

juga

tion

PnzU

pH 6.0 46×65•66, 30.9(+5.3) 46×67•68, 30.0(+10.7)

47×65•66, 20.9(-4.7)

48×65•66, 15.7(-9.9)

49×65•66, 32.0(+6.4)

L1 - linker pH 7.3

50×65•66, 24.6(+11.1)

53×65•66, 22.6(+9.1)

56×65•66, not formed 59×65•66, 15.8(+2.3)

L2 - linker pH 7.3

51×65•66, 20.9(+7.4)

54×65•66, 20.3(+6.8)

57×65•66, 20.6(+7.1) 60×65•66, not formed

DPP

Z - c

onju

gatio

n

L3 - linker pH 7.3

52×65•66, 17.3(+3.8)

55×65•66, 20.5(+7.0)

58×65•66, not formed 61×65•66, not formed

(phen)2Ru2+DPPZ - conjugation (L2 - linker) pH 7.3

62×65•66, 21.2(+7.7)§

63×65•66 19.7(+6.2)§

64×65•66, not formed§

# Tm(∆Tm) in ºC are shown after numbers of triplex strands (symbols • and × stand for Watson-Crick and Hoogsteen hydrogen bonding respectively). ∆Tm = Tm(modified triplex) - Tm(corresponding native triplex) at given pH $ For sequence composition of oligos see Table 2 (page 16) § Unpublished data

of a sequence assisted in only moderate triplex 46×65•66 stabilization which was maximal at pH 6.0

(∆Tm = 5.3°C) slightly decreasing upon adjusting pH to 7.3 (∆Tm = 4.0°C). Comparable stability was

also observed for modification with Pnz attached at 5'-hydroxyl of 5'-terminal thymidine (PnzT) of TFO

strand as in 45×65•66 (∆Tm = 4.7°C and 3.9°C at pH 6.0 and 7.3 respectively).53 No triplex formation

or destabilization (at low pH) was detected when Pnz-modification was introduced at the 3'-end or at

the middle of TFO sequence. As a consequence, double modification with Pnz at both 5'- and 3'-ends of

TFO gave only marginal improvement of triplex stability (∆Tm = 6.4°C and 5.2°C at pH 6.0 and 7.3

33

respectively) with respect to single Pnz-modification at the 5'-terminal. Insertion of the five extra A•T

base paires into the duplex target 65•66 at the triplex-duplex junction, as in triplexes 44×67•68 and

46×67•68, allowed to enhance the stabilization effect of 5'-tethered Pnz group in comparison with its

affinity to the parent duplex target 65•66. This testifies that Pnz binds more strongly at T×A•T - A•T

triplex-duplex junction then at T×A•T - C•G site. As expected, the stability of Pnz-modified triplexes as

well as their native counterparts was found to increase upon lowering the pH.

It has been shown that DPPZ-conjugated ODNs have more affinity towards ssDNA target

(ODN•DNA duplex formation) than Pnz-modified analogs. The same trend was also observed with the

triplex formation, yet there are significant variations depending on the ODN attachment site (5'-, 3'-, or

internucleotide position) and the length of the linker arm used. The most essential stabilization was seen

with DPPZ conjugation at the 5'-terminal through the shortest linker L1 (∆Tm = 11.1°C) whereas the

increase of the linker length led to a sharp decrease of stabilization (up to ∆Tm = 3.8°C for the longest

linker L3). This tendency was also found for 3'-terminal DPPZ conjugation, albeit more gradually (∆Tms

were changed from 9.1°C to 6.8°C). Such effect can be attributed to the entropy decrease in the course

of triplex formation which should be higher upon employment of more extended flexible linkers. On the

other hand, longer linkers may interfere with the triplex strands thus destabilizing triple helical

structure. Triplex formation by 18mers carrying DPPZ at the central position was especially

sensitive to the linker length: only 12-atom linker L2 was found to provide optimal DPPZ configuration

to stabilize the triplex (57×67•68, ∆Tm = 7.1°C) while with DPPZ tethered through the shorter or longer

linkers no triplex formation was detected. Triplex 57×67•68 was, in fact, the only one among other

middle-modified analogs, studied in this work, which showed improved stability.

Despite the extensive investigation of polypyridyl Ru2+ complex interaction with dsDNA, only

few reports were devoted to the elucidation of the binding affinity of those complexes towards triple

helical DNA.219,220 It has been shown that [Ru(phen)2DPPZ]2+ intermolecularly bound to poly(T×dA•T)

or poly(dC+×dG•dC) significantly increases the triplex stability. Another issue to be addressed is the

metal complex binding mode. Because the major groove in triplexes is blocked by the third strand, the

major groove interacting drugs should not bind to the triple-stranded DNA or their binding mode must

be changed. Since the spectral characteristics of [Ru(phen)2DPPZ]2+ bound to poly(T×dA•T) triplex

were similar to those bound to poly(dA•T) duplex, it was concluded that the DPPZ ligand of the metal

complex is intercalated with the two phen ligands located in the minor groove, thereby stabilizing the

third strand by expansion of the stacking interaction. From this point of view, the tethering of potential

metallointercalator to the different sites of a TFO strand could shed light on a still contraversial

question: from which direction (major or minor groove) are the metal complexes intercalated? To date,

the available literature data on triplex formation assisted with metal complex conjugated TFO strand are

34

limited by two works in which [Ru(phen)2DPPZ]2+ 189 or [Ru(bpy)3]2+ 183 complexes were tethered

eithert at the 3'-terminal or at the middle of TFO respectively.

In our [Ru(phen)2DPPZ]2+-ODNs the metal complex was uniformly tethered through the DPPZ

ligand to different positions of 18mer ODN strand (not published data). The metal complex conjugation

at the 5'- or 3'-terminals allowed us to enhance the stability of the resulting triplexes (∆Tm = 7.7°C and

6.2°C for 62×65•66 and 63×65•66 respectively, Table 5). The insertion of metal complex at the interior

of a TFO strand led to the complete loss of ability to form a triplex. Following comparisons are

noteworthy: (1) Ligation of Ru2+(phen)2 moiety to the DPPZ ligand did not improve the TFOs binding

affinity with respect to DPPZ-tethered TFOs, if the modification is located at terminals of TFO

(compare stability of 62×65•66 and 51×65•66 as well as 63×65•66 and 54×65•66 in Table 5), which is

in contrast with the trend observed for the duplexes. (2) Middle-modification of an ODN strand with the

metal complex allowed to stabilize ODN•DNA duplexes but, it does not make the ODN able to bind to

dsDNA and form a triplex. These observations only strengthen the fact that the binding modes of the

tethered metal complex in the metal conjugated duplexes and triplexes are different. These binding

modes are directly dictated by structural compatibility of the metal complex and various types of DNA

hybrids. Most probably the bulky Ru2+ complex can not intercalate into the triplex from the minor

groove as well, when it is conjugated to the TFO strand because of the high steric interference of the

linker which connects the metal complex and the third strand.

2.2.4 Circular dichroism (CD) and DNase I footprinting studies on [Ru(phen)2DPPZ]2+-

labeled ODN•DNA duplexes performed for the elucidation of the metallointercalation

binding mode in the DNA double helixes (Paper IV, in collaboration with Mr. P. I.

Pradeepkumar)

CD spectroscopy221-223 was employed to determine the difference in the global structure of the

ODN•DNA duplexes site-specifically tethered with the metal complex at various positions. The CD

spectra of the racemic Ru2+-ODN conjugates 21, 22, and 23 hybridized with the DNA target 29 were

recorded to exclude the contribution of the CD signals originated from each of the pure Ru2+complex

diastereomers in order to observe the global helical conformation of the DNA duplex. The spectral

characteristics for 5'- and middle-modified duplexes 21•29 and 23•29 were significantly different from

those of the native counterpart. Positive (267.5 nm) and negative (240.0 nm) bands of the native duplex

were shifted correspondingly to 260 nm and 232.5 nm of modified analogs, while CD spectra of 3'-

conjugated duplex 22•29 exhibited some features which are characteristic for natural duplex: i.e. the

same negative band wavelength (240 nm) and a shoulder at 267.5 nm which coincides with the native

duplex positive band. At the same time, crossover point (246.3 nm) and the wavelength shift of main

35

positive band for 3'-modified duplex 22•29 were identical with those of the 5'- and middle-modified

analogs. Generally, it appears that the metal conjugation to ODN applied in the present work allowed to

bring the metal complex in tight contact with nucleobases which leads to the essential global helical

structure alteration from the native B-type helix. This alteration was more pronounced in the case of 5'-

and middle-conjugation. The different structural changes in the enantiomerically pure Ru2+-ODNs 23a-

d upon its hybridization with DNA target 29 further proved that the metal complex strongly interact

within double helix: distinct CD changes upon hybridization of ∆-isomers 23c-d compared with those

of Λ-isomers 23a-b implies their different metal complex binding geometry in the duplex. This

situation is expected when ancillary phen ligands are rigidly located within helical grooves but not

when the metal complex is externally bound.224

DNase I provides simple and quick method for identifying or confirming the preferred dsDNA

binding sites for several ligands.225-227 We exploited this technique to evaluate the extent of alterations

from the B-type DNA duplex structure caused by tethered metal complex as well as to study the DNase I

cleavage pattern as a reflection of structural accessibility of phosphodiester groups which can be

protected from the enzyme digestion by the appending Ru2+ complex. The extent of the cleavage by

DNase I (Figure 1) decreases as follows: native duplex 1•29 (97%) ~ 3'-Ru2+-modified 22•29 (98%) >

middle-Ru2+-modified 23•29 (48%) > 5'-Ru2+-modified 21•29 (38%) signifying that in case of 5'- and

middle-modification the duplex structure is deeply altered from the canonical B-form which is ideal

for DNase I recognition.228 Nevertheless, conformational reorganization in the 5'- and middle-modified

duplexes do not completely preclude the recognition by the enzyme. DNase I is known to bind at least 4

bp to the 5'-end and 6 bp to the 3'-end from the scissile phosphodiester bond of the substrate duplex.228

Thus, the full length of studied 9mer + 11mer duplex should be covered by DNase I. Cleavage patterns of

5'- and 3'- ruthenated duplexes are not distinguishable from the native counterpart, while middle

modified analog has completely different digestion pattern with more or less one cleavage site only

(between 8G and 9T, Figure 1). Moreover, it was found that the cleavage sites of the middle-DPPZ-

modified duplex 16•29 were stretched from 5T to 8G (Figure 2). This shows that the bulky

[Ru(phen)2(dppz)]2+ moiety, tethered at the middle of ODN sequence, acts as a protection against the

enzyme cleaving site (E75-H131-H2O228), blocking those phosphodiester groups which are normally

cleaved when the metal entity is distal from the center of a duplex (as in case of 5'- and 3'-metal complex

conjugation) or when simple planar DPPZ group is used for the internucleotide conjugation. The

cleavage patterns of the diastereomerically pure middle-modified (Ru2+-ODN)•DNA duplexes were also

found (Figure 3) to be distinct from each other confirming the CD spectral data that binding geometries

of ∆- isomers 23c-d and Λ-isomers 23a-b are indeed different. Altogether, the CD and DNase I

36

Figure 1. DNase I digestion of duplexes formed with 5'-32P-labeled target 29 and native ODN 1 (B) or the racemic Ru2+-modified ODNs 21 (D), 22 (C), 23 (A). Time in minutes after the addition of the enzyme is shown at the top of each gel lane. The main cleavage sites after 30 min of reaction are indicated by arrows.

Figure 2. DNase I digestion of duplexes formed with 5'-32P-labeled target 29 and native ODN 1 (B) or internally modified DPPZ-ODN conjugate 16. Time in minutes after the addition of the enzyme is shown at the top of each gel lane. The main cleavage sites after 30 min of reaction are indicated by arrows.

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P(B)

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P(A)

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P(C)

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A 9G8G7T 6T5T4G3T2A1C-32P(D)

720' 180' 90' 30' 15' 720' 180' 90' 30' 15'

Midle Ru2+-mod. (A) Native (B) 3'- Ru2+-mod. (C) 5'-Ru2+-mod. (D)

60' 30' 15' 5' 60' 30'

11C10A9T8G7T6T5T4G3G5'

11C10A9T8G7T6T5T4G3G

5'2T

11C10A9T8G7T6T5T4G3G

5'2T

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P(A)

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P(B)

11C10A9T8G7T6T5T4G3G

2T5'

180' 90' 15' 180' 90' 15'

Native (B)Dppz-mod. (A)

11C10A9T8G7T6T5T4G3G5'

37

footprinting data performed on ruthenated duplexes point to some special binding mode which allow

the metal complex strongly interact with nucleobases which leads to (1) the high increase of duplex

stability, (2) conformational reorganization of DNA double helix, and (3) protecting phosphodiester

groups in helical grooves.

Figure 3. DNase I digestion of duplexes formed with 5'-32P-labeled target 29 and native ODN 1 or different diastereomers of Ru2+ middle-modified ODNs 23a-d. Time in minutes after addition of the enzyme is shown at the top of each gel lane. The main cleavage sites after 30 min of reaction are indicated by arrows.

The results of CD, DNase I footprinting, and thermal denaturation studies on ODN•DNA

duplexes tethered with [Ru(phen)2DPPZ]2+ complex and their comparison with the thermal stabilities of

analogous ODN•RNA duplexes and ODN×DNA•DNA triplexes, allowed us to suggest the most

probable mode of complex interaction within the DNA double helix. As it was shown, the DPPZ

moiety of [Ru(phen)2(DPPZ-CO-L2)]2+ complex intercalates into the native duplex 1•29. We proposed

a similar intercalation mode of DPPZ subunit in the [Ru2+(DPPZ)-ODN]•DNA duplexes. Tethering of

[Ru(phen)2DPPZ]2+ or [Ru(tpy)(DPPZ)(CH3CN)]2+ to an ODN through the DPPZ ligand leaves the

only possibility for DPPZ to intercalate in a duplex: via its threading across the double helix core thus

delivering Ru2+(phen)2 or Ru2+(tpy)(CH3CN) moieties to the final position in the opposite groove

(Figure 4). This type of binding229 can be envisaged to give a very rigid structure in which covalently

attached complex acts as a staple, holding the bases firmly stacked together near the intercalation site.

On the other hand, upon such “stapling” dsDNA is likely to undergo a much larger conformational

reorganization then in case of classical intercalation of a planar chromophore. With the above scenario,

the resulting ODN•DNA duplexes should be very stable, their global structure should be altered from

the parent B-type, and the position of the Ru2+(phen)2 moiety is restricted in the helical groove

protecting some nearest backbone phosphodiester groups. The protecting ability is also expected to be

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P23a , 23b

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P23c

5'-1T 2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C-32P23d

90' 180' 90' 180' 90' 180' 90' 180' 90' 180'

23a(Λ) 23b(Λ) 23c(∆) 23d(∆) Native

11C10A9T8G7T6T5T

4G3G

2T5'

11C10A9T8G7T6T

5T4G3G

2T5'

38

dependent on the ∆- or Λ-configuration of the two ancillary phen ligands around Ru2+ center. These

expectations were in agreement with the results obtained for our 5'- and middle-modified duplexes

21•29 and 23•29. The net free-energy gain which can be reached by metal complex threading is

Figure 4. Proposed metallointercalation mode shown for the ∆-[Ru(phen)2DPPZ]2+ complex tethered to the internal position of the duplex 23•29 through the linker L2. The linker-complex unit –L2-DPPZ Ru2+(phen)2 covalently linked to the glycerol moiety of the ODN 23 is depicted in bold.

determined by balancing between the positive deposit of additional π-π stacking interactions and

negative impact of structural reorganization of the DNA helix near the metallointercalation site. The

last factor is very sensitive to the initial helical conformation undergoing structural changes upon the

threading of metal complex. This explains the fact that Ru2+ middle-modification gave us highly

stabilized ODN•DNA duplex while analogous ODN•RNA duplex had the same stability as the native

counterpart. Finally, the presence of the third strand bound to the dsDNA in the major groove

completely inhibited triplex formation consisting of Ru2+ middle-modified TFO strand. Interesting is to

note also that comparison of Tms of [Ru(tpy)(DPPZ)(CH3CN)]2+-ODN•DNA duplexes 39•43 - 42•43

with Tms of structurally related amino-linker modified duplexes 35•43 - 38•43 allows to estimate the

stabilization effect inherent in [Ru(tpy)(DPPZ)(CH3CN)]2+ itself. Such effect is more pronounced in

case of internucleotide conjugation (∆∆Tm = 24.3°C for 41•43 and 37•43) which is more than for 5'- and

3'-metalated duplexes (in 1.5 and 3.5 times respectively). This observation can be interpreted by the

structural feature of the applied internucleotide modification: insertion of a bulged abasic site magnifies

the distance between adjacent base pairs providing more space for accommodation of the bulky metal

39

complex (Figure 5A and 5D). In the 5'-Ru2+-modified ODN•DNA duplexes the internucleotide

distance expected for metallointercalation is not structurally increased giving the less positive net

Ru

Ru

Ru

5'-1T2C3C4A 5A6A7C8A9T

3'-11C10A9G8G7T 6T5T4G3T2A1C

5'-1T 2C3C4A5A6A7C8A9T

3'-11C10A 9G8G7T6T5T4G3T2A1C

5'-1T2C3C4A5A6A7C8A9T

3'-11C10A9G8G7T6T5T4G3T2A 1C

(E)

(A)

(B)

(C)

(D)

(F)

5'-1C2T3T4A5C 6C7A8A9T10C

3'-11G10A9A8T7G 6G5T4T3A2G1TRu

Ru

Ru

5'-1C 2T3T4A5C6C7A8A9T10C

3'-11G 10A9A8T7G6G5T4T3A2G1T

5'-1C2T3T4A5C6C7A8A9T10C

3'-11G10A9A8T7G6G5T4T3A2G 1T

Figure 5. Schematic illustration of the positioning of the metal complex in the various ODN•DNA duplexes site-specifically labeled with [Ru(phen)2(DPPZ)]2+ (A – C) or [Ru(tpy)(DPPZ)(CH3CN)]2+ (D – F) metal complexes.

stabilization effect (Figure 5B and 5E). The relatively low stabilization of 3'-Ru2+-modified duplexes,

which is comparable with those of (DPPZ-ODN)•DNA duplexes 11•29 and 14•29 (Table 3), can be a

result of the less effective π-π stacking interactions between the metal complex and the terminal base

pair instead of intercalation between internal base pairs (Figure 5C and 5F).

2.3 RNase H degradation studies on the AON•RNA hybrid duplexes conjugated with various

intercalators at the ODN strand (Paper III, in collaboration with Dr. E. Zamaratski and Mr. P.

I. Pradeepkumar).

2.3.1 Physicochemical properties of RNA targets and their duplexes formed with AON conjugates.

To evaluate the potency of ODNs (abbreviated in this chapter as AONs) conjugated with intercalating

agents to form stable ODN•RNA duplexes, three RNA targets 31 – 33 (Table 1) have been chosen

consisting of the same central 9mer tract which is complementary to the AON 1 sequence and flanked

with different patches of deoxynucleotides with the expectation to give different self-folding capacities

of the resulting RNA molecules. The self-aggregation of these targets was examined by CD and UV-

spectroscopy. Thermal melting of the 11mer RNA 31 showed no sigmoidal change in UV absorbance

while clear concentration dependent transition was observed for the 17mer RNA 33 which allowed us

to determine Tm (29.6°C and 31.2°C at 1 µM and 3 µM concentration respectively) and the

thermodynamics of its self-aggregation: ∆H° = -342 kJ/mol, -T∆S° = 303.9 kJ/mol, and ∆G°298 = -38.3

40

kJ/mol. Melting experiments with another 17mer RNA 32 revealed only slight hyperchromic effect. The

temperature-dependent CD was found to be more sensitive tool230 to monitor structural transitions in

given RNA targets. The CD spectra recorded at three different temperatures (6°C, 20°C, and 50°C)

were almost identical for 11mer 31 whereas ellipticity of 17mer 33 strongly decreased upon heating,

suggesting a high degree of self-organization. Although 17mer 32 did not show any clear transition in

the UV melting experiments, it exhibited a definite (but less pronounced compared to RNA 33)

temperature dependency of its CD spectra. The ellipticity measured at 265 nm in the range of

temperatures from 5°C to 60°C was found to have the sigmoidal dependence from temperature for both

RNAs 32 and 33, giving the Tms values for RNA aggregation (24.7°C for target 32 and 28.8°C for

target 33 at 1 µM concentration). The concentration dependence of Tm found from the CD experiments

for both targets allowed us to tentatively propose the structures of self-aggregated RNAs 32 and 33:

Thus, the above RNA targets with different tertiary structures231,232 gave the possibility to study the

effect of intercalator tethered to the AON to influence on the two competing and reversible

hybridizations leading correspondingly to RNA•RNA and AON•RNA structures.

Native AON 1 and its conjugates tethered with Pnz and DPPZ chromophores at 5'- or 3'-terminals

as in 3, 8, 10, and 14 (Table 1) have been chosen to study the effect of covalently linked intercalator on

the antisense activity of the resulting ODN conjugates. The phosphorothioate (PS) analogs of the above

AONs have also been synthesized (PS-AONs 24 – 28) to compare the effect of chromophore

conjugation with their phosphodiester (PO) counterparts. All chromophore-conjugated (both PO and

PS) AON•RNA duplexes formed with non-aggregated RNA target 31 demonstrated higher stability

compared to the non-conjugated analogs (Table 6). The additional π-π stacking interactions between

the aromatic rings of the chromophore and the adjacent base pairs provide a significant gain in enthalpy

(∆H°) upon hybridization of conjugated AONs to the RNA target 31. On the other hand, the competing

decrease of entropy (∆S°) is much higher when conformationally flexible tethers participate in the

duplex formation which restrict their flexibility due to the positioning of a chromophore near the paired

nucleobases. The net change of free energy (∆G°298) was yet found to be more negative for the

hybridization of conjugated AONs to the non-aggregated target 31 compared to the native counterparts,

which correlates with the enhanced thermal stability of chromophore-tethered AON•RNA duplexes. As

expected, PS-AON•RNA duplexes revealed poorer stability then their PO-analogs (0.5 - 1°C Tm

decrease for each PS-backbone modification)71 as well as the stabilizing effect of a chromophore in the

case of PS-AON•RNA duplexes was less then for the corresponding PO-hybrids.

41

When the low-aggregated 17mer RNA target 32 was hybridized with PO-AONs 3, 8, 10, 14, the

stability of the resulting duplexes was close to that of duplexes formed with these PO-AONs and 11mer

RNA target 31 suggesting that the chromophore-conjugated PO-AONs effectively prevents self-

aggregation of RNA 32 upon targeting to it. In contrast, the Tms of PS-AON•RNA duplexes and the

native PO-AON•RNA hybrid were somewhat lower (by 1.1 – 3.8 °C) which can be due to the existence

of both RNA•RNA (Tm = 24.7°C) and AON•RNA structures in dynamic equilibria with single-stranded

molecules. Note that ∆G°298 of self-aggregation of RNA target 32 obtained from CD melting study

Table 6. Thermodynamic characteristics of the AON•RNA duplexes (1 µM) formed by PO- and PS-AONs with various RNA targets (non-aggregated 11mer 31, low-aggregated 17mer 32, or highly-aggregated 17mer 33)

Target 31

Target 32

Target 33

AON$ Tm,

(˚C) ∆Tm, (˚C)

∆Hº, (kJ mol-1)

∆Sº, (e.u)

∆Gº298, (kJ mol-1)

Tm, (˚C)

Tm, (˚C)

Native 1 22.1 - -226±8 -0.65±0.03 -33.8±0.5 20.2 28.6

5'-Pnz-mod. 3 28.7 6.5 -298±4.5 -0.87±0.01 -39.8±0.1 28.5 30.5

5'-DPPZ-mod. 10 27.6 5.5 -286±20 -0.83±0.07 -38.7±2.8 27.7 29.2

3'-Pnz-mod. 8 28.6 6.5 -256±10 -0.73±0.03 -39.2±1.5 28.1 30.4

PO-A

ON

s

3'-DPPZ-mod. 14 30.6 8.5 -316±23 -0.92±0.07 -41.9±3.2 30.7 30.3

Native 24 16.6 - -220±14 -0.64±0.05 -29.5±1.6 11 - 14 29.3

5'-Pnz-mod. 25 19.1 2.4 -245±20 -0.72±0.06 -30.9±2.7 15.7 28.9

5'-DPPZ-mod. 27 19.3 2.7 -279±14 -0.83±0.05 -30.6±1.3 18.2 29.1

3'-Pnz-mod. 26 19.7 3.1 -267±3.5 -0.79±0.01 -31.3±0.6 15.9 29.2

PS-A

ON

s

3'-DPPZ-mod. 28 22.0 5.3 -257±15 -0.75±0.05 -33.4±1.5 20.9 29.2

$ For sequence composition of oligos see Table 1 (page 15)

(-34.7 kJ/mol) is very close to those of the hybridization of PS-AONs 24 – 28 and native PO-AON 1

towards 11mer RNA 31 (29.5 – 33.8 kJ/mol). The difference in Tms between PS-AON•32 and PS-

AON•31 duplexes was minimal in the case of 5'- and 3'-DPPZ-conjugated PS-AONs 27 and 28,

indicating that DPPZ assists in the shift of equilibria towards AON•RNA duplex formation more

effectively.

When AONs were hybridized with highly-aggregated 17mer RNA target 33, all Tms were close to

the Tm of the target aggregation itself (28.8°C and 29.6°C obtained from CD and UV melting studies

respectively), which made it impossible to estimate the extent of the AON•RNA duplex formation.

Comparing free energies of the self-aggregation of RNA target 33 (-38.3 kJ/mol) with ∆G°298 of the

42

AON•31 duplex formation, we assumed that only a fraction of AON strands was able to form the

desired duplex with highly aggregated RNA 33. Generally, these data show that the thermodynamic

accessibility of RNA tertiary structure for AON•RNA duplex formation can be increased by tethering of

the intercalating agents to an AON strand.

Another important aspect of successful application of conjugated AONs in the RNase H mediated

degradation of target RNA molecules is the deviation of the global AON•RNA heteroduplex structure

from the native DNA•RNA conformation, in which RNA strand is of the A-type and the DNA strand

has a conformation close to the B-type.233,234 This type of the heteroduplex conformation was proven to

be essential for the substrate recognition by RNase H.233-236 CD spectra of our chromophore-conjugated

duplexes (both PO and PS) formed with 11mer RNA target 31 were very similar to the native duplexes

1•31 and 24•31, and rather different from those of the highly aggregated 17mer RNA 33 and natural

AON•DNA duplex 1•29 representing the canonical A- and B-type structures respectively. The shape of

all heteroduplex spectra supports that the global structure of AON•RNA duplexes is inremediate

between A- and B-type. Thus, terminal conjugation of the planar chromophores through the flexible

linkers does not alter the helical structure inherent in the native DNA•RNA duplex.

2.3.2 RNase H mediated cleavage studies on the conjugated AON•RNA duplexes

5'-32P-labeled RNA targets were hybridized with complementary AON strands in equimolar ratio (0.8

µM) and incubated with Escherichia coli RNase H at 21 °C and aliquots were taken after 15 and 120 min

intervals. All modified AONs (both PO and PS) promoted a higher degree of RNA hydrolysis compared

to the natural counterparts 1 and 24 (Table 7) irrespective of the nature of the RNA target (31, 32, or 33).

The following comparisons are noteworthy: (1) DPPZ conjugated PO-AONs and PS-AONs promoted

exceptionally higher extent of the cleavage of all RNA targets compared to the Pnz-modified analogs.

(2) Generally, 3'-conjugation of a chromophore assisted in slightly better hydrolytic activity of RNase H

relative to the corresponding 5'-conjugates (especially in the case of DPPZ-modification). (3) The extent

of hydrolysis for the PS-AON•RNA substrates was somewhat lower in comparison with the PO-

counterparts as expected from the lower thermal stability of these duplexes. (4) When PO- and PS-

AONs were hybridized with the low- or highly-aggregated 17mer RNAs 32 and 33, the hydrolysis yields

remained approximately the same as those with the non-aggregated 11mer RNA 31 despite different

accessibility of the single strand region in these RNA targets. This suggets that the rate of conversion of

the folded RNA structures to the single-stranded form is much faster than the rate limiting RNase H

promoted cleavage and the kinetic accessibility of the single strand region in all three targets becomes

equal. (5) No clear correlation between the thermal stability of AON•RNA duplexes and the extent of

their cleavage by RNase H can be drawn on the basis of present data. According to the Michaelis-Menten

43

mechanism of the enzyme catalysis, the extent of the target RNA cleavage in the AON•RNA duplexes

depends not only on the thermodynamic stability of a duplex (expressed by Tm) but also on the

recognition (expressed by Km) and catalytic (expressed by kcat) properties of RNase H towards the hybrid

duplex. As more recent studies showed,237,238 when RNA is completely hybridized with AON and the

effect of thermodynamics is thus excluded, the cleavage extent is determined only by Km and kcat, which

Table 7. Extent (after 2 h of incubation) of the PO-AONs and PS-AONs promoted RNase H hydrolysis of the RNA targets with different capacity to the self-aggregation.

Extent of hydrolysis with:

AON$

Non-aggregated RNA target 31

Low-aggregated RNA target 32

Highly-aggregated RNA target 33

Native 1 60% 65% 64%

5'-Pnz-mod. 3 72% 76% 69%

5'-DPPZ-mod. 10 80% 78% 72%

3'-Pnz-mod. 8 76% 76% 71%

PO-A

ON

s

3'-DPPZ-mod. 14 92% 85% 82%

Native 24 40% 41% 32%

5'-Pnz-mod. 25 61% 51% 39%

5'-DPPZ-mod. 27 75% 75% 60%

3'-Pnz-mod. 26 58% 59% 48%

PS-A

ON

s

3'-DPPZ-mod. 28 91% 87% 83%

$ For sequence composition of oligos see Table 1 (page 15)

have their own and complex dependency on the Tm values for various heteroduplex systems. On the

other hand, it was shown239 that the concentration of AON required to reach the maximal degree of the

RNA hydrolysis under the fixed RNA concentration indeed reflected the thermodynamics of

AON•RNA duplex formation. The more thermally stable the heteroduplex is, the less amount of

antisense oligonucleotide is necessary to achieve the saturation of the RNA target, after which the

cleavage extent becomes maximal and no more proportionally dependent on the exclusively

thermodynamic parameters of a hybrid duplex.239

As evident from the PAGE pictures (Figure 6), all modified AON•31 duplexes had the same

cleavage sites as the native counterparts. The exeption was found for the 3'-DPPZ-conjugated hybrid,

44

RNA 31RNA 31

RNA 32RNA 32

RNA 33RNA 33

G8G9

U7

G9G8U7

A13G12U10

A13G12

U10

A13

U10

G12

A13G12

U10

A

B

C

5'-r(CAUGUUUGGAC)-3'3' - TACAAACCT - 5'

5'-r(ACUCAUGUUUGGACUCU)-3'3' - TACAAACCT - 5'

5'-r(UAACAUGUUUGGACUCU)-3'3' - TACAAACCT - 5'

G9G8

U7

A13

A13

G12

G12

U10

U10

1 2 3 4 5 6 7 8 9 10

1 2 3 4 5

1 2 3 4 5

6 7 8 9 10

6 7 8 9 10

Figure 6. RNase H hydrolysis of: 11mer RNA 31 (A), 17mer RNA 32 (B), and 17mer RNA 33 (C) after 15 min of incubation with PO-AONs 10, 3, 1, 14, 8 (lanes 1 – 5 in each gel respectively) or PS-AONs 27, 25, 24, 28, 26 (lanes 6 – 10 in each gel respectively). Arrows indicate the major cleavage sites.

which showed site-specific hydrolysis at the U7 position. The digestion patterns of both aggregated

17mer RNAs 32 and 33 were slightly altered in comparison with the short non-aggregated 11mer RNA

31. Again for the 3'-DPPZ-modified duplexes formed with targets 32 or 33 only a single cleavage site

was observed at the U10 which corresponds to the U7 position in the RNA 31. These results show that

the nature of chromophore directly modulate the recognition and binding properties of the enzyme.

45

Thus, it has been clearly demonstrated that the tethering of the planar chromophores (Pnz and

DPPZ) at the terminal positions of the AON strand stabilized the resulting AON•RNA duplexes without

altering their global helical conformation. As a result, the extent of RNase H promoted hydrolysis was

improved compared to the native counterpart. The relatively poorer thermal stability and the RNase H

assisted cleavage efficiency of the PS-AON•RNA duplexes was greatly compenseted by tethering of a

suitable chromophore (as in 3'-DPPZ-conjugated hybrid). The nature of the chromophore appeared to

contribute to the antisense effect of the modified AON not only by the increase of the thermodynamic

stability of the corresponding hybrid duplexes but mainly by the influence on the recognition properties

of RNase H towards modifications in the AON•RNA duplexes.

In order to investigate the tolerance level of RNase H233-236,240 towards the various chromophore

conjugation in the antisense strand, causing the AON•RNA duplex conformational changes in a

different manner, the middle-DPPZ-conjugated AON 16 and AONs 21 – 23 tethered with

[Ru(phen)2DPPZ]2+ moiety at various sites of a sequence have been hybridized with low-aggregated

17mer RNA 32 and subjected to the enzymatic hydrolysis. All of these AONs have been shown to form

stable duplexes with non-aggregated RNA target 31, and the stability of 5'-Ru2+ or 3'-Ru2+-conjugated

duplexes (as in 21•31 and 22•31, Table 4) was even improved to a greater extent compared to that for

the 3'-DPPZ-modified duplex 14•31. Despite that 3'-DPPZ-modified duplex 14•32 showed the best

ability to activate RNase H cleavage, the extent of RNA 32 cleavage was found (Table 8) to be less for

all tested duplexes compared to the native counterpart except for the 3'-Ru2+-modified analog, whose

antisense activity remained the same as for the 3'-DPPZ-modified duplex 14•32. The introduction of the

Table 8. Extent of RNase H hydrolysis of the low-aggregated RNA 32 promoted by PO-AONs tethered with DPPZ- or Ru2+(phen)2DPPZ at different sites of AON sequence.$

DPPZ-modification at: Ru2+(phen)2dppz-modification at: Native duplex, 1•32

3'-terminal, 14•32

5'-terminal, 10•32

center, 16•32

3'-terminal, 22•32

5'-terminal, 21•32

center, 23•32

60%

91%

75%

50%

92%

49%

4%

$ For sequence composition of oligos see Table 1 (page 15)

modification at the middle of a sequence should decrease the enzyme binding which is more poor as

more bulky group placed and more helical structure changes can be induced by such a group. The

influence of the modification is less but still takes place when the conjugation is performed at the 5'-

terminal of a sequence: in our experiments the cleavage sites of the target RNA are in close proximity

46

to the 5'-end of the AON sequence and the presence of any tether at this position might interfere with

the enzyme active sites. Finally, the 3'-end of the AON seems to be the most suitable for the

introduction of different chemical functions as bulky as [Ru(phen)2DPPZ]2+ complex. The marked

difference in the activation of RNase H cleavage for the middle or 5'-metal Ru2+ complex tethered

heteroduplexes and their corresponding DPPZ-tethered analogs might be a consequence of the

specificity of metallointercalation which, as was shown on ODN•DNA duplexes, strongly alters the

conformation of a duplex.

2.3.3 Nuclease resistance of the 3'-Pnz and 3'-DPPZ-conjugated AONs

To be suitable for pharmaceutical application, the AONs are required to be resistant to the nucleic acid

degrading enzymes.240,241 The 3'-exonuclease activity was found to be responsible for most of the AON

degradation,240-242 therefore, modification of AONs at the 3'-end has been used to minimize the extent

of degradation.243,244 Under incubation of our AONs with snake venom phosphodiesterase, the non-

modified PO-AON 1 had a half-life time of 4 min, however, 3'-Pnz and 3'-DPPZ-modified PO-AONs 8

and 14 did not show any sign of degradation even after 2 h of incubation. Similar enhancement of

stability was observed with the phosphorothioate analogs. Most probably the steric hindrance caused by

the tethered polyaromatic system as well as the flexible linker prevents the enzyme binding to the

modified AON.

2.4 Specific chemical modifications of the complementary DNA strand by DPPZ and Ru2+-

DPPZ tethered ODNs in ODN•DNA duplexes.

2.4.1 Sequence-specific cleavage of target DNA by metal-binding DPPZ-group linked to an

internucleotide position of the complementary ODN strand (unpublished data)

The intercalating ability of some chromophores can be endowed with chemical properties such that they

can be used to induce irreversible modifications in their target sequence. These chromophore-ODN

conjugates appear particularly attractive for those ODNs which do not allow cleavage of mRNA by

RNase H and, for this reason, often have little or no antisense activity.9,68,70,75 Moreover, the reactivity

of interactive groups can be directed to specific sites in both single-stranded mRNA and double-

stranded genomic DNA allowing to improve the potency of synthetic ODNs as antisense and antigene

agents.11,245

The DPPZ ligand can be considered as the extended structural analog of the phenanthroline, with

two chelating nitrogens which are able to coordinate transition metal ions. Metal complexes formed

with ODN-tethered ligands have been shown to act as sequence-specific cleavers of RNA101-106 and

DNA targets.110-115,117-121 Such polypyridyl ligands as terpyridine (tpy),101,246-249 phenanthroline

47

(phen),249,250 and bypyridine (bpy)251 have been linked to ODNs or monomeric 2'-deoxynucleosides to

give artificial ribonucleases which successfully hydrolzed RNA in ODN•RNA duplex. On the other

hand, nuclease activity of Cu+(phen)2 and the related complexes towards dsDNA has been

comprehensively studied252-263 and found to proceed through the generation of hydroxyl radicals which

probably abstract H atom from the deoxyribose of DNA with subsequent break of the phosphodiester

backbone:

M(n-1)+ + O2 → Mn+ + O2• (HO2

•)

O2• (HO2

•) + H2A → HO2 (H2O2) + HA•

M(n-1)+ + H2O2 → Mn+ + HO• + OH

Mn+ + H2A or HA• → M(n-1)+ + HA• or A + H+

HO• + H-DNA → H2O + DNA•; DNA• → scission products,

where M(n-1)+/ Mn+ corresponds to the Cu+/Cu2+ or Fe2+/Fe3+ redox active pairs and H2A is the auxiliary

reducing agent.264 Correspondingly, phen ligand was covalently tethered to sequence-specific DNA

binding protein,265,266 minor groove binder agents,266 short DNA113-117,267 or RNA molecules268,269 for

directing Cu+(phen)2 nuclease activity in a sequence specific manner towards RNA,266,116 ssDNA,113-

115,267 and dsDNA.115,117,265,266,268 From this point of view, the ODN conjugates covalently linked to the

metal chelating DPPZ group are expected to sequence specifically hydrolyze RNA (artificial

ribonucleases and ribozymes) or oxidatively cleave dsDNA when hybridized to their targets. In this

sense linking of DPPZ group to ODNs would allow to prepare new antisense or antigene agents which

act through the chemical inactivation of genetic information.

To test the ability of the DPPZ moiety to participate in the formation of redox-active complex

with the subsequent cleavage of target DNA in the ODN•DNA duplex, the middle-modified DPPZ-

ODN conjugates 15 – 17 (Table 1) were hybridized with 5'-32P-labeled DNA target 29 and then

incubated with ferrous ammonium sulfate and DTT as a reducing agent. The presence of DPPZ-ODN

conjugates led to the highly specific formation of scission products at the positions of the expected

ligand intercalation (T6 and T7, Figure 7) while the native ODN 1 did not induce any target degradation

after 5 h of incubation. The extent of cleavage was greater for the internucleotide DPPZ-conjugation

through the linker L2 (28% of the target 29 was cleaved with 66% of the cleavage occurring at T6 and

T7). The cleavage was less for DPPZ-L1- and DPPZ-L3-ODN conjugates which correlated with the

lower stability of their duplexes. It should be noted that the extent of cleavage with our DPPZ-ODN

conjugate is of the same order of magnitude compared to that, which was detected with phen-tethered

ODN under the same conditions (38%).267 These experiments proved that our DPPZ-modified ODNs

have the potential to act as sequence-specific redox-based artificial nucleases, almost as good as phen-

ODN conjugates.267

48

Figure 7. Scission of the duplexes formed with 5'-32P-labeled 11mer DNA 29 and ODNs covalently linked to the DPPZ chromophore at the middle position of a sequence. (A) Target DNA 29 (1 µM) was hybridized with complementary ODN (5 µM) and then incubated for 5 h at r.t. with 10 µM Fe2+ and 4 mM DTT in a buffer containing 40 mM Tris•HCl and 0.2 M NaCl. Lanes 2 – 5 correspond to the experiments with native ODN 1 and DPPZ-ODN conjugates 15, 16, 17, respectively. Lane 1: footprinting of the native duplex 1•29 (10 µM) with Fe2+-MPE (50 µM) in the presence of 4 mM DTT in a buffer containing 10 mM Tris•HCl and 50 mM NaCl. (B) Sites of primary cleavage.

2.4.2 Photochemical generation of Ru2+(tpy)(DPPZ)(H2O)-ODN (Paper V)

[Ru(tpy)(DPPZ)(CH3CN)]2+-ODN conjugates represent another type of ODNs attached to reactive

groups inducing chemical modifications in its target nucleic acids in a sequence-specific manner. Basing

on the photochemical properties of Ru2+-CH3CN complexes172,193,194 one can expect the dissociation of

the CH3CN ligand in the [Ru(tpy)(DPPZ)(CH3CN)]2+-ODNs upon light exposure. This has been indeed

observed by MALDI-TOF MS analysis (see chapter 2.1.6). The photochemical behavior of

[Ru(tpy)(DPPZ)(CH3CN)]2+-ODN conjugates has also been examined by UV-vis spectroscopy in

aqueous solutions not containing any potential nucleophiles. Under light irradiation (1 h, λ > 300 nm) the

MLCT absorption bands initially centered at 454 nm shifted to longer wavelength (482 nm) exhibiting

three isobestic points in the 300 – 600 nm region (309, 387, and 465 nm respectively). The same change

in the UV-vis characteristics was also observed in the course of light irradiation of the not tethered

[Ru(tpy)(DPPZ-CO-L2)(CH3CN)]2+ complex. The photosubstitution of CH3CN ligand by solvent

molecule has been established for [Ru(tpy)(bpy)(CH3CN)]2+ or related complexes193-195 which allowed us

to assume that [Ru(tpy)(DPPZ-CO-L2)(CH3CN)]2+ complex should undergo photoaquation affording

[Ru(tpy)(DPPZ-CO-L2)(H2O)]2+ if no any potential nucleophiles are present in the mixture. To confirm

this assumption, [Ru(tpy)(DPPZ-CO-L2)(H2O)]2+ has been chemically synthesized from its

[Ru(tpy)(DPPZ-CO-L2)Cl]+ precursor by treatment with a silver p-toluenesulphonate270,271 and the

product was found by UV-vis and 1H-NMR spectroscopy to be identical to that which was photolytically

obtained from the corresponding Ru2+-CH3CN analog. The similarity of the spectroscopic changes

DNA 29

T7

T6

G8

G9

A10

T5

G4

T7

T6

A B

5'-CATGTT TGGAC-3'

3'-TACAA ACCT-5'

T7T61 2 3 4 5

49

occurring upon photoirradiation of the non-tethered complex [Ru(tpy)(DPPZ-CO-L2)(CH3CN)]2+ and

[Ru(tpy)(DPPZ)(CH3CN)]2+-ODN conjugates enable us to conclude that the photosubstitution of CH3CN

ligand by H2O molecule takes place also on an oligonucleotide level affording [Ru(tpy)(DPPZ)(H2O)]2+-

ODN conjugates. The metallated ODNs were analyzed by PAGE before and after photolysis to show that

no ODN decomposition occurs upon the irradiation conditions applied. The above observations based

on MS and UV-vis spectroscopic data thus clearly point that our [Ru(tpy)(DPPZ)(CH3CN)]2+-ODN

conjugates 39 – 42 (Table 1) are indeed thermally stable, and can be activated by light through the

dissociation of the CH3CN ligand. Acetonitrile thus serves as a protective group for the ODN tethered

rective monofunctional [Ru(tpy)(DPPZ)(H2O)]2+ complex. This also suggests that CH3CN is expected to

be replaced by other potential coordinating species if they exist in the reaction mixture.

2.4.3 Photochemical cross-linking of Ru2+(tpy)(DPPZ)(CH3CN)-ODN conjugates to the

complementary DNA strand in ODN•DNA duplexes (Paper V)

Since the initial reports by Rosenberg at al. on the biological affects of some platinum complexes, in

particular cisplatin,272,273 there has been a great deal of interest in the development of transition metal

complexes for use in medicine.274 Most research on potentially chemotherapeutic transition metal

complexes has involved thermal aquation.135-145 Particularly, it has been established that mono- and

diaqua polypyridyl Ru2+ complexes covalently binds to DNA (presumably at N7 of guanine residue)

with different efficiency depending on the steric effects caused by polypyridyl ligands in the octahedral

geometry.275,276 A striking increase of dsDNA thermal stability was detected upon going from mono-

aqua complex binding to the binding of diaqua complexes pointing out that the difunctional metal

complexes are able to cross-link the two DNA strands.276 On the other hand, all of the complexes

requiring thermal activation are toxic because of the facile ligand substitution. Photochemotherapy has

the potential to minimize such toxicity due to that drug activation can be highly controlled in time and

space. Most work in this area has focused on the development of photosensitizing drugs which produce

DNA-damaging oxygen species.277 Recently, some Rh3+ and Cr3+ complexes have been shown to

undergo aquation and form covalent adducts with calf thymus DNA upon irradiation with UV light.278-

285 However, the photoreactivity of such complexes is still not sequence specific and it is desirable to

design ODN conjugates covalently linked to the photoactivatable metal complexes to precisely direct

the drug reactivity towards the sequence of interest.

From this point of view, [Ru(tpy)(DPPZ)(CH3CN)]2+-ODN conjugates can be considered as the

molecules which are activated by light owing to the photoaquation of their Ru2+-label and site

specifically cross-link to the dG residue in the complementary DNA strand after ODN•DNA duplex

formation. To prove this proposal, first, the reactivity of the non-tethered [Ru(tpy)(DPPZ)(H2O)]2+ and its

50

derivatives towards nucleic acid components and natural dsDNA has been investigated. The

monofunctional complex [Ru(tpy)(DPPZ-CONHEt)(H2O)]2+ has been photolytically prepared from its

Ru2+-CH3CN precursor and showed no reaction with thymidine, 2'-deoxycytidine, and 2'-deoxyadenosine

5'-monophosphates (TMP, dCMP, and dAMP respectively). Mixing of equimolar amounts of the

complex with 2'-deoxyguanosine 5'-monophosphate (dGMP) exhibited the formation of a product, which

is characterized by upfield shifts of 1H NMR resonances from 8.2 to 6.8 ppm for H8 and 6.2 to 6.0 for

H1'. The composition of this mixture was identified by positive mode MALDI-TOF MS (Figure 8A) as a

mixture of the starting complex, which is ionized with the loss of H2O molecule [Ru2+(tpy)(DPPZ-

CONHEt)-H+] and the product. The isotopic pattern observed for the product corresponds to a singly

charged species containing Ru2+(tpy)(DPPZ-CONHEt) (688 Da) plus the mass of the singly protonated

dGMP ([C10H13N5O7P], 346 Da). Thus, the product composition fits the expected adduct of the formula

(DPPZ-CONHEt)(tpy)Ru2+-dGMP. The photolysis of [Ru(tpy)(DPPZ)(CH3CN)]2+ in the presence of the

native duplex 34•43 revealed that the photoactivated monofunctional Ru2+ complexes bind to the

dsDNA. From the MALDI-TOF MS analysis of the photolyzed mixture (Figure 8B) it is seen that the

10mer 34 (containing only dA, dC, and T nucleotides) remained completely unaltered (calculated mass is

2945.5, the observed mass is 2944.9), while the target 11mer 43 was modified giving rise to new

m/z2500 3000 3500 4000 4500 5000 55000

3000

6000

9000

12000

15000

3500 4000 4500 50000

1000

(2944.9)34

43 (3409.2)

43+Ru2+

(4024.0)

43+2Ru2+

(4642.6)43+3Ru2+

(5263.4)

43+Ru2+

43+2Ru2+ 43

m/z600 700 800 900 1000 11000

2000

4000

6000

8000

1010 1020 1030 10400

400

800

1200

1600

Figure 8. (A) MALDI-TOF MS (positive mode) of the 1:1 reaction between the 5'-dGMP and complex [Ru(tpy)(DPPZ-CONHEt)(H2O)]2+ in the acetone-H2O mixture (5.4 ×10-3 mol l-1). (B) Negative ion MALDI-TOF MS spectra of the native duplex 34•43 (10-5 mol l-1, reaction volume - 1600 µl) photolyzed with 5 equivalent excess of [Ru(tpy)(dppz)(CH3CN)]2+ for 2 h in 20 mM PO4

3-, 0.1M NaClO4, pH 6.6 buffer and passed after photolysis through Sephadex G-25 column to remove salts. The peaks corresponding to the target ODN 43 bound with Ru2+(tpy)(dppz) residues are depicted as 43+nRu2+ (n = 1,2,3).

687.08 1034.04

Intensity IntensityA B

51

products with the observed m/z values corresponding to the molecular weight of the [43 +

Ru2+(tpy)(DPPZ)] (m/z 4024.0) and [43 + 2×Ru2+(tpy)(DPPZ)] (m/z 4642.6) adducts. It is of interest that

only target ODN 43 containing dG nucleotides is the substrate for the covalent binding of the mono-aqua

Ru2+ complexes in accordance with our observation that dAMP, dCMP, and TMP do not react with such

complexes. Thus, these data clearly esiablish that [Ru(tpy)(DPPZ)(CH3CN)]2+ and its derivatives, upon

light activation, can bind to the guanine containing compounds and to the dG nucleotides of the double

stranded DNA.

From the above experiments, it is clear that when a monofunctional Ru2+ complex is tethered to an

ODN strand of an ODN•DNA duplex it should form a cross-linkage, upon photoactivation, to the

complementary strand containing dG residue in the direct proximity to the appended Ru2+ center. Figure

9 shows the autoradiogram from the photolysis experiments immediately analyzed after irradiation (2 h)

by PAGE in denaturing conditions. All of the Ru2+-ODN conjugates 39 – 42 produced a higher molecular

1 2 3 4 5 6 7 8 9 Figure 9. Autoradiogram of a 20% denaturing polyacrylamide gel showing the formation of interstrand photo cross-linked product (P) upon irradiation of duplexes (10-5 mol l-1) formed with 5'-32P-labeled 11mer 43 and 10mer 34 or different Ru2+ labeled ODNs 39 – 42 in 20 mM PO4

3-, 0.1M NaClO4, pH 6.6 buffer. Lane 1: 42•43 without irradiation. Duplexes irradiated for 2 h: 42•43 (lane 2), 41•43 (lane 3), 39a•43 (lane 4), 39b•43 (lane 5), 40a•43 (lane 6), 40b•43 (lane 7), 34•43 (lane 8). Lane 9: 5'-32P-labeled 11mer 43. 39a and 39b stand for two fractions obtained in the course of HPLC purification of ODN 39. Corresponding two fractions of ODN 40 are designated as 40a and 40b.

weight band under light exposure, indicating the formation of a cross-linked product. As expected, the

highest yield (34%) was obtained with the ODN 42 carrying reactive centers both at the 5'- and 3'-

terminals. For the ODNs carrying only one Ru2+ complex the cross-coupling efficiency is decreased in

the order of 3'-(22%) > 5'-(9%) > middle-(7%) modified duplex which is reversible to the duplex

stabilization effect caused by the tethered metal complex (see Table 3). This trend can be explained in

P→

32P-labeled ODN 43 →

52

terms of structural rigidity of the metallointercalation site in the duplex. The more rigid structure arising

from the intercalation of Ru2+ complex by its threading through the duplex core (see chapter 2.2.4), the

higher stabilization effect and less the metal complex flexibility which dictates the accessibility of the

target dG moiety. As a consequence, the probability of the favorable structural geometry resulting in

cross-coupling reaction is less for the duplexes in which Ru2+ center is more rigidly packed within the

double helix. It is noteworthy that the electrophoretic mobility of the target strand is not effected by the

irradiation of the native duplex (lane 8, Figure 9) as well as without irradiation of Ru2+ labeled duplexes

(lane 1, Figure 9), indicating that indeed the [Ru(tpy)(DPPZ)(CH3CN)]2+ moiety exposed to the light

source is responsible for the cross-coupling effect. MALDI-TOF MS analysis of the irradiated 3'-Ru2+

modified duplex 40•43 confirmed the formation of the cross-linked product generating 1- charged ions at

m/z 7282.1 which corresponds to the sum of the m/z values observed for the non-reacted target strand 43

(m/z 3410.3) and singly Ru2+ modified ODN 40 with decoordinated CH3CN ligand (m/z 3872.8).

The time course of photoreaction examined for the 5',3'-bis-Ru2+ modified ODN 42 (Figure 10A,

lanes 1 - 6) showed that the yield of the adduct is almost steady after 1 h of irradiation achieving the

maximal value of ~33% (Figure 10B). The reason that the cross-coupling reaction proceeds till

saturation at such low yield can be two-fold: (1) the reversibility of the cross-coupling because of the low

product stability, (2) the cross-coupling is accompanied by side reactions deactivating the intermediate

aquaruthenium-ODN species. Note that both undesired processes could take place in the present system.

To explore the stability of the cross-coupling product, the reaction mixture obtained after irradiation for

5 h was kept at room temperature or 37°C and analyzed by PAGE (Figure 10A, lanes 7 – 12). One can

see that the high molecular weight band continuously disappears faster as the incubation temperature is

higher indicating that the cross-coupling product is not stable. When the 32P-labeled slow-migrating band

corresponding to the cross-coupled duplex (as in lane 5, Figure 10A) was excised and subsequently

extracted from the gel with sodium acetate (0.3 M), the analytical PAGE of the resulting extract revealed

a partial regeneration of the non-modified 32P-labeled target ODN 43 (Figure 10C).

The reversibility of the coss-coupling was also proven by MS analysis of the material obtained in

the course of isolation of the PAGE low-migrated band. For this purpose, two photo-cross-linking

reactions were performed on a 17 nmol scale for 3'-Ru2+ modified duplex 40•43 and 5',3'-bis-Ru2+

modified duplex 42•43. The photolyzed duplexes were separated by denaturing PAGE (Figure 11) and

the low-migrated bands were isolated by excision and extraction from gel analogously as it has been

done in the analytical photolysis experiments with 32P-labeled target ODN 43. Optimal separation of the

photoproduct was achived in the case of 3'-Ru2+-labeled duplex (lane 1 in Figure 11), while in the case of

5',3'-bis-Ru2+ modified duplex the electrophoretic mobility of the cross-coupled duplex was very similar

to that of the ODN 42 (lane 2 in Figure 11). UV-Shadowing PAGE at 366 nm clearly exhibits

53

lane 1 2 3 4 5 6 7 8 9 10 11 12 irradiation time 5' 10' 30' 1h 2h 5h 5h 5h 5h 5h 5h 5h

yield (%) 16.8

23.9

29.4

30.1

30.8

33.0

27.9

25.1

9.8

32.8

31.5

25.2

that the photoproducts in each case contain Ru2+ complex. MALDI-TOF MS of the isolated low-

migrated bands showed that the individual cross-coupled duplexes decompose in the course of

purification and MS fragmentation affording a mixture of three components, as was found in MS

analysis of the crude photolyzed duplex 40•43. These results clearly demonstrate that the duplexes

cross-coupled through the (DPPZ)(tpy)Ru2+-dG bond [where (DPPZ)(tpy)Ru2+ belongs to the antisense

min0 50 100 150 200 250 300

Yiel

d (%

)

0

20

40

60

80

100BA

C 1 2 3

Figure 10. (A) Autoradiogram of a 20% denaturing polyacrylamide gel showing the cross-linked duplex Pobtained after photoirradiation of duplexes (10-5 mol l-1) formed with 5'-32P-labeled 11mer 43 and 5',3'-bis-Ru2+ labeled ODN 42 in 20 mM PO4

3-, 0.1M NaClO4, pH 6.6 buffer. Lanes 1 – 6: duplex 42•43 irradiated for 5 min, 10 min, 30 min, 1 h, 2 h, or 5 h, and immediately analyzed by PAGE after photolysis. Lanes 7 – 9: duplex 42•43 irradiated for 5 h and then incubated at 37˚C for 1 h, 2h, or 15 h respectively. Lanes 10 – 12: duplex 42•43 irradiated for 5 h and then incubated at room temperature for 1 h, 2h, or 15 h respectively. (B) Plot of the yield of cross-coupling product P versus irradiation time for duplex 42•43 (10-5 mol l-1) immediately analyzed by PAGE after photolysis experiments, showing a maximum yield of 33% at the plateau. (C) Autoradiogram of a 20% denaturing polyacrylamide gel showing the high molecular weight band (P) after photolysis (2 h) of the duplex formed with 5'-32P-labeled 11mer 43 and 5',3'-bis-Ru2+ labeled ODN 42 and photolyzed for 2 h in 20 mM PO4

3-, 0.1M NaClO4, pH 6.6 buffer (lane 1), followed by excision of the high-molecular weight band (i.e. P), extraction from the gel with sodium acetate (0.3 M) overnight at room temperature, and passing of the extract through Sephadex G-25 column (lane2). Lane 2 displays the presence of P and 5'-32P-labeled 11mer 43 in ca 1:1 ratio. Lane 3: 5'-32P-labeled 11mer 43.

P→ 32P-labeled ODN 43 →

P→

32P-labeled ODN 43 →

54

Oligo T

Oligo VI

Photoproduct

1 2 21

254 nm 366 nm

Photoproduct + Oligo VIII

Figure 11. UV-Shadowing 20% denaturing polyacrylamide gel (λ = 254 nm and 366 nm) of the reaction mixtures obtained after photolysis of 3'-Ru2+ labeled duplex 40•43 (lane 1) and 5',3'-bis-Ru2+ labeled duplex 42•43 (lane 2) in 20 mM PO4

3-, 0.1M NaClO4, pH 6.6 buffer (duplex concentration in each experiment was 10-5 mol l-1, reaction volume: 1600 µl) and passed through Sephadex G-25 column to remove salts. Comparison of the bands visualized at 254 nm and 366 nm shows that the slow-migrating bands (i.e.high molecular weight photoproduct) indeed show fluorescence at 366 nm, which indicates that the photoproduct consists of Ru2+ cross-linkage. The slow-migrating photoproduct bands in each lane were excised and then extracted from the gel with sodium acetate (0.3 M). The extracts were passed through Sephadex G-25 column and analyzed by MALDI-TOF MS.

strand and dG belongs to the target strand] are not stable and can reversibly decomposed liberating the

target strand without any secondary modifications.

To understand the reason of instability of cross-coupling products formed with the target

DNA strand 43 and photochemically generated [Ru(tpy)(DPPZ)(H2O)]2+-ODN conjugates, the time

dependence of the 1:1 reaction between [Ru(tpy)(dppz-CONHEt)(H2O)]2+ and 5'-dGMP in

acetone-d6/D2O mixture was monitored by 1H NMR at various temperatures. It was found that this

reaction never goes to completion, and in fact reaches an equilibrium depending upon the reaction

condition (Figure 12A). Thus, the experimental data pointed that coordination of dGMP to the mono-

aqua Ru2+ complex belongs to reversible reaction of type, A + B ←→ X + Y, where X is [Ru(tpy)(dppz-

CONHEt)(dGMP)] and A and B are starting complex [Ru(tpy)(dppz-CONHEt)(H2O)]2+ and dGMP

respectively. The equilibrium was only slightly shifted to the product side with the decrease of

temperature, which indicates that the reaction is poorly exothermic. Fitting of the observed equilibrium

constants K to the van’t-Hoff equation (Figure 12B) gave 39.3±1.7 kJ/mol for ∆Hº and –0.038±0.006

kJ/(mol K) for ∆Sº. Our experimental results show that the product-complex, [Ru(tpy)(dppz-

CONHEt)(dGMP)], thus formed suffers reverse aquation and exists in an equilibrium with

[Ru(tpy)(dppz-CONHEt)(H2O)]2+. Next, we added an excess of CH3CN to the reaction mixture when it

has already reached the equilibrium, and incubated in the dark at room temperature. After 17 h of

55

incubation with CH3CN, 1H NMR spectra indicated the presence of starting dGMP and [Ru(tpy)(dppz-

CONHEt)(CH3CN)](PF6)2 only, which confirms the reversibility of the reaction. The poor stability of the

adduct could be a consequence of high distortions of the octahedral geometry which arise to minimize

the steric constraints between the bulky guanine and polypyridyl ligands (tpy and dppz). The present data

Figure 12. (A) Kinetic study of the 1:1 reaction between complex [Ru(tpy)(dppz-CONHEt)(H2O)]2+ and 5'-dGMP in CD3COCD3-D2O (1:1, v/v) solution at 20˚C (●), 33˚C (○), 44˚C (▼), and 56˚C (∇ ), which shows that the yield of the photo adduct varies from 80% to 60% in the temperature range of 20-56˚C. (B) Plot of RlnK vs 1/T obtained from the corresponding kinetic studies, giving the enthalpy ∆Hº, and entropy ∆Sº from the slope and intercept respectively: RlnK = ∆Sº - ∆Hº(1/T).

are in accordance with the low efficiency of the cross-coupling of the two strands in Ru2+ labeled

duplexes 39•43 - 42•43, and correlate with the thermal instability of cross-coupled duplexes. The present

study also explains the very low binding efficiency of the polypyridyl mono-aquaruthenium(II)

complexes towards the double-stranded DNA observed by Thorp.275

2.4.4 The extension of the idea of the Ru2+ complex activation via photoaquation (unpublished data)

The idea to activate the thermally stable polypyridyl Ru2+ complexes by quantative exclusion of the

labile ligands with subsequent aquation of the metal complexes under light irradiation can also be

applied for the bifunctional Ru2+ analogs. It is expected that increasing the number of sites available in

the Ru2+ coordination sphere for photoaquation and subsequent binding to nucleobases, one can

influence on tethered complex reactivity and strikingly improve the stability of the cross-linked product.

We utilized that the mixed Ru2+ complexes containing sterically congested bidentate ligands such as

dmbpy, pypz, or biq undergo clean and selective ligand photosubstitution by solvent molecules.286-294

1/T (K-1)

0.0030 0.0032 0.0034R

ln(K

)80

85

90

95

100

time (h)

Con

cent

ratio

n of

5'-d

GM

P (m

ol/l)

0.000

0.001

0.002

0.003

0.004

0.005

0.0000.0010.0020.0030.0040.005

0 6 12 18 24

0 20 40 60 80 100 120 140 160 180

BA

56

Indeed, we found that the photolysis of [Ru(bpy)2X]2+ (X = dmbpy, pypz, or biq) or

[Ru(bpy)2(CH3CN)2]2+ in the acetone-H2O mixture leads to the formation of [Ru(bpy)2(H2O)2]2+

(Scheme 6). A time course of the photolysis reactions (Figure 13) revealed that CH3CN and dmbpy

HN

NN

NRu

N

N

N

N

NCMe

O

RuN

N NCMe

NCMe

N

N

RuN

N OH2

OH2

N

N

HN

NN

NRu

N

N

N

N

H2O

O

RuN

N N

N

N

N

N

RuN

N N

N

N

N

RuN

N N

N

N

N

H2O

H2O H2O

H2O H2O

2+

2+

2+

2+

2+

2+ 2+

hνννν

hνννν hνννν

hνννν hνννν

Ru2+(bpy)2(dmbpy) Ru2+(bpy)2(H2O)2 Ru2+(bpy)2(biq)

Ru2+(bpy)2(pypz)Ru2+(bpy)2(CH3CN)2

Ru2+(tpy)(DPPZ-CONHEt)(CH3CN) Ru2+(tpy)(DPPZ-CONHEt)(H2O)

Scheme 6. Photoaquation reactions performed on mono- and bifunctional complexes.

ligands are rapidly and quantitatively substituted by water molecules while pypz and biq complexes of

Ru2+ showed a reversible kinetic decay with a characteristic equilibrium plateau. The extent of

fotolabilization in these particular systems depends on the rate constants ratio of the direct reaction of the

ligand exclusion and the back reaction of X ligand (X = CH3CN, dmbpy, biq, pypz) re-coordination to

the resulting Ru2+(H2O)2 species. Thus, pypz and biq ligands were found to re-coordinate effectively

which indicates that the strength of the ligand field along the weakest Ru2+-X axis is greater for

[Ru(bpy)2(biq)]2+ and [Ru(bpy)2(pypz)]2+ complexes. It should be noted, however, that in the presence of

a competing nucleophile, which forms a stable adduct with Ru2+ complex, the photolysis of

[Ru(bpy)2X]2+ (X = biq, pypz) should go to completion because the reactive [Ru(bpy)2(H2O)2]2+ species

is led out of the equilibrium by binding to this nucleophile. The important aspect of this photolysis

experiments is that they shows that the principle Ru2+ complexes activation by photoaquation can be

applied for the generation of the more reactive bis-aqua Ru2+ complexes. For instance, it has been shown

that [Ru(bpy)2(H2O)2]2+ reacts with 9-ethylguanine in 1:1 ratio affording the stable adduct [cis-

Ru(bpy)2(9-ethylguanine-κN7)(H2O)]2+.145 Rigid Ru2+ complexes of the form [Ru(diimine)2(H2O)2]2+ are

57

quite stable with respect to the dissociation of diimine ligands on one hand, and on the other hand, they

enantioselectively binds to B-DNA.295,296 As a matter of fact, covalent binding of cis-[Ru(bpy)2(H2O)2]2+

time (min)0 200 400 600 800 1000

% o

f sta

rting

Ru2+

com

plex

0

20

40

60

80

100

0 20 40 600

30

60

90

Figure 13. Time course study of the photolysis of metal complex solutions in CD3COCD3-D2O (7:4, v/v) mixture: Ru2+(bpy)2(CH3CN)2 (●), Ru2+(tpy)(DPPZ)(CH3CN) (○), and Ru2+(bpy)2X, where X stands for dmbpy (■), biq(▼), or pypz (∇ ) liberating ligand. Complex concentration in each experiment was 6.1×10-3 mol l-1. Solutions were irradiated in a NMR tube positioned in 3 cm in front of lens of a Kodak slide projector (250-W halogen lamp) and analyzed by 1H NMR spectroscopy in definite time intervals.

to dsDNA has been reported as an undesired side effect of probing DNA with a derivative of

[Ru(bpy)3]2+.177 However, covalent binding of polypyridyl mono- and bis-aqua Ru2+ complexes is not yet

very well developed, and therefore it is of considerable necessity to comparatively study the binding of

mono- and bifunctional Ru2+ compounds to guanine derivatives. Further work is in progress to evaluate

the stability of binding products of bifunctional Ru2+ complexes to guanosine nucleosides and to tether

the stable precursors of bis-aqua Ru2+ compounds to ODNs.

58

3. Acknowledgements This work was carried out at the Department of Bioorganic Chemistry, Biomedical Center, Uppsala University, Uppsala, during the years 1996-2002. I would like to express my appreciation to my supervisor Professor Jyoti Chattopadhyaya for giving me this opportunity to work and learn in his well-equipped laboratory, for introducing me to the field of nucleic acids chemistry and its biological applications, for stimulating and constructive criticism, thoughtful guidance throughout the years and valuable discussions. I wish to express my deep gratitude to all the former and present colleagues at the Department of Bioorganic Chemistry: Parag Acharya, Sandipta Acharya, Peter Agback, Jharna Barman, Somer Bekiroglu, Alexei Denisov, Krishna Kumar Dubey, Melker Holmer, Johan Isaksson, Anil Kumar, Mrinal Kanti Kundu, Brian Lawrence, Xiaoguang Luo, Ingrid Luyten, Tatiana Maltseva, Jan Milecki, Frans Peder Nilson, Dimitry Ovcharenko, Catherine Petit, Janez Plavec, Nitin Puri, Anders Sandström, Christian Sund, Christophe Thibaudeau, Anna Trifonova, Andrei Vasiliev, Irina Velikian, Shun-ichi Yamakage, Peter Weiler, Pei Wen and Edouard Zamaratski for their contribution in creation of exiting scientific atmosphere on the Department. I am specially grateful to: Pradeep Kumar for his splendid human character and readiness to help András Földesi who helped me a lot with synthesis, instruments and in chemicals search Nariman Amirkhanov for the fruitful scientific discussions which allowed me to understand many not trivial aspects of my work Oles Plashkevich for solution of computer problems Gopal Datta for his brotherhood and the same excitement by magic of mountains Kristiina Komminaho, Pernilla Brohage, Rabab Elkarib and Jaana Öst for their kind assitance, availability and competent secretarial work. Alexander Papchikhin for importing me to Sweden Suresh Gohil (Department of Chemistry, Swedish University of Agricultural Sciences) for the rare ability to share difficulties, which I had a lot with my last project, and for the qualified MS analysis. I am deeply thankful to Konstantin Kovalev, the only person I’ve ever met abroad, with whom I felt myself most easy and comfortable – for his attractively ironic point of view on things. My warmest thanks I express to Alex and Olga Shebanov for polemic discussions throughout where we sat and drank, drank, and drank. My work in Uppsala would never be somehow possible without that warm and re-animating atmosphere created by my friends, Yuri Khotyaintsev and Tania Leonova, Maxim and Larisa Smirnov, Oleg Travnikov, Svjatoslav Korepov and Lena Vlasova, Konstantin Matveev, Tatyana Tentler, Mikis and Mette Larsen and many others. I am very grateful to Line Strand and Joel Sohlberg for taking care of my son and exporting of Swedish traditions to my family. I would like to thank my wife Elena for love, patience, creating the home comfort which served as a buffer against aggressive “chemical medium” present in a chemist life, and for my son Daniil, who is specially acknowledged for his love and that endless energy reservoir which I have now in his person on my back. Last but not least, I thank my mother and father who gave me life, love, and everything to be the person who I am. I thank my sister Sasha, mother-in-law, and all relatives for their love, support, and waiting, waiting, and waiting for me all these years.

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4. References

1 Zamecnik, P. C.; Stephenson, M. L. Proc. Natl. Acad. Sci. USA 1978, 75, 280 2 Stephenson, M. L.; Zamecnik, P. C. Proc. Natl. Acad. Sci. USA 1978, 75, 285 3 Uhlmann, E.; Peyman, A. Chem. Rev. 1990, 90, 543 4 De Mesmaeker, A.; Häner, R.; Martin, P.; Moser, H. E. Acc. Chem. Res. 1995, 28, 366 5 Agrawal, S.; Kandimalla, E. Mol. Med. Today 2000, 6, 72 6 Toulme, J.-J. Nat. Biotechnol. 2001, 19, 17 7 Braasch, D. A.; Corey, D. R. Biochemistry 2002, 41, 4503 8 Watson, J. D.; Crick, F. H. C. Nature 1953, 171, 737 9 Oligodeoxynucleotides. Antisense Inhibitors of Gene Expression, Ed. Cohen, J. S.; Topics in Molecular and Structural Biology, MacMillan, 1989 10 Hélène, C.; Saison-Behmoaras, T. Medicine/Science, 1994, 10, 257 11 Thuong, N. T., Hélène, C. Angew. Chem. Int. Ed. Engl. 1993, 32, 666 12 Sun, J.-S.; Garestier, T., Hélène, C. Curr. Opin. Struct. Biol. 1996, 6, 327 13 Vasquez, K. M.; Wilson, J. Trends Biochem. Sci. 1998, 23, 4 14 Hoogsteen, K. Acta Crystallogr. 1959, 12, 822 15 Stain, C. A.; Cheng, Y.-C. Science, 1993, 261, 1004 16 Moser, H. E.; Dervan, P. B. Science, 1987, 238, 645 17 Povsic, T. J.; Dervan, P. B. J. Am. Chem. Soc. 1990, 112, 9428 18 Bertrand, J.-R.; Imbach, J.-L.; Paoletti, C. Biochem. Biophys. Res. Commun. 1989, 164, 311 19 Blacke, K. R.; Murakami, A.; Spitz, S. A. Biochemistry 1985, 24, 6139 20 Boiziau, C.; Kurfurst, R.; Cazenave, C. Nucleic Acids Res. 1991, 19, 1113 21 Johansson, H. E.; Belsham, G. J.; Sproat, B. S. Nucleic Acids Res. 1994, 22, 4591 22 Haeuptle, M. T.; Frank, R.; Dobberstain, B. Nucleic Acids Res. 1986, 14, 1427 23 Minshull, J.; Hunt, T. Nucleic Acids Res. 1986, 14, 6433 24 Knorre, D. G.; Vlassov, V. V. Progr. Nucleic Acids Res. Mol. Biol. 1985, 32, 291 25 Goodchild, J. Bioconjugate Chem. 1990, 1, 165 26 Lerman, L. S. J. Mol. Biol. 1961, 3, 18 27 Asseline, U.; Thuong, N. T.; Hélène, C. New J. Chem. 1997, 21, 5 28 Lokhov, S.; Podyminogin, M. A.; Sregeev, D. S.; Silnikov, V. N.; Kutyavin, I. V.; Sishkin, G. V.; Zarytova, V. P. Bioconjugate Chem. 1992, 3, 414 29 Balbi, A.; Sottofattori, E.; Grandi, T.; Mazzei, M. Tetrahedron 1994, 50, 4009 30 Asseline, U.; Delarue, M.; Lancelot, G.; Toulme, F.; Thuong, N. T.; Montenay-Garestier, T.; Hélène, C. Proc. Natl. Acad. Sci. USA 1984, 81, 3297 31 Asseline, U.; Bonfils, E.; Dupret, D.; Thuong, N. T. Bioconjugate Chem. 1996, 7, 369 32 Fukui, K.; Morimoto, M.; Segawa, H.; Tanaka, K.; Shimidzu, T. Bioconjugate Chem. 1996, 7, 349 33 Fukui, K.; Iwane, K.; Shimidzu, T.; Tanaka, K. Tetrahedron Lett. 1996, 37, 4983 34 Fukui, K.; Tanaka, K.; Nucleic Acids Res. 1996, 24, 3962 35 Asseline, U.; Thuong, N. T. Nucleosides & Nucleotides 1991, 10, 359 36 Lin, K.-Y.; Matteuchi, M. Nucleic Acids Res. 1991, 19, 3111 37 Ono, A.; Dan, A.; Matsuda, A. Bioconjugate Chem. 1993, 4, 499 38 Dan, A.; Yoshimura, Y.; Ono, A.; Matsuda, A. Bioorg. Med. Chem. Lett. 1993, 3, 615 39 Chen, J. K.; Carlson, D. V.; Weith, H. L.; O’Brien, J. A.; Goldman, M. E.; Cushman, M. Tetrahedron Lett. 1992, 33, 2275 40 Mann, J. S.; Shibata, Y.; Meehan, T. Bioconjugate Chem. 1992, 3, 554 41 Yamana, K.; Ohashi, Y.; Nunota, K.; Kitamura, M.; Nakana, H.; Sangen, O.; Shimidzu,

60

T. Tetrahedron Lett. 1991, 32, 6347 42 Telser, J.; Cruickshank, K. A.; Morrison, L. E.; Netzel, T. L.; J. Am. Chem. Soc. 1989, 111, 9428 43 Dikalov, S. I.; Rumyantseva, G. V.; Weiner, L. M.; Sergejev, D. S.; Frolova, E. I.; Godovikova, T. S.; Zarytova, V. F. Chem. Biol. Interactions 1991, 77, 325 44 Kasai, H.; Goto, M.; Ikeda, K.; Zama, M.; Takemura, S.; Matsuura, S.; Sugimoto, T.; Goto, T. Biochemistry 1976, 15, 898 45 Yamana, K.; Aota, R.; Nakano, H. Tetrahedron Lett. 1995, 46, 8427 46 Sun, J. S.; François, J.-C.; Montenay-Garestier, T.; Saison-Behmoaras, T.; Roig, V.; Thuong, N. T.; Hélène, C. Proc. Natl. Acad. Sci. USA, 1989, 86, 9198 47 Collier, D. A.; Thuong, N. T.; Hélène, C. J. Am. Chem. Soc. 1991, 113, 1457 48 Collier, D. A.; Mergny, J.-L.; Thuong, N. T.; Hélène, C. Nucleic Acids Res. 1991, 19, 4219 49 Montenay-Garestier, T.; Sun, J. S.; Chomilier, J.; Mergny, J. L.; Takasugi, M.; Asseline, U.; Thuong, N. T.; Rougée, M.; Hélène, C. In Molecular Basis of Specificity in Nucleic Acid-Drug-Interactions, Pullman, B.; Lortner, J. Eds.; Kluwer, Dordrecht, 1990, 275 50 Takasugi, M.; Guendouz, A.; Chassignol, M.; Decout, J. L.; Lhomme, J.; Thuong, N. T.; Hélène, C. Proc. Natl. Acad. Sci. USA, 1991, 88, 5602 51 Garbesi, A.; Bonazzi, S.; Zanella, S.; Capobianco, M. L.; Giannini, G.; Arcamone, F. Nucleic Acids Res. 1997, 25, 2121 52 Gianolio, D. A.; McLaughlin, L. W. J. Am. Chem. Soc. 1999, 121, 6334 53 Puri, N.; Zamaratski, E.; Sund, C.; Chattopadhyaya, J. Tetrahedron 1997, 53, 10409 54 Silver, G. C.; Nguyen, C. H.; Boutorine, A. S.; Bisagni, E.; Garestier, T.; Hélène, C. Bioconjugate Chem. 1997, 8, 15 55 Silver, G. C.; Sun, J.-S.; Nguyen, C. H.; Boutorine, A. S.; Bisagni, E.; Hélène, C. J. Am. Chem. Soc. 1997, 119, 263 56 Horne, D. A.; Dervan, P. B. J. Am. Chem. Soc. 1990, 112, 2435 57 Ono, A.; Chen, L. S.; Kan, L. S. Biochemistry 1991, 30, 9914 58 Froehler, B. C.; Terhorst, T.; Shaw, J. P.; McCurdy, S. N. Biochemistry 1992, 31, 1603 59 Sun, J. S.; Giovannangeli, C.; François, J. C.; Kurfurst, R.; Montenay-Garestier, T.; Asseline, U.; Saison-Behmoaras, T.; Thuong, N. T.; Hélène, C. Proc. Natl. Acad. Sci.

USA, 1991, 88, 6023 60 Horne, D. A.; Dervan, P. B. Nucleic Acids Res. 1991, 19, 4963 61 Rahakrishnan, I.; Patel, D. J. J. Mol. Biol. 1994, 241, 600 62 Huang, C.-Y.; Bi, G.; Miller, P. S. Nucleic Acids Res. 1996, 24, 2606 63 Zhou, B. W.; Puga, E.; Sun, J. S.; Garestier, T.; Hélène, C. J. Am. Chem. Soc. 1995, 117, 10425 64 Kukreti, S.; Sun, J. S.; Garestier, T.; Hélène, C. Nucleic Acids Res. 1997, 25, 4264 65 Milligan, J. F.; Matteucci, M. D.; Martin, J. C. J. Med. Chem. 1993, 36, 1923 66 Zamaratski, E.; Pradeepkumar, P. I.; Chattopadhyaya, J. J. Biochem. Biophys. Methods 2001, 48, 189 67 Cazenave, C.; Stain, C. A.; Loreau, N.; et al. Nucleic Acids Res. 1989, 17, 4255 68 Gagnor, C.; Bertrand, J.-R.; Thenet, S.; Lemaitre, M.; Morvan, F.; Rayner, B.; Malvy, C.; Lebleu, B.; Imbach, J.-L.; Paoletti, C. Nucleic Acids Res. 1987, 15, 10419 69 Crooke, S. T. Biochimica et Biophysica Acta 1999, 1489, 31 70 Lesnik, E. A.; Guinosso, C. J.; Kawasaki, A. M.; Sasmor, H.; Zounes, M.; Cummins, L. L.; Ecker, D. J.; Cook, P. D.; Freier, S. M. Biochemistry 1993, 32, 7832 71 Monoharan, M. Biochimica et Biophysica Acta 1999, 1489, 117 72 Martin, P. Helv. Chim. Acta 1995, 78, 486 73 Kawasaki, A. M.; Casper, M. D.; Freier, S. M.; Lesnik, E. A.; Zounes, M. C.;

61

Cummins, L. L.; Gonzalez, C.; Cook, P. D. J. Med. Chem. 1993, 36, 831 74 Wengel, J. Acc. Chem. Res. 1999, 32, 301 75 Furdon, P. J.; Dominski, Z.; Kole, R. Nucleic Acids Res. 1989, 17, 9193 76 Agrawal, S.; Mayrand, S. H.; Zamecnik, P. C.; Pederson, T. Proc. Natl. Acad. Sci. USA 1990, 87, 1401 77 Morvan, F.; Sanghvi, Y. S.; Perbost, M.; Vasseur, J.-J.; Bellon, L. J. Am. Chem. Soc. 1996, 118, 255 78 Gryaznov, S. M. Biochimica et Biophysica Acta 1999, 1489, 131 79 Rait, V. K.; Shaw, B. R. Antisense Nucleic Acid Drug Dev. 1999, 9, 53 80 Damha, M. J.; Wilds, C. J.; Noronha, A.; Brukner, I.; Borkow, G.; Arion, D.; Parniak, M. A. J. Am. Chem. Soc. 1998, 120, 12976 81 Noronha, A. M.; Wilds, C. J.; Lok, C.-N.; Viazovkina, K.; Arion, D.; Parniak, M. A.; Damha, M. J. Biochemistry 2000, 39, 7050 82 Manisov, G.; Teplova, M.; Nielsen, P.; Wengel, J.; Egli, M. Biochemistry 2000, 39, 3525 83 Giles, R. V.; Tidd, D. M. Nucleic Acids Res. 1992, 20, 763 84 Agrawal, S.; Jiang, Z.; Zhao, Q.; Shaw, D.; Cai, Q.; Roskey, A.; Channavajjala, L.; Saxinger, C.; Zhang, R. Proc. Natl. Acad. Sci. USA 1997, 94, 2620 85 Argawal, S. Biochimica et Biophysica Acta 1999, 1489, 53 86 Altmann, K.-H.; Fabbro, D.; Dean, N. M.; Geiger, T.; Monia, B. P.; Nicklin, P. Biochem. Soc. Trans. 1996, 24, 630 87 Lok, C.-N.; Viazovkina, E.; Min, K.-L.; Nagy, E.; Wilds, C. J.; Damha, M. J.; Parniak, M. A. Biochemistry 2002, 41, 3457 88 Wahlestedt, C.; Salmi, P.; Good, L.; Kela, J.; Johnsson, T.; Hökfelt,T.; Broberger, C.;

Porreca, F.; Lai, J.; Ren, K.; Ossipov, M.; Koshkin, A.; Jakobsen, N.; Skouv, J.; Oerum, H.; Jacobsen, M. H.; Wengel, J. Proc. Natl. Acad. Sci. USA 2000, 97, 5633 89 Kurreck, J.; Wyszko, E.; Gillen, C.; Erdmann, V. A. Nucleic Acids Res. 2002, 30, 1911 90 Pradeepkumar, P. I.; Zamaratski, E.; Földesi, A.; Chattopadhyaya, J. Tetrahedron Lett. 2000, 41, 8601 91 Pradeepkumar, P. I.; Zamaratski, E.; Földesi, A.; Chattopadhyaya, J. J. Chem. Soc., Perkin Trans. 2, 2001, 402 92 Pradeepkumar, P. I.; Chattopadhyaya, J. J. Chem. Soc., Perkin Trans. 2, 2001, 2074 93 Puri, N.; Chattopadhyaya, J. Nucleosides & Nucleotides, 1999, 18, 2785 94 Cazenave, C.; Loreau, N.; Thoung, N. T.; Toulme, J.-J.; Hélène, C. Nucleic Acids Res. 1987, 15, 4717 95 Godard, G.; François, J.-C.; Duroux, I.; Asseline, U.; Chassignol, M.; Thuong, N.; Hélène, C.; Saison-Behmoaras, T. Nucleic Acids Res. 1994, 22, 4789 96 Boutorine, A. S.; Brault, D.; Takasugi, M.; Delgado, O.; Hélène, C. J. Am. Chem. Soc. 1996, 118, 9469 97 Kutyavin, I. V.; Gamper, H. B.; Gall, A. A.; Meyer, R. B.; J. Am. Chem. Soc. 1993, 115, 9303 98 Maruenda, H.; Tomasz, M. Bioconjugate Chem. 1996, 7, 541 99 Tsubouchi, A.; Bruice, T. C. J. Am. Chem. Soc. 1995, 117, 7399 100 Trawick, B. N.; Daniher, A. T.; Bashkin, J. K. Chem. Rev. 1998, 98, 939 101 Bashkin, J. K.; Frolova, E. I.; Sampath, U. J. Am. Chem. Soc. 1994, 116, 5981 102 Matsumura, K.; Endo, M.; Komiyama, M. J. Chem. Soc., Chem. Commun. 1994, 2019 103 Magda, D.; Miller, R. A.; Sessler, J. L.; Iverson, B. L. J. Am. Chem. Soc. 1994, 116, 7439 104 Magda, D.; Crofts, S.; Lin, A.; Miles, D.; Wright, M.; Sessler, J. L. J. Am. Chem. Soc. 1997, 119, 2293

62

105 Magda, D.; Wright, M.; Crofts, S.; Lin, A.; Sessler, J. L. J. Am. Chem. Soc. 1997, 119, 6947 106 Baker, B. F.; Ramasamy, K.; Keily, J. Bioorg. Med. Chem. Lett. 1996, 6, 1647 107 Takeda, N.; Imai, T.; Irisawa, M.; Sumaoka, J.; Yashiro, M.; Shigekawa, H.; Komiyama, M. Chem. Lett. 1996, 599 108 Chu, B. C.; Orgel, L. E. Proc. Natl. Acad. Sci. USA 1985, 82, 963 109 Lin, S.-B.; Blake, K. R.; Miller, P. S.; Ts’o, P. O. Biochemistry 1989, 28, 1054 110 Moser, H. E.; Dervan, P. B. Science 1987, 238, 645 111 Strobel, S. A.; Dervan, P. B. Science 1990, 249, 73 112 Han, H.; Dervan, P. B. Proc. Natl. Acad. Sci. USA 1993, 90, 3806 113 François, J.-C.; Saison-Behmoaras, T.; Chassignol, M.; Thuong, N. T.; Sun, J.-S.; Hélène, C. Biochemistry 1988, 27, 2272 114 Chen, C. B.; Sigman, D. C. Proc. Natl. Acad. Sci. USA 1986, 83, 7147 115 François, J.-C.; Saison-Behmoaras, T.; Chassignol, M.; Thuong, N. T.; Hélène, C.; J. Biol. Chem. 1989, 264, 5891 116 Chen, C. B.; Sigman, D. C. J. Am. Chem. Soc. 1988, 110, 6570 117 François, J.-C.; Saison-Behmoaras, T.; Barbier, C.; Chassignol, M.; Thuong, N. T.; Hélène, C.; Proc. Natl. Acad. Sci. USA 1989, 86, 9702 118 Le Doan, T.; Perrouault, L.; Chassignol, M.; Thuong, N. T.; Hélène, C. Nucleic Acids Res. 1987, 15, 8643 119 Frolova, E. I.; Fedorova, O. S.; Knorre, D. G. Biochimie, 1993, 75, 5 120 Sergeev, D. S.; Zarytova, V. F.; Mamaev, S. V.; Godovikova, T. S.; Vlassov, V. V. Antisense Res. Dev. 1992, 2, 235 121 Sergeev, D. S.; Vorobjev, P. E.; Zarytova, V. F. Nucleosides & Nucleotides 1997, 16,

1575 122 Carter, P. J.; Cheng, C.-C.; Thorp, H. H. J. Am. Chem. Soc. 1998, 120, 632 123 Carter, P. J.; Cheng, C.-C.; Thorp, H. H. Inorg. Chem. 1996, 35, 3348 124 Cheng, C.-C.; Goll, J. G.; Neyhart, G. A.; Welch, T. W.; Singh, P.; Thorp, H. H. J. Am. Chem. Soc. 1995, 117, 2970 125 Pyle, A. M.; Chiang, M. Y.; Barton, J. K. Inorg. Chem. 1990, 29, 4487 126 Murphy, C. J.; Arkin, M. R.; Jenkins, Y.; Ghatlia, N. D.; Bossmann, S. H.; Turro, N. J.;

Barton, J. K. Science 1993, 262, 1025 127 Hall, D. B.; Holmlin, R. E.; Barton, J. K. Nature 1996, 382, 731 128 Daniel, B. H.; Barton, J. K. J. Am. Chem. Soc. 1997, 119, 5045 129 Tossi, A. B.; Kelly, J. M. Photochem. Photobiol. 1989, 49, 545 130 Sentagne, C.; Chambron, J.-C.; Sauvage, J.-P.; Paillous, N. J. Photochem. Photobiol. B 1994,

26, 165 131 Moucheron, C.; Kirsch-De Mesmaeker, A.; Kelly, J. M. J. Photochem. Photobiol. B 1997, 40,

91 132 Kirsch-De Mesmaeker, A.; Moucheron, C.; Boutonnet, N. J. Phys. Org. Chem. 1998, 11, 566 133 Ortmans, I.; Content, S.; Boutonnet, N.; Kirsch-De Mesmaeker, A.; Bannwarth, W.; Constant,

J.-F.; Defrancq, E.; Lhomme, J. Chem. Eur. J. 1999, 5, 2712 134 Arkin, M. R.; Stemp, E. D. A.; Pulver, S. C.; Barton, J. K. Chem. Biol. 1997, 4, 389 135 Gelasco, A.; Lippard, S. J. Anticancer Activity of Cisplatin and Related Complexes; Clarke, M.

J.; Sadler, P. J., Ed.; “Metallopharmaceuticals I : DNA Interactions”, Springer-Verlag Berlin Heidelberg: Berlin, 1999, p117

136 Mestroni, G.; Alessio, E.; Calligaris, M.; Attia, W. M.; Quadrifoglio, F.; Cauci, S.; Sava, G.; Zorzet, S.; Pacor, S.; Monti-Bragadin, C.; Tamaro, M.; Dolzani, L. In Ruthenium and Other Non-Platinum Metal Complexes in Cancer Chemotherapy (Clarke, M. J. ed.), Springer-Verlag, Heidelberg, 1989, p74

63

137 Nováková, O.; Kašpárková, J.; Vrána, O.; van Vliet, P. M.; Reedijk, J.; Brabec, V. Biochemistry 1995, 34, 12369

138 Cheng, C.-C.; Lee, W.-L.; Su, J.-G.; Liu, C.-L. J. Chin. Chem. Soc. 2000, 47, 213 139 van Vliet, P. M.; Toekimin, S. M. S.; Haasnoot, J. G.; Reedijk, J.; Nováková, O.; Vrána, O.;

Brabec, V. Inorg. Chim. Acta 1995, 231, 57 140 Armor, J. personal communication 1970 141 Bottomley, F. Can. J. Chem. 1977, 55, 2788 142 Cauci, S.; Viglino, P.; Esposito, G.; Quadrifoglio, F. J. Inorg. Biochem. 1991, 43, 739 143 Esposito, G.; Cauci, S.; Fogolari, F.; Alessio, E.; Scocchi, M.; Quadrifoglio, F.; Viglino, P.

Biochemistry 1992, 31, 7094 144 Anagnostopoulou, A.; Moldrheim, E.; Katsaros, N.; Sletten, E. J. Biol. Inorg. Chem. 1999, 4,

199 145 van Vliet, P. M.; Haasnoot, J. G.; Reedijk, J. Inorg. Chem. 1994, 33, 1934 146 Vlassov, V. V.; Gorn, V. V.; Ivanova, E. M.; Kazakov, S. A.; Mamaev. S. V.; FEBS Lett.

1983, 162, 286 147 Gruff, E. S.; Orgel, L. E. Nucleic Acids Res. 1991, 19, 6849 148 Colombier, C.; Lippert, B.; Leng, M. Nucleic Acids Res. 1996, 24, 4519 149 Bernal-Méndez, E.; Sun, J.-S.; Francisco, G.-V.; Leng, M. New J. Chem. 1998, 1479 150 Manchanda, R.; Dunham, S. U.; Lippard, S. J. J. Am. Chem. Soc. 1996, 118, 5144 151 Schliepe, J.; Berghoff, U.; Lippert, B.; Cech, D. Angew. Chem. Int. Ed. Engl. 1996, 35, 646 152 Berghoff, U.; Schmidt, K.; Janik, M.; Schröder, G.; Lippert, B. Inorg. Chim. Acta 1998, 269,

135 153 Schmidt, K. S.; Filippov, D. V.; Meeuwenoord, N. J.; van der Marel, G. A.; van Boom, J. H.;

Lippert, B.; Reedijk, J. Angew. Chem. Int. Ed. Engl. 2000, 39, 375 154 Zoltewicz, J. A.; Clark, D. F.; Sharpless, T. W.; Grahe, G. J. Am. Chem. Soc. 1970, 92, 1741 155 Gaucheron, F.; Malinge, J.-M.; Blacker, A. J.; Lehn, J.-M.; Leng, M. Proc. Natl. Acad. Sci.

USA 1991, 88, 3516 156 Payet, D.; Gaucheron, F.; Sip, M.; Leng, M. Nucleic Acids Res. 1993, 21, 5846 157 Kehrmann, F.; Havas, E. Ber. 1913, 46, 341 158 Sinha, N. D.; Biernat, J.; McManus, J.; Köster, H. Nucleic Acids Res. 1984, 12, 4539 159 McBride, L.; Caruthers, M. Tetrahedron Lett. 1983, 24, 245 160 Holy, A. Nucleic Acids Res. 1974, 1, 289 161 Oligonucleotide Synthesis. A Practical Approach; Gait, M. J., Ed.; IRL Press: Oxford, 1984, p 45 162 Misiura, K.; Durrant, I.; Evans, M. R.; Gait, M. J. Nucleic Acids Res. 1990, 18, 4345 163 Theisen, P.; McCollum, C.; Upadhya, K.; Jacobson, K.; Vu, H.; Andrus, A. Tetrahedron Lett. 1992, 33, 5033 164 Willis, M. C.; Collins, B.; Zhang, T.; Green, L. S.; Sebesta, D. P.; Bell, C.; Kellogg, E.; Gill, S. C.; Magallanez, A.; Knauer, S.; Bendele, R. A.; Gill, P. S.; Janjić, N.

Bioconjugate Chem. 1998, 9, 573 165 Ing; Manske J. Chem. Soc. 1926, 2348 166 Pon, R. T.; Yu, S. Tetrahedron Lett. 1997, 38, 3331 167 Sullivan, B. P.; Salmon, D. J.; Meyer, T. J. Inorg. Chem. 1978, 17, 3334 168 Hartshorn, R. M.; Barton, J. K. J. Am. Chem. Soc. 1992, 114, 5919 169 Sullivan, B. P.; Calvert, J. M.; Meyer, T. J. Inorg. Chem. 1980, 19, 4104 170 Takeuchi, K. J.; Thompson, M. S.; Pipes, D. W.; Meyer, T. J. Inorg. Chem. 1984, 23, 1845 171 Pramanik, N. C. ; Pramanik, K.; Ghosh, P.; Bhattacharya, S. Polyhedron 1998, 17, 1525 172 Laemmel, A.-C.; Collin, J.-P.; Sauvage, J.-P.; C. R. Acad. Sci. Paris 2000, 3, 43

64

173 Paul, R.; Anderson, G. W. J. Am. Chem. Soc. 1960, 82, 4596 174 Hu, X.; Smith, G. D.; Sykora, M.; Lee, S. J.; Gristaff, M. W. Inorg. Chem. 2000, 39, 2500 175 Khan, S. I.; Beilstein, A. E.; Sykora, M.; Smith, G. D.; Hu, X.; Gristaff, M. W. Inorg. Chem. 1999, 38, 3922 176 Kelly, J. M.; Tossi, A. B.; McConnel, D. J.; OhUigin, C.; Hélène, C.; Le Doan, T. In Free Radicals, Metal Ions and Biopolymers (Beaumont, P. C.; Deedle, D. J.; Parsons, B. J.; Rice-Evans, C., eds.), Richelieu Press, London, 1989, p 143 177 Telser, J.; Cruickshank, K. A.; Schanze, K. S.; Netzel, T. N. J. Am. Chem. Soc. 1989, 111, 7221 178 Bannwarth, W.; Schmidt, D.; Stallard, R. L.; Hornung, C.; Knorr, R.; Müller, F. Helv. Chim. Acta 1988, 71, 2085 179 Bannwarth, W.; Schmidt, D. Tetrahedron Lett. 1989, 30, 1513 180 Jenkins, Y.; Barton, J. K.; J. Am. Chem. Soc. 1992, 114, 8736 181 Meggers, E.; Kursch, D.; Giese, B. Helv. Chim. Acta 1997, 80, 640 182 Hurley, D. J.; Tor, Y. J. Am. Chem. Soc. 1998, 120, 2194 183 Wiederholt, K.; McLaughlin, L. W. Nucleic Acids Res. 1999, 27, 2487 184 Khan, S. I.; Beilstain, A. E.; Gristaff, M. W. Inorg. Chem. 1999, 38, 418 185 Khan, S. I.; Beilstain, A. E.; Sykora, M.; Smith, G. D.; Hu, X.; Gristaff, M. W. Inorg. Chem. 1999, 38, 3922 186 Khan, S. I.; Beilstain, A. E.; Tierney, M. T.; Sykora, M.; Gristaff, M. W. Inorg. Chem. 1999, 38, 5999 187 Lewis, F. D.; Helvoigt, S. A.; Letsinger, R. L. Chem. Commun. 1999, 327 188 Rack, J. J.; Krider, E. S.; Meade, T. J. J. Am. Chem. Soc. 2000, 122, 6287 189 Grimm, G. N.; Boutorine, A. S.; Lincoln, P.; Hélène, C. Nucleosides & Nucleotides 2001, 20, 909 190 Vargas-Baca, I.; Mitra, D.; Zulyniak, H. J.; Banerjee, J.; Sleiman, H. F. Angew. Chem. Int. Ed. 2001, 40, 4629 191 Dandliker, P. J.; Holmlin, R. E.; Barton, J. K. Science, 1997, 275, 1465 192 Holmlin, R. E.; Dandliker, P. J.; Barton, J. K. Bioconjugate Chem. 1999, 10, 1122 193 Suen, H.-F.; Wilson, S. W.; Pomerantz, M.; Walsh, J. L. Inorg. Chem. 1989, 28, 786 194 Hecker, C. R.; Fanwick, P. E.; McMillin, D. R. Inorg. Chem. 1991, 30, 659 195 Hartshorn, S. M.; Maxwell, K. A.; White, P. S.; DeSimone, J. M.; Meyer, T. J. Inorg.

Chem. 2001, 40, 601 196 Serron, S. A.; Aldridge III, W. S.; Danell, R. M.; Meyer, T. J. Tetrahedron Lett. 2001, 41, 4039 197 Sinha, N. D.; Biernat, J.; McManus, J.; Koster, H. Nucleic Acids Res. 1984, 12, 4539. 198 Sinha, N. D.; Biernat, J.; Koster, H. Tetrahedron Lett. 1983, 24, 5843. 199 Ren, S.; Cai, L.; Segal, B. M. J. Chem. Soc., Dalton Trans. 1999, 1413 200 Cai, L.; Lim, K.; Ren, S.; Cadena, R. S.; Beck, W. T. J. Med. Chem. 2001, 44, 2959 201 Fagalde, F.; Lis de Katz, N. D.; Katz, N. E. Polyhedron 1997, 16, 1921 202 Naal, Z.; Tfouni, E.; Benedetti, A. V. Polyhedron 1994, 13, 133 203 Weizman, H.; Tor, Y. J. Am. Chem. Soc. 2001, 123, 3375 204 Potier, N.; Van Dorsselaer, A.; Cordier, Y.; Roch, O.; Bischoff, R. Nucleic Acids Res. 1994, 22, 3895 205 Apffel, A.; Chakel, J. A.; Fischer, S.; Lichtenwalter, K.; Hancock, W. Anal. Chem. 1997, 69, 1320 206 Huber, C. G.; Buchmeiser, M. R. Anal. Chem. 1998, 70, 5288 207 De Bellis, G.; Salani, G.; Battaglia, C.; Pietta, P.; Rosti, E.; Mauri, P. Rapid Commun. Mass Spectrom. 2000, 14, 243

65

208 Koomen, J. M.; Russell, W. K.; Tichy, S. E.; Russell, D. H. J. Mass. Spectrom. 2002, 37, 357 209 Hiort, C.; Norden, B.; Rodger, A. J. Am. Chem. Soc. 1990, 112, 1971 210 Kelly, J. M.; Tossi, A. B.; McConnel, D. J.; OhUigin, C. Nucleic Acids Res. 1985, 13, 6017 211 Rehmann, J.; Barton, J. K. Biochemistry 1990, 29, 1701 212 Barton, J. K. J. Biol. Struct. Dyn. 1983, 1, 621 213 Long, E. C.; Barton, J. K. Acc. Chem. Res. 1990, 13, 271 214 Hiort, C.; Linkoln, P.; Norden, B. J. Am. Chem. Soc. 1993, 115, 3448 215 Friedman, A. E.; Chambron, J. C.; Sauvage, J.-P.; Turro, N. J.; Barton, J. K. J. Am. Chem. Soc. 1990, 112, 4960 216 Dupureur, C. M.; Barton, J. K. J. Am. Chem. Soc. 1994, 116, 10286 217 Jenkins, Y.; Friedman, A. E.; Turro, N. J.; Barton, J. K. Biochemistry 1992, 31, 10809 218 Barawkar, D.; Rajeev.; Kumar, V.; Ganesh, K. Nucleic Acids Res. 1996, 24, 1229 219 Choi, S.-D.; Kim, M.-S.; Linkoln, P.; Tuite, E.; Norden, B. Biochemistry 1997, 36, 214 220 Cho, C.-B.; Cho, T.-S.; Kim, S. K.; Kim, B.-J.; Han, S. W.; Jung, M. J. Bull. Korean Chem. Soc. 2000, 21, 995 221 Tinoco, I.; Sauer, K.; Wang, J. C. In Physical Chemistry: Principles and Applications in Biological Scinces (Young, D.; Cavanaugh, D., Eds.), Prentice Hall, Upper Saddle River, NJ, 1995, p 559 222 Allen, F. S.; Gray, D. M.; Roberts, G. P.; Tinoco, I. Biopolymers 1972, 11, 853 223 Fasman, G. Handbook of Biochemistry and Molecular Biology, CRC Press: Ohio, 1975, Vol. 1, p 589 224 Eriksson, M.; Leijon, M.; Hiort, C.; Norden, B.; Gräslund, A. Biochemistry 1994, 33, 5031 225 Lomonossoff, G. P.; Butler, P. J. G.; Klug, A. J. Mol. Biol. 1981, 149, 745 226 Fox, K. R.; Waring, M. J. Nucleic Acids Res. 1984, 12, 9271 227 Fox, K. R. In Methods in Molecular Biology (Fox, K. R., Ed.), Humana Press Inc., Totowa, NJ, Vol. 90, p 1 228 Suck, D.; Oefner, C. Nature 1986, 321, 620 229 Önfelt, B.; Linkoln, P.; Norden, B. J. Am. Chem. Soc. 1999, 121, 10846 230 Davis, T. M.; McFail-Isom, L.; Keane, E.; Williams, L. D. Biochemistry 1998, 37, 6975 231 Li, J.; Wartell, R. M. Biochemistry 1998, 37, 5154 232 Lima, W. F.; Mohan, V.; Crooke, S. T. J. Biol. Chem. 1997, 29, 18191 233 Salazar, M.; Fedoroff, O. Y.; Miller, J. M.; Riberio, N. S.; Reid, B. R. Biochemistry 1993, 32, 4207 234 Fedoroff, O. Y.; Salazar, M.; Reid, B. R. J. Mol. Biol. 1993, 233, 509 235 Uchiyama, Y.; Iwai, S.; Ueno, Y.; Ikehara, M.; Ohtsuka, E. J. Biochem. 1994, 116, 1322 236 Uchiyama, Y.; Miura, Y.; Inoue, H.; Ohtsuka, E.; Ueno, Y.; Ikehara, M.; Iwai, S. J. Mol. Biol. 1994, 4, 782 237 Amirkhanov, N. V.; Chattopadhyaya, J. J. Chem. Soc., Perkin Trans. 2 2002, 271 238 Amirkhanov, N. V.; Pradeepkumar, P. I.; Chattopadhyaya, J. J. Chem. Soc., Perkin Trans. 2 2002, in press 239 Amirkhanov, N. V.; Zamaratski, E.; Chattopadhyaya, J. Tetrahedron Lett. 2001, 42, 489 240 Temsamani, J.; Tang, J.-Y.; Padmapriya, A.; Kubert, M.; Argawal, S. Antisense Res. Dev. 1993, 3, 277 241 Nicklin, P. L.; Ambler, J.; Mitchelson, A.; Bayley, D.; Phillips, J. A.; Craig, S. J.;

66

Monia, B. P. Proc. Natl. Acad. Sci. USA 1997, 16, 1145 242 Shaw, J.-P.; Kent, K.; Fishback, J.; Froehler, B. Nucleic Acids Res. 1991, 19, 747 243 Boutorin, A. S.; Guskova, L. V.; Ivanova, E. M.; Kobetz, N. D.; Zarytova, V. F.; Ryte, A. S.; Yurchenko, L. V.; Vlassov, V. V. FEBS Lett. 1989, 254, 129 244 Ryte, A. S.; Karamyshev, V. N.; Nechaeva, M. N.; Guskova, L. V.; Ivanova, E. M.; Zarytova, V. F.; Vlassov, V. V. FEBS Lett. 1992, 299, 124 245 Hélène, C. Current Opinion of Biotechnology 1993, 4, 29 246 Hall, J.; Hüsken, D.; Häner, R. Nucleic Acids Res. 1996, 24, 3522 247 Daniher, A. T.; Bashkin, J. K. J. Chem. Soc., Chem. Commun. 1998, 1077 248 Trawick, B. N.; Osiek, T. A.; Bashkin, J. K. Bioconjugate Chem. 2001, 12, 900 249 Putnam, W. C.; Bashkin, J. K. J. Chem. Soc., Chem. Commun. 2000, 767 250 Åström, H.; Strömberg, R. Nucleosides, Nucleotides & Nucleic Acids, 2001, 20, 1385 251 Kodak, A. S.; Gard, J. K.; Merriman, M. C.; Winkeler, K. A.; Bashkin, J. K.; Stern, M. K. J.

Am. Chem. Soc. 1991, 113, 283 252 Spassky, A.; Sigman, D. S. Biochemistry 1985, 24, 8050 253 Hélène, C.; Thuong, N. T.; Saison-Behmoaras, T.; François, J. C. Trends Biotechnol. 1989, 7,

310 254 Sigman, D. S. Biochemistry 1990, 29, 9097 255 Sigman, D. S.; Mazumder, A.; Perrin, D. M. Chem. Rev. 1993, 93, 2295 256 Sigman, D. S.; Bruice, T. W.; Mazumder, A.; Sutton, C. L. Acc. Chem. Res. 1993, 26, 98 257 Patviel, G.; Bernadou, J.; Meunier, B. Angew. Chem. Int. Ed. Engl. 1995, 34, 746 258 Cowan, J. A. Curr. Opin. Chem. Biol. 2001, 5, 634 259 Baker, A. D.; Morgan, R. J.; Strekas, T. C. J. Chem. Soc., Chem. Commun. 1992, 1099 260 Gallagher, J.; Chen, C. B.; Pan, C. Q.; Perrin, D. M.; Cho, Y.-M.; Sigman, D. S. Bioconjugate Chem. 1996, 7, 413 261 Meijler, M. M.; Zelenko, O.; Sigman, D. S. J. Am. Chem. Soc. 1997, 119, 1135 262 Baudoin, O.; Teulade-Fichou, M.-P.; Vigneron, J.-P.; Lehn, J.-M. Chem. Commun. 1998, 2349 263 Pitie, M.; Meunier, B. Bioconjugate Chem. 1998, 9, 604 264 Knorre, D. G.; Vlassov, V. V.; Zarytova, V. F.; Lebedev, A. V.; Fedorova, O. S. Design and targeted reactions of oligonucleotide derivatives; CRC Press: 1994 265 Ebright, R. H.; Ebright, Y. W.; Pendergrast, P. S.; Gunasekera, A. Proc. Natl. Acad. Sci. USA 1990, 87, 2882 266 Chen, C. B.; Mazumder, A.; Constant, J.-F.; Sigman, D. S. Bioconjugate Chem. 1993, 4,

69 267 Bergstrom, D. E.; Gerry, N. P. J. Am. Chem. Soc. 1994, 116, 12067 268 Chen, C. B.; Gorin, M. B.; Sigman, D. S. Proc. Natl. Acad. Sci. USA 1993, 90, 4206 269 Chen, C. B.; Milne, L.; Landgraf, R.; Perrin, D. M.; Sigman, D. S. ChemBioChem 2001, 2, 735 270 Gupta, N.; Grover, N.; Neyhart, G. A.; Liang, W.; Singh, P.; Thorp, H. H. Angew. Chem. Int. Ed. Engl. 1992, 31, 1048 271 Ho, C.; Che, C.-M.; Lau, T.-C. J. Chem. Soc., Perkin Trans. 1990, 967 272 Rosenberg, B.; Vancamp, L.; Krigas, T. Nature 1965, 205, 689 273 Rosenberg, B.; Vancamp, L.; Trosko, J. E.; Mansour, V. H. Nature 1969, 222, 385 274 Clarke, M. J.; Stubbs, M. Interactions of Metallopharmaceuticals with DNA; Sigel, A.; Sigel, H., Ed.; “Metal Ions in Biological Systems”, Marcel Dekker Inc.: New York, 1996, vol. 32, p 727 275 Grover, N.; Gupta, N.; Thorp, H. H. J. Am. Chem. Soc. 1992, 114, 3390 276 Grover, N.; Welch, T. W.; Fairlay, T. A.; Cory, M.; Thorp, H. H. Inorg. Chem. 1994, 33, 3544

67

277 Oseroff, A. R.; Ara, G.; Ohuoha, D.; Aprille, J.; Bommer, J. C.; Yarmush, M. L.; Foley, J.; Cincotta, L. Photochem. Photobiol. 1987, 46, 83 278 Billadeau, M. A.; Morrison, H. Photolytic Covalent Binding of Metal Complexes to DNA; Sigel, A.; Sigel, H., Ed.; “Metal Ions in Biological Systems”, Marcel Dekker Inc.: New York, 1996, vol. 33, p 269 279 Mahnken, R. E.; Bina, M.; Deibel, R. M.; Luebke, K.; Morrison, H. Photochem. Photobiol. 1989, 49, 519 280 Mahnken, R. E.; Billadeau, M. A.; Niconowicz, E. P.; Morrison, H. J. Am. Chem. Soc. 1992, 114, 9253 281 Harmon, H. L.; Morrison, H. Inorg. Chem. 1995, 34, 4937 282 Billadeau, M. A.; Wood, K. V.; Morrison, H. Inorg. Chem. 1994, 33, 5780 283 Mohammad, T.; Tessman, I.; Morrison, H.; Kennedy, M. A.; Simmonds, S. W. Photochem. Photobiol. 1994, 59, 189 284 Mohammad, T.; Chen, C.; Guo, P.; Morrison, H. Bioorg & Med. Chem. Lett. 1999, 9, 1703 285 Billadeau, M. A.; Morrison, H. J. Inorg. Biochem. 1995, 57, 249 286 Steel, P. J.; Lahousse, F.; Lerner, D.; Marzin, C. Inorg. Chem. 1983, 22, 1488 287 von Zelewski, A.; Gremaud, G. Helv. Chim. Acta 1988, 71, 1108 288 Laemmel, A.-C.; Collin, J.-P.; Sauvage, J.-P. Eur. J. Inorg. Chem. 1999, 383 289 Collin, J.-P.; Laemmel, A.-C.; Sauvage, J.-P. New J. Chem. 2001, 25, 22 290 Baranoff, E.; Collin, J.-P.; Furusho, Y.; Laemmel, A.-C.; Sauvage, J.-P. Chem. Commun. 2000, 1935 291 Wang, R.; Vos, J. G.; Schmehl, R. H.; Hage, R. J. Am. Chem. Soc. 1992, 114, 1964 292 Buchanan, B. E.; Degn, P.; Pavon Velasco, J. M.; Hughes, H.; Creaven, B. S.; Long, C.; Vos, J. G.; Howie, R. A.; Hage, R.; van Diemen, J. H.; Haasnoot, J. G.; Reedijk, J. J. Chem. Soc., Dalton Trans. 1992, 1177 293 Buchanan, B. E.; Hughes, H.; van Diemen, J. H.; Hage, R.; Haasnoot, J. G.; Reedijk, J.; Vos, J. G. Chem. Commun. 1991, 300 294 Baranoff, E.; Collin, J.-P.; Furusho, J.; Furusho, Y.; Laemmel, A.-C.; Sauvage, J.-P. Inorg. Chem. 2002, 41, 1215 295 Barton, J. K.; Lolis, E. J. Am. Chem. Soc. 1985, 107, 708 296 Danishefsky, A. Ph.D. Thesis, Columbia University, 1987