UC San Diego UC San Diego Electronic Theses and Dissertations

Home , Jerry Donohue

Title Design, synthesis and application of novel fluorescent nucleosides

Permalink https://escholarship.org/uc/item/5w84v342

Author Greco, Nicholas Joseph

Publication Date 2008

Peer reviewed|Thesis/dissertation

eScholarship.org Powered by the California Digital Library University of California UNIVERSITY OF CALIFORNIA, SAN DIEGO

Design, synthesis and application of novel fluorescent nucleosides.

A Dissertation submitted in partial satisfaction of the requirements for the degree

Doctor of Philosophy

Chemistry

Nicholas Joseph Greco

Committee in charge:

Professor Yitzhak Tor, Chair Professor Simpson Joseph Professor David Kleinfeld Professor Charles L. Perrin Professor Jerry Yang

2008

Nicholas Joseph Greco, 2008

The dissertation of Nicholas Joseph Greco is approved, and is acceptable in quality and form for publication on microfilm:

______

______Chair

University of California, San Diego

2008

iii DEDICATION

to those whom without I would be nothing, but with I am everything my parents, Joseph and Barbara my brother, Anthony

iv EPIGRAPH

“There is a time in every man's education when he arrives at the conviction that envy is ignorance; that imitation is suicide.” Ralph Waldo Emerson

v TABLE OF CONTENTS

SIGNATURE PAGE ...... iii

DEDICATION...... iv

EPIGRAPH ...... v

TABLE OF CONTENTS...... vi

LIST OF FIGURES...... ix

LIST OF SCHEMES ...... xiii

LIST OF TABLES ...... xiv

ACKNOWLEDGEMENTS...... xv

VITA ...... xvii

ABSTRACT...... xviii

Chapter 1. Nucleic acids: The need for fluorescent nucleosides...... 1

1.1 History of nucleic acids ...... 1

1.2 Primary structure of nucleic acids ...... 2

1.3 Watson-Crick base pairing...... 4

1.4 Secondary structure of nucleic acids ...... 5

1.5 Biological roles of the 3 major polymorphs...... 11

1.6 Using fluorescence spectroscopy to probe DNA ...... 12

1.7 Applications of fluorescent nucleosides...... 12

1.8 Major families of fluorescent nucleosides...... 13

1.9 Aims of the thesis ...... 33

1.10 References...... 34

Chapter 2. Design, synthesis and application of novel fluorescent nucleosides ...... 51

2.1 Fluorescent nucleoside design principles...... 51

2.2 Inspiration ...... 53

vi 2.3 Synthesis of thymidine analogues...... 54

2.4 Isomorphicity of modified nucleosides ...... 55

2.5 Photophysical properties of thymidine analogues...... 56

2.6 Comparison to 2-aminiopurine ...... 58

2.7 Fluorescence quenching...... 60

2.8 Furanyl cytidine, adenosine and guanosine analogues...... 61

2.9 Synthesis of furanyl cytidine analogue...... 62

2.10 Photophysical properties of furanyl cytidine analogue ...... 63

2.11 Synthesis of furanyl adenosine and guanosine analogues ...... 64

2.12 Photophysical properties of furanyl adenosine and guanosine analogues ...... 65

2.13 Effect of temperature on the emission of furanyl decorated nucleosides...... 66

2.14 Photophysical evaluation of furanyl decorated nucleosides...... 67

2.15 Phosphoramidite synthesis of furanyl thymidine nucleoside...... 69

2.16 Site specific incorporation of furanyl thymidine nucleoside...... 69

2.17 Characterizing the furanyl modified oligonucleotide ...... 72

2.18 Stability of modified duplexes ...... 75

2.19 Absorption spectra of furanyl dT containing oligonucleotides...... 77

2.20 Steady state emission spectra of furanyl dT containing oligonucleotides ...... 77

2.21 Investigation of additional duplexes ...... 79

2.22 Thermal denaturation of additional duplexes ...... 80

2.23 Photophysical properties of additional duplexes...... 82

2.24 Effects of temperature on the photophysical properties of furanyl containing

duplexes...... 85

2.25 Enzymatic incorporation of furanyl dT analogue...... 87

2.26 Site specific incorporation of furanyl dC analogue...... 88

2.27 Improving the photophysical properties of furanyl dT analogue ...... 92

vii 2.28 Concluding remarks ...... 96

2.29 Acknowledgements ...... 97

Appendix - Experimental information ...... 97

2.30 References...... 122

Chapter 3. Exploring the polarity of DNA grooves ...... 128

3.1 Introduction ...... 128

3.2 Expression of polarity...... 130

3.3 Probe design...... 132

3.4 Polarity reference scale ...... 141

3.5 Polarity of DNA grooves using furanyl dT analogue ...... 145

3.6 Polarity conversion graphs ...... 151

3.7 Conclusion and future plans ...... 154

3.8 Acknowledgements ...... 160

Appendix - Experimental information ...... 160

3.9 References...... 168

viii LIST OF FIGURES

Figure 1.1 Structures of nucleotides ...... 3

Figure 1.2 Sugar puckering in nucleic acids...... 4

Figure 1.3 Hydrogen bonding pattern of DNA bases...... 5

Figure 1.4 Major and minor grooves of DNA...... 6

Figure 1.5 The three major polymorphs of DNA ...... 7

Figure 1.6 B-form DNA ...... 8

Figure 1.7 A-form DNA ...... 9

Figure 1.8 Z-form DNA...... 10

Figure 1.9 Isomorphic bases ...... 14

Figure 1.10 Hydrogen bonding pattern of 2-aminopurine...... 15

Figure 1.11 Emission energy and quantum yield verses microenvironment polarity ...... 15

Figure 1.12 Mispairing of 2-aminopurine and cytidine...... 16

Figure 1.13 Pteridines ...... 18

Figure 1.14 Expanded nucleobases ...... 21

Figure 1.15 Structures of NAD+, wyosine and C...... 22

Figure 1.16 Chromophoric base analogues ...... 25

Figure 1.17 Base pairing of coumarin nucleoside...... 27

Figure 1.18 Conjugated base analogues...... 28

Figure 1.19 Base pairing of pyrene nucleoside...... 30

Figure 1.20 Size expanded BDFs ...... 31

Figure 1.21 Base pairing of BPP with dG and dA...... 32

Figure 2.1 Absorption and emission spectra of furan, benzene and 2-phenylfuran ...... 53

Figure 2.2 Thymidine analogues ...... 54

ix Figure 2.3 X-ray structures of furanyl, thienyl, oxazolyl and thiazolyl dT analogues...... 56

Figure 2.4 Absorption and emission spectra of furanyl, thienyl, oxazolyl and thiazolyl dT

analogues ...... 57

Figure 2.5 Emission energy and Stokes shift verses microenvironment polarity for 2-

aminopurine...... 59

Figure 2.6 Stern-Volmer plot of furanyl dT analogue and 2-aminopurine ...... 61

Figure 2.7 Structures of furanyl decorated nucleosides ...... 62

Figure 2.8 Absorption and emission spectra of furanyl dT and dC analogues ...... 64

Figure 2.9 Absorption and emission spectra of furanyl A and G analogues ...... 66

Figure 2.10 Temperature dependent emission spectra of furanyl decorated nucleosides ...... 67

Figure 2.11 Emission area and energy verses microenvironment polarity for furanyl decorated

nucleosides...... 68

Figure 2.12 Sequences of modified and control oligonucleotides ...... 70

Figure 2.13 Solid phase DNA phosphoramidite synthesis cycle ...... 71

Figure 2.14 Digest of modified oligonucleotide visualized by MALDI-TOF MS ......

...... 73

Figure 2.15 Digest of modified oligonucleotide visualized by HPLC ...... 74

Figure 2.16 Thermal denaturation of modified and control duplexes ...... 76

Figure 2.17 Absorption spectra of furanyl dT containing oligonucleotide...... 77

Figure 2.18 Emission spectra of furanyl dT containing duplexes ...... 79

Figure 2.19 Thermal denaturation of modified and additional control duplexes ...... 81

Figure 2.20 Emission spectra of furanyl dT containing duplexes at high salt concentrations ..83

Figure 2.21 Site specific detection of an abasic site via emission enhancement of furanyl dT

containing oligonucleotide ...... 84

Figure 2.22 Base pairing of the furanyl dT analogue...... 85

x Figure 2.23 Thermal denaturation of modified duplexes monitored by absorption and emission

...... 86

Figure 2.24 Thermal denaturation of modified duplexes monitored by emission ...... 87

Figure 2.25 Sequences of primer and template ...... 88

Figure 2.26 DNA polymerase reaction...... 88

Figure 2.27 Sequence modified oligonucleotide ...... 90

Figure 2.28 Digest of modified oligonucleotide visualized by HPLC ...... 91

Figure 2.29 Retrosynthetic analysis of cyclized furanyl dT analogue...... 93

Figure 3.1 Visual description of the reference scale approach...... 129

Figure 3.2 Lippert-Mataga equation...... 130

Figure 3.3 Jablonski diagram ...... 130

Figure 3.4 Crystal structure of Hoechst 33258 bound to a DNA duplex...... 133

Figure 3.5 Structures of fluorescent nucleosides used to determine groove polarity...... 134

Figure 3.6 Models of DNA duplexes containing dT and amidodansyl dU ...... 136

Figure 3.7 Stokes shift verses microenvironment polarity for groove polarity probes...... 138

Figure 3.8 Structure of dT, furanyl dT and acetylated furanyl dT ...... 139

Figure 3.9 Models of DNA duplexes containing dT and furanyl dT ...... 140

Figure 3.10 Absorption and emission spectra of furanyl dT and acetylated furanyl dT in various

solvent mixtures ...... 141

Figure 3.11 Probes for ET(30) polarity determination of solvents ...... 142

Figure 3.12 Polarity reference scale ...... 144

Figure 3.13 Corrected polarity reference scale ...... 144

Figure 3.14 Sequences of modified and unmodified oligonucleotides ...... 145

Figure 3.15 Thermal denaturation of modified and control duplexes ...... 146

Figure 3.16 Absorption, emission and CD spectra of furanyl dT containing and control

duplexes...... 148

xi Figure 3.17 Absorption spectra of furanyl dT containing and control duplexes ......

...... 149

Figure 3.18 Polarity conversion plots...... 152

Figure 3.19 Comparison of major groove polarities...... 153

Figure 3.20 Structures of novel nucleoside furanyl analogues ...... 155

Figure 3.21 Models of DNA duplexes containing furanyl dT and extended furanyl dT

analogues ...... 156

Figure 3.22 Absorption and emission spectra of thienyl extended dT analogues ...... 159

Figure 3.23 X-ray structures of extended thiophene dT analogues...... 160

xii LIST OF SCHEMES

Scheme 2.1 Synthesis of furanyl and thiophene dT analogues...... 54

Scheme 2.2 Synthesis of oxazolyl and thiazolyl dT analogues ...... 55

Scheme 2.3 Synthesis of furanyl dC analogue ...... 63

Scheme 2.4 Synthesis of furanyl A and G analogues ...... 65

Scheme 2.5 Synthesis of furanyl dT phosphoramidite ...... 69

Scheme 2.6 Synthesis of the triphosphate form of furanyl dT analogue...... 88

Scheme 2.7 Synthesis of furanyl dC phosphoramidite...... 89

Scheme 2.8 Synthesis towards the cyclized furanyl dT analogue...... 95

Scheme 3.1 Synthesis of novel thienyl extended dT analogues ...... 157

xiii LIST OF TABLES

Table 1.1 Helical parameters for the three major DNA polymorphs ...... 11

Table 2.1 Photophysical properties of furanyl, thienyl, oxazolyl and thiazolyl dT analogues ...58

Table 2.2 Photophysical properties of furanyl dT analogue...... 58

Table 2.3 Photophysical properties of 2-aminopurine ...... 59

Table 2.4 Photophysical properties of furanyl decorated nucleosides ...... 68

Table 2.5 MALDI-TOF MS of modified and control oligonucleotides...... 72

Table 2.6 Modified furanyl dT containing oligonucleotide composition ...... 75

Table 2.7 Tm values for modified and control duplexes ...... 76

Table 2.8 Tm values for modified and additional control duplexes ...... 82

Table 2.9 Tm values for modified and additional control duplexes at high salt concentrations.83

Table 2.10 Modified furanyl dC containing oligonucleotide composition...... 90

Table 3.1 Photophysical properties of furanyl dT in dioxane-water mixtures ...... 143

Table 3.2 Photophysical properties of acetylated furanyl dT MCH-iPrOH mixtures ...... 143

Table 3.3 Tm values for modified and control duplexes ...... 146

Table 3.4 Photophysical properties of furanyl dT containing duplexes ...... 151

xiv ACKNOWLEDGEMENTS

It has been said that no one accomplishes anything on their own, but only with the help and support of others. This is especially true in ones graduate career. I would like to thank my parents, Joseph and Barbara, my stepmother, Roz, and my brothers, Anthony, Michael and

Timothy for their constant support and unwavering love.

I would like to thank my “partner in crime” Victor Tam for his constant support and advise over my entire graduate career. I have been lucky enough to work with some extremely talented and driven Post-Docs over the years that still inspire me to this very day. They are

Dr. Kenneth Blount, Dr. Michelle Hysell, Dr. Renatus Sinkeldam and Dr. S. G. Srivatsan. The following people have provided support through-out my graduate career, both in and out of the lab and to them I am indebted: Matthew Belousoff, Dr. Paola Castaldi, Maia Carnevali, Dr.

Pradip Chakraborty, Derek Fischer, Qin Qin Gao, Dr. Phoebe Glazer, Dr. David Jaramillo, Dr.

Damien Jouvenot, Dr. Charles Liu, Erin Olson, Dr. Susan Del Valle, Dr. Beth Wilson, Yun Xie and Dr. Grace Yang. I have had the opportunity to work with some very talented undergrads, specifically Kay Buchner, Joe Goldenberg, Denise Kwong, Baia Lasky, Sing Lam, Paul

Marcus, Dan Palacios and Samar Yalda. I would like to thank them for their support over the years. I am indebted to Prof. Arnold Rheingold, Dr. Antonio Dipasquale and Dr. Lev for their crystallography work.

I would like to thank my committee for their stimulating classes and support during my graduate work. I would like to thank Prof. Yitzhak Tor for the environment, guidance and support that allowed me to grow into the scientist I am today. His focus and chemical understanding is a constant inspiration.

In regards to the material in this dissertation, please note the following:

Portions of Chapter 2 have been previously published in the following articles: (1) "Furan decorated nucleoside analogues as fluorescent probes: synthesis, photophysical evaluation,

xv and site-specific incorporation" Tetrahedron 2007, 63, 3515–3527 by N. J. Greco and Y. Tor.

(2) "Synthesis and site-specific incorporation of a simple fluorescent pyrimidine" Nat. Protoc.

2007, 2, 305–316 by N. J. Greco and Y. Tor. (3) "Simple fluorescent pyrimidine analogues detect the presence of DNA abasic sites" J. Am. Chem. Soc. 2005, 127, 10784–10785 by N.

J. Greco and Y. Tor. The dissertation author was the primary investigator and author of these papers. While Portions of Chapter 3 have been previously published in the article "Polarity of major grooves explored using an isosteric emissive nucleosides" ChemBioChem in Press

2007 by R. W. Sinkeldam, N. J. Greco and Y. Tor. The dissertation author was one of the primary investigators and author of this paper.

xvi VITA

Education

University of California, San Diego – La Jolla, CA Ph.D. Chemistry – March 2008

University of California, San Diego – La Jolla, CA M.S. Chemistry – March 2004

Stonehill College – North Easton, MA B.S. Chemistry – May 2002

Honors and Awards

Teddy Traylor Graduate Fellowship, University of California, San Diego, 2007 Excellence in Teaching Award; University of California, San Diego, 2003 William C. LaPlante Memorial Scholarship; Stonehill College, 2002 Undergraduate Research Experience Scholar; Stonehill College, 1999

Publications

1. Sinkeldam, R. W.; Greco, N. J.; Tor, Y. "Polarity of major grooves explored using an isosteric emissive nucleosides" ChemBioChem in Press 2008.

2. Greco, N. J.; Tor, Y. "Furan decorated nucleoside analogues as fluorescent probes: synthesis, photophysical evaluation, and site-specific incorporation" Tetrahedron 2007, 63, 3515–3527.

3. Greco, N. J.; Tor, Y. "Synthesis and site-specific incorporation of a simple fluorescent pyrimidine" Nat. Protoc. 2007, 2, 305–316.

4. Greco, N. J.; Hysell, M.; Goldenberg, J. R.; Rheingold, A. L.; Tor, Y. "Alkyne- containing chelating ligands: synthesis, properties and metal coordination of 1,2- di(quinolin-8-yl)ethyne" Dalton Trans. 2006, 2288–2290.

5. Greco, N. J.; Tor, Y. "Simple fluorescent pyrimidine analogues detect the presence of DNA abasic sites" J. Am. Chem. Soc. 2005, 127, 10784–10785.

xvii ABSTRACT OF THE DISSERTATION

Design, synthesis and application of novel fluorescent nucleosides.

Nicholas Joseph Greco

Doctor of Philosophy in Chemistry

University of California, San Diego, 2008

Professor Yitzhak Tor, Chair

Nucleic acids, the biomolecules of life, have demanded the attention of chemists and biologists for greater than a century. The ability to investigate, in detail, the structure, function and the interplay between the two has driven the development of novel approaches for the investigation of these concepts. Fluorescence spectroscopy, a technique that protein chemists have utilized for many years due to the inherent favorable photophysical properties of select naturally occurring amino acids, has gained tremendous interest by nucleic acids scientists in recent decades. The lack of naturally occurring nucleosides with favorable photophysical properties has driven the design and application of novel fluorescent nucleosides. Towards this pursuit, we have described guidelines that will enable us to systematically identify novel heterocycles with both favorable photophysical and structural properties.

Utilizing our design principles, we have explored the idea of conjugating small five membered ring heterocycles, namely furan, thiophene, oxazole and thiazole, to both the pyrimidine and purine nuclei. This has resulted in seven emissive nucleosides, of which two

xviii have emerged with desirable photophysical properties after incorporation into oligonucleotides. The furanyl thymidine analogue has been shown to positively and non- destructively detect the presence of abasic sites. This is an important advancement since most methods used to detect abasic sites result in the destruction of the target strand. The furanyl thymidine analogue has also redefined the polarity of the grooves in various DNA polymorphs. In addition, it has been shown that DNA polymerase accepts the triphosphate form of the furanyl thymidine analogue, thereby making it attractive to a wider biochemical audience. In the push to investigate more than just thymidine analogues, a furanyl cytidine analogue has been site specifically incorporated into an oligonucleotide, where its photophysical properties are currently being evaluated. These results, identified above and expanded on in the following chapters, clearly indicate the utility of and need for novel fluorescent nucleosides.

xix Chapter 1

Nucleic acids: The need for fluorescent nucleosides.

1.1 – History of nucleic acids

For over a century, scientists have been intrigued by the relationship between structure and

function in nucleic acids. Their central role in passing hereditary information was the most

attractive and compelling motivation for their investigation. In the past 140 years, numerous

scientists drove nucleic acid research from discovery to the forefront of modern science. The

discovery of nucleic acids is attributed to Friedrich Miescher,1 who in 1868 discovered a

phosphorus containing substance that he termed “nuclein”. Phoebus A. Levene and Walter A.

Jacobs proposed the primary structure of nucleic acids.2-10 Their predicted cyclic structure

where four nucleotides were connected in a large ring, was incorrect but did establish the

phosphodiester linkage. It was only later that crystal structures by Klein and Thannhauser11-13

and chemical synthesis by Sir Alexander Todd14-20 solidified the primary structure of nucleic

acids. Erwin Chargaff,21 not persuaded that experimental error resulted in variations from

equimolar base ratios, diligently determined that the proportions of A and T, being equal to

each other, was higher than C and G, thus discrediting the cyclic hypothesis suggested by

Levene. This set the stage for the discovery of the secondary structure of DNA. The first diffraction pattern seen by William Astbury identified DNA as having a regular structure,22

whereby only 16 years later, Francis Crick and James Waston23,24 utilizing diffraction patterns

from Marurice Wilkins25 and Rosalind Franklin26 determined the anti-parallel double helical structure of DNA. The function of nucleic acids, first hypothesized by Edmund B. Wilson in

1895,27 was later confirmed by the work of many research groups, most notably those of

Marshall Nirenberg28 and Severo Ochoa.29,30 The multidisciplinary idea, that chemists have a unique skill-set to solve biological questions is one that was pioneered in the 1940/50s by

1 2

Fredrick Sanger31 and Gobind Khorana.32,33 It is with their monumental work that we now have entire fields that thrive at the interface of chemistry and biology.

1.2 – Primary structure of nucleic acids

Nucleic acids are long polymers composed of repeating units called nucleotides, which in

turn are composed of (1) a nitrogen containing heterocyclic base, (2) a pentose sugar ring and

(3) a phosphate group.

Heterocyclic bases The five nitrogen rich heterocyclic bases found in DNA/RNA (figure 1.1 in blue) are divided

into the monocyclic heterocycles, known as the pyrimidines (thymine, uracil, and cytosine) and

the bicyclic heterocycles, known as the purines (adenine and guanine). DNA is composed of

guanine (G), adenine (A), cytosine (C) and thymine (T), while RNA sees the substitution of

uracil (U) for thymine. This substitution (T U) in RNA, while not exclusive, does have

various biological implications.35,37 The heterocyclic bases are connected to the furanose

sugars via a -glycosidic bond, placing the base on the same side as the 5 hydroxyl (figure

1.1).

Pyrimidines H N H O O O O O O 5 O O P 3 N P N H P N H O 5’ 1 O O O N O N O N O 1’ O O O O O 2 Na 3’ 2 Na 2 Na HO R HO R HO R Cytidine 5'-phosphate Thymidine 5'-phosphate Uridine 5'-phosphate

Purines H O O O O N N H O N 7 P O P O 5 N O N 1 N O N H O 3 O N N 2 Na 2 Na N H HO R HO R H Adenosine 5'-phosphate Guanosine 5'-phosphate

Figure 1.1. Structures of nucleotides in DNA (R=H) and RNA (R=OH).

Pentose sugar The pentose sugar is found in either the D-ribose form (RNA) or the deoxy-D-ribose form

(DNA) as seen in figure 1.1 (red). The furanose sugars in DNA and RNA exist in a “puckered”

conformation minimizing non-bonded interactions and maximizing H-bonding interactions with

neighboring substituents.34,35 This type of puckering is distinguished by denoting the atom,

normally C2 or C3, that is out of the plane created by the remaining atoms, namely C1-O4-

C4 (figure 1.2). These two conformations, in solution, are in equilibrium with an energy

separation of less than 25 kJ mol1.36 Despite this minimal energetic separation between conformations, C2-endo is normally seen in B-form DNA, while C3-endo is typically seen in

A-form DNA/RNA and a mixture of both is seen in Z-form DNA (table1.1).34,35

C5' C2' C3' B B C5'

C3' C2'

C2'-endo C3'-endo

Figure 1.2. Depiction of the preferred sugar puckered conformations seen in nucleic acids.

Phosphate group The nucleosides (sugar and base) are connected to each other via phosphodiester bonds

from the 5-hydroxyl of one nucleoside to the 3-hydroxyl of the next nucleoside (figure 1.2 in

green). The phosphate group connects nucleosides in a polymer-like chain where the

termination of the chain ends in either the 5-OH or 3-OH of the nucleoside, thus

distinguishing the direction of the chain.

1.3 – Watson-Crick base pairing

Early crystal structures of nucleotides (dTMP and dGMP) were depicted in their enolic

tautomer form (figure 1.3b), making it difficult to clearly identify the hydrogen-bonding pattern

that exists between two DNA strands. However, upon the suggestion by Jerry Donohue,38,39

dTMP and dGMP were manipulated into their keto forms, thus making the dC•dG and dT•dA

hydrogen bonding pattern evident to James Watson. Figure 1.3a depicts Watson-Crick type

hydrogen bonding interactions between complementary bases, where the hydrogen bond

donors, highlighted in red, are paired with the hydrogen bond acceptors, highlighted in blue.

In the canonical base pairing of dC•dG, three hydrogen bonds are formed via the interaction of

N1 and N2 of dG that donate hydrogens to the electronegative O2 and N3 of dC, with the N4

of dC donating a hydrogen to the electronegative O6 of dG to complete the pattern. In a

similar fashion, a dA•dT base pair, having only two hydrogen bonds, is formed via each base

(N6 of dA and N3 of dT) donating a hydrogen to the electronegative N1 of dA and O4 of dT.

Although depicted below in the deoxy-form, the same canonical hydrogen bonding patterns

exist for RNA, with U replacing dT. Typical N-N and N-O hydrogen bond distances range from

2.8–2.95Å with average strengths of 610 kJ mol1.35

(a) H H N H O N O H N N 7 5 5 3 N H N 1 N N H N N 1 3 N N dR N N dR O dR O H N dR H dC dG keto dT keto dA

(b) H H H H N H O N O H N N

N N N N N N N N dR N N dR dR O H N dR O H dC dG enolic dT enolic dA

Figure 1.3. Hydrogen bonding pattern of DNA bases in their (a) keto form and (b) enolic form where the hydrogen bond donors are highlighted in red and the hydrogen bond acceptors are highlighted in blue.

1.4 – Secondary structure of nucleic acids

With the correct hydrogen bonding pattern in hand and a diffraction pattern depicting a

monoclinic C2 symmetry, Watson and Crick were able to determine the secondary structure of

DNA, specifically an anti-parallel double helix.24 In their native conformation, the heterocyclic bases are found in an anti conformation, where the hydrogen bonding face is projected away

from the sugar moiety. This allows for the negatively charged phosphate groups to be pointed

outward toward the polar media, while the aromatic bases point inwards toward each other

creating a hydrophobic cavity. The anti-parallel helix generates two grooves that run along the

helical axis, appropriately named the major and minor grooves. The major and minor grooves

are identified by the distance around the helical width from the C1 on one strand to the C1 on

the second strand (figure 1.4).

H H N H O N O H N N

N H N N N H N N N N dR N N dR O dR O H N dR H C•G T•A

Figure 1.4. Depiction of the major and minor grooves found in C•G and T•A base pairs.

Double helical structures of DNA exist in many different polymorphs, where the three major

ones are A-, B- and Z-form (figure 1.5). The B-form is the most stable under physiological

conditions and can be interconverted by manipulating the surrounding environment. The

conversion between the A- and B-forms was established early on by Rosalind Franklin, who

showed that two forms of DNA, later identified as A and B, could be interconverted depending

upon the hydration level.26 The conversion between B- and Z-forms was shown to be salt

dependent, where a high salt concentration (2.5 M NaCl or 0.7 M MgCl2) stabilizes the Z-form of poly(dG-dC).40 The phosphate groups are pushed closer together resulting in the smallest

helical diameter among the three major polymorphs. Chemical modifications, such as

alkylations of guanosine41 or cytosine,42,43 can also facilitate the B- to Z-form conversion,

allowing the transition to occur at lower salt concentrations.

A-Form B-Form Z-Form

Figure 1.5. The three major polymorphs of DNA visualized by Chimera44 from Glactone (Chancellor of the University System of Georgia) generated pdb file.45

B-Form DNA The structure predicted by Watson and Crick in 1953 was that of B-form DNA, where the

two anti-parallel strands form a right-handed helix with a twist of 36° per base pair that rises

3.3 Å and is composed of 10 base pairs per full turn. The close packing of each base pair

results in a helical diameter of 20 Å, which is larger than Z-form but smaller than A-form DNA

(Table 1.1). B-form DNA, having the most identifiable grooves (figure 1.6) has a major groove

width (11.7 Å) that is approximately double its minor groove width (5.7 Å), while the depths of

both grooves are almost equal with the major groove being slightly deeper than the minor

groove (8.8 Å and 7.5 Å respectively).

Major Groove Minor Groove

Figure 1.6. B-Form DNA depicting both major and minor grooves visualized by Chimera44 from Glactone (Chancellor of the University System of Georgia) generated pdb file.45

A-Form DNA A-form DNA, originally seen in diffraction patterns26 in the early 1950s, is also a right handed helix, but has one extra base pair (11) per helical turn more than B-form DNA, which results in a shorter twist angle (32.7°) and rise per base pair (2.56 Å). The consequence of a shorter rise results in a 20° tilt in order to maintain minimum van der Waals interactions of 3.4

Å between adjacent base pairs.35 A-form DNA, as a result of the base stacking distance and

twist, has a minor groove that is extremely wide (11 Å) in comparison to its very narrow major

groove (2.7 Å).

Major Groove Minor Groove

Figure 1.7. A-Form DNA depicting both major and minor grooves visualized by Chimera44 from Glactone (Chancellor of the University System of Georgia) generated pdb file.45

Z-Form DNA Z-form DNA, the polymorph of the first single crystal X-ray structure,46 is a left handed helix where sequential bases alternate their orientation from anti to syn resulting in a narrow helical

diameter (18 Å) and an extended distance (3.7 Å) between adjacent base pairs. Unlike A- and

B-form DNA, where the major and minor grooves are easily identified, the grooves of Z-form

DNA are more difficult to identify (figure 1.8). Z-form DNA, denoted because of its backbone

that zigzags due to the syn–anti base flipping that occurs, has a shallow and wide major groove (8.8 and 3.7 Å, respectively) with a narrow and deep minor groove (2.0 and 13.8 Å respectively). Z-form DNA is typically seen with alternating sequences of purines and pyrimidines, typically dG-dC sequences, because there is very little energetic penalty associated with an anti to syn flip of a purine residue.47

Minor Major Groove Groove

Figure 1.8. Z-Form DNA depicting both major and minor grooves visualized by Chimera44 from Glactone (Chancellor of the University System of Georgia) generated pdb file.45

Table 1.1 Helical parameters for the three major DNA polymorphs.35 Helical Twist Rise Groove Groove DNA Residues Base Sugar Helix diameter per bp per bp width / Å depth / Å form per turn tilt / ° pucker / Å / ° / Å major minor major minor

A Right 11 26 32.7 2.56 20 C3-endo 2.7 11.0 13.5 2.8

B Right 10 20 36 3.3 6 C2-endo 11.7 5.7 8.8 7.5 C3-endo Z Left 12 18 9,51 3.7 7 C2-endo 8.8 2.0 3.7 13.8

1.5 – Biological roles of the 3 major DNA polymorphs

The varied structural features of the three polymorphs of DNA are related to their biological

function. B-form DNA is the most stable of all the DNA polymorphs at physiological conditions

and is involved in many biological functions that require DNA. However, B-form DNA when

composed of mainly oligopurines-oligopyrimidines can also form higher order structures, such

as DNA triplexes. These triplexes, known since 1957,48,49 result from two poly A-poly U

duplexes that transform to a triple stranded poly(U-A-U) structure and a single strand of poly

A. Specifically, H-DNA, a triplex found in vivo, is thought to play an important role in gene regulation, due in part to the abundance of oligopurine-oligopyrimidine units in proximity to gene regulatory regions.50-55 Z-form DNA has been shown to exist in the unwrapping of DNA from nucleosomes,56 where it prevents reformation of the nucleosome57 and thereby stimulates

transcription.58,59 Sequences that have the propensity to morph from B-form to Z-form have frequently been identified near transcription start sequences.60 In addition, proteins whose

function is directly related to viral pathogenesis, have also been shown to bind Z-form

DNA.61,62 Although A-form DNA does not exist free in solution under physiological conditions

for it is a higher energy polymorph, it has been seen in certain polymerases.63,64 For example, the human immunodeficiency virus type 1 (HIV-1) reverse transcriptase, which transcribes single-stranded RNA into the corresponding DNA duplex via formation of a DNA/RNA hybrid, is known to bind the resulting DNA duplex in such a fashion that the DNA duplex near the

active site is A-form while the DNA duplex near the RNase H site is B-form.65-69 The three major DNA polymorphs clearly have biological functions whose investigation is of interest to both the scientific and public communities.

1.6 – Using fluorescence spectroscopy to probe DNA

While the active role in biology of the various polymorphs clearly addresses the need for

their study, it however does not depict the approach. Classical methods still serve to provide

relevant and important information concerning the effects of these DNA polymorphs. They

tend to provide global information while fluorescence spectroscopy, a more rapid and less

invasive technique, can provide both global and local information about the structure and

dynamics of nucleic acids. Due to the occurrence of amino acids with favorable photophysical

properties, fluorescence spectroscopy has provided protein chemists useful information over

the past decades. However, it is the poor photophysical properties of the common nucleic

acids (common pyrimidines and purines are practically non-emissive70-78) that present a challenge to develop novel nucleosides that have enhanced photophysical properties.

1.7 – Applications of fluorescent nucleosides

Presented below are two examples depicting the utility of fluorescent nucleosides in

answering critical biological questions. Fluorescent nucleosides have been utilized to monitor

binding events, specifically the binding of small molecules to RNA constructs. While, it is

possible to monitor small molecule-RNA binding via gel shift assays or isothermal titration

calorimetry (ITC), the power of fluorescence spectroscopy is its ability to rapidly and

accurately access, with very small quantities, the binding of numerous small molecules in a

day, rather than weeks. This approach has been demonstrated independently by both T.

Hermann and D. Pilch, where they monitor the binding of aminoglycosides to the ribosomal A-

site using 2-aminopurine (2-AP), an environmentally sensitive fluorescent nucleoside.79,80 The ribosomal A-site, which is responsible for the fidelity of protein biosynthesis,81 utilizes two

conserved adenine residues to discriminate against noncognate codon-anticodon

interactions.82 Separate replacement of these two adenines with 2-AP results in a detectable

method to discern the binding of aminoglycosides that cause the fluorescent 2-APs to bulge

out, thereby inducing a fluorescence intensity change. The advantages of this system are the

need for extremely small amounts of very expensive RNA, the robustness to quickly screen

numerous compounds for binding, and the precision to determine the binding affinity of

molecules that previously resulted in positive hits.

This second example illustrates the unique ability of fluorescent nucleosides to answer

important biological questions that could not be answered by traditional techniques. Saito,

Majima, Ganesh and Tor have utilized novel fluorescent nucleosides to determine the polarity

of the grooves in a DNA helix (for a more in depth discussion of this field, see chapter 3).83-91

The polarity of a DNA groove is determined by correlating the photophysical properties,

specifically the Stokes shift, of the free nucleosides in known polarity solvents to the read-out

determined upon incorporation of the fluorescent nucleoside into a DNA duplex. Fluorescent

nucleosides can provide an understanding of these unique environments, namely the local

polarity of specific cavities in nucleic acids, in order to facilitate the design of novel low

molecular weight ligands. Therefore, fluorescent nucleosides are vital to the understanding of

nucleic acid structure and function.

1.8 – Major families of fluorescent nucleosides

The above two examples highlight the importance of fluorescent nucleosides as tools for

investigating key biochemical questions. Fluorescent nucleosides can be classified into

several major families, including: (a) isomorphic base analogues (b) pteridines (c) expanded

nucleobases, (d) chromophoric base analogues, and (e) extended base analogues (review articles on fluorescent nucleosides92-100).

Isomorphic base analogues Isomorphic base analogues are heterocycles that closely resemble the corresponding natural nucleobases with respect to their overall dimensions, hydrogen bonding face, and ability to form isostructural W-C base pairs (selected examples are depicted in figure 1.9). An advantage of isomorphic fluorescent nucleosides is their similarity to the native nucleobases, normally resulting in minimal structural perturbation from the native system. Since favorable photophysical characteristics (e.g., red shifted absorption and high emission quantum efficiencies) are typically associated with significant structural perturbation, isomorphic fluorescent nucleosides are the most challenging to design.

NH2 NH2 N N H N N N N N N N N NH N O 2 N R N NH2 N R R R 2-aminopurine (2-AP) 2,6-diaminopurine formycin 5-methyl-2-pyrimidinone λ abs (nm) 303 280 295 280 λ em (nm) 370 350 340 400 θ F 0.68 0.01 0.06 --- τ 7.0 --- <1 --- (ns) Figure 1.9. Selected examples of isomorphic base analogues where R = 2-deoxyribose or ribose. Data obtained in water.94,97

One of the first and most widely utilized fluorescent nucleosides, 2-AP, is a constitutional isomer of adenine with enhanced photophysical properties over the native nucleoside. Since the initial publication in 1969, describing the fluorescence properties of 2-AP both in

nucleoside and oligonucleotide form,101 2-AP has generated over 1,600 journal references

(SciFinder). This monumental paper by Reich and Stryer describes the ideal properties of 2-

AP, namely its high quantum efficiency (0.68 in water), isolated absorption band (303 nm), minimal fluorescence change due to pH changes, ability to form W-C like hydrogen bonds with dT (figure 1.10), acceptable substrate for RNA polymerase and most importantly, its sensitivity to the surrounding environment.

R2 O H N N R2 O N

N H N N N H N N N N R1 N N R1 O O H N R1 R1 H T/U A T/U 2-AP

Figure 1.10. Hydrogen bonding of T/U•A and T/U•2-AP where R1 = 2deoxyribose or ribose and R2 = Me (T) or H (U).

The remarkable sensitivity of 2-AP can be seen in figure 1.11, where hyperchromic and bathochromic shifts are seen upon an increase in polarity of the surrounding environment.

a) b) 2.9 0.8 ) -1 cm 4 2.8 F φ

0.4

2.7 Energy (x 10

0 40 50 60 40 50 60

ET(30) Figure 1.11. Relationship between (a) emission energy, (b) quantum yield and microenvironment polarity. For both (a) and (b) data was plotted from Nordlund102 (red squares), Wierzchowski103 (blue circles) and Stryer101 (green diamonds).

While 2-AP pairs with adenine in a way that does not disturb the secondary structure of either

A- or B-form DNA/RNA, it can also pair with cytosine in various forms depending upon the pH

of the surrounding environment (figure 1.12), being the origin for its mutagenicity104-106.

H N H N

N H N N N N R1 H N O H N R1 N H H + N N C 2-AP + N R H N H N 1 N H H O R1 N H N C 2-AP N H N N N N R1 O H N R1 H C+ 2-AP neutral wobble pairing Ionized WC pairing

Figure 1.12. Mispairing of 2-AP with cytosine and the effect of pH upon its pairing.

Due to the extensive use of 2-AP, the following section will describe select examples of 2-

APs utility as a fluorescent probe. The environmental sensitivity of 2-AP allows it to detect minor structural changes in nucleic acids, such as the binding of small molecules to the A-site

(Section 1.7). In a different RNA system, Tor and coworkers have utilized 2-AP to monitor the activity of a catalytic RNA, namely the hammerhead rybozyme.107 The hammerhead ribozyme, a RNA enzyme that catalyzes the selective hydrolysis of one of its phosphodiester bonds, has been well studied for over 20 years, resulting in a large amount of knowledge concerning its structure, mechanism and kinetics. Y. Tor and coworkers utilized 2-AP to monitor the influence of small molecules on the activity of the hammerhead ribozyme. It was possible, but laborious, to monitor the influence of small molecules on the cleavage rate via

radiolabeling. However, the introduction of 2-AP near the cleavage site allowed for rapid

monitoring of the ribozyme as a result of the different environments experienced before and

after cleavage by 2-AP. As seen in the A-site assay (Section 1.7), 2-AP in an intrahelical

position, the native hammerhead ribozyme form, is quenched and upon exposure to a more

polar environment, the cleavage of the phosphodiester bond results in an increased emission

intensity.

The use of 2-AP has not been limited to that of small molecule-RNA interactions. 2-AP,

due to its dramatic photophysical changes as a result of a minor structural change (A to 2-AP)

has led to numerous ab initio calculations concerning the reason behind its enhanced

photophysical properties over adenine108-110 and other phenomenon, specifically quenching effects.111-118 2-AP has been used to investigate the structure and dynamics of abasic sites,119-

121 the strength of base pairing interactions,122 the rate of helicase activity,123 and it has been used as a mutagen in vivo and then as a probe of DNA polymerase fidelity in vitro.124

While 2-AP has been the isomorphic base of choice for the biophysical community, 5- methyl-2-pyrimidione has also been investigated as a fluorescent nucleobase (figure 1.9).

Synthesis of 5-methyl-2-pyrimidinone (d5 or m5K) and its incorporation into oligonucleotides occurred in the late 1980s.125,126 Since then, there have been but a handful of papers, mostly

ab initio calculations,127,128 discussing its utility as a fluorescent probe.129 Initial investigation of d5 described its isolated absorption (280 nm) and lifetime studies that investigated the stacking ability of d5 in a single stranded oligonucleotide.129 A more recent account utilizes d5

to monitor the binding of ssDNA to Escherichia coli RecA protein.130 However, the recent interest in d5 as a non-natural base pair for enzymatic incorporation131-133 or as a base in a novel triple helix motif,134,135 and not as a fluorescent probe, possibly stems from its inability to form an adequate W-C base pair with adenine.

Pteridines

Pteridines are naturally occurring intensely fluorescent (F = 0.39–0.88) aromatic compounds whose structure is similar to that of the purines (selected examples are depicted in figure 1.13). Their intense fluorescence, located in the visible at 430 nm with a rather long lifetime ( = 3.8–6.5 ns), results from an isolated absorption band located above 300 nm. The development of the pteridines as fluorescent nucleoside analogues was initiated and extended almost exclusively by Mary E. Hawkins and coworkers. After initial screening of potential pteridine nucleosides, a few were selected whose photophysical and chemical properties were suitable for use as fluorescent probes.

O O NH2 NH2 N N N N N NH N N

O N N NH2 O N N NH2 O N N O N N R R R R 3-MI 6-MI DMAP 6-MAP λ abs (nm) 348 340 330 310 λ em (nm) 430 431 430 430 θ F 0.88 0.70 0.48 0.39

τ 6.5 6.4 4.8 3.8 (ns) Figure 1.13. Selected examples of pteridines, purine analogues, where R = 2-deoxyribose or ribose.93

All four pteridine analogues, namely the G analogues (3-MI and 6-MI) and the A analogues

(DMAP and 6-MAP) are able to be site specifically incorporated into oligonucleotides using standard automated DNA synthesis whereby they retain their overall absorption and emission bands. Initial studies revealed a large quenching effect upon incorporation into an oligonucleotide (ranging from 64 to 98%) with a major dependence upon the flanking bases, namely a greater quenching effect when spaced between purines than pyrimidines. There is

no general trend seen when pteridines are flanked by both a pyrimidine and a purine, only that

the quantum efficiency is lower than the two pyrimidine flanked case, thus generating a

sequence specific quenching effect (see ref136 for a more detailed discussion). Thermal denaturation experiments revealed that the incorporation of these modified nucleosides, except for 6-MI, resulted in a destabilizing effect similar to that of a single base pair mismatch.137 Although a thermal destabilization was seen with 6-MAP and DMAP, their overall stability was selective for thymidine, suggesting the formation of hydrogen bonds when in a helical structure.

The minimal destabilization and large quenching effect seen upon incorporation were utilized in the development of a novel assay for monitoring intergrase activity. Intergrase is a retrovirally-encoded protein in HIV-1 that is responsible for integrating the newly reverse transcribed double stranded DNA into the host cells genome. The first step of this process results in the cleavage of a dinucleotide fragment from both of the 3-ends. The incorporation of 3-MI as the second base in from the 3-end of the double stranded DNA resulted in a quenching effect in comparison to the isolated nucleoside. Upon cleavage of the dinucleotide containing the pteridine, part of the fluorescence intensity was recovered because the probe was flanked on only one side.138 This allowed for the monitoring of the intergrase protein in real-time as opposed to previous methods that typically employed radiolabeling the substrate and visualization via gel electrophoresis139 or high throughput plates.140 This technique has also been applied to other enzymes, specifically P1 nuclease and exonuclease III.100

Again, relying upon the quenching properties of the pteridine analogues, Hawkins and

coworkers developed a fluorescent probe that recognizes PCR products without the need to

separate annealed or labeled products.100 A bulge hybridization mechanism was utilized, where a sequence complementary to the product, except for the fluorescent probe (3-MI), is designed. Then upon hybridization to the product strand, the fluorescent probe will bulge out, thus unstacking from its neighboring bases, resulting in an increase in fluorescence intensity

(typically 1.5 to 27 fold). This technique allowed the probe strand to be added to the PCR

mixture prior to amplification, where a weak background fluorescence signal is seen due to the

presence of non-complementary sequences. Then continuation of the PCR cycle and production of the product strand sees an increase in fluorescence intensity due to the hybridization of the probe strand to the target PCR product.

Pteridines, such as 3-MI, have also been utilized to monitor the binding of HU, a multifunctional histone-like protein found only in prokaryotes, to duplexed DNA.141 As

fluorescent nucleosides are typically smaller in size than large fluorescent tags, 3-MI was also

employed to monitor the cellular uptake of oligonucleotides.100 Most recently, the adenosine analogue 6-MAP was used to investigate the structure of long sequences of consecutive adenosine residues, known as A tracts.142 The pteridine class of heterocycles is a well-

established class of novel fluorescent nucleosides that will continue to be used as fluorescent

probes do to their high quantum efficiency, minimal structural perturbations, commercial

availability and well-documented quenching effects.

Expanded nucleobases The expanded nucleobases are characterized by their extended conjugation of the natural

nucleobases through the fusing of additional aromatic rings (selected examples are depicted

in figure 1.14). While the addition of rather large aromatic rings to the natural nucleobases

normally induces a structural perturbation to the system of study, it will also result in enhanced

photophysical properties. As can be seen from figure 1.14, the modified nucleobases retain

their W-C hydrogen bonding face (the exception being etheno-A), thus allowing them, despite

their large aromatic rings, to interact in normal W-C base pairing. All the expanded

nucleobases pictured below have red shifted absorption bands in comparison to their natural

nucleobase derivatives, which results in an emission band near the visible range and rather

high quantum efficiencies, ranging from 0.2–0.82.

O NH NH NH NH2 S N N N N N N NH N O N O N N N N N O R R R R R

aaaa b Pyrrolo-dC Etheno-A (ε A) Benzo-A BgQ tC λ abs (nm) 350 294 356 360 375 λ em 460 415 358, 379, 395 434 500 (nm) θ F 0.2 0.6 0.44 0.82 0.2

τ --- 20 ------3.7 (ns) Figure 1.14. Selected examples of expanded nucleobase analogues where R = 2- deoxyribose or ribose. Photophysical properties measured in buffer at: a pH = 7.0 and b 7.5.94,97,99

Nelson Leonard and coworkers first investigated etheno-A (A)143,144 and benzo-A145 in the early 1970s following an initial report that showed adenine and cytosine could be cyclized to produce nucleobases146 with red shifted absorptions bands. While the fused structure of A,

reminiscent of the naturally occurring fluorescent nucleoside wyosine,147 masked the hydrogen bonding face, it also enhanced its photophysical properties, most notably with a red shifted

absorption (294 nm), an intense (F = 0.56) emission band in the visible range (415 nm) that was associated with a rather large stokes shift (121 nm) and a very long lifetime ( = 20 ns) for a small organic chromophore. A, identified as a fluorescent replacement for ATP, showed activity in AMP/ATP binding enzymes similar to that of the natural substrates, which shed light on the minimal importance of the exocyclic amine in enzymatic binding events. Leonard extended his initial investigation into exploring the cytosine version, namely C, and other

compounds that contained ATP, specifically NAD+, where the ATP portion was replaced by

A (figure 1.15).144

O O H H2N N N H3C N N N O O N P O CH3 O O HO OH Wyosine (Yt)

O P O H N N (HCl) O N N N O N N N HO OH R O ε + ε NAD etheno- C ( C)

Figure 1.15. Structures of a fluorescent analogue of NAD+ (NAD+), naturally occurring fluorescent nucleoside wyosine and the HCl salt of ethano-C where R = ribose.

Benzo-A retains the hydrogen bonding face of adenine with an extended body, whereby

the photophysical properties are improved over adenosine. A structural and photophysical

comparison of A and benzo-A reveals the intricacies involved in novel fluorescent

nucleosides. The addition of a benzene ring in benzo-A results in a tremendous red shift in

the absorption spectra with a rather small stokes shift (~40 nm) and a lower quantum

efficiency in comparison to A. Similar to the impact that Reich and Stryer had with their

publication of 2-AP, Leonards two Science publications described two novel fluorescent nucleosides that are still being studied today, most notably benzo-A by E. T. Kool148,149 and A

by A. Holmén.150 In developing a novel expanded genetic alphabet, Kool has published a

large number of expanded fluorescent nucleosides whose basic photophysical properties have

been explored, but not utilized in any application.151-155

Pyrrolo-dC, a novel fluorescent nucleoside that has gained much attention in recent years,

was originally discovered as a side product in the Sonogashira coupling of terminal alkynes to

5-iodonucleosides.156-158 While initial investigations were focused on the enzymatic

activity,159,160 it was not long before its fluorescence properties were explored.161 The

excitement concerning pyrrolo-dC results its photophysical properties, W-C type hydrogen

bonding face and commercial availability. The absorption band of pyrrolo-dC (350 nm) is

considerably isolated from that of the native nucleobases, which results in a sufficiently

intense ( F = 0.2) long wavelength emission band (460 nm). These photophysical properties along with a minimally perturbing structure have led to a variety of applications. A rigorous study confirmed the utility of pyrrolo-dC as a fluorescent probe by concluding that under various ionic strengths, pHs, addition of cosolvents and temperatures, minimal effect to the fluorescence spectra was observed. In addition, a typical quenching effect was seen where a

60% reduction in quantum efficiency upon incorporation into an oligonucleotide and additional quenching (75% total) upon hybridization to form a duplex paved the way for kinetic measurements of association and disassociation of an RNA/DNA hybrid.162 Due to its

minimally perturbing structure, pyrrolo-dC was used to investigate the structure of the poly-

purine tract in HIV-1. Both in the presence and absence of reverse transcriptase, pyrrolo-dC

responded with markedly different fluoresce intensities depending upon its position in the

DNA/RNA hybrid, thereby suggesting the degree to which the replaced base is intrahelical.163

In a similar fashion, pyrrolo-dC was utilized to investigate the structure and dynamics of the T7

RNA polymerase from initiation to elongation.164 N. Turro and coworkers explored the utility of

molecular beacons containing both 2-AP and pyrrolo-dC. Both fluorophores were strategically

placed where upon stem loop formation, in the absence of target strand, 2-AP would be

unpaired in the loop portion, thus generating an intense emission, while the pyrrolo-dC would

be paired in the stem portion, generating a weak emission. Then upon duplex formation, in

the presence of the target strand, the 2-AP would be base paired, resulting in a lowering of its

fluorescence intensity while the pyrrolo-dC would be unpaired outside the duplex, resulting in

a markedly higher fluorescence intensity. Due to the general overlap of the absorption bands

(ideal excitation at 325 nm) and the isolated emission bands of both chromophores (2-AP =

370 nm and pyrrolo-dC = 450 nm), a dual readout resulted whereby target detection can be

measured in a ratiometric way.165 Much like 2-AP, where its isolated absorption band has lead to the study of DNA structure and dynamics via low energy CD spectra,166 pyrrolo-dC with a

more red shifted absorption band has also been utilized in similar studies.167 Pyrrolo-dC has

also seen interest from the computational community.168

The last two nucleosides seen in figure 1.14, BgQ and tC, are recent additions to this class of fluorescent nucleosides. BgQ and a structural isomer of BgQ, due to their large surface area, were studied in duplex and triplex forming systems.169 Later the utility of BgQ was expanded to determine, much like 2-AP, the binding of a protein (tat) to its RNA construct

(TAR) whose function is important in the life cycle of HIV-1.170 The synthesis of a tricyclic (tC) nucleoside for antisense research was initially reported by M. Matteucci171 (Giled Sciences

Inc.), and later developed into a PNA strand172 by P. Nielsen. A collaboration between P.

Nielsen and L. Wilhelmsson finally saw the investigation of its photophysical properties.173 tC

is a cytidine analogue that forms W-C like base pairs with guanosine174 and surprisingly does not suffer from a dramatic reduction in quantum efficiency upon in corporation into PNA173 or

DNA,175 unlike most other fluorescent nucleosides. The expanded nucleobases class of

fluorescent nucleosides has been there from the beginning, with Leonards A, and continues

to provide novel structures that result in excellent photophysical properties for use as probes

in nucleic acids.

It should be noted that some fluorescent bases in the expanded base class were not

discussed here, most notably the work of I. Saito in single nucleotide polymorphism detection,

which is discussed in the extended base section.

Chromophoric base analogues Chromophoric base analogues comprise a class of fluorescent nucleoside analogues

where the natural heterocycle is replaced with a fluorescent aromatic residue that typically

lacks the W-C type hydrogen bonding face (selected examples are depicted in figure 1.16).

This replacement by a known chromophore results in positive photophysical properties and

normally larger overall structures than the native nucleobases.

O O N

R R R R

Pyrenea Perylenea Benzopyrenea Coumarinb λ abs (nm) 345 440 394 400 λ em (nm) 375, 395 443, 472 408 515 θ F 0.12 0.88 0.98 --- τ ------7.4 (ns) Figure 1.16. Selected examples of chromophoric base analogues where R = 2-deoxyribose a b or ribose. Data obtained in methanol and aqueous buffer pH=7.2 where F was determined in oligonucleotide form.94,97

E.T. Kool and R. S. Coleman have dominated this class of fluorescent nucleobases with

their work on C-linked nucleosides, such as pyrene, perylene, benzopyrene, and coumarin

moieties. All of these fluorescent nucleobases have isolated absorption bands ( 345 nm) that

allow for selective excitation in the presence of natural nucleobases. Although the

photophysical properties of the bases is known and those with high quantum efficiencies and

emission bands in the visible are typically chosen, most (benzopyrene retains its quantum

efficiency in protic solvents)97 still suffer from quenching effects upon incorporation into oligonucleotides.

Although Kool has utilized chromophoric base analogues mainly for the investigation of enzyme-substrate recognition, proving that size and shape is a major factor in polymerase- substrate recognition,176 and for the discovery of novel sensors through the development of

chromophoric arrays utilizing DNA as a scaffold,177,178 his introduction (synthesis and basic

photophysical properties) of these novel nucleosides179,180 has lead to applications by other researchers. Christensen and Pedersen have utilized an intercalating pseudo nucleotide

(IPN) that upon incorporation into a DNA strand forming an intercalating nucleic acids strand

(IDN), was able to distinguish between ssDNA and ssRNA by both thermal denaturation and fluorescence intensity measurements.181 This detection method is based upon the degree of

intercalation the pyrene moiety undergoes, where a greater intercalation results in a higher Tm

and a weaker emission intensity (seen for ssDNA) and a lower degree of intercalation results

in a lower Tm and a higher emission intensity (seen for ssRNA). Aubert and Asseline have utilized perylene connected either directly to the 2-deoxy sugar (C-linked nucleoside) or through a propyl linker, where upon incorporation into an oligonucleotide is shown to stabilize duplex and triplet formation. Introduction of the propyl linked perylene unit at both the 5 and

3 ends saw the greatest stabilization of +13°C for duplex and +22°C for triplex.182 An increase in fluorescence intensity is seen upon both duplex and triplet formation, thus allowing for hybridization monitoring by fluorescence spectroscopy.

Coleman and coworkers in 1998 described the synthesis of a coumarin containing nucleoside, whereby the base was replaced by coumarin 102.183 The coumarin nucleoside

analogue, having similar photophysical properties to its parent compound (Coumarin 102: abs

184 = 388nm, em = 470, F = 0.65 and = 4.7 ns), has an absorption band at 400 nm that results in a short lived ( = 7 ns) emission band near 515 nm due to a charge transfer transition in the excited state.185,186 The coumarin nucleoside analogue was designed to be

paired with an abasic site mimic (tetrahydrofuran) where it would be intrahelical and minimally

perturb a DNA duplex (figure 1.17).

H N O H N O N H3OC O O N N H N N O O N O O O Figure 1.17. Proposed position of coumarin nucleoside when paired with an abasic site mimic. A transparent W-C A•T base pair is provided for spatial comparisons.

A collaboration between R. S. Coleman, C. J. Murphy and M. A. Berg utilized the coumarin

nucleoside to probe the relaxation dynamics of a DNA duplex, whereby time-resolved Stokes-

shift (TRSS) measurements indicated: (1) that the interior of the DNA helix was neither fluid

nor crystal like,187 (2) that sequence dependent relaxation occurs faster than 40 ps,188 (3) that counter ions can be bound to a DNA duplex and measured using TRSS189,190 and (4) that

TRSS can monitor dynamic changes that occur due to structural changes or probe

positioning.191 The addition of N. P. Ernsting to this collaboration saw two recent publications

in which TRSS measurements investigated how DNA dynamics varied based upon the

position of the coumarin nucleoside (middle or end) within the DNA duplex.192,193 The

coumarin nucleoside is not limited to studies of standard DNA duplexes, but has also been

used to study the dynamics of DNA bound to certain endonucleases (APE1).194 Coleman has also described the synthesis of two anthracene containing nucleosides connected either via the 1 or 2 position, although complete photophysical properties or application of these nucleosides have yet to be described.195

Extended base analogues The final class of fluorescent nucleoside analogues is distinguished by fluorescent aromatic

moieties that are linked to the natural nucleobases (selected examples are depicted in figure

1.18). These probes have the advantage of utilizing known chromophores whose

photophysical properties are well studied and connecting them to the native nucleobases

whose hydrogen bonding ability is also well studied, resulting in a florescent nucleoside whose

fluorescent moiety is mostly, but not always, in conjugation with the natural nucleobase (figure

1.18).

N H NH2 N R2 N N X R N N 2 Y R1 a bb 8-aza-7-deaza-7-phenylethynyl X=N 1,10-Phenanthroline pyU/C Y=H py (7-deaza-phenylethynyl X=CH) (DMAP- U/C Y=PhN(CH3)2)

5 5 5 --- R2 = dU R2 = dU R2 = dC λ abs (nm) 294 (296) 333 341 (303) 329 (---) λ em (nm) 360 (412) 408 397 (440 LE) 393(---) (540 ICT) θ F 0.08 (0.02) 0.16 0.2 (0.01) 0.15 (---)

τ 0.55 (0.69) ------(---) --- (---) (ns) Figure 1.18. Selected examples of conjugated base analogues where R1 = 2-deoxyribose. Data collected in a double distilled water and b aqueous phosphate buffer (pH=7.0).97,196-198

The 5 position of U/C, being synthetically accessible minimally destabilizing when using rigid linkers199 has generated a large number of fluorescent nucleosides whose applications are wide ranging. Initial investigations explored a direct, carbonyl and amide linked pyrene to the 5-position of 2-deoxyuridine.200,201 These detailed studies, reporting absorption spectra, emission spectra, lifetimes and quenching effects seen upon attaching pyrene to the uridine core provided evidence that such nucleosides can be responsive to changes in their environment. Y. Tor and coworkers, having shown that 1,10-phenanthroline ethynyl

compounds respond to polarity changes202 and previously investigated polypyridine Ru and Os containing nucleosides,203-205 decided to investigate the metal free nucleoside, namely 5-

[(1,10-phenanthrolin-3-yl)ethynyl]-2-deoxyuridine (dUphen). This novel nucleoside displays an absorption band at 333 nm, well isolated from the native nucleobases that was insensitive to the polarity of its environment. Excitation of dUphen resulted in an emission band ranging from

385 nm (dichloromethane) to 408 nm (water) depending upon the polarity of its environment.

This polarity sensitivity directly translated to the emission seen when dUphen was incorporated into an oligonucleotide, where a multi-component emission spectra, deconvoluted through ratiometric treatment of the emission maximum, was able to discriminate between the four natural bases it was paired with in the DNA duplex (base discriminating fluorosides –

BDFs).196

In 2003, I. Saito started publishing novel BDFs that could be used for single nucleotide

polymorphism (SNPs) analysis. The approach that a single emission band intensity of a

fluorescent nucleobase analogue can detect SNPs is only one of many.206 These fluorescent nucleosides can be spilt into two categories: nucleobases with pendent fluorophores and expanded ring system nucleobases, both of which will be discussed in this section for simplicity. The first of these fluorescent nucleoside analogues utilized pyrene, previously shown to be sensitive to its environment, and connected to 2-deoxyuridine or cytosine via a ridged alkyne amide linker (see PyU/C in figure 1.18). This retained the hydrogen bonding face of U/C and the photophysical properties of pyrene, namely an isolated absorption band at

~335 nm and a relatively intense (F = ~0.2) emission band at ~400 nm. Despite the large

distance and unconjugated nature between the pyrene and the uracil/cytidine core, these

BDFs have been shown to clearly identify their complementary base via a markedly higher

emission in comparison to mismatch pairs or single stranded cases.96,198 The reason for such

drastic emission properties is likely structural in nature. Upon W-C hydrogen bond formation

(perfect complement), the rigid ethynyl linkage forces the pyrene into a polar environment

resulting in an intense emission band, while without a base pairing partner (ss or mismatch),

the nucleoside flips syn thus stacking the hydrophobic pyrene into the minor groove (a hydrophobic environment) and producing a quenched emission (figure 1.19). Molecular modeling also supports these structural predictions.198

O PyU•dA PyU•dG HN H O H N N H O O N N N N H N O H N N N N dR N NNH dR O O dR dR H N H

Figure 1.19. The suggested structure showing the extrahelical pyrene in the PyU•dA base pair and the intrahelical pyrene in the PyU•dG case.

The reporting ability of PyU/C has been shown to be most sensitive when flanked by

cytidines, where the flanking C/Gs better stabilize the syn conformation resulting in a greater

quenching effect, thereby accentuating the emissive (perfect complement) and quenched

states (ss or mismatch). However, in the perfect complement case, there is no additional

quenching seen when the probe is flanked by C/Gs over A/Ts as has been seen for some

expanded BDFs.207,208 The amide moiety is vital to the sensitivity of the PyU/C probes,

whereby removal of the carbonyl is completely detrimental.198 This is probably due to quenching by oxidative electron transfer from G209,210 or reductive electron transfer from C or

T.209,211 Amine removal results in a weaker effect, where a red shifted emission spectra (462 nm) showed varied intensities for each base pair.212 Interestingly, B. H. Kim and coworkers

have shown that the complete removal of the amide moiety, thus conjugating the pyrene to the

uridine core through an ethynyl linker also proves to be a good BDF. The fluorescent probe

selectively detects the perfect complement resulting in a higher emission intensity than the

mismatch or ss cases.213 Saito has expanded this approach to include detection of T•Hg•T

base pairs,214 a DMAP-PyU probe with dual emission properties,215 and an A analogue where a flexible linker was used generating a quenched spectra only in the presence of the perfect complement.216

The second type of BDFs designed by I. Saito are of the expanded base class (figure

1.20). These fused analogues tend to have similar photophysical properties to their pyrene

cousins, namely an absorption band near 350 nm and a nominally intense (F = ~0.1) emission band near 400 nm.

O O O Y NH NH X N N N N N O N O R R R MD A Y=NH2 and X=N

BPP NPP (MDI Y=O and X=NH ( λ abs (nm) 347 365 327 (315) λ em 390 395 397, 427 (442) (nm) θ F 0.04 0.26 0.12 (0.12)

τ ------(ns) Figure 1.20. Photophysical properties of size expanded BDFs. Spectra taken in aqueous buffer (pH = 7.0).97,208

The benzopyridopyrimidine (BPP) is a pseudo cytidine analogue that when incorporated into an oligonucleotide forms stable bases pairs with both dA and dG as seen by thermal

denaturation experiments. The fluorescence intensity of BPP is quenched in the presence of

dG, while it remains emissive when paired with dA. 15N NMR determined that BPP pairs with dG in a pseudo W-C type fashion, while it forms a wobble base pair with dA (figure 1.21).207

H O O H N N

N NH O N N H N N dR N H N N N N N dR N O H O dR N dR H BPP•dG BPP•dA W-C base pair wobble base pair

Figure 1.21. The base-pairing pattern of BPP of dG and dA as determined by 15N NMR.

The low quantum efficiency (0.04) and interference with ssBPP emission intensity lead to

the design of the naphthopyridopyrimidine (NPP) fluorescent nucleoside analogue. NPP

retains similar absorption and emission wavelengths, while yielding a substantially higher

quantum efficiency (F = 0.2). The surprisingly very weak ss emission intensity of NPP does not interfere with the purine discrimination (A vs. G).217 With an acceptable purine

discriminating fluorescent nucleoside in hand, Saito went on to design novel pyrimidine

discriminating fluorescent nucleosides, namely adenosine and inosine analogues that were

used to distinguish between dT and dC in a similar manner as BPP/NPP (figure 1.20).208

F. Seela and coworkers have diligently investigated the photophysical effects of extending the conjugation of 7-deaza-adenosine and 8-aza-7-deaza-adenosine by attaching numerous functionalized alkynes and alkenes to the 7-position (figure 1.18). Although the absorption spectra of the parent compounds, 7-deaza-adenosine and 8-aza-7-deaza-adenosine, are slightly red shifted in comparison to adenosine (270 nm vs. 261 nm respectively), they are not

emissive, unlike the conjugated, extended A analogues (alkyne and alkene). The

phenylethynyl analogues, depicted in figure 1.18, show some of the most red shifted

absorption and emission bands, albeit with relatively weak quantum yields (some analogues

197 have higher quantum efficiencies, F 0.3). Seela and coworkers have extended their system to include fluorescent guanosine analogues that are site-specifically incorporated into

DNA using standard solid phase synthesis, while their triphosphate forms are accepted by

DNA polymerases.218 These modified bases show minimal perturbation to DNA duplexes (7- postion is projected toward the major groove).219

In recent years, several groups have investigated the use of rather large fluorophores

(FAM, TAMRA, etc.) connected to either the 5-position of the pyrimidines or the 7-position of the purines. Both positions are known to be minimally perturbing to DNA secondary structure.

T. Brown and coworkers have studied the use of these fluorescent bases in various PCR methods220-222 and recently utilized naphthalenyl- and anthracenyl-ethynyl extended dT analogues as BDFs.223 F. Seela and coworkers have investigated the enzymatic acceptance of such fluorescent nucleosides and their resulting stability.218,224,225 Most interestingly, I. Hirao

and coworkers have demonstrated site-specific fluorescent labeling of RNA via an unnatural

base pair, in which both components are fluorescent nucleobases analogues.226,227 The extended fluorescent nucleosides are an ever-growing class of probes that have seen an abundance of interest from varied fields in recent years.

1.9 – Aims of the thesis

The interface of chemistry and biology requires not only the expertise of both, but also the

understanding of their complex relationship. The complexity of this field has lead to the

generation of numerous fluorescent nucleoside probes to investigate and answer questions in

an efficient and real-time manner that only fluorescence spectroscopy can provide. As is

evident from the diverse structures and un-related diverse photophysical properties of the

presented fluorescent nucleosides, there still lacks an understanding between the structure

and photophysical properties of modified nucleosides. A detailed investigation of all

fluorescent nucleosides clearly reveals the lack of isomorphic pyrimidine, specifically dT,

analogues with desirable photophysical properties. This thesis will demonstrate that minimal

structural modifications of the pyrimidine core can result in isomorphic dT nucleosides whose

desirable photophysical properties make them useful as probes, namely for the non-

destructive detection of abasic sites and the estimation of the polarity in DNA grooves.

1.10 References

1. Miescher, F. "Ueber die chemische Zusammensetzung der Eiterzellen." Die Histochemischen und physiologischen Arbeiten. 1897, 2, 323.

2. Jacobs, W. A.; Levene, P. A. "Further Studies on the Constitution of Inosinic Acid" Proc. Soc. Exp. Biol. Med. 1909, 6.

3. Jacobs, W. A.; Levene, P. A. "Isonic Acid" Proc. Am. Soc. Biol. Chem. 1910, 25.

4. Jacobs, W. A.; Levene, P. A. "Isonic Acid" J. Biol. Chem. 1912, 9.

5. Levene, P. A.; Jacobs, W. A. "Inosinic Acid" Ber. Deutschen Chem. Ges. 1908, 41, 27032707.

6. Levene, P. A.; Jacobs, W. A. "Inosinic Acid III" Ber. Deutschen Chem. Ges. 1909, 42, 11981203.

7. Levene, P. A.; Jacobs, W. A. "Inosinic Acid II" Ber. Deutschen Chem. Ges. 1909, 42, 335338.

8. Levene, P. A.; Jacobs, W. A. "Inosinic Acid IV" Ber. Deutschen Chem. Ges. 1911, 44, 746753.

9. Levene, P. A.; Jacobs, W. A. "Structure of Thymus Nucleic Acid" J. Biol. Chem. 1912, 12, 411420.

10. Levene, P. A. "The structure of yeast nucleic acid. IV. Ammonia hydrolysis" J. Biol. Chem. 1919, 40, 415424.

11. Klein, W.; Thannhauser, S. J. "Experimental studies on nuclein metabolism. XXXII. Ribodesose-guanylic acid" Hoppe-Seyler's Z. Physiol. Chem. 1933, 218, 173180.

12. Klein, W.; Thannhauser, S. J. "Experimental studies on nuclein metabolism. XXXIV. Ribodesoseadenylic acid from thymonucleic acid" Hoppe-Seyler's Z. Physiol. Chem. 1934, 224, 252260.

13. Klein, W.; Thannhauser, S. J. "Experimental studies on nuclein metabolism. XXXV. The pyrimidine nucleotides from thymus nucleic acid" Hoppe-Seyler's Z. Physiol. Chem. 1935, 231, 96103.

14. Brown, D. M.; Todd, A. R.; Varadarajan, S. "Nucleotides. XL. O2,5'-Cyclouridine and a synthesis of isocytidine" J. Chem. Soc. 1957, 868872.

15. Michelson, A. M.; Todd, A. R. "Nucleotides. XXXVIII. An improved synthesis of uridine diphosphate-glucose (UDPG)" J. Chem. Soc. 1956, 34593463.

16. Michelson, A. M.; Szabo, L.; Todd, A. R. "Nucleotides. XXXVI. Adenosine-2' uridine-5' phosphate" J. Chem. Soc. 1956, 15461549.

17. Chase, B. H.; Kenner, G. W.; Todd, A. R.; Webb, R. F. "Nucleotides. XXXV. Cyclopentanone oxime p-nitrobenzenesulfonate as an intermediate in the synthesis of nucleotide derivatives-an alternative synthesis of adenosine 5'-triphosphate" J. Chem. Soc. 1956, 13711376.

18. Michelson, A. M.; Todd, A. R. "Nucleotides. XXXII. Synthesis of a dithymidine dinucleotide containing a 3',5'-internucleotidic linkage" J. Chem. Soc. 1955, 26322638.

19. Hayes, D. H.; Michelson, A. M.; Todd, A. R. "Nucleotides. XXX. Mononucleotides derived from deoxyadenosine and deoxyguanosine" J. Chem. Soc. 1955, 808815.

20. Todd, A. R. "Chemical structure of the nucleic acids" Proc. Nat. Acad. Sci. U.S.A. 1954, 40, 748755.

21. Chargaff, E.; Magasanik, B.; Doniger, R.; Vischer, E. "The nucleotide composition of ribonucleic acids" J. Am. Chem. Soc. 1949, 71, 15131514.

22. Astbury, W. T. "X-ray studies of nucleic acids" Symp. Soc. Exp. Biol. 1947, 6676.

23. Watson, J. D.; Crick, F. H. "Genetical implications of the structure of deoxyribonucleic acid" Nature 1953, 171, 964967.

24. Watson, J. D.; Crick, F. H. "Molecular structure of nucleic acids; a structure for deoxyribose nucleic acid" Nature 1953, 171, 737738.

25. Wilkins, M. H.; Stokes, A. R.; Wilson, H. R. "Molecular structure of deoxypentose nucleic acids" Nature 1953, 171, 738740.

26. Franklin, R. E.; Gosling, R. G. "Molecular configuration in sodium thymonucleate" Nature 1953, 171, 740741.

27. Wilson, E. B.; Leaming, E. An atlas of the fertilization and karyokinesis of the ovum; Pub. for the Columbia university press by Macmillan and co.: New York, London, 1895.

28. Nirenberg, M. W.; Martin, R. G.; Jones, O. W.; Barondes, S. H.; Mathhaei, J. H. "Approximation of Genetic Code Via Cell-Free Protein Synthesis Directed by Template Rna" Fed. Proc. 1963, 22, 55.

29. Kaziro, Y.; Ochoa, S.; Grossman, A. "Identification of Peptides Synthesized by Cell- Free E. Coli System with Polynucleotide Messengers" Proc. Nat. Acad. Sci. U.S.A. 1963, 50, 5461.

30. Ochoa, S. "Synthetic Polynucleotides and Genetic Code" Fed. Proc. 1963, 22, 6274.

31. Sanger, F. "Sequences, Sequences, and Sequences" Annu. Rev. Biochem. 1988, 57, 128.

32. Khorana, H. G. "Polynucleotide Synthesis and Genetic Code" Fed. Proc. 1965, 24, 14731487.

33. Khorana, H. G. "Total Synthesis of a Gene" Science 1979, 203, 614625.

34. Saenger, W. Principles of nucleic acid structure; Springer-Verlag: New York, 1984.

35. Blackburn, G. M.; Gait, M. J. Nucleic acids in chemistry and biology; 2nd ed.; Oxford University Press: Oxford, England ; New York, 1996.

36. Neidle, S. Nucleic acid structure and recognition; Oxford University Press: Oxford ; New York, 2002.

37. Takahashi, I.; Marmur, J. "Replacement of Thymidylic Acid by Deoxyuridylic Acid in Deoxyribonucleic Acid of Transducing Phage for Bacillus Subtilis" Nature 1963, 197, 794795.

38. Judson, H. F. The eighth day of creation : makers of the revolution in biology; Simon and Schuster: New York, 1979.

39. Watson, J. D. The double helix; a personal account of the discovery of the structure of DNA; 1st ed.; Atheneum: New York,, 1968.

40. Pohl, F. M.; Jovin, T. M. "Salt-Induced Cooperative Conformational Change of a Synthetic DNA - Equilibrium and Kinetic Studies with Poly(Dg-Dc)" J. Mol. Biol. 1972, 67, 375396.

41. Möller, A.; Nordheim, A.; Nichols, S. R.; Rich, A. "7-Methylguanine in Poly(Dg- Dc).Poly(Dg-Dc) Facilitates Z-DNA Formation" Proc. Nat. Acad. Sci. U.S.A. 1981, 78, 47774781.

42. Behe, M.; Felsenfeld, G. "Effects of Methylation on a Synthetic Polynucleotide - the B- Z Transition in Poly(Dg-M5dc).Poly(Dg-M5dc)" Proc. Nat. Acad. Sci. U.S.A. 1981, 78, 16191623.

43. Behe, M.; Zimmerman, S.; Felsenfeld, G. "Changes in the Helical Repeat of Poly(Dg- M5dc).Poly(Dg-M5dc) and Poly(Dg-Dc).Poly(Dg-Dc) Associated with the B-Z Transition" Nature 1981, 293, 233235.

44. Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. "UCSF chimera - A visualization system for exploratory research and analysis" J. Comput. Chem. 2004, 25, 16051612.

45. http://chemistry.gsu.edu/glactone/index.shtml

46. Wang, A. H. J.; Quigley, G. J.; Kolpak, F. J.; Crawford, J. L.; Vanboom, J. H.; Vandermarel, G.; Rich, A. "Molecular-Structure of a Left-Handed Double Helical DNA Fragment at Atomic Resolution" Nature 1979, 282, 680686.

47. Rich, A.; Nordheim, A.; Wang, A. H. J. "The Chemistry and Biology of Left-Handed Z- DNA" Annu. Rev. Biochem. 1984, 53, 791846.

48. Felsenfeld, G.; Davies, A. G.; Rich, A. "Formation of the three-stranded polynucleotide molecule" J. Am. Chem. Soc. 1957, 79, 20232024.

49. Felsenfeld, G.; Rich, A. "Studies on the formation of two- and three-stranded polyribonucleotides" Biochim. Biophys. Acta 1957, 26, 457468.

50. Agazie, Y. M.; Burkholder, G. D.; Lee, J. S. "Triplex DNA in the nucleus: Direct binding of triplex-specific antibodies and their effect on transcription, replication and cell growth" Biochem. J. 1996, 316, 461466.

51. Kinniburgh, A. J. "A Cis-Acting Transcription Element of the C-Myc Gene Can Assume an H-DNA Conformation" Nucleic Acids Res. 1989, 17, 77717778.

52. Ohno, M.; Fukagawa, T.; Lee, J. S.; Ikemura, T. "Triplex-forming DNAs in the human interphase nucleus visualized in situ by polypurine/polypyrimidine DNA probes and antitriplex antibodies" Chromosoma 2002, 111, 201213.

53. Pestov, D. G.; Dayn, A.; Siyanova, E. Y.; George, D. L.; Mirkin, S. M. "H-DNA and Z- DNA in the Mouse C-Ki-Ras Promoter" Nucleic Acids Res. 1991, 19, 65276532.

54. Schroth, G. P.; Ho, P. S. "Occurrence of Potential Cruciform and H-DNA Forming Sequences in Genomic DNA" Nucleic Acids Res. 1995, 23, 19771983.

55. Wang, G. L.; Vasquez, K. M. "Naturally occurring H-DNA-forming sequences are mutagenic in mammalian cells" Proc. Nat. Acad. Sci. U.S.A. 2004, 101, 1344813453.

56. Liu, R.; Liu, H.; Chen, X.; Kirby, M.; Brown, P. O.; Zhao, K. J. "Regulation of CSF1 promoter by the SWI/SNF-like BAF complex" Cell 2001, 106, 309318.

57. Garner, M. M.; Felsenfeld, G. "Effect of Z-DNA on Nucleosome Placement" J. Mol. Biol. 1987, 196, 581590.

58. Oh, D.-B.; Kim, Y.-G.; Rich, A. "Z-DNA-binding proteins can act as potent effectors of gene expression in vivo" Proc. Nat. Acad. Sci. U.S.A. 2002, 99, 1666616671.

59. Rich, A.; Zhang, S. G. "Z-DNA: the long road to biological function" Nat. Rev. Genet. 2003, 4, 566572.

60. Champ, P. C.; Maurice, S.; Vargason, J. M.; Camp, T.; Ho, P. S. "Distributions of Z- DNA and nuclear factor I in human chromosome 22: a model for coupled transcriptional regulation" Nucleic Acids Res. 2004, 32, 65016510.

61. Brandt, T. A.; Jacobs, B. L. "Both carboxy- and amino-terminal domains of the vaccinia virus interferon resistance gene, E3L, are required for pathogenesis in a mouse model" J. Virol. 2001, 75, 850856.

62. Kim, Y. G.; Muralinath, M.; Brandt, T.; Pearcy, M.; Hauns, K.; Lowenhaupt, K.; Jacobs, B. L.; Rich, A. "A role for Z-DNA binding in vaccinia virus pathogenesis" Proc. Nat. Acad. Sci. U.S.A. 2003, 100, 69746979.

63. Pelletier, H.; Sawaya, M. R.; Kumar, A.; Wilson, S. H.; Kraut, J. "Structures of Ternary Complexes of Rat DNA-Polymerase-Beta, a DNA Template-Primer, and Ddctp" Science 1994, 264, 18911903.

64. Sawaya, M. R.; Pelletier, H.; Kumar, A.; Wilson, S. H.; Kraut, J. "Crystal-Structure of Rat DNA-Polymerase-Beta - Evidence for a Common Polymerase Mechanism" Science 1994, 264, 19301935.

65. Ding, J. P.; Hughes, S. H.; Arnold, E. "Protein-nucleic acid interactions and DNA conformation in a complex of human immunodeficiency virus type 1 reverse transcriptase with a double-stranded DNA template-primer" Biopolymers 1997, 44, 125138.

66. Jacobomolina, A.; Ding, J. P.; Nanni, R. G.; Clark, A. D.; Lu, X. D.; Tantillo, C.; Williams, R. L.; Kamer, G.; Ferris, A. L.; Clark, P.; Hizi, A.; Hughes, S. H.; Arnold, E. "Crystal-Structure of Human-Immunodeficiency-Virus Type-1 Reverse-Transcriptase Complexed with Double-Stranded DNA at 3.0 Angstrom Resolution Shows Bent DNA" Proc. Nat. Acad. Sci. U.S.A. 1993, 90, 63206324.

67. Saenger, W.; Hunter, W. N.; Kennard, O. "DNA Conformation Is Determined by Economics in the Hydration of Phosphate Groups" Nature 1986, 324, 385388.

68. Salazar, M.; Fedoroff, O. Y.; Zhu, L. M.; Reid, B. R. "The Solution Structure of the R(Gcg)D(Tataccc)-D(Gggtatacgc) Okazaki Fragment Contains 2 Distinct Duplex Morphologies Connected by a Junction" J. Mol. Biol. 1994, 241, 440455.

69. Sarafianos, S. G.; Das, K.; Tantillo, C.; Clark, A. D.; Ding, J.; Whitcomb, J. M.; Boyer, P. L.; Hughes, S. H.; Arnold, E. "Crystal structure of HIV-1 reverse transcriptase in complex with a polypurine tract RNA : DNA" EMBO J. 2001, 20, 14491461.

70. Albinsson, B.; Norden, B. "Electronic-Transition Moment Directions and Identification of Low-Energy N-Pi-Asterisk States in Weakly Perturbed Purine Chromophores" J. Am. Chem. Soc. 1993, 115, 223–231.

71. Andreasson, J.; Holmén, A.; Albinsson, B. "The photophysical properties of the adenine chromophore" J. Phys. Chem., B 1999, 103, 9782–9789.

72. Callis, P. R. "Electronic States and Luminescence of Nucleic-Acid Systems" Annu Rev Phys Chem 1983, 34, 329–357.

73. Daniels, M.; Hauswirth, W. "Fluorescence of the purine and pyrimidine bases of the nucleic acids in neutral aqueous solution at 300 degrees K" Science 1971, 171, 675– 677.

74. Morgan, J. P.; Daniels, M. "Excited-States of DNA and Its Components at Room- Temperature .4. Spectral, Polarization and Quantum Yield Studies of Emissions from Cpc and Poly-Rc" Photochem. Photobiol. 1980, 31, 207–213.

75. Nir, E.; Kleinermanns, K.; Grace, L.; de Vries, M. S. "On the photochemistry of purine nucleobases" J. Phys. Chem., A 2001, 105, 5106–5110.

76. Onidas, D.; Markovitsi, D.; Marguet, S.; Sharonov, A.; Gustavsson, T. "Fluorescence properties of DNA nucleosides and nucleotides: A refined steady-state and femtosecond investigation" J. Phys. Chem., B 2002, 106, 11367–11374.

77. Pecourt, J. M. L.; Peon, J.; Kohler, B. "Ultrafast internal conversion of electronically excited RNA and DNA nucleosides in water" J. Am. Chem. Soc. 2000, 122, 9348– 9349.

78. Wilson, R. W.; Callis, P. R. "Fluorescent Tautomers and the Apparent Photophysics of Adenine and Guanine" Photochem. Photobiol. 1980, 31, 323–327.

79. Kaul, M.; Barbieri, C. M.; Pilch, D. S. "Fluorescence-based approach for detecting and characterizing anti biotic-induced conformational changes in ribosomal RNA: Comparing aminoglycoside binding to prokaryotic and eukaryotic ribosomal RNA sequences" J. Am. Chem. Soc. 2004, 126, 34473453.

80. Shandrick, S.; Zhao, Q.; Han, Q.; Ayida, B. K.; Takahashi, M.; Winters, G. C.; Simonsen, K. B.; Vourloumis, D.; Hermann, T. "Monitoring molecular recognition of the ribosomal decoding site" Angew. Chem. Int. Ed. 2004, 43, 31773182.

81. Ogle, J. M.; Carter, A. P.; Ramakrishnan, V. "Insights into the decoding mechanism from recent ribosome structures" Trends Biochem. Sci. 2003, 28, 259266.

82. Ogle, J. M.; Brodersen, D. E.; Clemons, W. M.; Tarry, M. J.; Carter, A. P.; Ramakrishnan, V. "Recognition of cognate transfer RNA by the 30S ribosomal subunit" Science 2001, 292, 897902.

83. Barawkar, D. A.; Ganesh, K. N. "Exploring DNA Minor-Groove Interactions through a Probe Conjugate in Major Groove - Fluorescence Studies on Netrosin Complexation with Du-5-Aminodansyl-DNA" Biochem. Biophys. Res. Commun. 1994, 203, 5358.

84. Barawkar, D. A.; Ganesh, K. N. "Fluorescent D(Cgcgaattcgcg) - Characterization of Major Groove Polarity and Study of Minor-Groove Interactions through a Major Groove Semantophore Conjugate" Nucleic Acids Res. 1995, 23, 159164.

85. Jadhav, V. R.; Barawkar, D. A.; Ganesh, K. N. "Polarity sensing by fluorescent oligonucleotides: First demonstration of sequence-dependent microenvironmental changes in the DNA major groove" J. Phys. Chem., B 1999, 103, 73837385.

86. Kimura, T.; Kawai, K.; Majima, T. "Monitoring of microenvironmental changes in the major and minor grooves of DNA by dan-modified oligonucleotides" Org. Lett. 2005, 7, 58295832.

87. Kimura, T.; Kawai, K.; Majima, T. "Probing the microenvironments in the grooves of Z- DNA using dan-modified oligonucleotides" Chem. Commun. 2006, 15421544.

88. Okamoto, A.; Tainaka, K.; Kamei, T. "Sequence-selective osmium oxidation of DNA: efficient distinction between 5-methylcytosine and cytosine" Org. Biomol. Chem. 2006, 4, 16381640.

89. Okamoto, A.; Tainaka, K.; Saito, I. "A dielectric-sensitive fluorescent DNA probe for monitoring polarities on the interior of a DNA-binding protein" Bioconjugate Chem. 2005, 16, 11051111.

90. Tainaka, K.; Tanaka, K.; Ikeda, S.; Nishiza, K.; Unzai, T.; Fujiwara, Y.; Saito, I.; Okamoto, A. "PRODAN-conjugated DNA: Synthesis and photochemical properties" J. Am. Chem. Soc. 2007, 129, 47764784.

91. Sinkeldam, R. W.; Greco, N. J.; Tor, Y. "Polarity of major grooves explored using an isosteric emissive nucleoside" ChemBioChem 2008.

92. Asseline, U. "Development and applications of fluorescent oligonucleotides" Curr. Org. Chem. 2006, 10, 491–518.

93. Hawkins, M. E. "Fluorescent pteridine nucleoside analogs: a window on DNA interactions" Cell. Biochem. Biophys. 2001, 34, 257–281.

94. Rist, M. J.; Marino, J. P. "Fluorescent nucleotide base analogs as probes of nucleic acid structure, dynamics and interactions" Curr. Org. Chem. 2002, 6, 775–793.

95. Cobb, A. J. A. "Recent highlights in modified oligonucleotide chemistry" Org. Biomol. Chem. 2007, 5, 32603275.

96. Okamoto, A.; Saito, Y.; Saito, I. "Design of base-discriminating fluorescent nucleosides" J. Photochem. Photobiol., C 2005, 6, 108122.

97. Wilson, J. N.; Kool, E. T. "Fluorescent DNA base replacements: reporters and sensors for biological systems" Org. Biomol. Chem. 2006, 4, 42654274.

98. Jameson, D. M.; Eccleston, J. F. "Fluorescent nucleotide analogs: synthesis and applications" Methods Enzymol. 1997, 278, 363390.

99. Cremo, C. R. "Fluorescent nucleotides: Synthesis and characterization" Methods Enzymol. 2003, 360, 128177.

100. Hawkins, M. E. "Fluorescent nucleoside analogs as DNA probes" Top. Fluoresc. Spectrosc. 2003, 7, 151175.

101. Ward, D. C.; Reich, E.; Stryer, L. "Fluorescence studies of nucleotides and polynucleotides. I. Formycin, 2-aminopurine riboside, 2,6-diaminopurine riboside, and their derivatives" J. Biol. Chem. 1969, 244, 1228–1237.

102. Evans, K.; Xu, D.; Kim, Y.; Nordlund, T. M. "2-Aminopurine optical spectra: solvent, pentose ring, and DNA helix melting dependence" J. Fluor. 1992, 2, 209–216.

103. Smagowicz, J.; Wierzchowski, K. L. "Lowest excited states of 2-aminopurine" J. Lumin. 1974, 8, 210232.

104. Sowers, L. C.; Boulard, Y.; Fazakerley, G. V. "Multiple structures for the 2- aminopurine-cytosine mispair" Biochemistry 2000, 39, 76137620.

105. Sowers, L. C.; Fazakerley, G. V.; Eritja, R.; Kaplan, B. E.; Goodman, M. F. "Base- Pairing and Mutagenesis - Observation of a Protonated Base Pair between 2- Aminopurine and Cytosine in an Oligonucleotide by Proton Nmr" Proc. Nat. Acad. Sci. U.S.A. 1986, 83, 54345438.

106. Sowers, L. C.; Shaw, B. R.; Veigl, M. L.; Sedwick, W. D. "DNA-Base Modification - Ionized Base-Pairs and Mutagenesis" Mutat. Res. 1987, 177, 201218.

107. Kirk, S. R.; Luedtke, N. W.; Tor, Y. "2-aminopurine as a real-time probe of enzymatic cleavage and inhibition of hammerhead ribozymes" Bioorg. Med. Chem. 2001, 9, 22952301.

108. Serrano-Andres, L.; Merchan, M.; Borin, A. C. "Adenine and 2-aminopurine: paradigms of modern theoretical photochemistry" Proc. Nat. Acad. Sci. U.S.A. 2006, 103, 86918696.

109. Mishra, S. K.; Shukla, M. K.; Mishra, P. C. "Electronic spectra of adenine and 2- aminopurine: an ab initio study of energy level diagrams of different tautomers in gas phase and aqueous solution" Spectrochim. Acta, Part A 2000, 56A, 13551384.

110. Mennucci, B.; Toniolo, A.; Tomasi, J. "Theoretical Study of the Photophysics of Adenine in Solution: Tautomerism, Deactivation Mechanisms, and Comparison with the 2-Aminopurine Fluorescent Isomer" J. Phys. Chem., A 2001, 105, 47494757.

111. Zhang, R.-B.; Ai, X.-C.; Zhang, X.-K.; Zhang, Q.-Y. "Solvent effects on the excited state properties of 2-aminopurine- A theoretical study by the ONIOM and supramolecular method" THEOCHEM 2004, 680, 2127.

112. Jean, J. M.; Hall, K. B. "Theoretical Study of the Excited State Properties and Transitions of 2-Aminopurine in the Gas Phase and in Solution" J. Phys. Chem., A 2000, 104, 19301937.

113. Holmen, A.; Broo, A.; Albinsson, B.; Norden, B. "Assignment of Electronic Transition Moment Directions of Adenine from Linear Dichroism Measurements" J. Am. Chem. Soc. 1997, 119, 1224012250.

114. Broo, A.; Holmen, A. "Ab initio MP2 and DFT calculations of geometry and solution tautomerism of purine and some purine derivatives" Chem. Phys. 1996, 211, 147161.

115. Jean, J. M.; Hall, K. B. "2-Aminopurine fluorescence quenching and lifetimes: role of base stacking" Proc. Nat. Acad. Sci. U.S.A. 2001, 98, 3741.

116. Rachofsky, E. L.; Ross, J. B. A.; Krauss, M.; Osman, R. "CASSCF Investigation of Electronic Excited States of 2-Aminopurine" J. Phys. Chem., A 2001, 105, 190197.

117. Rachofsky, E. L.; Ross, J. B. A.; Krauss, M.; Osman, R. "Spectroscopy of 2- aminopurine: an MCSCF study" Acta Phys. Pol., A 1998, 94, 735748.

118. Broo, A. "A Theoretical Investigation of the Physical Reason for the Very Different Luminescence Properties of the Two Isomers Adenine and 2-Aminopurine" J. Phys. Chem., A 1998, 102, 526531.

119. Rachofsky, E. L.; Seibert, E.; Stivers, J. T.; Osman, R.; Ross, J. B. A. "Conformation and Dynamics of Abasic Sites in DNA Investigated by Time-Resolved Fluorescence of 2-Aminopurine" Biochemistry 2001, 40, 957967.

120. Stivers, J. T. "2-Aminopurine fluorescence studies of base stacking interactions at abasic sites in DNA: metal-ion and base sequence effects" Nucleic Acids Res. 1998, 26, 38373844.

121. Sagher, D.; Strauss, B. "Abasic sites from cytosine as termination signals for DNA synthesis" Nucleic Acids Res. 1985, 13, 42854298.

122. Guest, C. R.; Hochstrasser, R. A.; Sowers, L. C.; Millar, D. P. "Dynamics of Mismatched Base-Pairs in DNA" Biochemistry 1991, 30, 32713279.

123. Raney, K. D.; Sowers, L. C.; Millar, D. P.; Benkovic, S. J. "A Fluorescence-Based Assay for Monitoring Helicase Activity" Proc. Nat. Acad. Sci. U.S.A. 1994, 91, 66446648.

124. Goodman, M. F.; Fygenson, D. K. "DNA polymerase fidelity: from genetics toward a biochemical understanding" Genetics 1998, 148, 14751482.

125. Connolly, B. A.; Newman, P. C. "Synthesis and properties of oligonucleotides containing 4-thiothymidine, 5-methyl-2-pyrimidinone-1-beta-D(2'-deoxyriboside) and 2- thiothymidine" Nucleic Acids Res. 1989, 17, 49574974.

126. Gildea, B.; McLaughlin, L. W. "The synthesis of 2-pyrimidinone nucleosides and their incorporation into oligodeoxynucleotides" Nucleic Acids Res. 1989, 17, 22612281.

127. Kistler, K. A.; Matsika, S. "The fluorescence mechanism of 5-methyl-2-pyrimidinone: an ab initio study of a fluorescent pyrimidine analog" Photochem. Photobiol. 2007, 83, 611624.

128. Kistler, K. A.; Matsika, S. "Radiationless Decay Mechanism of Cytosine: An Ab Initio Study with Comparisons to the Fluorescent Analogue 5-Methyl-2-pyrimidinone" J. Phys. Chem., A 2007, 111, 26502661.

129. Wu, P.; Nordlund, T. M.; Gildea, B.; McLaughlin, L. W. "Base stacking and unstacking as determined from a DNA decamer containing a fluorescent base" Biochemistry 1990, 29, 65086514.

130. Singleton, S. F.; Shan, F.; Kanan, M. W.; McIntosh, C. M.; Stearman, C. J.; Helm, J. S.; Webb, K. J. "Facile Synthesis of a Fluorescent Deoxycytidine Analogue Suitable for Probing the RecA Nucleoprotein Filament" Org. Lett. 2001, 3, 39193922.

131. Rappaport, H. P. "The fidelity of replication of the three-base-pair set adenine/thymine, hypoxanthine/cytosine and 6-thiopurine/5-methyl-2-pyrimidinone with T7 DNA polymerase" Biochem. J. 2004, 381, 709717.

132. Rappaport, H. P. "Replication of the base pair 6-thioguanine/5-methyl-2-pyrimidinone with the large Klenow fragment of Escherichia coli DNA polymerase I" Biochemistry 1993, 32, 30473057.

133. Rappaport, H. P. "The 6-thioguanine/5-methyl-2-pyrimidinone base pair" Nucleic Acids Res. 1988, 16, 72537267.

134. Buchini, S.; Leumann, C. J. "2'-O-Aminoethyl oligoribonucleotides containing novel base analogues: synthesis and triple-helix formation at pyrimidine/purine inversion sites" Eur. J. Org. Chem. 2006, 31523168.

135. Buchini, S.; Leumann, C. J. "Stable and selective recognition of three base pairs in the parallel triple-helical DNA binding motif" Angew. Chem. Int. Ed. 2004, 43, 39253928.

136. Driscoll, S. L.; Hawkins, M. E.; Balis, F. M.; Pfleiderer, W.; Laws, W. R. "Fluorescence properties of a new guanosine analog incorporated into small oligonucleotides" Biophys. J . 1997, 73, 32773286.

137. Hawkins, M. E.; Pfleiderer, W.; Balis, F. M.; Porter, D.; Knutson, J. R. "Fluorescence properties of pteridine nucleoside analogs as monomers and incorporated into oligonucleotides" Anal. Biochem. 1997, 244, 8695.

138. Hawkins, M. E.; Pfleiderer, W.; Mazumder, A.; Pommier, Y. G.; Falls, F. M. "Incorporation of a Fluorescent Guanosine Analog into Oligonucleotides and Its Application to a Real-Time Assay for the Hiv-1 Integrase 3'-Processing Reaction" Nucleic Acids Res. 1995, 23, 28722880.

139. Brown, P. O.; Bowerman, B.; Varmus, H. E.; Bishop, J. M. "Retroviral integration: structure of the initial covalent product and its precursor, and a role for the viral IN protein" Proc. Nat. Acad. Sci. U.S.A. 1989, 86, 25252529.

140. Vink, C.; Banks, M.; Bethell, R.; Plasterk, R. H. "A high-throughput, non-radioactive microtiter plate assay for activity of the human immunodeficiency virus integrase protein" Nucleic Acids Res. 1994, 22, 21762177.

141. Wojtuszewski, K.; Hawkins, M. E.; Cole, J. L.; Mukerji, I. "HU binding to DNA: Evidence for multiple complex formation and DNA bending" Biochemistry 2001, 40, 25882598.

142. Augustyn, K. E.; Wojtuszewski, K.; Hawkins, M. E.; Knutson, J. R.; Mukerji, I. "Examination of the Premelting Transition of DNA A-Tracts Using a Fluorescent Adenosine Analogue" Biochemistry 2006, 45, 50395047.

143. Secrist, J. A.; Barrio, J. R.; Leonard, N. J. "Fluorescent Modification of Adenosine- Triphosphate with Activity in Enzyme-Systems - 1,N6-Ethenoadenosine Triphosphate" Science 1972, 175, 646–647.

144. Secrist, J. A.; Weber, G.; Leonard, N. J.; Barrio, J. R. "Fluorescent Modification of Adenosine-Containing Coenzymes - Biological-Activities and Spectroscopic Properties" Biochemistry 1972, 11, 34993506.

145. Scopes, D. I. C.; Barrio, J. R.; Leonard, N. J. "Defined Dimensional Changes in Enzyme Cofactors - Fluorescent Stretched-out Analogs of Adenine-Nucleotides" Science 1977, 195, 296298.

146. Kochetko, N. K.; Shibaev, V. N.; Kost, A. A. "New Reaction of Adenine and Cytosine Derivatives, Potentially Useful for Nucleic Acids Modification" Tetrahedron Lett. 1971, 12, 19931996.

147. RajBhandary, U. L.; Chang, S. H.; Stuart, A.; Faulkner, R. D.; Hoskinson, R. M.; Khorana, H. G. "Studies on polynucleotides. LXVIII. The primary structure of yeast phenylalanine transfer RNZ" Proc. Nat. Acad. Sci. U.S.A. 1967, 57, 751758.

148. Gao, J.; Liu, H.; Kool, E. T. "Expanded-size bases in naturally sized DNA: evaluation of steric effects in Watson-Crick pairing" J. Am. Chem. Soc. 2004, 126, 11826–11831.

149. Liu, H.; Gao, J.; Maynard, L.; Saito, Y. D.; Kool, E. T. "Toward a new genetic system with expanded dimensions: size-expanded analogues of deoxyadenosine and thymidine" J. Am. Chem. Soc. 2004, 126, 1102–1109.

150. Holmén, A.; Albinsson, B.; Nordén, B. "Electronic-Transition Dipole-Moments of the 1,N-6-Ethenoadenine Chromophore" J. Phys. Chem. 1994, 98, 13460–13469.

151. Lee, A. H.; Kool, E. T. "Novel benzopyrimidines as widened analogues of DNA bases" J. Org. Chem. 2005, 70, 132–140.

152. Lee, A. H.; Kool, E. T. "Exploring the limits of DNA size: naphtho-homologated DNA bases and pairs" J. Am. Chem. Soc. 2006, 128, 9219–9230.

153. Liu, H.; Gao, J.; Kool, E. T. "Helix-forming properties of size-expanded DNA, an alternative four-base genetic form" J. Am. Chem. Soc. 2005, 127, 1396–1402.

154. Liu, H.; Gao, J.; Kool, E. T. "Size-expanded analogues of dG and dC: synthesis and pairing properties in DNA" J. Org. Chem. 2005, 70, 639–647.

155. Lu, H.; He, K.; Kool, E. T. "yDNA: a new geometry for size-expanded base pairs" Angew. Chem. Int. Ed. 2004, 43, 5834–5836.

156. Crisp, G. T.; Flynn, B. L. "Palladium-Catalyzed Coupling of Terminal Alkynes with 5- (Trifluoromethanesulfonyloxy)Pyrimidine Nucleosides" J. Org. Chem. 1993, 58, 66146619.

157. Robins, M. J.; Barr, P. J. "Nucleic-Acid Related-Compounds .31. Smooth and Efficient Palladium-Copper Catalyzed Coupling of Terminal Alkynes with 5-Iodouracil Nucleosides" Tetrahedron Lett. 1981, 22, 421424.

158. Robins, M. J.; Barr, P. J. "Nucleic-Acid Related-Compounds .39. Efficient Conversion of 5-Iodo to 5-Alkynyl and Derived 5-Substituted Uracil Bases and Nucleosides" J. Org. Chem. 1983, 48, 18541862.

159. McGuigan, C.; Yarnold, C. J.; Jones, G.; Velazquez, S.; Barucki, H.; Brancale, A.; Andrei, G.; Snoeck, R.; De Clercq, E.; Balzarini, J. "Potent and selective inhibition of varicella-zoster virus (VZV) by nucleoside analogues with an unusual bicyclic base" J. Med. Chem. 1999, 42, 44794484.

160. Loakes, D.; Brown, D. M.; Salisbury, S. A.; McDougall, M. G.; Neagu, C.; Nampalli, S.; Kumar, S. "Synthesis and some biochemical properties of a novel 5,6,7,8- tetrahydropyrimido[4,5-c]pyridazine nucleoside" HeIv. Chim. Acta 2003, 86, 1193– 1204.

161. Berry, D. A.; Jung, K. Y.; Wise, D. S.; Sercel, A. D.; Pearson, W. H.; Mackie, H.; Randolph, J. B.; Somers, R. L. "Pyrrolo-dC and pyrrolo-C: fluorescent analogs of cytidine and 2 '-deoxycytidine for the study of oligonucleotides" Tetrahedron Lett. 2004, 45, 2457–2461.

162. Tinsley, R. A.; Walter, N. G. "Pyrrolo-C as a fluorescent probe for monitoring RNA secondary structure formation" RNA 2006, 12, 522529.

163. Dash, C.; Rausch, J. W.; Le Grice, S. F. J. "Using pyrrolo-deoxycytosine to probe RNA/DNA hybrids containing the human immunodeficiency virus type-1 3 ' polypurine tract" Nucleic Acids Res. 2004, 32, 15391547.

164. Liu, C. H.; Martin, C. T. "Promoter clearance by T7 RNA polymerase - Initial bubble collapse and transcript dissociation monitored by base analog fluorescence" J. Biol. Chem. 2002, 277, 27252731.

165. Marti, A. A.; Jockusch, S.; Li, Z. M.; Ju, J. Y.; Turro, N. J. "Molecular beacons with intrinsically fluorescent nucleotides" Nucleic Acids Res. 2006, 34, e50.

166. Johnson, N. P.; Baase, W. A.; von Hippel, P. H. "Low-energy circular dichroism of 2- aminopurine dinucleotide as a probe of local conformation of DNA and RNA" Proc. Nat. Acad. Sci. U.S.A. 2004, 101, 34263431.

167. Johnson, N. P.; Baase, W. A.; von Hippel, P. H. "Investigating local conformations of double-stranded DNA by low-energy circular dichroism of pyrrolo-cytosine" Proc. Nat. Acad. Sci. U.S.A. 2005, 102, 71697173.

168. Thompson, K. C.; Miyake, N. "Properties of a new fluorescent cytosine analogue, pyrrolocytosine" J. Phys. Chem., B 2005, 109, 60126019.

169. Godde, F.; Toulmé, J. J.; Moreau, S. "Benzoquinazoline derivatives as substitutes for thymine in nucleic acid complexes. Use of fluorescence emission of benzo[g]quinazoline-2,4-(1H,3H)-dione in probing duplex and triplex formation" Biochemistry 1998, 37, 13765–13775.

170. Arzumanov, A.; Godde, F.; Moreau, S.; Toulmè, J. J.; Weeds, A.; Gait, M. J. "Use of the fluorescent nucleoside analogue benzo[g]quinazoline 2 '-O-methyl-beta-D- ribofuranoside to monitor the binding of the HIV-1 Tat protein or of antisense oligonucleotides to the TAR RNA stem-loop" HeIv. Chim. Acta 2000, 83, 1424–1436.

171. Lin, K. Y.; Jones, R. J.; Matteucci, M. "Tricyclic 2'-Deoxycytidine Analogs - Syntheses and Incorporation into Oligodeoxynucleotides Which Have Enhanced Binding to Complementary Rna" J. Am. Chem. Soc. 1995, 117, 38733874.

172. Eldrup, A. B.; Nielsen, B. B.; Haaima, G.; Rasmussen, H.; Kastrup, J. S.; Christensen, C.; Nielsen, P. E. "1,8-Naphthyridin-2(1H)-ones - Novel bicyclic and tricyclic analogues of thymine in peptide nucleic acids (PNAs)" Eur. J. Org. Chem. 2001, 17811790.

173. Wilhelmsson, L. M.; Holmén, A.; Lincoln, P.; Nielsen, P. E.; Nordén, B. "A highly fluorescent DNA base analogue that forms Watson-Crick base pairs with guanine" J. Am. Chem. Soc. 2001, 123, 2434–2435.

174. Engman, K. C.; Sandin, P.; Osborne, S.; Brown, T.; Billeter, M.; Lincoln, P.; Nordén, B.; Albinsson, B.; Wilhelmsson, L. M. "DNA adopts normal B-form upon incorporation of highly fluorescent DNA base analogue tC: NMR structure and UV-Vis spectroscopy characterization" Nucleic Acids Res. 2004, 32, 5087–5095.

175. Sandin, P.; Wilhelmsson, L. M.; Lincoln, P.; Powers, V. E.; Brown, T.; Albinsson, B. "Fluorescent properties of DNA base analogue tC upon incorporation into DNA-- negligible influence of neighbouring bases on fluorescence quantum yield" Nucleic Acids Res. 2005, 33, 5019–5025.

176. Matray, T. J.; Kool, E. T. "A specific partner for abasic damage in DNA" Nature 1999, 399, 704708.

177. Gao, J. M.; Strassler, C.; Tahmassebi, D.; Kool, E. T. "Libraries of composite polyfluors built from fluorescent deoxyribosides" J. Am. Chem. Soc. 2002, 124, 1159011591.

178. Gao, J. M.; Watanabe, S.; Kool, E. T. "Modified DNA analogues that sense light exposure with color changes" J. Am. Chem. Soc. 2004, 126, 1274812749.

179. Ren, R. X. F.; Chaudhuri, N. C.; Paris, P. L.; Rumney, S.; Kool, E. T. "Naphthalene, phenanthrene, and pyrene as DNA base analogues: Synthesis, structure, and fluorescence in DNA" J. Am. Chem. Soc. 1996, 118, 7671–7678.

180. Strässler, C.; Davis, N. E.; Kool, E. T. "Novel nucleoside analogues with fluorophores replacing the DNA base" HeIv. Chim. Acta 1999, 82, 2160–2171.

181. Christensen, U. B.; Pedersen, E. B. "Intercalating nucleic acids containing insertions of 1-O-(1-pyrenylmethyl)glycerol: stabilisation of dsDNA and discrimination of DNA over RNA" Nucleic Acids Res. 2002, 30, 49184925.

182. Aubert, Y.; Asseline, U. "Synthesis and hybridization properties of oligonucleotide perylene conjugates: influence of the conjugation parameters on triplex and duplex stabilities" Org. Biomol. Chem. 2004, 2, 34963503.

183. Coleman, R. S.; Madaras, M. L. "Synthesis of a novel coumarin C-riboside as a photophysical probe of oligonucleotide dynamics" J. Org. Chem. 1998, 63, 5700– 5703.

184. Kitamura, N.; Fukagawa, T.; Kohtani, S.; Kitoh, S.; Kunimoto, K. K.; Nakagaki, R. "Synthesis, absorption, and fluorescence properties and crystal structures of 7- aminocoumarin derivatives" J. Photochem. Photobiol., A 2007, 188, 378386.

185. Horng, M. L.; Gardecki, J. A.; Papazyan, A.; Maroncelli, M. "Subpicosecond Measurements of Polar Solvation Dynamics - Coumarin-153 Revisited" J. Phys. Chem. 1995, 99, 1731117337.

186. Moog, R. S.; Davis, W. W.; Ostrowski, S. G.; Wilson, G. L. "Solvent effects on electronic transitions in several coumarins" Chem. Phys. Lett. 1999, 299, 265271.

187. Brauns, E. B.; Madaras, M. L.; Coleman, R. S.; Murphy, C. J.; Berg, M. A. "Measurement of local DNA reorganization on the picosecond and nanosecond time scales" J. Am. Chem. Soc. 1999, 121, 11644–11649.

188. Brauns, E. B.; Madaras, M. L.; Coleman, R. S.; Murphy, C. J.; Berg, M. A. "Complex local dynamics in DNA on the picosecond and nanosecond time scales" Phys. Rev. Lett. 2002, 88, 158101158104.

189. Gearheart, L. A.; Somoza, M. M.; Rivers, W. E.; Murphy, C. J.; Coleman, R. S.; Berg, M. A. "Sodium-ion binding to DNA: detection by ultrafast time-resolved stokes-shift spectroscopy" J. Am. Chem. Soc. 2003, 125, 11812–11813.

190. Sen, S.; Gearheart, L. A.; Rivers, E.; Liu, H.; Coleman, R. S.; Murphy, C. J.; Berg, M. A. "Role of monovalent counterions in the ultrafast dynamics of DNA" J. Phys. Chem. B 2006, 110, 13248–13255.

191. Somoza, M. M.; Andreatta, D.; Murphy, C. J.; Coleman, R. S.; Berg, M. A. "Effect of lesions on the dynamics of DNA on the picosecond and nanosecond timescales using a polarity sensitive probe" Nucleic Acids Res. 2004, 32, 2494–2507.

192. Andreatta, D.; Perez Lustres, J. L.; Kovalenko, S. A.; Ernsting, N. P.; Murphy, C. J.; Coleman, R. S.; Berg, M. A. "Power-law solvation dynamics in DNA over six decades in time" J. Am. Chem. Soc. 2005, 127, 7270–7271.

193. Andreatta, D.; Sen, S.; Perez Lustres, J. L.; Kovalenko, S. A.; Ernsting, N. P.; Murphy, C. J.; Coleman, R. S.; Berg, M. A. "Ultrafast dynamics in DNA: "fraying" at the end of the helix" J. Am. Chem. Soc. 2006, 128, 6885–6892.

194. Sen, S.; Paraggio, N. A.; Gearheart, L. A.; Connor, E. E.; Issa, A.; Coleman, R. S.; Wilson, D. M., 3rd; Wyatt, M. D.; Berg, M. A. "Effect of protein binding on ultrafast DNA dynamics: characterization of a DNA:APE1 complex" Biophys. J . 2005, 89, 4129–4138.

195. Coleman, R. S.; Mortensen, M. A. "Stereocontrolled synthesis of anthracene beta-C- ribosides: fluorescent probes for photophysical studies of DNA" Tetrahedron Lett. 2003, 44, 1215–1219.

196. Hurley, D. J.; Seaman, S. E.; Mazura, J. C.; Tor, Y. "Fluorescent 1,10-phenanthroline- containing oligonucleotides distinguish between perfect and mismatched base pairing" Org. Lett. 2002, 4, 2305–2308.

197. Seela, F.; Zulauf, M.; Sauer, M.; Deimel, M. "7-substituted 7-deaza-2 '- deoxyadenosines and 8-aza-7-deaza-2 '-deoxyadenosines: Fluorescence of DNA- base analogues induced by the 7-alkynyl side chain" HeIv. Chim. Acta 2000, 83, 910– 927.

198. Okamoto, A.; Kanatani, K.; Saito, I. "Pyrene-labeled base-discriminating fluorescent DNA probes for homogeneous SNP typing" J. Am. Chem. Soc. 2004, 126, 4820– 4827.

199. Ahmadian, M.; Zhang, P. M.; Bergstrom, D. E. "A comparative study of the thermal- stability of oligodeoxyribonucleotides containing 5-substituted 2 ' deoxyuridines" Nucleic Acids Res. 1998, 26, 31273135.

200. Kerr, C. E.; Mitchell, C. D.; Headrick, J.; Eaton, B. E.; Netzel, T. L. "Synthesis and photophysics of a 1-pyrenyl substituted 2 '-deoxyuridine-5-carboxamide nucleoside: Electron transfer products as CISINDO/S excited states" J. Phys. Chem., B 2000, 104, 1637–1650.

201. Netzel, T. L.; Zhao, M.; Nafisi, K.; Headrick, J.; Sigman, M. S.; Eaton, B. E. "Photophysics of 2'-Deoxyuridine (Du) Nucleosides Covalently Substituted with Either 1-Pyrenyl or 1-Pyrenoyl - Observation of Pyrene-to-Nucleoside Charge-Transfer Emission in 5-(1-Pyrenyl)-Du" J. Am. Chem. Soc. 1995, 117, 9119–9128.

202. Joshi, H. S.; Jamshidi, R.; Tor, Y. "Conjugated 1,10-phenanthrolines as tunable fluorophores" Angew. Chem. Int. Ed. 1999, 38, 27222725.

203. Hurley, D. J.; Tor, Y. "Donor/Acceptor Interactions in Systematically Modified RuII-OsII Oligonucleotides" J. Am. Chem. Soc. 2002, 124, 1323113241.

204. Hurley, D. J.; Tor, Y. "Ru(II) and Os(II) Nucleosides and Oligonucleotides: Synthesis and Properties" J. Am. Chem. Soc. 2002, 124, 37493762.

205. Hurley, D. J.; Tor, Y. "Metal-Containing Oligonucleotides: Solid-Phase Synthesis and Luminescence Properties" J. Am. Chem. Soc. 1998, 120, 21942195.

206. Suh, Y.; Cantor, C.; ed. "Single nucleotide polymorphisms (SNPs): detection, interpretation, and application" Mutat. Res. 2005, 573, 1220.

207. Okamoto, A.; Tainaka, K.; Saito, I. "Clear distinction of purine bases on the complementary strand by a fluorescence change of a novel fluorescent nucleoside" J. Am. Chem. Soc. 2003, 125, 49724973.

208. Okamoto, A.; Tanaka, K.; Fukuta, T.; Saito, I. "Design of base-discriminating fluorescent nucleoside and its application to T/C SNP typing" J. Am. Chem. Soc. 2003, 125, 92969297.

209. Manoharan, M.; Tivel, K. L.; Zhao, M.; Nafisi, K.; Netzel, T. L. "Base-Sequence Dependence of Emission Lifetimes for DNA Oligomers and Duplexes Covalently Labeled with Pyrene - Relative Electron-Transfer Quenching Efficiencies of a- Nucleoside, G-Nucleoside, C-Nucleoside, and T-Nucleoside toward Pyrene" J. Phys. Chem. 1995, 99, 1746117472.

210. Mayer, E.; Valis, L.; Huber, R.; Amann, N.; Wagenknecht, H. A. "Preparation of pyrene-modified purine and pyrimidine nucleosides via Suzuki-Miyaura cross- couplings and characterization of their fluorescent properties" Synthesis 2003, 23352340.

211. Shafirovich, V. Y.; Courtney, S. H.; Ya, N. Q.; Geacintov, N. E. "Proton-Coupled Photoinduced Electron-Transfer, Deuterium-Isotope Effects, and Fluorescence Quenching in Noncovalent Benzo[a]Pyrenetetraol - Nucleoside Complexes in Aqueous-Solutions" J. Am. Chem. Soc. 1995, 117, 49204929.

212. Saito, Y.; Miyauchi, Y.; Okamoto, A.; Saito, I. "Synthesis and properties of novel base- discriminating fluorescent (BDF) nucleosides: a highly polarity-sensitive fluorophore for SNP typing" Tetrahedron Lett. 2004, 45, 78277831.

213. Hwang, G. T.; Seo, Y. J.; Kim, S. J.; Kim, B. H. "Fluorescent oligonucleotide incorporating 5-(1-ethynylpyrenyl)2 '-deoxyuridine: sequence-specific fluorescence changes upon duplex formation" Tetrahedron Lett. 2004, 45, 35433546.

214. Okamoto, A.; Ochi, Y.; Saito, I. "Modulation of base selectivity for a base- discriminating fluorescent nucleobase by addition of mercury ion" Bioorg. Med. Chem. Lett. 2005, 15, 42794281.

215. Okamoto, A.; Tainaka, K.; Nishiza, K.; Saito, I. "Monitoring DNA structures by dual fluorescence of pyrene derivatives" J. Am. Chem. Soc. 2005, 127, 1312813129.

216. Saito, Y.; Miyauchi, Y.; Okamoto, A.; Saito, I. "Base-discriminating fluorescent (BDF) nucleoside: distinction of thymine by fluorescence quenching" Chem. Commun. 2004, 17041705.

217. Okamoto, A.; Tainaka, K.; Saito, I. "Synthesis and properties of a novel fluorescent nucleobase, naphthopyridopyrimidine" Tetrahedron Lett. 2003, 44, 68716874.

218. Seela, F.; Feiling, E.; Gross, J.; Hillenkamp, F.; Ramzaeva, N.; Rosemeyer, H.; Zulauf, M. "Fluorescent DNA: the development of 7-deazapurine nucleoside triphosphates

applicable for sequencing at the single molecule level" Journal of Biotechnology 2001, 86, 269–279.

219. Seela, F.; Zulauf, M. "7-deazaadenine-DNA: Bulky 7-iodo substituents or hydrophobic 7-hexynyl chains are well accommodated in the major groove of oligonucleotide duplexes" Chemistry-a European Journal 1998, 4, 1781-1790.

220. Ranasinghe, R. T.; Brown, T. "Fluorescence based strategies for genetic analysis" Chemical Communications (Cambridge, United Kingdom) 2005, 5487-5502.

221. May, J. P.; Brown, L. J.; van Delft, I.; Thelwell, N.; Harley, K.; Brown, T. "Synthesis and evaluation of a new non-fluorescent quencher in fluorogenic oligonucleotide probes for real-time PCR" Org. Biomol. Chem. 2005, 3, 2534-2542.

222. McKeen, C. M.; Brown, L. J.; Nicol, J. T. G.; Mellor, J. M.; Brown, T. "Synthesis of fluorophore and quencher monomers for use in Scorpion primers and nucleic acid structural probes" Org. Biomol. Chem. 2003, 1, 2267-2275.

223. Xiao, Q.; Ranasinghe, R. T.; Tang, A. M. P.; Brown, T. "Naphthalenyl- and anthracenyl-ethynyl dT analogues as base discriminating fluorescent nucleosides and intramolecular energy transfer donors in oligonucleotide probes" Tetrahedron 2007, 63, 3483-3490.

224. Seela, F.; Jawalekar, A. M.; Sun, L.; Leonard, P. "Oligonucleotides Containing Pyrazolo[3,4-d]Pyrimidines: 8-Aza-7-deazaadenines With Bulky Substituents in the 2- or 7-Position" Nucleosides, Nucleotides & Nucleic Acids 2005, 24, 1485-1505.

225. Seela, F.; Sirivolu, V. R. "DNA containing side chains with terminal triple bonds: base- pair stability and functionalization of alkynylated pyrimidines and 7-deazapurines" Chemistry & Biodiversity 2006, 3, 509-514.

226. Kimoto, M.; Mitsui, T.; Harada, Y.; Sato, A.; Yokoyama, S.; Hirao, I. "Fluorescent probing for RNA molecules by an unnatural base-pair system" Nucleic Acids Res. 2007, 35, 5360-5369.

227. Kawai, R.; Kimoto, M.; Ikeda, S.; Mitsui, T.; Endo, M.; Yokoyama, S.; Hirao, I. "Site- Specific Fluorescent Labeling of RNA Molecules by Specific Transcription Using Unnatural Base Pairs" J. Am. Chem. Soc. 2005, 127, 17286-17295.

Chapter 2

Design, synthesis and application of novel fluorescent nucleosides.

2.1 – Fluorescent nucleoside design principles

Fluorescence spectroscopy provides biophysicists with incredible opportunities to explore

the structure, dynamics, and recognition of biomolecules. Nucleic acids, however, present a

challenge since the common pyrimidines and purines are practically non-emissive.1-9

Adenosine, for example, displays a quantum yield of 0.00005.6 To remedy this limitation, numerous fluorescent nucleoside analogues have been synthesized and implemented (see chapter 1 for more details).

Favorable photophysical characteristics are typically associated with significant structural perturbation when compared to the heterocycles found in the native nucleosides. A large majority of fluorescent nucleosides (see chapter 1) utilize large structural motifs, thus generating the desired photophysical properties, although simultaneously deviating from the size and shape of the natural nucleobases. It is with this in mind that we have set out to define some guiding principles for the design of novel fluorescent nucleosides.

These novel fluorescent nucleosides must:

(a) maintain the highest structural similarity to the natural nucleobases and ensure the

formation of W-C like base pairs. The size, shape and hydrogen-bonding face of a

modified nucleoside are vital considerations, for the insertion of any probe will

inevitably change the system of interest, thereby tainting the readout.

(b) display an absorption band isolated from that of the native nucleobases (~260 nm)

thus resulting in long wavelength emission, preferably in the visible range. Although

natural nucleic acids evolved to have very short lifetimes and weak quantum

51 52

efficiencies, they are still known to undergo photochemistry in the excited state,

albeit with minimal efficiency. Therefore it is ideal to selectively excite the novel

probe in the presence of all other chromophores.

phenomenon that upon incorporation into an oligonucleotide the quantum efficiency

of a fluorescent nucleoside will be severely attenuated.10 The lack of a correlation

between quantum efficiency of nucleosides before and after incorporation makes

the quantum efficiency of the isolated nucleosides informative, while adequate

quantum efficiency after incorporation is vital.

(d) respond to changes in their microenvironment via marked differences in their basic

photophysical parameters (e.g. emission wavelength em, quantum efficiency F,

and/or or excited state lifetime ). A fluorescent probe must have the ability to

distinguish between different environments and report these findings if it is to be

utilized. It is the last of these requirements that is so difficult to predict, yet so vital

to the development of biologically relevant nucleosides.

Although the above four ideal properties seem straightforward, they are in conflict with each other, thereby denoting the challenge associated with designing relevant novel relevant fluorescent nucleosides. For example, it is well accepted that extended conjugation will red shift the absorption spectra of a chromophore, thus satisfying requirement b. Yet any serious structural addition to the conjugation of the natural nucleobases will ultimately violate requirement a. This creates a situation in which it is virtually impossible to design a probe that will fulfill all of the above-mentioned requirements, but instead presents the idea that certain 53

fluorescent nucleosides because of their fulfillment of these requirements will be useful in some situations and not in others.

2.2 – Inspiration

The basic design principle underlining the fluorescent nucleosides contained within this

chapter relies on conjugating aromatic five membered rings to natural nucleosides.

Examination of the photophysical properties of simple conjugated heterocycles reveals

intriguing fluorescent properties. Comparing the absorption and emission spectra of benzene

and furan to that of the corresponding 2-phenylfuran, for example, clearly illustrates the

favorable features of the conjugated system (figure 2.1). While benzene and furan have

relatively weak absorption bands above 250 nm, the conjugated 2-phenylfuran, under the

same experimental conditions, displays an intense absorption around 280 nm ( = 20,000).

This is associated with a strong emission band (F = 0.4) centered around 320 nm, a feature

that is completely absent in either of the aromatic precursors. Thus, we have chosen to

explore a series of nucleoside analogues where an aromatic 5 membered ring (furan,

thiophene, thiazole and oxazole) is conjugated at the 5 position of uridine/thymidine (figure

2.2).

6 benzene cps -1 6 O cm -1 4 M 1 4 furan 10 O / x

ε 2 PL Intensity / x 10 PL

0 0 2-phenylfuran 200 250 300 350 400 λ / nm Figure 2.1. Absorption and steady state emission spectra of furan (red), benzene (blue) and 2-phenylfuran (green) in hexanes (1.0 10-6 M). 54

O O N O N O

O NH S NH O NH S NH

N O N O HO HO HO N O HO N O O O O O

HO 1 HO 2 HO 3 HO 4

Figure 2.2. Thymidine analogues where a five-membered heterocycle is attached at the 5- position.

2.3 – Synthesis of thymidine analogues

Access to 5-(furyl-2-yl) and 5-(thiophen-2-yl) 2-deoxyuridine analogues, nucleosides 1 and

2 respectively, was accomplished in a similar fashion to previously published procedures.11,12

Stille coupling of commercially available 5-iodo-2-deoxyuridine under standard conditions

results in the desired products 1 and 2 in good yields (Scheme 2.1).

O O I NH X NH

HO N O HO N O O a O

HO 5 HO 1 X=O 2 X=S

Scheme 2.1. (a) 1: 2-(tributylstannyl)furan, PdCl2(PPh3)2, dioxane, 94%; 2: 2- (tributylstannyl)thiophene, PdCl2(PPh3)2, dioxane, 80%.

Extension of this procedure to include the oxazolyl and thiazolyl analogues via Stille

coupling to 5-iodo-2-deoxyuridine was unsuccessful. Protection of 3 and 5 hydroxyls as

toluoyl esters was required for synthesis of the oxazolyl and thiazolyl dT analogues (Scheme

2.2). Stille coupling to the now protected 5-iodo-2-deoxyuridine resulted in desired

nucleosides 3 and 4, albeit in lower yield than the corresponding furanyl or thienyl analogues. 55

Despite the low yielding synthesis of the oxazolyl and thiazolyl analogues, sufficient quantities for photophysical evaluation were easily achieved.

O O N O I I NH O NH X NH

HO N O O N O HO N O O a O b O

O HO 5 O 6 HO 3 X=O 4 X=S

Scheme 2.2. (a) p-toluoyl chloride, pyridine, 85%; (b) 3: (i) 2-(tributylstannyl)oxazole,

Pd(PPh3)4, toluene; (ii) potassium carbonate, 5% tetrahydrofuran/methanol, 10%; 4: (i) 2- (tributylstannyl)thiazole, PdCl2(PPh3)2, dioxane; (ii) potassium carbonate, 5% tetrahydrofuran/methanol, 34%.

2.4 – Isomorphicity of modified nucleosides

Retaining the size and shape of the native nucleobases is very important in our design.

Crystallization of all four modified nucleosides allowed for comparison to the native thymidine

structure (figure 2.3). Visual inspection identifies that in the solid state the modification of

conjugating a five membered ring to the 5-position does not induce any major structural

changes. The positions of the sugars and heterocycles of all four modified nucleosides are

almost identical to thymidine itself. In the solid state, there is an interaction between the C-4

carbonyl and the corresponding hetero-atom in the five membered ring. The oxygen

containing heterocycles, furanyl and oxazolyl, crystallize with the ring planar to the dU core,

but with the oxygen opposed to the C-4 carbonyl, while the sulfur containing heterocycles,

thienyl and thiazolyl, crystallize again in a planar fashion to the dU core, but with the sulfur

facing the C-4 carbonyl. All four nucleosides are the result of one structure with the

heterocyclic five membered ring in one orientation, thereby suggesting a disfavorable 56

interaction with the oxygen species and a favorable one with the sulfur species. This interaction between sulfur and a proximal carbonyl has been documented in other thienyl containing molecules.13 It is interesting to note that extension of the thienyl moiety away from

the dU core (see chapter 3) results in a 50/50 mixture of structures, presumably due to the

loss of the sulfur-carbonyl interaction.

dT Furanyl dU (1) Thienyl dU (2)

4 Oxazolyl dU (3) Thiazolyl dU ( ) Figure 2.3. X-ray structures of the furanyl, thienyl, oxazolyl and thiazolyl dU analogues in comparison to dT.14

2.5 – Photophysical properties of thymidine analogues

Although synthetic procedures of nucleosides 1, 2 and 4 were previously reported in the

11,12 literature as early as 1991, no photophysical data, excluding their max of absorption, were reported. Rewardingly, all modified nucleosides were found to be emissive. They differ, however, in their quantum efficiency of emission and sensitivity to their microenvironment polarity (figure 2.4 and Table 2.1). The absorption spectra of each modified nucleoside reveals a low energy absorption band at ~315 nm for 1, 2 and 4 and ~295 nm for 3, in addition 57

to the typical higher energy pyrimidine absorption band at ~268 nm. The max of absorption for all four nucleosides remains practically unchanged upon changing solvent polarity (Table 2.1).

In contrast, the emission profile of the conjugated nucleosides is much more sensitive to the chromophores microenvironment, with bathochromic (7–36 nm) and hyperchromic (1.0–5.6 fold) shifts upon increase in solvent polarity from ether to water. In a non-polar environment such as ether, all four nucleosides show emission at the higher energy end of the visible range

(390–421 nm), with relatively large Stokes shifts (79–101 nm). In water, however, all four nucleosides show emission in the visible range (400–434 nm), with even larger Stokes shifts

(88–115 nm). Notably, the quantum efficiency of all four nucleosides is relatively low (0.01–

0.03; Table 2.1) although sufficient enough, due to the sensitivity of modern

spectrofluorometers, for use as fluorescent probes in DNA/RNA systems. Comparing the

quantum efficiencies of the model system, 2-phenylfuran (F = 0.4), to nucleosides 14 (F =

0.03–0.01), reveals a ~13–40 fold decrease in quantum efficiency. This is not surprising since

15 the quantum efficiency of benzene (F = 0.07), a component of the model system, is ~500

7 times more emissive than thymidine (F = 0.000132).

12 8 –1 cm

–1 8 M 3 4 PL Intensity PL

/ × 10 4 ε

0 0 250 350 450 550 λ / nm Figure 2.4. Absorption (5.0 10-5 M) and steady state emission (1.0 10-5 M) spectra of 1 (black), 2 (red), 3 (green) and 4 (blue) in water.

Table 2.1. Photophysical data of nucleosides 1–4.

cmpd max Et2O max H2O em Et2O em H2O I F (nm) (nm) (nm) (nm) H2O/Et2O H2O 1 314 316 395 431 5.6 0.03 2 320 314 421 434 1.6 0.01 3 292 296 390 400 1.0 <0.01 4 318 316 397 404 2.1 <0.01

Table 2.2. Photophysical data of furanyl dT analogue 1. ET(30) values were experimentally determined using Reichardts salt.18

Solvent ET(30) / abs / nm em / nm em Energy / Stokes F kcal mol-1 cm-1 Shift / cm-1 Water 63.1 316 431 23200 8400 0.03 Methanol 55.2 316 422 23700 7900 --- Acetonitrile 45.9 316 402 24900 6800 --- Dichloromethane 41.4 316 401 24900 6700 --- Ethyl Ether 34.9 316 395 25300 6300 ---

2.6 – Comparison to 2-aminopurine

The comparison of the furanyl dT analogue 1, having the most desirable photophysical properties of the four thymidine analogues, to existing fluorescent probes allows a true evaluation of its photophysical properties. 2-aminopurine (2-AP), being one of the most commonly used fluorescent nucleosides to date as well as its isomorphic character, make it a great comparison to the furanyl thymidine analogue 1. While the quantum efficiency of 2-AP is remarkably greater (0.68 vs. 0.03) than the furanyl dT analogue, its sensitivity to its surrounding environment and lack of an isolated absorption band make it less ideal than the furanyl dT analogue 1. The furanyl dT analogue is more sensitive to polarity changes in its surrounding environment than 2-AP, as seen by the greater negative slope and the larger spectral window (figure 2.5, table 2.2 and table 2.3). It should be noted that this comparison is simply noting the effectiveness that one can design novel fluorescent nucleosides with comparable photophysical parameters to those already being used as biological probes. It is 59

not however suggesting that the furanyl dT analogue can nor should be utilized in the exact same manner that 2-AP can, albeit in select examples.16,17

18 Table 2.3. Photophysical data of 2-AP. ET(30) values are from Reichardts Review. Absorption and emission maxima without parentheses were taken from Nordlund19 and those 20 in parentheses from Wierzchowski . F values without parentheses are for 9-ethyl-2- aminopurine, taken from Stryer21 and those in parentheses were taken from Wierzchowski20. E (30) / Energy / Stokes Solvent T / nm / nm em kcal mol-1 abs em cm-1 Shift / cm-1 F 305 369 27100 5687 0.68 Water 63.1 (305) (377) (26500) (6300) (0.86) Methanol 55.4 309 367.5 27211 5152 --- (---) Ethanol 51.9 (310) (370) (27050) (5250) --- (0.55) 1-Propanol 50.7 308.5 366.5 27285 5130 --- (---) Butanol 49.7 305 354 28249 4538 --- (---) Acetonitrile 45.6 (304) (360) (27800) (5100) --- (0.08) DMSO 45.1 311 369 27100 5054 --- (---) DMF 43.2 (311) (369) (27100) (5050) --- (0.12) 305.5 354 28249 4485 Ethyl Acetate 38.1 --- (0.05) (306) (358) (27920) (4730) 1,4-Dioxane 36.0 306 359 27855 4825 0.13 (---) 305.5 354 28249 4485 Ethyl Ether 34.5 --- (0.10) (307) (358) (27900) (4650)

a) b)

0.8 )

-1 2.7 cm 4

2.5 0.6 Energy (x 10

40 50 60 40 50 60 E (30) / kcal mol-1 T Figure 2.5. Comparing the (a) energy change of emission or (b) Stokes shift change for 2-AP (green squares from Nordlund19 and blue circles from Wierzchowski20) and furanyl dT analogue 1 (red squares) as the polarity of the surrounding environment increases. 60

2.7 – Fluorescence quenching

A quenching effect is typically associated with the incorporation of a fluorescent nucleoside into an oligonucleotide, except in the case of tC (see chapter 1). A major contributing factor to the quenching effect is the neighboring bases stacked above and below the modified nucleoside. In order to probe the quenching effect the natural nucleobases have on the emission spectra of the furanyl dT analogue 1, quenching experiments with all four natural nucleobases were performed. The furanyl dT analogue 1 is most noticeably quenched by dCMP, dGMP and dT, while titration of dAMP shows a most interesting increase in emission intensity (figure 2.6a). To gain perspective, a common fluorescent nucleoside, 2AP, was also subjected to the same experimental conditions. 2AP, whose quantum efficiency is known to be quenched up to 100-fold upon incorporation into an oligonucleotide,10,22 is drastically quenched in the presence of all four nucleobases, with dAMP showing the least effect (figure

2.6b). Comparing these results, 2AP is quenched to a greater degree than the furanyl dT analogue 1, suggesting that nucleobase quenching effects will be far less detrimental to the overall quantum efficiency upon incorporation into an oligonucleotide.

2 30 a) b)

20 /F 0 F

1 10

0 0 0.1 0.2 0.1 0.2

[Q] / M

Figure 2.6. Stern-Volmer plots of (a) furanyl dT analogue 1 (filled) and (b) 2-deoxy-2- aminopurine (open) along with furanyl dT analogue 1 (filled) where Q = dCMP (red circles), dGMP (blue squares), dAMP (green diamonds) and dT (orange triangles). Fluorophores were measured at 1.0 10-5 M in water.

2.8 – Furanyl cytidine, adenosine, and guanosine analogues

The furanyl dT analogue 1 displays the most desirable photophysical properties of the four

thymidine analogues in terms of emission wavelength, quantum efficiency and sensitivity to its

microenvironment. It has therefore become of interest to probe the photophysical changes

that result from conjugating a furanyl moiety onto cytidine as well as its effect on the purine

nucleus (figure 2.7). 62

O NH2

O NH O N

N O N O HO HO O O

HO 17HO

NH2 O N N N NH

O N O N HO N HO N NH2 O O

HO OH HO OH 8 9 Figure 2.7. Furanyl decorated nucleoside analogues where a furanyl moiety is attached to either the 5 position of the pyrimidine core (1 and 7) or the 8 position of the purine core (8 and 9).

2.9 – Synthesis of furanyl cytidine analogue

Conversion of the furanyl 2-deoxyuridine analogue 1 to its corresponding 2-deoxycytidine

analogue 7 can be accomplished by activation of the 4 position followed by a displacement

reaction with aqueous ammonia. To facilitate this conversion, the 3 and 5 hydroxyl groups

were protected as acetate esters. This type of protection was chosen because the use of

strongly basic conditions during the aminolysis reaction would also remove the acetate

protecting groups, generating the desired free nucleoside 7. Acetate protection was achieved

in quantitative yield by treating 1 with acetic anhydride under basic conditions. The protected

nucleoside 10 was then activated using 2,4,6-triisopropylbenzenesulfonyl chloride in the

presence of 4-dimethylaminopyridine and triethylamine. Treatment of the activated nucleoside

with aqueous ammonia and in situ acetate deprotection furnished the desired 2-deoxycytidine

nucleoside 7 in 95 % yield (scheme 2.3). 63

O O NH2 I NH O NH O N

N O N O N O HO RO HO O acO O

HO RO HO 5 1 R=H 7 b 10 R=Ac

Scheme 2.3. (a) 2-(tributylstannyl)furan, PdCl2(PPh3)2, dioxane, 94%; (b) acetic anhydride, pyridine, 99%; (c) (i) 2,4,6-triisopropylbenzenesulfonyl chloride, triethylamine, 4- dimethylaminopyridine, acetonitrile; (ii) ammonium hydroxide, 95%.

2.10 – Photophysical properties of furanyl cytidine analogue

The photophysical characteristics of the 2-deoxycytidine analogue 7 were compared to the

corresponding precursor, furanyl dT analogue 1. Both pyrimidine analogues have distinct

absorption bands (~315 and ~310 nm for 1 and 7, respectively) in addition to the natural

pyrimidine absorption band at ~260 nm. While nucleoside 1 shows two defined absorption

bands of approximately equal intensity at ~260 and ~315 nm, the low energy absorption

bands in the cytidine analogue 7 appear as well-defined shoulders in various solvents. The

absorption of each nucleoside, 1 and 7, is minimally sensitive to changes in solvent polarity

(figure 2.8). The emission spectra, all in the visible range, display much higher sensitivity to

changes in the microenvironment. Similar to nucleoside 1, the cytidine analogue 7, shows

bathochromic (12 nm) and hyperchromic (4.4 fold) shifts upon increasing solvent polarity.

Note, however, the quantum efficiency of emission for the furanyl dC analogue 7, is

approximately 3-fold lower than that of the furanyl dT analogue 1 (table 2.4 on page 68). 64

–1 8

cm 10 –1 M 3 4 5 PL Intensity PL / × 10 ε

0 0 300 400 500 300 400 500 λ / nm

Figure 2.8. Absorption (5.0 10-5 M) and steady state emission (1.0 10-5 M) spectra of 1 (a) and 7 (b) in water (black), methanol (red), acetonitrile (green), dichloromethane (dark blue) and ethyl acetate (light blue).

2.11 – Synthesis of furanyl adenosine and guanosine analogues

The furanyl-containing purine analogues, 8 and 9, were prepared in their ribose form (due

to commercial availability) according to previously published synthetic procedures (Scheme

2.4).23 Temporary protection of the free hydroxyls as their TMS derivatives, followed by a

Stille coupling reaction with 2-(tributylstannyl)furan and standard deprotection using potassium

carbonate in methanol afforded the purine analogues 8 and 9 (Scheme 2.4). Despite the

relatively low and unoptimized yields of this synthetic path, sufficient quantities for

photophysical studies were easily obtained. 65

NH2 NH2 N N N N Br N O N HO N HO N O a O

HO OH HO OH 11 8 O O N N NH NH Br N O N HO N NH2 HO N NH2 O a O

HO OH HO OH 12 9 Scheme 2.4. (a) (i) 1,1,1,3,3,3-hexamethyldisilazane, ammonium sulfate, pyridine (ii) 2-

(tributylstannyl)furan, PdCl2(PPh3)2, THF (iii) K2CO3, methanol, 8: 9%; 9: 10%.

2.12 – Photophysical properties of furanyl adenosine and guanosine

analogues

Unlike the pyrimidine analogues, the conjugated purines (8 and 9) lack a separate and

distinct absorption band, but instead display one major red shifted transition around ~300 nm.

These absorptions bands are largely unaffected by changes in solvent polarity, except in the

case of guanosine analogue 9, where the less polar solvents (dichloromethane and ethyl

acetate) show a slightly lower intensity and red shift in comparison to the other more polar

solvents (figure 2.9). Intriguingly, the furanyl-conjugated purines are highly emissive (F =

0.57 and 0.69 for 8 and 9, respectively). In contrast to the pyrimidine analogues that typically

emit in the visible range, the emission maxima of nucleosides 8 and 9 are centered around

375 nm with an average Stokes shift of 70 and 80 nm, respectively. Somewhat disappointing,

however, is the minimal bathochromic and hyperchromic effects the purine derivatives display

upon increasing solvent polarity, with the G derivative 9 being more sensitive than the A

analogue 8. 66

–1 8 15 cm –1

M 10 3 4

5 Intensity PL / × 10 ε

0 0 300350 400 450 300 350 400 450 λ / nm Figure 2.9. Absorption (5.0 10-5 M) and steady state emission (1.0 10-5 M) spectra of 8 (a) and 9 (b) in water (black), methanol (red), acetonitrile (green), dichloromethane (dark blue) and ethyl acetate (light blue).

2.13 – Effect of temperature on the emission of furanyl decorated

nucleosides

The furanyl-heterocycle being the emitting species of nucleosides 1, 7, 8 and 9 whereby

increased temperatures would increase the free rotation of the furanyl ring suggests that their

emission is likely to be temperature dependent. Indeed, all nucleosides display a significant

hypochromic effect when their spectra are taken as function of increasing temperatures (figure

2.10). The emission bands of nucleosides 1, 7 and 9 display significant susceptibility to

increasing temperatures, while the A analogue 8 is affected to a lesser degree. Importantly,

this temperature dependent behavior is completely reversible and therefore does not reflect

any heat induced chemical transformation. The sensitivity of the emission quantum yield to

temperature is thought to arise from the relatively low barrier for rotation around the furanyl–

pyrimidine/purine single bond that conjugates the two heterocycles. Thermal population of

non-radiative decay pathways hence lowers the emission quantum yield at elevated

temperatures.

1 aa*bb*

0 400 450 500 550 400 450 500 550 400 450 500 550 400 450 500 550 1 cc*dd* Relative PL Intensity Relative PL

0 350 400 450 350 400 450 350 400 450 350 400 450 λ / nm

Figure 2.10. Steady-state emission spectra of 1 (a) 1.0 10-5 M, 7 (b) 1.0 10-5 M, 8 (c) 1.0 10-6 M and 9 (d) 1.0 10-6 M at various temperatures (°C): 25 (black), 35 (dark blue), 45 (red), 55 (orange), 65 (light blue), 75 (green). Unstared letters represent the increase of temperature and the stared letters represent the decrease in temperature.

2.14 – Photophysical evaluation of furanyl decorated nucleosides

Evaluation of all four nucleosides is necessary for the selection of the most responsive probe. As discussed above, absorption bands of all four nucleosides remain largely unaffected by changes in solvent polarity (table 2.4), whereas changes in the emission spectra with increasing polarity are seen for all four nucleosides. A graphical comparison of the bathochromic and hyperchromic shifts that take place when the polarity of the microenvironment is increased is given in figure 2.11. Plotting the change in emission area

(relative to most apolar solvent) versus ET(30) values illustrates that all four nucleosides undergo a dramatic hyperchromic shift, as denoted by the positive slope. Nucleosides 1 and 7

are the most responsive (hyperchromic), followed closely by nucleoside 8 and then 9 (figure

2.11a) as denoted by the steepness of the slope and y-axis intercept.

Table 2.4. Photophysical Data of Furanyl Nucleosides 1, 2, 8 and 9.

max cmpd EtOAc max H2O em EtOAc em H2O F H O (nm) (nm) (nm) (nm) H2O/EtOAc 2 1 314 316 397 431 4.4 0.03 7 306 310 431 443 4.4 0.01 8 302 304 372 374 1.6 0.69 9 294 294 370 378 2.5 0.57

Graphing the emission energy versus ET(30) values clearly shows that nucleoside 1 experiences the largest bathochromic shift of all four nucleosides upon increasing solvent polarity (figure 2.11b). In contrast, nucleosides 7, 8 and 9 all show very shallow dependency on solvent polarity. It is also apparent that the furanyl-conjugated purines exhibit relatively insensitive emission at significantly higher energies compared to the conjugated pyrimidines.

Photophysically, nucleoside 1 is the most responsive nucleoside of the entire family and was therefore selected for site-specific incorporation into oligonucleotides.

ab 2.7

0.8 ) -1 cm 4

2.5

0.4 Energy (x 10 Change in Emission Area Change in Emission

2.3

0 40 50 60 40 50 60

ET(30) Figure 2.11. (a) Relationship between change in emission area (relative to the most apolar solvent) and microenvironment polarity. (b) Relationship between emission energy and microenvironment polarity. ET(30) values were experimentally determined for each solvent using Reichardts dye. For both (a) and (b) 1 (black circles), 7 (red squares), 8 (green diamonds) and 9 (blue triangles).

2.15 – Phosphoramidite synthesis of furanyl thymidine nucleoside

Standard solid-phase oligonucleotide synthesis requires 5-protected and 3-activated building blocks. Preparation of the required phosphoramidite 14, was achieved through standard 5-OH protection of nucleoside 1 with 4,4-dimethoxytrityl chloride in the presence of triethylamine and pyridine (scheme 2.5). The 3-OH was then phosphitylated under standard conditions, resulting in the required modified building block for DNA oligonucleotide synthesis.

O O O

O NH O NH O NH

N O N O N O HO DMTO DMTO O abO O

HO 113HO O i P N( Pr)2 O 14 NC Scheme 2.5. (a) 4,4-dimethoxytrityl chloride, pyridine, triethylamine, 71%; (b)

(iPr2N)2POCH2CH2CN, 1H-tetrazole, acetonitrile, 65%.

2.16 – Site-specific incorporation of furanyl thymidine nucleoside

Phosphoramidite 14 was site specifically incorporated into a non-self complementary singly modified DNA oligonucleotide 15 (figure 2.12) with an optimized coupling efficiency of 99%.

This was accomplished by synthesizing the natural nucleotide via standard trityl-off synthesis

(figure 2.13) and then the modified nucleoside was manually coupled to the growing strand.

Upon successful coupling, the modified nucleoside was oxidized and capped on the synthesizer followed by completion of the DNA oligonucleotide again via trityl-off synthesis.

15 5 GCG – ATG – 1GT – AGC – G 3 16 5 CGC – TAC – ACA – TCG – C 3 17 5 CGC – TAC – GCA – TCG – C 3 18 5 CGC – TAC – CCA – TCG – C 3 19 5 CGC – TAC – TCA – TCG – C 3 20 5 CGC – TAC – CAT – CGC 3 21 5 CGC – TAC – YCA – TCG – C 3 22 5 TGA – CGC – TAC – ACA – TCG – CAG – T 3 23 5 TGA – CGC – TAC – CCA – TCG – CAG – T 3 24 5 GCG – ATG – TGT – AGC – G 3 25 5 GCG – ATG – dUGT – AGC – G 3

Figure 2.12. Sequences of oligonucleotides: 15 modified, 16–23 complement (Y=abasic site mimic) and 24, 25 control sequences

O O B1 O O O O Step 5 O O B2 O Capping

O O O Bn O O P O Step 4 O End Cycle O NC B1 O Oxidation O P O O O B2 O O O O O O P O O B2 O Step 6 O B1 Cleavage and O O O Deprotection O P HO O NC B1 O

O Step 1' O Cycle Repeat O

Step 3 Coupling Step 1 HO B1 Detritylation O

Step 2 O O Start Cycle Phosphoramidite O Activation O

O O B1 O O B2 O O O O O O O P O N

Figure 2.13. Key steps in solid phase DNA phosphoramidite synthesis cycle. Step 1: removal of the 5-DMT protecting group using dichloroacetic acid; Step 2: activation of the

phosphoramidite using 1H-tetrazole; Step 3: coupling of the 5-hydroxyl on B1 to the activated phosphoramidite B2; Step 4: oxidation of the trivalent phosphorous to the pentavalent phosphorus using iodine in a tetrahydrofuran/pyridine/water solution; Step 5: all unreacted

nucleotides (B1) are capped using acetic anhydride; Step 6: cleavage of the oligonucleotide from the CPG resin and deprotection of protected nucleoside functional groups using concentrated ammonium hydroxide; Step 1: repetition of the cycle continues until the desired DNA length is reached.

2.17 – Characterizing the furanyl modified oligonucleotide

Characterization of the modified oligonucleotide 15 was accomplished by MALDI-TOF mass spectrometry as well as HPLC digest experiments. These MALDI-TOF experiments provide vital information that typical HPLC digest experiments cannot, i.e. the exact incorporation position of the modified nucleoside. The masses of purified oligonucleotide 15 and all other commercially available oligonucleotides 16–25 were observed to be in good agreement with the calculated masses (Table 2.5). Enzymatic digestion experiments utilized two endonucleases, bovine spleen (bsp) and snake venom phosphodiesterases (svp) to cleave the modified oligonucleotide 15 in a systematic fashion that could be monitored by

MALDI-TOF mass spectrometry. Bsp, cleaving from the 5 to 3 direction and svp cleaving in the opposite direction (3 to 5) resulted in overlapping spectra that clearly confirmed the sequence of the modified oligonucleotide (figure 2.14).

Table 2.5. MALDI-TOF of oligonucleotides 15–25.

Oligo Calculated Mass Found Mass

15 4081.59 4081.98 16 3878.70 3880.07 17 3894.69 3894.31 18 3854.69 3855.57 19 3869.68 3870.32 20 3565.64 3565.23 21 3744.65 3745.49 22 5771.01 5770.35 23 5747.00 5746.30 24 4029.67 4029.71 25 4014.70 4015.61

100 100 dT dA dT 305.5 314.6 305.2 dG 1 mono dC phosphonate dA 330.2 314.3 dG 290.2 330.3 80 80 dG 330.6

60 dG 60 dG 330.2 330.5 dG dG dC 330.7 330.0 290.7 dT 304.4

40 40 % Intensity

20 20

0 0 1684 2830 3976 830 2331 3833 Mass (m/z) Mass (m/z) BSP Digestion (5’ to 3’) SVP Digestion (3’ to 5’)

Figure 2.14. MALDI-monitored digestion of oligonucleotide 15. Bovine spleen phosphodiesterase (bsp), cleaving in the 5 to 3 direction, identifies the first six bases in oligonucleotide 15, while snake venom phosphodiesterase (svp), cleaving in the 3 to 5 direction, identifies nine nucleotides in sequence, including a two nucleotide overlap with the bsp digestion and identification of incorporated nucleotide 1 (see experimental).

All four 2-deoxynucleosides and the modified furanyl dT analogue 1 were separated via

HPLC to allow for identification of the digested nucleosides. Oligonucleotide 15 was digested

and separated by HPLC, resulting in identification of the furanyl dT analogue at ~14 min. The

correct composition was verified by integration of the peak area (Table 2.6). These

experiments (MALDI-TOF MS and HPLC) clearly identify that nucleoside 1 has not undergone

any chemical modification that may have occurred during the numerous coupling, capping and

oxidation steps involved in solid-phase DNA synthesis.

a) dG 0.4 dA dT dT 0.3 Furan

0.2 dC

0.1

1.0 b)

Absorption / a.u. 0.8

0.6

0.4

0.2

0 051015 Time / min. Figure 2.15. HPLC trace of (a) standard 2-dexoynucleosides, the furanyl dT 1 analogue and (b) digest of modified oligonucleotide 15 monitored by UV absorption at 260 nm (black) and 316 nm (red).

Table 2.6. Determination of modified oligonucleotide 15 composition.

Nucleoside Normalized Area at 260 nm Area dC 1177 7300 2.0 dG 5413 11700 5.7 dT 1409 8800 2.0 dA 2170 15400 1.7 1 801 13000 0.8

2.18 – Stability of modified duplexes

One of the design principles outlined in section 2.1, to maintain the highest structural similarity to the natural nucleobases and ensure the formation of WC-like base pairs, was experimentally investigated through thermal denaturation experiments of both unmodified

(24•16–24•23 and 25•16–25•23) and modified (15•16–15•23) oligonucleotide duplexes. In

order to gain insight into the role the furan ring may play concerning duplex stability, a

comparison to 2-deoxythymidine and 2-deoxyuridine containing duplexes was made. When

the furanyl modified oligo 15 was paired with its perfect complement 16, a stable duplex

(15•16) with no detectable shape (figure 2.16) or Tm (Table 2.7) difference than the dT 24•16

or dU 25•16 control duplexes was seen. Upon identifying that nucleoside 1 does not disturb

duplex formation other mismatch cases were investigated. Furanyl containing oligo 15 was

paired with complementary strands that contained a single mismatch (dG, dC and dT) at the

base pair site of modified dT analogue 1. All mismatch cases resulted in a lower melting

temperature (~8–12 °C) than the corresponding perfect duplex. Specifically, pairing furanyl dT

analogue with mismatches dC and dT results in the same destabilizing effect seen with the

control duplexes containing dT and dU. However, when furanyl dT analogue is paired with dG

mismatch, it results in a greater destabilizing effect than dT or dU containing oligos that are

3.1 and 1.0 °C more stable respectively. This however is not strictly a steric issue, for the

control dT•dG (24•18) is the most stable of the three, yet has similar steric bulk as the furanyl- 76

dT•dG duplex (15•18) while both have increased steric specifically at the 5 position over the dU•dG duplex (25•17). Investigation into the absorption and emission spectra of these duplexes is of interest.

1 ab

0.5 Normalized Absorption

0 30 50 70 30 50 70 Temperature / °C

Figure 2.16. Thermal denaturation of furanyl modified oligonucleotide 15 (open) and (a) control dT oligonucleotide 24 (closed) or (b) control dU oligonucleotide 25 (closed) paired with its perfect complement dA 16 (black), mismatch dG 17 (blue), mismatch dC 18 (green) and mismatch dT 19 (red).

fu Table 2.7. Tm values of modified and control duplexes where dT = 1.

Tm from 24 / °C Tm from 25 / °C Duplex Central Average Tm / Base Pair °C (duplex – corresponding dT (duplex – corresponding dU duplex) duplex) 15•16 dTfu•dA 55.8 0.2 0.1 +0.0 15•17 dTfu•dG 45.0 0.3 3.1 1.0 15•18 dTfu•dC 43.8 0.2 +0.9 +0.4 15•19 dTfu•dT 46.3 0.4 +0.0 +0.6 24•16 dT•dA 56.0 0.3 --- +0.2 24•17 dT•dG 48.1 0.4 --- +2.1 24•18 dT•dC 42.9 0.3 --- 0.5 24•19 dT•dT 46.3 0.3 --- +0.5 25•16 dU•dA 55.8 0.8 0.2 --- 25•17 dU•dG 46.0 0.7 2.1 --- 25•18 dU•dC 43.4 0.7 +0.5 --- 25•19 dU•dT 45.7 0.6 0.5 ---

2.19 – Absorption spectra of furanyl dT containing oligonucleotides

Absorption spectra of modified duplex 15•16 in comparison to the unmodified duplex 16•17

reveals normal absorption bands at 260 nm, but more importantly show a small intensity

difference in the 315 nm range (figure 2.17). Subtraction of these two absorption spectra, their

nucleoside composition being identical except for the furanyl dT analogue 1 in duplex 15•16,

clearly reveals the second absorption band (~320 nm) of nucleoside 1 (figure 2.17). The max

of absorption for furanyl dT analogue 1 is relatively unchanged upon incorporation into an

oligonucleotide.

0.4 0.06

0.2

Absorption / a.u. 0.02 Absorption / a.u.

0 0 240 320 λ / nm

Figure 2.17. Absorption spectra of oligonucleotide duplexes 15•16 (blue) and 16•17 (black) at 1.0 10-6 M and the subtraction of oligonucleotide duplexes, 15•1616•17 (red).

2.20 – Steady state emission spectra of furanyl dT containing

oligonucleotides

Importantly, furanyl dT analogue 1 retains its wavelength of emission upon incorporation into an oligonucleotide. Upon excitation at 320 nm, emission is observed at ~430 nm, as 78

previously seen for furanyl dT analogue 1. The emission curve of the oligonucleotide is sharper than the one observed for the free nucleoside in solution and lacks the fine structure observed for 1 (comparison of figure 2.18 to figure 2.4).

The ability of the fluorescent dT probe 1 to interrogate the base opposite it in a duplex was investigated. We have thus measured the furanyl dT analogue in the single strand case, when it is paired with its perfect complement, thus a duplex case, as well as the three other possible single mismatch cases (dG, dC and dT). The furanyl thymidine analogue is most emissive in the single strand case and weakest when paired with its perfect complement. These results parallel those seen in the solvent polarity studies of the free nucleoside (section 2.5), where the furanyl dT analogue is most emissive in a polar environment and least emissive in a non- polar environment. In the single strand case, where the furanyl dT is the most exposed to the aqueous environment, it shows the highest intensity and in a perfect complement case when the modified nucleoside is protected from the aqueous environment via base staking in the

DNA double helix, it shows the lowest emission intensity. The mismatch cases all result in some intermediary emission intensity, but closer to that of the perfect duplex than to the single strand intensity. This suggests that upon mismatch pairing, the furanyl dT is pushed more outside the duplex, exposing it to a more polar environment and thus resulting in an increased emission. The inability for the mismatch cases to return the emission intensity back to the single strand case provides evidence that the furanyl dT analogue in a mismatch case, while it is more exposed to an aqueous environment, it is still somewhat protected. The furanyl dT does not have the ability to distinguish between each mismatch case individually, but can distinguish between a mismatch, perfect complement and single strand cases. There is also

no direct correlation that can be drawn between the intensity seen in figure 2.19 and the Tm of the same construct noted in table 2.7. The insights gained here led us to investigate the furanyl dT oligonucleotide base paired with 4 other complementary constructs.

1 Relative PL Intensity Relative PL

0 400 450 500 550 λ / nm Figure 2.18. Steady state emission spectra of furanyl dT single strand (15 red) and it hybridized to its perfect complement (15•16 blue), mismatch dG (15•17 black) mismatch dC (15•18 green) and mismatch dT (15•19 orange).

2.21 – Investigation of additional duplexes

In exploring the reporting ability of the furanyl dT analogue further we have investigated two new scenarios, where the furanyl dT has no pairing partner and where hybridization ability is investigated. Two constructs were imagined where the furanyl dT has no pairing partner: (1) when the furanyl dT oligonucleotide is paired with a 12 bp sequence that lacks the corresponding dA (24) across from the furanyl dT position. (2) when the furanyl dT oligonucleotide is paired with a 13 bp sequence in which its corresponding base is absent, hence an abasic site (25).

The interest in the detection of abasic sites stems from their biological presence and function. Nucleic acids undergo a variety of chemical transformations, including depurination and depyrimidination. These abasic sites can be generated either spontaneously or via enzymatic base excision of damaged nucleosides. Several methods have been developed for detecting the presence of these cytotoxic DNA lesions. Most techniques require irreversible modifications of isolated DNA,24-59 whereas others use non-natural nucleobase analogues60-71

and only a few use 2AP to investigate the stacking ability of a base opposite an abasic site.72- 80

74 Fluorescent nucleobase analogues that positively sense the presence of abasic sites could provide useful nondestructive tools for the detection of such important DNA defects.

Hybridization of the furanyl dT containing oligonucleotide 15 was investigated with two

19mer complementary oligonucleotides (a perfect complement 22 and a single mismatch 23) were paired with the furanyl dT containing oligonucleotide 15, resulting in hanging ends. The additional bases (3 on each end) would provide extra stability ensuring a greater degree of hybridization. Besides hanging ends, increased hybridization can also be achieved via increased ionic strength of the solution. Both of these approaches have been explored.

2.22 – Thermal denaturation of additional duplexes

Investigation into the thermal stability of the furanyl modified oligonucleotide 15 in comparison to dT/dU control oligonucleotides paired with some non-natural complements revealed some interesting results. There is a noticeable enhancement of stability (2.9–4.1°C) provided by the furanyl dT when it has no direct base to pair with or when it is paired across from an abasic site (figure 2.19 and Table 2.8). Specifically, the furanyl containing modified duplex 15•20 is 3.6 °C and 2.9 °C more stable than the corresponding control dT/dU duplexes

24•20 and 25•20 respectively. It appears that the 5 position of uridine when extended with steric bulk containing - stacking ability offers greater stability than steric bulk without - stacking ability (i.e. dT) or the lack of any modification (i.e. dU). A similar trend is seen when the complement oligonucleotide contains an abasic site, where the furanyl modified duplex

15•21 is the most stable, followed by the uridine 24•21 and thymidine 25•21 containing duplexes respectively. The furanyl thymidine analogue must provide the extra 4.1 or 3.0 °C of stability through - stacking interactions, therefore it is thought to be in a syn conformation, allowing for both the furanyl and thymidine moieties to interact with the bases above and below (figure 2.22). It is this conformational change that will be mentioned again in the 81

description of the fluorescence readings of these duplexes. Increased stability of ~3.0 °C was seen by the 13/19 mer duplexes (table 6) in comparison to the 13/13mer duplexes (table 2.8).

As seen with the 13/13mer duplexes, the modified furanyl does not perturb the stability of the duplex over that of uridine or thymidine. The thermal stability of the 12mer and abasic containing duplexes suggests that the furanyl moiety is behaving in a different manner than

the uridine or thymidine controls while additional nucleotides increase the overall Tm, no

change is seen when compared to their control duplexes. It is with these results in mind that

the photophysical properties of the above mentioned duplexes were investigated.

1 ab

0.5 Normalized Absorption

0 30 50 70 30 50 70 Temperature / °C

Figure 2.19. Thermal denaturation of furanyl modified oligonucleotide 15 (open) and (a) control dT oligonucleotide 24 (closed) or (b) control dU oligonucleotide 25 (closed) paired with a short 12mer oligonucleotide 20 (black), an abasic site containing oligonucleotide 21 (blue), a perfect complement dA 19mer 22 (green) and a mismatch dC 19mer 23 (red).

fu Table 2.8. Tm values of modified and control duplexes where dT = 1, __ = no pairing nucleoside present, and AP = stable abasic site mimic (THF residue).

Tm from 24 / °C Tm from 25 / °C Central Average Tm / Duplex (duplex – corresponding dT (duplex – corresponding dU Base Pair °C duplex) duplex) 15•20 dTfu•__ 47.6 0.8 +3.6 +2.9 15•21 dTfu•AP 39.0 0.8 +4.1 +3.0 15•22 dTfu•dA 58.9 0.2 0.2 0.5 15•23 dTfu•dC 46.9 0.4 +1.1 +0.4 24•20 dT•__ 44.0 0.4 --- 0.8 24•21 dT•AP 35.0 0.4 --- 1.1 24•22 dT•dA 59.1 0.3 --- 0.3 24•23 dT•dC 45.8 0.1 --- 0.7 25•20 dU•__ 44.7 0.7 +0.8 --- 25•21 dU•AP 36.1 0.8 +1.1 --- 25•22 dU•dA 59.4 0.3 +0.3 --- 25•23 dU•dC 46.5 0.7 +0.7 ---

2.23 – Photophysical properties of additional duplexes

Investigation into the UV-Vis spectra reveals similar profiles to those seen before with the

furanyl dT containing duplexes, where there is an intense band at ~260 nm as well as the

more red shifted absorption band at ~320 nm. The 320 nm band is very weak in comparison

to the 260 nm band. The 320 nm band does not greatly change max or in the various

constructs. It is however, the emission properties of this fluorescent nucleobase that results in

markedly different emission intensities when paired with the 12mer 20 and abasic site 21

complements (figure 2.20). When the furanyl dT analogue is paired with the 12mer

complement (15•20), the probe reveals an emission intensity about half that of the single

strand, suggesting that the probe is partially exposed to the polar environment. While the

duplex with containing the abasic site (15•21) results in an emission intensity that is

remarkably higher than that of the single strand. There was very little difference in emission

intensity between the hanging end duplexes, 15•22 and 15•23, and their corresponding blunt

end duplexes 15•16 and 15•18 (data not shown). To confirm the emission spectra were a

result of duplexed DNA and not small amounts of single strand DNA, these three constructs 83

were tested under higher salt conditions (1 M). The same relative relationship is seen upon an increase in salt concentration, depicting the emission are results from the duplexes and not from denatured strands. There is an increase in the intensity difference between the furanyl dT ss (15) and when it is paired across from an abasic site and a sharpening of the signal,

suggesting a flattening of the chromophore. Thermal denaturation also shows a greater

increase in Tm for the abasic case than for the 12mer system, suggesting the furanyl dT analogue increases its stacking ability with increased salt concentration (Table 2.9).

1 a) b) Relative PL Intensity Relative PL

0 400 450 500 400 450 500 λ / nm

Figure 2.20. Steady state emission spectra of furanyl dT single strand (15 red), it hybridized to the 12mer strand (15•20 blue) and to the abasic site containing strand (15•21 green) at (a) 100 mM NaCl, 10 mM phosphate buffer, pH = 7.0 and (b) 1 M NaCl, 100 mM phosphate buffer, pH = 7.0.

fu Table 2.9. Tm values of modified and control duplexes where dT = 1, __ = no pairing nucleoside present, and AP = stable abasic site mimic (THF residue).

Tm from 24 / °C Duplex Central [NaCl] M Average Tm / Base Pair °C (duplex – corresponding dT duplex) 15•20 dTfu•__ 0.1 47.6 0.8 +2.9 15•21 dTfu•AP 0.1 39.0 0.8 +3.0 24•20 dT•__ 0.1 44.0 0.4 0.8 24•21 dT•AP 0.1 35.0 0.4 1.1 15•20 dTfu•__ 1.0 54 --- 15•21 dTfu•AP 1.0 48 --- 84

The furanyl dT analogue 1 can be used to distinguish between perfect complement and abasic site pairing, by a dramatic increase in fluorescence intensity. When oligo 15 is hybridized to the tetrahydrofuran-containing oligo 21 a duplex containing an abasic site is

formed. Remarkably, the emission of duplex 15•21 is enhanced 7-fold when compared to the

duplex 15•16, formed upon hybridization to the perfect complement (figure 2.11).

1 Relative PL Intensity Relative PL

0 400 450 500 λ / nm

Figure 2.21. Steady state emission spectra of furanyl dT single strand (15 red) and it hybridized to the perfect complement strand (15•16 blue) and to the abasic site-containing strand (15•21 green).

An unpaired base opposite an abasic site can be intrahelical or extrahelical depending on the sequence context. Our current working hypothesis is that the furanyl dT analogue is intrahelical, assuming a syn conformation (figure 2.22). This stacked conformation protects the hydrophobic furanyl moiety, while projecting the hydrogen bonding face toward the major groove. Support is offered by the following: (a) duplex 15•21 is more stable than the control

duplex 24•21 or 25•21 that contains a dT or dU residue opposite the abasic site. The

increased stability of the modified abasic duplex (Tm = +3–4 °C) suggests a favorable

accommodation of the modified nucleobase by the duplex, and (b) the emission band

observed for duplex 15•21 decays sharper (> 500 nm) than when compared to the emission

exhibited by the free nucleoside in solution. This is consistent with flattening of the 85

chromophore that can be associated with the restricted rotation of the conjugated furanyl ring upon inclusion within the DNA duplex. Therefore, the furanyl dT analogue 1, when incorporated into a reporter oligonucleotide, positively signals the presence of a DNA abasic site.

H N O O H N O N O O O O N HN N H N O O O O O N N O O O O O O

1•dA 1•Abasic site

Figure 2.22. The furanyl dT analogue base pairing with dA in a W-C manner and the proposed syn conformation upon recognition of an abasic site.

2.24 – Effects of temperature on the photophysical properties of furanyl

containing duplexes.

The furanyl dT analogue is emissive in polar environments and quenched in apolar

environments, which allows the furanyl dT analogue to clearly detect the difference between

ss and duplexed DNA (figure 2.24). This ability can be used to monitor the melting of a duplex

containing the furanyl dT analogue by fluorescence intensity. The Tm value determined by fluorescence is very similar to the value determined by absorption (figure 2.23). The noticeable decreases in the fluorescence intensity initially at 25–40°C and towards the end of the melting profile at 65–75°C are a result of the inherent temperature sensitivity of the furanyl dT analogue. 86

1 1 Normalized absorption Normalized fluorescence 0 0

30 50 70 Temperature / °C

Figure 2.23. Denaturation of duplex 15•16 monitored by absorption or fluorescence.

Monitoring the denaturation of DNA duplexes via fluorescence can be applied to the ss and all duplex cases (figure 2.24). The melting of the ss and abasic site containing duplex both decrease in emission intensity with no inflection point. This is expected in the single strand case, where the probe is in a very polar environment to begin with, thus upon heating the chromophore cannot become drastically more exposed the surrounding polar environment. In the abasic site containing duplex, the initial fluorescence intensity is higher than the single strand case, thus upon denaturation, exposure to a more polar environment results in a decrease in emission intensity, not an increase as seen in the perfect complement or mismatch cases. All denaturation curves result in the same intensity fluorescence above the melting temperature of the duplex ( 60°C) and all suffer from the inherent temperature dependence of the furanyl analogue itself (figure 2.10). 87

1 Relative PL intensity Relative PL

0 30 40 50 60 70 80 Temperature / °C

Figure 2.24. Denaturation of ss 15 (red triangles) and duplexes 15•16 (blue circles), 15•18 (orange squares) and 15•21 (green diamonds) monitored by fluorescence.

2.25 – Enzymatic incorporation of furanyl dT analogue

To extend the utility of the furanyl dT analogue, the triphosphate form was synthesized in a

one-pot reaction starting from the previously synthesized furanyl dT analogue 1 (scheme 2.6).

The product resulted in relatively low yield, albeit with sufficient quantity to carryout preliminary studies. Initial investigation determined that the Klenow fragment (exo-) accepted the furanyl dTTP analogue 26 in ~80% efficiency in comparison to the natural system (figure 2.26). The

primer/template duplex was designed so that incorporation of a dA was last, whereby using -

32P ATP, only the full-length products would be visualized. Current work is directed toward the

exploration of enzymatically incorporating multiple furanyl dT analogues into one sequence

with the interest of studying the photophysical properties of these resulting oligonucleotides.

O O

O NH O O O O NH HO O P O P O P O N O N O O a O O O O

(Et NH+) HO 3 4 HO 1 26 Scheme 2.6. (a) (i) proton sponge, (MeO)3PO, POCl3, 0 °C (ii) tributylammonium pyrophosphate, Bu3N, 0 °C, 5%.

27 5 CGA – CTC – ACT – ATA – G 3 28 3 GCT – GAG – TGA – TAT – CGC – GGC – AGC – T 5

Figure 2.25. Template and primer sequence for DNA extension experiment.

Figure 2.26. DNA polymerase reactions utilizing primer 27 and template 28. Lane 1: natural dNTPs. Lane 2: dATP, dGTP, dCTP and 26. All lanes contained Klenow fragment (exo-) and -32P ATP.

2.26 – Site-specific incorporation of furanyl dC analogue

The results seen with the furanyl dT analogue inspired the incorporation of the cytidine

analogue. Starting with the previously prepared furanyl dT analogue 1, the 3 and 5 hydroxyl

groups were protected as the TBDMS ethers allowing for easy conversion from the dT face to

the dC face in good yield (80% scheme 2.7). Benzoyl protection of the amine, followed by

quantative deprotection of the silyl groups afforded nucleoside 31. Protection of the 5

hydroxyl as the DMT ester and phosphitylation of the 3 hydroxyl afforded the appropriate

building block for solid phase DNA synthesis.

O O NH2 I NH O NH O N

N O N O N O HO RO TBDMSO O acO O

HO RO TBDMSO 5 1 R=H 30 b 29 R=TBDMS

O O O

HN HN HN

O N O N O N

N O N O N O DMTO DMTO RO O g O f O

O HO RO PN(iPr) 2 33 31 R=TBDMS e O 32 R=H 34 NC Scheme 2.7. (a) 2-(tributylstannyl)furan, PdCl2(PPh3)2, dioxane, 94%; (b) TBDMS-Cl, imidazole, DMF, 96%; (c) (i) 2,4,6-triisopropylbenzenesulfonyl chloride, triethylamine, 4- dimethylaminopyridine, acetonitrile; (ii) ammonium hydroxide, 80%; (d) benzoyl chloride, 4- dimethylaminopyridine, dichloromethane, 96%; (e) triethylamine trihydrogen fluoride, tetrahydrofuran, 99%; (f) dimethoxytrityl chloride, 4-dimethylaminopyridine, triethylamine, pyridine, 66%; 2-cyanoethyl diisopropyl-chlorophosphoramidite, diisopropylethylamine, 1,2- dichloroethane, 70%.

Phosphoramidite 34 was site specifically incorporated into a non-self complementary singly

modified DNA oligonucleotide 35 (figure 2.27) with an optimized coupling efficiency of 97%.

This was accomplished in a similar manner to the furanyl dT analogue, where the natural

nucleotides were synthesized via standard trityl-off synthesis followed by manually coupling of

the modified nucleoside into the growing strand. Upon successful coupling, the modified

nucleoside was oxidized and capped on the synthesizer, followed by completion of the DNA

oligonucleotide again via trityl-off synthesis (figure 2.13).

35 5 GCG – ATG – 7GT – AGC – G 3

Figure 2.27. Sequence of modified oligonucleotide 35.

Characterization of the modified oligonucleotide 35 was accomplished by MALDI-TOF mass spectrometry and HPLC digest experiments. MALDI-TOF MS of oligonucleotide 35, resulted in a mass of 4079.55, similar to the calculated mass of 4080.67. However, MALDI-

TOF MS cannot distinguish between the dC analogue and the dT analogue, being only 1 mass unit different. Therefore an HPLC digest was preformed to identify that the furanyl dC moiety was intact after incorporation, deprotection and purification. All four 2-deoxy-nucleosides and the modified dT and dC furanyl analogues (1 and 7 respectively) were separated via HPLC to

allow for identification of the digested nucleosides. Oligonucleotide 35 was digested and separated by HPLC, resulting in identification of the dC analogue at ~12 min. The correct composition was verified by integration of the peak area (Table 2.10). Photophysical evaluation of modified oligonucleotide 35 is presently being investigated.

Table 2.10. Determination of modified oligonucleotide 35 composition.

Nucleoside Normalized Area at 260 nm Area dC 792 7300 2.0 dG 3762 11700 5.9 dT 937 8800 2.0 dA 1494 15400 1.8 7 693 9600 1.3

0.5 a) dG 0.4 dA dT dC dT Furan Furan 0.3

0.2 dC

0.1

0 0.7 b)

0.6

Absorption / a.u.

0.5

0.4

0.3

0.2

0.1

0 0 5 10 15 Time / min.

Figure 2.28. HPLC trace of (a) standard 2-dexoynucleosides, furanyl dT 1 analogue, furanyl dC 7 analogue and (b) digest of modified oligonucleotide 35 monitored by UV absorption at 260 nm (black) and 304 nm (blue).

2.27 – Improving the photophysical properties of furanyl dT analogue

The mechanism by which the furanyl dT analogue selectively detects the presence of abasic sites most likely involves the free rotation of the furanyl moiety. Since rigidifying conjugated moieties, such as furanyl dT moiety, can often increase quantum efficiency we have decided to investigate locking the furanyl moiety in a planar position to the uridine core.

The ability to attach either the 2- or 3-furanyl to the 5 position of uridine allows for the construction of three separate locked furanyl dT analogues as depicted in figure 2.29.

Cyclizing the furanyl moiety onto the C-4 carbonyl of the uridine core would result in a non W-

C hydrogen bonding face, therefore the C-4 carbonyl is first converted to the amine, resulting in a cytosine hydrogen bonding face and allowing for intermolecular amination to close the ring. 93

O I NH

N O NH2 NH2 R O O N N

N O N O R R

Br Br NH2 NH2 NH2 O O O N N N Br N O N O N O R R R Br O NH2

N O R O O NH NH NH O N N N

N O N O N O R R R 36 37 38

Figure 2.29. Retrosynthetic analysis of 3 cyclized versions (36, 37 and 38) of the furanyl dT analogue where R = 2-deoxyribose.

Synthesis was first investigated with the furanyl dT analogue 1, because this allowed for

large amounts of material to be rapidly acquired. However, low yields (24%) during

bromination of the furanyl ring resulted in the use of the thienyl dT analogue 2, whose

bromination yield was better (73%) (scheme 2.8). It should be noted that most of the steps

completed with the thienyl analogue can and have been completed with the furanyl analogue

as well, but for a clear presentation, only the thienyl will be presented completely. Removal of 94

the more reactive bromine (5 position) in acceptable yield, can be accomplished either before or after cyclization. Conversion of the dT analogue (41) to the dC analogue (43) proved to be problematic with the large 2,4,6-triisopropylbenzenesulfonamide intermediate and the proximity of the bromine moiety, while a smaller activating group (1,2,4-triazole) resulted in a much higher and acceptable yield. The free amine can be protected, thus allowing cyclization to take place from either the amine or amide functionality. A few amination procedures were unsuccessful and continuation of this work is presently ongoing.

Br Br O O O Br X NH X NH O NH

N O N O N O RO AcO c AcO O b O O

RO AcO AcO 1 R=H, X=O 40 X=O 42 2 41 a R=H, X=S X=S 10 R=Ac, X=O 39 R=Ac, X=S d

O Br Br HN NH2 Br Br S N S N

N O N O AcO AcO O O e

AcO AcO 44 43

f g

O Br Br N NH S S N N

N O N O AcO AcO O O

AcO AcO 45 46 Scheme 2.8. (a) acetic anhydride, pyridine, 10: 99%, 39: 97%; (b) bromine, carbon tetrachloride, 1,2-dichloroethane, 40: 34%, 41: 73%; (c) 40, zinc dust, acetic anhydride, acetic acid, dimethylformamide, 42%; (d) (i) 41, 2,4,6-triisopropylbenzenesulfonyl chloride, triethylamine, 4-dimethylaminopyridine, acetonitrile; (ii) ammonium hydroxide, 27% or (i) 41, 1,2,4-triazole, phosphorus oxychloride, pyridine; (ii) ammonium hydroxide, 62%; (e) benzoyl chloride, 4-dimethylaminopyridine, dichloromethane, 55%; (f) copper iodide, cesium acetate, dimethylsulfoxide;75 (g) palladium (II) acetate, 1,1-Bis(diphenylphosphino)ferrocene, sodium t- butoxide, toluene.76

2.28. Concluding remarks

Conjugating a five membered ring to the 5 position of dU has proven to be successful in

identifying a novel class of fluorescent nucleosides. The furanyl moiety, proving to be the

modification with the most desired photophysical properties, when conjugated to the 5 position

of the pyrimidines and 8 position of the purines has provided a family of fluorescent

nucleobase analogues. The pyrimidines show emission in the visible range (400–440 nm)

while the purines emit at higher energies (375 nm). Analysis of the solvatochromic behavior of

these fluorophores identifies the conjugated dT derivative as the most promising nucleobase

for uses in biophysical assays due to the sensitivity of its emission to solvent polarity.

Specifically, the furanyl dT analogue 1 exhibits strong visible emission in aqueous

environments (em 430 nm, F = 0.03) and significantly weaker emission in apolar media (em

~400 nm, F ~ 0.005). A furanyl-modified oligonucleotide forms stable duplexes with its perfect complement, and importantly retains its absorption (~320 nm) and emission properties

(em 430 nm). The furanyl dT analogue has been shown to positively detect the presence of abasic sites with a significantly increased emission. The utility of the furanyl dT analogue has been extended by the synthesis of the corresponding triphosphate and initial studies show acceptance of the modified nucleotide by DNA polymerase. Although specific photophysical results have yet to be seen, the furanyl dC analogue was site specifically incorporated into an oligonucleotide where it forms stable duplexes (data not shown) and represents a novel dC fluorescent probe whose applications are currently being explored. Although the furanyl dT analogue 1, having the desired photophysical properties, has been shown to be a useful fluorescent probe, its low quantum efficiency can possibly be improved by fusing the furanyl ring to the dU core. The penultimate step of such synthetic explorations is currently being explored. In conclusion, it has been shown that simple modifications to the native 97

nucleobases results in novel isomorphic nucleosides that are highly useful as probes for the study of nucleic acids.

2.29. Acknowledgements

Portions of Chapter 2 have been previously published in the following articles: (1) "Furan

decorated nucleoside analogues as fluorescent probes: synthesis, photophysical evaluation,

and site-specific incorporation" Tetrahedron 2007, 63, 3515–3527 by N. J. Greco and Y. Tor.

(2) "Synthesis and site-specific incorporation of a simple fluorescent pyrimidine" Nat. Protoc.

2007, 2, 305–316 by N. J. Greco and Y. Tor. (3) "Simple fluorescent pyrimidine analogues

detect the presence of DNA abasic sites" J. Am. Chem. Soc. 2005, 127, 10784–10785 by N.

J. Greco and Y. Tor. The dissertation author was the primary investigator and author of these

papers.

Appendix – Experimental information

A.1 – General procedures

All chemicals were obtained from commercial suppliers and used without further purification unless otherwise specified. Anhydrous pyridine and acetonitrile were obtained from Fluka. Anhydrous dioxane and triethylamine were obtained from Acros. Anhydrous toluene and tetrahydrofuran were obtained using a two-column purification system (Glasscontour System, Irvine, CA). Analytical thin-layer chromatography was performed on pre-coated silica gel aluminum-backed plates (Kieselgel 60 F254, E. Merck & Co., Germany). Flash chromatography was performed using silica gel (230–400 mesh) from E.M. Science or Silicycle. NMR solvents were purchased from Cambridge Isotope Laboratories (Andover, MA). All NMR spectra were recorded on a Varian Mercury 400MHz instrument with chemical shifts reported relative to residual deuterated solvent peaks. Chemical shifts () are reported in parts per million (ppm); multiplicities are indicated by s (singlet), d (doublet), t (triplet), q (quartet), dd (doublet of doublet), td (triplet of doublet), or m (multiplet); coupling constants (J) are reported in Hertz. Mass spectra were recorded at the UCSD Mass Spectrometry Facility, utilizing either a LCQDECA (Finnigan) ESI with a quadrpole ion trap or a MAT900XL (ThermoFinnigan) FAB double focusing mass spectrometer. UV-Visible experiments were carried out at ambient temperature in a quartz micro cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Hewlett Packard 8452A or 8453 diode array spectrometer. Steady State fluorescence experiments were carried out at ambient temperature in a micro fluorescence cell with a path length of 1.0 cm (Hellma GmH & Co KG, 98

Mullenheim, Germany) on a Perkin Elmer LS 50B luminescence spectrometer (figures 5, 7–9 and 15) or a Horiba fluoromax-3 luminescence spectrometer (figures 2 and 16).

A.2 – Synthetic procedures and X-ray structure data

3,5-Di-O-p-toluoyl-(+)-5-iodo-2-deoxyuridine (6). I O Anhydrous pyridine (5 ml) was added to IdUrd (1.00 g, 2.82 mmol) and stirred under an Ar atmosphere until NH all IdUrd was dissolved. The solution was cooled to 0 O N ºC and p-toluoyl chloride (785 ml, 918 mg, 5.94 mmol) O O was added dropwise. Reaction was allowed to warm O to room temperature and stirred under Ar overnight. O Water was added to the reaction solution and the O precipitate filtered off. Resulting solid was washed with cold water, cold ethanol and then cold ether. Product: white solid (1.42 g, 2.41 mmol, 85 % yield). 1 H NMR (400 MHz, DMSO-d6): 11.76 (s, NH, 1H), 8.09 (s, H-6, 1H), 7.90 (t, J = 7.8 Hz, Tol, 4H), 7.34 (t, J = 8.8 Hz, Tol, 4H), 6.22 (t, J = 7.0 Hz, H-1, 1H), 5.57–5.56 (m, H-3, 1H), 4.60 (d, J = 4.8

Hz, H-5, 2H), 4.52–4.49 (m, H-4, 1H), 2.67–2.60 (m, H-2, 2H), 2.39 (s, Tol-CH3, 3H), 2.38 13 (s, Tol-CH3, 3H); C NMR (100 MHz, DMSO-d6): 165.5 (Tol-carbonyl), 165.2 (Tol-carbonyl), 160.4 (C-4), 150.0 (C-2), 144.6 (C-6), 144.1 (Tol - C-4), 143.9 (Tol - C-4), 129.4 (Tol - C-2,3,5, and 6), 126.5 (Tol - C-1), 85.3 (C-1), 81.5 (C-4), 74.5 (C-3), 70.1 (C-5), 64.2 (C-5), 36.4 (C- + 2), 21.2 (Tol-CH3); ESI-MS calculated for C25H23IN2NaO7 [M+Na] 613.04, found 612.97.

5-(fur-2-yl)-2-deoxyuridine (1). To a suspension of IdUrd (520 mg, 1.47 mmol) and dichlorobis(triphenylphosphine)Pd(II) (21 mg, O O 0.03 mmol) in anhydrous dioxane (40 ml) was added 2- (tributylstannyl)furan (2 ml, 2.26 g, 6.35 mmol). The suspension NH was heated to 90 ºC under argon for 2 h, cooled, and filtered HO N through celite 545. The solvent was removed under reduced O O pressure where the resulting oil was triturated with hexanes (3x) to produce a solid. The resulting solid was taken up in a minimum of HO hot solvent (1/1 methanol/chloroform) and precipitated from hexanes. Product: white solid (407.7 mg, 1.39 mmol, 94 % yield). 1H NMR (400 MHz, DMSO-

d6): 11.63 (s, NH, 1H), 8.33 (s, H-6, 1H), 7.62 (s, H-5, 1H), 6.86 (d, J = 2.8 Hz, H-3, 1H), 6.52 (m, 1H, H-4), 6.22 (t, J = 6.6 Hz, 1H, H-1), 5.27 (d, J = 4.4 Hz, 1H, 3-OH), 5.08 (t, J = 4.4 Hz, 1H, 5-OH), 4.28 (m, 1H, H-3), 3.84 (m, 1H, H-4), 3.61 (m, 2H, H-5), 2.18 (t, J = 5.4 13 Hz, 2H, H-2); C NMR (100 MHz, DMSO-d6): 160.1 (C-4), 149.4 (C-2), 146.5 (C-2), 141.5 (C-5), 134.7 (C-6), 111.6 (C-4), 107.9 (C-3), 105.6 (C-5), 87.6 (C-4), 84.7 (C-1), + 70.4 (C-3), 61.1 (C-5), 40.1 (C-2); HR-FAB calculated for C13H15N2O6 [M+H] 295.0925, found: 295.0928; UV (buffer) max = 316 nm ( = 11,000), 260 = 13,000. Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 1. 99

Crystal data and structure refinement for 5-(fur-2-yl)-2-deoxyuridine (1). Identification code mich2

Empirical formula C13H14N2O6 Formula weight 294.26 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Monoclinic Space group P2(1) Unit cell dimensions a = 5.9450(4) Å a= 90° b = 20.5525(12) Å b= 91.2720(10)° c = 10.3217(6) Å g = 90° Volume 1260.84(13) Å3 Z 4 Density (calculated) 1.550 Mg/m3 Absorption coefficient 0.124 mm-1 F(000) 616 Crystal size 0.35 x 0.15 x 0.10 mm3 Theta range for data collection 1.97 to 27.48°. Index ranges -6<=h<=7, -26<=k<=26, -10<=l<=13 Reflections collected 7951 Independent reflections 5393 [R(int) = 0.0149] Completeness to theta = 27.48° 99.5 % Absorption correction Semi-empirical from equivalents Max. and min. transmission 1.000 and 0.854 Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 5393 / 1 / 492 Goodness-of-fit on F2 1.052 Final R indices [I>2sigma(I)] R1 = 0.0424, wR2 = 0.1088 R indices (all data) R1 = 0.0443, wR2 = 0.1104 Absolute structure parameter 0.3(7) Largest diff. peak and hole 1.149 and -0.228 e.Å-3

Atomic coordinates ( x 104) and equivalent isotropic displacement parameters (Å2x 103) for 5-(fur-2-yl)-2-deoxyuridine (1). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) O(1) 2841(3) -770(1) 3546(2) 16(1) O(2) 7093(3) -203(1) 7066(2) 19(1) O(3) 7720(3) -71(1) 1184(2) 16(1) O(4) 3912(3) -299(1) -836(2) 24(1) O(5) 7093(4) 1338(1) 1021(2) 32(1) O(6) 11694(3) 767(1) 4576(2) 18(1) N(1) 5039(3) -471(1) 5274(2) 15(1) N(2) 6079(3) -199(1) 3202(2) 14(1) C(1) 4548(4) -497(1) 3982(2) 14(1) C(2) 6875(4) -176(1) 5885(2) 15(1) C(3) 8417(4) 140(1) 5005(2) 14(1) C(4) 7946(4) 106(1) 3716(2) 16(1) 100

C(5) 5709(4) -259(1) 1778(2) 15(1) C(6) 3877(4) 188(1) 1237(2) 18(1) C(7) 4635(4) 266(1) -155(2) 18(1) C(8) 7209(4) 291(1) 6(2) 16(1) C(9) 8198(4) 967(1) 73(2) 23(1) C(10) 10429(4) 477(1) 5518(2) 19(1) C(11) 11304(4) 579(1) 6682(3) 21(1) C(12) 13278(4) 969(1) 6512(3) 22(1) C(13) 13434(4) 1066(1) 5238(2) 19(1) O(1) 12408(3) 8839(1) 6901(2) 19(1) O(2) 7258(3) 8290(1) 3780(2) 19(1) O(3) 9957(3) 7515(1) 9300(1) 14(1) O(4) 12070(3) 8305(1) 11564(2) 21(1) O(5) 5390(3) 7752(1) 10098(2) 31(1) O(6) 3594(3) 7222(1) 6575(2) 19(1) N(1) 9756(3) 8568(1) 5376(2) 15(1) N(2) 9495(3) 8190(1) 7481(2) 14(1) C(1) 10670(4) 8550(1) 6601(2) 15(1) C(2) 7819(4) 8251(1) 4921(2) 15(1) C(3) 6611(4) 7905(1) 5924(2) 15(1) C(4) 7483(4) 7899(1) 7156(2) 15(1) C(5) 10506(4) 8140(1) 8801(2) 15(1) C(6) 9606(4) 8640(1) 9747(2) 17(1) C(7) 9815(4) 8279(1) 11035(2) 16(1) C(8) 9331(4) 7572(1) 10656(2) 16(1) C(9) 6918(4) 7349(1) 10813(2) 22(1) C(10) 4531(4) 7559(1) 5578(2) 19(1) C(11) 3356(4) 7479(1) 4463(2) 19(1) C(12) 1531(4) 7057(1) 4768(3) 20(1) C(13) 1739(4) 6918(1) 6043(2) 21(1)

5-(thiophen-2-yl)-2-deoxyuridine (2). To a suspension of IdUrd (521 mg, 1.47 mmol) and dichlorobis(triphenylphosphine)Pd(II) (20 S O mg, 0.028 mmol) in anhydrous dioxane (40 ml) was added 2- (tributylstannyl)thiophene (2.5 ml, 2.9 g, 7.9 mmol). The suspension NH was heated to 95 ºC under argon for 5 h, cooled, and filtered through HO N celite 545. The solvent was removed under reduced pressure and O O the resulting oil was triturated with hexanes (3x) to produce a solid. The resulting solid was taken up in a minimum of hot solvent (1/1 HO methanol/chloroform) and precipitated from hexanes. Product: white 1 solid. (242 mg, 0.779 mmol, 53 % yield). H NMR (400 MHz, DMSO-d6): 11.7 (s, NH, 1H), 8.57 (s, H-6, 1H), 7.45 (d, J = 5.2 Hz, H-5, 1H), 7.40 (d, J = 3.2 Hz, H-3, 1H), 7.05 (t, J = 4.4 Hz, H-4), 6.22 (t, J = 6.2 Hz, H-1, 1H), 5.28 (m, 5-OH and 3-OH, 2H), 4.31 (m, H-3, 1H), 3.85 (m, H-4, 1H), 3.66 (m, H-5, 2H), 2.21 (m, H-2, 2H); 13C NMR (100 MHz, DMSO-

d6): 161.3 (C-4), 149.4 (C-2), 135.7 (C-6), 134.0 (C-2), 126.4, 125.7, and 122.5 (C-3, C- 4 and C-5), 108.3 (C-5), 87.6 (C-4), 84.8 (C-1), 70.0 (C-3), 60.9 (C-5), 40.4 (C-2),; + HR-FAB calculated for C13H15N2O5S [M+H] 311.0696, found: 311.0701; UV (buffer) max = 314 nm ( = 9,000). Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 2. 101

Crystal data and structure refinement for 5-(thiophen-2-yl)-2-deoxyuridine (2). Identification code mich03

Empirical formula C13H15N2O5.50S Formula weight 319.33 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Monoclinic Space group P2(1) Unit cell dimensions a = 7.7173(12) Å a= 90° b = 20.730(3) Å b= 97.428(2)° c = 8.7295(14) Å g = 90° Volume 1384.8(4) Å3 Z 4 Density (calculated) 1.532 g/cm3 Absorption coefficient 0.263 mm-1 F(000) 668 Crystal size 0.25 x 0.20 x 0.20 mm3 Theta range for data collection 1.96 to 27.57° Index ranges -9<=h<=9, -18<=k<=26, -9<=l<=11 Reflections collected 8780 Independent reflections 5290 [R(int) = 0.0205] Completeness to theta = 27.57° 98.8 % Absorption correction Semi-empirical from equivalents Max. and min. transmission 1.000 and 0.873 Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 5290 / 1 / 392 Goodness-of-fit on F2 1.059 Final R indices [I>2sigma(I)] R1 = 0.0420, wR2 = 0.1165 R indices (all data) R1 = 0.0430, wR2 = 0.1177 Absolute structure parameter 0.03(7) Largest diff. peak and hole 0.468 and -0.642 e Å-3

Atomic coordinates (x 104) and equivalent isotropic displacement parameters (Å2x 103) for 5- (thiophen-2-yl)-2-deoxyuridine (2). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) S(1) 4793(1) 3568(1) 9171(1) 20(1) N(1) 6404(3) 5364(1) 5408(2) 16(1) N(2) 6810(3) 4292(1) 4788(2) 15(1) O(1) 6241(2) 3469(1) 6346(2) 21(1) O(2) 7413(2) 5095(1) 3153(2) 19(1) O(3) 6810(3) 6406(1) 6415(2) 24(1) O(4) 5769(3) 7408(1) 3671(3) 34(1) O(5) 3289(3) 6501(1) 7145(3) 36(1) C(1) 4056(4) 3796(2) 10858(3) 22(1) C(2) 4031(4) 4448(2) 11014(3) 21(1) C(3) 4621(3) 4786(1) 9747(3) 18(1) C(4) 5089(3) 4361(1) 8629(3) 15(1) 102

C(5) 5709(3) 4540(1) 7166(3) 14(1) C(6) 5805(3) 5166(1) 6741(3) 17(1) C(7) 6923(3) 4927(1) 4368(3) 16(1) C(8) 6249(3) 4054(1) 6132(3) 14(1) C(9) 6537(4) 6055(1) 5021(3) 19(1) C(10) 4870(4) 6317(1) 4123(3) 23(1) C(11) 4843(4) 7012(2) 4638(3) 26(1) C(12) 5786(4) 6988(1) 6292(3) 25(1) C(13) 4560(4) 6992(2) 7520(4) 32(1) S(2) 9243(1) 10067(1) 3377(1) 24(1) N(3) 10702(3) 8234(1) -333(2) 16(1) N(4) 11081(3) 9291(1) -1047(2) 16(1) O(6) 10415(3) 10138(1) 396(2) 23(1) O(7) 11829(2) 8471(1) -2573(2) 19(1) O(8) 9115(2) 7279(1) -632(2) 19(1) O(9) 11954(3) 6121(1) -303(2) 22(1) O(10) 8716(3) 7085(1) 2563(2) 32(1) C(14) 8561(4) 9857(2) 5083(3) 26(1) C(15) 8392(4) 9218(2) 5246(3) 24(1) C(16) 8846(3) 8839(1) 3961(2) 12(1) C(17) 9359(3) 9275(1) 2800(3) 17(1) C(18) 9970(3) 9078(1) 1347(3) 16(1) C(19) 10092(3) 8451(1) 981(3) 17(1) C(20) 11256(3) 8651(1) -1407(3) 15(1) C(21) 10480(3) 9548(1) 250(3) 17(1) C(22) 10825(3) 7534(1) -575(3) 17(1) C(23) 11944(3) 7186(1) 750(3) 18(1) C(24) 11109(3) 6522(1) 719(3) 18(1) C(25) 9179(3) 6650(1) 97(3) 18(1) C(26) 8021(4) 6656(2) 1366(3) 26(1) O(1S) 746(6) 6369(3) 4609(6) 105(2)

103

5-(oxazol-2-yl)-2-deoxyuridine (3). To a solution 6 (98 mg, 0.17 N mmol) and tetrakis(triphenylphosphine)Pd(0) (20 mg, 0.017 mmol) in O O anhydrous toluene (3 ml) was added 2-(tributylstannyl)oxazole (106 ml, 181 mg, 0.506 mmol). The solution was refluxed under argon for NH 8h, cooled and the solvent removed under reduced pressure. The resulting solid was taken up in 20% methanol/chloroform and run HO N O O through a silica plug where fractions were collected. Those containing the desired product (fluorescent spot) were consolidated, HO solvent evaporated under reduced pressure, and resulting solid was taken up in 5% THF/methanol (3 ml) and K2CO3 (23 mg, 0.17 mmol) was added. The suspension stirred at rt for 12 h. The solvent was removed under reduced pressure and product was isolated by flash chromatography (85/15 chloroform/methanol). Resulting solid was triturated with hexanes. Product: white solid (5 mg, 0.02 mmol, 10 % 1 yield). H NMR (400 MHz, DMSO-d6): 11.65 (s, NH, 1H), 8.58 (s, H-6, 1H), 8.11 (s, H-5, 1H), 7.26 (s, H-4, 1H), 6.17 (t, J = 6.6 Hz, H-1, 1H), 5.24 (d, J = 3.6 Hz, 3-OH, 1H), 5.00 (t, J = 4.6 Hz, 5-OH, 1H), 4.25 (m, H-3, 1H), 3.85 (m, H-4, 1H), 3.58 (m, H-5, 2H), 2.19 (m, H- 13 2, 2H); C NMR (100 MHz, DMSO-d6): 159.6 (C-4), 156.6 (C-2), 149.6 (C-2), 142.1 (C-6), 139.6 (C-5), 127.7 (C-4), 102.9 (C-5), 87.8 (C-4), 85.2 (C-1), 70.3 (C-3), 61.0 (C-5), 40.3 + (C-2); ESI-MS calculated for C12H13N3NaO6 [M+Na] 318.07, found 317.99. UV (buffer) max = 296 nm ( = 10,000). Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 3.

Crystal data and structure refinement for 5-(oxazol-2-yl)-2-deoxyuridine (3). Identification code tor1

Empirical formula C12H14N3O6.50 Formula weight 304.26 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Triclinic Space group P1 Unit cell dimensions a = 6.6557(13) Å a= 65.158(3)° b = 10.1584(19) Å b= 83.596(3)° c = 10.397(2) Å g = 85.909(3)° Volume 633.7(2) Å3 Z 2 Density (calculated) 1.595 Mg/m3 Absorption coefficient 0.132 mm-1 F(000) 318 Crystal size 0.28 x 0.14 x 0.10 mm3 Theta range for data collection 2.17 to 27.53°. Index ranges -8<=h<=8, -13<=k<=12, -13<=l<=13 Reflections collected 5352 Independent reflections 4786 [R(int) = 0.0146] Completeness to theta = 27.53° 93.2 % Absorption correction Semi-empirical from equivalents Max. and min. transmission 1.000 and 0.670 Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 4786 / 3 / 488 Goodness-of-fit on F2 1.039 104

Final R indices [I>2sigma(I)] R1 = 0.0326, wR2 = 0.0842 R indices (all data) R1 = 0.0334, wR2 = 0.0856 Absolute structure parameter -0.3(6) Largest diff. peak and hole 0.327 and -0.269 e.Å-3

Atomic coordinates ( x 104) and equivalent isotropic displacement parameters (Å2x 103) for 5-(oxazol-2-yl)-2-deoxyuridine (3). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) O(1) 2210(2) 644(2) 4769(2) 22(1) O(2) 2950(2) -2322(1) 9385(1) 19(1) O(3) 2898(2) 1381(1) 10330(1) 16(1) O(4) 3974(2) 3914(1) 5278(2) 19(1) O(5) 1429(2) 6002(2) 2846(2) 23(1) O(6) 3480(2) 7236(1) 5855(1) 19(1) N(1) 2315(2) 1715(2) 6306(2) 15(1) N(2) 2495(2) -802(2) 7107(2) 15(1) N(3) 2488(2) -1006(2) 11435(2) 17(1) C(1) 2332(3) 524(2) 5968(2) 14(1) C(2) 2447(3) 1540(2) 7668(2) 14(1) C(3) 2602(3) 226(2) 8777(2) 14(1) C(4) 2697(3) -1077(2) 8505(2) 14(1) C(5) 2689(3) 117(2) 10212(2) 14(1) C(6) 2816(3) 992(2) 11783(2) 17(1) C(7) 2568(3) -437(2) 12443(2) 17(1) C(8) 2334(3) 3182(2) 5130(2) 17(1) C(9) 461(3) 4135(2) 5135(2) 18(1) C(10) 1316(3) 5654(2) 4332(2) 17(1) C(11) 3415(3) 5435(2) 4880(2) 15(1) C(12) 3470(3) 5719(2) 6206(2) 18(1) O(1B) 7813(2) 856(1) -617(1) 19(1) O(2B) 7032(2) -2206(1) 4076(1) 18(1) O(3B) 7279(2) 1439(1) 5078(1) 17(1) O(4B) 9354(2) 4119(1) 311(1) 18(1) O(5B) 8200(2) 5945(2) -2668(2) 23(1) O(6B) 6613(2) 7229(2) 706(2) 20(1) N(1B) 8070(2) 1791(2) 1003(2) 14(1) N(2B) 7510(2) -662(2) 1746(2) 14(1) N(3B) 7577(2) -972(2) 6130(2) 17(1) C(1B) 7797(3) 662(2) 625(2) 14(1) C(2B) 7915(3) 1603(2) 2390(2) 14(1) C(3B) 7557(3) 293(2) 3481(2) 14(1) C(4B) 7349(3) -965(2) 3182(2) 14(1) C(5B) 7463(3) 166(2) 4939(2) 14(1) C(6B) 7304(3) 1037(2) 6525(2) 18(1) C(7B) 7484(3) -406(2) 7161(2) 18(1) C(8B) 8411(3) 3246(2) -191(2) 16(1) C(9B) 6453(3) 4043(2) -741(2) 18(1) C(10B) 7064(3) 5620(2) -1335(2) 18(1) 105

C(11B) 8407(3) 5571(2) -204(2) 17(1) C(12B) 7316(3) 5754(2) 1087(2) 19(1) O(1S) 12805(3) 4148(2) 1646(2) 35(1)

5-(thiazol-2-yl)-2-deoxyuridine (4). To a solution of 6 (104 mg, N 0.176 mmol) and dichlorobis(triphenylphosphine)Pd(II) (24 mg, S O 0.033 mmol) in anhydrous dioxane (3 ml) was added 2- (tributylstannyl)thiazole (160 ml, 190 mg, 0.509 mmol). The solution NH was refluxed at 90 ºC under argon for 20h, cooled and the solvent was removed under reduced pressure. The resulting solid was HO N O O triturated with methanol. The precipitate was taken up in 5% THF/methanol (3 ml) and K2CO3 (33 mg, 0.24 mmol) was added. HO The suspension stirred at rt for 24 h. The solvent was removed under reduced pressure and product was isolated by flash chromatography (90/10 chloroform/methanol). The resulting solid was triturated with hexanes. 1 Product: white solid (19 mg, 0.061 mmol, 34 % yield). H NMR (400 MHz, DMSO-d6): 11.93 (s, NH, 1H), 8.82 (s, H-6, 1H), 7.85 (d, J = 3.2 Hz, H-5, 1H), 7.65 (d, J = 3.2 Hz, H-4, 1H), 6.21 (t, J = 6.6 Hz, H-1, 1H), 5.29 (d, J = 4.4 Hz, 3-OH, 1H), 5.01 (t, J = 5.0 Hz, 5-OH, 1H), 4.29–4.25 (m, H-3, 1H), 3.88–3.86 (m, H-4, 1H), 3.61–3.58 (m, H-5, 2H), 2.23–2.19 (m, H- 13 2, 2H); C NMR (100 MHz, DMSO-d6): 161.3 (C-4), 158.1 (C-2), 149.4 (C-2), 141.8 (C- 4), 138.5 (C-6), 119.8 (C-5), 107.6 (C-5), 87.8 (C-4), 85.3 (C-1), 70.5 (C-3), 61.3 (C-5), + 40.1 (C-2); ESI-MS calculated for C12H13N3NaO5S [M+Na] 334.05, found 333.94; UV (buffer) lmax = 316 nm (e = 11,500). Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 4.

Crystal data and structure refinement for 5-(thiazol-2-yl)-2-deoxyuridine (4). Identification code tor30

Empirical formula C12H13N3O5S Formula weight 311.31 Temperature 296(2) K Wavelength 1.54178 Å Crystal system Monoclinic Space group P2(1) Unit cell dimensions a = 5.9563(7) Å a= 90° b = 20.591(3) Å b= 90.762(9)° c = 10.3793(11) Å g = 90° Volume 1272.9(3) Å3 Z 4 Density (calculated) 1.624 g/cm3 Absorption coefficient 2.544 mm-1 F(000) 648 Crystal color, size Colorless, 0.33 x 0.04 x 0.02 mm3 Theta range for data collection 4.29 to 63.48° Index ranges -5<=h<=6, -23<=k<=21, -11<=l<=8 Reflections collected 4450 Independent reflections 2785 [R(int) = 0.0407] Completeness to theta = 52.50° 97.9 % Absorption correction Multi-scan 106

Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 2785 / 1 / 379 Goodness-of-fit on F2 0.994 Final R indices [I>2sigma(I)] R1 = 0.0424, wR2 = 0.0982 R indices (all data) R1 = 0.0542, wR2 = 0.1053 Absolute structure parameter 0.05(3) Largest diff. peak and hole 0.263 and -0.314 e Å-3

Atomic coordinates (x 104) and equivalent isotropic displacement parameters (Å2x 103)for 5- (thiazol-2-yl)-2-deoxyuridine (4). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) S(1) -3543(2) 2425(1) 4179(1) 21(1) S(1) 945(2) 4383(1) 3040(1) 22(1) O(1) -7321(6) 1664(2) 3912(3) 20(1) O(1) -2999(6) 5130(2) 2983(3) 21(1) O(2) -12303(6) 1103(2) 6966(3) 20(1) O(2) -7217(6) 5759(2) 6366(3) 20(1) O(3) -9803(6) 2445(2) 9330(3) 19(1) O(3) -2449(6) 5041(2) 8833(3) 19(1) O(4) -11801(6) 1679(2) 11617(3) 26(1) O(4) -6005(6) 5286(2) 10784(3) 28(1) O(5) -5246(7) 2211(2) 10140(4) 39(1) O(5) -3566(9) 3682(2) 8627(4) 47(1) N(1) -9734(8) 1386(2) 5464(4) 16(1) N(1) -5052(7) 5432(2) 4700(4) 15(1) N(2) -9389(7) 1756(2) 7565(4) 16(1) N(2) -4035(7) 5172(2) 6786(4) 17(1) N(3) 1686(7) 4211(2) 5453(4) 19(1) N(3) -3333(7) 2695(2) 6588(4) 20(1) C(1) -1394(9) 2896(3) 4729(5) 21(1) C(1) 3251(9) 3951(3) 3515(5) 22(1) C(2) -1544(9) 2987(3) 6021(5) 17(1) C(2) 3371(9) 3912(3) 4809(5) 20(1) C(3) -4545(8) 2380(3) 5718(4) 14(1) C(3) 278(9) 4470(3) 4629(5) 16(1) C(4) -7833(9) 1697(3) 5049(5) 18(1) C(4) -3234(9) 5122(3) 4149(5) 18(1) C(5) -6595(9) 2041(3) 6045(5) 15(1) C(5) -1741(9) 4816(3) 5062(5) 18(1) C(6) -7400(9) 2048(3) 7249(5) 18(1) C(6) -2195(9) 4861(3) 6333(5) 16(1) C(7) -10558(9) 1393(3) 6685(5) 15(1) C(7) -5537(9) 5468(3) 5975(5) 18(1) C(8) -10363(9) 1815(3) 8853(5) 17(1) C(8) -4400(8) 5252(3) 8199(4) 16(1) C(9) -9476(10) 1338(3) 9816(5) 23(1) C(9) -6329(9) 4856(3) 8696(5) 22(1) C(10) -9623(10) 1692(3) 11065(5) 24(1) 107

C(10) -5541(9) 4722(3) 10072(5) 22(1) C(11) -9149(9) 2392(3) 10677(4) 21(1) C(11) -3019(9) 4625(3) 9903(5) 20(1) C(12) -2286(11) 3937(3) 9654(6) 34(2) C(12) -6738(10) 2612(3) 10839(5) 30(2)

3,5-di-O-acetyl-5-(fur-2-yl)-2-deoxyuridine (10).To a solution of 1 (1.0 g, 3.5 mmol) in anhydrous pyridine (15 ml) was added O O acetic anhydride (825 l, 891 mg, 8.73 mmol). The reaction was allowed to stir under argon at room temperature overnight. NH Solvent removed under reduced pressure. Resulting oil taken up in chloroform (200ml) and washed with 1M HCl (3x 100 ml) and O N O O water (100 ml). The organic layer was dried over sodium sulfate and the solvent removed under reduced pressure. Product: light O O orange foam (1.34 g, 3.5 mmol, 99 % yield). 1H NMR (400 MHz, O CDCl3): 8.60 (s, NH, 1H), 7.97 (s, H-6, 1H), 7.32 (d, J = 1.6 Hz, H-5, 1H), 7.05 (d, J = 3.2 Hz, H-3, 1H), 6.45 (q, J = 2 and 3.2 Hz, H-3, 1H), 6.41 (td, J = 8.8 and 5.6 Hz, H-1, 1H), 5.28–5.26 (m, H-3, 1H), 4.38–4.37 (m, H-5, 2H), 4.29 (q, J = 3 and 5 Hz, H-4, 1H), 2.55–2.50 (m, H-2, 1H), 2.28–2.19 (m, H-2, 13 1H), 2.14 (s, Ac, 3H), 2.11 (s, Ac, 3H); C NMR (100 MHz, CD3Cl): 170.1 (Ac carbonyl), 170.0 (Ac carbonyl), 159.2 (C-4), 148.9 (C-2), 145.4 (C-2), 140.9 (C-5), 132.2 (C-6), 112.0 (C- 4), 109.7 (C-3), 107.7 (C-5), 85.2 (C-4), 82.5 (C-1), 74.6 (C-3), 64.1 (C-5), 38.1 (C-2), + 21.0 (Ac methyl), 20.8 (Ac methyl); ESI-MS calculated for C17H19N2O8 [M+H] 379.11, found + 378.84 and for C17H18N2NNaO8 [M+Na] 401.10, found 401.00.

5-(fur-2-yl)-2-deoxycytidine (7). To a solution of 10 (800 mg, 2.1 mmol), 2,4,6-triisopropylbenzenesulfonyl chloride (2.0 g, 6.72 mmol) and 4-dimethylaminopyridine (825 mg, 6.72 mmol) in O NH 2 anhydrous acetonitrile (70 ml) was added triethylamine (937 l, 680 mg, 6.72 mmol). The reaction was allowed to stir under argon at N room temperature for 36 h. Once all starting material was N HO consumed (monitored by tlc), concentrated ammonium hydroxide O O (90 ml) was added to the reaction flask. The reaction was again HO allowed to stir at room temperature overnight. Solvent removed under reduced pressure. Resulting solid taken up in cold chloroform and solid filtered off. Product: off white solid (590 mg, 2.0 mmol, 95% yield). 1H

NMR (400 MHz, DMSO-d6): 8.25 (s, H-6, 1H), 7.68–7.67 (m, H-5, 1H), 7.64 (s br, 4-NH2, 1H), 6.65 (s br, 4-NH2, 1H), 6.57–6.54 (m, H-4 and H-3, 2H), 6.16 (t, J = 6.4 Hz, H-1, 1H), 5.22 (d, J = 4 Hz, 3-OH, 1H), 5.08 (t, J = 4.8 Hz, 5-OH, 1H), 4.24–4.20 (m, H-3, 1H), 3.08– 3.78 (m, H-4, 1H), 3.64–3.54 (m, H-5, 2H), 2.20–2.15 (m, H-2, 1H), 2.07–2.01 (m, H-2, 1H); 13 C NMR (100 MHz, DMSO-d6): 161.5 (C-4), 153.5 (C-2), 147.2 (C-2), 142.3 (C-5), 139.6 (C-6), 111.3 (C-4), 106.5 (C-3), 98.0 (C-5), 87.3 (C-4), 85.2 (C-1), 69.9 (C-3), 60.9 (C-5), + 40.8 (C-2); ESI-MS calculated for C13H16N3O5 [M+H] 294.11, found 293.84; UV (buffer) max = 306 nm ( = 5,000), 260 nm ( = 9,600). Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 7.

108

Crystal data and structure refinement for 5-(fur-2-yl)-2-deoxycytidine (7). X-ray ID tor19

Empirical formula C13H15N3O5 Formula weight 293.28 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Orthorhombic Space group P2(1)2(1)2(1) Unit cell dimensions a = 4.5420(2) Å a= 90. b = 10.2410(4) Å b= 90° c = 26.9210(10) Å g = 90° Volume 1252.22(9) Å3 Z 4 Density (calculated) 1.556 Mg/m3 Absorption coefficient 0.121 mm-1 F(000) 616 Crystal size 0.40 x 0.20 x 0.10 mm3 Crystal color/habit colorless needle Theta range for data collection 2.13 to 28.19°. Index ranges -3<=h<=6, -10<=k<=13, -35<=l<=35 Reflections collected 8150 Independent reflections 2850 [R(int) = 0.0177] Completeness to theta = 25.00° 99.9 % Absorption correction Semi-empirical from equivalents Max. and min. transmission 0.9880 and 0.9530 Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 2850 / 0 / 192 Goodness-of-fit on F2 1.080 Final R indices [I>2sigma(I)] R1 = 0.0439, wR2 = 0.1126 R indices (all data) R1 = 0.0441, wR2 = 0.1129 Absolute structure parameter 0.5(12) Largest diff. peak and hole 0.744 and -0.444 e.Å-3

Atomic coordinates ( x 104) and equivalent isotropic displacement parameters (Å2x 103) for 5- (fur-2-yl)-2-deoxycytidine (7). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) C(1) 8909(4) 8890(2) 3135(1) 16(1) C(2) 10006(4) 8135(2) 3551(1) 14(1) C(3) 8860(4) 6912(2) 3594(1) 16(1) C(4) 5806(4) 7206(2) 2879(1) 15(1) C(5) 12081(5) 8614(2) 3923(1) 16(1) C(6) 13822(6) 9678(2) 3969(1) 31(1) C(7) 15342(6) 9532(3) 4436(1) 32(1) C(8) 14386(6) 8463(3) 4636(1) 36(1) C(9) 5708(4) 5099(2) 3333(1) 14(1) C(10) 8034(4) 4037(2) 3298(1) 15(1) C(11) 6629(4) 2957(2) 3608(1) 15(1) 109

C(12) 5205(4) 3723(2) 4032(1) 15(1) C(13) 7111(5) 3943(2) 4485(1) 18(1) N(1) 6828(4) 6434(2) 3271(1) 14(1) N(2) 6910(4) 8419(2) 2822(1) 17(1) N(3) 9829(4) 10108(2) 3047(1) 20(1) O(1) 3921(3) 6752(1) 2592(1) 19(1) O(2) 12379(5) 7856(2) 4335(1) 37(1) O(3) 4493(3) 4993(1) 3827(1) 16(1) O(4) 4325(3) 2319(1) 3344(1) 19(1) O(5) 7616(3) 2746(2) 4745(1) 21(1)

8-(fur-2-yl)-adenosine (8). To a suspension of 11 (208 mg, NH2 0.6 mmol) and ammonium sulfate (43 mg) in anhydrous N 1,1,1.3,3,3-hexamethyldisilazane (20 ml) was added O N anhydrous pyridine (2 ml). The reaction was refluxed under HO N O N argon overnight. Solvent was removed under reduced pressure and the resulting oil was used without further HO purification. To a solution of the resulting trimethylsilyl protected nucleoside and dichlorobis(triphenylphosphine)Pd(II) (28 mg, 0.04 mmol) in anhydrous tetrahydrofuran (10 ml) was added 2-(tributylstannyl)furan (950 l, 1.0 g, 3.0 mmol) in a pressure tube and was heated to 90 °C for 24 h. Solvent removed under reduced pressure and the resulting oil run through a silica plug with 5% methanol/dichloromethane. The resulting oil was taken up in methanol (5 ml) to which potassium carbonate (500 mg, 3.6 mmol) was added. The reaction was allowed to stir at room temperature overnight. Solid was filtered off and the product was recrystallized from methanol. Resulting white solid was run through a silica plug using 50/50 water/acetonitrile to remove any remaining potassium carbonate. Product: white solid (18.6 mg, 0.056 mmol, 9% 1 yield). H NMR (400 MHz, DMSO-d6): 8.13 (s, H-2, 1H), 8.01 (d, J = 2.4 Hz, H-5, 1H), 7.58 (s br, 6-NH2, 2H), 7.13 (d, J = 4.8 Hz, H-3, 1H), 6.77–6.76 (m, H-4, 1H), 6.10 (d, J = 8.8 Hz, H-1, 1H), 5.77 (dd, J = 4.4 and 12 Hz, 5-OH, 1H), 5.46 (d, J = 8.4 Hz, 2-OH, 1H), 5.21 (d, J = 5.6 Hz, 3-OH, 1H), 5.14–5.08 (m, H-2, 1H), 4.12–4.17 (m, H-3, 1H), 3.98 (d, J = 2.4 Hz, H- 13 4, 1H), 3.72–3.67 (m, H-5, 1H), 3.51–3.49 (m, H-5, 1H); C NMR (100 MHz, DMSO-d6): 156.3 (C-6), 152.3 (C-2), 149.6 (C-4), 145.6 and 143.3 (C-2 and C-5), 141.5 (C-8), 119.4 (C-5), 113.9 and 112.1 (C-3 and C-4), 89.3 (C-1), 86.7 (C-4), 71.6 (C-2), 71.0 (C-3), + 62.2 (C-5); ESI-MS calculated for C14H16N5O5 [M+H] 334.12, found 334.02; UV (buffer) max = 304 nm ( = 18,000).

110

8-(fur-2-yl)-guanosine (9). To a suspension of 12 (139 mg, O N 0.4 mmol) and ammonium sulfate (21 mg) in anhydrous 1,1,1,3,3,3-hexamethyldisilazane (10 ml) was added O NH HO N anhydrous pyridine (1 ml). The reaction was stirred under O N argon at reflux overnight. Solvent was removed under NH 2 reduced pressure and the resulting oil was used without HO further purification. To a solution of the resulting trimethylsilyl protected nucleoside and dichlorobis(triphenylphosphine)Pd(II) (8.5 mg, 0.01 mmol) in anhydrous tetrahydrofuran (10 ml) was added 2-(tributylstannyl)furan (600 l, 680 mg, 1.9 mmol) in a pressure tube and was heated to 90 °C for 24 h. Solvent removed under reduced pressure and the resulting oil run through a silica plug with 5% methanol/dichloromethane. The resulting oil was taken up in methanol (5 ml) to which potassium carbonate (300 mg, 2.2 mmol) was added. The reaction was allowed to stir at room temperature overnight. Solid was filtered off and the product was recrystallized from methanol. Resulting white solid was run through a silica plug using 50/50 water/acetonitrile to remove any remaining potassium carbonate. Product: white solid (13.8 mg, 0.04 mmol, 10% 1 yield). H NMR (400 MHz, DMSO-d6): 10.83 (s, 1-NH, 1H), 7.90 (d, J = 1.6 Hz, H-5, 1H), 6.94 (d, J = 3.2, H-3, 1H), 6.68 (q, J = 1.7 Hz, H-4, 1H), 6.46 (s br, 2-NH2, 2H), 5.91 (d, J = 6.4 Hz, H-1, 1H), 5.38 (d, J = 6.0 Hz, 2-OH, 1H), 5.04–4.96 (m, 3-OH, 5-OH and H-2, 3H), 4.13–4.08 (m, H-3, 1H), 3.84 (dt, J = 4.8 and 8.8, H-4, 1H), 3.68–3.63 (m, H-5, 1H), 3.54– 13 3.48 (m, H-5, 1H); C NMR (100 MHz, DMSO-d6): 156.5 (C-6), 153.3 (C-2), 151.7 (C-4), 144.4 and 143.9 (C-2 and C-5), 138.2 (C-8), 117.3 (C-5), 112.1 and 111.7 (C-3 and C- 4), 89.0 (C-1), 85.7 (C-4), 70.6 and 70.5 (C-2 and C-3), 62.1 (C-5); ESI-MS calculated + for C14H16N5O6 [M+H] 350.11, found 349.80; UV (buffer) max = 294 nm ( = 16,000).

5-dimethoxytrityl-5-(fur-2-yl)-2-deoxyuridine (13). To O a solution of 1 (163 mg, 0.554 mmol) and DMT-Cl (226 O O mg, 0.669 mmol) in anhydrous pyridine (3 ml) was added triethylamine (60 ml). The reaction was stirred at room NH temperature under argon for 4 hr and evaporated under reduced pressure. The product was purified by flash O N O O column chromatography (1/1 ethylaceate/hexanes, 1% Et3N). Product: light brown foam (234 mg, 0.393 mmol, 1 HO 71% yield). H NMR (400 MHz, DMSO-d6): 11.70 (s, NH, 1H), 7.96 (s, H-6, 1H), 7.27 – 7.16 (m, DMT and H- O 5, 10H), 6.84 – 6.81 (m, DMT and H-3, 5H), 6.46 – 6.44 (m, H-4, 1H), 6.18 (t, J = 6.6 Hz, H-1, 1H), 5.36 (d, J = 4.4 Hz, 3-OH, 1H), 4.23 – 4.19 (m, H-3, 1H), 3.96 – 3.93 (m, H-4, 1H), 3.69 (s, DMT- 13 OCH3, 6H), 3.24 – 3.15 (m, H-5, 2H), 2.27 – 2.24 (m, H-2, 2H); C NMR (100 MHz, DMSO- d6): 160.1, 158.0, 149.3, 146.1, 144.7, 141.2, 135.5, 133.9, 129.6, 127.7, 127.6, 126.6, 113.1, 111.4, 107.9, 105.6, 85.8, 85.1, 70.3, 63.6, 55.0, 40.2; ESI-MS calculated for + C34H32N2NaO8 [M+Na] 619.21, found 619.08.

111

3-2-cyanoethyldiisopropylphosphoramidite-5- O dimethoxytrityl-5-(fur-2-yl)-2-deoxyuridine (14). 13 was taken up in anhydrous acetonitrile and solvent was O O removed under reduced pressure (5x). To a solution of 13 (58 mg, 0.10 mmol) and 1-H tetrazole (0.45 M NH solution in acetonitrile: 215 ml, 0.0968 mmol) in O N anhydrous acetonitrile (2 ml) was added 2-cyanoethyl O O tetraisopropylphosphorodiamidite (37 ml, 35 mg, 0.12 mmol). The solution was stirred at room temperature O under argon for 4 hrs. The reaction mixture was diluted P N with cold 1% triethylamine/dichloromethane (100 ml) O O and washed with 1M NaHCO3 (2x 10 ml) and brine (2x NC 10 ml). The organic layer was dried over sodium sulfate and evaporated under reduced pressure. The product was purified by flash column chromatography (1/1 ethylaceate/hexanes, 1% Et3N). Product: 1 light brown foam (50 mg, 0.063 mmol, 65% yield) H NMR (400 MHz, DMSO-d6): 11.60 (s, H-6, 1H), 7.39 – 7.17 (m, DMT and H-5, 10H), 6.82 – 6.79 (m, DMT and H-3, 5H), 6.45 – 6.43 (m, H-4, 1H), 6.17 (t, J = 6.4 Hz, H-1, 1H), 4.50 – 4.44 (m, H-3, 1H), 4.11 – 4.08 (m,

H-4, 1H), 3.69 – 3.39 (m, DMT-OCH3, isopropyl and cyanoethyl, 10H), 3.24 – 3.19 (m, H-5, 2H), 2.64 – 2.61 (m, cyanoethyl, 2H), 1.23 – 1.08 (m, isopropyl, 12H); 13C NMR (100 MHz,

DMSO-d6): 160.1, 158.1, 149.2, 146.1, 144.6, 141.2, 135.3, 134.0, 129.7, 127.7, 127.6, 126.6, 118.7, 113.1, 111.5, 108.0, 105.7, 85.9, 85.2, 72.3, 63.5, 58.5, 55.0, 42.6, 42.5, 40.2, 31 24.3, 24.3, 24.2, 19.8; P NMR (162 MHz, DMSO-d6, referenced to H3PO4): 148.7, 148.3; + ESI-MS calculated for C43H49N4NaO9P [M+Na] 819.31, found 819.14.

O 5-(fur-2-yl)-5-triphosphate-2-deoxyuridine (26). To a cold solution (0°C) of nucleoside 1 (0.0586 g, 0.2 O O O O NH mmol) in trimethyl phosphate (0.8 mL) was added freshly distilled POCl (30 L, 0.32 mmol). After O P O P O P O 3 μ N O stirring the reaction mixture for 1 h at 0°C, a solution O O O O of bis-tributylammoniumpyrophosphate (0.5 M in DMF, 2 mL, 1.0 mmol) and tributylamine (475 L, 2.0 (Et NH+) μ 3 4 HO mmol) were added simultaneously. The reaction mixture was quenched after 15 min. with 1 M triethylammonium bicarbonate buffer (TEAB, pH 7.5, 20 mL) and allowed to stir for 15 min. The reaction mixture was evaporated under reduced pressure at ~23°C. The residue was purified first on a DEAE sephadex-A25 anion exchange column (0.01–1.0 M, TEAB buffer, pH 7.5) followed by reversed-phase HPLC (Vydac C18 column, 1.0 cm 25 cm, 5 μm TP silica, 8-15% acetonitrile in 100 mM triethylammonium acetate buffer over 15 min). Appropriate fractions were repeatedly 1 lyophilized. Product: off white solid (9.2 mg, 5%). H NMR (400 MHz, D2O): 8.16 (s, H-6, 1H), 7.55 (s, H-5, 1H), 6.83 (d, J = 3.2 Hz, H-3, 1H), 6.50–6.49 (m, H-4, 1H), 6.33 (t, J = 6.8 Hz, H-1, 1H), 4.23–4.17 (m, H-3 and H-4, 2H), 3.04–2.96 (m, H-5, 2H), 2.45–2.35 (m, 13 H-2, 2H) ; C NMR (100 MHz, D2O): 162.6, 150.8, 145.5, 142.8, 135.9, 111.6, 109.0, 31 107.4, 85.9, 85.8, 71.0, 65.7, 38.8; P NMR (162 MHz, D2O, referenced to H3PO4): 10.15 (d, J = 20.6 Hz, P), 10.88 (d, J = 19.4 Hz, P), 22.54 (t, J = 20.1 Hz, P); ESI-MS calculated - for C13H16N2O15P3 [M-H] 532.98, found 533.01.

112

O 3,5-tertbutyldimmethylsilyl-5-(fur-2-yl)-2-deoxyuridine (29). A solution of 1 (0.5 g, 1.7 mmol), tertbutyldimethylsilyl O NH chloride (1.6 g, 10.6 mmol) and imidazole (1.4 g, 20.4 mmol) in anhydrous dimethylformamide (20 ml) was allowed to stir at Si O N O room temperature for 20 h. Solvent removed under reduced O pressure. Resulting oil taken up in dichloromethane (100ml) and washed with saturated sodium bicarbonate (2x 50 ml) and brine (50 ml). The organic layer was dried over sodium sulfate Si O and the solvent removed under reduced pressure. The product was purified by gravity column chromatography (3/7 ethyl acetate/hexanes). Product: white foam (0.857 g, 1.6 mmol, 96 % yield). 1H NMR (400

MHz, CDCl3): 8.94 (s, NH, 1H), 8.04 (s, H-6, 1H), 7.31 (d, J = 2.0 Hz, H-5, 1H), 7.03 (d, J = 3.2, H-3, 1H), 6.43 (dd, J = 3.4 and 1.8 Hz, H-3, 1H), 6.34 (td, J = 8.2 and 5.8 Hz, H-1, 1H), 4.44–4.42 (m, H-3, 1H), 4.00–3.97 (m, H-5, 1H) 3.88–3.76 (m, H-4, 2H), 2.35–2.30 (m, H-2, 1H), 2.10–2.02 (m, H-2, 1H), 0.88 (s, t-butyl, 9H), 0.87 (s, t-butyl, 9H), 0.08–0.06 (m, 13 Me, 12H); C NMR (100 MHz, CD3Cl): 159.8, 149.1, 145.7, 140.7, 133.5, 111.7, 109.2, 107.0, 88.2, 85.6, 72.5, 63.0, 41.6, 26.0, 25.9, 18.5, 18.2, 4.5, 4.6, 5.3, 5.4; ESI-MS calculated + + for C25H43N2O6Si2 [M+H] 523.27, found 522.97 and for C25H42N2NaO6Si2 [M+Na] 545.25, found 545.04.

3,5-tertbutyldimmethylsilyl-5-(fur-2-yl)-2-deoxycytidine NH2 (30). To a solution of 29 (0.498 g, 0.95 mmol), 2,4,6- triisopropylbenzenesulfonyl chloride (0.997 g, 3.3 mmol) and O N 4-dimethylaminopyridine (0.373 g, 3.05 mmol) in anhydrous acetonitrile (30 ml) was added triethylamine (420 l, 3.0 Si O N O O mmol). The reaction was allowed to stir under argon at room temperature overnight. TLC showed starting material was still present, so a solution of 2,4,6-triisopropylbenzenesulfonyl Si O chloride (0.579 g, 1.9 mmol) and 4-dimethylaminopyridine (0.177 g, 1.45 mmol) and triethylamine (400 l, 2.9 mmol) in 5 mL of anhydrous acetonitrile was cannulated into the reaction mixture. The reaction was again allowed to stir under argon at room temperature overnight. Concentrated ammonium hydroxide (50 ml) was added to the reaction flask and reaction was allowed to stir at room temperature for 24 h. Solvent was concentrated under reduced pressure resulting solution was extracted with dichloromethane. Solvent was removed under reduced pressure. Product was purified by flash column chromatography (5% methanol/dichloromethane). Product: white 1 foam (396 mg, 0.76 mmol, 80% yield). H NMR (400 MHz, CDCl3): 8.60 (broad, NH2, 1H), 8.06 (s, H-6, 1H), 7.46 (d, J = 2.0 Hz, H-5, 1H), 6.57 (d, J = 3.2, H-3, 1H), 6.48 (dd, J = 3.4 and 1.8 Hz, H-3, 1H), 6.30 (t, J = 6.4 Hz, H-1, 1H), 4.38–4.35 (m, H-3, 1H), 4.02–4.00 (m, H-5, 1H) 3.90–3.86 (m, H-4, 1H), 3.77–3.73 (m, H-4, 1H), 2.53–2.47 (m, H-2, 1H), 2.07– 2.00 (m, H-2, 1H), 0.88 (s, t-butyl, 9H), 0.80 (s, t-butyl, 9H), 0.07 (s, Me, 3H), 0.06 (s, Me, 13 3H), 0.01 (s, Me, 3H), 0.02 (s, Me, 3H); C NMR (100 MHz, CDCl3): 161.1, 153.3, 146.3, 142.6, 139.6, 111.6, 108.2, 99.3, 88.4, 86.9, 72.1, 62.8, 42.5, 25.8, 25.7, 18.3, 18.0, 4.6, + 4.9, 5.6; ESI-MS calculated for C25H44N3O5Si2 [M+H] 522.28, found 521.99 and for + C25H43N3NaO5Si2 [M+Na] 544.26, found 544.14.

113

3,5-tertbutyldimmethylsilyl-N-benzoyl-5-(fur-2-yl)-2- O deoxycytidine (31). To a solution of 30 (0.114 g, 0.22 mmol), and 4-dimethylaminopyridine (0.058 g, 0.47 HN mmol) in anhydrous dichloromethane (10 ml) was added benzoyl chloride (40 L, 0.33 mmol). The reaction was O N μ stirred at room temperature overnight. Methanol (1 mL) was added to quench the reaction. Solvent was removed Si O N O O under reduced pressure. The product was purified by gravity column chromatography (dichloromethane). Product: light yellow foam (0.133 g, 0.21 mmol, 96 % 1 Si O yield). H NMR (400 MHz, CDCl3): 8.31–8.28 (m, H-6 and Bz, 2H), 7.55–7.46 (m, Bz, 4H), 7.40 (d, J = 1.2 Hz, H-5, 1H), 7.35 (d, J = 3.2, H-3, 1H), 6.55 (dd, J = 3.0 and 1.6 Hz, H-3, 1H), 6.35 (td, J = 7.6 and 5.6 Hz, H-1, 1H), 4.46–4.45 (m, H-3, 1H), 4.07–4.05 (m, H-5, 1H) 3.93–3.90 (m, H- 4, 1H), 3.84–3.80 (m, H-4, 1H), 2.47–2.42 (m, H-2, 1H), 2.13–2.04 (m, H-2, 1H), 0.91 (s, t- butyl, 9H), 0.87 (s, t-butyl, 9H), 0.10 (s, Me, 3H), 0.10 (s, Me, 3H), 0.09 (s, Me, 3H), 0.07 (s, 13 Me, 3H); C NMR (100 MHz, CDCl3): 179.9, 156.3, 146.7, 146.0, 141.0, 137.1, 136.2, 132.5, 129.9, 128.2, 111.9, 110.7, 107.0, 88.6, 86.5, 72.6, 63.0, 41.9, 25.9, 25.8, 25.7, 18.3, + 18.0, 4.7, 4.9, 5.6, 5.7; ESI-MS calculated for C32H48N3O6Si2 [M+H] 626.31, found 625.96 + and for C32H47N3NaO6Si2 [M+Na] 648.29, found 648.09.

N-benzoyl-5-(fur-2-yl)-2-deoxycytidine (32). To a solution of O 31 (0.383 g, 0.61 mmol) in anhydrous tetrahydrofuran (6 ml) was added triethylamine trihydrogenfluoride (210 μL, 1.28 mmol). HN The reaction was stirred at room temperature for 8 h. Starting material still seen by tlc, so more triethylamine O N trihydrogenfluoride (150 μL, 0.92 mmol) was added to the reaction. The reaction solution was run through a silica column, HO N O O then flushed with dichloromethane followed by 5% methanol/dichloromethane. A yellow product spot was collected. Product: yellow foam (0.242 g, 0.61 mmol, 99 % yield). 1H NMR

HO (400 MHz, CD3OD): 8.71 (s, H-6, 1H), 8.22 (d, J = 7.2, Bz, 2H), 7.56–7.43 (m, Bz and H-5, 3H), 7.25 (d, J = 2.4, H-3, 1H), 6.51 (s, H-4, 1H), 6.33 (t, J = 6.2 Hz, H-1, 1H), 4.47–4.46 (m, H-3, 1H), 4.02–4.00 (m, H-5, 1H) 3.89–3.78 (m, H-4, 2H), 13 2.44–2.34 (m, H-2, 2H); C NMR (100 MHz, CD3OD): 146.4, 141.7, 132.5, 129.5 128.2, + 111.4, 88.2, 86.8, 70.7, 61.3, 40.8; ESI-MS calculated for C20H19N3NaO6 [M+Na] 420.12, + found 420.03 and for C20H19N3KO6 [M+K] 436.09, found 436.09; HR-MS calculated for C20H19N3O6 397.1268, found 397.1261.

114

5-dimethoxytrityl-N-benzoyl-5-(fur-2-yl)-2- O deoxycytidine (33). To a solution of 32 (0.242 g, O 0.61 mmol), 4-dimethylaminopyridine (8.5 mg, 0.07 HN mmol) and DMT-Cl (0.270 g, 0.8 mmol) in anhydrous pyridine (2 ml) was added triethylamine O N (75 l). The reaction was stirred at room temperature under argon for 6 h and evaporated O N O O under reduced pressure. The product was purified by flash column chromatography (1% methanol, 1% triethylamine, 98% dichloromethane). Product: light HO yellow foam (282 mg, 0.4 mmol, 66% yield). 1H NMR (400 MHz, DMSO-d ): ; 13C NMR (100 MHz, O 6 CDCl3): 8.45 (s, H-6, 1H), 8.30 (d, J = 8.0, Bz, 2H), 7.55–7.12 (m, DMT, Bz, H-5 and H-3, 14H), 6.78–6.71 (m, DMT, 4H), 6.4–6.36 (m, H-

4 and H-1, 2H), 4.48– 4.50 (m, H-3, 1H), 4.16–4.08 (m, H-4, 1H), 3.74 (s, DMT-OCH3, 6H), 3.56–3.53 (m, H-5, 1H), 3.36–3.30 (m, H-5, 1H), 2.60–2.54 (m, H-2, 1H), 2.36–2.29 (m, H- 13 2, 1H); C NMR (100 MHz, CDCl3): 180.3, 158.7, 148.9, 147.2, 147.1, 145.8, 144.7, 141.5, 137.3, 135.9, 135.8, 132.8, 130.3, 130.1, 128.8, 128.5, 128.3, 128.1, 127.2, 113.4, 111.7, 110.8, 107.6, 106.8, 86.8, 86.4, 72.2, 64.7, 63.6, 55.4, 41.8; ESI-MS calculated for + C41H37N3NaO8 [M+Na] 722.24, found 722.16 and for C41H37N3KO8 738.22, found 738.07. HR- MS calculated for C41H37N3O8 699.2575, found 699.2556.

O 3-2-cyanoethyldiisopropylphosphoramidite-5- O dimethoxytrityl-N-benzoyl-5-(fur-2-yl)-2- HN deoxycytidine (34). 33 was taken up in anhydrous acetonitrile (5 mL) and solvent was removed under O N reduced pressure (3x). To a solution cold solution (0°C) of 33 (67 mg, 0.95 mmol) and O N O diisopropylethylamine (200 l, 1.14 mmol) in O anhydrous 1,2-dichloroethane (0.65 ml) was added 2-cyanoethyldiisopropylchloro phosphorodiamidite (31 l, 0.14 mmol) dropwise. The solution was O PN(iPr) allowed to warm to room temperature and stirred for O 2 2 h. Solvent was removed under reduced pressure. O The product was purified by gravity column NC chromatography (1% Et3N in dichloromethane). Resulting solid precipitated from a dichloromethane solution by adding hexanes. Product: light 1 yellow solid (60 mg, 0.67 mmol, 70% yield) H NMR (400 MHz, CDCl3): 8.51 and 8.46 (s, H- 6, 1H), 8.30 (d, J = 9.2, Bz, 2H), 7.58–7.18 (m, 13H), 6.77 (d, J = 10.8, DMT, 4H), 6.62 (d, J = 14.8, 1H), 6.43–6.34 (m, H-4 and H-1, 2H), 4.57– 4.54 (m, H-3, 1H), 4.27–4.23 (m, H-4,

1H), 3.75 and 3.74 (s, DMT-OCH3, 6H), 3.63–3.49 (m, isopropyl and cyanoethyl, 4H), 3.31– 3.28 (m, H-5, 2H), 2.73–2.29 (m, H-2 and cyanoethyl, 4H), 1.29–1.04 (m, isopropyl 12H); 13C

NMR (100 MHz, CDCl3): 180.2, 158.5, 156.2, 146.7, 145.5, 144.4, 144.3, 141.2, 137.1, 135.6, 135.5, 135.4, 135.3, 132.5, 130.1, 130.0, 129.8, 128.2, 128.1, 128.0, 127.8, 126.9, 117.5, 117.3, 113.0, 111.3, 110.5, 107.4, 86.7, 86.2, 86.0, 85.9, 85.8, 73.8, 73.6, 73.4, 73.1, 63.0, 62.8, 58.4, 58.2, 58.1, 57.9, 55.2, 55.1, 43.3, 43.2, 43.0, 24.5, 24.4, 24.3, 20.4, 20.1; 31P

NMR (162 MHz, CDCl3, referenced to H3PO4): 149.0, 148.2; ESI-MS calculated for + C50H55N5O9P [M+H] 900.37, found 899.95

115

3,5-di-O-acetyl-5-(thiophen-2-yl)-2-deoxyuridine (39). To a O solution of 2 (3.2 g, 10.3 mmol) in anhydrous pyridine (45 ml) was added acetic anhydride (2.2 mL, 2.4 g, 23.3 mmol). The reaction S NH was allowed to stir under argon at room temperature overnight. O Solvent removed under reduced pressure. Resulting oil taken up N O O in dichloromethane (400ml) and washed with 1M HCl (3x 100 ml) O and water (100 ml). The organic layer was dried over sodium sulfate and the solvent removed under reduced pressure. O 1 O Product: light orange foam (3.88 g, 9.8 mmol, 96 % yield). H NMR (400 MHz, CDCl3): 9.60 (s, NH, 1H), 7.84 (s, H-6, 1H), 7.40 (d, J = 3.6 Hz, H-5, 1H), 7.26 (m, H-3, 1H), 7.02 (t, J = 4.4 Hz, H-3, 1H), 6.34 (td, J = 8.4 and 5.6 Hz, H-1, 1H), 5.23 (d, J = 6.0 Hz, H-3, 1H), 4.43– 4.30 (m, H-5 and H-4, 3H), 2.59–2.54 (m, H-2, 1H), 2.23–2.15 (m, H-2, 1H), 2.11 (s, Ac, 13 3H), 2.01 (s, Ac, 3H); C NMR (100 MHz, CD3Cl): 170.2 (Ac carbonyl), 170.0 (Ac carbonyl), 160.7 (C-4), 149.2 (C-2), 133.7 (C-2), 133.1 (C-5), 127.0 (C-6), 125.4 (C-4), 124.7 (C-3), 110.6(C-5), 85.5 (C-4), 82.6 (C-1), 74.2 (C-3), 64.0 (C-5), 38.2 (C-2), 21.0 (Ac methyl), + 20.9 (Ac methyl); ESI-MS calculated for C17H18N2NaO7S [M+Na] 417.07, found 416.93.

3,5-di-O-acetyl-5-(3,5-bromofur-2-yl)-2-deoxyuridine (40). Br To a cold (0°C) solution of 10 (0.191 g, 0.5 mmol) in anhydrous O 1,2-dichloroethane (5 ml) was added a carbon tetrachloride (1.5 Br O NH mL) bromine (60 L, 0.187 g, 1.17 mmol) solution over 10 min. via a dropping funnel. The reaction was allowed to stir under argon at 0°C for 1 h. Dichloromethane (15 mL) was added to the reaction O N O solution. The organic solution was washed with saturated sodium O O bicarbonate solution followed by water. The organic layer was dried over sodium sulfate and the solvent removed under reduced O pressure. Product purified via flash column chromatography (3% methanol/dichloromethane). Product: yellow oil (0.092 g, 0.17 1 O mmol, 34 % yield). H NMR (400 MHz, CDCl3): 9.33 (s, NH, 1H), 7.84 (s, H-6, 1H), 6.42 (s, H-4, 1H), 6.34 (t, J = 6.8 Hz, H-1, 1H), 5.22 (d, J = 6.0 Hz, H-3, 1H), 4.37–4.25 (m, H-5 and H-4, 3H), 2.59–2.54 (m, H-2, 1H), 2.23–2.15 (m, H-2, 1H), 2.10 + (s, Ac, 3H), 2.00 (s, Ac, 3H); ESI-MS calculated for C17H16Br2N2NaO8 [M+Na] 556.92, 558.92 and 560.91 found 556.75, 558.73 and 560.75.

116

Br 3,5-di-O-acetyl-5-(3,5-bromothiophen-2-yl)-2- O deoxyuridine (41). To a cold (0°C) solution of 39 (3.35 g, 8.5 Br mmol) in anhydrous 1,2-dichloroethane (90 ml) was added a S NH carbon tetrachloride (25 mL) bromine (1 mL, 3.1 g, 19.5 mmol) N O solution over 30 min. via a dropping funnel. The reaction was O allowed to stir under argon at 0°C for 1 h. Dichloromethane (500 O O mL) was added to the reaction solution. The organic solution was washed with saturated sodium bicarbonate solution (3) followed O by water. The organic layer was dried over sodium sulfate and the solvent removed under reduced pressure. Product purified via O flash column chromatography (2% methanol/dichloromethane). Product: light yellow foam (3.4 g, 6.1 mmol, 73 % yield). 1H NMR

(400 MHz, CDCl3): 9.10 (s, NH, 1H), 8.2 (s, H-6, 1H), 6.98 (s, H-4, 1H), 6.36 (td, J = 8.2 and 5.8 Hz, H-1, 1H), 5.25–5.23 (m, H-3, 1H), 4.34–4.27 (m, H-5 and H-4, 3H), 2.56–2.54 (m, H-2, 1H), 2.24–2.18 (m, H-2, 1H), 2.11 (s, Ac, 3H), 2.00 (s, Ac, 3H); ESI-MS calculated + for C17H16Br2N2NaO7S [M+Na] 572.89, 574.89 and 576.89 found 572.72, 574.74 and 576.70.

Br 3,5-di-O-acetyl-5-(3-bromofur-2-yl)-2-deoxyuridine (42). To O a suspension of 40 (0.135 g, 0.25 mmol) and zinc dust (0.041 g, 0.63 mmol) in anhydrous dimethylformamide (1 ml) was added O NH acetic acid (25 L, 0.44 mmol) and acetic anhydride (12 L, 0.13 N O mmol) successively. The reaction was heated to 90°C for 3.5 h. O Ether (5 mL) was added to the reaction solution. The organic O O solution was washed with saturated sodium bicarbonate solution followed by water. The organic layer was dried over sodium O sulfate and the solvent removed under reduced pressure. Product purified via flash column chromatography (1% O methanol/dichloromethane). Product: yellow film (0.048 g, 0.10 1 mmol, 42% yield). H NMR (400 MHz, CDCl3): 9.04 (s, NH, 1H), 7.87 (s, H-6, 1H), 7.44 (d, J = 2.0 Hz, H-5, 1H), 6.49 (d, J = 2.0 Hz, H-4, 1H), 6.36 (td, J = 8.2 and 5.48 Hz, H-1, 1H), 5.24–5.22 (m, H-3, 1H), 4.35–4.25 (m, H-5 and H-4, 3H), 2.58– 2.53 (m, H-2, 1H), 2.25–2.15 (m, H-2, 1H), 2.10 (s, Ac, 3H), 1.95 (s, Ac, 3H); 13C NMR (100

MHz, CD3Cl): 170.1, 169.9, 159.7, 149.2, 143.3, 143.0, 139.6, 114.8, 105.3, 99.5, 85.3, 82.4, + 74.0, 64.0, 38.3, 21.0, 20.7; ESI-MS calculated for C17H17BrN2NaO8 [M+Na] 479.01 and 481.00 found 478.85 and 480.83.

117

3,5-di-O-acetyl-5-(3,5-bromothiophen-2-yl)-2- Br deoxycytidine (43). To a solution of 41 (0.312 g, 0.56 mmol) and NH2 Br 1,2,4triazole (0.465 g, 6.7 mmol) in anhydrous pyridine (25 ml) S N was added phosphorus oxychloride (151 l, 1.6 mmol). The reaction was allowed to stir under argon at room temperature O N O overnight. The reaction was cooled to 0°C and concentrated O O ammonium hydroxide (3 mL) was added dropwise to the reaction flask. The reaction was allowed to stir at 0°C for 15 min, whereby the solvent removed under reduced pressure. Resulting solid O taken up in dichloromethane and washed with 1M HCl (3 x) and water. The resulting organic layer was dried over anhydrous O sodium sulfate. Solvent was removed under reduced pressure. Product purified via gravity column chromatography (5% methanol/dichloromethane). Product: 1 light brown solid (0.193 g, 0.35 mmol, 62% yield). H NMR (400 MHz, CDCl3): 8.64 (broad, NH2, 1H), 7.70 (s, H-6, 1H), 7.06 (s, H-4, 1H), 6.24 (td, J = 7.6 and 6.0 Hz, H-1, 1H), 5.64 (broad, NH2, 1H), 5.18 (d, J = 6.4 Hz, H-3, 1H), 4.32–4.26 (m, H-5 and H-4, 3H), 2.72–2.66 (m, H-2, 1H), 2.16–2.08 (m, H-2, 1H), 2.09 (s, Ac, 3H), 1.93 (s, Ac, 3H); 13C NMR (100 MHz,

CDCl3): 170.3, 170.1, 162.9, 154.1, 142.2, 133.0, 129.8, 114.6, 112.8, 98.7, 86.3, 82.5, 74.1, + 63.7, 38.9, 20.9, 20.5; ESI-MS calculated for C17H18Br2N3O6S [M+H] 549.93, 551.93, and 553.92 found 549.68, 551.63 and 553.66.

O 3,5-di-O-acetyl-N-benzoyl-5-(3,5-bromothiophen-2-yl)- Br 2-deoxycytidine (44). To a solution of 43 (0.089 g, 0.16 HN mmol), and 4-dimethylaminopyridine (0.044 g, 0.36 mmol) in Br anhydrous dichloromethane (8 ml) was added benzoyl S N chloride (28 μL, 0.24 mmol). The reaction was stirred at room O temperature overnight. Solvent was removed under reduced O N O pressure. The product was purified by gravity column O chromatography (dichloromethane). Product: light yellow solid 1 (0.059 g, 0.09 mmol, 55 % yield). H NMR (400 MHz, CDCl3): O 8.26–8.23 7.70 (m, Bz and H-6, 3H), 7.63–7.53 (m, Bz, 1H), 7.5–7.45 (m, Bz, 2H), 7.05 (s, H-4, 1H), 6.40 (td, J = O 8.0 and 5.2 Hz, H-1, 1H), 5.27 (d, J = 6.0 Hz, H-3, 1H), 4.3– 4.29 (m, H-5 and H-4, 3H), 2.67–2.62 (m, H-2, 1H), 2.29– 13 2.22 (m, H-2, 1H), 2.12 (s, Ac, 3H), 1.96 (s, Ac, 3H); C NMR (100 MHz, CD3Cl): 179.6, 170.3, 170.1, 157.2, 146.7, 140.7, 138.5, 136.3, 132.9, 131.9, 130.2, 128.3, 127.4, 115.3,

108.8, 107.7, 85.8, 82.6, 73.9, 63.8, 38.4, 20.9, 20.6; ESI-MS calculated for C17H19N2O8 + + [M+H] 379.11, found 378.84 and for C24H21Br2N3NaO7S [M+Na] 675.94, 677.94 and 679.93, found 675.78, 677.78 and 679.75. 118

A.3 – Solvent polarity measurements UV-Vis experiments were carried out at ambient temperature (unless otherwise noted) in a micro fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Hewlett Packard 8453 or 8452A diode array spectrometer. Steady State emission experiments were carried out at ambient temperature (unless otherwise noted) in a micro fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Perkin Elmer LS 50B luminescence spectrometer or a Horiba fluoromax-3 luminescence spectrometer. All nucleoside samples were measured at the noted concentration (1% DMSO) in the appropriate spectroscopic grade solvent. The relationship

between emission energy and microenvironment solvent polarity [ET(30)] was determined for nucleoside 1. All emission spectra were determined at 10 M with 1% DMSO in five different solvents (buffer, methanol, acetonitrile, dichloromethane and ether or ethyl acetate). Emission 1 spectra were converted from wavelength (nm) to wavenumbers (cm ). ET(30) values for each solvent mixture were experimentally determined using Reichardts salt.18

A.4 – Quantum efficiency determination Quantum yields were determined using anthracene as a standard15 for nucleosides 1, 2 and 4. Quantum yield of nucleoside 3 was determined with respect to nucleoside 1. Quantum yields of all nucleosides (1–4, and 7–9) were determined using the following equation:

2 F(x) = (As/Ax) (Fx/Fs) (x/s) F(s) (1)

Where s is the standard, x is the unknown, A is the absorbance at excitation wavelength, F is the area under the emission curve, is the refractive index of the solvent and F is the quantum yield77

A.5 – Temperature dependent nucleoside emission spectra Temperature dependent fluorescence experiments were carried out at 10 degree intervals from 25 to 75 °C in a micro fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) using a temperature controlled cell holder connected to an external water-based temperature control unit on either a Perkin Elmer LS 50B luminescence spectrometer or a Horiba fluoromax-3 luminescence spectrometer. Nucleoside samples were measured at 10 M (1 and 7) and 1 M (8 and 9) in water (1% DMSO). Spectra at the appropriate temperature were recorded after allowing the sample to equilibrate for 14 min.

A.6 – Oligonucleotide Synthesis and Purification. Unmodified oligonucleotides were purchased from Intergraded DNA Technologies (Coralville, Iowa) and purified by PAGE as described below. The THF residue in oligonucleotide 21 has the following structure:

O O

119

Modified oligonucleotides were synthesized on a 1.0 mole scale (500 Å CPG column) using a Biosearch Cyclone Plus DNA synthesizer. Nucleoside 1 was site specifically incorporated into the oligonucleotides by trityl-off synthesis of the base oligonucleotide, followed by manual coupling of phosphoramidite 14. Typically, the modified phosphoramidite was dissolved in 100L of anhydrous acetonitrile to give a final concentration of 0.1M. The phosphoramidite solution was pushed into the CPG column via syringe and then 200 L of 0.45M 1H-tetrazole was pushed into the other end of the column via syringe. Coupling reactions, performed twice, were allowed to proceed for 5 minutes (99% optimized coupling efficiency) and were subsequently followed by standard oxidation and capping steps. The rest of the oligonucleotide was synthesized via the standard trityl-off procedure. Upon completion of the oligonucleotide synthesis, the CPG column was treated with 3 mL of 30% aqueous ammonium hydroxide for 2 h at room temperature, mixing via syringe every 1 h. The resulting solution was removed and the CPG column was treated with 1 mL of 30% aqueous ammonium hydroxide at room temperature for 15 min, mixing via syringe every 5 min. The resulting aqueous ammonium hydroxide solutions were consolidated and stored at room temperature for 24 h. The aqueous ammonium hydroxide solutions were freeze-dried and purified by 20% polyacrylamide gel electrophoresis. The oligonucleotide was visualized by UV shadowing, bands were excised from the gel, and extracted with 0.5M sodium aceate buffer overnight. The resulting solution was filtered (Bio Rad poly-prep chromatography column) and desalted using a Sep-Pak cartridge (Waters Corporation, MA). The following 260nm extinction coefficients were used to determine concentration of oligonucleotides: dG = 11,700, dC = 7,300, dA = 15,400, dT = 8,800, and 1 = 13,000.

A.7 – MALDI-TOF of control and modified oligonucleotides The MW of each control and modified oligonucleotide was determined via MALDI-TOF MS. 1 L of a 200 M stock solution of each oligonucleotide was combined with 1 M ammonium citrate buffer (PE Biosystems), 1 L of a 75 M DNA standard (5-GCTGAATACATAAGACG - 3) and 6 L of saturated 3-hydroxypiccolinic acid. The samples were desalted with an ion- exchange resin (PE Biosystems) and spotted onto a gold-coated MALDI plate where they were dried on a 55ºC heat block. The resulting spectra were calibrated relative to the +1 and +2 ions of the internal DNA standard, thus the observed oligonucleotides should have a resolution of ±2 mass units. Modified oligonucleotide 15 was sequenced via onplate digestion (Sequazyme Oligonucleotide Sequencing Kit - Applied Biosystems). 2 L of water and 1 L of a 200 M stock solution of oligonucleotide 5 were combined and spotted eight times onto a gold-coated MALDI plate. 2 L of SVP dilutions (1 L of ammonium citrate buffer and 1 L of SVP dilution) were added to the first five wells and 2 L of BSP dilutions (1 L of BSP reaction buffer and 1 L of BSP dilution) were added to the remaining three wells. The MALDI plate was incubated in a humidity chamber (Applied Biosystems) at 37ºC for 20 min. 7L of saturated 3-hydroxypiccolinic acid was added to each well. The samples were desalted with an ion-exchange resin (PE Biosystems) and then respotted onto the same wells of the gold- coated MALDI plate where they were subsequently dried on a 55ºC heat block. All MALDI- TOF spectra were collected on a PE Biosystems Voyager-DE STR MALDI-TOF spectrometer in positive-ion, delayed-extraction mode. The following nucleoside masses were used to confirm the sequence: dG = 329.053, dC = 289.046, dA = 313.058, dT = 304.046, and the modified furanyl dT = 356.041.

120

A.8 – Enzymatic digestion of furanyl dT oligonucleotide 15 and furanyl dC oligonucleotide 35

A mixture of 10 L of 1.0 M MgCl2, 10 L of 10x alkaline phosphatase buffer, 3 L of alkaline phosphatase, 3 L of phosphodiesterase I, 4 L nuclease P1 and 70 L of water was added to a tube containing 2 nmoles of the appropriate oligonucleotide. The mixture was left at 37 °C overnight. The nucleoside mixture was analyzed by reversed-phase analytical HPLC using a Vydac C18 column (0.46 cm 25 cm, 5 m TP silica). Mobile phase A: 100 mM triethyl ammonium acetate buffer (pH 7.0). Mobile phase B: acetonitrile. Flow rate: 1 mL/min. Gradient: 0-20% acetonitrile in 15 min and 20-100% in 5 min. The mixtures were monitored by UV at 260 nm (15 and 35), 304 nm (35) and 316 nm (15).

A.9 – Thermal denaturation studies All hybridizations and UV melting experiments were carried out in 100 M sodium chloride, 10 M sodium phosphate buffer at pH 7.0 using a Beckman-Coulter DU® 640 spectrometer with a high performance temperature controller and micro auto six holder. Samples (double- stranded concentrations: 1 M) were heated to 95 ºC for 5 min and cooled to room temperature over 2-3 h prior to measurements. Samples were placed in a stoppered 1.0-cm path length cell and a background spectra (buffer) was subtracted from each sample. Denaturation runs were performed between 26 and 75 ºC at a scan rate of 0.5 ºC min-1 with optical monitoring every minute at 260 nm. Beckman-Coulter software (provided with Tm Analysis Accessory for DU® Series 600 Spectrometers) determined the melting temperatures utilizing the first derivative from the melting profile. All reported standard deviations were calculated using STDEVP in Microsoft Excel.

A.10 - Oligonucleotide Spectroscopy Studies UV-Vis spectra were recorded on a Hewlett Packard 8453 or 8452A diode array spectrometer in a 400 L quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) at ambient temperature. Steady state emission experiments were carried out at 23°C in a 400 L quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on either a Perkin Elmer LS 50B luminescence spectrometer or a Jobin Yvon Horiba FluoroMax-3 luminescence spectrometer. DNA samples were hybridized by heating to 95 ºC for 5 min and subsequently allowed to cool to room temperature over 2 h prior to measurements. All DNA samples, unless otherwise noted, were measured at 1 M in in 100 M sodium chloride, 10 M sodium phosphate buffer at pH 7.0. A blank (buffer) was subtracted for each DNA fluorescence spectra unless otherwise noted. Thermal denaturation experiments monitored via fluorescence spectroscopy were recorded after allowing the sample to equilibrate for 14 min.

A.11 – Enzymatic incorporation of furanyl dTTP analogue 26 Single-strand DNA templates were annealed to the primer sequence in TE buffer (10 mM Tris-HCl, 1 mM EDTA, 100 mM NaCl, pH 7.8) by heating a 1:1 mixture (5 M) at 90 °C for 5 min and cooling the solution slowly to room temperature. Typical polymerase reactions were

performed in 50 mM Tris-HCl (pH 7.2), 10 mM MgSO4, 0.1 mM DTT containing 0.5 M annealed template (IDT), 100 M dGTP (Fermentas), 100 M dCTP (Fermentas), 300 M 26, 5.0 Ci -32P dATP, and 3.0 U/L of Klenow fragment, exo minus (Promega) in a total volume of 10 L. Reactions were typically run for 5, 15 or 30 min at 37 °C and quenched by adding 121

10 L of loading buffer (7 M urea in 10 mM Tris-HCl, 100 mM EDTA, pH 8 and 0.05% bromophenol blue). 5 L of the quenched sample was loaded onto an analytical 20% denaturing polyacrylamide gel (1,000 volts for 5h). The products on the gel were analyzed using a phosphorimager. Transcription efficiencies are reported with respect to transcription in the presence of natural nucleotides.

2.30 – References

1. Albinsson, B.; Norden, B. "Electronic-Transition Moment Directions and Identification of Low-Energy N-Pi-Asterisk States in Weakly Perturbed Purine Chromophores" J. Am. Chem. Soc. 1993, 115, 223–231.

2. Andreasson, J.; Holmén, A.; Albinsson, B. "The photophysical properties of the adenine chromophore" J. Phys. Chem. B 1999, 103, 9782–9789.

3. Callis, P. R. "Electronic States and Luminescence of Nucleic-Acid Systems" Annu Rev Phys Chem 1983, 34, 329–357.

4. Daniels, M.; Hauswirth, W. "Fluorescence of the purine and pyrimidine bases of the nucleic acids in neutral aqueous solution at 300 degrees K" Science 1971, 171, 675– 677.

5. Morgan, J. P.; Daniels, M. "Excited-States of DNA and Its Components at Room- Temperature .4. Spectral, Polarization and Quantum Yield Studies of Emissions from Cpc and Poly-Rc" Photochem. Photobiol. 1980, 31, 207–213.

6. Nir, E.; Kleinermanns, K.; Grace, L.; de Vries, M. S. "On the photochemistry of purine nucleobases" J. Phys. Chem. A 2001, 105, 5106–5110.

7. Onidas, D.; Markovitsi, D.; Marguet, S.; Sharonov, A.; Gustavsson, T. "Fluorescence properties of DNA nucleosides and nucleotides: A refined steady-state and femtosecond investigation" J. Phys. Chem. B 2002, 106, 11367–11374.

8. Pecourt, J. M. L.; Peon, J.; Kohler, B. "Ultrafast internal conversion of electronically excited RNA and DNA nucleosides in water" J. Am. Chem. Soc. 2000, 122, 9348– 9349.

9. Wilson, R. W.; Callis, P. R. "Fluorescent Tautomers and the Apparent Photophysics of Adenine and Guanine" Photochem. Photobiol. 1980, 31, 323–327.

10. Rist, M. J.; Marino, J. P. "Fluorescent nucleotide base analogs as probes of nucleic acid structure, dynamics and interactions" Curr. Org. Chem. 2002, 6, 775–793.

11. Gutierrez, A. J.; Terhorst, T. J.; Matteucci, M. D.; Froehler, B. C. "5-Heteroaryl-2'- Deoxyuridine Analogs - Synthesis and Incorporation into High-Affinity Oligonucleotides" J. Am. Chem. Soc. 1994, 116, 5540–5544.

12. Wigerinck, P.; Pannecouque, C.; Snoeck, R.; Claes, P.; Declercq, E.; Herdewijn, P. "5- (5-Bromothien-2-Yl)-2'-Deoxyuridine and 5-(5-Chlorothien-2-Yl)-2'-Deoxyuridine Are 122

Equipotent to (E)-5-(2-Bromovinyl)-2'-Deoxyuridine in the Inhibition of Herpes-Simplex Virus Type-I Replication" J. Med. Chem. 1991, 34, 2383–2389.

13. Pomerantz, M.; Amarasekara, A. S.; Dias, H. V. R. "Synthesis and solid-state structures of dimethyl 2,2 '-bithiophenedicarboxylates" J. Org. Chem. 2002, 67, 6931– 6937.

14. Young, D. W.; Tollin, P.; Wilson, H. R. "Crystal and Molecular Structure of Thymidine" Acta Crystallographica Section B-Structural Crystallography and Crystal Chemistry 1969, B 25, 1423-&.

15. Berlman, I. B. Handbook of fluorescence spectra of aromatic molecules; 2d ed.; Academic Press: New York,, 1971.

16. Srivatsan, S. G.; Tor, Y. "Fluorescent pyrimidine ribonucleotide: Synthesis, enzymatic incorporation, and utilization" J. Am. Chem. Soc. 2007, 129, 2044-2053.

17. Srivatsan, S. G.; Tor, Y. "Using an emissive uridine analogue for assembling fluorescent HIV-1 TAR constructs" Tetrahedron 2007, 63, 3601–3607.

18. Reichardt, C. "Solvatochromic Dyes as Solvent Polarity Indicators" Chem. Rev. 1994, 94, 2319–2358.

19. Evans, K.; Xu, D.; Kim, Y.; Nordlund, T. M. "2-Aminopurine optical spectra: solvent, pentose ring, and DNA helix melting dependence" J. Fluoresc. 1992, 2, 209–216.

20. Smagowicz, J.; Wierzchowski, K. L. "Lowest excited states of 2-aminopurine" J. Luminesc. 1974, 8, 210-232.

21. Ward, D. C.; Reich, E.; Stryer, L. "Fluorescence studies of nucleotides and polynucleotides. I. Formycin, 2-aminopurine riboside, 2,6-diaminopurine riboside, and their derivatives" J. Biol. Chem. 1969, 244, 1228–1237.

22. Ward, D. C.; Reich, E.; Stryer, L. "Fluorescence studies of nucleotides and polynucleotides. I. Formycin, 2-aminopurine riboside, 2,6-diaminopurine riboside, and their derivatives" J. Biol. Chem. 1969, 244, 1228–1237.

23. Persson, T.; Gronowitz, S.; Hornfeldt, A. B.; Johansson, N. G. "Synthesis and Antiviral Effects of 2-Heteroaryl Substituted Adenosine and 8-Heteroaryl Substituted Guanosine Derivatives" Bioorg. Med. Chem. 1995, 3, 1377-1382.

24. Adamczyk, M.; Mattingly, P. G.; Moore, J. A. "O-(fluoresceinylmethyl)hydroxylamine (OFMHA): a fluorescent reagent for detection of damaged nucleic acids" Bioorg. Med. Chem. Lett. 1998, 8, 3599–3602.

25. Asaeda, A.; Ide, H.; Terato, H.; Takamori, Y.; Kubo, K. "Highly sensitive assay of DNA abasic sites in mammalian cells optimization of the aldehyde reactive probe method" Anal. Chim. Acta 1998, 365, 35–41. 123

26. Atamna, H.; Cheung, I.; Ames, B. N. "A method for detecting abasic sites in living cells: age-dependent changes in base excision repair" Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 686–691.

27. Belmont, P.; Boudali, A.; Constant, J. F.; Demeunynck, M.; Fkyerat, A.; Michon, P.; Serratrice, G.; Lhomme, J. "Efficient and versatile chemical tools for cleavage of abasic sites in DNA" New J. Chem. 1997, 21, 47–54.

28. Berthet, N.; Boudali, A.; Constant, J. F.; Decout, J. L.; Demeunynck, M.; Fkyerat, A.; Garcia, J.; Laayoun, A.; Michon, P.; Lhomme, J. "Design of molecules that specifically recognize and cleave apurinic sites in DNA" J. Mol. Recognit. 1994, 7, 99–107.

29. Berthet, N.; Constant, J. F.; Demeunynck, M.; Michon, P.; Lhomme, J. "Search for DNA repair inhibitors: selective binding of nucleic bases-acridine conjugates to a DNA duplex containing an abasic site" J. Med. Chem. 1997, 40, 3346–3352.

30. Berthet, N.; Michon, J.; Lhomme, J.; Teulade-Fichou, M. P.; Vigneron, J. P.; Lehn, J. M. "Recognition of abasic sites in DNA by a cyclobisacridine molecule" Chem. Eur. J. 1999, 5, 3625–3630.

31. Boturyn, D.; Boudali, A.; Constant, J. F.; Defrancq, E.; Lhomme, J. "Synthesis of fluorescent probes for the detection of abasic sites in DNA" Tetrahedron 1997, 53, 5485–5492.

32. Boturyn, D.; Constant, J. F.; Defrancq, E.; Lhomme, J.; Barbin, A.; Wild, C. P. "A simple and sensitive method for in vitro quantitation of abasic sites in DNA" Chem. Res. Toxicol. 1999, 12, 476–482.

33. Boturyn, D.; Defrancq, E.; Ducros, V.; Fontaine, C.; Lhomme, J. "Quantitative one step derivatization of oligonucleotides by a fluorescent label through abasic site formation" Nucleosides Nucleotides 1997, 16, 2069–2077.

34. Bowman, K. J.; Pla, R. L.; Guichard, Y.; Farmer, P. B.; Jones, G. D. "Evaluation of phosphodiesterase I-based protocols for the detection of multiply damaged sites in DNA: the detection of abasic, oxidative and alkylative tandem damage in DNA oligonucleotides" Nucleic Acids Res. 2001, 29, E101.

35. Chakravarti, D.; Badawi, A. F.; Venugopal, D.; Meza, J. L.; Crandall, L. Z.; Rogan, E. G.; Cavalieri, E. L. "Improved measurement of dibenzo[a,l]pyrene-induced abasic sites by the aldehyde-reactive probe assay" Mutat. Res. 2005, 588, 158–165.

36. Chen, B. X.; Kubo, K.; Ide, H.; Erlanger, B. F.; Wallace, S. S.; Kow, Y. W. "Properties of a monoclonal antibody for the detection of abasic sites, a common DNA lesion" Mutat. Res. 1992, 273, 253–261.

37. Constant, J. F.; Fkyerat, A.; Demeunynck, M.; Laval, J.; O'Connor, T. R.; Lhomme, J. "Design of molecules which specifically cleave abasic sites in DNA" Anti-Cancer Drug Des. 1990, 5, 59–62. 124

38. Dolinnaya, N. G.; Jan, M. R.; Kawde, A.-N.; Oretskaya, T. S.; Tashlitsky, V. N.; Wang, J. "Electrochemical detection of abasic site-containing DNA" Electroanalysis 2006, 18, 399-404.

39. Fkyerat, A.; Demeunynck, M.; Constant, J. F.; Michon, P.; Lhomme, J. "A New Class of Artificial Nucleases That Recognize and Cleave Apurinic Sites in DNA with Great Selectivity and Efficiency" J. Am. Chem. Soc. 1993, 115, 9952–9959.

40. Georgakilas, A. G.; Bennett, P. V.; Sutherland, B. M. "High efficiency detection of bi- stranded abasic clusters in gamma-irradiated DNA by putrescine" Nucleic Acids Res. 2002, 30, 2800–2808.

41. Hirose, T.; Ohtani, T.; Muramatsu, H.; Tanaka, A. "Direct visualization of abasic sites on a single DNA molecule using fluorescence microscopy" Photochem. Photobiol. 2002, 76, 123–126.

42. Huang, W.; Morita, K.; Sankaran, N. B.; Nishizawa, S.; Teramae, N. "Electrochemical detection at low temperature for a specific nucleobase of target nucleic acids by an abasic site-containing DNA binding ligand" Electrochem. Commun. 2006, 8, 395–398.

43. Ide, H.; Akamatsu, K.; Kimura, Y.; Michiue, K.; Makino, K.; Asaeda, A.; Takamori, Y.; Kubo, K. "Synthesis and Damage Specificity of a Novel Probe for the Detection of Abasic Sites in DNA" Biochemistry 1993, 32, 8276–8283.

44. Ide, H.; Kurisu, S.; Tanaka, R.; Asaeda, A.; Kubo, K. "Detection of DNA damage by the aldehyde reactive probe (ARP) assay: Principles and its application" Recent Res. Devel. Nucleosides Nucleotides 2003, 1, 33–47.

45. Kim, J.; Muramatsu, H.; Lee, H.; Kawai, T. "Near-field optical imaging of abasic sites on a single DNA molecule" FEBS Lett. 2003, 555, 611–615.

46. Kubo, K.; Ide, H.; Wallace, S. S.; Kow, Y. W. "A novel, sensitive, and specific assay for abasic sites, the most commonly produced DNA lesion" Biochemistry 1992, 31, 3703–3708.

47. Lhomme, J.; Constant, J. F.; Demeunynck, M. "Abasic DNA structure, reactivity, and recognition" Biopolymers 1999, 52, 65–83.

48. Makrigiorgos, G. M.; Chakrabarti, S.; Mahmood, A. "Fluorescent labelling of abasic sites: a novel methodology to detect closely-spaced damage sites in DNA" Int. J. Radiat. Biol. 1998, 74, 99–109.

49. Martelli, A.; Jourdan, M.; Constant, J. F.; Demeunynck, M.; Dumy, P. "Photoreactive threading agent that specifically binds to abasic sites in DNA" Bioorg. Med. Chem. Lett. 2006, 16, 154–157.

50. Morita, K.; Sankaran, N. B.; Huang, W.; Seino, T.; Sato, Y.; Nishizawa, S.; Teramae, N. "Electrochemical SNPs detection using an abasic site-containing DNA on a gold electrode" Chem. Commun. 2006, 2376–2378. 125

51. Roberts, K. P.; Sobrino, J. A.; Payton, J.; Mason, L. B.; Turesky, R. J. "Determination of apurinic/apyrimidinic lesions in DNA with high-performance liquid chromatography and tandem mass spectrometry" Chem. Res. Toxicol. 2006, 19, 300–309.

52. Sato, K.; Greenberg, M. M. "Selective detection of 2-deoxyribonolactone in DNA" J Am Chem Soc 2005, 127, 2806–2807.

53. Sun, H. B.; Qian, L.; Yokota, H. "Detection of abasic sites on individual DNA molecules using atomic force microscopy" Anal. Chem. 2001, 73, 2229–2232.

54. Teulade-Fichou, M. P.; Vigneron, J. P.; Lehn, J. M.; Berthet, N.; Michon, J.; Garcia, J.; Jourdan, M.; Lhomme, J. "Specific recognition and stabilization of an abasic site- containing DNA duplex by a macrocyclic bisacridine" Nucleosides Nucleotides 1999, 18, 1351–1353.

55. Weinfeld, M.; Liuzzi, M.; Paterson, M. C. "Response of phage T4 polynucleotide kinase toward dinucleotides containing apurinic sites: design of a 32P-postlabeling assay for apurinic sites in DNA" Biochemistry 1990, 29, 1737–1743.

56. Yanagisawa, H.; Hirano, A.; Sugawara, M. "A dot-blot method for quantification of apurinic/apyrimidinic sites in DNA using an avidin plate and liposomes encapsulating a fluorescence dye" Anal. biochem. 2004, 332, 358–367.

57. Zhang, L. K.; Gross, M. L. "Location of abasic sites in oligodeoxynucleotides by tandem mass spectrometry and by a chemical cleavage initiated by an unusual reaction of the ODN with MALDI matrix" J. Am. Soc. Mass. Spectrom. 2002, 13, 1418–1426.

58. Zhang, L. K.; Rempel, D.; Gross, M. L. "Matrix-assisted laser desorption/ionization mass spectrometry for locating abasic sites and determining the rates of enzymatic hydrolysis of model oligodeoxynucleotides" Anal. Chem. 2001, 73, 3263–3273.

59. Zhao, C.; Dai, Q.; Seino, T.; Cui, Y.-Y.; Nishizawa, S.; Teramae, N. "Strong and selective binding of amiloride to thymine base opposite AP sites in DNA duplexes: simultaneous binding to DNA phosphate backbone" Chem. Commun. 2006, 1185– 1187.

60. Alarcon, K.; Demeunynck, M.; Lhomme, J.; Carrez, D.; Croisy, A. "Potentiation of BCNU cytotoxicity by molecules targeting abasic lesions in DNA" Bioorg. Med. Chem. 2001, 9, 1901–1910.

61. Belmont, P.; Alarcon, K.; Demeunynck, M.; Lhomme, J. "Synthesis of an imidazo[1,2- e]purine-acridine heterodimer for targeting abasic sites in DNA" Bioorg. Med. Chem. Lett. 1999, 9, 233–236.

62. Brotschi, C.; Häberli, A.; Leumann, C. J. "A Stable DNA Duplex Containing a Non- Hydrogen-Bonding and Non-Shape-Complementary Base Couple: Interstrand Stacking as the Stability Determining Factor" Angew. Chem. Int. Ed. Engl. 2001, 40, 3012–3014. 126

63. Fukui, K.; Morimoto, M.; Segawa, H.; Tanaka, K.; Shimidzu, T. "Synthesis and properties of an oligonucleotide modified with an acridine derivative at the artificial abasic site" Bioconjugate Chem. 1996, 7, 349–355.

64. Fukui, K.; Tanaka, K. "The acridine ring selectively intercalated into a DNA helix at various types of abasic sites: double strand formation and photophysical properties" Nucleic Acids Res. 1996, 24, 3962–3967.

65. Fukui, K. I., Kazunori; Shimidzu, T.; Tanaka, K. "Oligonucleotides covalently linked to an acridine at artificial abasic site: influence of linker length and the base-sequence" Tetrahedron Lett. 1996, 37, 4983–4986.

66. Langenegger, S. M.; Haener, R. "A simple, non-nucleosidic base surrogate increases the duplex stability of DNA containing an abasic site" Chem. Biodiversity 2004, 1, 259–264.

67. Langenegger, S. M.; Haener, R. "Remarkable stabilization of duplex DNA containing an abasic site by non-nucleosidic phenanthroline and pyrene building blocks" ChemBioChem 2005, 6, 848–851.

68. Matray, T. J.; Kool, E. T. "A specific partner for abasic damage in DNA" Nature 1999, 399, 704–708.

69. Singh, I.; Beuck, C.; Bhattacharya, A.; Hecker, W.; Parmar, V. S.; Weinhold, E.; Seitz, O. "Abasic site stabilization by aromatic DNA base surrogates: High-affinity binding to a baseflipping DNA-methyltransferase" Pure Appl. Chem. 2004, 76, 1563–1570.

70. Valis, L.; Amann, N.; Wagenknecht, H. A. "Detection of single base mismatches and abasic sites using phenanthridinium as an artificial DNA base and charge donor" Org. Biomol. Chem. 2005, 3, 36–38.

71. Yoshimoto, K.; Nishizawa, S.; Minagawa, M.; Teramae, N. "Use of abasic site- containing DNA strands for nucleobase recognition in water" J. Am. Chem. Soc. 2003, 125, 8982–8983.

72. Pompizi, I.; Häberli, A.; Leumann, C. J. "Oligodeoxynucleotides containing conformationally constrained abasic sites: a UV and fluorescence spectroscopic investigation on duplex stability and structure" Nucleic Acids Res. 2000, 28, 2702– 2708.

73. Rachofsky, E. L.; Seibert, E.; Stivers, J. T.; Osman, R.; Ross, J. B. "Conformation and dynamics of abasic sites in DNA investigated by time-resolved fluorescence of 2- aminopurine" Biochemistry 2001, 40, 957–967.

74. Stivers, J. T. "2-Aminopurine fluorescence studies of base stacking interactions at abasic sites in DNA: metal-ion and base sequence effects" Nucleic Acids Res. 1998, 26, 3837–3844.

75. Yamada, K.; Kubo, T.; Tokuyama, H.; Fukuyama, T. "A mild copper-mediated intramolecular amination of aryl halides" Synlett 2002, 231-234. 127

76. Song, J. J.; Yee, N. K. "A novel synthesis of 2-aryl-2H-indazoles via a palladium- catalyzed intramolecular amination reaction" Org. Lett. 2000, 2, 519-521.

77. Fery-Forgues, S.; Lavabre, D. "Are fluorescence quantum yields so tricky to measure? A demonstration using familiar stationery products" J. Chem. Educ. 1999, 76, 1260– 1264.

Chapter 3

Exploring the polarity of DNA groove

3.1 – Introduction

The discovery of the double helical structure of DNA in 1953 by Watson and Crick has led

to enormous insights into the chemistry and biology of nucleic acids, although there still

remains certain fundamental features of this magnificent macromolecular assembly whose

details have not been identified. DNA exists in various polymorphs that under specific

environmental conditions drastically change their size and shape, which in turn modulates

their function (see chapter 1). As described in chapter 1, B-DNA whose structure is heavily

hydrated.1-4 is the most dominant conformation under physiological conditions. The first

hydration shell, consisting of approximately 20 water molecules per base pair, is practically

unfreezable while the second layer is impermeable to ions.5-7 It is in the grooves of DNA

where key biological recognition events take place. The understanding of the weak molecular

forces that operate in these grooves8-11 can facilitate the design of low MW ligands that impact the structure and thus the biological function of these key biopolymers.12,13

Breslauer and coworkers made the first experimental investigation into the polarity of DNA

grooves.14 Their attractive supramolecular approach correlated the photophysical properties of a bisbenzimide probe (Hoechst 33258) in known polarity solvents to the read-out determined upon the tight binding of the probe to the minor groove of a poly[d(AT)•poly[d(AT)

B-DNA duplex (figure 3.4).15-18 A reference scale (figure 3.1) is typically created by

determining the Stokes shift of the isolated probe in known polarity solvents which allows for

the correlation from an unknown Stokes shift read-out (bound to DNA) through orientational

polarizability to a polarity value, namely dielectric constant in this case.

128 129

Figure 3.1. Visual description of the reference scale approach to determine the polarity in DNA grooves (adapted from19).

This approach relies on the Lippert-Mataga equation (figure 3.2) that describes a linear

relationship between the Stokes shift (AF or absem) of a chromophore as the polarity of

the surrounding environment changes, if only general solvent effects, namely polarity, are

present. An increase in the refractive index () of the solvent tends to stabilize both the

ground and excited states, while changes in dielectric constant () effects the excited state

(figure 3.3). Most organic chromophores have a lifetime that is ~1ns, substantially longer than

the time it takes for the surrounding solvent to accommodate the newly formed, and commonly

more polar, excited state. Therefore, a longer emission wavelength or lower energy emission

state is generally seen in more polar solvents that can better accommodate the polar excited

state, while apolar solvents lack this ability resulting in emissions at higher energies or shorter

wavelengths (figure 3.3).

130

æ - 2 - ö - 2 - = 2 ç 1 - n 1 ÷ ( E G ) + abs em ç ÷ const hc è 2 +1 2n2 +1ø a3

Figure 3.2. The Lippert-Mataga equation describing solvent influence on ground and excited 29-31 states where absem = Stokes shift, h = Plancks constant, c = speed of light, a = radius of the cavity occupied by the chromophore and E and G represent the dipole moment of the excited and ground state, respectively. In large parentheses is the orientational polarizability portion.

μ E Less Polar Solvent Solvent Relaxation (10-10s) More Polar Solvent

ν ν ν ν ν´́ -9 A F A F F (10 s) (10-15s)

μ G ∆F

Gas Phase In Solution

Figure 3.3. Jablonski diagram depicting the effects of solvent on absorption and emission energies.

This approach, also utilized by Ganesh,20-22 Kunugi,23 Majima24,25 and Saito26-28 is informative, but bears several predicaments that are in part responsible for the dramatically different estimates reported for the polarity of the major groove in nucleic acids (ranging from an of < 20 to 70). The following sections will discuss the multiple parameters that impact the determination of the polarity in a biocavity, specifically grooves of DNA and our scholarly advance to expand the current approach.

3.2 – Expression of polarity

Polarity, particularly when describing the environment in biocavities, is a multi-dimensional

term where the correlation to simple organic solvents is a difficult task. Thus, the 131

representative value by which one chooses to define polarity is of the utmost importance. The following terms are typically used for the description of biocavities: (a) Dielectric constant or relative permittivity, a parameter reflecting the attenuating ability of solvent molecules in response to an applied macroscopic electric field relative to vacuum ( = 1, by definition); (b)

Solvent orientational polarizability, f, a parameter that expresses the interaction of the dipole

moment of the fluorophore in the excited state with the electrical field induced by surrounding

solvent dipoles (figure 3.2). This value is the result of a mathematical equation that combines

the dielectric constant and the refractive indexes of a given solvent; (c) ET(30) value (given in kcal/mol), an experimentally determined value based on the spectral shift of the absorption maximum of the Reichardts dye 3 (see page 138) in a solvent or solvent mixture, thereby representing the solvent influence on the molecular level.

The polarity of biocavities, specifically DNA grooves, has typically utilized the dielectric constant as the polarity parameter, due to its presence in the Lippert-Mataga equation29-31

(figure 3.2). The experimentally determined Stokes shift is stated to be linear with the

orientational polarizability of the solvent, only for general solvent effects, which is in turn

composed of the refractive index and the dielectric constant. Although other solvent polarity

parameters have been described for many years, and even utilized in groove polarity

determination, it was only recently that the striking discrepancy between the dielectric constant

and the ET(30) value in probing the polarity of biocavities was definitively presented (for a

19 detailed discussion on the use of or ET(30) parameters ). ET(30) values provide an insight into the environment directly surrounding the probe, similar to the environment in question, rather than the dielectric constant, whose description is based upon the bulk of the solvent.

132

3.3 – Probe design

General concerns It is important to state that any probe placed within the cavity to be assessed inherently modifies the molecular architecture of the native environment and therefore taints the readout.

This dilemma should draw attention to the details involved in “accurately” determining the readout and not serve to discredit the approach in general.

Probe size and shape The pioneering studies of Breslauer and coworkers utilized a rather large chromophore that spanned 5 base pairs in the minor groove of a DNA duplex (figure 3.4). Therefore, the determination that the minor groove has a polarity of 20 D, similar to 60% dioxane in water solution, is undoubtedly drastically influenced by the disruption of chromophore (Hoechst

33258). 133

Figure 3.4. Crystal structure of Hoechst 33258 bound to a DNA duplex.32 Image created using Chimera.33

The studies that followed focused their efforts on utilizing fluorescent nucleosides as

probes, whose structure is smaller than that of Hoechst 33258. Ganesh, Majima and Saito all

utilized either the dansyl or dan chromophores (figure 3.5) with various linkers connecting

them to natural nucleobases. The dan and dansyl chromophores are larger in size than the

native nucleobases and the dimethyl functional group adds steric bulk to the rather flat

chromophore. Major destabilizing effects have been seen by the introduction of these bases

in comparison to the natural sequence (up to 12°C) presumably due to the large size of these

modified nucleosides.21 134

O O

R N 1 H R2 N N

PRODAN - Saito DNC - Saito

O N R3 N O H H S N N O R2 DAN - Majima amidodansyl - Ganesh

Figure 3.5. Structures of the chromophores used for groove polarity determination where R1 = dU/dC connected at position 5 or dA/dG connected at position 7, R2 = dU connected at position 5, and R3 = dC connected at the N4 position or dG connected at the N2 position.

Linker effects In some instances, very minimal perturbation to the global stability is achieved, although

this typically requires the use of long flexible linkers, e.g. 5-PRODAN-dU.28 All four

chromophores (figure 3.5) are connected via flexible linkers, whereby they can possibly

populate multiple conformers each sensing a different microenvironment. This is evident by

comparing the polarity of the major groove determined by the Prodan and DNC probes. The

polarity of the major groove in a DNA duplex with similar base composition near the

modification site was determined to be 70, ~90 v % water in dioxane with the DNC probe,

while the prodan probe, having a shorter, yet dramatically more flexible linker, determined a

polarity of ~40 kcal/mol-1 or >5 v% water in dioxane. Therefore, the use of a flexible linker

drastically affects the position of the probe and ultimately the read-out. The length of the linker

is also a considerable concern, no matter how rigid the linker is, if the length is very long, the

probe will be monitoring an environment far outside the groove in a DNA duplex. The probes

utilized by Ganesh, Saito and Majima have 3–5 atoms in their linker, resulting in a minimal

distance of ~3 Å, not including the probe itself. These distances are rather short, but

computational studies have shown that the polarity in a DNA groove drastically changes after 135

2–3 Å.11 A visual comparison clearly shows how the aminodansyl probe, one of the smallest

with the shortest linker, is drastically larger than the natural dT it is supposed to be mimicking

(figure 3.6). A top view of the duplex seen in figure 3.6b clearly reveals that the dansyl moiety

extends past the diameter of the helix itself. Clearly these probes, with their rather long and

flexible linkers are not probing the groove surface, but rather environment(s) in the bulk

solvent.

136

0 2 4 6 8 1010Å 0 2 4 6 8 1010Å

Figure 3.6. Models of DNA duplexes (blue) (a) from the side and (b) from the top showing the location and surface area (orange) of the amidodansyl group (right - green) in comparison to the methyl group of T (left - green).

137

Probe sensitivity The photophysical properties of the fluorescent probes are important for their ability to

appropriately determine the polarity of DNA grooves. While an isolated absorption for

selective excitation is important, the sensitivity or responsiveness of the probe is vital. All

probes are measured in known polarity solvents where their Stokes shifts are recorded and

plotted against the corresponding solvent polarity, ET(30) in this case. An appropriate

sensitivity comparison can be derived from the slope, namely the change in Stokes shift in

regards to a change in polarity. A larger slope expresses a larger Stokes shift change per

ET(30) value, resulting in a greater sensitivity. A comparison reveals the slope of Hoechst

33258, dansyl nucleoside and dan nucleoside have similar slopes while the sensitivity of the

PRODAN species is dramatically less (figure 3.7). It should be noted that attachment of known chromophores does affect their sensitivity to changes in polarity. For example, the

PRODAN monomer has a slope of ~140, compared to the nucleoside form of 63, while the opposite effect is seen with the dansyl, where the monomer is ~105 and the nucleoside is 154

(figure 3.5).19 138

9.5 Breslauer 11.5 Ganesh

9.0 11.0

8.5

-1 10.5 8.0

cm 3 slope = 157 slope = 154 R2 = 0.993 10.0 R2 = 0.969 50 52 54 56 58 60 62 52 54 56 58 60 62

9.5 Majima Saito 7.0 9.0

Stokes shifts / x 10 6.5

8.5 6.0

8.0 slope = 124 slope = 63 R2 = 0.974 5.5 R2 = 0.637 52 54 56 58 60 62 45 50 55 60 E (30) / kcal mol-1 T

Figure 3.7. Stokes shift verses microenvironment polarity (ET(30)) for Breslauer (Hoechst 33258), Ganesh (dansyl nucleoside), Majima (DAN nucleoside) and Saito (PRODAN nucleoside).

Our approach In light of the above discussion, we have defined the following requirements for a fluorescent probe to accurately investigate the polarity of a DNA groove. The probes (a) size and shape must be such that only the groove is examined; a linker, if used, must be as short and rigid as possible; b) presence must not hamper W-C base pairing or native helix formation; c) absorption maximum must allow for selective excitation; and d) fluorescence maximum must be sensitive to polarity changes while maintaining sufficient quantum yield under all conditions. To meet these requirements, we avoid the conjugation of large fluorophores but rather develop new emissive nucleoside analogues where a natural 139

nucleobase fragment is an integral electronic element of the chromophore. In this fashion, small and minimally invasive probes, capable of engaging in normal W-C base pairing within unaltered duplexes, are employed. We have set out to investigate the groove microenvironment in B-, A- and abasic-duplex DNA using the previously studied furanyl dT analogue 1 (see chapter 2).

The furanyl dT nucleoside analogue 1 nicely fulfils the criteria listed above. It is an isosteric nucleobase mimic of dT, that upon base pairing with A in a W-C fashion, forms stable duplexes whose stability is the same as unmodified control duplexes (table 3.3).34 The rigid modification at the 5-position projects toward the major groove with a well-defined trajectory, while probing the grooves inner surface (figures 3.9). The direct conjugation of the furanyl moiety to the pyrimidine core creates a biaryl chromophore, resulting in an isolated absorption band (316 nm) that is virtually insensitive to changes in polarity, while its emission spectra are significantly impacted by the surrounding environment.34-36 In addition to these attractive photophysical features, the furanyl dT analogue 1 is prepared in only one step from available precursors, and can be site-specifically incorporated into oligonucleotides using standard solid-phase phosphoramidite-based chemistry (see chapter 2).34-36 Acetylation of nucleoside 1

generated the more lipophilic analogue 2 that could be utilized for photophysical

measurements in more apolar solvents (figure 3.8).

O O O O O O NH NH NH HO N HO N O N O O O O O O O HO HO O dT 12

Figure 3.8. Furanyl thymidine analogue in the free hydroxyl 1 and diacetylated form 2.

140

0 2 4 6 8 1010Å 0 2 4 6 8 1010Å

Figure 3.9. Models of DNA duplexes (blue) (a) from the side and (b) from the top showing the location and surface area (orange) of the furanyl dT analogue 1 (right - green) in comparison to the methyl group of T (left - green).

141

3.4 – Polarity reference scale

Probing the polarity contained in a biocavity is always referenced to the photophysical

properties, often the Stokes shift, obtained with the same probe in solvents or solvent mixtures

of known polarity. Although previous researchers have utilized the Lippert-Mataga equation to

correlate Stokes shift with the dielectric constant, previous discussion has shown a better

description of polarity in this case is use of the ET(30) scale.

1.0 12 a

8 water 0.5

-1 dioxane 4

cm -1 0 0.0

3 i 1.0 12 PrOH b × / a.u. PL

/ 10

ε 8 0.5 4 MCH 0 250 300 350 400 450 500 550 600 λ / nm

Figure 3.10. Absorption (dashed lines) and emission (solid lines) spectra of (a) furanyl dT analogue 1 in dioxane (red) water (blue) mixtures (9.0 10-6 M) and (b) nucleoside 2 in MCH (red) iPrOH (blue) mixtures (1.1 10-5 M).

To generate an expanded polarity scale for the furanyl dT analogue, four series of

absorption and emission spectra (figure 3.10) were measured in different dioxane-water

mixtures with nucleoside 1 (table 3.1) and in MCH-iPrOH mixtures with the diacetylated

nucleoside 2 (table 3.2). In order to accurately determine the ground state absorption

maximum of the emissive nucleoside within the hybridized duplexes, the absorption spectrum

of a control, unmodified oligonucleotide, containing the same base composition, was

subtracted from the absorption of the modified duplex. Then from the absorption and emission 142

spectra, accurate Stokes shifts (absem) were calculated. It should be noted that most

studies calculate the Stokes shifts from the emission and excitation maxima and do not extract

the actual ground state absorption maximum of the probe within the specific duplex. The

polarity of the dioxane-water mixtures were determined using the standard Reichardt dye 3,

i while the more lipophilic version 4, was used for the MCH- PrOH mixtures. The ET(30) value for pure water and 90 v% water in dioxane cannot be determined experimentally with

Reichardts dye 3 due to solubility issues, so the value for pure water was taken from

37 Reichardts review. Only the averaged ET(30) values are listed in tables 3.1 and 3.2 because

of the minimal changes seen in their determination.

N N

O O

Figure 3.11. Reichardts dye 3 and the more lipophilic version 4.

143

–1 –1 Table 3.1. Photophysical data of 1 in dioxane-water mixtures, abs (cm ), em (cm ), Stokes –1 –1 shift (SS, cm ), standard deviation (sd) and ET(30) values (kcal mol ). vol. frac. Avg. duplo triplo quadruplo Avg. sd.

water ET(30) Abs Em SS Abs Em SS Abs Em SS Abs Em SS SS

0.0 36.4 31847 24510 7337 31847 24510 7337 31847 24510 7337 31847 24390 7457 7367 52 0.1 45.9 31646 23866 7780 31746 23810 7936 31746 23810 7936 31746 23753 7993 7911 79 0.2 48.3 31646 23419 8227 31746 23474 8272 31746 23474 8272 31646 23419 8227 8250 23 0.3 50.3 31646 23310 8336 31746 23256 8490 31646 23256 8390 31646 23095 8551 8442 84 0.4 51.6 31447 23041 8406 31646 23041 8605 31546 22989 8557 31546 22989 8557 8531 75 0.5 53.2 31646 22831 8815 31546 22831 8715 31546 22831 8715 31546 22831 8715 8740 43 0.6 55.0 31447 22676 8771 31546 22624 8922 31447 22727 8720 31546 22676 8870 8821 80 0.7 56.0 31447 22573 8874 31746 22573 9173 31546 22624 8922 31646 22573 9073 9011 119 0.8 57.5 31646 22523 9123 31546 22472 9074 31646 22523 9123 31546 22472 9074 9099 25 0.9 - 31646 22472 9174 31746 22422 9324 31746 22523 9223 31746 22422 9324 9261 65 1.0 63.1 31646 22422 9224 31746 22422 9324 31746 22422 9324 31746 22422 9324 9299 43

i –1 –1 Table 3.2. Photophysical data of 2 in MCH- PrOH mixtures, abs (cm ), em (cm ), Stokes –1 –1 shift (SS, cm ), standard deviation (sd) and ET(30) values (kcal mol ). vol. frac. Avg. duplo triplo quadruplo Avg. sd. i PrOH ET(30) Abs Em SS Abs Em SS Abs Em SS Abs Em SS SS 0.000 32.2 31646 24722 6924 31646 24814 6832 31646 24876 6770 31646 24814 6832 6840 55 0.001 33.8 31646 24752 6894 31646 24722 6924 31646 24752 6894 31646 24814 6832 6886 33 0.002 35.0 31746 24691 7055 31746 24722 7024 31646 24691 6955 31646 24722 6924 6990 52 0.003 35.6 31746 24631 7115 31746 24661 7085 31646 24570 7076 31646 24691 6955 7058 61 0.010 38.7 31746 24540 7206 31746 24540 7206 31746 24510 7236 31746 24510 7236 7221 15 0.050 40.8 31746 24420 7326 31746 24361 7385 31746 24331 7415 31646 24301 7345 7368 35 0.200 43.6 31746 24096 7650 31746 24067 7679 31746 23981 7765 31746 24038 7708 7701 43 0.300 44.6 31746 23952 7794 31746 23952 7794 31746 23981 7765 31746 23981 7765 7780 15 0.500 46.1 31746 23866 7880 31746 23781 7965 31746 23669 8077 31746 23810 7936 7965 72 0.800 47.1 31746 23641 8105 31746 23669 8077 31746 23641 8105 31746 23725 8021 8077 34 8168 23 1.000 48.4 31746 23585 8161 31746 23613 8133 31746 23557 8189 31746 23557 8189

The Stokes shifts and corresponding ET(30) values were plotted resulting in a polarity reference scale seen in figure 3.12, thereby revealing the overlap and consistency between the two very different solvent mixtures. The correlation appears to be linear, with the exception of two obvious outliers, namely the data points representing pure water and pure dioxane (circled in red). The amalgamation of both data sets (dioxane-water and MCH- iPrOH), after the removal of data points representing pure water and pure dioxane, resulted in an enhanced polarity reference scale allowing correlation of virtually any probe readout to a microenvironmental polarity (figure 3.13). 144

-1 9.0 cm 3 8.5

8.0

7.5 Stokes shift / x 10 7.0

35 40 45 50 55 60 E (30) / kcal mol-1 T Figure 3.12. Polarity reference scale: averaged Stokes shifts of isolated nucleoside 2 in MCH-iPrOH (filled circles) and nucleoside 1 in dioxane-water (open circles) vs. the corresponding ET(30) value. Pure dioxane and water data points are circled in red.

9.0 -1 cm

3 8.5

8.0

7.5 Stokes shift / x 10 7.0

35 40 45 50 55 E (30) / kcal mol-1 T Figure 3.13. Correlation between Stokes shifts and microscopic polarity (ET(30)) for nucleosides 1 and 2. Shown are the averaged data points (filled circles) including error bars and a linear fit (solid orange line).

145

3.5 – Polarity of DNA grooves using furanyl dT analogue

To probe the polarity of major grooves in nucleic acids, a non self-complementary

oligonucleotide 5, that contains the furanyl dT analogue 1 in a central position, was prepared

(see chapter 2 for experimental details). It was then hybridized to different single stranded

oligonucleotides, including a perfect complement 6, an identical RNA complement 7, and an

abasic site containing oligonucleotide 8 (figure 3.14).

5 5' – GCG – ATG – 1GT – AGC – G –3' 6 5' – CGC – TAC – ACA – TCG – C –3' 7 5'–r(CGC – UAC – ACA – UCG – C) –3' 8 5' – CGC – TAC – YCA – TCG – C –3' 9 5' – GCG – ATG – TGT – AGC – G –3'

Figure 3.14. Single stranded oligonucleotides used. Y stands for an abasic site mimic (THF residue).

Thermal denaturation experiments confirm that the emissive oligonucleotides are fully hybridized under the experimental conditions used for photophysical evaluation. The modified

DNA (6•5) and DNA/RNA (7•5) duplexes are indistinguishable from the unmodified versions

6•9 and 7•9 (figure 3.13 and Table 3.3). As previously seen and discussed in chapter 2, the

furanyl dT analogue 1 results in an enhanced stabilization (+ ~4°C) over its unmodified version

when it is paired with an abasic site mimic (8•5 vs. 8•9).

146

0.8

0.4 Normalized Absorption

30 50 70 Temperature / °C

Figure 3.15. Thermal denaturation plot of control duplexes 6•9 (closed red circles) 7•9 (closed blue squares), 8•9 (closed green triangles) and modified duplexes 6•5 (open red circles) 7•5 (open blue squares), 8•5 (open green triangles) all at 1.0 106 M.

fu Table 3.3. Tm values of duplexes where dT = 1 and AP = stable abasic site mimic (THF residue).

Tm from 9 / °C Duplex Central Base Pair Average Tm / °C (duplex – corresponding dT duplex) 6•9 dT•dA 55.8 0.5 --- 7•9 dT•rA 45.0 0.6 --- 8•9 dT•AP 43.8 0.8 --- 6•5 dTfu•dA 46.3 0.5 0.0 7•5 dTfu•rA 56.0 0.6 0.8 8•5 dTfu•AP 48.1 1.1 3.9

CD spectroscopy was utilized to confirm the presence of the expected duplex polymorphs

(figure 3.16). All modified duplexes show identical CD curves as compared to their unmodified

versions. The single stranded oligonucleotides (5 and 9) showed a CD signal denoting the

formation of a defined secondary structure, although denaturation measurements denoted no 147

two-state melting. The resulting double stranded oligonucleotides represent a perfect B-form duplex DNA (6•5), an A-form DNA/RNA hybrid duplex (7•5), and an abasic-containing B-form

DNA duplex (8•5), where the fluorescent nucleoside is placed opposite to the defect position.

The B-form polymorph is identified by a cotton effect where the negative peak is at ~255nm,

the positive at ~280 nm with the crossing point at ~265 nm. The A-polymorph however, is

identified by an almost exclusive positive signal seen at ~275 nm and a weak negative band

below 250 nm. The isolated absorption band of the furanyl dT analogue 1 (316 nm) results in

a weak (26:1 ratio of native to modified) negative band at ~320 nm that can be seen in the

DNA B-form (6•5) and DNA/RNA hybrid A-form (7•5) duplexes while is absent from the single

stranded (5) and abasic duplex spectra (8•5). The implications of these observations are

unclear, although it does suggest the structural orientation of furanyl dT analogue 1, when

paired across from an abasic site is different than when paired with its complement dA.

148

5 a 8 4

2 3 0 4 -2

1 -4 6

5 b 8 5 P / × 10 L

0 3 4 -5 a.u. -2 1 -10

5 8 15 c

Absorption / × 10 3 5 4 -1 0 cm

1 -1 -5 L m o l

5 1 d 8 5 E / × 10

3 0 Δ 4 -5 1

0 0 250 300 350 400 450 500 550 λ / nm

Figure 3.16. Absorption (red) and emission (blue) spectra for (a) 5, (b) 6•5, (c) 7•5, and (d) 8•5 along with CD spectra for modified (open green circles) and control oligonucleotides (filled green circles): (a) 5 vs. 9, (b) 6•5 vs. 6•9, (c) 7•5 vs. 7•9, and (d) 8•5 vs. 8•9. 149

The single stranded oligonucleotide 5 and corresponding duplexes 6•5–8•5 were then

photophysically evaluated for their absorption and emission maxima, whereby Stokes shifts

were calculated (figure 3.16 and table 3.4). To accurately determine the ground state

absorption maximum of the emissive nucleoside within the hybridized duplexes, the absorption

spectrum of a control, unmodified oligonucleotide, containing the same base composition, was

subtracted from the absorption of the modified duplex (see figure 3.16 for subtracted and

figure 3.17 for non-subtracted absorption spectra). It should be noted, that most studies

calculate the Stokes shifts from the emission and excitation maxima and do not extract the

actual ground state absorption maximum of the probe within the specific duplex.

0.6

0.4 Absorption

0.2

0 240 280 320 360 400 λ / nm

Figure 3.17 Absorption spectra of single stranded oligonucleotides 9 (solid red line), 5 (dashed red line), and duplexes 6•9 (solid blue line), 6•5 (dashed blue line), 7•9 (solid green line), 7•5 (dashed green line), 8•9 (solid orange line), and 8•5 (dashed orange line) all at 25.0 106 M.

150

The calculated Stokes shifts could then be converted to corresponding polarity values utilizing the linear fit of the reference scale (figure 3.13 and table 3.4). As expected, the single strand 5 shows the largest Stokes shift, indicating the environmentally sensitive nucleoside is

exposed to a relatively polar aqueous environment. Its calculated polarity of ET(30) = 48.3 is

significantly lower than that of water [ET(30) = 63.1 kcal/mol], suggesting partial shielding of

the furanyl dT analogue 1 by neighbouring nucleobases, which is supported by the CD

spectrum showing a noteworthy secondary structure (figure 3.16). Upon perfect duplex

formation, a significant drop in Stokes shift suggesting a more apolar environment [ET(30) =

46.2 kcal/mol] consistent with encapsulation of the probe within a double helical B-DNA is seen. Where upon hybridizing 5 to its RNA complement 7, forming an A-form hybrid duplex

(figure 3.18) yields a lower major groove polarity [ET(30) = 44.8 kcal/mol]. This is consistent

with concealing the probe in a much deeper major groove found in A-form duplexes (see

chapter 1 for structural differences in DNA polymorphs). Interestingly, placing the fluorescent

probe opposite an abasic site shows the smallest Stokes shift, corresponding to an apolar

environment [ET(30) = 44.6 kcal/mol] similar that of A-form DNA. This is consistent with the previous hypothesis (see chapter 2) that the furanyl dT nucleoside 1 assumes a syn

conformation stacked between the neighbouring base pairs when paired across from an

abasic site.34,38-40 The polarity difference between B-form and A-form DNA is rather small in comparison to previous studies.24 However, due to the difference in position of previous

probes and the furanyl dT analogue 1, it is difficult to determine whether it is the probe that

cannot respond to these changes, or whether the difference at the surface of both polymorphs

is truly similar in polarity.

151

–1 –1 Table 3.4. Photophysical properties of 5, 6•5, 7•5, and 8•5; abs (cm ), em (cm ), Stokes shift –1 (SS, cm ), standard deviation (sd) and the corresponding ET(30) value determined using figure 3.10. duplo Triplo abs em SS Oligos abs em SS abs em SS abs em SS avg sd avg sd avg sd ET(30) 5 31250 23068 8182 31250 23041 8209 31153 23041 8111 31218 46 23050 13 8167 41 48.3 ± 0.44 6 5 30960 23095 7865 31153 23121 8031 31153 23148 8004 31088 91 23121 22 7967 73 46.2 ± 0.78 75 30960 23202 7758 31056 23229 7827 31153 23229 7924 31056 79 23220 13 7836 68 44.8 ± 0.73 8 5 30960 23148 7812 31056 23095 7961 30960 23095 7865 30992 45 23113 25 7879 62 44.6 ± 0.65

3.6 – Polarity conversion graphs

To accurately relate our finding with those of Ganesh, Majima and Saito, several conversions from their polarity systems to a common one, namely volume percent water in dioxane was needed. The following graphs, namely to fraction of water in dioxane41,42 (figure

43 3.18a), ET(30) to the fraction of water in glycol (figure 3.18b), and ET(30) to fraction of water in dioxane (figure 3.18c), allowed for these conversions. Breslauers 1988 publication14 made use of a reference from 1966 for the relation between and the fraction of water in dioxane.44

However, the data was not tabulated, thus we utilized the data from a more recent publication

41,42 for the construction of figure 3.18a. Multiple ET(30) determinations of dioxane-water

mixtures (containing 0.4 v% DMSO) proved to be very reproducible, allowing for the

construction of a plot revealing the relation between the ET(30) value and the ratio of water in

dioxane (figure 3.18c). Plotting a couple of dioxane-water samples without DMSO on the

same graph (stars) clearly shows there is a negligible influence of the DMSO. All three graphs

were curve fitted to facilitate the estimation of intermediate values. 152

80 a)

/ D

ε 40

0 0.0 0.2 0.4 0.6 0.8 1.0 Fraction of water in dioxane

62 60 b) c)

-1 61 55 60 50 59 45 58

(30) / kcal mol T 57 40

56 0.0 0.2 0.4 0.6 0.8 1.0 0.0 0.2 0.4 0.6 0.8 1.0 Fraction water in glycol Fraction of water in dioxane

Figure 3.18. (a) The relationship between and the fraction of water in dioxane. The orange 41,42 line represents an exponential curve fit. (b) The ET(30) dependence of the fraction water in 43 glycol. (c) A plot of the ET(30) vs. the fraction water in dioxane containing 0.4 v% DMSO (open circles) and select pure dioxane-water samples without DMSO (stars).

The following calculations utilizing the above graphs were then used to create figure 3.19.

Tor – ss 48.3 kcal/mol (figure 3.18c) 19 v% water in dioxane Tor – B-form 46.2 kcal/mol (figure 3.18c) 10 v% water in dioxane Tor – A-form 44.8 kcal/mol (figure 3.18c) 5 v% water in dioxane Ganesh:21 55 (figure 3.18a) 75 v% water in dioxane Majima:24 62 (figure 3.18a) 83 v% water in dioxane Saito:27 70 (figure 3.18a) 92 v% water in dioxane (used for figure 3.18)

An alternate conversion method: 70D 0.83 mole fraction water in (55.56 M) in glycol (17.93 M) equals 61 v%

water in glycol (figure 3.18b) ET(30) = 58.3 (figure 3.18c) 85 v% water in dioxane

153

The differences (92 v% verses 85 v%) in conversion calculations for the groove polarity

reported by Saito indicate the intricacies when conversion from one polarity reference scale to

the other is required.

100 70 48 80

47 65 60

46 40 60

45 20

55 44 E (30) v% water in T [kcal mol-1] dioxane

Figure 3.19. Comparison of major groove polarity as determined using furanyl dT analogue 1 to reported values. For perspective, both ET(30) values (left, this work) and published work (right, reported as in D) are converted to v% water in dioxane (center). Key: single strand (open circles), perfect duplex (triangles), A-form DNA (squares), data presented herein (black), Ganesh21 (orange), Majima24 (green) and Saito27 (blue).

The comparison of our observations to previously reported values for the polarity of the

DNA major groove can be seen in figure 3.19. Ganesh reported a value of 55 D for a DNA

duplex with a dansyl probe placed close to the center of a self-complementary 12-mer,21

thereby corresponding to 75 v% water in dioxane. While Majima reported a major groove polarity of 61 D,24 which converts to ~83 v% water in dioxane. Saito estimated the major groove to be even more polar, with a value of 70 D,27 which on their reference scale equals

61% water in ethylene glycol, or 92 v% water in dioxane.43 In contrast, all of the results determined using the furanyl dT analogue 1 have been more apolar than those reported by

Ganesh, Majima and Saito. Although these results seem to be in contradiction, it is our 154

contention that because all other probes are connected through relatively long and flexible linkers, they interrogate an environment farther away from the groove wall, thus a dramatically more polar environment. While the relatively rigid and linkerless furanyl-containing probe 1, is located deeper in the major groove, while its readout suggests a rather apolar environment.

This is consistent with a relatively low polarity proposed for the interior of the groove by distance-dependent dielectric constant correlations showing a steep increase in polarity as one moves away from the groove wall toward the groove exterior.9,11 Investigation of the distance dependent nature of the polarity found in DNA grooves can easily be experimentally explored with extended furanyl probes such as those suggested in section 3.7. Identification of a very apolar groove surface may explain the high binding affinity of very hydrophobic moieties while also suggesting the addition of hydrophobic moieties to current groove binders could increase their affinity.

3.7 – Conclusion and future plans

As presented above, the numerous considerations that must be addressed in determination of groove polarity via fluorescent probes is considerable. The probes size, shape and linker length will inevitably evoke changes within the biomolecular cavity that might taint the readout. The sensitivity of the probe may result in the inability to accurately determine slight differences in polarity. Infinitesimally small probes are unrealistic, thus small and minimally perturbing novel fluorophores, such as the furanyl dT analogue 1, that when judiciously placed and systematically applied in conjunction with the expression of polarity

based on ET(30) are likely to shed new light on fundamentally important questions regarding

biopolymeric microenvironments.

Two novel fluorescent nucleosides have been designed to explore the distance dependent

polarity relationship of DNA grooves (figure 3.20). The extension of the furanyl moiety via an

ethynyl or diethynyl linker will extend out the probe in a linear fashion and when base paired 155

with its perfect complement dA, will maintain a clearly defined position (figure 3.21). The exploration of sequence composition is also of interest. Thus, utilizing all three probes (1, 5 and 6) will allow for a three-dimensional exploration of the extent of sequence on polarity.

These novel ethynyl extended thymidine analogues 5 and 6 can be cyclized to allow the probing of a similar distance from the groove wall, but at a different angle resulting in dC analogues 7 and 8.

O O O O

NH NH NH HO N HO N HO N O O O O O O

HO HO HO 1 10 11

NH NH

N N HO N HO N O O O O

HO HO 12 13

Figure 3.20. Structures of novel nucleoside analogues who structure systematically extends (10 and 11) and reorients (12 and 13) the probe from the current analogue 1.

156

0 2 4 6 8 1010Å 0 2 4 6 8 1010Å

Figure 3.21. Models of the furanyl analogue 1 and ethynyl thienyl analogue 17 (meant to be structurally similar to the furanyl ethynyl 10) in a DNA duplex viewed (a) from the side with surfaces and (b) from the top.

To identify that these suggested nucleosides have desirable photophysical properties, the

thienyl versions of 10 and 12, being synthetically simpler, have been synthesized (scheme

3.1). Standard Sonogashira coupling of 2-ethenylthiophene resulted in the extended thienyl

12 in 40% yield over two steps. Cyclization of the ethyne moiety onto the C-4 carbonyl

produced the furano derivative whereby exposure to ammonia under elevated temperatures

resulted in the pyrrolo or dC face. 157

S S 14 Si I O O a NH NH HO N HO N b S O O H O O

15 HO 16 HO 17

S S

NH O

N N HO N HO N d O O O O

HO HO 19 18 Scheme 3.1. (a) sodium hydroxide, 1:1 methanol:tetrahydrofuran; (b) Pd(dppf)Cl2, copper iodide, triethylamine, dimethylformamide, 40% over two steps; (c) copper iodide, triethylamine, methanol, 57%; (d) ammonia, methanol.

All three nucleosides are fluorescent with absorption bands ( 320 nm) that are isolated from the native nucleobases (figure 3.22). The extended ethyne, while having the lowest quantum yield (general comparison of 17–19 under similar conditions) is the most sensitive to polarity changes. Similar to other nucleosides we have synthesized, 17 and 18 show bathochromic and hyperchromic shifts upon an increase in the polarity of its environment. The extended ethyne is the most dramatic, resulting in a ~45 nm difference between polar and apolar environments, while the cyclized furano 18 results in only a ~25 nm change.

Conversion from the ethyne thienyl 17 to the cyclized furano 18 shows only a minor red shift in emission maximum (446 nm 445 nm in water) with a dramatic shortening of the corresponding stokes shift from 8,828 cm-1 to 5,955 cm-1. The pyrrolo dC 19 was synthesized 158

in only small quantities and thus only a excitation and emission spectra were recorded, although it appears that the absorption and emission maximum do not shift upon conversion from the furano to the pyrrolo. These future plans are not free from the concerns addressed in the above chapter, but do represent the extension on novel investigations whose results have shed new light on the polarity of DNA grooves. 159

a) 4 0.8

2 0.4

0 0

b) 12 0.8 PL Intensity PL Absoprtion / a.u. 8

0.4 4

0 0 c) 3

2 PL Intensity PL 1

0 300 400 500 600 λ / nm

Figure 3.22. Absorption and emission spectra of (a) 17 and (b) 18 in water (black), acetonitrile (blue) and dioxane (red). (c) Excitation and emission spectra of 19 in water.

160

dT Thienyl dU Thienyl ethynyl dU (17) Thienyl furano dU (18)

Figure 3.23. X-ray structures of the thienyl analogues in comparison to dT.45

3.8. Acknowledgements

Portions of Chapter 3 have been previously published in the article "Polarity of major

grooves explored using an isosteric emissive nucleosides" ChemBioChem in Press 2007 by

R. W. Sinkeldam, N. J. Greco and Y. Tor. The dissertation author was one of the primary investigators and author of this paper.

Appendix – Experimental information

A.1 – General procedures See “A.1 – General procedures” located in the appendix of chapter 2.

A.2 – Synthetic procedures. For synthetic procedures for nucleosides 1 and 2, see “A.2 – Synthetic procedures” located in the appendix of chapter 2. Solvatochromatic dye 3 was purchased from sigma and used without further purification. Solvatochromatic dye 4 was a generous gift from Prof. Dr. Chr. Reichardt.

2-ethenylthiophene (15). To a solution of 2-trimethylsilylethenyl thiophene (500 L, 545 mg, 3.0 mmol) in 1:1 methanol:tetrahydrofuran (10 ml) was S H added sodium hydroxide (1 N solution, 3.3 mmol). The reaction was stirred at room temperature for 15 min. Then pentane was added to the reaction solution and the organic layer was washed with water (2 ). Organic layer was dried over sodium sulfate and then the solvent was removed under reduced pressure. Product used 1 without further purification. H NMR (400 MHz, CDCl3): 7.29–7.26 (m, 2H), 6.98 (dd, J = 5.2 Hz, 1H), 3.34 (s, 1H).

161

5-ethenyl(thiophen-2-yl)-2-deoxythymidine (17). To a solution of IdUrd (0.350 g, 1.0 mmol), copper iodide (0.01 g, 0.05 mmol) and S Pd(dppf)Cl2 (0.04 g, 0.05 mmol) in anhydrous and argon degassed 1/3 trietheylamine/dimethylformamide (2 ml) was added via cannula O an argon degassed solution of 2-ethenylthiophene (0.216 g, 2.0 mmol) in 1/3 trietheylamine/dimethylformamide (2 ml). The solution NH was stirred at room temperature overnight. Solvent removed under HO N reduced pressure. Resulting solid taken up in 1:1 O O methanol:dichloromethane where upon dowex-1 1 8 200 exchange resin (1 g) was added. The suspension was stirred at HO room temperature for 45 min. The resin was filtered off and the solvent removed under reduced pressure. Product purified by flash column chromatography (5% methanol/dichloromethane) followed by precipitation from boiling 20% methanol/dichloromethane. Product: white solid (0.133 g, 0.4 mmol, 40 % yield). 1H NMR

(400 MHz, DMSO-d6): 11.72 (s, NH, 1H), 8.35 (s, H-6, 1H), 7.64 (dd, J = 5.2 and 5.2 Hz, H- 5'', 1H), 7.34 (dd, J = 3.6 and 3.6 Hz, H-3'', 1H), 7.10 (dd, J = 4.8 and 4.8 Hz, H-4''), 6.11 (t, J = 6.6 Hz, H-1', 1H), 5.25 (d, J = 4 Hz, 3'-OH, 1H), 5.14 (t, J = 5 Hz, 5'-OH, 1H), 4.24 (m, H-3', 13 1H), 3.79 (m, H-4', 1H), 3.59 (m, H-5', 2H), 2.15 (m, H-2', 2H); C NMR (100 MHz, DMSO-d6): 161.1, 149.1, 143.8, 132.3, 1218.6, 137.5, 121.8, 97.7, 87.5, 85.9, 84.9, 84.8, 69.9, 60.8; + ESI calculated for C15H15N2O5S [M+H] 335.07, found: 334.82; UV (water) max = 320 nm ( =

17,600), 260 = 11,750. Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 17.

Crystal data and structure refinement for 5-ethenyl(thiophen-2-yl)-2-deoxythymidine (17). Identification code tor23

Empirical formula C15H14N2O5S Formula weight 334.34 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Monoclinic Space group P2(1) Unit cell dimensions a = 6.7936(15) Å a= 90° b = 15.611(3) Å b= 97.065(4)° c = 13.795(3) Å g = 90° Volume 1451.8(5) Å3 Z 4 Density (calculated) 1.530 g/cm3 Absorption coefficient 0.252 mm-1 F(000) 696 Crystal size 0.30 x 0.20 x 0.15 mm3 Theta range for data collection 2.61 to 28.13° Index ranges -8<=h<=, -20<=k<=19, -17<=l<=17 Reflections collected 12526 Independent reflections 6388 [R(int) = 0.0296] Completeness to theta = 27.57° 94.6 % Absorption correction multi scan Max. and min. transmission 0.9632 and 0.9282 Refinement method Full-matrix least-squares on F2 Data / restraints / parameters 5290 / 9 / 434 162

Goodness-of-fit on F2 1.111 Final R indices [I>2sigma(I)] R1 = 0.0501, wR2 = 0.1286 R indices (all data) R1 = 0.0532, wR2 = 0.1310 Absolute structure parameter 0.04(9) Largest diff. peak and hole 0.596 and -0.281 e Å-3

5-furano(thiophen-2-yl)-2-deoxythymidine (18). A solution of 17 (0.048 g, 0.14 mmol) and copper iodide (0.018 g, 0.09 mmol) in S anhydrous 1:4 triethylamine:methanol (5 ml) was refluxed for 5 h. Solvent removed under reduced pressure and solid washed triturated O with 5% EDTA solution. Product: light brown solid (0.0.27 g, 0.82 1 mmol, 57 % yield). H NMR (400 MHz, DMSO-d6): 8.81 (s, H-6, 1H), N 7.72 (d, J = 5.2 Hz, H-5'', 1H), 7.63 (d, J = 2.8 Hz, H-3'', 1H), 7.20 (t, J = 4.2 Hz, H-4''), 6.16 (t, J = 5.8 Hz, H-1', 1H), 5.29 (d, J = 4.4 Hz, 3'- HO N O O OH, 1H), 5.16 (t, J = 5.2 Hz, 5'-OH, 1H), 4.23 (m, H-3', 1H), 3.92 (m, H-4', 1H), 3.65 (m, H-5', 2H), 2.40 (m, H-2', 1H), 2.08 (m, H-2', 1H); + HO ESI calculated for C15H14N2NaO5S [M+Na] 357.05, found: 356.92; UV (water) max = 358 nm ( = 14,500), 260 = 5,200. Suitable crystals were obtained by the slow diffusion of chloroform into a methanol solution containing nucleoside 18.

Crystal data and structure refinement for 5-furano(thiophen-2-yl)-2-deoxythymidine (18). X-ray ID tor24

Empirical formula C15H14N2O5S Formula weight 334.34 Temperature 100(2) K Wavelength 0.71073 Å Crystal system Orthorhombic Space group P2(1)2(1)2(1) Unit cell dimensions a = 6.644(3) Å a= 90. b = 6.680(3) Å b= 90° c = 31.034(15) Å g = 90° Volume 1377.3(11) Å3 Z 4 Density (calculated) 1.612 Mg/m3 Absorption coefficient 0.266 mm-1 F(000) 696 Crystal size 0.20 x 0.08 x 0.04 mm3 Crystal color/habit colorless blade Theta range for data collection 2.63 to 25.03°. Index ranges -7<=h<=7, 0<=k<=7, 0<=l<=36 Reflections collected 2417 Independent reflections 2417 [R(int) = 0.0826] Completeness to theta = 25.00° 99.9 % Absorption correction Semi-empirical from equivalents Max. and min. transmission 0.9895 and 0.9488 Refinement method Full-matrix least-squares on F2 163

Data / restraints / parameters 2417 / 0 / 211 Goodness-of-fit on F2 1.170 Final R indices [I>2sigma(I)] R1 = 0.0785, wR2 = 0.1567 R indices (all data) R1 = 0.0899, wR2 = 0.1619 Absolute structure parameter 0.2(3) Largest diff. peak and hole 0.488 and -0.412 e.Å-3

Atomic coordinates ( x 104) and equivalent isotropic displacement parameters (Å2x 103) for 5- furano(thiophen-2-yl)-2-deoxythymidine (18). U(eq) is defined as one third of the trace of the orthogonalized Uij tensor. x y z U(eq) C(1) 1810(7) -42(9) 3474(2) 18(1) C(2) 1883(8) -206(9) 3040(2) 18(1) C(3) 3373(8) -226(9) 2705(2) 18(1) C(4) 2378(7) -255(8) 2331(2) 13(1) C(5) -2(8) -275(8) 2825(2) 13(1) C(6) -1860(7) -238(8) 3441(2) 14(1) C(7) 3007(8) -286(9) 1891(2) 17(1) C(8A) 1382(3) -249(3) 1488(1) 21(1) C(8B) 5369(4) -381(4) 1758(1) 29(1) C(9) 4983(10) -439(9) 1266(2) 26(1) C(10) 3127(9) -382(9) 1120(2) 25(1) C(11) -188(8) 260(8) 4136(2) 17(1) C(12) 95(8) -1628(8) 4397(2) 13(1) C(13) 1040(7) -854(7) 4811(2) 10(1) C(14) 2292(8) 886(8) 4666(2) 13(1) C(15) 4510(8) 365(9) 4592(2) 18(1) N(1) -26(6) -35(7) 3669(1) 16(1) N(2) -1775(6) -298(7) 2999(1) 14(1) O(1) 264(5) -327(6) 2399(1) 15(1) O(2) -3414(5) -353(6) 3637(1) 15(1) O(3) 1409(6) 1562(6) 4262(1) 17(1) O(4) -446(5) -188(6) 5109(1) 16(1) O(5) 4715(6) -1471(5) 4368(1) 16(1) S(1A) 1382(3) -249(3) 1488(1) 21(1) S(1B) 5369(4) -381(4) 1758(1) 29(1) 164

5-pyrrolo(thiophen-2-yl)-2-deoxythymidine (19). A solution of 12 S (0.011 g, 0.03 mmol) in ammonia saturated anhydrous methanol (2 ml) was stirred at 50°C overnight. Solvent removed under reduced pressure. Product purified by flash column chromatography (10% NH methanol/dichloromethane). Product: yellow oil; fluorescent spot that is more polar than starting material on tlc. N

HO N O O

A.3 - Photophysical Studies of the isolated nucleosides 1 and 2. UV-Vis spectra were recorded on a Hewlett Packard 8453 Diode Array Spectrometer in a 1 mL quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) at ambient temperature (23°C). Steady state fluorescence experiments were carried out at ambient temperature (23°C) in a 1 mL quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Jobin Yvon Horiba FluoroMax-3 luminescence spectrometer with excitation and emission slit-widths of 6 nm. All dioxane-water samples were prepared from a stock solution in DMSO, hence, the resulting measured samples contain 0.4 v% DMSO. All MCH-iPrOH solutions were prepared from a 1,2- dichloroethane stock solution resulting in samples containing 0.2 v% 1,2-dichloroethane. The

ET(30) values of polar (dioxane-water) and apolar (methylcyclcohexane-isopropanol) solvent mixtures were determined experimentally by taking the long wavelength absorption maximum of the dissolved Reichardts dye (3)37 and the lipophilic penta t-butyl substituted pyridinium-N- phenolate betaine dye[4,5] (4).

All emission maxima in cm–1 were determined after correction of the intensity according:

Intensity [] = 2 Intensity [].46

Curve fits were generated using OriginPro 7.5. All reported standard deviations were calculated using STDEVP in Microsoft Excel.

A.4 - Photophysical Studies of the isolated nucleosides 12–14. UV-Vis spectra were recorded on a Hewlett Packard 8453 Diode Array Spectrometer in a 1 mL quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) at ambient temperature (23°C). Steady state fluorescence experiments were carried out at ambient temperature (23°C) in a 1 mL fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Jobin Yvon Horiba FluoroMax-3 luminescence spectrometer with excitation and emission slit-widths of 4 nm for 12 and 2 nm for 13. All samples were prepared from a stock solution in DMSO, hence, the resulting measured samples contain 0.5 v% DMSO with a nucleoside concentration of 5.0 10-5 M.

165

A.5 - Oligonucleotide Synthesis and Purification Unmodified oligonucleotides (including RNA) were purchased from Intergraded DNA Technologies (Coralville, Iowa) and purified by PAGE. Modified oligonucleotides were synthesized on a 1.0 mole scale (500 Å CPG column) using a Biosearch Cyclone Plus DNA synthesizer. Phosphoramidite 8 was site specifically incorporated into the oligonucleotide by trityl-off synthesis of the base oligonucleotide, followed by manual coupling of phosphoramidite 8. Typically, the modified phosphoramidite was dissolved in 100 L of anhydrous acetonitrile to give a final concentration of 0.1M. The phosphoramidite solution was pushed into the CPG column via syringe and then 200 L of 0.45M 1H-tetrazole was pushed into the other end of the column via syringe. Coupling reactions, performed twice, were allowed to proceed for 5 minutes (99% coupling efficiency) and were subsequently followed by standard oxidation and capping steps. The rest of the oligonucleotide was synthesized via the standard trityl-off procedure. Upon completion of the oligonucleotide synthesis, the CPG column was treated with 3 mL of 30% aqueous ammonium hydroxide for 2 h at room temperature, mixing via syringe every 1 h. The resulting solution was removed and the CPG column was treated with 1 mL of 30% aqueous ammonium hydroxide at room temperature for 15 min, mixing via syringe every 5 min. The resulting aqueous ammonium hydroxide solutions were consolidated and stored at room temperature for 24 h. The aqueous ammonium hydroxide solutions were freeze-dried and purified by 20% polyacrylamide gel electrophoresis. The oligonucleotide was visualized by UV shadowing, bands were excised from the gel, and extracted with 0.5M sodium acetate buffer overnight. The resulting solution was filtered (Bio Rad poly-prep chromatography column) and desalted using a Sep-Pak cartridge (Waters Corporation, MA). The following 260 nm extinction coefficients were used to determine the concentration of oligonucleotides: rG/dG = 11,700, rC/dC = 7,300, rA/dA = 15,400, dT = 8,800, rU = 10,100, and 1 = 13,000.

A.6 – Oligonucleotide Sequencing Using MALDI-TOF MS See “A.7 – MALDI-TOF of control and modified oligonucleotides” located in appendix of chapter 2.

A.7 – Oligonucleotide Spectroscopy Studies UV-Vis spectra were recorded on a Hewlett Packard 8453 Diode Array Spectrometer in a 350 L quartz UV cell with a path length of 0.1 cm (Hellma GmbH & Co KG, Müllheim, Germany) at ambient temperature. The absorption of the unique modified nucleobase was determined by subtracting a control, unmodified oligonucleotide, containing the same base composition, from the modified duplex. Steady state fluorescence experiments were carried out at 23°C in a 100 L quartz fluorescence cell with a path length of 1.0 cm (Hellma GmbH & Co KG, Müllheim, Germany) on a Jobin Yvon Horiba FluoroMax-3 luminescence spectrometer with excitation and emission slit-widths of 10 nm. DNA samples were hybridized by heating to 90 ºC for 5 min and subsequently allowed to cool to room temperature over 2 h prior to measurements. DNA samples were measured at 25.0 106 M concentration (ss and ds) in 1.0 101 M sodium chloride, 1.0 102 M sodium phosphate buffer at pH 7.0. All emission maxima in cm-1 were determined after correction of the intensity according:

Intensity [] = 2 Intensity [].46

All reported standard deviations were calculated using STDEVP in Microsoft Excel.

166

A.8 – Thermal denaturation studies See “A.8 – Thermal denaturation studies” located in the appendix of chapter 2.

A.9 - Circular Dichroism Studies CD spectra were recorded on a Aviv 215 Circular Dichroism Spectrometer in a 350 L quartz UV cell with a path length of 0.1 cm (Hellma GmbH & Co KG, Müllheim, Germany) at 23°C. DNA samples were hybridized by heating to 90 ºC for 5 min and subsequently allowed to cool to room temperature over 2 h prior to measurements. DNA samples were measured at 25.0 106 M concentration (ss and ds) in 1.0 101 M sodium chloride, 1.0 102 M sodium phosphate buffer at pH 7.0. Spectra were recorded as the average of two scans from 220– 380nm with a reading time of 5.0 sec every nm.

A.10 - Oligonucleotide Duplex Models Models were generated by manually docking the coordinates obtained from x-ray structures of the free nucleotide 1 into published coordinates of a short DNA duplex[15] using Swiss Pdb-Viewer.47 Surfaces were visualized using Chimera.33

3.9 – References

1. Falk, M.; Hartman, K. A.; Lord, R. C. "Hydration of Deoxyribonucleic Acid .3. A Spectroscopic Study of Effect of Hydration on Structure of Deoxyribonucleic Acid" J. Am. Chem. Soc. 1963, 85, 391-&.

2. Hearst, J. E. "Determination of Dominant Factors Which Influence Net Hydration of Native Sodium Deoxyribonucleate" Biopolymers 1965, 3, 57-&.

3. Neidle, S. Nucleic acid structure; VCH Publishers: New York, N.Y., (USA), 1987.

4. Saenger, W. Principles of nucleic acid structure; Springer-Verlag: New York, 1984.

5. Falk, M.; Poole, A. G.; Goymour, C. G. "Infrared Study of State of Water in Hydration Shell of DNA" Can. J. Chem. 1970, 48, 1536-&.

6. Tunis, M. J. B.; Hearst, J. E. "On Hydration of DNA .I. Preferential Hydration and Stability of DNA in Concentrated Trifluoroacetate Solution" Biopolymers 1968, 6, 1325-&.

7. Wolf, B.; Hanlon, S. "Structural Transitions of Deoxyribonucleic-Acid in Aqueous- Electrolyte Solutions .2. Role of Hydration" Biochemistry 1975, 14, 1661-1670.

8. Jayaram, B.; Sharp, K. A.; Honig, B. "The Electrostatic Potential of B-DNA" Biopolymers 1989, 28, 975–993.

9. Lamm, G.; Pack, G. R. "Calculation of dielectric constants near polyelectrolytes in solution" J. Phys. Chem., B 1997, 101, 959–965. 167

10. Mazur, J.; Jernigan, R. L. "Distance-dependent dielectric constants and their application to double-helical DNA" Biopolymers 1991, 31, 1615–1629.

11. Wang, L. H.; Hingerty, B. E.; Srinivasan, A. R.; Olson, W. K.; Broyde, S. "Accurate representation of B-DNA double helical structure with implicit solvent and counterions" Biophys. J . 2002, 83, 382–406.

12. Lu, X. F.; Heilman, J. M.; Blans, P.; Fishbein, J. C. "The structure of DNA dictates purine atom site selectivity in alkylation by primary diazonium ions" Chem. Res. Toxicol. 2005, 18, 1462–1470.

13. Szekely, J.; Gates, K. S. "Noncovalent DNA binding and the mechanism of oxidative DNA damage by fecapentaene-12" Chem. Res. Toxicol. 2006, 19, 117–121.

14. Jin, R.; Breslauer, K. J. "Characterization of the Minor Groove Environment in a Drug DNA Complex - Bisbenzimide Bound to the Poly[D(at)].Poly[D(at)]Duplex" Proc. Nat. Acad. Sci. U.S.A. 1988, 85, 8939–8942.

15. Harshman, K. D.; Dervan, P. B. "Molecular Recognition of B-DNA by Hoechst 33258+" Nucleic Acids Res. 1985, 13, 4825-4835.

16. Mikhailov, M. V.; Zasedatelev, A. S.; Krylov, A. S.; Gurskii, G. V. "Mechanism of the Recognition of at Pairs in DNA by Molecules of the Dye Hoechst 33,258" Mol. Biol. 1981, 15, 541-554.

17. Pjura, P. E.; Grzeskowiak, K.; Dickerson, R. E. "Binding of Hoechst-33258 to the Minor Groove of B-DNA" J. Mol. Biol. 1987, 197, 257-271.

18. Zimmer, C.; Wahnert, U. "Nonintercalating DNA-Binding Ligands - Specificity of the Interaction and Their Use as Tools in Biophysical, Biochemical and Biological Investigations of the Genetic Material" Prog. Biophys. Mol. Biol. 1986, 47, 31-112.

19. Sinkeldam, R. W.; Tor, Y. "To D or not to D?On estimating the microenvironment polarity of biomolecular cavities" Org. Biomol. Chem. 2007, 5, 2523–2528.

20. Barawkar, D. A.; Ganesh, K. N. "Exploring DNA Minor-Groove Interactions through a Probe Conjugate in Major Groove - Fluorescence Studies on Netrosin Complexation with Du-5-Aminodansyl-DNA" Biochem. Biophys. Res. Commun. 1994, 203, 53–58.

21. Barawkar, D. A.; Ganesh, K. N. "Fluorescent D(Cgcgaattcgcg) - Characterization of Major Groove Polarity and Study of Minor-Groove Interactions through a Major Groove Semantophore Conjugate" Nucleic Acids Res. 1995, 23, 159–164.

22. Jadhav, V. R.; Barawkar, D. A.; Ganesh, K. N. "Polarity sensing by fluorescent oligonucleotides: First demonstration of sequence-dependent microenvironmental changes in the DNA major groove" J. Phys. Chem., B 1999, 103, 7383–7385.

23. Tang, G. Q.; Tanaka, N.; Kunugi, S. "Salt effects on fluorescence spectral shifts of DNA-bound Hoechst 33258 and reaction volumes of the minor groove binding" Bull. Chem. Soc. Jpn. 1999, 72, 1129–1137. 168

24. Kimura, T.; Kawai, K.; Majima, T. "Monitoring of microenvironmental changes in the major and minor grooves of DNA by dan-modified oligonucleotides" Org. Lett. 2005, 7, 5829–5832.

25. Kimura, T.; Kawai, K.; Majima, T. "Probing the microenvironments in the grooves of Z- DNA using dan-modified oligonucleotides" Chem. Commun. 2006, 1542–1544.

26. Okamoto, A.; Tainaka, K.; Fujiwara, Y. "Nile Red nucleoside: Design of a solvatofluorochromic nucleoside as an indicator of micropolarity around DNA" J. Org. Chem. 2006, 71, 3592–3598.

27. Okamoto, A.; Tainaka, K.; Saito, I. "A dielectric-sensitive fluorescent DNA probe for monitoring polarities on the interior of a DNA-binding protein" Bioconjugate Chem. 2005, 16, 1105–1111.

28. Tainaka, K.; Tanaka, K.; Ikeda, S.; Nishiza, K.; Unzai, T.; Fujiwara, Y.; Saito, I.; Okamoto, A. "PRODAN-conjugated DNA: Synthesis and photochemical properties" J. Am. Chem. Soc. 2007, 129, 4776–4784.

29. Kawski, A. "Solvent dependence of the wave number of electron bands of luminescent molecules and determination of the electrical dipole moments in the excited state" Acta Phys. Pol. 1966, 29, 507–518.

30. Lippert, E. "Habilitationsschrift Zur Erlangung Der Lehrberichtigung (Venia Legendi) Fur Das Fach Physikalische Chemie Ander Techischen-Hochschule-Stuttgart - Spektroskopische Bestimmung Des Dipolmomentes Aromatischer Verbindungen Im Ersten Angeregten Singulettzustand" Zeitschrift Fur Elektrochemie 1957, 61, 962– 975.

31. Mataga, N.; Kaifu, Y.; Koizumi, M. "Solvent Effects Upon Fluorescence Spectra and the Dipolemoments of Excited Molecules" Bull. Chem. Soc. Jpn. 1956, 29, 465–470.

32. Quintana, J. R.; Lipanov, A. A.; Dickerson, R. E. "Low-Temperature Crystallographic Analyses of the Binding of Hoechst-33258 to the Double-Helical DNA Dodecamer C- G-C-G-a-a-T-T-C-G-C-G" Biochemistry 1991, 30, 10294-10306.

33. Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. "UCSF chimera - A visualization system for exploratory research and analysis" J. Comput. Chem. 2004, 25, 1605–1612.

34. Greco, N. J.; Tor, Y. "Simple fluorescent pyrimidine analogues detect the presence of DNA abasic sites" J. Am. Chem. Soc. 2005, 127, 10784–10785.

35. Greco, N. J.; Tor, Y. "Synthesis and site-specific incorporation of a simple fluorescent pyrimidine" Nat. Protoc. 2007, 2, 305–316.

36. Greco, N. J.; Tor, Y. "Furan decorated nucleoside analogues as fluorescent probes: synthesis, photophysical evaluation, and site-specific incorporation" Tetrahedron 2007, 63, 3515–3527. 169

37. Reichardt, C. "Solvatochromic Dyes as Solvent Polarity Indicators" Chem. Rev. 1994, 94, 2319–2358.

38. Brauns, E. B.; Madaras, M. L.; Coleman, R. S.; Murphy, C. J.; Berg, M. A. "Measurement of local DNA reorganization on the picosecond and nanosecond time scales" J. Am. Chem. Soc. 1999, 121, 11644–11649.

39. Somoza, M. M.; Andreatta, D.; Murphy, C. J.; Coleman, R. S.; Berg, M. A. "Effect of lesions on the dynamics of DNA on the picosecond and nanosecond timescales using a polarity sensitive probe" Nucleic Acids Res. 2004, 32, 2494–2507.

40. Stivers, J. T. "2-aminopurine fluorescence studies of base stacking interactions at abasic sites in DNA: metal-ion and base sequence effects" Nucleic Acids Res. 1998, 26, 3837–3844.

41. Hasted, J. B. Water: A Comprehensive Treatise 1973, 2, 405–458.

42. Wohlfarth, C.; Madelung, O. Statische Dielektrizitätskonstanten reiner Flüssigkeiten und binärer flüssiger Mischungen; Springer-Verlag: Berlin ; New York, 1991.

43. Skwierczynski, R. D.; Connors, K. A. "Solvent effects on chemical processes. Part 7. Quantitative description of the composition dependence of the solvent polarity measure ET(30) in binary aqueous-organic solvent mixtures" J. Chem. Soc. Faraday Trans. 2 1994, 467–472.

44. Franks, F.; Ives, D. J. G. "Structural Properties of Alcohol-Water Mixtures" Quarterly Rev. 1966, 20, 1–44.

45. Young, D. W.; Tollin, P.; Wilson, H. R. "Crystal and Molecular Structure of Thymidine" Acta Crystallographica Section B-Structural Crystallography and Crystal Chemistry 1969, B 25, 1423-&.

46. Lakowicz, J. R. Principles of fluorescence spectroscopy; 2nd ed.; Kluwer Academic/Plenum: New York, 1999.

47. http://expasy.org/spdbv/