SYNTHESIS AND CHARACTERIZATION OF C8 ANALOGS OF C-DI-GMP;
NEW SYNTHETIC METHOD FOR 5’-CAPPED OLIGORIBONUCLEOTIDES
by
ELIZABETH VELIATH
A Dissertation submitted to the
Graduate School-New Brunswick
Rutgers, The State University of New Jersey
in partial fulfillment of the requirements
for the degree of
Doctor of Philosophy
Graduate Program in Chemistry
written under the direction of
Roger A. Jones
and approved by
______
______
______
______
New Brunswick, New Jersey
January, 2011
ABSTRACT OF THE DISSERTATION
Synthesis and Characterization of C8 Analogs of c-di-GMP; New Synthetic Method
for 5’-Capped Oligoribonucleotides
By ELIZABETH VELIATH
Dissertation Director: Roger A. Jones
This research is centered on two projects of biological importance composed of
RNA –based compounds: C8 analogs of cyclic diguanosine monophosphate (c-di-GMP)
and 5’-capped oligoribonucleotides.
c-di-GMP is an important bacterial second messenger molecule that is critical for the transition between a biofilm-protected sessile state and a virulent, single cellular motile state in species like V. Cholerae and S.Typhimurium. The synthesis and solution phase structural analysis of a family of C8-modified analogs is described. Starting from unmodified c-di-GMP, elaboration at the C8 position of both guanine moieties of c-di-
GMP resulted in the bromo, thio, methylthio, phenyl, and meta-acetylphenyl analogs.
Biophysical studies of all five compounds was performed using 1D and (1H, 31P,
13C) and 2D (DOSY, HMBC/HMQC) NMR to ascertain the level of G-quadruplex formation. It was found that only c-di-Br-GMP as the K+ salt form adopts the formation
of higher order complexes containing guanine quartet structures. All analogs have an
NMR-visible amino resonance that is most prominent at low temperatures due to
protection from exchange by the self-stacking, and that disappears at elevated
ii
temperatures, which does not occur in with the 8-bromo-GMP monomer illustrating the
special structural characteristics of the symmetric dimer.
Capped RNA has a unique 5’-end structure containing a terminal N7-methylated
guanosine that is joined via a triphosphate bridge to the 5’-OH of all eukaryotic mRNAs.
This structure plays a vital role in regulating RNA maturation, processing, transport and
translation, but capped RNA is extremely difficult to synthesize chemically due to its
instability.
A new protection strategy was devised that uses a lipophilic dimethoxytrityl
(DMT) group on the amino group of the N7-methylated guanine activated capping
reagent. This allows efficient purification of the final capped RNA on reverse-phase
HPLC. Additionally, the DMT group increases the capping reagent solubility in organic solvents which improves the final coupling step.
The synthetic method is general to include a variety of mixed sequence oligonucleotides, and is compatible with reverse-phase HPLC. This method has been
used to synthesize and purify the unmodified cap structure m7GpppG, the individual
7 7 diastereomers of the α-thiophosphate analog m Gppp(s)G, m GpppT8 and mixed
7 7 7 sequences of m Gppp-(2’-O-Me-GAUGC), m Gppp-(2’-O-Me-GAUGC)2, m Gppp-(2’-
7 O-Me-GUAUC)4 and m Gppp-(GUAUC)4.
iii
DEDICATION
This work is dedicated to my husband Aaron, whose continued support, encouragment and understanding, has been absolutely invaluable to me throughout my graduate career; to my brother, Andrew, for his friendship and generosity; and to my parents, George Veliath and Ginger Cyr, for always believing in me. I also wish to extend my deepest thanks to my extended family and friends for their encouragement and support as well.
iv
ACKNOWLEDGEMENTS
I am extremely grateful to my advisor, Professor Roger A. Jones, for his
guidance, inspiration and continuous support, throughout my graduate research in the field of nucleic acid chemistry. I thank him for the countless conversations that have helped hone my skills in becoming a better scientist.
I am extremely grateful to Professor Barbara L. Gaffney for many useful conversations pertaining to chemistry, for her daily role as a mentor and for her friendship and encouragment.
I would like to thank the members of my thesis committee, Professor Jeehiun K.
Lee, Professor Lawrence Williams from the Department of Chemistry at Rutgers and
Professor Ed LaVoie from the Department of Pharmacy at Rutgers for their advice, time
and helpful suggestions.
I would like to thank Professor Kenneth Breslauer for help with
spectrophotometric instrumentation; and Dr. Jens Volker for many useful conversations
pertaining to spectroscopy of oligonucleotides.
I would like to thank the past Jones group lab members, as well as the friends I
have forged strong friendships with over the years at Rutgers, for making my graduate
study a very enriching experience.
v
I am grateful for the significant financial support provided by the Rutgers
Excellence Fellowship, the GAANN Fellowship, NIH and the Department of Chemistry and Chemical Biology at Rutgers, the State University of New Jersey.
vi
TABLE OF CONTENTS
Abstract ..……………………………………………………………………………. ii
Dedication ………………………………………………………………………….. iv
Acknowledgements ………………………………………………………………….. v
Table of Contents ………………………………………………………………….. viii
List of Figures ………………………………………………………………………. xii
List of Schemes ……………………………………………………………………. xiv
List of Tables ……………………………………………………………………….. xvii
List of Abbreviations …………………………………………………………...…. xviii
Chapter 1
A Brief Overview of Synthetic Ribonucleoside Chemistry ………………………....1
References …………………………………………………………………………. 2
Chapter 2
Synthesis and Characterization of C8 Analogs of c-di-GMP ………………………3
1. Biological Background of c-di-GMP ………………………………………….. 3
1.1 c-di-GMP as Bacterial Second Messenger Signaling Molecule ………….. 3
1.2 Discovery of c-di-GMP …………………………………………………... 4
1.3 c-di-GMP Regulates Biofilm Production ………………………………… 4
1.4 c-di-GMP Regulates Virulence and Pathogenesis …………………………6
1.5 Enzymes that Synthesize and Degrade c-di-GMP …………………………8
1.6 c-di-GMP Target Receptors ………………………………………………..8
1.7 Temporal and Spatial Regulation of c-di-GMP Levels …………………...12
1.8 Effects of c-di-GMP on Mammalian Cells ………………………………. 14
vii
1.9 References ………………………………………………………………..15
2. Synthetic Background of c-di-GMP and Analogs ……………………………18
2.1 c-di-GMP Synthetic Introduction ………………………………………..18
2.2 Phosphotriester Chemistry ……………………………………………….19
2.3 H-Phosphonate Chemistry ……………………………………………….21
2.4 Combination of Phosphotriester, H-Phosphonate, and Phosphoramidite
Chemistry ………………………………………………………………...23
2.5 Other Methods ……………………………………………………………28
2.6 References ………………………………………………………………..29
3. Synthesis of C8 Analogs of c-di-GMP …………………………….………….31
3.1 Introduction ……………………………………………………………....31
3.2 Synthesis of c-di-Br-GMP ………………………………………………..31
3.3 Synthesis of c-di-thio-GMP ………………………………………………34
3.4 Synthesis of c-di-methylthio-GMP ……………………………………….38
3.5 Synthesis of c-di-phenyl-GMP and c-di-acetylphenyl-GMP …….……….39
3.6 Conclusions ……………………………………………………….………45
3.7 Experimental Procedures …………………………………………………46
3.8 References ……………………………………………………….………..52
3.9 Appendix …………………………………………………………………54
4. Biophysical Studies of C8 Analogs of c-di-GMP …………………………….78
4.1 Introduction ………………………………………………………………78
4.2 Previous Work on the Biophysical Studies of c-di-GMP ………………..78
4.3 Biophysical Studies of c-di-Br-GMP Salt Forms ………………………..81
viii
4.3.1 NMR Studies of c-di-Br-GMP ………………………………..81
4.3.1.1 Introduction …………………………………………...81
4.3.1.2 K+ form of c-di-Br-GMP ……………………………...82
4.3.1.3 Na+ form of c-di-Br-GMP …………………………….86
4.3.1.4 Li+ form of c-di-Br-GMP ……………………………..90
4.3.1.5 TEA+ form of c-di-Br-GMP ……………………….….91
4.3.1.6 Salt Forms of 8-Br-GMP Monomer ……………….….93
4.3.1.7 Conclusions for NMR Studies of c-di-Br-GMP ….…..96
4.3.2 UV Studies of c-di-Br-GMP ……………………………….…96
4.3.2.1 UV Melting Study for K+ and Na+ Forms of
c-di-Br-GMP ………………………………………….96
4.3.2.2 Results ………………………………………………..97
4.3.3 Further Investigation of NMR-Visible N2-Amino Resonance .98
4.3.3.1 Heteronuclear 2D NMR of K+ and Na+ Forms of
c-di-Br-GMP …………………………………………99
4.3.3.2 Effect of pH on UV Profile of Na+ Form of
c-di-Br-GMP ………………………………….……..101
4.4 Biophysical Studies of c-di-thio-GMP: K+, Na+, and TEA+ forms …..102
4.5 Biophysical Studies of c-di-methylthio-GMP: K+, Na+,
and TEA+ forms ………………………………………………………105
4.6 Biophysical Studies of c-di-phenyl-GMP: K+, Na+, and TEA+ forms ..107
4.7 Biophysical Studies of c-di-acetylphenyl-GMP: K+ and Na+ forms ….110
4.8 NMR Re-examination of c-IMP-GMP: K+ form ……………………..112
ix
4.9 Conclusions ……………………………………………………………114
4.10 References ……………………………………………………………..116
Chapter 3
New Synthetic Method for 5’-Capped Oligoribonucleotides ……………………..118
1. Capped RNA Biological Background …………………………………………….118
1.1 Introduction ……………………………………………………………...118
1.2 Biological Function of Capped RNA ……………………………………118
1.3 Chemical Structure of Capped RNA …………………………………….119
1.4 Role of Capped RNA in Regulating RNA Decay ……………………….122
1.4.1 Deadenylation-Independent Decay Mechanisms ………………122
1.4.2 Deadenylation-Dependant Decay Mechanisms ………………..124
1.5 References ………………………………………………………………..126
2. Synthetic Background of Capped RNA ……………………………….………….129
2.1 Introduction ………………………………………………….…………..129
2.2 Enzymatic Methods ……………………………………………………..129
2.3 Combination of Enzymatic and Chemical Methods …………………….130
2.4 Chemical Methods ………………………………………………………132
2.5 References ……………………………………………………………….145
3. Synthesis of 5’-Capped Oligoribonucleotides …………………………………….149
3.1 Introduction ………………………………………………………………149
3.2 Synthesis of Capping Reagent m7GDPDMT Imidazolide …………………151
3.2.1 Synthesis of GMPDMT: Transient Protection and Tritylation …..151
x
3.2.2 Synthesis of GMPDMT Imidazolide: Activation ………………..154
3.2.3 Synthesis of GDPDMT: Pyrophosphate Bond Formation ……….155
3.2.4 Synthesis of m7GDPDMT Imidazolide: N7-Methylation and
Activation ………………………………………………………156
3.2.5 Conclusions ……………………………………………………...159
3.3 Synthesis of the Cap Structure and 5’-Capped Oligoribonucleotides ……160
7 7 3.3.1 Synthesis of m GpppG and m Gppp(s)G ……………………….160
3.3.2 Synthesis of 5’-Capped Oligonucleotides ………………………163
3.3.2.1 Capping Reaction Model Systems: pT8mer and 2’-O-Me-
pGAUGC ……………………………………………...166
3.3.2.2 Capping Reaction for p(2’-O-Me-GAUGC)2 , p(2’-O-Me-
GUAUC)4 and p(2’-OH-GUAUC)4 ...... 168
3.4 Conclusions ……………………………………………………………….172
3.5 Experimental Procedures …………………………………………………173
3.6 References ………………………………………………………………..185
3.7 Appendix …………………………………………………………………187
xi
List of Figures
Figure Page
Chapter 2
2-1 PilZ Domain of VCA0042 and PP4387 ……………………………………………9
2-2 V. cholerae vc2 riboswitch in complex with c-di-GMP …………………………..12
2-3 Three main synthetic methods for internucleotide couplings ……………………..18
2-4 Condensing reagent in phosphotriester chemistry ………………………………...20
2-5 Catalytic cycle for the Suzuki-Miyaura aryl coupling …………………………….41
2-6 Different G-quartets: unmodified vs. modified …………………………………...44
2-7 Equilibrium between bi-, tetra- and octamolecular
c-di-GMP complexes ……………………………………………………………..79
2-8 Anti and Syn conformers of c-di-GMP ……………………………………………80
2-9 UV melting profiles of c-di-GMP ………………………………………………...80
2-10 1H NMR of different salt forms of c-di-GMP ……………………………………81
2-11 1H NMR of c-di-Br-GMP (K+ form) ……………………………………………..83
2-12 31P NMR of c-di-Br-GMP (K+ form) …………………………………………….85
2-13 1H NMR of c-di-Br-GMP (Na+ form) ……………………………………………88
2-14 31P NMR of c-di-Br-GMP (Na+ form) ……………………………………………89
2-15 1H and 31P NMR of c-di-Br-GMP (Li+ form) …………………………………….90
2-16 1H NMR of c-di-Br-GMP (TEA+ form) ………………………………………….92
2-17 1H NMR of 8-Br-GMP (K+ form) ………………………………………………..94
2-18 1H NMR of 8-Br-GMP (TEA+ form) …………………………………………….95
2-19 UV melting profiles for c-di-Br-GMP (K+ and Na+ forms) ……………………...98
xii
Figure Page
2-20 1H-13C HMBC/HMQC of c-di-Br-GMP (K+ and Na+ forms) …………………..100
2-21 UV wavelength scan of c-di-Br-GMP at pH 3.0 and 7.0 ……………………….102
2-22 1H NMR of c-di-thio-GMP (K+ form) …………………………………………..104
2-23 1H NMR of c-di-methylthio-GMP (K+ form) …………………………………...106
2-24 1H NMR of c-di-phenyl-GMP (K+ form) ……………………………………….108
2-25 1H NMR of c-di-phenyl-GMP (Na+ form) ……………………………………...109
2-26 1H NMR of c-di-acetylphenyl-GMP (K+ and Na+ forms) vs.
c-di-Br-GMP (K+ form) …………………………………………………………111
2-27 1H NMR of c-IMP-GMP (K+ form) ……………………………………………..113
Chapter 3
3-1 Structure of m7GpppG or capped RNA …………………………………………120
3-2 In vivo enzymatic synthesis of capping on 5’ end of transcribing RNA ………..121
3-3 Two possible RNA degradation pathways ………………………………………125
3-4 Key capping intermediate, m7GDPDMT imidazolide …………………………….151
3-5 LCMS of partially detritylated 2’-O-Me-cap20mer reaction mixture …………..169
3-6 Crude HPLC of optimized 2’-O-Me-cap20mer reaction mixture ………………170
xiii
List of Schemes
Scheme Page
Chapter 2
2-1 Phosphotriester method for synthesis of cyclic ribonucleosides ………………….19
2-2 Phosphortriester method for synthesis of c-di-GMP ………………………………21
2-3 H-phosphonate tautomeric equilibrium …………………………………………...22
2-4 H-phosphonate method for synthesis of c-di-GMP ………………………………23
2-5 Phosphoramidite/phosphotriester methods
for synthesis of c-di-GMP …………………………………………………………24
2-6 Solid-phase synthesis of c-di-GMP ……………………………………………….25
2-7 Phosphoramidite/H-phosphonate methods
for synthesis of c-di-GMP …………………………………………………………26
2-8 One-flask method for synthesis of c-di-GMP ……………………………………..28
2-9 Synthesis of 8-Br-GMP ……………………………………………………………32
2-10 Mechanism for the bromination of GMP ………………………………………..33
2-11 Synthesis of c-di-Br-GMP ……………………………………………………….33
2-12 Tautomeric equilibrium of 8-thioguanine ……………………………………….34
2-13 Initial attempts for thiolation of 8-Br-GMP …………………………………….35
2-14 Proposed mechanism of pyr-TFA catalyzed thiolation of 8-Br-GMP ………….36
2-15 Synthesis of 8-thio-GMP ……………………………………………………….37
2-16 Synthesis of c-di-thio-GMP …………………………………………………….38
2-17 Synthesis of c-di-methylthio-GMP ……………………………………………..39
xiv
Scheme Page
2-18 Synthesis of 8-Ph-GMP ………………………………………………………….42
2-19 Synthesis of c-di-phenyl-GMP …………………………………………………..42
2-20 Synthesis of c-di-acetylphenyl-GMP …………………………………………….45
2-21 Divergent synthesis for a family of C8 analogs of c-di-GMP …………………...45
Chapter 3
3-1 Synthesis of m7(LNA)GpppG cap analog ……………………………………….132
3-2 Synthesis of bismethylene cap analogs GpppG and ApppA …………………….134
3-3 Synthesis of α:β and β:γ methylene cap analogs ………………………………...135
7 7 7 3-4 Synthesis of α-m Gppp(s)G, γ-m Gp(s)ppG, and β-m Gpp(s)pG thiophosphate cap
analogs …………………………………………………………………………...137
7 3-5 Synthesis of β-m Gpp(Se)pG cap analog …………………………………………138
3-6 Synthesis of m7GpppG using S-thiophenyl activated GDP ……………………..140
3-7 Synthesis of m7GpppG using 4-methoxythiophenyl activated GDP ……………140
7,3’-O 3-8 Synthesis of m2 GpppG using morpholidate activated GDP ………………..141
3-9 Synthesis of capped GACU using 5-chloroquinoyl activated m7GDP ………….141
3-10 Synthesis of m7GpppG and capped 11mer using imidazolide activated m7GDP .142
7,3’-O 7 3-11 Synthesis of m2 GpppG and m dGpppG using imidazolide activated GMP ..143
3-12 Retrosynthetic analysis for final coupling of capped RNA ……………………..151
3-13 Transient silylation and amino tritylation of GMP ………………………………153
3-14 Imidazolide activation of GMPDMT ………………………………………………155
3-15 Mechanism of phosphorimidazolide formation for GMPDMT imidazolide ………155
3-16 Pyrophosphate bond formation resulting in GDPDMT ……………………………156
xv
3-17 Optimized reaction conditions for N7-methylation of GDPDMT ………………….158
3-18 Imidazolide activation of m7GDPDMT …………………………………………….159
3-19 Synthetic route for key capping intermediate, m7GDPDMT imidazolide …………160
3-20 Synthesis of the unmodified cap structure, m7GpppG …………………………..161
7 3-21 Synthesis of α-thiophosphate cap structure, m Gppp(s)G ………………………..162
3-22 5’-Phosphorylation using a non-nucleoside phosphoramidite …………………..163
3-23 Purification and deprotection sequence for 5’-PO4 oligonucleotides ……………164
3-24 5’-Phosphorylation with a non-nucleoside phosphoramidite ……………………165
3-25 Purification and deprotection sequence for 5’-PO4 oligonucleotides …………...166
xvi
List of Tables
Tables Page
Chapter 2
2-1 Diffusion Coefficients (m2/s) of c-di-GMP C8 analogs …………………………115
Chapter 3
3-1 Summary conversion yields for cap structures and capped oligonucleotides ……172
xvii
List of Abbreviations
AdCl adamantoyl chloride
ARCA anti-reverse cap analog
BPO butanone peroxide
8-Br-GMP 8-bromoguanosine monophosphate c-di-AMP cyclic-di-adenosine-monophosphate c-di-CMP cyclic-di-cytidine-monophosphate c-di-dGMP cyclic-di-deoxyguanosine-monophosphate c-di-GMP cyclic-di-guanosine-monophosphate c-di-TMP cyclic-di-thymidine-monophosphate c-IMP-GMP cyclic-inosine-monophosphate-guanosine-
monophosphate
Cpep 2’-O-(1-(4-chlorophenyl)-4-ethoxypiperidine-4-yl)
CBC cap-binding complex
CBP cap-binding protein
CPG controlled-pore glass c-XMP-GMP cyclic-xanthosine-monophosphate-guanosine-
monophosphate
DCA dichloroacetic acid
DGC diguanylate cyclase
DNA deoxyribonucleic acid
DMF dimethylformamide
DMT dimethoxytrityl
xviii
DMOCP 5,5-dimethyl-2-oxo-2-chloro-1,3,2-
dioxaphosphinane
DMSO dimethylsulfoxide
DOSY diffusion-ordered spectroscopy dsRNA double-stranded RNA
DTT dithiothreitol
EtOH ethanol
GMP guanosine monophosphate
GDP guanosine diphosphate
GTP guanosine triphosphate
HIT histidine triad
HMBC heteronuclear multiple-bond coherence
HMPA hexamethyl phosphoramide
HMQC heteronuclear multiple-quantum coherence
HPLC high performance liquid chromatography
IMP inosine monophosphate
I-site inhibition site
LNA locked nucleic acid
MeCN acetonitrile
MeNH2 methylamine
MeOH methanol miRNA micro RNA
MSNT 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4-triazole
xix
NBS N-bromosuccinimide
NMI N-methylimidazole
NMR nuclear magnetic resonance
PABP poly-A-binding protein
PDE phosphodiesterase
8-Ph-GMP 8-phenylguanosine monophosphate
PvCl pivaloyl chloride
Px 5’-O-(9-phenyl-xanthen-9-yl) pyr-TFA pyridinium trifluoroacetate
RNAi RNA interference
RP reverse phase
SAME S-adenosylmethionine siRNA small-interfering RNA snRNA small-nuclear RNA
SPS solid-phase synthesis tBuNH2 tert-butylamine tBuOOH tert-butylhydroperoxide
TEA triethylamine
TEAA triethylammonium acetate
TEA-HF triethylamine-hydrofluoride
TPS triisopropylbenzene sulfonyl chloride
TXPTS tri(4.6-dimethyl-3-sulfonatophenyl)phosphine
xx 1
Chapter 1
A Brief Overview of Synthetic Ribonucleoside Chemistry
Over fifty years ago, Watson and Crick published their results on the structure of deoxyribonucleic acid (DNA), which sparked a new era in the understanding of the organization of genetic material.1 Since then, the need for synthetically made DNA and
RNA has risen and as a result the synthetic methods to meet that challenge has also
evolved.2-4 The advent of phosphoramidite chemistry to create the internucleotide
linkages was an improvement over the phosphotriester or the H-phosphonate couplings
due to it’s facile adaptation to solid-phase synthesis in the high-throughput synthesis of
DNA and RNA for biochemical studies. The discovery of RNA interference (RNAi) as
an endogenous means of controlling gene expression in 19985 has lead to a surge of
research in the area of RNAi by double-stranded RNA (dsRNA), and consequently lead
to a high demand for synthetic RNA for probing biological activity.
RNA is utilized by viruses, bacteria and eukaryotes in a variety of ways to
modulate cellular activity. It has a multitude of functions aside from being an
intermediary mRNA molecule that relays the genetic code from DNA to protein primary
structure. Some of RNAs roles include self-catalysis as ribozymes, protein synthesis
machinery in the ribosome and tRNA, bacterial signaling molecules, and regulatory
elements of gene expression or riboswitches.
This thesis focuses on two distinct areas of synthetic RNA chemistry. The first
project, Chapter 2, is centered on the synthesis and characterization of a family of C8
analogs of the cyclic ribonucleotide, cyclic-di-guanosine monophosphate (c-di-GMP),
2
which in its unmodified state is an important bacterial signaling molecule. This research entails the synthesis of the analogs, and explores the biophysical properties of the compounds using nuclear magnetic resonance spectroscopy (NMR). The second project,
Chapter 3, is centered on a new synthetic method for 5’-capped oligoribonucleotides.
Capped RNA is a critical structural element on the 5’ ends of mRNA that controls mRNA transport, processing and degradation. This method uses a new protection strategy that
also serves as a purification handle to efficiently separate the chemically synthesized
capped RNA as well as the any unreacted oligonucleotide.
References
1. Watson, J.D., Crick, F.H.C., "Molecular Structure of Nucleic Acids. A Structure for Deoxyribose Nucleic Acid". Nature, 1953, 171, 737-738.
2. Letsinger, R.L., Finnan, J.L., Heavner, G.A., Lunsford, W.B., "Nucleotide Chemistry. Phosphite Coupling Procedure for Generating Internucleotide Links". Journal of the American Chemical Society, 1975, 97, 3278-3279.
3. Beaucage, S.L., Caruthers, M.H., "Deoxynucleoside Phosphoramidites. A New Class of Key Intermediates for Deoxypolynucleotide Synthesis". Tetrahedron Letters, 1981, 1981(22), 1859-1862.
4. McBride, L.J., Caruthers, M.H., "Nucleotide Chemistry. An Investigation of Several Deoxynucleoside Phosphoramidites Useful for Synthesizing Deoxyoligonucleotides". Tetrahedron Letters, 1983, 1983(24), 245-248.
5. Fire, A., Xu, S., montgomery, M.K., Kostas, S.A., Driver, S.E., Mello, C.C., "Potent and Specific Genetic Interference by Double-Stranded RNA in Caenorhabditis elegans". Nature, 1998, 391, 806-811.
3
Chapter 2
Synthesis and Characterization of C8 Analogs of c-di-GMP
1. Biological Background of c-di-GMP
1.1 c-di-GMP as Bacterial Second Messenger Signaling Molecule
c-di-GMP is a cytoplasmic bacterial second messenger molecule found in a wide
variety of human pathogenic bacteria. This compound relays extracellular signals from
the environment through the cell membrane to target effector receptors inside the cell as part of the signaling transduction cascade.1 The biological processes regulated are
numerous, including extracellular polysaccharide biofilm formation, the expression of associated factors for virulence, organelle formation for motility, cell-cycle differentiation, antibiotic resistance and quorum sensing.2, 3
Quorum sensing is a mode of bacterial cell-to-cell communication mediated by
extracellular first messenger signaling molecules that elicit an internal response. It
enables the local bacterial population to transition between stationary and planktonic
states depending on the population. This is a critical ability for adaptation to the
environmental stimuli.4, 5
A primary physiological function of c-di-GMP is acting as an allosteric activator
of Glucoacetobacter xylinus cellulose synthase. This compound is also a ligand for a
variety of other protein targets.1, 2, 6 Thus c-di-GMP is a key compound in a conserved regulatory mechanism for either a sessile or motile phenotype.4
4
1.2 Discovery of c-di-GMP
c-di-GMP was first reported as a regulator of bacterial cellulose production in a landmark paper by Benziman in 1987.7 G. xylinum is a gram-negative bacteria that
produces pure cellulose as part of its biofilm extracellular matrix. c-di-GMP binds to the
membrane bound cellulose synthase enzyme, which stimulates the production of the
cellulose. The rate of cellulose production is directly correlated with the intracellular
concentration of c-di-GMP.
In order to further probe the binding site and activation of cellulose synthase by c-
di-GMP, van Boom and coworkers synthesized 13 analogs of c-di-GMP, including
dimers and trimers using different nucleobases, and monothiophosphate analogs.8 Upon
examination of the binding affinity to cellulose synthase and degradation rate by a
phosphodiesterase (PDE), it was shown that 7 out of 13 had affinity for the synthase and
stimulated its activity. c-di-GMP, c-dGMP-GMP and the monothiophosphate [Rp]-c-di-
GMP had the strongest binding. The cyclic-di-xanthosine monophosphate analog (c-di-
XMP) was not found to be an allosteric activator of the synthase, but was degraded by the specific PDE for c-di-GMP.
1.3 c-di-GMP Regulates Biofilm Production
It is now known that c-di-GMP is ubiquitous among many different species of
bacteria, particularly gram-negative type. Echerichia coli,9 Salmonella typhimurium,
Caulobacter cresentus,10 Vibrio cholerae,11 Yersinia pestis12 and Pseudomonas
aeruginosa4 all possess the cellular machinery that uses c-di-GMP to regulate the
5
transition between a stationary state and a motile one, among many other functions that
will be described.
Despite being a known regulator of biofilm formation, among other associated
cellular functions, the control mechanisms of c-di-GMP are not fully understood. The
small molecule exerts control through different stages of gene expression, including
transcription, translation, post-translational activities with regulatory and metabolic
enzymes.1 c-di-GMP regulates the transition between the sessile state and the motile state
in specific differentiated cells within a growing colony. High cellular concentrations
results in exopolysaccharide production for biofilms, where cells can assume a stationary
state with exterior stalk or pili formation. Low concentrations have been associated with the motile state with the development of motility surface organelles, such as flagella, and the removal of surface attachment organelles.10 Thus the bacterial phenotype expressed
depends on its microenvironment, which is mediated through dynamic c-di-GMP cellular
concentrations.
Biofilm can serve as a bacterial adaption to the environment, depending on the
infection stage of the host. For example, the biofilm of certain species is associated with the non-infectious state. Vibrio cholerae and Xanthomonas campestris secrete biofilm to
adhere to environmental surfaces outside the host as a protection measure; but then
decrease biofilm production to switch to a motile, planktonic state during host infection.4
Other species use biofilms as a means to increase their virulence in the infectious
state. For example, Staphylococci use biofilms as a defense against host defenses and
antibiotics with infections involving prosthetic medical implants in vivo. Pseudomonas
6
aeroginosa also use antibiotic-resistant biofilms during the chronic pulmonary infections
of cystic fibrosis patients.4
1.4 c-di-GMP Regulates Virulence and Pathogenesis
The role of c-di-GMP in virulence and pathogenesis is to mediate between acute
and chronic pathogenic infections.1 c-di-GMP affects the expression of virulence factors
in Vibrio cholerae, Salmonella typhimurium, and Pseudomonas aeruginosa. In these
examples, the expression of these factors are inhibited by increased levels of c-di-GMP.
For example, Salmonella contains a gene encoding for an EAL-domain
phosphodiesterase (PDE) enzyme that is required against the host’s defensive phagocytic
oxidase. This enzyme specifically hydrolyzes c-di-GMP, which in turn shifts the bacteria
to the motile, more virulent state.13 Similarly, Vibrio cholerae has a multi-component
response regulator VieSAB that is required for the expression of factors that encode
cholera toxin. This protein complex contains an EAL-domain protein that when activated
ensures the low concentrations of c-di-GMP needed for virulence factor upregulation.10
1.5 Enzymes that Synthesize and Degrade c-di-GMP
Cellular levels of c-di-GMP are controlled by the enzymes that synthesize and
degrade the molecule. Diguanylate cyclase (DGC) synthesizes c-di-GMP from two molecules of guanosine triphosphate (GTP), using a conserved GGDEF catalytic domain.
This enzyme’s GGDEF domain is 180 residues in length, and is named after the conserved amino acid sequence (Gly-Gly-Asp-Glu-Phe) contained in the active site. The active enzyme consists of a protein homo-dimer, each containing the GGDEF domain.14
7
This class of cyclase also contains an inhibition site (I-site) that enables allosteric product
inhibition, which prevents the overconsumption of the reactant GTP. The mechanism of
DGC catalysis in forming c-di-GMP involves bringing 2 moles GTP close together in the
active site with the use of Mg2+, followed by the cyclization to form c-di-GMP. The 3’-
OH of one of the GTP is then deprotonated via assistance by a neighboring glutamic acid residue, to attack the alpha-phosphate of the other GTP moiety.1
PDEs assigned to hydrolyze c-di-GMP contain an EAL domain, so named after the conserved amino acid sequence required for activity (Glu-Ala-Leu). The c-di-GMP is first cleaved to linear dimer pGpG, which requires Mg2+ or Mn2+ for activity, and is then cleaved to guanosine monophosphate (GMP) by non-specific cellular PDEs.14 This reaction is strongly inhibited by the presence of Ca2+ or Zn2+.2 Another PDE enzyme is
the HD-GYP-domain-containing hydrolase, which can directly cleave c-di-GMP to pGpG, then to 2 moles of GMP. This enzyme is so named from the histidine-dependant subgroup of metal dependent phosphodiesterases.1, 14 Here is a case where nature displays
a functional redundancy with the existence of two separate PDE enzymes for the same
substrate.
These GGDEF and EAL domain proteins regulate the intracellular levels of c-di-
GMP, which in turn controls the phenotype for motility. Both domains are commonly
found in many phylogenetically different bacterial species, usually occurring in tandem
on the same protein, with only one as the working output domain.4 These domains
typically display the least amount of amino acid variation, as shown by bacterial genome
sequencing.
8
The GGDEF/EAL proteins usually have an N-terminal sensor domain that is
responsive to extracellular signals or stimuli. For example, auxiliary sensor domains that
are responsive to oxygen and redox conditions, UV light, nutrient scarcity, polyamines,
and heme or flavin cofactors have been associated with these proteins.2, 10, 14 In contrast,
some of these proteins have lost their catalytic ability, having variations in the key amino acid residues. For example, E. coli possesses an EAL-domain containing protein, YcgF, that has lost the functioning PDE activity; however, it serves to direct macromolecular interactions. In C. crescentus, a GEDEF domain protein lacks DGC activity, but binds to
GTP which in turn enhances the PDE activity located on the same proteins C-terminal domain.10 All known degenerate GGDEF/EAL proteins are still connected with biofilm
and motility functions of the cell.14
1.6 c-di-GMP Target Receptors
There are five biomacromolecular receptors that are known to be a target for c-di-
GMP for signal transduction: the PilZ family proteins, FleQ transcription factor, PelD, the I-site in GGDEF proteins, and the GEMM riboswitch.14 The PilZ family of proteins is
the most widely studied c-di-GMP receptor since it is the most abundant, and is found in
most bacterial species.15 It contains an allosteric binding site that induces a
conformational shift of the holo-protein, which then can affect the receptor proteins output signal, such as cellulose or alginate production.14
The crystal structure of two PilZ domain proteins, one in Vibrio cholerae
VCA0042, and one in Pseudomonas putida PP4397 was recently reported that illustrated the diversity of c-di-GMP binding modes (Figure 1-1, a-b).15 In the VCA0042 dimer
9
protein, there is a single binding site for one c-di-GMP molecule, that is sandwiched
between the PilZ and the YcgR-N domains (Figure 2-1, a). This protein exists as a dimer
in an open conformation, and upon binding to c-di-GMP closes to surround the ligand. A
leucine residue within the binding site serves as a hydrophobic “cap”, which possibly
excludes another molecule of c-di-GMP from binding.
In contrast, the PP4397 dimer of P. putida contains two self-intercalated c-di-
GMP molecules in its binding site (Figure 2-1, b). The binding event causes a dimer to
monomer transition with the protein subunits. There are two arginine residues, whose
positively charged side chains cause favorable interactions with the c-di-GMP aromatic
bases.
a) b)
Figure 2-1: PilZ Domains of VC0042 and PP4387: a) VCA0042 PilZ domain showing unimolecular binding mode of c-di-GMP; b) PP4387 PilZ domain showing the bimolecular binding mode of c-di-GMP.15 Images from Protein Data Bank at Rutgers University.
Another c-di-GMP receptor is the FleQ transcription factor in P. aeruginosa,
which is known to upregulate the genes encoding for flagella formation, as well as
represses those proteins required for exopolysaccharide synthesis. The PelD receptor in
10
the same P. aeruginosa also contains a c-di-GMP binding site, and induces
exopolysaccharide synthesis upon binding.14 The PelD receptor in Caulobacter
crescentus, a GGDEF domain protein, upon c-di-GMP binding was found to cause a loss
of motility and a change in morphology for the swarmer-type cells.16 The crystal structure
of the PelD was also found to be a high affinity binding site for non-competitive product
inhibition (I-site) of the DGC activity.1 The I-site is an allosteric binding site present on
DGC for negative feedback control during c-di-GMP synthesis.2
The most recently discovered receptor for c-di-GMP is the GEMM riboswitch in bacteria.17, 18 Riboswitches are regulatory sequences of RNA found on the upstream 5’-
untranslated region of a gene that can bind to a ligand directly without protein factor
intermediaries for translational control.19 They contain two areas: a metabolite sensing
domain that is the ligand binding site, and the gene expression signal that changes
conformation on ligand binding for controlling protein synthesis. The GEMM riboswitch
is found in bacterial species, like anthrax and cholerae, usually preceding genes that code for DGCs, PDEs, pilus assembly, motility, virulence factors and chemotaxis sensing.6, 20
A reported study using a 110 nucleotide length RNA aptamer construct of vc2 in V. cholerae was used to determine the binding affinity to c-di-GMP.17 The Vc2 riboswitch is
upstream from a gene coding for a sugar-binding protein that is a key factor for V.
cholerae’s colonization of mammalian intestines. Based on their results using an “in-line”
probing technique that monitored the extent of cleavage at RNA specific sites, the paper
reports that the riboswitch construct and c-di-GMP form a 1:1 saturable complex, that has
a Kd of ~1 nM. This binding affinity is three orders of magnitude stronger than c-di-GMP
21 binding to a PilZ domain (Kd = 840 nM).
11
The crystal structure of the Vc2 riboswitch construct shows one molecule of c-
di-GMP ligand binds to the site located at the edge of the three helical junction aptamer
(Figure 2-2).6, 20 This gene is associated with the rugose-phenotype capable of increased
biofilm production. Both reports indicate that each guanine interacts differently to the
RNA, where one guanine forms Watson-Crick base pairs with the aptamer C92 and the
other one forms a Hoogsteen base pair with G20 (Figure 2-2). The two c-di-GMP
guanines participate in an extended stacking with RNA purine bases from the P1 and P2
helices, but not with each other.20 These stacking interactions with the RNA purines,
particularly A47 between the two guanines from c-di-GMP, are more conserved than the
base-pairing nucleotides.18 The 12-membered ring, containing the two phosphodiester
bonds, is mostly exposed to solvent in the major groove of the helix; however upon
binding, the majority of the solvent-accessible surface of c-di-GMP becomes buried
within the riboswitch.6 In one report, the kinetics of binding were determined using a gel shift assay to measure the RNA-c-di-GMP interaction using radio-labeled c-di-GMP.20 It
was found the complex had vastly different on and off rates with a calculated Kd of 10
pM. The group hypothesized that the riboswitch must be kinetically controlled since the equilibrium is strongly shifted to the ligand-riboswitch complex.20
12
Figure 2-2: V. cholerae vc2 riboswitch in complex with c-di-GMP, 3.2 Ǻ crystal structure.6 Images from Protein Data Bank at Rutgers University.
1.7 Temporal and Spatial Regulation of c-di-GMP Levels
As a part of a complicated system of bacterial signal transduction, the cellular concentration of c-di-GMP must be tightly controlled. The presence of numerous receptors with varying degrees of binding affinity, as well as a number of DGC and PDEs contribute to the temporal and spatial control of c-di-GMP concentration.3 A recent publication used a genetically coded fluorescence resonance energy transfer biomarker attached onto a PilZ domain protein to monitor the c-di-GMP binding event. The unbound PilZ displayed a large detected emission while the c-di-GMP-bound PilZ showed a fluorescence quenching, presumably due to the conformational shift of the protein upon the binding event. During cellular mitosis, C. cresentus was shown to have
13
an asymmetric distribution of c-di-GMP in the daughter cells, resulting in a c-di-GMP-
rich stalked cell and a c-di-GMP-poor swarmer cell. This is a non-random event and is
seen in other species, such as Pseudomonas, Salmonella, and Klebsiella.3 The spatial
segregation of c-di-GMP within a single cell can allow for separate and parallel operating
c-di-GMP dependent pathways affecting different targets. For example, C. crescentus
uses spatial compartmentalization during the transition from motile swarmer to sessile
state by localizing the PelD DGC enzyme from the cytoplasm to the cell pole for the
flagellum removal and stalk formation.2
Spatial distribution of c-di-GMP involving its storage as the protein-bound form can be a mechanism by which the bacterial cell controls the concentration, in addition to
regulating its synthesis and degradation. Benziman reported the isolation and
characterization of a c-di-GMP binding protein found in the cell membrane in G.
xylinum, whose equilibrium is strongly dependent on the presence of K+.22 It was discovered when c-di-GMP was in the bound state with K+, it was unavailable to act as
the activator of cellulose synthase, which points to a potential storage mechanism. The
nature of the binding was found to be non-cooperative and Mg2+-dependent, having a Hill coefficient of ~1. Additionally, the Kd was determined in this study to be 25 nM, which indicates most of the c-di-GMP in G. xylinum would be in the bound state since the
cellular K+ concentration is ~100 mM.
Additional details of note are that in the absence of K+, 45% of the bound c-di-
GMP was released; and in its presence, only 8% was free. This phenomenon was not seen
with other monovalent cations, such as Na+, Li+, Cs+, and Rb+. Since it is widely known
that guanosine and guanosine-rich ribonucleotides, like c-di-GMP, can readily form
14
stable quartet structures in the presence of K+ metal,23, 24 it is possible that c-di-GMP is
stored in higher order structures with K+ within the cell that is bound to a storage protein.
1.8 Effects of c-di-GMP on Mammalian Cells
Independent reports have shown that in vitro cancer cells treated with c-di-GMP
undergo proliferation inhibition. Amikam demonstrated that c-di-GMP caused a marked increase in (H3) thymidine uptake in human lymphoblastic and T-cell leukemia cells.25
These cells were shown to have been locked into the S-phase of the cell cycle, when
DNA is replicated. This in turn hampered cell division, leading to smaller cells with greater DNA content. In a later study in 2005, Karaolis reported that c-di-GMP exerted a negative impact on basal cell proliferation and epidermal- and acetylcholine growth factor-induced proliferation in human colon cancer cell line H508.26
c-di-GMP has also been shown effective in stimulating mammalian hosts innate
immunity against extracellular bacterial pathogens. This has potential for using c-di-GMP
as a vaccine adjuvant for immunostimulation. In a 2007 study, mice that were pretreated
with intranasal or subcutaneous c-di-GMP 24 hours prior to exposure to Klebsiella
pneumoniae had an increased survival rate due to increased levels of neurtophils, alpha-
beta-T cells and -lymphocytes.27 Similar findings were found with mice that underwent c-
di-GMP pre-treatment experienced a decrease in pnuemonialcoccal loading in the lung
after serotype 2 and 3 challenge, and overall lower lung titer after serotype 4 challenge.28
In a separate study, coadministration of c-di-GMP and pnuemonialcoccal surface adhesion A elicited a strong inflammatory response in mice due to the activation of the antigen-specific serum immunoglobulin G and the secretory IgA response.29 As the
15
above data indicate, c-di-GMP has been shown to hold promise for use in host defense
against pathogenic bacterial mucosal infections.
1.9 References
1. Jenal, U., Malone, J., "Mechanisms of Cyclic-di-GMP Signaling in Bacteria". Annual Review of Genetics, 2006, 40, 385-407.
2. Tamayo, R., Pratt, J., Camilli, A., "Roles of Cyclic Diguanylate in the Regulation of Bacterial Pathogenesis". Annual Reviews of Microbiology, 2007, 61, 131-148.
3. Christen, M., Kulasekara, H., Christen, B., Kulasekara, B., Hoffman, L, Miller, S. , "Asymmetrical Distribution of the Second Messenger c-di-GMP upon Bacterial Cell Division". Science, 2010, 328, 1295-1297.
4. Cotter, P., Stibitz, S., , "c-di-GMP-Mediated Regulation of Virulence and Biofilm Formation". Current Opinion in Microbiology, 2007, 10, 17-23.
5. Hammer, B., Bassler, B., "Distinct Sensory Pathways in Vibrio Cholerae El Tor and Classical Biotypes Modulate Cyclic Dimeric GMP Levels To Control Biofilm Formation". Journal of Bacteriology, 2009, 191, 169-177.
6. Kulshina, N., Baird, N.J., Ferre-D'Amare, A.R., "Recognition of the Bacterial Second Messenger Cyclic Diguanylate by it's Cognate Riboswitch". Nature Structural and Molecular Biology, 2009, 16(12), 1212-1217.
7. Ross, P., Weinhouse, H., Aloni, Y., Michaeli, D., Weinberger-Ohana, P., Mayer, R., Sraun, S., de Vroom, E., van der Marel, G.A., van Boom, J.H., Benziman, M., "Regulation of Cellulose Synthesis in Acetobacter xylinum by Cyclic Diguanylic acid". Nature, 1987, 325, 279-281.
8. Ross, P., Mayer, R., Weinhouse, H., Amikam, D., Huggirat, Y., Benziman, M., de Vroom, E., Fidder, A., de Paus, P., Slieregt, L., van der Marl, G., van Boom, J., "The Cyclic Diguanylic Acid Regulatory System of Cellulose Synthesis in Acetobacter xylinum". Journal of Biological Chemistry, 1990, 265(31), 18933- 18943.
9. Simm, R., Morr, M., Kader, A., Nimtz, M, Romling, U., "GGDEF and EAL Domains Inversely Regulate Cyclic di-GMP Levels and Transition from Sessility to Motility". Molecular Microbiology, 2004, 53(1123-1134).
16
10. Romling, U., Amikan, D., "Cyclic di-GMP as a Second Messenger". Current Opinion in Microbiology, 2006, 9, 218-228.
11. Tischler, A., Camilli, A., "Cyclic Diguanylate (c-di-GMP) Regulates Vibrio cholerae Biofilm Formation." Molecular Microbiology, 2004, 53(3), 857-869.
12. Bobrov, A., Kirillina, O., Perry, R., "The Phosphodiesterase Activity of the HmsP EAL Domain is Required for Negative Regulation of Biofilm Formation in Yersinia pestis". FEMS Microbiology Letters, 2005, 247, 123-130.
13. Hisert, K.B., MacCoss, M., Shiloh, M.U., Darwin, H., Singh, S., Jones, R.A., Sabine, E., Zhang, Z., Gaffney, B.L., Gandotra, S., Holden, D.W., Murray, D., Nathan, C., "A Glutamate-Alanine-Leucine (EAL) Domain Protein of Salmonella Controls Bacterial Survival in Mice, Antioxidant Defence and Killing of Macrophages: Role of cyclic diGMP". Molecular Microbiology, 2005, 56(5), 1234-1245.
14. Hengge, R., "Principles of c-di-GMP Signalling in Bacteria". Nature Reviews. Microbiology, 2009, 7, 263-273.
15. Ko, J., Ryu, K., Kim, H., Shin, J., Lee, J., Cheong, C., Choi, B., "Structure of PP4397 Reveals the Molecular Basis for Different c-di-GMP Binding Modes by Pilz Domain Proteins". Journal of Molecular Biology, 2010, 398, 97-110.
16. Hecht, G.B., Newton, A., "Identification of a Novel Response Regulator Required for the Swarmer-to-Stalked-Cell Transition in Caulobacter crescentus. " Journal of Bacteriology, 1995, 177, 6223-6229.
17. Sudarsan, N., Lee, E. R., Weinberg, Z., Moy, R. H., Kim, J. N., Link, K. H., Breaker, R. R., "Riboswitches in Eubacteria Sense the Second Messenger Cyclic Di-GMP". Science, 2008, 321, 411-413.
18. Smith, K.D., Lipchock, S.V., Livingston, A.L., Shanahan, C.A., Strobel, S.A., "Structural and Biochemical Determinants of Ligand Binding by the c-di-GMP Riboswitch". Biochemistry, 2010, 49, 7351-7359.
19. Batey, R.T., "Structures of Regulatory Elements in mRNAs". Current Opinion in Structural Biology, 2006, 16, 1-8.
20. Smith, K.D., Lipchock, S.V., Ames, T.D., Wang, J., Breaker, R.R., Strobel, S.A., "Structural Basis of Ligand Binding by a c-di-GMP Riboswitch". Nature Structural and Molecular Biology, 2009, 16(12), 1218-1223.
21. Ryjenkov, D.A., Simm, R., Romling, U., Gomelsky, M., "The PilZ Domain Is a Receptor for the Second Messenger c-di-GMP". Journal of Biological Chemistry, 2006, 281(41), 30310-30315.
17
22. Weinhouse, H., Sapir, S., Amikam, D., Shilo, Y., Volman, G., Ohana, P., "c-di- GMP-Binding Protein, a New Factor Regulating Cellulose Synthesis in Acetobacter xylinum". FEBS Letters, 1997, 416, 207-211.
23. Davis, J.T., "G-Quartets 40 Years Later: From 5'-GMP to Molecular Biology and Supramolecular Chemistry". Angewandte Chemie, 2004, 43, 668-698.
24. Zhang, Z., Kim, S., Gaffney, B.L., Jones, R.A., "Polymorphism of the Signaling Molecule c-di-GMP". Journal of the American Chemical Society, 2006, 128(21), 7015-7024.
25. Amikam, D., Steinberger, O., Shkolnik, T., Ben-Ishai, Z., "The Novel Cyclic Dinucleotide 3'-5' Cyclic Diguanylic Acid Binds to p21ras and Enhances DNA Synthesis But Not Cell Replication in the Molt 4 Cell Line". Biochemical Journal, 1995, 311, 921-927.
26. Karaolis, D.K., Cheng, K., Lipsky, M., Elnabawi, A., Catalano, J., Hyodo, M., Hayakawa, Y., Raufman, J.P., "3'-5'-Cyclic Diguanylic Acid (c-di-GMP) Inhibits Basal and Growth Factor-Stimulated Human Colon Cancer Cell Proliferation." Biochemical Biophysical Research Communications, 2005, 329, 40-45.
27. Karaolis, D.K., Newstead, M.W., Zeng, X.Y., Hyodo, M., Hayakawa, Y., Bhan, U., Lian, H., Standiford, T.J., "Cyclic Di-GMP Stimulates Protective Innate Immunity in Bacterial Pneumonia". Infection and Immunity, 2007, 75(10), 4942- 4950.
28. Ogunniyi, A.D., Paton, J.C., Kirby, A.C., McCullers, J.A., Cook, J, Hyodo, M., Hayakawa, Y., Karaolis, D.K., "C-di-GMP is an Effective Immunomodulator and Vaccine Adjuvant Against Pneumococcal Infection". Vaccine, 2008, 26(36), 4676-4685.
29. Yan, H.B., LuoLee, R., Tram, K., Qui, H.Y., Zhang, J.B., Patel, G.B., Chen, W.X., "3'-5'-Cyclic Diguanylic Acid Elicits Mucosal Immunity Against Bacterial Infection". Biochemical Biophysical Research Communications, 2009, 387(3), 581-584.
18
2. Synthetic Background of c-di-GMP and Analogs
2.1 c-di-GMP Synthetic Introduction
The internucleotide phosphodiester linkages of c-di-GMP are formed primarily via three different intermediates: phosphotriesters, H-phosphonates, and phosphoramidites (Figure 2-3). The phosphotriester method is the oldest, relying on a condensing reagent to activate the coupling of a ribose hydroxyl to a hard, electrophilic
P(V) phosphorus center. The H-phosphonate and the phosphoramidite methods are faster, owing to the P(III) oxidation state of the phosphorus center.1 There are a few reports of
other methods to synthesize c-di-GMP and analogs that will be described further on.
a) Phosphotriester b) H-phosphonate c) Phosphoramidite
B' B' B' DMTO DMTO DMTO O O O
O R O R O R Ar O P O H P O P CN (iPr)2N O O- O-
1) coupling: 1) coupling: 1) coupling: e.g. tetrazole, pyr-TFA e.g. sulfonyl chloride e.g. acid chloride 2) oxidation 2) oxidation 3) P-deprotection 2) P-deprotection
B' O O R=H,OH
O R B' O P O O O
O R
Figure 2-3: The three main synthetic methods of internucleotide couplings: a) phosphotriester, b) H- phosphonate, c) phosphoramidite.
19
2.2 Phosphotriester Chemistry
The phosphotriester methodology is the older of the methods toward making phosphodiester bond connections and is quite common in the earliest synthetic methods for preparation of cyclic ribonucleotides.2, 3 This approach is used infrequently due in part
to the P(V) oxidation state of the phosphorus center, which leads to slow reaction kinetics
and, by extension, longer reaction times and the generation of unwanted side products.
The linear dimer 3 (Scheme 2-1) was prepared via an initial coupling of a free 5’-OH nucleotide 2 and a 3’-protected phosphate diester nucleotide 1 using a condensing agent
(e.g. triisopropylbenzene sulfonyl chloride (TPS) and tetrazole).2 This was followed by
similar conditions as shown in Scheme 2-1 to cyclize the dimer to generate 4 containing
the 12-membered ring. These conditions were also used to synthesize cyclic-di- adenosine-monophosphate (c-di-AMP), cyclic-di-uridine-monophosphate (c-di-UMP), and the unsymmetric dimer c-AMP-UMP.
Scheme 2-1: Phosphotriester method for synthesis of cyclic ribonucleosides. B = adenine or uracil
20
Alternatively, a P(V) bifunctional benzotriazole-based phosphorylating reagent
was used to install the protected phosphate or thiophosphate between the free 5’OH of
one nucleoside and the 3’-OH of the second nucleoside moiety for the linear dimer.3, 4
Cyclization was achieved with this condensing reagent (Figure 2-4, 5 or 6) that closed the 5’-OH with the 3’-phosphate diester to synthesize a variety of dimers and trimers including cyclic-inosine-monophosphate-guanosine-monophosphate (c-IMP-GMP), cyclic-di-deoxyguanosine-monophosphate (c-di-dGMP), cyclic-xanthine- monophosphate-guanosine-monophosphate (c-XMP-GMP) and the monothiophosphate of c-di-GMP.3
A bifunctional phosphorylation reagent (Figure 2-4, 7) was reportedly used to
simultaneously install the phosphates and cyclize the abasic intermediate (Scheme 2-2,
12→13), followed by addition of the nucleobases via a modified Vorbruggen reaction
(13→15), resulted in c-di-GMP 16, cyclic-adenosine monophosphate (c-di-AMP), cyclic- thymidine-monophosphate (c-di-TMP) and c-di-theophylline-monophosphate (not shown).4
Figure 2-4: Condensing reagents used in phosphotriester chemistry: 5-7) hydroxybenzotriazole phosphate derivatives for phosphate addition and coupling for linear dimer, 8) 1-(mesitylene-2-sulfonyl)-3-nitro-1,2,4- triazole (MSNT) used for triester cyclization, 9) bisfunctional phosphorylation reagent for phosphate addition and cyclization.
21
TBSO O Cl HO OH O O O O O 10 O O P O O O O 1) 7,THF O P O 9, pyridine O + O O O 2) CAN, MeOH O O Cl O O O P HO O O O OH O O OH O Cl O 12 13 11
Cl O -O G 1) AcOH, Ac O, H SO O P O O 2 2 4 O G' 1) pyridine-2-carbaldoxime, 2) 14,BSA P O O O tetramethylguanidine OH O OH OAc O 3)TMSOTf 2) NH OH O O OAc 4 O P - G O O O P O G' O O 16 Cl 15 NO2
O BSA = bis(trimethylsilyl)acetamide N NH 14 = N N NH H 2
Scheme 2-2: Phosphotriester method for synthesis of c-di-GMP: formation of an abasic cyclic backbone, followed by modified Vorbruggen reaction to install the nucleobases, and deprotection to yield c-di-GMP, 16.
2.3 H-Phosphonate Chemistry
The H-phosphonate method is another class of reactions to form the phosphate
linkages between nucleotides.1 This requires the use of the H-phosphonate reactive
center, generally found on the 3’-OH terminus of the reacting nucleotide. The H-
phosphonate can be linked to a free 5’-OH via activation by an acyl chloride (e.g. pivaloyl chloride (PvCl), adamanoyl chloride (AdCl) and others in pyridine). The
advantage of this method is the increased reactivity over that of the phosphotriester
chemistry. The phosphate functionality exists in a tautomeric equilibrium, that is shifted
22
to the mildly electrophilic tetracoordinate phosphonate species (Scheme 2-3, 17) from the
nucleophilic tricoordinate phosphonate species (Scheme 2-3, 18).1 Species 18 is slightly
basic and a soft nucleophile; however, when protonated it can react with other nucleophiles. Species 17 is slightly more stable because of the presence of the phosphoryl group, making this tautomer less prone to oxidation since it lacks the lone electron pair.
Therefore, the H-phosphonate is both a soft nucleophile and soft electrophile, and the subsequent reactions with either form are fairly rapid.
In general for reactions involving H-phosphonate chemistry, it has been empirically observed that the reaction time is significantly shorter compared to the triester method, and that the formation of side products is reduced. The H-phosphonate species is fairly stable toward self-oxidation, therefore it is a slightly more forgiving method than the phosphoramidite methodology that will be described in the following section.
Scheme 2-3: H-phosphonate tautomeric equilibrium
There are reports in the literature for the synthesis of cyclic ribonucleotides that exclusively use the H-phosphonate method for both the linear dimer coupling and for the
23
cyclization.5-7 One report discloses the synthesis of the dithiophosphate analog for cyclic-
di-cytosine monophosphate (c-di-CMP) that uses PvCl as an activator.5 Two recent publications from Yan et. al disclose the use of H-phosphonate chemistry to synthesize c-
di-GMP 16 as well as the mono- and dithiophosphate analogs. Other notable features of
this method include using the 5’-O-(9-phenyl-xanthen-9-yl) protecting group (Px) along with the 2’-O-(1-(4-chlorophenyl)-4-ethoxypiperidine-4-yl) (Cpep) protecting group on the guanosine moieties (Scheme 2-4).6, 7
G''
PxO O O G'' G'' - G O P O O HO O OCpep PxO O 1) PvCl, 21 1) TFA O O PhS P O G'' OH OH 2) 3'-OH deprotection O O O O OCpep OR OCpep O O O 2) PhO P OPh O P O- HO P O G 3) PvCl, PhO P O Cl O H O OCpep 3) 21 H 4) deprotection HO P O 20 19 H 16 22
Scheme 2-4: H-phosphonate method used for synthesis of c-di-GMP for both linear dimer coupling, and cyclization using an acid chloride as an activator.
2.4 Combination of Phosphotriester, Phosphoramidite and H-Phosphonate
Chemistry
The majority of methods to synthesize c-di-GMP, its analogs and other cyclic
ribonucleotides involve the phosphoramidite chemistry for the initial coupling which is
followed by either the phosphotriester method8-10 or H-phosphonate method to achieve
cyclization.11-14
24
Hayakawa reported the use of the phophoramidite/phosphotriester method to
synthesize the unmodified c-di-GMP 16. The initial protected linear dimer 25 was
assembled via coupling of protected guanosine phosphoramidite 23 coupled to a free 5’-
OH on 3’-allylic-protected guanosine nucleotide 24 with imidazolium perchlorate (IMP)
as an activator (Scheme 2-5). Oxidation was accomplished using 2-butanone peroxide
(BPO) followed by 5’ detritylation with dichloroacetic acid (DCA). The subsequent
triester cyclization was accomplished using TPS and N-methylimidazole (NMI) to
generate the protected triester of c-di-GMP. c-di-GMP 16 was then produced via
cleavage of the allylic groups and reverse-phase high performance liquid chromatography
(RP HPLC) purification.8, 9
In a subsequent paper using similar chemistry, the monothiophosphate of c-di-
GMP, c-GMP-AMP, and c-GMP-IMP was synthesized using tert-butyl hydroperoxide
(tBuOOH) as the oxidant and bis[3-(triethoxysilyl)propyl] tetrasulfide as the
corresponding sulfurizing reagent.10 A few years later, this was followed by the synthesis of c-di-UMP, c-di-IMP, and c-IMP-UMP.15
Scheme 2-5: Phosphoramidite/phosphotriester methods for synthesis of c-di-GMP: phosphoramidite coupling followed by phosphotriester cyclization.9
25
There is one report of a solid-phase synthesis of c-di-GMP that utilizes a phosphoramidite coupling followed by a phosphotriester cyclization (Scheme 2-6). The linear dimer possessed methyl phosphate protection on the 3’ terminus bound to the controlled-pore glass (CPG) support, while the 5’ moiety possessed a cyanoethyl phosphate protecting group. After selectively deprotecting the 3’-guanosine methyl phosphate using S2Na2 leaving the 5’ guanosine cyanoethyl group intact, the cyclization was achieved with 1-mesitylenesulfonyl-3-nitro-1,2,4-triazole (MSNT) in pyridine while attached to the CPG, resulting in c-di-GMP after cleavage from the support.16 A second generation solid-phase method used symmetrically protected methyl-phosphates, using a phosphoramidite coupling producing the linear dimer, followed by cleavage from the support, then cyclization in solution-phase which increased the yields from the first iteration (Scheme 2-6, 26 → 16).
G' HO O O O OTBS G MeO P O O O O P OMe G' O ODMT SPS 1) TEA in MeCN O S OH O OH O O O 2) MSNT in pyr O O P OMe 3) TEA-HF G O OTBS O O P OMe O 16 O O S O
26
Scheme 2-6: Solid-phase synthesis of c-di-GMP.
26
The Jones lab reported using the phosphoramidite/H-phosphonate method for the synthesis of c-di-GMP initially in 2004 (Scheme 2-7).11 This method began from the
protected guanosine 27, from which the standard cyanoethyl-protected phosphoramidite
29 and the 3’-O-phosphonate 30 was formed. The coupling of 29 and 30 required
activation with pyridinium trifluoroacetate (pyr-TFA) followed by oxidation with t-
BuOOH to give the linear dimer 31. 16 was prepared via sequential detritylation of the 5’
end, cyclization using AdCl, oxidation with methanol (MeOH)/N-bromosuccimide (NBS)
and lastly deprotection of the amino groups and the 2’-hydroxyls.
G' DMTO O (iPr) N OCE 2 P G' pyr-TFA O OTBS DMTO N(iPr)2 O G' P O (iPr) N OCE G 2 O -O DMTO OTBS 1) sulfonic acid resin P O O O 29 1) pyr-TFA O P OCE G' 2) AdCl O 1) 28 OH O OH O OH OTBS 2) detritylation 2) tBuOOH O 3) MeOH/NBS 4) deprotection O O P O- G' G 27 O OTBS O HO O O P OH 16 O OTBS H O O P OH 31 O H 28 = P O 30 Cl
Scheme 2-7: Phosphoramidite/H-phosphonate methods for synthesis of c-di-GMP.11
In a later paper using similar chemistry, c-GMP-IMP and a tethered c-di-GMP
dimer were synthesized. Biophysical studies were done using NMR and UV, which
characterized the cation- and concentration- dependant quartet formation in solution.12 A few years later, a procedure was published that synthesized the seven thiophosphate
27
analogs of c-di-GMP, using the phosphoramidite/H-phosphonate chemistry with
elemental sulfur to sulfurize the phosphorus center at the appropriate steps.13
More recently, an improved one-flask method to synthesize the c-di-GMP, as well
as the dithiophosphate analogs, was reported that can produce c-di-GMP on a gram scale
using the phosphoramidite/H-phosphonate methodology (Scheme 2-8).14 Cyclic dimerization of 29 to give 35 began with hydrolysis of the reactive guanosine phosphoramidite 29 to the H-phosphonate diester 32 using pyr-TFA and water (Scheme
2-8). The β-elimination of the cyanoethyl phosphate protection on 32 was accomplished with tBuNH2 and 5’ detritylation with DCA. The phosphoramidite linear coupling of 29
and 32, using in situ formed pyridinium dichloroacetate as the activator, resulted in
linear dimer 33. This intermediate was oxidized using tBuOOH to convert the
phosphorus center from P(III) to P(V), which was followed by detritylation using DCA to
yield 34. Cyclization was achieved using an acid chloride (e.g. 5,5-dimethyl-2-oxo-2-
chloro-1,3,2-dioxaphosphinane (DMOCP), PvCl or AdCl) to join the 5’-OH with the 3’-
H-phosphonate, followed by iodine/water oxidation to give 35. Treatment of the
protected dimer 35 with tBuNH2 to remove the remaining cyanoethyl phosphate
protection gave intermediate 36, which was isolated by crystallization from methylene
2 chloride in 40% overall yield. After removal of the N -isobutryl using MeNH2 and the removal of the 2’-O-TBS groups using and TEA-HF, c-di-GMP 16 is isolated by crystallization from acetone in 79% stepwise yield, and 32% overall yield from 29.
28
Scheme 2-8: One-flask method for synthesis of c-di-GMP 16 (32 % overall yield) using the phosphoramidite/H-phosphonate method.14
2.5 Other Methods
The bis-carbamate analog of c-di-dGMP is an isosteric neutral analog that is able
to permeate membranes more readily.17 It was synthesized with N-hydroxyl succinimide
carbonate as a carbonyl donor between the 5’-NH2 and the 3’-OH of protected deoxyguanosine. The cyclization was accomplished using carbonyl diimidazole as the carbonyl donor to form the 12-membered ring, and following deprotection yielded the bis-carbamate c-di-dGMP analog.
Additionally, an enzymatic method to synthesize c-di-GMP was reported that used a thermophilic diguanylate cyclase domain that was more stable than previously reported enzymatic syntheses.18 This catalytic DGC contained a mutated I-site that
allowed for the production of c-di-GMP to continue without the drawback of product
inhibition.
29
2.6 References
1. Stawinski, J., Kraszewski, A., "How To Get the Most Out of Two Phosphorus Chemistries. Studies on H-Phosphonates". Accounts of Chemical Research, 2002.
2. Hsu, C.J., Dennis, D., Jones, R.A., "Synthesis and Physical Characterization of Bis 3'-5' Cyclic Dinucleotides (NpNp): RNA Polymerase Inhibitors". Nucleosides & Nucleotides, 1985, 4(3), 377-389.
3. Ross, P., Mayer, R., Weinhouse, H., Amikam, D., Huggirat, Y., Benziman, M., de Vroom, E., Fidder, A., de Paus, P., Slieregt, L., van der Marl, G., van Boom, J., "The Cyclic Diguanylic Acid Regulatory System of Cellulose Synthesis in Acetobacter xylinum". Journal of Biological Chemistry, 1990, 265(31), 18933- 18943.
4. Amiot, N., Heintz, K., Giese, B., "New Approach for the Synthesis of c-di-GMP and Its Analogues". Synthesis, 2006, 24, 4230-4236.
5. Battistini, C., Fustinoni, S., Brusca, M.G., Borghi, D., "Stereoselective Synthesis of Cyclic Dinucleotide Phosphorothioates". Tetrahedron Letters, 1993, 49(5), 1115-1132.
6. Yan, H., Aguilar, A.L., "Synthesis of 3',5'-Cyclic Diguanylic Acid (cdiGMP) Using 1-(4-chlorophenyl)-4-ethoxypiperdin-4-yl as a Protecting Group for 2'- hydroxy Functions of Ribonucleotides". Nucleosides, Nucleotides, and Nucleic Acids, 2007, 26, 189-204.
7. Yan, H., Wang, X., Kuolee, R., Chen, W., "Synthesis and Immunostimulatory Properties of the Phosphorothioate Analogues of cdiGMP". Bioorganic & Medicinal Chemistry Letters, 2008, 18, 5631-5634.
8. Hayakawa, Y., Nagata, R., Hirata, A., Hyodo, M., Kawai, R., "A Facile Synthesis of Cyclic bis(3'-5')Diguanylic Acid". Tetrahedron, 2003, 59, 6465-6471.
9. Hyodo, M., Hayakawa, Y., "An Improved Method for Synthesizing Cyclic Bis(3'- 5')diguanylic Acid (c-di-GMP)". Bulletin of the Chemical Society of Japan, 2004, 77, 2089-2093.
10. Hyodo, M., Sato, Y., Hayakawa, Y., "Synthesis of Cyclic Bis(3'-5')diguanylic Acid (c-di-GMP) Analogs". Tetrahedron, 2006, 62, 3089-3094.
30
11. Zhang, Z., Gaffney, B.L., Jones, R.A., "c-di-GMP Displays A Monovalent Metal- Ion Dependent Polymorphism". Journal of the American Chemical Society, 2004, 126, 16700-16701.
12. Zhang, Z., Kim, S., Gaffney, B.L., Jones, R.A., "Polymorphism of the Signaling Molecule c-di-GMP". Journal of the American Chemical Society, 2006, 128(21), 7015-7024.
13. Zhao, J., Veliath, E., Kim, S., Gaffney, B., Jones, R.A., "Thiophosphate Analogs of c-di-GMP: Impact on Polymorphism". Nucleosides, Nucleotides, and Nucleic Acids, 2009, 28(5-7), 352-378.
14. Gaffney, B.L., Veliath, E., Jones, R.A., "One-Flask Synthesis of c-di-GMP and the [Rp,Rp] and [Rp,Sp] Dithiophosphate Analogs". Organic Letters, 2010, 12(14), 3269-3271.
15. Ching, S.M., Tan, W.J., Chua, K.L, Lam, Y., "Synthesis of Cyclic Di-Nucleotidic Acids as Potential Inhibitors Targeting Diguanylate Cyclase". Bioorganic & Medicinal Chemistry, 2010, 18, 6657-6665.
16. Kiburu, I., Shurer, A., Yan, L., Sintim, H.O., "A Simple Solid-Phase Synthesis of the Ubiquitous Bacterial Signaling Molecule, c-di-GMP and Analogues". Molecular Biosystems, 2008, 4, 518-520.
17. Kline, T., Jackson, S.R., Deng, W., Verlinde, C.L., Miller, S.I., "Design and Synthesis of bis-Carbamate Analogs of Cyclic bis-(3'-5')-Diguanylic Acid (c-di- GMP) and the Acyclic Dimer PGPG". Nucleosides, Nucleotides, and Nucleic Acids, 2008, 27, 1282-1300.
18. Rao, F., Pasunooti, S., Ng, Y., Zhuo, W., Lim, L., Liu, A.W., Liang, Z., "Enzymatic Synthesis of c-di-GMP Using a Thermophilic Diguanylate Cyclase". Analytical Biochemistry, 2009, 389, 138-142.
31
3. Synthesis of C8 Analogs of c-di-GMP
3.1 Introduction
Our interest in synthesizing analogs of c-di-GMP in a divergent manner stems from investigating the guanosine quartet equilibrium with C8 base modifications of c-di-
GMP. Substituents possessing different steric and electronic properties that could alter
the formation of quartets in solution were chosen not just from a physical property
standpoint, but also from a synthetic standpoint.
The fact that c-di-GMP can be synthesized in a larger quantity using the recently
reported one-flask method facilitates this goal by providing the parent compound on a
gram scale without any chromatographic steps.1 Therefore, the utility of using the parent
compound as starting material, as opposed to using modified guanosine monomers, is
appealing for the potential to generate many compounds from c-di-GMP synthon.
3.2 Synthesis of c-di-Br-GMP
It has been long known that guanosine is regiospecifically brominated at
the C8 position using a variety of brominating reagents, such as NBS, N- bromoacetamide, and bromine in either organic or aqueous solvents.2, 3 The use of
aqueous bromine was attractive as the reagent of choice for the derivatization of both the
model compound GMP and c-di-GMP since both compounds are water-soluble.
Additionally, the incorporation of 8-bromoguanosine (8-Br-G) is known to induce the syn conformer about the glycosidic bond due to the steric bulk of the bromine substituent 4, 5, and has been found to stabilize the Z-form of DNA (left-handed DNA helices).6 Aside
32
from providing the all syn conformer of c-di-Br-GMP, the dibrominated analog is a versatile synthon enroute to a variety of other downstream analogs.
Initial work on the 8-bromo analog started with the bromination of GMP (Na+ form) (Scheme 2-9). A freshly prepared solution of aqueous bromine (0.24 M) was necessary since the reagent lost potency over the course of a few days. The 8-Br-GMP derivative 38 was cleanly formed at room temperature on slow, small additions of the aqueous bromine (Scheme 2-9). If an excess of aqueous bromine was added initially, 38 did form, however, degradation products appeared followed by loss of UV absorption.
On prolonged exposure to the aqueous bromine, it is known that C8-bromination occurs followed by rapid oxidative-imidazole ring opening and final degradation of the chromophore altogether.3
Scheme 2-9: Synthesis of 8-Br-GMP 38.
Mechanistically, the bromination of the C8 position is presumed to be
accomplished by guanosine N7 attacking molecular bromine (Scheme 2-10, 39), with
subsequent addition of bromide at C8 40. The guanosine N7 is the most basic nitrogen on
the guanine moiety, having a pKa of 2.5 in water, which initiates the reaction sequence.
The unstable anti-aromatic intermediate containing 8 π electrons 41 is formed, which
33
rapidly undergoes elimination of HBr to generate the C8-brominated guanosine 42.
Optimization of the bromination of GMP and HPLC purification generated 8-Br-GMP in
54% yield in the ammonium form.
Scheme 2-10: Mechanism for the bromination of GMP.
Bromination of c-di-GMP was achieved in a similar manner to 8-Br-GMP using
small additions of the aqueous bromine, requiring twice the equivalents due to the two
reaction sites (Scheme 2-11). After quenching with cyclohexene and extraction with
methylene chloride, the product was purified and desalted via HPLC resulting in 82% isolated yield of c-di-Br-GMP 43 in the triethylammonium form.
O N O HN Br N NH H N N N O O 2 -O N - P O O N NH2 O P O O O O OH OH aq. Br2 (3.5 eq) OH OH O O O H2O, RT O O - O P - P O H2N N N O O 82% N N NH2 O HN Br N NH O N O 16 43
Scheme 2-11: Synthesis of c-di-Br-GMP, using aqueous bromine.
34
3.3 Synthesis of c-di-thio-GMP
The di-thiolated cyclic dimer offers a unique substituent that can exist as two
tautomers (Scheme 2-12), in addition to inducing the syn conformer, similar to the di-
bromo analog. The equilibrium is strongly shifted to the thio tautomer 45 in both
dimethylsulfoxide (DMSO) and in aqueous solvents, that was shown by 15N NMR in a
previous study.7 Therefore, the presence of this substituent could serve as a useful probe
for the formation of guanine quartets in solution.
O H O N N NH NH HS S N N N NH N NH2 2 R R 44 45
Scheme 2-12: Tautomeric equilibrium of 8-thioguanine, shifted toward the thio form.
Previous research showed that treatment of 8-Br-G with thiourea in refluxing
ethanol (EtOH) generated the 8-thioguanosine product readily and in good yield, 92%.3
More recently was reported the use of NaS2O3 in a solvent mixture of BuOH/EtOH/H2O; however, the reaction required an elevated reaction temperature of 125°C over 18 hours.8
Since the mildest conditions are the most desirable for use on the c-di-Br-GMP starting material, the method employing thiourea in EtOH was preferentially chosen due to the milder reaction conditions with that particular system.
Working with 8-Br-GMP 38 as the model system, the initial reactions were unsuccessful in generating the 8-thio-GMP product (Scheme 2-13). Refluxing at 90°C in aq. EtOH using 1.3 eq of thiourea over 20 hours resulted in dephosphorylation of the 8-
35
Br-GMP with only trace formation of the desired 8-thio-GMP product. Using H2O as the
solvent and increasing the thiourea to 2 eq with a gradual increase of temperature from
50°C to 100°C reflux only resulted in incremental increase in the dephosphorylated
starting material. Moving to a polar aprotic solvent dimethylformamide (DMF) there was
no sign of product formation at 85°C over 23 hours.
Scheme 2-13: Initial attempts for thiolation of 8-Br-GMP model system using thiourea in a variety of solvents.
Using the conditions reported as being successful for the thiolation of the
uncharged guanosine, it was possible that the failure of these initial attempts was due to
the anionic nature of 8-Br-GMP in terms of solubility and the repulsive forces inhibiting
the nucleophilic attack by the thiourea. Any loss of the phosphate in the 8-Br-GMP
system could indicate a ring-opening degradation on the c-di-Br-GMP substrate.
In order to make the C8 a more electrophilic center susceptible to nucleophilic attack, the protonation of the neighboring N7 with a mild acid could induce the desired reaction to occur. Pyridinium trifluoroacetate (pyr-TFA) is a commonly used activator for
nucleoside phosphoramidite couplings, where the pKa of the pyridinium cation is 5.2 in
water.9 In addition, the resultant conjugate base pyridine is nucleophilic to displace the
bromo- leaving group on 47, moving through the pyridinium intermediate 49, to then be
36
attacked by the sulfur from the thiourea for formation of the desired product (46 → 52,
Scheme 2-14).
Scheme 2-14: Proposed mechanism of the pyr-TFA catalyzed thiolation of 8-Br-GMP.
The reaction of 8-Br-GMP with 2.0 eq of thiourea and 0.5 eq of pyr-TFA in 1:1
EtOH:MeOH at reflux showed the formation of 40% of 8-thio-GMP 55 at 3 hours,
converting to 86% over 19 hours with minimal side product formation (Scheme 2-15).
In order to test the hypothesis whether an acidic pH alone would be sufficient in
catalyzing the reaction, the reaction was repeated using an aqueous MeOH solution
whose pH was adjusted to 4.0 which was 1.5 pH units lower than the original reaction.
No reaction was observed on refluxing overnight; however on addition of 1 eq of pyr-
TFA the reaction progressed to 88% complete after an additional 24 hour period.
Additionally, the most efficient solvent system was a binary solvent system of EtOH:H2O
(2:1) over any non aqueous combination of EtOH/MeOH/DMF, presumably due to solubility reasons. The optimized reaction conditions were determined to be 2 eq
37
thiourea, 0.25 eq pyr-TFA in 60% aqueous EtOH at 80-85 °C, which resulted in clean
conversion to the 8-thio-GMP 55. After HPLC purification, the isolated yield for the 8-
thio model system was 68% in the ammonium form (Scheme 2-15).
Scheme 2-15: Synthesis of 8-thio-GMP, 55.
Based on the successful results with the use of a catalytic amount of pyr-TFA to generate the 8-thio analog, similar reaction conditions were used on c-di-Br-GMP 43 for
the transformation to c-di-thio-GMP 56. Initial attempts using 4 eq of thiourea, 0.5 eq of
pyr-TFA in 85% aqueous EtOH at 85 °C over 6 hours showed product formation albeit
with a significant amount of ring-opened dithiolated linear dimer.
In order to determine the most efficient reaction temperature, a reaction using similar reagents was set up that started at room temperature and then incrementally increasing the temperature. Using LCMS to monitor both product formation and any possible accumulation of side products, the optimal temperature was observed to be 60-
65 °C, with the onset of significant degradation at 70 °C. After optimization, the final reaction conditions to produce c-di-thio-GMP 56 were 4 eq of thiourea, 0.4 eq of pyr-
TFA in 30% aqueous EtOH heated at 60 °C for 22 hours. After HPLC for purification
38
and desalting, the c-di-thio-GMP was synthesized in 65% isolated yield as the
triethylammonium form (Scheme 2-16).
O N HN SH N O H2N N - O P O O thiourea (4.0 eq), O pyr·TFA (0.4 eq), OH OH 43 O aq. ETOH, 60°C O O P O-
65% N N NH2 O HS N NH O 56
Scheme 2-16: Synthesis of c-di-thio-GMP, 56.
3.4 Synthesis of c-di-methylthio-GMP
The methylation of the thiol groups prevents the tautomerization shown in
Scheme 2-12, and therefore, should hypothetically allow the N7 to participate as a hydrogen bond acceptor. The preparation of the c-di-methylthio-GMP 57 was
accomplished by the methylation of 56 with Me2SO4 in anhydrous DMF (Scheme 2-17).
Care was taken with the addition of the methylation reagent since over methylation was possible due to the numerous nucleophilic sites on the starting material. After HPLC purification and desalting, 57 was formed in a 32% isolated yield in the triethylammonium form.
39
Scheme 2-17: Synthesis of c-di-methylthio-GMP, 57.
3.5 Synthesis of c-di-phenyl-GMP and c-di-acetylphenyl-GMP
Aryl halides serve as a convenient precursor to biaryl compounds using proven
palladium-catalyzed aryl couplings with an arylboronic acid, such as with the Suzuki-
Miyaura coupling reaction.10, 11 The Suzuki coupling reaction has several advantages that
lend itself to regiospecific reaction of a multifunctional substrate like nucleotides: 1) it
can be carried out under mild conditions 2) the reaction is tolerant to a variety of
substrate functional groups while being regiospecific, 3) many arylboronic acids are
commercially available, and 4) the reaction proceeds smoothly in aqueous solvents. All
of these aforementioned aspects are highly desirable for most synthetic conditions;
however, the last point is particularly attractive for nucleotides whose solubility is limited
to high boiling point polar aprotic solvents and water.
The arylation of 8-Br-G and 8-Br-dG has been reported on the nucleoside and
nucleotide monomeric units.12-15, 16 An aryl substituent for the C8 position was chosen to observe the effect of the additional aromatic ring on any potential π-stacking that the c-di-
GMP analog may assume in solution. Since the aforementioned publication12 referenced the use of a water-soluble phosphine ligand, tri(4.6-dimethyl-3-
40
sulfonatophenyl)phosphine (TXPTS), this became the starting point for initial synthetic
work on the model system 8-Br-GMP.
The Suzuki coupling catalytic cycle is shown in Figure 2-5. The resting state of
the Pd catalyst is oxidation state (0), with two TXPTS ligands. Oxidative addition occurs
when the catalyst inserts between the Br-C bond at the C8 position of guanine, driving
the oxidation state of Pd to (II). The metathesis step follows whereby the base exchanges
with the bromide ligand on the metal center, which is followed by transmetallation where
the aryl substituent from the boronic acid switches places with the base ligand on the Pd with concomitant loss of a ligand from the metal center. The remaining reductive elimination step involves the C-C bond forming step joining the two aromatic moieties,
and the return of one ligand to the Pd which returns the catalyst to the initial step in the
cycle and an oxidation state of (0).
41
Figure 2-5: Catalytic cycle for the Suzuki-Miyaura aryl coupling of an 8-bromoguanine moiety and an arylboronic acid. L = tri(4.6-dimethyl-3-sulfonatophenyl)phosphine (TXPTS)
The initial Suzuki reactions for preparing 8-phenyl-guanosine monophosphate (8-
Ph-GMP, Scheme 2-18, 58) found that water was superior to a binary solvent system of
water and acetonitrile. Additionally, temperatures closer to 100 °C, in refluxing water,
resulted in higher conversion to product over the cited 80 °C reaction temperatures with
8-Br-GMP and 8-Br-GTP in water.14, 16 After several rounds of optimization, the reaction
for forming 8-Ph-GMP 58 required 1.5 eq of phenylboronic acid, 4 mole % of Pd(OAc)2,
12 mole % of TXPTS ligand, 3 eq of Na2CO3 in refluxing water for 22 hours, which
resulted in 42% isolated yield of the product (Scheme 2-18).
42
O
HN N
PhB(OH)2 (1.5 eq), Pd(OAc)2 (4%) H N N N O 2 TXPTS (12%), Na2CO3 (3 eq) 38 O O - P O - H O, 100°C, 22h O 2 OH OH
42% 58
Scheme 2-18: Synthesis of 8-Ph-GMP 58.
Turning to the target c-di-phenyl-GMP (59, Scheme 2-19), the model system reaction conditions proved too harsh at 100 °C reflux, due to presence of over 40% of ring opened diphenylated dimer. The Suzuki coupling was successful, however the temperature and alkalinity of the medium hydrolyzed the phosphodiester bond of the 12- membered ring.
After exploring a range of temperatures from 50 °C up to the 100 °C endpoint, it was found that 90-95 °C was optimal in forming product while minimizing degradation.
Using conditions shown in Scheme 2-19, 59 was formed in 42% isolated yield after purification.
Scheme 2-19: Synthesis of c-di-phenyl-GMP 59 using a tandem Suzuki-Miyaura aryl coupling.
43
Synthesis of the fifth analog, c-di-3-acetylphenyl-GMP was chosen after the
biophysical NMR studies were completed on the earlier described four analogs. It was
revealed that only c-di-Br-GMP as the K+ salt form exhibited formation of higher-order
guanine quartets in solution, and to a much lesser extent as the Na+ form (described in
detail in the Biophysical Study section).
In an effort to provide additional intermolecular stabilizing interactions for
possible quartets, the 3-acetylphenyl analog was chosen. In lipophilic guanosine derivatives, this modification was shown to extend the Hoogsteen line of H-bond acceptors (carbonyl oxygen from the acetyl group on the phenyl ring) with the neighboring Watson-Crick line of H-bond donors (N2 amino group). The general overall effect strengthened the quartet stability in organic solvents (Figure 2-6).13, 15 Although
the substrates in this reference and those of this current study are very different in terms
of ionizable groups and solubility, this analog was chosen to probe the effect of this
added structural element.
44
Figure 2-6: Different G-quartets: a) unmodified guanine quartet stabilized by a network of H-bonds and ion-dipole interactions, b) arylated guanine quartet stabilized by an extended network of H-bonds provided by the additional acetyl acceptor on the C8 substituent.13
Using the synthetic conditions for 59 with 3-acetylphenyl boronic acid as a reactant, it was shown that the 95 °C temperature was much too harsh for this analog.
The diarylated compound 60 was formed but was readily degraded to the ring-opened modified linear dimer. After optimizing the temperature conditions, it was found that 80
°C over 20 hours was successful with product conversion (60, Scheme 2-20). The large proportion of the diacetylphenylated analog in the optimized reaction mixture readily crystallized from the aqueous reaction mixture after filtration of the inactivated palladium catalyst in a modest 12% isolated yield. This simplified the purification and isolation tremendously, however posed other consequences, as the product was found to be not soluble in water, buffer, acetonitrile, or a combination of these three solvents. It was found this analog was only soluble in 50% aqueous MeOH, which then had repercussions for the NMR studies that ensued.
45
Scheme 2-20: Synthesis of c-di-acetylphenyl-GMP, 60.
3.6 Conclusions
In conclusion, a divergent synthetic method was established that generated a family of C8-modified analogs of c-di-GMP (Scheme 2-21).
Scheme 2-21: Divergent synthesis for a family of C8 analogs of c-di-GMP.
46
3.7 Experimental Procedures
General Methods
Semi-preparative reverse phase HPLC purification was performed on a Waters
Novapak C18 19 x 300 mm column using gradients of acetonitrile with 0.1M
triethylammonium acetate (TEAA) buffer (pH 6.8). Desalting of pure samples was performed on a Waters Novapak C18 19 x 300 mm column using gradients of acetonitrile
with degassed Millipore water. Analytical reverse phase HPLC was performed on a
Waters 2960 system, with an Atlantis C18 column, 100 Ǻ, 4.6 mm x 50 mm, 3μm using gradients of acetonitrile with 0.1M TEAA (pH 6.8) at a flow rate of 1.0 mL/min. ESI-MS was acquired using a Waters Micromass single quadrupole LCZ system.
Triethylammonium, sodium and potassium salts of 43, 56, 57, 59, 60 were obtained by ion exchange using 15 mL of AG 50W-X2 sulfonic acid resin, which had been converted
to the TEA+, Na+ and K+ forms respectively. Maximum UV absorbance for all analogs
was determined in TEAA buffer (pH 6.8) at 25°C and 75°C on a Cary Varian
spectrometer. The extinction coefficients were determined by dissolving a known mass
of lyophilized C8-analog sample in 10mM sodium citrate-phosphate buffer (pH 6.8),
after having Karl Fischer analysis done for determining the water content. After the
appropriate dilution was made, the absorbance of the sample and the blank was measured
in a 1 cm path length quartz cell at 25°C in triplicate. The average value was used to
calculate the extinction coefficient based on Beer’s Law, ε = A/lc.
47
NMR
All the NMR spectra were acquired on a Varian Inova 500 MHz spectrometer.
1 13 The H and C NMR were referenced to sodium trimethylsilylpropylsulfonate in D2O,
31 1 and the P NMR was referenced to 10% phosphoric acid in D2O. The H NMR utilized
frequency presaturation for water suppression. Samples were ~30mM within 0.30 mL
volume, containing 10% D2O. The pH of all samples was adjusted to 6.8, using either
HCl or the corresponding base hydroxide, i.e. NaOH, KOH, or aq. TEA. Each sample
was heated to 75 °C for 10 minutes, then allowed to cool at room temperature prior to transferring to the NMR tube.
The DOSY spectra were acquired on a Varian Inova 500 MHz spectrometer at
25°C, using presaturation pulse for water suppression. The data were collected using a
2.5 second relaxation delay over 8000 Hz spectral width, containing 16 repetitions over
256 increments with a diffusion delay of 0.1 seconds.
Preparation of Cyclo–8-bromoguanosinylyl (3’Æ 5’)-8’-bromoguanosinylyl (3’Æ
5’), triethylammonium salt (c-di-Br-GMP, 43). To a stirred solution of 16 (0.19 g,
0.22 mmol, TEA+ form) in 2 mL water, was added a freshly prepared solution of 0.24 M
aqueous bromine (2.7 mL, 0.65 mmol, 3.0 equiv). After 1 hour, additional aqueous
bromine was added (0.5 mL, 0.11 mmol, 0.50 equiv). After 1 hour, 3 mL of cyclohexene
was added and stirred vigorously. The reaction mixture was diluted with 8 mL of water,
and then extracted with three 5 mL portions of methylene chloride. The aqueous layer
was concentrated on a speedvac to remove traces of organic solvent, and then
48
lyophilized. The product was purified by semi-preparative reverse-phase HPLC and then
desalted to give 0.19 g of 43 (0.182 mmol, 82%) in the triethylammonium form. UV λmax
-1 -1 1 (25°C) 263 nm; ε (25°C, pH 6.8) 32,800 OD M cm ; H NMR (H2O:D2O, 500 MHz): δ
6.46 (br s), 5.90 (d, J = 3.5 Hz, 2H), 5.21-5.15 (m, water suppression reduces intensity),
4.26-4.23 (m, water suppression reduces intensity), 4.10-4.07 (m, 2H), 3.11 (q, J = 7.5
13 Hz, 12H), 1.19 (t, J = 7.5 Hz, 18H); C NMR (H2O:D2O, 500 MHz): δ 160.1, 156.3,
155.2, 126.4, 119.3, 93.3, 82.8 (app t, JCP = 8.6 Hz), 74.1 (d, JCP = 5.0 Hz), 73.4, 65.1
31 (JCP = 5.0 Hz), 49.3, 10.9; P NMR (H2O:D2O, 500 MHz): δ -0.83 (s). The mass was
confirmed by ESI-MS in negative mode as m/z (M-1) 847.4 (calculated for
- C20H21Br2N10O14P2 : 847.2)
Preparation of Cyclo–8-thioguanosinylyl (3’Æ 5’)-8’-thioguanosinylyl (3’Æ 5’),
triethylammonium salt (c-di-thio-GMP, 56). To a flask containing 43 (0.180g, 0.17
mmol, TEA+ form), thiourea (0.052 g, 0.68 mmol, 4.0 equiv), and pyridinium trifluoroacetate (0.013 g, 0.68 mmol, 0.4 equiv) was added 2.5 mL of degassed water and
1.0 mL of degassed ethanol under a nitrogen atmosphere. A condenser was affixed and the flask was heated in a 60°C oil bath for 22 hours. The cooled reaction mixture was diluted with 3 mL water, and then lyophilized. The product was purified by semi- preparative reverse-phase HPLC, and then desalted to give 0.105 g of 56 (0.110 mmol,
65%) in the triethylammonium form. UV λmax (25°C) 302, 286 (shoulder) nm; ε (25°C,
-1 -1 1 pH 6.8) 38,200 OD M cm ; H NMR (H2O:D2O, 500 MHz): δ 6.49 (br s), 6.37 (d, J =
2.5 Hz, 2H), 5.24-5.15 (m, water suppression reduces intensity), 4.27-4.21 (m, water
suppression reduces intensity), 4.10-4.07 (m, 2H), 3.11 (q, J = 7.5 Hz, 12H), 1.20 (t, J =
49
13 7.5 Hz, 18H); C NMR (H2O:D2O, 500 MHz): δ 167.6, 156.6, 155.2, 153.0, 107.2, 92.4,
31 82.3 (app t, JCP = 9.5 Hz), 74.2 (d, JCP = 4.0 Hz), 65.1 (d, JCP = 4.0 Hz), 49.3, 10.9; P
NMR (H2O:D2O, 500 MHz): δ -0.78 (s). The mass was confirmed by ESI-MS in
- negative mode as m/z (M-1) 753.4 (calculated for C20H23N10O14P2S2 : 753.5)
Preparation of Cyclo-8-methylthioguanosinylyl (3’Æ 5’)-8’-methylthioguanosinylyl
(3’Æ 5’), triethylammonium salt (c-di-methylthio-GMP, 57). To a flask containing 56
(0.089g, 0.093 mmol, TEA+ form) was added 1.0 mL of dry DMF, which was then dried
via coevaporation with acetonitrile, and placed under a nitrogen atmosphere. A solution
of 0.7 M dimethylsulfate in dry DMF (0.60 mL, 0.42 mmol, 4.5 equiv) was added to 56
and allowed to react 4 hours. The reaction was quenched with 1 mL aqueous MeOH,
diluted with 5 mL of 0.1M TEAA buffer (pH 6.8), then extracted with three 5 mL
portions of ethyl ether. The aqueous layer was concentrated on a speedvac to remove
traces of organic solvent, and then lyophilized. The product was purified by semi-
preparative reverse-phase HPLC, and then desalted to give 0.030 g of 57 (0.029 mmol,
32%) in the triethylammonium form. UV λmax (25°C) 274 nm; ε (25°C, pH 6.8) 41,100
-1 -1 1 OD M cm ; H NMR (H2O:D2O, 500 MHz): δ 5.89 (d, J = 4.0 Hz, 2H), 5.13-5.10 (m, water suppression reduces intensity), 4.25-4.23 (m, water suppression reduces intensity),
4.10-4.08 (m, 2H), 3.11 (q, J = 7.5 Hz, 12H), 2.59 (s, 6H), 1.20 (t, J = 7.5 Hz, 18H); 13C
NMR (H2O:D2O, 500 MHz): δ 160.4, 156.1, 155.8, 150.6, 119.2, 91.9, 82.8 (app t, JCP =
31 9.0 Hz), 74.3 (d, JCP = 5.0 Hz), 73.7, 65.1 (d, JCP = 5.0 Hz), 49.4, 18.0, 10.9; P NMR
(H2O:D2O, 500 MHz): δ -0.73 (s).The mass was confirmed by ESI-MS in negative mode
- as m/z (M-1) 781.5 (calculated for C22H27N10O14P2S2 : 781.6)
50
Preparation of Cyclo–8-phenylguanosinylyl (3’Æ 5’)-8’-phenylguanosinyly (3’Æ 5’),
triethylammonium salt (c-di-Ph-GMP, 59). To a flask containing 43 (0.12 g, 0.19
mmol, TEA+ form), phenylboronic acid (0.046 g, 0.38 mmol, 3.2 equiv), palladium
acetate (0.002g, 0.0094 mmol, 0.08 equiv), tri(4,6-dimethyl-3-sulfonatophenyl)phosphine
(0.019 g, 0.028 mmol, 0.24 equiv), and sodium carbonate (0.038 g, 0.35 mmol, 3 equiv) was added 2 mL freshly degassed water under a nitrogen atmosphere. A condenser was affixed, and the flask was heated in a 95°C oil-bath for 20 hours. The cooled reaction
mixture was neutralized with 1 M HCl, and then extracted with three 5 mL portions of
ethyl acetate. The aqueous layer was concentrated on a speedvac to remove traces of
organic solvent and lyophilized. A Waters PoraPak 20cc reverse-phase pre-column was
used to separate the palladium catalyst. Further purification was done by semi-
preparative reverse-phase HPLC, and then desalted to give 0.052 g of 59 (0.050 mmol,
42%) in the triethylammonium form. UV λmax (25°C) 280 nm; ε (25°C, pH 6.8) 43,800
-1 -1 1 OD M cm ; H NMR (H2O:D2O, 500 MHz): δ 7.66-7.65 (m, 4H), 7.55-7.54 (m, 6H),
6.51 (br s), 5.70 (d, J = 4.0 Hz, 2H), 5.30-5.28 (m, water suppression reduces intensity)
5.17-5.12 (m, water suppression reduces intensity), 4.31-4.29 (m, water suppression reduces intensity), 4.16 – 4.11 (m, 2H), 3.07 (q, J = 7.5 Hz, 12H), 1.16 (t, J = 7.5 Hz,
13 18H); C NMR (H2O:D2O, 500 MHz): δ 161.0, 156.2, 155.0, 152.6, 133.3, 132.0, 131.6,
31 130.4, 118.6, 92.9, 82.3 (app t, JCP = 10.0 Hz), 73.7, 73.0, 65.1, 49.3, 10.9; P NMR
(H2O:D2O, 500 MHz): δ -0.77 (s). The mass was confirmed by ESI-MS in negative mode
- as m/z (M-1) 841.5 (calculated for C32H31N10O14P2 : 841.6)
51
Preparation of Cyclo–8-acetylphenylguanosinylyl (3’Æ 5’)-8’-
acetylphenylguanosinyly (3’Æ 5’), triethylammonium salt (c-di-acetylphenyl-GMP,
60). To a flask containing 43 (0.91 g, 0.087 mmol, TEA+ form), 3-acetylphenylboronic
acid (0.046 g, 0.28 mmol, 3.2 equiv), palladium acetate (0.002g, 0.007 mmol, 0.08
equiv), tri(4,6-dimethyl-3-sulfonatophenyl)phosphine (0.014 g, 0.021 mmol, 0.24 equiv),
and sodium carbonate (0.028 g, 0.26 mmol, 3.0 equiv) was added 2 mL freshly degassed
water under a nitrogen atmosphere. A condenser was affixed, and the flask was heated in
a 80°C oil-bath for 20 hours. The cooled reaction mixture was neutralized with 1 M HCl,
and then extracted with three 5 mL portions of ethyl acetate. The aqueous layer was
concentrated on the speedvac to remove traces of organic solvent and lyophilized. The
crude solid was dissolved in 15 mL of 50% aqueous MeOH, filtered through a 0.45 μm
filter, and then placed on the speedvac to remove the MeOH to crystallize the product.
The product was isolated as a white solid pellet, after decanting the supernatant, which
formed 0.012 g of 60 (0.011 mmol, 12%) in the triethylammonium form. UV λmax (25°C)
-1 -1 1 281 nm; ε (25°C, pH 6.8) 36,500 OD M cm ; H NMR (H2O:D2O, 500 MHz): δ 8.25
(s, 2H), 8.08 (d, J = 7.0 Hz, 2H), 7.96 (d, J = 8.0 Hz, 2H), 7.70 (app t, J = 7.5 Hz, 2H),
6.50 (br s), 5.70 (d, J = 4.0 Hz, 2H), 5.33-5.31 (m, water suppression reduces intensity)
5.21-5.18 (m, water suppression reduces intensity), 4.33-4.31 (m, water suppression reduces intensity), 4.17-4.12 (m, water suppression reduces intensity), 3.10 (q, J = 7.5
13 Hz, 12H), 2.70 (s, 6H), 1.19 (t, J = 7.5 Hz, 18H); C NMR (H2O:CD3OD, 500 MHz): δ
204.2, 161.4, 156.5, 155.8, 151.6, 140.0, 137.2, 132.8, 132.5, 132.4, 131.9, 119.3, 92.8,
31 83.8, 75.9, 73.4, 65.849.5, 29.0, 11.0; P NMR (H2O:D2O, 500 MHz): δ -0.90 (s). The
52
mass was confirmed by ESI-MS in negative mode as m/z (M-1) 925.7 (calculated for
- C36H35N10O16P2 : 925.7)
3.8 References
1. Gaffney, B.L., Veliath, E., Jones, R.A., "One-Flask Synthesis of c-di-GMP and the [Rp,Rp] and [Rp,Sp] Dithiophosphate Analogs". Organic Letters, 2010, 12(14), 3269-3271.
2. Shapiro, R., Agarwal, S., "Oxidation of Guanine and Guanosine by Bromine". Biochemical and Biophysical Research Communications, 1966, 24, 401-404.
3. Holmes, R., Robins, R., "Purine Nucleosides. VII. Direct Bromination of Adenosine, Deoxyadenosine, Guanosine, and Related Purine Nucleosides." Journal of the American Chemical Society, 1964, 86, 1242-1245.
4. Dias, E., Battiste, J.L., Williamson, J.R., "Chemical Probe for Glycosidic Conformation in Telomeric DNAs". Journal of the American Chemical Society, 1994, 116, 4479-4480.
5. Esposito, V., Randazzo, A., Piccialli, G., Petraccone, L., Giancola, C., Mayol, L., "Effects of an 8-Bromodeoxyguanosine Incorporation on the Parallel Quadruplex Structure [d(TGGGT)4]". Organic and Biomolecular Chemistry, 2004, 2, 313- 318.
6. Nadler, A., Diederichsen, U., "Guanosine Analog with Respect to Z-DNA Stabilization: Nucleotide with Combined C8-Bromo and C2'-Ethynyl Modifications". European Journal of Organic Chemistry, 2008, 1544-1549.
7. Cho, B.P., Kadlubar, F.F., Culp, S.J., Evans, F.E., "15N Nuclear Magnetic Resonance Studies on the Tautomerism of 8-Hydroxy-2'-deoxyguanosine, 8- Hydroxyguanosine, and Other C8-Substituted Guanine Nucleosides". Chemical Research in Toxicology, 1990, 3, 445-452.
8. Jankowski, A., Wise, D., Townsend, L., "Sodium Thiosulfate and Potassium Selenosulfate as Reagents to Prepare Thio- and Selenopurine Nucleosides". Nucleosides & Nucleotides, 1989, 8(3), 339-348.
9. Nurminen, E., Lonnberg, H., "Mechanisms of the Substitution Reactions of Phosphoramidites and Their Congeners". Journal of Physical Organic Chemistry, 2004, 17, 1-17.
53
10. Suzuki, A., "Recent Advances in the Cross-Coupling Reactions of Organoboron Derivatives with Organic Electrophiles, 1995-1998". Journal of Organometallic Chemistry, 1999, 576, 147-168.
11. Pierre Genet, J., Savignac, M., "Recent Developments of Palladium(0) Catalyzed Reactions in Aqueous Medium". Journal of Organometallic Chemistry, 1999, 576, 305-317.
12. Western, E., Daft, J., Johnson, E., Gannett, P., Shaughnessy, K., "Efficient One- Step Suzuki Arylation of Unprotected Halonucleosides, Using Water-Soluble Palladium Catalysts". Journal of Organic Chemistry, 2003, 68, 6767-6774.
13. Gubala, V., Betancourt, J., Rivera, J., "Expanding the Hoogsten Edge of 2'- Deoxyguanosine: Consequences for G-Quadruplex Formation". Organic Letters, 2004, 6, 4735-4738.
14. Collier, A., Wagner, G. , "A Facile Two-Step Synthesis of 8-Arylated Guanosine Mono- and Triphosphates (8-Aryl GXPs)". Organic and Biomolecular Chemistry, 2006, 4, 4526-4532.
15. Gubala, V., De Jesus, D., Rivera, J. M., "Self-Assembled Ionophores Based on 8- Phenyl-2'-deoxyguanosine Analogues". Tetrahedron Letters, 2006, 47, 1413- 1416.
16. Collier, A., Wagner, G., "A Fast Synthetic Route to GDP-sugars modified at the nucleobase". Chemical Communications, 2008, 178-180.
54
3.9 Appendix
55
c-di-Br-GMP (TEA+ form), 43
1H NMR
13C NMR
31P NMR
56
HPLC 2-20% acetonitrile with 0.1M triethylammonium acetate buffer (pH 6.8), 280 nm 1.20
1.00 0.80
0.60 AU 0.40
0.20 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 13.00 Minutes
LRMS
847.4 100
423.3 %
1271.7
0 m/z 200 300 400 5 00 600 700 800 900 1 000 1100 1200 130 0 1400 1500 1600 1700 1800 1900 2000
UV
57
DOSY of c-di-Br-GMP, K+ form
58
DOSY of c-di-Br-GMP, Na+ form
59
DOSY of c-di-Br-GMP, TEA+ form
60
c-di-thio-GMP (TEA+ form), 56
1H NMR
13C NMR
31P NMR
61
HPLC 2-20% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
1.50
1.00
AU 0.50
0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 Min ut es
LRMS
753.4 100 376.0
%
207.2 1005.1
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000
UV
62
DOSY of c-di-thio-GMP, K+ form
63
DOSY of c-di-thio-GMP, Na+ form
64
DOSY of c-di-thio-GMP, TEA+ form
65
c-di-methylthio-GMP (TEA+ form), 57
1H NMR
13C NMR
31P NMR
66
HPLC 2-20% acetonitrile in 0.1 M triethylammonium acetate (pH 6.8), 280 nm
0.40
AU 0.20
0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 Minutes
LRMS
100 781.5
%
207.2 299.9
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000
UV
67
DOSY of c-di-methylthio-GMP, K+ form
68
DOSY of c-di-methylthio-GMP, Na+ form
69
DOSY of c-di-methylthio-GMP, TEA+ form
70
c-di-Ph-GMP (TEA+ form), 59
1H NMR
13C NMR
31P NMR
71
HPLC 2-40% acetronitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
1.20
1.00 0.80 AU 0.60 0.40 0.20 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes
LRMS
841.5 100
420.3 %
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000
UV
72
DOSY c-di-phenyl-GMP, K+ form
73
DOSY of c-di-phenyl-GMP, Na+ form
74
DOSY of c-di-phenyl-GMP, TEA+ form
75
c-di-acetylphenyl-GMP (TEA+ form), 60
1H NMR
13C NMR
31P NMR
76
HPLC 2-40% acetonitrile in 0.1 M triethylammonium acetate (pH 6.8) 1.40 1.20
1.00
0.80
AU 0.60
0.40
0.20
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes
LRMS
92 5 .7 100
% 462.6
1389.1
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000
UV
77
DOSY of c-di-acetylphenyl-GMP, K+ form
78
4. Biophysical Studies of C8 Analogs of c-di-GMP
4.1 Introduction
NMR and UV spectroscopy are well suited to study G-quartets in solution.1, 2 1H
NMR provides insight to the local phenomena of the guanine base protons participating in H-bonding within quartets, as evidenced by downfield resonances from 9-12 ppm. UV spectroscopy can assess the more global phenomena of base stacking at more dilute concentrations (< 0.5 mM vs. ~30 mM for NMR) by tracking absorbance with thermal denaturation. Taken together, these techniques can provide valuable insight to the solution-phase structures of species containing G-quadruplexes.
4.2 Previous Work on the Biophysical Studies of c-di-GMP
In two previous publications from Zhaoying Zhang et al. from the Jones lab involving the biophysical studies of different salt forms of c-di-GMP 3, 4, it was shown
that the Li+, Na+ and K+ salt forms of c-di-GMP formed intermolecular G-quadruplex
complexes in solution to differing degrees. All showed a concentration-dependent
formation of these higher-order structures as shown by 1H and 31P NMR. On heating the
NMR samples to 55°C, the Li+ and Na+ forms showed disruption of the quartet structures
toward lower-ordered bimolecular and tetramolecular structures; whereas the K+ form exhibited much more stable higher-order structures such as tetramolecular and octamolecular species (Figure 2-6). From the diffusion-ordered spectroscopy (DOSY)
2D NMR, it was shown that the K+ form formed two populations of higher-order
complexes: octamolecular with smaller amounts of tetramolecular species. There existed
also a combination of syn and anti conformers (Figure 2-7) within the quartet complexes
79
which was established by the presence of NOE crosspeaks for the syn form between the
H1’ and the H8 and the absence of a crosspeak for the anti form. Overall, there existed an equilibrium between the bimolecular, tetramolecular and octamolecular species at NMR concentrations (Figure 2-7). The lower-order forms were favored by more dilute concentrations (0.05-0.5 mM) used in UV melting studies (Figure 2-9), the Li+ counter- ion salt form and higher temperatures that disrupted higher-ordered forms. The Na+ form exhibited a singular transition in the UV melt, with a Tm of 8 °C (Figure 2-9, a). The
higher-order forms were favored by higher concentrations (~35 mM) used in NMR, the
K+ counter-ion salt form and lower temperatures (Figure 2-10). The K+ form exhibited
two transitions with the UV melt, with Tm of 30 °C and 45 °C respectively, indicating an
equilibrium among three species (Figure 2-9, b).
Figure 2-7: Equilibrium of different ordered structures dependant on the counter-cation identity and concentration of unmodified c-di-GMP.4
80
Figure 2-8: anti and syn conformers of c-di-GMP showing the 3- bonds contructing the torsion angle about the glycosidic bond. Torsion angles for the glycosidic bond are determined from the angle (χ) between C4- N9-C1’-O4’: anti- 150° ≥ χ ≥ 210° and syn- -30° ≥ χ ≥ 30°.5
a)
b)
Figure 2-9: UV melting profiles of c-di-GMP a) Na+ form concentrations were 0.0367, 0.0646, 0.162, and 0.338 mM respectively in 10 mM sodium phosphate buffer, 0.1 M NaCl pH 7; b) K+ form concentrations were 0.0242, 0.0462, 0.118, and 0.241 mM respectively in 10 mM potassium phosphate buffer, 0.1 M KCl pH 7.3
81
Figure 2-10: 1H NMR of the different salt forms of c-di-GMP at different temperatures illustrating the relative stability of the quartet structures, indicated by the downfield resonances from 8.5- 12 ppm.3
4.3 Biophysical Studies of c-di-Br-GMP
4.3.1 NMR studies of c-di-Br-GMP
4.3.1.1 Introduction
1H and 31P NMR studies of c-di-Br-GMP consisting of the K+, Na+ and the TEA+ form (~30 mM ) with 0.1 M of the corresponding chloride salt were prepared in H2O:D2O
(9:1). Additional salt ensured an excess of cation was present to participate in any
potential quartet in a host-guest manner, and is within the concentration range found in
bacterial cells.6 Variable temperature NMR was used to probe the substituent effect on
complexation, and 2D Diffusion Ordered Spectroscopy (DOSY) NMR was used to
determine the diffusion coefficients of any potential complexes. In addition, the
82
remaining analog NMR samples were prepared in a similar manner for the associated
biophysical studies.
4.3.1.2 K+ form of c-di-Br-GMP
The K+ form of c-di-Br-GMP showed clear quartet formation at 25 °C (Figure
2-11, b). The 1H NMR shows that the region between 5.5-6.5 ppm contains multiple H1’
resonances. The 2D DOSY NMR clearly indicates the presence of predominantly higher
order octamolecular species from the three main peaks (6.05, 6.09, 6.25 ppm respectively). From the remaining peaks, some tetramolecular (6.14 and 6.24 ppm) is observed, and to a lesser extent, some bimolecular (5.97 ppm) is also observed. The diffusion coefficents (D) to these corresponding peaks from the DOSY NMR are 1.5, 1.7, and 3.5 m2/s, respectively. The non-hydrogen-bonded amino peak region (7-8 ppm) and
the hydrogen-bonded amino peak region (8-9 ppm) are dominated by three main sets of
peaks that also have a D = 1.5 m2/s, which are the resonances that correspond to the octamolecular complexes. The most downfield region (10.5-11.5 ppm) displays the amido protons, which contain two major peaks (10.60, 10.82 ppm) with a D = 1.5 m2/s, and two minor peaks (10.85, 11.03 ppm) with a D = 1.7 m2/s. At 25 °C, this data shows
the existence of an equilibrium among three species, which is predominantly
octamolecular complexes with a small amount of tetramolecular species and a minor
portion of bimolecular species.
The variable temperature conditions illustrate the shifting of the complex
equilibrium to higher-order complexes at lower temperatures (5 °C, Figure 2-11, a) and
toward more mid- to lower-order complexes at elevated temperatures (55 °C, Figure 2-
11, b). This process has been shown to be reversible.
83
1H NMR of c-di-Br-GMP (K+ form) a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-11: 1H NMR of c-di-Br-GMP (K+ form, 31 mM), showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C . The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M KCl, H2O:D2O (9:1), pH 6.8.
84
The 31P NMR spectra of c-di-Br-GMP show similar phenomena with a
simplified spectra (Figure 2-12, a-c). The 25 °C spectrum is dominated by three main
peak resonances (-3.02, -0.84, and 0.11 ppm), which increase in proportion at lower
temperatures, and that decrease on heating. This is analogous to the set of three
resonances of octamolecular complexes found in the 1H NMR spectra. The other minor
peaks match to the lower proportions of tetramolecular (-1.44 and -0.50 ppm) and bimolecular (-0.68 ppm). At elevated temperatures, the bimolecular resonances increase
in intensity and the tetramolecular resonances decrease, which is in agreement with the disruption of the H-bonding network with increased thermal energy inputted into the
system. The pattern of the three sets of resonances for the octamolecular, two sets of
resonances for the tetramolecular and one set for the bimolecular are repeated in the 31P spectra as was in the 1H spectra for the c-di-Br-GMP in the K+ form.
85
31P NMR of c-di-Br-GMP (K+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-12: 31P NMR of c-di-Br-GMP (K+ form, 31 mM) with annotated molecularity at a) 5 °C, b) 25 °C, and c) 55 °C. Samples were 0.1 M KCl, H2O:D2O (9:1), pH 6.8.
86
4.3.1.3 Na+ form of c-di-Br-GMP
The Na+ form of c-di-Br-GMP under similar sample conditions as the K+ form, illustrates a lesser tendency to form G-quadruplexes in solution. It has been proposed that the Na+ cation (ionic radii of 1.02 Å)7 is less apt to stabilize quartet formation due to the
relatively high free energy cost of dehydration, compared to K+ (ionic radii of 1.38 Å),
that is required for inner sphere contacts with the guanine ligands. This results in a tighter
sphere of hydration that hinders inner sphere coordination by the Lewis basic sites on the
guanine O6.8 Overall, the sodium form of c-di-Br-GMP can form a small proportion of
G-quadruplexes when the sample is cooled to 5 °C, which illustrates the level of
instability of the complex with this particular salt form.
The 1H NMR at 25 °C of the region between 5-12 ppm shows only a trace
amount of tetramolecular species present (Figure 2-13, b). The most predominant peak in
the H1’ region is a singlet peak (5.76 ppm, J = 3.0 Hz) that corresponds to the bimolecular
form with a diffusion coefficient of D = 3.2 m2/s. The next smaller downfield peaks (6.15
ppm and 6.24 ppm) correspond to the tetramolecular form, having a D = 1.7 m2/s. The
second most prominent peak is the broad amino resonance (6.52 ppm) that has a
corresponding D = 3.2 m2/s, which indicates it is associated with the bimolecular species.
The absence of any further downfield resonances indicates no quartet formation under
these conditions for the sodium form of c-di-Br-GMP at this temperature.
Variable temperature 1H NMR on the sodium form of c-di-Br-GMP illustrates the instability of G-quadruplex complexation under these conditions, which is in stark
contrast to its facile formation as the K+ form. Upon cooling the sample to 5 °C, solvent-
protected proton resonances appear in the quartet region, albeit in a small percentage to
87
the predominantly bimolecular form (Figure 2-13, a). Within the H1’ region, the
tetramolecular peaks increase in intensity, as well as the other downfield markers of
quartet formation. The non-hydrogen bonded amino peaks (~7 ppm) appear along with
the hydrogen-bonded peaks (~8.5-9 ppm). The H1 amido peaks also appear at 5 °C (10.5-
11 ppm). Additionally, the broad singlet of the amino (6.36 ppm) has increased in
intensity. Upon heating to 55 °C, any minor quartet complexation is disrupted
completely, and the broad amino peak (6.75 ppm) has virtually disappeared which
illustrates the special protection from solvent exchange that the dibrominated cyclic
dimer offers at lower temperatures.
The 31P NMR spectra for the Na+ form of c-di-Br-GMP confirm the existence
of a predominantly bimolecular form under the observed temperatures (Figure 2-14, a-c).
At 25 °C, the main resonance (-0.73 ppm) is assigned as being the bimolecular form by
analogy to the 1H NMR spectrum at the same temperature (Figure 2-14, b). Upon cooling to 5 °C, smaller tetramolecular resonances (-1.41 and -0.61 ppm) appear adjacent to the main bimolecular peak (-0.96 ppm) (Figure 2-14, a). Upon heating, the spectrum appears almost identical to the 25 °C spectrum, with only one bimolecular resonance (-
0.50 ppm) (Figure 2-14, c).
88
1H NMR of c-di-Br-GMP (Na+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-13: 1H NMR of c-di-Br-GMP (Na+ form, 31 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C. The a) 5 °C) and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M NaCl, H2O:D2O (9:1), pH 6.8.
89
31P NMR of c-di-Br-GMP (Na+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-14: 31P NMR of c-di-Br-GMP (Na+ form, 31 mM) with annotated molecularity at a) 5 °C, b) 25 °C, and c) 55 °C. Samples were 0.1 M NaCl, H2O:D2O (9:1), pH 6.8.
90
4.3.1.4 Li+ form of c-di-Br-GMP
The Li+ form of c-di-Br-GMP does not support any higher order quadruplex
complexation at all, even at 5 °C. The only phenomenon observed is the increasing
intensity of the amino proton broad singlet (6.37 ppm) on sample cooling to 5 °C, similar
to the Na+ form. Both the 1H and 31P NMR illustrate the absence of higher order complexes at 5 °C, since the spectra lack any characteristic downfield resonances for G- quartet structures (Figure 2-15, a-b).
a)
b)
+ 1 Figure 2-15: 5° C NMR of c-di-Br-GMP (Li form, 31 mM), 0.1 M LiCl, H2O:D2O (9:1) pH 6.8. a) H NMR , b) 31P NMR
91
4.3.1.5 TEA+ form of c-di-Br-GMP
The TEA+ form of c-di-Br-GMP does not support any quadruplex complexation
of any kind, even at the 5 °C temperature. The triethylammonium cation is sterically
bulky and cannot interact with the guanine O6 in an ion-dipole electrostatic interaction in a manner similar to a metal cation. In the variable temperature NMR study, the 1H and
31P NMR indicate a strictly bimolecular arrangement of the molecule that is in fast
exchange with the monomeric unit (Figure 2-16, a-c). The amino group of the compound
is strikingly visible at both 5 °C (6.60 ppm) and 25 °C (6.50 ppm), similar to the Na+ form at 25 °C which increases in intensity at lower temperature. The structural characteristics of this symmetric dimer allows for increased protection of the amino protons from solvent exchange.
92
1H NMR of c-di-Br-GMP (TEA+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-16: 1H NMR of c-di-Br-GMP (TEA+ form, 31 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C . The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M TEACl, H2O:D2O (9:1), pH 6.8.
93
4.3.1.6 Salt Forms of 8-Br-GMP Monomer
In an effort to understand the observed amino proton resonances of the Na+ and
TEA+ form of c-di-Br-GMP and to asses the boundary of quartet formation, the monomer
8-Br-GMP was prepared for an NMR study in a similar manner to c-di-Br-GMP as the
+ + + K , Na and TEA salts with 0.1 M chloride salt in H2O:D2O (9:1) at pH 6.8.
In all cases, quartet formation is not supported across the different temperatures
with the different counter-cations (Figure 2-17 for K+, Figure 2-18 for TEA+, Na+ not shown). The diffusion coefficients (D) for the K+, Na+ and TEA+ forms are 4.3, 4.0, and
3.7 m2/s respectively. Additionally, the amino peak is not visible, with the exception of a
trace amount at 5 °C (~6.60 ppm, Figures 2-17a, 2-18a). This illustrates the special
structural characteristics of the cyclic dimer whereby the amino protons are protected
against solvent exchange that is not seen with the monomeric unit.
94
1H NMR of 8-Br-GMP (K+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-17: 1H NMR of 8-Br-GMP (K+ form, 30 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C . The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M KCl, H2O:D2O (9:1), pH 6.8. There is no evidence of quartet formation for this compound as indicated by the absence of downfield resonance peaks.
95
1H NMR of 8-Br-GMP (TEA+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-18: 1H NMR of 8-Br-GMP (TEA+ form, 30 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C . The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M TEACl, H2O:D2O (9:1), pH 6.8. There is no evidence of quartet formation for this compound as indicated by the absence of downfield resonance peaks.
96
4.3.1.7 Conclusions for NMR Studies of c-di-Br-GMP
Variable temperature 1H and 31P 1D NMR and DOSY 2D NMR was used to
assess the solution-phase structures of c-di-Br-GMP as the K+, Na+ and the TEA+ salt form. Only the K+ form supports G-quadruplex formation across all observed
temperatures. The Na+ form only exhibits a minor proportion of quadruplex formation at
lower temperatures, but also exhibits a special structural conformation that protects the
amino protons from solvent exchange allowing them to be visible under NMR conditions.
The TEA+ form does not support quadruplex formation, but does show the amino proton resonances similar to the Na+ form. Additionally, the NMR visible amino protons are
specific to the cyclic dimer since the 8-Br-GMP monomer does not show this resonance
under similar NMR sample conditions.
4.3.2 UV Studies of c-di-Br-GMP
4.3.2.1 UV Melting study for K+ and Na+ Forms of c-di-Br-GMP
UV absorption of a conjugated chromophore, such as guanine, is sensitive to the
the aromatic stacking effects often seen in solution. Chromophores in close proximity, as
in π-stacking, exhibit a decrease in in absorption of UV light. Upon application of heat to
the system, the stacking effects are disrupted and there is an observable increase in absorpance, that can be correlated to the Tm of the complex. If there are higher order
structures contained within the sample, hypothetically two or more transitions can be
97
observed. This phenmonenon was observed with the parent molecule, c-di-GMP, as
previously described in the introduction of this section.
Both the K+ and the Na+ form of c-di-Br-GMP were prepared in a solution containing 0.1 M of the chloride salt in 10 mM K+ or Na+ citrate-phosphate buffer (pH
6.8). A series of dilutions were prepared in differing path length quartz cells to assess any
possible concentration-dependence of complexation equilibrium.
4.3.2.2 Results
Both the K+ and the Na+ form of c-di-Br-GMP containing excess chloride salt in
the appropriate buffer did not exhibit any hypochromic indication of self-stacking at UV
concentrations. The concentration series that underwent the UV melting from 4 °C to 80
°C was a slightly hyperchromic curve, indicating a minor dilution effect on the expansion
of the solvent volume upon heating within the confined cell (Figure 2-19, a-b). The
absence of an increase in absorption, or hypochromicity with an increase in temperature,
would indicate no inter- or intramolecular self stacking of the aromatic bases has occurred. The lack of this phenmonemon reveals the stacking of c-di-Br-GMP is much
less stable compared to the parent unmodified compound at UV concentrations.
98
a) b)
Figure 2-19: Normalized data for UV melting profile for a) c-di-Br-GMP (K+ form), b) c-di-Br-GMP (Na+ form).
4.3.3 Further Investigation of NMR-visible Amino Resonance
The appearance of the broadened singlet from 6.0-6.5 ppm prompted a more in-
depth investigation to the protons responsible on the guanine bases. The fact that the peak
is only visible with the C8-brominated or otherwise modified cyclic dimer and not the
brominated monomer, is a statement to the special structural characteristics of the
molecule.
99
4.3.3.1 Heteronuclear 2D NMR of K+ and Na+ forms of c-di-Br-GMP
A 1H-13C Heteronuclear Multiple Bond Correlation (HMBC)/ Heteronuclear
Multiple Quantum Correltaion (HMQC) NMR of both the K+ and Na+ forms of c-di-Br-
GMP was performed to assess the longer two and three-bond couplings using the HMBC method, and to detect any direct, single bond couplings using the HMQC method. The combined spectra for both salt forms of c-di-Br-GMP is shown in Figure 2-20.
Overall, the data indicate a syn-conformation about the glycosidic bond between the C1’ and N9. When the conformation is syn, a stronger cross peak is seen between the
H1’-C4; if the conformation is anti, then a stronger crosspeak is seen between the H1’-
C8. In both salt forms of c-di-Br-GMP, a stronger crosspeak is seen between H1’ and C4, confirming the syn conformer (Figure 2-20, a-b). Most significantly, the lack of any crosspeak between the broadened singlet downfield at ~6.5 ppm from the typical H1’ region to any carbon confirms this proton is not directly attached to a carbon on the guanosine base.
100
a)
b)
Figure 2-20: 1H-13C HMBC/HMQC NMR of a) K+ and b) Na+ forms of c-di-Br-GMP. Samples were 31 mM of c-di-Br-GMP, 0.1 M in chloride salt, H2O:D2O (9:1). In both samples, the stronger crosspeak between the H1’-C4 indicate a syn conformation about the glycosidic bond.
101
4.3.3.2 Effect of pH on UV profile of Na+ Form of c-di-Br-GMP
To rule out the possibility that the prominent broad singlet peak is due to a shifted
pKa of the N7 due to chemical modification at the adjacent C8, a series of wavelength
scans at different pH points was performed to track any discernible changes in the
wavelength scan. If the N7 is being protonated, the electronic transitions of the
chromophore should change drastically due to the full positive charge residing on the
imidazole portion of the guanine moiety. This would shift the overall wavelength scan to
a different maxima and switches the position of a shoulder for the N7-protonated
guanosine.9
A series of UV samples from pH 3.0 to 7.0 in 0.25 increment steps were prepared
and had wavelength scans performed at 25 °C. The pKa of the unmodified guanosine N7
is typically 2.4. Since this low pH would likely degrade c-di-Br-GMP, the chosen pH range for study was 3.0 to 7.0, which should give some indication of a pKa shift.
Overall, there was no significant change in the shape of the UV profile or the
positions of the UVmax or the shoulder, as seen in Figure 2-21, which shows the pH
endpoints of the study. Therefore, the broadened singlet is clearly not from a protonated
guanosine N7.
In conclusion, the results from 2D NMR and UV studies support the hypothesis that
the broad singlet at ~6.5 ppm of the c-di-GMP analogs is the amino peak shielded from
solvent exchange.
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Figure 2-21: UV wavelength scan of c-di-Br-GMP (Na+ form, 27 mM) in 10 mM sodium citrate-phosphate buffer at pH 3.0 and 7.0. The lack of a significant change in the UV profile indicates the chromophore is not being protonated to a discernable extent.
4.4 Biophysical Studies of c-di-thio-GMP: K+ , Na+ and TEA+ forms
The 1H and 31P NMR of the K+ form illustrate that c-di-thio-GMP does not
undergo any quadruplex complexation of any sort at the observed NMR concentration,
even at lower temperatures. The only noticeable phenomenon in the 1H NMR is the
appearance of amino resonance at 25 °C as a broad singlet (6.48 ppm). This peak increases in intensity at 5 °C and completely disappears at 55 °C (Figure 2-22, a-c). The
DOSY NMR shows the diffusion coefficient of this bimolecular species of 2.9 m2/s. The
31P NMR of the three temperature points also show only a sharp singlet (data not shown),
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only varying slightly in chemical shift due to the temperature change, confirming the
existence of only one species in solution.
Both the Na+ and TEA+ forms of c-di-thio-GMP resemble the K+ form in that
both salt forms lack any appearance of quadruplex complexation. The 1H NMR spectra
for these salt forms resemble those of the K+ form in Figure 2-22. Again, the amino peaks are visible at 25 °C and to a larger extent at 5 °C , and which completely disappears at 55 °C in a similar manner to the data for K+. The DOSY NMR indicates the diffusion
coefficients for the Na+ and TEA+ forms are 2.5 and 3.0 m2/s respectively. The 31P NMR also supports the data from the 1H NMR showing only one species present in solution
(data not shown).
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1H NMR of c-di-thio-GMP (K+ form)
Figure 2-22: 1H NMR of c-di-thio-GMP (K+ form, 27 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C. The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of + + the resonances. Samples were 0.1 M KCl, H2O:D2O (9:1), pH 6.8. The spectra for the Na and the TEA forms are similar to above.
105
4.5 Biophysical Studies of c-di-methylthio-GMP: K+ , Na+ and TEA+ forms
The 1H and 31P NMR of the K+ form of c-di-methylthio-GMP indicates that this analog does not undergo complexation to quadruplex structures, in a similar manner to the thio analog. The 1H NMR at the 5 °C, 25 °C and 55 °C temperature points show the lack of any downfield quartet resonances which supports a strictly a bimolecular form
(Figure 2-23, a-c). The amino resonance is observed at 25 °C (6.36 ppm) and more prominently at 5 °C (6.48 ppm) in a similar way to the other analogs. The diffusion coefficient for the bimolecular form from DOSY NMR is 3.6 m2/s. The 31P NMR show only a sharp singlet with each of the three temperature points illustrating the singular species present (data not shown).
The 1H and 31P NMR for the Na+ and TEA+ forms of c-di-methylthio-GMP are virtually identical to the K+ form in terms of the amino peak and the absence of any higher order complexes. The diffusion coefficients for the Na+ and TEA+ forms are 3.7 and 3.0 m2/s respectively, which differ from the K+ form slightly, showing the different hydrodynamic radius of the compound in solution. Thus, c-di-methylthio-GMP does not support quadruplex formation, similar to c-di-thio-GMP.
106
1H NMR of c-di-methylthio-GMP (K+ form)
a) 5 °C
b) 25 °C
c) 55 °C
Figure 2-23: 1H NMR of c-di-methylthio-GMP (K+ form, 26 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C. The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 26 mM c-di-methylthio-GMP, 0.1 M KCl, H2O:D2O (9:1), pH 6.8. The spectra for the Na+ and the TEA+ forms are similar to above.
107
4.6 Biophysical studies of c-di-phenyl-GMP: K+, Na+ and TEA+ forms
The different salt forms of c-di-phenyl-GMP exhibit similar biophysical
phenomenon as compared to the thio- and the methylthio- analogs, in that there is no
quadruplex formation under the observed NMR sample conditions. The amino peak
follows the similar trend whereby the amino protons are protected from solvent exchange
particularly at lower temperatures; however the K+ and the TEA+ forms show this peak at
55 °C (Figure 2-24) whereas the Na+ form does not at this elevated temperature (Figure
2-25). The K+ spectra is virtually identical to the TEA+ spectra, therefore only the K+ is shown in Figure 2-24. The additional phenyl ring substituents provide an extended conjugated aromatic system that can provide additional protection, and potentially offer an amino-aromatic ring hydrogen bonding interaction.10 It is not clear why the amino peak with the Na+ sample is lower in intensity as compared to the K+ and TEA+ forms at the various temperatures.
DOSY NMR provided the diffusion coefficients for the three salt forms. The
K+, Na+ and the TEA+ form have a bimolecular value of D = 2.6, 2.9, and 2.5 m2/s
respectively.
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1H NMR of c-di-phenyl-GMP (K+ form)
Figure 2-24: 1H NMR of c-di-phenyl-GMP (K+ form, 28 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C . The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of + + the resonances. The TEA form is very similar to the above K spectra. Samples were 0.1 M KCl, H2O:D2O (9:1), pH 6.8.
109
1H NMR of c-di-phenyl-GMP (Na+ form)
Figure 2-25: 1H NMR of c-di-phenyl-GMP (Na+ form, 28 mM) showing the diffusion coefficients (D, m2/s) and molecularity at b) 25 °C. The a) 5 °C and c) 55 °C temperatures are annotated with the molecularity of the resonances. Samples were 0.1 M NaCl, H2O:D2O (9:1), pH 6.8.
110
4.7 Biophysical studies of c-di-acetylphenyl-GMP: K+ and Na+ forms
As mentioned earlier in Section 3.5, the c-di-acetylphenyl-GMP analog was
selected as a target for biophysical studies after it was discovered that c-di-Br-GMP was
the only compound of the C8-modified analogs that could undergo quadruplex formation.
The additional H-bond acceptor on the carbonyl of the meta-acetyl group of the phenyl substituent could provide an H-bonding site for the exocyclic amino group in a quartet arrangement.
The solubility of this analog posed a challenge in that is it only freely soluble in
50% aqueous MeOH. The NMR samples had to be prepared using 0.05 M chloride salt in MeOD: H2O to avoid the sample precipitating completely in the NMR tube. In
addition, the changed solvent composition prompted reexamination of the c-di-Br-GMP
(K+ form) to assess the effect of the aqueous MeOH on quadruplex formation.
The results show that the K+ form of c-di-acetylphenyl-GMP does not support
quadruplex formation at all under NMR sample conditions. The only species present is the bimolecular form, showing only those resonance peaks associated with the
uncomplexed compound (Figure 2-26, a). The DOSY NMR shows the diffusion
coefficient as being 2.0-2.1 m2/s at 25 °C. The Na+ form does not form any higher order
complexes as well (Figure 2-26, b). For comparison, the c-di-Br-GMP (K+ form) in 50%
aqueous MeOD does form quadruplexes, as shown by the characteristic downfield
resonances from 8-12 ppm (Figure 2-26, c). Therefore, the binary solvent system should not adversely affect the potential for complexation. The lack of G-quadruplex formation is inherent to the c-di-acetylphenyl-GMP structure and not the aqueous MeOH solution.
111
1H NMR of c-di-acetylphenyl-GMP (K+ and Na+ form) and c-di-Br-GMP (K+ form)
a)
b)
c)
1 + Figure 2-26: a) H NMR of c-di-acetylphenyl-GMP (K form, 26 mM) at 25 °C, 0.05 M KCl, MeOD:H2O (1:1) with annotated diffusion coefficients (m2/s), b) 1H NMR of c-di-acetylphenyl-GMP (Na+ form, 26 1 + mM) a 25 °C, 0.05 M NaCl, MeOD:H2O (1:1), c) H NMR of c-di-Br-GMP (K form, 30 mM) at 25 °C, 0.05 M KCl, MeOD:H2O (1:1).
The inability of the c-di-acetylphenyl to form quadruplexes could stem from the adverse enthalpic penalty of bulky groups coming in close proximity, as well as the negative entropic effect of fixing many degrees of freedom for a large tetramolecular complex. The original reference11, 12 studied the use of this added structural element to
112
extend the Hoogsteen base pairing edge on lipophilic guanosine derivatives in organic
solvents like chloroform and acetonitrile. However, this system is an aqueous-organic
solvent system, that has water and MeOH competing as H-bonding donors and acceptors
against the cyclic dimer itself.
4.8 NMR Re-examination of c-IMP-GMP: K+ Form
Previously, Zhaoying Zhang from the Jones lab published a method to synthesize
c-IMP-GMP (K+ form) and study its solution-phase structure via NMR.4 It was found
from this research the absence of one exocyclic amino group due to the incorporation of
inosine into the cyclic ribonucleotide excluded quadruplex formation. Therefore, this
asymmetric cyclic dimer exists solely as the bimolecular form in solution at NMR
concentrations.
In light of the fact that the majority of the C8 analogs of c-di-GMP exist also as
the uncomplexed, bimolecular form, the 1H NMR of c-IMP-GMP (K+ form) was re-
examined. It exhibited a broadened singlet (5.92 ppm) at 25 °C that appeared slightly downfield to the H1’ resonance region (Figure 2-27, b). This peak was more prominent
at 5 °C (Figure 2-27, a), having an integration almost twice that at 25 °C (Figure 2-27, b). When this sample was lyophilized and reconstituted in all D2O, this peak disappeared
confirming it is an exchangeable proton on the nucleobase moiety (Figure 2-27, c). In
conclusion, c-IMP-GMP also possesses an NMR-visible guanine amino peak that is
characteristic of a cyclic ribonucleotide dimer. This phenomenon has been shown to be
independent of C8 modification on the purine base as well.
113
1H NMR of c-IMP-GMP (K+ form)
a)
b)
c)
1 + Figure 2-27: H NMR of c-IMP-GMP (K form, 61 mM), 0.1 M KCl, H2O:D2O (9:1) at a) 5 °C, b) 25 °C
c) 25 °C in D2O.
114
4.9 Conclusions for Biophysical Studies of C8 analogs of c-di-GMP
All five analogs were studied using NMR for evidence of quadruplex formation as the K+, Na+ and the TEA+ forms with excess chloride salt. Of the five, only c-di-Br-GMP was the only one that spontaneously formed higher order octamolecular and tetramolecular species as the K+ form, and bimolecular and tetramolecular as the Na+ form. A summary of the diffusion coefficients D for all compounds at 25 °C are shown in
Table 2-1.
All compounds that did not form higher order structures did exhibit a prominent broadened amino singlet peak that was more intense at lower temperatures.
115
Diffusion coefficients (m2/s) of c-di-GMP C8 analogs Bi- (m2/s) Tetra- (m2/s) Octa- (m2/s) 8-Br-GMP
TEA+ 3.6-3.8 nd nd Na+ 3.9-4.1 nd nd K+ 4.2-4.4 nd nd monomer c-di-Br- GMP TEA+ 2.8-3.0 nd nd Na+ 3.1-3.3 1.6-1.7 nd K+ 3.4-3.5 1.6-1.7 1.4-1.5
c-di-thio- GMP TEA+ 2.9-3.0 nd nd Na+ 2.4-2.6 nd nd K+ 3.0-3.2 nd nd
c-di-S-Me- GMP TEA+ 2.9-3.1 nd nd Na+ 3.7-3.8 nd nd K+ 3.4-3.6 nd nd
c-di-Ph- GMP TEA+ 2.3-2.5 nd nd Na+ 2.8-2.9 nd nd K+ 2.5-2.6 nd nd
c-di-AcPh- GMP*
K+ 1.9-2.1 nd nd
Table 2-1: Diffusion coefficients (m2/s) of c-di-GMP C8 analogs as a function of substituent and counter- ion at 25°C. *All samples are in H2O:D2O (9:1), except for c-di-AcPh-GMP which is in MeOD:H2O (1:1). nd = not detected.
116
4.10 References
1. Hardin, C., Perry, A., White, K., "Thermodynamic and Kinetic Characterization of the Dissociation and Assembly of Quadruplex Nucleic Acids". Biopolymers (Nucleic Acid Sciences), 2001, 56, 147-194.
2. Keniry, M., "Quadruplex Structures in Nucleic Acids". Biopolymers (Nucleic Acid Sciences), 2001, 56, 123-146.
3. Zhang, Z., Gaffney, B.L., Jones, R.A., "c-di-GMP Displays A Monovalent Metal- Ion Dependent Polymorphism". Journal of the American Chemical Society, 2004, 126, 16700-16701.
4. Zhang, Z., Kim, S., Gaffney, B.L., Jones, R.A., "Polymorphism of the Signaling Molecule c-di-GMP". Journal of the American Chemical Society, 2006, 128(21), 7015-7024.
5. Saenger, W., Structures and Conformational Properties of Bases, Furanose Sugars, and Phosphate Groups, in Principles of Nucleic Acid Structure, Cantor, C.R., Editor. 1984, Springer-Verlag: New York. p. 51-101.
6. Nakamura, T., Tokuda, H., Unemoto, T., "Effects of pH and Monovalent Cations on the Potassium Ion Exit from the Marine Bacterium, Vibrio Alginolyticus, and the Manipulation of Cellular Cation Contents". Biochimica et Biophysica Acta, 1982, 692, 389-396.
7. Feig, A.L., Uhlenback, O.C., The Role of Metal Ions in RNA Biochemistry, in The RNA World, Gesteland, C., Atkins, Editor. 1999, Cold Spring Harbor Laboratory Press: New York.
8. Hud, N.V., Smith, F.W., Anet, F.A.L, Feigon, J., "The Selectivity for K+ versus Na+ in DNA Quadruplexes is Dominated by Relative Free Energies of Hydration: A Thermodynamic Analysis by 1H NMR." Biochemistry, 1996, 35, 15383-15390.
9. Pandey, K.S., Mishra, P.C., "Electronic Spectra of 8-Bromoguanosine and 8- Bromoadenosine". Journal of Photochemistry and Photobiology, 1991, 62(1), 107-115.
10. Levitt, M., Perutz, M.F., "Aromatic Rings Act as Hydrogen Bond Acceptors". Journal of Molecular Biology, 1988, 201, 751-754.
11. Gubala, V., Betancourt, J., Rivera, J., "Expanding the Hoogsten Edge of 2'- Deoxyguanosine: Consequences for G-Quadruplex Formation". Organic Letters, 2004, 6, 4735-4738.
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12. Gubala, V., De Jesus, D., Rivera, J. M., "Self-Assembled Ionophores Based on 8- Phenyl-2'-deoxyguanosine Analogues". Tetrahedron Letters, 2006, 47, 1413- 1416.
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Chapter 3
New Synthetic Method for 5’-Capped Oligoribonucleotides
1. Capped RNA Biological Background
1.1 Introduction
RNA can have major modifications made to it after transcription, such as 1) the
addition of a polyadenosine tail (poly (A) tail) on the 3’ end, 2) the splicing out of intron
sequences within the nucleus, and 3) the addition of a 7-methylguanosine triphosphate
cap on the 5’ end of the nascent transcript.1, 2 This last modification, known as RNA
capping, occurs in the cell nucleus on any RNA polymerase II transcript3, and is enzymatically attached during transcription when the transcript strand is 25-30 nucleotides in length.4
1.2 Biological Function of Capped RNA
The presence of the 5’ cap on RNA transcripts is critical for a variety of important
cellular functions to occur. Some of these functions are: 1) to facilitate the efficient
splicing and removal of intron sequences by binding to the cap binding protein (CBC),
which then initiates the assembly of the spliceosome 5, 6, 2) to assist the binding of protein
factors that allow the pre-RNA to be shuttled out of the nucleus to then participate in
translation in the cytoplasm1, 3, 3) to provide a site where protein factors bind which
assists in the start of protein synthesis, or translation initiation7, 8, and 4) to provide resistance to exonuclease enzyme degradation via regulated RNA decay pathways.1, 2
RNA degradation control sets the level of gene expression and serves as a point of
regulation in this process.1 Responses to environmental stimuli, like temperature,
119
nutrients, cytokines and hormones affect the coordination of RNA decay of clinically
relevant sequences.9 It has also been recently shown that RNA decay is localized to a
central area within the cytoplasm, known as P-bodies, where certain genes can be repressed or expressed in a coordinated manner separate from translational activities.2
Thus, RNA decay is vital toward maintaining stringent quality control for recognizing and degrading aberrant RNAs that could be deleterious to the cell.
Moreover, RNA decay plays a vital role in defending against viral infections within the cell. Consequently, certain viruses have adapted to exploit the cap recognition machinery in order to proliferate. The influenza virus has an RNA polymerase that synthesizes viral RNA from short capped primers which were ‘stolen’ from the host RNA via a ‘cap-snatching’ endonuclease. The capped viral RNA then serves as a template for viral RNA replication10
Disruption of the normal dynamics of RNA turnover can have consequences that
manifest in certain disease states. For example, neoplasia, thalassemia and Alzheimer’s
disease, have abnormal RNA decay mechanisms. These conditions result in the
accumulation of aberrant RNAs within the cell and subsequently affect the levels of
proteins present.9
1.3 Chemical Structure of Capped RNA
The cap chemical structure on the 5’ end of capped RNA contains a 5’-N7- methylguanosine triphosphate linked through a 5’-5’ dinucleotide linkage, or m7GpppX
(Figure 3-1). The X nucleotide is most often guanosine, but adenosine is occasionally found.11 Cap (0) structure contains only one methylation on the N7 of the terminal
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guanosine for the above mentioned structure. The cap (1) structure has the first
nucleotide, either a G or an A, methylated at the 2’-OH position by a methyltransferase.
Following this progression, the cap (2) structure contains methylation on the 2’-OH
positions of the first two nucleotides from the 5’ end.6 A unique cap (4) structure is found in the protozoan Trypanosomatid, with 2’-O-methylations on the first ribose moieties, in addition to methylations on the first adenosine and the fourth uridine.12 This over-
methylated cap structure functions on small nuclear RNAs (snRNA), which are recognized by the nuclear transport protein Snuportin 1,13 for efficient transport to the
nucleus, where the snRNAs play a role in the splicing of nascent mRNAs.14
O
OH OH N NH O O O N O N NH2 O P O P O P O O O O O- H2N N N O OH N N CH O 3 Oligoribonucleotide
7-methylguanosine triphosphate bridge
Figure 3-1: Structure of m7GpppG, or capped RNA consisting of the dinucleotide triphosphate with a 5’-5’ linkage.
The in vivo cap synthesis is enzymatically accomplished by 3 distinct steps
(Figure 3-2). First, RNA triphosphatase cleaves the 5’-triphosphate of the nascent pre-
RNA to a diphosphate. The next step is done by guanylyltransferase which attaches a
GMP moiety to the terminal phosphate, creating a GpppN terminus. Lastly, a methyltransferase methylates the N7 of the terminal G using S-adenosyl-L-methionine
(SAME) as the methyl donor.6 The positive charge residing on the imidazole ring
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stemming from the methylation of the N7-guanosine is critical for recognition with a
variety of receptors that are involved with processing, transport and translation.1, 15
O O O O O HO P P P O B RNA triphosphatase O O O P P O B GMP•guanylyltransferase - HO O O O O - O O O O OH -3 PO4 O OH RNA RNA
OH OH OH OH O O O O O O O O B methyltransferase O P O P O P O OP P P O B N O O O O H2N N - H2N N N - O O O SAME O O O HN O OH O OH N N NH O O CH3 RNA RNA
Figure 3-2: In vivo enzymatic synthesis of capping on 5’ end of transcribing RNA. The RNA strand is at least 25 bases long when the cap is attached.
The RNA cap serves as a binding site for the cap-binding protein (CBP) within
the nucleus. This serves to facilitate the maturation of the RNA by splicing out of 5’-end
intronic sequences, and is required for splicing accuracy.6 The heterodimer CBP80-
CBP20 binds to the cap and shuttles the transcript from the nucleus into the cytoplasm.
After transport to the site of protein synthesis, the CBP is replaced by initiation factors
for translation initiation. The export process is enhanced by CBP binding to the cap, but
not strictly required.5
The cap structure of capped RNA provides a critical binding site for factors
during protein translation.7 The eukaryotic initiation factor, eIF4E, binds to the cap during the 48S initiation complex formation for translation. This rate-determining step is the critical step toward subsequent recruitment of other ribosomal subunits for translation.16 The binding of eIF4E to the m7GpppG moiety actually stabilizes the RNA
against degradation. In fact, there is a direct correlation between RNAs associated with
the ribosome and a decreased rate of decapping.1
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Certain 3’ elements can interact with the 5’ cap structure to coordinate the start of
RNA decay. The poly(A) tail is a 3’ modification to RNA transcripts, whose binding
factors indirectly interact with the cap via cap-binding proteins. This, in turn, is a
stabilizing interaction that inhibits decapping.1 The poly-A-binding protein (PABP)
interacts with eIF4E, thereby enhancing translation initiation. Once the poly(A) tail is
cleaved to about 12 bases long, the protection is lost, and the RNA transcript becomes
vulnerable to RNA decay. The idea is that once the binding site is removed for PABP,
then the stabilizing interactions are lost. Thus there is a synergistic effect of both the 5’
and 3’ elements in RNA stability.1, 7
1.4 Role of Capped RNA in Regulating RNA Decay
RNA decay or degradation is important for cell vitality in maintaining homeostasis, and as such, the lifetime of mRNA is a highly regulated process. It has been found that mRNAs that code for proteins involved in similar processes have a similar half-life.2 RNA decay can be categorized into two main pathways: deadenylation-
independent and deadenylation-dependent. The three deadenylation-independent decay
mechanisms include endonucleolytic RNA decay, or RNA interference (RNAi), nonsense
mediated decay, and non-stop decay.
1.4.1 Deadenylation-Independent Decay Mechanisms
RNAi is a decay pathway where cleavage occurs in the middle of the targeted
RNA strand by an endonuclease prior to poly(A) tail removal. The cleavage sites are usually sequence specific, targeting a particular gene.1 This is a critical event, where the
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ATP-dependent endonuclease Dicer, cleaves double-stranded RNA to generate small
RNAs roughly 23 nucleotides long, endogenous microRNAs (miRNA), that are
complementary or partially complementary to specific sequences targeted for destruction.
The RNAi-induced silencing complex then recruits an endonuclease, the Argonaut
protein, to degrade the target mRNA. miRNAs have been also reported to have a dual
mechanism for gene repression: translation repression and inducing poly(A) tail loss
leading to degradation.17
This combined effect on RNA degradation leading to targeted gene repression has
lead a surge in research to chemically synthesize sequence-specific short-interfering
RNAs (siRNA) to be exogenously administered for the purposes of silencing genes in
therapeutic applications.17-20 A series of modified siRNAs containing an internal ribo-2,4-
difluorotolyl nucleotide were synthesized and found to be slightly destabilizing in terms
of thermal denaturation compared to other non-canonical base pairs, to impart luciferase
gene-silencing activity, and to have improved resistance to serum nuclease in vitro.18
Another study showed that a systemic administration of lipid-encapsulated siRNA targeting apolipoprotein B mRNA had an in vivo dose-dependent effect in non-human primates, which lead to significant reductions in serum cholesterol, Apoprotein B, and low-density lipoprotein.19 These results illustrate that the potential of this class of
theraputics is warranted.
Nonsense-mediated decay is a deadenylation independent decay mechanism
whereby aberrantly transcribed RNAs that contain a premature stop codon are decapped
prior to the removal of the 3’ poly (A) tail. This prevents the cell from accumulating
truncated proteins that could be deleterious to the cell. This quality control measure
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occurs in the nucleus as a ‘proofreading’ mechanism. It is estimated that roughly one- third of all intron splicing results in RNAs that contain these premature stop codons.1
Non-stop decay is another deadenylation independent decay pathway that occurs when there is an absence of a stop codon within the reading frame of the RNA transcript.
The protein factor Ski7p interacts with the stalled ribosome, which then recruits the exosome for decay.1
1.4.2 Deadenylation-Dependent Decay Mechanisms
Research has shown that in yeast and mammalian decay mechanisms, that
deadenylation-dependent decay is the most prevalent category of degradation pathway.2
The deadenylation-dependent decay mechanisms are divided into two categories: 5’ to 3’ decay and 3’ to 5’ decay (Figure 3-3). Of these two, the 5’ to 3’ has been reported to be the predominant one in eukaryotes.21 The preferential substrate of the 5’ to 3’ decay
pathway are ribonucleotides that are at least 25 bases long, as it is thought that the RNA
strand may allosterically interact with the decapping enzyme.1 In this pathway, after the
poly(A) tail is shortened to roughly 12 bases long, the decapping holoenzyme Dcp1/Dcp2
cleaves the pyrophosphate bond between the α and β phosphates of the cap structure,
resulting in m7GDP and the free 5’-phosphate oligoribonucleotide. The Dcp2 is the
catalytic subunit whose activity is enhanced by Dcp1, but is not strictly required for
activity. This enzyme is also conserved among eukaryotic organisms.1 After decapping,
the 5’-phosphate ribonucleotide is hydrolyzes to the monomeric nucleotides from the 5’
to 3’ direction by an exonuclease enzyme. The m7GDP fragment product of the
decapping reaction is then cleaved at the pyrophosphate bond leaving m7GMP and a free
125
phosphate by DcpS or the scavenger enzyme.1, 7 It was found that unmodified capped
RNA tends to undergo this degradation pathway since their 5’ sequences disappeared faster. Assays containing RNA with chemically modified cap structures underwent degradation by the alternative 3’ to 5’ decay pathway.7
3'-deadenylation 7 7 m GpppNNNnAAAn m GpppNNNn 3 ' t ex o o 5 p so ' /2 m hy 1 e d cp ro D (1) (2) l ys is
7 7 + npN m Gpp + pNNNn m Gpp
5' to 3' hydrolysis DcpS exonuclease
7 npN m Gp + p
Figure 3-3: Two possible RNA degradation pathways: 1) Deadenylation followed by decapping, then 5’ to 3’ hydrolysis of the 5’ phosphate oligonucleotide by exonucleases, 2) Deadenylation followed by 3’ to 5’ hydrolysis resulting in m7GDP, which then is hydrolyzed by DcpS to m7GMP and phosphate.
As mentioned, the 3’ to 5’ decay mechanism represents the second type of
deadenylation-dependent decay pathway. Removal of the poly(A) tail initiates the
degradation in a similar way to the 5’ to 3’ decay mode. This is then followed by
hydrolysis of the RNA chain into the monomeric nucleotides by the exosome, which is a large barrel-shaped collection of exonucleases, in the 3’ to 5’ direction. The remaining m7GDP is cleaved between the original β and γ phosphates by DcpS enzyme.21 This
enzyme is part of a class of nucleotide hydrolyases/pyrophosphatases that act on the α-
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phosphate of ribonucleotides, which uses a histidine triad (HIT) in the active site for
catalysis.1 It has been shown that this pathway is dominant with multiple-methylated
(7,3’) capped structure substrates m2 GpppG, or methylene bridged cap structures
(7,3’) m2 GppCH2pG, since the chemical modification slows the enzyme kinetics of the 5’ to
3’ decay pathway.7
1.5 References
1. Coller, J., Parker, R., "Eukaryotic mRNA Decapping". Annual Reviews in Biochemistry, 2004, 73, 861-890.
2. Wilusz, C.J., Wilusz, J., "Bringing the Role of mRNA Decay in the Center of Gene Expression". Trends in Genetics, 2004, 20(10), 491-497.
3. Lewis, J.D., Izaurralde, E., "The Role of the Cap Structure in RNA Processing and Nuclear Export". European Journal of Biochemistry, 1997, 247, 461-469.
4. Grudzien, E., Stepinski, J., Jankowska-Anyszka, M., Stolarski, R., Darzynkiewicz, E., Rhoads, R., "Novel Cap Analogs for In Vivo Synthesis of mRNAs with High Translational Efficiency". RNA, 2004, 10, 1479-1487.
5. Worch, R., Niedzwiecka, A., Stepinski, J., Mazza, C., Jankowska-Anyszka, M., Darzynkiewicz, E., Cusack, S., Stolarski, R., "Specificity of Recognition of mRNA 5' Cap by Human Nuclear Cap-Binding Complex". RNA, 2005, 11, 1355- 1363.
6. Schoenberg, D.R., Maquat, L.E., "Re-Capping the Message". Trends in Biochemical Sciences, 2009, 34(9), 435-442.
7. Grudzien, E., Kalek, M., Jemielity, J., Darzynkiewicz, E., Rhoads, R., "Differential Inhibition of mRNA Degradation Pathways by Novel Cap Analogs". Journal of Biological Chemistry, 2006, 281, 1857-1867.
8. Kowalska, J., Jukaszewicz, M., Zubrek, J., Ziemniak, M., Darzynkiewicz, E., Jemielity, J., "Phosphorothioate Analogs of m7GTP are Enzymatically Stable Inhibitors of Cap-dependent Translation". Bioorganic & Medicinal Chemistry Letters, 2009, 19, 1921-1925.
9. Guhaniyogi, J., Brewer, G., "Regulation of mRNA Stability in Mammalian Cells". Gene, 2001, 265, 11-23.
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10. Dias, A., Bouvier, D., Crepin T., McCarthy, A., Hart, D., Baudin, F., Cusack, S., Ruigrok, R., "The Cap-Snatching Endonuclease of Influenza Virus Polymerase Resides in the PA Subunit". Nature 2009, 458, 914-918. 11. Peyrane, F., Selisko, B., Decroly, E., Vasseur, J.J., Benarroch, D., Canard, B., Alvarez, K., "High-Yield Production of Short GpppA- and m7GpppA-Capped RNAs and HPLC-Monitoring of Methyltransfer Reactions at the Guanine-N7 and Adenosine-2'O positions". Nucleic Acids Research, 2007, 35(4), e26.
12. Lewdorowicz, M., Yoffe, Y., Zuberek, J., Jemielty, J., Stepinski, J., Kierzek, R., Stolarski, R., Shapira, M., Darzynkiewicz, E., "Chemical Synthesis and Binding Activity of the Trypanosomatid Cap-4 Structure". RNA, 2004, 10, 1469-1478.
13. Sekine, M., Ushioda, M., Wada, T., Seio, K., "Synthesis of TMG-Capped RNA- DNA Chimeric Oligonucleotides". Tetrahedron Letters, 2003, 44, 1703-1707.
14. Moore, M.J., Query, C.C., Sharp, P.A., Splicing of Prescursors to mRNA by the Spliceosome, in The RNA World, Gestland, R.F., Atkins, J.F., Editor. 1993, Cold Spring Harbor Laboratory Press: Cold Spring Harbor, NY. p. 303-357.
15. Mikkola, S., Salomaki, S., Zhang, Z., Maki, E., Lonnberg, H., "Preparation and Properties of mRNA 5'-cap Structure". Current Organic Chemistry, 2005, 9, 999- 1022.
16. Kowalska, J., Lukaszewicz, M., Zuberek, J., Darzynkiewicz, E., Jemielity, J., "Phosphoroselenoate Dinucleotides for Modification of mRNA 5' End". ChemBioChem, 2009, 10, 2469-2473.
17. Roush, S.F., Slack, F.J., "Micromanagement: A Role for MicroRNAs in mRNA Stability". ACS Chemical Biology, 2006, 1(3), 132-134.
18. Xia, J., Noronha, A., Toudjarska, I., Li, F., Akinc, A. Braich, R., Frank- Kamenetsky, M., Rajeev, K.G., Egli, M., Manoharan, M., "Gene Silencing Activity of siRNAs with a Ribo-difluorotoluyl Nucleotide". ACS Chemical Biology, 2006, 1(3), 176-183.
19. Zimmermann, T.S., Lee, A.C.H., Akinc, A., Bramlage, B., Bumcrot, D., Fedoruk, M.N., Harborth, J., Heyes, J.A., Jeffs, L.B., John, M., Judge, A.D., Lam, K., McClintock, K., Nechev, L.V., Palmer, L.R., Racie, T., Rohl, I., Seiffert,j S., Shanmugam, S., Sood, V., Soutschek, J., Toudjarska, I., Wheat, A.J., Yaworski, E., Zedalis, W., Koteliansky, V., Manorharan, M., Vornlocker, H., MacLachlan, I., "RNAi-Mediated Gene Silencing in Non-Human Primates". Nature, 2006, 441, 111-114.
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20. Hall, A.H.S., Wan, J., Spesock, A., Sergueeva, Z., Shaw, B.R., Alexander, K.A., "High Potency Silencing by Single-Stranded Boranophosphate siRNA". Nucleic Acids Research, 2006, 34(9), 2773-2781.
21. Parker, R., Song, H., "The Enzymes and Control of Eukaryotic mRNA Turnover". Nature Structural and Molecular Biology, 2004, 11(2), 121-127.
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2. Capped RNA Synthetic Introduction
2.1 Introduction
The synthesis of capped RNA can be grouped into categories that are either
mainly enzymatic, chemical, or a combination of both. Due to the instability of the N7-
methyl guanosine moiety on the terminal end of the molecule, this compounds synthesis
and purification has proven difficult.1 In general, chemical methods have been
inadequate to supply the material for research to investigate capped RNAs binding
interactions with the variety of protein factors and enzymes for cellular processes.2
2.2 Enzymatic Methods
Previous reports of the enzymatic synthesis of capped RNA relied on a primarily non-chemical route to produce capped RNA, having used in vitro enzymatic methods with polymerases. One drawback in using polymerases for the incorporation of the cap moiety onto the oligomer from the DNA template is the potential for the reverse orientation of the cap moiety.2 Much research has been undertaken toward developing cap analogs that hinder the reverse incorporation by variety of chemical modifications.3-6
A small scale enzymatic synthesis of short capped RNA was reported using a bacteriophage T7 DNA primase fragment for the transcription of GpppACn and
7 7 m GpppACn in nanomolar quantities. Past methods for synthesizing capped RNA
containing a 5’ terminal A utilized a bacteriophage T7, T3 or SP3 DNA-dependent RNA
polymerase, which resulted in lower yields. The polymerase enzymes have a specificity
for guanosine as the first transcribed base position. The alternate primase system uses the cap dinucleotide triphosphate, m7GpppA, as the first incorporated nucleotide during
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transcription, which was convenient for this synthesis since there was only one adenosine
in the template.
This adapted enzymatic system was a truncated T7 DNA primase helicase
fragment that contained the active domain, which was similar in activity to the full-length enzyme, but was superior in expression yields, solubility and for purification of the
desired product. Using different DNA templates and varying reaction times, substrates
7 GpppACn and m GpppACn were obtained up to 42% (2-84 nmol) and 52% (3-103 nmol)
yields respectively.7
Another enzymatic method was reported by Chung et al. that uses a DNA plasmid transcription of a 5’-triphosphate RNA.8 This was followed by an enzymatic treatment
using guanylyl transferase and GTP to form GpppN, which was followed by enzymatic
methylation using a methyltransferase and S-adenosylmethionine as the methyl donor.
7 The resulting capped RNA product was m GpppNn, which was 67 residues in length. No
yields are reported, as the focus of this paper was to generate substrates to study the
cleavage activity of viral RNA polymerase on various lengths of capped RNA substrates,
the product of which subsequently formed primers for viral transcription.
2.3 Combination of Enzymatic and Chemical Methods
A majority of the reported methods involved a combination of chemical and
enzymatic means to synthesize capped RNA. 4, 6, 9-12. Brownlee et al. reported the solid-
phase chemical synthesis of 5’-diphosphate oligoribonucleotides, that were 11-13
residues in length, that were then capped using an enzymatic system containing
guanylyltransferase, methyltransferase and SAME to give the crude capped RNA
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oligomers in nanomolar quantities (no yields reported).9 Subsequent analysis and
purification of the capped RNA oligonucleotides were performed by gel electrophoresis and autoradiography of 32P-labeled products. These products were then used as primers
for viral RNA polymerase assays.
Matsuo et al. reported the synthesis of the capped structure, m7GpppA that was enzymatically condensed with a dimer using a bacteriophage gene 4 T7 primase, which formed the isotopically-labeled capped trimer, m7GpppACC (no yields reported).10 Here, the primase enzyme could accommodate an adenosine as the first transcribed nucleotide in the capped sequence. As a result, there was no risk of a reversely incorporated m7G as the first residue. Additionally, GTP was not requried for chain elongation.10
Another report from Peng et al. described the T7 polymerase enzymatic capping
3’,7 3’,7 of a m2 GpppGA3 or m2 GpppGA5, using a chemically synthesized cap structure as
the starting nucleotide on the DNA template.4 In combination with 3’-O-methyl and N7-
methyl modifications on the cap structure, the correctly-orientated incorporation of the
capped structure on the oligomers was achieved.
Sekine et al. then reported the enzymatic ligation of a capped RNA-DNA hybrid
oligonucleotide, having relied on the chemical solid-phase synthesis of a capped RNA, as
well as the chemical synthesis of the DNA fragment App7mer.11 The method for the
capped RNA portion was disclosed earlier with having an isolated yield of 20% for the
2,2,7 capped 2’-O-methylated RNA trimer, m3 GpppAmUmAm. This capped RNA trimer was synthesized via solid-phase methodology, using an acid-labile phosphoramidate
linkage to the CPG. The details of the solid-phase reaction conditions and reagents are
described later in this chapter. The crude capped RNA was then cleaved under acidic
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conditions, and the removal of the 3’ terminal phosphate was accomplished
enzymatically with alkaline phosphatase.13 No further purification was reported.
A further example of a combined methodology was reported by Kore et al., who described a chemical synthesis of locked-nucleic-acid (LNA) cap structure, m7(LNA)GpppG 1, that was used in a T7 polymerase transcription to form the hexamer
7 6 m (LNA)GpppGA6. This locked nucleic acid cap analog 61, containing a methylene
bridge joining the 2’-O and the 4’-C on the N7-methylated guanosine moiety (Scheme 3-
61, avoids the reverse enzymatic incorporation into the newly transcribed oligomer,
which would otherwise occur in equal proportions if the 2’-O were unblocked on the
guanosine moiety. The enzymatic transcription with T7 polymerase resulted in 54%
capping efficiency with the LNA-capped transcript illustrates that the cap analog is
recognized as a substrate by the polymerase.6
Scheme 3-1: Synthesis of m7(LNA)GpppG cap analog to be used in transcription.
2.4 Chemical Methods
The area of purely chemical synthesis covers a variety of methods toward the
preparation of capped RNA, the dinucleotide triphosphate cap structure, and a host of
modified analogs of both.1, 2 These methods involve modifications of various structural elements as probes for biological assays, various different methods in pyrophosphate bond formation, solution-phase vs. solid-phase synthetic methods, and the inclusion or exclusion of protecting groups in the synthetic intermediates.
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The most common modification is the multiple-methylations of the capped
structure beyond the singular N7-methylation. These additional methylations do occur
naturally within the cell, serving a variety of functions.14-16 The 3’-O-methylation on the
N7-methyl-guanosine of the oligoribonucleotide has been frequently reported as being
the “anti-reverse cap analog” (ARCA), whose main purpose is to avoid the reverse
incorporation onto the growing chain of RNA during enzymatic transcription. A variety
of methylated analogs have been synthesized and tested as biological probes in a number
of assays targeted toward mRNA nuclear transport, transcription and translational
functionality.3, 5, 11, 16
Modification at the 2’-OH position on the N7-methyl-guanosine provides
sufficient steric bulk to also avoid reverse incorporation, despite not being directly
involved in the new covalent bond during chain elongation.17 Kore et al. described the
synthesis of the 2’-fluoro cap analogs, one with a singular N7-methyl guanosine and
another that is symmetric containing two N7-methyl guanosines.12 The cap structure
analogs had the 2’ modification on the guanosine moiety far from the phosphodiester backbone, which did not interfere with the phosphodiester bond formation during subsequent enzymatic transcription. Despite the small size of the fluoro substituent, the modification resulted in 70% and 52% respective capping efficiency with using a T7 polymerase in vitro transcriptional system.
Klein et al. reported the synthesis of hydrolysis-resistant unmethylated cap
analogues, Gp3G and Ap3A, that contain bismethylene bridges that replaced the pyrophosphate bridging oxygens (Scheme 3-2).18 The key intermediate that contained the
bisphosphonate bridge stemmed from reaction of two equivalents of lithium
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methanselenophosphonate anion with N,N-dimethylphosphonamidous dichloride in a
double Michaelis-Arbuzov reaction (Scheme 3-2, 64→66). After oxidation and methyl
ester hydrolysis, a bifunctional phosphinic phosphonic acid 67 is formed. This
intermediate then reacts to join the two 5’ hydroxyls on either protected guanosine 68 or
protected 6-azidoadenosine 70 in a tandem Mitsunobu reaction that forms the benzyl-
protected dinucleotide triphosphonate (not shown). Deprotection by hydrogenolysis
yielded GpCH2pCH2pG 69 or ApCH2pCH2pA 71 in 37% and 43% respectively (Scheme
3-2).
Scheme 3-2: Synthesis of bismethylene cap analogs GpppG 69 and ApppA 71.
A singular methylene replacement of a bridging oxygen was reported either
between the α and β or β and γ phosphorus on a series of analogs that also contained the
ARCA 3’-O-methyl modification on the unmethylated guanosine moiety (Scheme 3-3).19,
20 The P-C bond is extremely stable and would be resistant toward decapping enzymes if
135
the modification was placed at the scissile location. The incorporation of the methylene
bridge was accomplished by reacting the 5’-OH of guanosine 72 with
methylenebis(phosphonic dichloride) to form a 5’-methylenebisphosphonate guanosine
73, which was based on a related method from Yoshikawa’s 5’ phosphorylation of
nucleosides in trimethylphosphate.21 The synthesis of the activated intermediate involved
the formation of an electrophilic phosphorimidazolide using the
PPh3/dithiodipyridine/imidazole system on one of the guanosines (74 or 78). Subsequent
formation of the pyrophosphate bridge involved the reaction of this activated electrophile with a nucleophilic guanosine intermediate (75 or 77) in the presence ZnCl2 in DMF to
form the final cap structure. The α:β 76 or β:γ 79 cap isomers were obtained in 47% and
37% isolated yield (Scheme 3-3).20 When tested for enzymatic stability, the β:γ isomer
was found to be slowly hydrolyzed by DcpS,20, and the α:β isomer was found to be
resistant to Dcp2 hydrolysis.5
Scheme 3-3: Synthesis of α:β 76 or β:γ 79 methylene cap analogs.
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Thiophosphate cap analogues have an incorporated sulfur for a non-bridging
oxygen in the pyrophosphate chain of the triphosphate bridge. Since thiophosphate
modifications on nucleotides are isosteric, isoelectronic and possess similar charge
distribution to the unmodified oxygen, but are potentially resistant to enzymatic
hydrolysis, they are suitable probes for biological activity.22, 23 Analogs of this sort were
synthesized in the α, β, and γ positions by utilizing PSCl3 as a thiophosphorylation
reagent on guanosine, based on a method by first Yoshikawa 21 and then Whitesides24
(Scheme 3-4). The mixed pyrophosphate bond formation was achieved through an imidazolide-activation process similar to that in Scheme 3-3, that was then displaced by a incoming nucleophilic phosphate from GMP or m7GMP. This reaction was enhanced
23 with the Lewis acid ZnCl2 in DMF. The two diastereomers of α, β, and γ
7 7 7 thiodinucleotide triphosphates, (m Gppp(s)G 82 mGpp(s)pG 83, m Gp(s)ppG 86) were
synthesized also with 3’-O-methyl modifications to allow for correct positioning during
enzymatic transcription (10-30% isolated yield, Scheme 3-4).23, 25 These compounds
were then assessed for binding affinity to eIF4E by fluorescence titration, tolerance to the
DcpS decapping enzyme, as well as effectiveness with in vitro translational system.23, 25,
26 In general, all 3 thiophosphate analogs had higher binding to eIF4E as determined by
fluorescence titration, and were subsequently better inhibitors of cap-dependant
translation.25 The ARCA D2 β-isomer (later eluting diastereomer on reverse-phase
column) was resistant to hydrolysis by human Dcp2 enzyme.27 The γ analog was resistant to human DcpS enzymatic hydrolysis, regardless of the stereochemistry of the P-center.25
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7 7 7 Scheme 3-4: Synthesis of α-m Gppp(s)G 82, γ-m Gp(s)ppG 83, and β-m Gpp(s)pG 86 thiophosphate cap analogs.
Selenophosphate modification are useful as biological probes in crystallographic studies over sulfur-containing analogs due to selenium’s defraction of radiation in
multiwavelength anomalous diffraction phasing techniques.28 The selenophosphate
analog in the β position of the pyrophosphate cap bridge was synthesized by generation
of an in situ selenating reagent 88, made from trimethylsilylphosphite and elemental
selenium in pyridine (Scheme 3-5, 87→88). The selenophosphate displaced the
7,2’-O imidazolide group on the activated m2 GMP imidazolide 89 in the presence of ZnCl2
7,2’-O 7,2’- in DMF to form the product m2 Gpp(Se) 90. The coupling to form the final m2
O Gpp(Se)pG occurred by reaction of GMP imidazolide 62 with 90 to give seleno-cap analog 91 in 40% conversion and 30% overall isolated yield (Scheme 3-5).28
Both ARCA isomers, D1 and D2, of the β-selenophosphate cap analog had
respective 2x and 4x higher affinity to eIF4E as determined by fluorescence titration. In addition, under in vitro transcription using a rabbit reticulocyte lysate system, both β-
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ARCA analogs were transcribed as a 5mer and a 1750mer 2.2x and 2.4x more efficiently
than the non-selenated ARCA cap analog.28
O O N NH N NH Me SiO OSiMe O O ZnCl ,DMF O O 3 3 1. Se, pyridine N 2 P + N N O P O N P O N NH2 O P O P O N NH2 OSiMe 3 2. TEA, MeOH Se O O Se O O
OH OCH 3 OH OCH3 87 88 89 90 O O H3CO OH N NH N NH O O O O N N O P P N P O N NH2 O P O N N NH 90,ZnCl2, O O 2 O O H2N N N O Se O O DMF HN OH OH N OH OH O 62 91
7,2’-O Scheme 3-5: Synthesis of β-m2 Gpp(Se)pG cap analog 91.
Modifications at the N7 on the guanosine, with groups other than the methyl, have
been performed on the dinucleotide triphosphate cap analogs. The N7-benzylated cap
analog was synthesized by using benzyl bromide for formation of the benzylated
guanosine intermediate, analogously to the use of methyl iodide used for the N7-methyl compound.29 The b7GpppG and its ARCA analog were in vitro transcribed as capped
mRNAs coding for Luciferase protein, and showed greater translational efficiency
compared to the N7-methyl.5, 30
The N7-ethyl analog was synthesized using ethyl bromide as the alkylated
reagent, in a similar manner as methyl iodide. This modification showed a decrease in
binding affinity for translational factor eIF4E5, but an increase in affinity to the cap-
binding protein.31 In comparison, the benzyl substituent on the N7 was shown to be
sandwiched between two tryptophan aromatic side-chain moieties of the protein eIF4E.
This benzyl moiety can stack with the aromatic rings, whereas the ethyl causes steric
clashes with those groups.5
139
The reactions for forming the pyrophosphate bond on the triphosphate bridge of
the cap structure involve primarily an activated nucleotide intermediate, containing a
suitable leaving group on the terminal phosphate, and a separate nucleophilic phosphate
oxygen that attacks the electrophilic phosphorus center displacing the leaving group. This
reaction often includes the use of a cationic metal as a Lewis acid to reduce the charge
build-up of converging negatively-charged phosphates, although not strictly necessary.1, 2
Depending on the desired target, the activated intermediate could be the nucleotide monophosphate or diphosphate whose coupling to the other synthon results in the cap structure, or capped RNA. The activated forms include a variety of leaving groups using a variety of Lewis acids and reaction conditions. The leaving groups include: S-thiophenylate, methoxyphenylthioate, morpholidate, chloroquinoylate and imidazolide.1
The thiophenyl group has been used in conjunction with iodine for pyrophosphate
bond formation. The S-thiophenyl activated m7GDP 92 (Scheme 3-6) was reacted with
+ 7 + GMP (Bu3NH form) to form the cap structure m GpppG (NH4 form) 93 in the presence
of iodine in pyridine resulting in 48% yield after paper electrophoresis purification.32
2,2,7 Additionally, the S-thiophenyl activated m3 GMP coupled with GDP in the presence
2,2,7 of iodine with a pyridine/DMF solvent system to form m3 GpppG in 59% yield after paper electrophoresis (not shown).33
140
Scheme 3-6: Synthesis of m7GpppG 93 using S-thiophenyl activated m7GDP.
A report showed the protected and 4-methoxythiophenyl-activated m7GDP was reacted with either AMP or protected GMP in the presence of AgNO3 to form the cap structure 93 (Scheme 3-7).34 The presence of the methoxy group on the phenyl ring
helped to stabilize the P-S during the synthetic route. This route used a trimethoxytrityl
group on the exocyclic guanosine N2 amino group and a methoxymethylene ortho ester
connecting the 2’ and 3’ hydroxyl groups which increased the solubility of the reactants
during the synthesis. After acidic deprotection and paper electrophoresis purification, the
final products m7GpppA and m7GpppG 93 were obtained in 57% and 45% yield
respectively.34
Me O O N NH N NH TMTr TMTr O O N N N O N N N H H MeO S P O P O O O P O O O- O- O- 1. AgNO 93 + 3 O O O O 2. 80% AcOH H Me H Me 94 95
Scheme 3-7: Synthesis of m7GpppG 93 using 4-methoxythiophenyl activated m7GDP.
Activation by a morpholidate group on GMP 96 was reported in a synthesis in
7,3’-O 7,3-O conjunction with m2 GDP 97 to synthesize the cap structure m2 GpppG 98 in
141
DMSO (Scheme 3-8).4 The reaction used a stoichiometric amount of tetrazole that shortened the overall reaction time from 1 week to 3 days. The final cap structure was obtained in 60% yield after purification.
7,3’-O Scheme 3-8: Synthesis of m2 GpppG 98 using morpholidate activated GDP.
The 5-chloroquinoyl group has been used as another form of activated nucleotide for the triphosphate bridge formation.35, 36 This group was used to synthesize a capped
7 tetramer using m Gppchloroquinolide 99 and 5’-PO4-GACU in HMPA/MPD with CuCl2 for 23 hours (Scheme 3-9).2 The final product m7GpppGACU 100 was obtained in 37%
yield after reverse-phase and ion-exchange chromatography.
Scheme 3-9: Capped GACU synthesis using 5-chloroquinoyl activated m7GDP 99.
The most prevalent leaving group used for activation of a nucleotide mono- or
diphosphate has been the imidazolide group, that has been utilized under either aqueous
or anhydrous conditions with a variety of metallic Lewis acids. 1, 3, 16, 17, 37, 38 Sawai et al.
reported the coupling m7GDP imidazolide 101 with GMP in the presence of a variety of
142
divalent metal chloride salts in the aqueous solvent system 0.2M N-ethylmorpholine-HCl
buffer (pH 7) to form the cap structure m7GpppG 93 (Scheme 3-10). The Lewis acid metals screened in these series of reactions included Mn2+, Mg2+, and Cd2+. Of these ion
catalysts, Mn2+ had the highest conversion as seen via HPLC of 77% over the course of 4
days at 30°C and using 5 equivalents of activated capping reagent.37 This method was
then applied to the 5’-PO4-oligoribonucleotide, pACACUUGCUUU, to form the capped
RNA product 102. It was reported that the best HPLC yield was 35% conversion using
Mn2+ over 6 days; however, no isolated yield was reported. Attempts at a longer 72mer
capped RNA product was problematic, as the 40% conversion to capped RNA product
was not successfully separated from the starting oligonucleotide during purification.
Scheme 3-10: Synthesis of m7GpppG 93 and capped 11mer 102 using imidazolide activated m7GDP.
7,3’-O 7 The cap analogs m2 GpppG 98 and m dGpppG 103 were synthesized by
7 7 coupling GMP imidazolide 62 with m GDP or m dGDP in dry DMF with ZnCl2, which resulted in 78% and 88% yields respectively (Scheme 3-11).3 In a similar manner, the
cap analogs m7GpppG 93 and m7GpppA were synthesized by coupling a m7GDP
38 imidazolide 101 with either GMP or AMP in dry DMF with ZnCl2 (no reported yields).
The use of ZnCl2 was reported as assisting to increase the solubility of the reactants, as
143 well as serve as a catalyst to reduce the negative charge buildup upon the condensation of the anionic reactants.3
7,3’-O 7 Scheme 3-11: Synthsis of m2 GpppG 98 and m dGpppG 103 using imidazolide activated GMP. 7,3’-O 7 NDP = m2 GDP or m dGDP
This methodology was used to synthesize a capped 2’-O-methyl tetramer
7 6,6,2’ 2’ 2’ 3,2’ 14 m Gpppm3 Am Am Cm2 U in 40% yield, as well as a capped 2’-O-methyl trimer m7GpppAUA in 48% yield.31 These conditions were also used in the synthesis of a cap
7,3’-O 7 analog, m2 Gpppm G that contains both of the guanosine moieties methylated at the
N7 position in addition to one position on a 3’-OH. This over-methylated cap analog was furnished in 61% yield.16
There have been reports of solid-phase synthesis of capped RNA, however application of any general solid-phase synthesis methodology is hampered by the fact that the m7G moiety is highly unstable under basic conditions.1 Sekine et al. used a piecemeal approach in the synthesis of a capped trimer that starts with a 2-cyanoethyl- phosphoramidate linkage that is chain-elongated via standard phosphoramidite chemistry.13 After the oligonucleotide is 3 residues long, the 5’ end is phosphorylated twice to yield the 5’-diphosphate trimer. The base protections and phosphate cyanoethyl groups are then removed, making the linkage to the solid-phase acid labile. This is
2,2,7 followed by the addition of the activated 2’,3’-phenylboronylated m3 GMP
144
imidazolide in pyridine, which adds the cap moiety to the terminal end. Acidic cleavage
from the support is then followed by an enzymatic treatment with a calf intestinal alkaline
phosphatase to remove the 3’-phosphate from the capped RNA. The resulting yield for
the capped trimer was 20% from the 5’-diphosphate trimer.13
Another report of a solid-phase approach to capped RNA used a disulfide linkage
to a solid support for the synthesis of a capped trimer 2’-O-methyl-m7GpppApUpAp.39
The use of the disulfide linkage between the solid support and the oligoribonucleotide allows the use of a mild set of conditions for the final cleavage of the capped trimer from the support. The disulfide resin was prepared in four steps to yield a dimethoxytritylated disulfide containing an eleven carbon alkyl chain that is attached to the amide functionalized resin via a conventional amide bond. The detritylation of the resin can then undergo normal phosphoramidite chemistry to grow the RNA chain. After chain elongation, the 5’ terminal was phosphitylated using a DMT-protected phosphoramidite
40, which was then oxidized to a phosphate triester. Upon detritylation followed by base
and phosphate deprotection, the cap moiety was introduced via m7GDP imidazolide 101
in DMF with an 8x excess of ZnCl2. Finally, the capped oligonucleotide was cleaved from the support with treatment of DTT and triethylamine. The thioethyl group on the 3’-
PO4 terminus was allowed to eliminate as episulfide in aqueous MeOH with a small
amount of triethylamine. The final product obtained was 2’-O-methyl-m7GpppApUpAp
(no yields reported). No further purification is reported.39
145
2.5 References
1. Mikkola, S., Salomaki, S., Zhang, Z., Maki, E., Lonnberg, H., "Preparation and Properties of mRNA 5'-cap Structure". Current Organic Chemistry, 2005, 9, 999-1022.
2. Koukhareva, I.I., Lebedev, A.V., "Chemical Route to the Capped RNAs". Nucleosides, Nucleotides, and Nucleic Acids, 2004, 23(10), 1667-1680.
3. Stepinski, J., Waddell, C., Stolarski, R., Darzynkiewicz, E., Rhoads, R., "Synthesis and Properties of mRNA Containing the Novel "Anti-Reverse" Cap Analogs 7-Methyl(3'-O-methyl)GpppG and 7-Methyl(3'-deoxy)GpppG". RNA, 2001, 7, 1486- 1495.
4. Peng, A., Sharma, V., Singleton, S.F., Gershon, P.D., "Synthesis and Application of a Chain-Terminating Dinucleotide mRNA Cap Analog". Organic Letters, 2002, 4(2), 161-164.
5. Grudzien, E., Kalek, M., Jemielity, J., Darzynkiewicz, E., Rhoads, R., "Differential Inhibition of mRNA Degradation Pathways by Novel Cap Analogs". Journal of Biological Chemistry, 2006, 281, 1857-1867.
6. Kore, A.R., Shanmugasundaram, M., Irudaya, C., Vlassov, A., Barta, T., "Locked Nucleic Acid (LCA)-Modified Dinucleotide mRNA Cap Analogue: Synthesis, Enzymatic Incorporation, and Utilization". Journal of the American Chemical Society, 2009, 131, 6364-6365.
7. Peyrane, F., Selisko, B., Decroly, E., Vasseur, J.J., Benarroch, D., Canard, B., Alvarez, K., "High-Yield Production of Short GpppA- and 7meGpppA-Capped RNAs and HPLC-Monitoring of Methyltransfer Reactions at the Guanine-N7 and Adenosine- 2'O positions". Nucleic Acids Research, 2007, 35(4), e26.
8. Chung, T.D., Cianci, C., Hagen, M., Terry, B., Matthews, J.T., Krystal, M., Colonno, R.J., "Biochemical Studies on Capped RNA Primers Identify a Class of Oligonucleotide Inhibitors of the Influenza Virus RNA Polymerase". PNAS, 1994, 91, 2372-2376.
9. Brownlee, G.G., Fodor, E., Pritlove, D.C., Gould, K.G., Kalluge, J.J., "Solid Phase Synthesis of 5'-Diphosphorylated Oligoribonucleotides and Their Conversion to Capped m7Gppp-Oligoribonucleotides For Use as Primers For Influenza A virus RNA Polymerase in vitro". Nucleic Acid Research, 1995, 23, 2641-2647.
146
10. Matsuo, H., Moriguchi, T., Takagi, T., Kusakabe, T., Suratowski, S., Sekine, M., Kyogoku, Y., Wagner, G., "Efficient Synthesis of 13C, 15N-Labeled RNA Containing the Cap Structure m7GpppA". Journal of the American Chemical Society, 2000, 122, 2417- 2421.
11. Sekine, M., Ushioda, M., Wada, T., Seio, K., "Synthesis of TMG-Capped RNA- DNA Chimeric Oligonucleotides". Tetrahedron Letters, 2003, 44, 1703-1707.
12. Kore, A.R., Shanmugasundaram, M., Charles, I., Cheng, A.M., Barta, T.J., "Synthesis and Application of 2'-Fluoro-Substituted Cap Analogs". Bioorganic & Medicinal Chemistry Letters, 2007, 17, 5295-5299.
13. Kadodura, M., Wada, T., Seio, K., Moriguchi, T., Huber, J., Luhrmann, R., Sekine, M., "Solid-phase Synthesis of a 5'-terminal TMG-capped Trinucleotide Block of U1 snRNA". Tetrahedron Letters, 2001, 42, 8853-8856.
14. Lewdorowicz, M., Yoffe, Y., Zuberek, J., Jemielty, J., Stepinski, J., Kierzek, R., Stolarski, R., Shapira, M., Darzynkiewicz, E., "Chemical Synthesis and Binding Activity of the Trypanosomatid Cap-4 Structure". RNA, 2004, 10, 1469-1478.
15. Grudzien, E., Stepinski, J., Jankowska-Anyszka, M., Stolarski, R., Darzynkiewicz, E., Rhoads, R., "Novel Cap Analogs for In Vivo Synthesis of mRNAs with High Translational Efficiency". RNA, 2004, 10, 1479-1487.
16. Kore, A.R., Shanmugasundaram, M., "Synthesis and Biological Evaluation of Trimethyl-Substituted Cap Analogs". Bioorganic & Medicinal Chemistry, 2008, 18, 880- 884.
17. Jemielity, J., Fowler, T., Zuberek, J., Stepinski, J., Lewdorowicz, M., Niedzwiecka, A., Stolarski, R., Darzynkiewicz, E., Rhoads, R., "Novel "Anti-Reverse" Cap Analogs with Superior Translational Properties". RNA, 2003, 9, 1108-1122.
18. Klein, E., Mons, S. Valleix, A., Mioskowski, C., Lebeau, L., "Synthesis of Enzymatically and Chemically Non-hydrolyzable Analogues of Dinuleotide Triphosphates and Ap3P and Gp3G". Journal of Organic Chemistry, 2002, 67, 146-153.
19. Kalek, M., Jemielity, J., Stepinski, J., Stolarski, R., Darzynkiewicz, E., "A Direct Method for the Synthesis of Nucleoside 5'-Methylenebis(phosphonate)s from Nucleosides". Tetrahedron Letters, 2005, 46, 2417-2421.
20. Kalek, M., Jemielity, J., Darzynkiewicz, Z. M., Bojarska, E., Stepinski, J., Stolarski, R., Davis, R., Darzynkiewicz, E. , "Enzymatically Stable 5' mRNA Cap Analogs: Synthesis and Binding Studies with Human DcpS Decapping Enzyme". Bioorganic & Medicinal Chemistry, 2006, 14, 3223-3230.
147
21. Yoshikawa, M., Kato, T., Takenishi, T.,, "A Novel Method For Phosphorylation of Nucleosides to 5'-Nucleotides". Tetrahedron Letters, 1967, 50, 5065-5068.
22. Frey, P.A., Sammons, R.D., "Bond Order and Charge Localization in Nucleoside Phosphorothioates". Science, 1985, 228, 541-545.
23. Kowalska, J., Lewdorowicz, M., Zuberek, J., Bojarska, E., Wojcik, J., Cohen, L., Davis, R., Stepinski, J., Stolarski, R., Darzynkiewicz, E., Jemielty, J., "Synthesis and Properties of mRNA Cap Analogs Containing Phosphorothioate Moiety in 5', 5'- Triphosphate Chain". Nucleosides, Nucleotides, and Nucleic Acids, 2005, 24, 595-600.
24. Moran, J., Whitesides, G., "A Practical Enzymatic Synthesis of (Sp)-Adenosine 5'-O-(1-Thiophosphate) ((Sp)-ATP-a-S)". Journal of Organic Chemistry, 1984, 49, 704- 706.
25. Kowalska, J., Lewdorowicz, M., Zuberek, J., Grudzien-Nogalska, E., Bojarska, E., Stepinski, J., Rhoads, R., Darzynkiewicz, E., Davis, R., Jemielty, J. , "Synthesis and Characterization of mRNA Cap Analogs Containing Phosphorothioate Substitutions that bind tightly to eIF4e and Are Resistant to the Decapping Pyrophosphatase DcpS". RNA, 2008, 14, 1119-1131.
26. Kowalska, J., Jukaszewicz, M., Zubrek, J., Ziemniak, M., Darzynkiewicz, E., Jemielity, J., "Phosphorothioate Analogs of m7GTP are Enzymatically Stable Inhibitors of Cap-dependent Translation". Bioorganic & Medicinal Chemistry Letters, 2009, 19, 1921-1925.
27. Grudzien-Nogalska, E., Jemielty, J., Kowalska, J., Darzynkiewicz, E., Rhoads, R., "Phosphorothioate Cap Analogs Stabilize mRNA and Increase Translational Efficiency in Mammalian Cells". RNA, 2007, 13, 1745-1755.
28. Kowalska, J., Lukaszewicz, M., Zuberek, J., Darzynkiewicz, E., Jemielity, J., "Phosphoroselenoate Dinucleotides for Modification of mRNA 5' End". ChemBioChem, 2009, 10, 2469-2473.
29. Darzynkiewicz, E., Stepinski, J., Tahara, S.M., Stolarski, R., Ekiel, I., Haber, D., Heuvonen, K., Lehikoinen, P., Labadi, I., Lonnberg, H., "Synthesis, Conformation, and Hydrolytic Stability of 1,P3 Dinucleotide Triphosphates Related to mRNA 5'-cap, andComparative Kinetic Studies on Their Nucleoside and Nucleoside Monophosphate Analogs". Nucleosides Nucleotides, 1990, 9(599-618).
30. Darzynkiewicz, E., Stepinski, j., Ekiel, I., Goyer, C., Sonenberg, N., Temeriusz, A., Jin, Y., Sijuwade, T., Haber, D., Tahara, S., "Inhibition of Eukaryotic Translation by Nucleoside 5'-Monophosphate Analogues of mRNA 5'-Cap: Changes in N7 Substituent Affect Analogue Activity". Biochemistry, 1989, 28, 4771-4778.
148
31. Worch, R., Niedzwiecka, A., Stepinski, J., Mazza, C., Jankowska-Anyszka, M., Darzynkiewicz, E., Cusack, S., Stolarski, R., "Specificity of Recognition of mRNA 5' Cap by Human Nuclear Cap-Binding Complex". RNA, 2005, 11, 1355-1363.
32. Nakagawa, I., Konya, S., Ohtani, S., Hata, T., "A "Capping" Agent: P1-S-Phenyl P2-7-Methylguanosine-5' Pyrophosphorothioate". Synthesis, 1980, 556-557.
33. Iwase, R., Sekine, M., Tokumoto, Y., Ohshima, Y., Hata, T., "Synthesis of N2, N2, 7-trimethylguanosine Cap Derivatives". Nucleic Acid Research, 1989, 17, 8979- 8989.
34. Kamimura, T., Osaki, Y., Sekine, M., Hata, T., "An Effective Method for the Synthesis of the Cap Structure of Eukaryotic Messenger Ribonucleic Acids". Tetrahedron Letters, 1984, 25(25), 2683-2686.
35. Sekine, M., Iwase, R., Hata, T., Miura, K., "Synthesis of Capped Oligoribonucleotides by Use of Protected 7-Methylguanosine 5'-Diphosphate Derivatives." Journal of the American Chemical Society, 1989, 1, 969-978.
36. Fukuoka, D., Suda, F., Suzuki, R., Takaku, H., Ishikawa, M., Hata, T., Tetrahedron Letters, 1994, 35, 1063.
37. Sawai, H.W., H., Nakamura-Ozaki, A., "Synthesis and Reactions of Nucleoside 5'-Diphosphate Imidazolide. A Nonenzymatic Capping Agent for 5'-Monophosphorylated Oligoribonucleotides in Aqueous Solution." Journal of Organic Chemistry, 1999, 64, 5836-5840.
38. Stepinski, J., Zuberek, J., Jemielty, J., Kalek, M., Stolarski, R., Darzynkiewicz, E., "Novel Dinucleoside 5',5'-Triphosphate Cap Analogues. Synthesis and Affinity for Murine Translation Factor eIF4E." Nucleosides, Nucleotides, and Nucleic Acids, 2005, 24, 629-633.
39. Jemielity, J., Heinonen, P., Lonnberg, H., Darzynkiewicz, E., "A Novel Approach to Solid Phase Chemical Synthesis of Oligonucleotide mRNA Cap Analogs". Nucleosides, Nucleotides, and Nucleic Acids, 2005, 24(5-7), 601-605.
40. Guzaev, A., Salo, H., Azhayev, A, Lonnberg, H., "A New Approach for Chemical Phosphorylation of Oligonucleotides at the 5'-Terminus". Tetrahedron, 1995, 51(34), 9357-9384.
149
3. Synthesis of 5’-Capped Oligoribonucleotides
3.1 Introduction
In an effort to develop a synthetic method to produce 5’-capped oligoribonucleotides, there are a number of obstacles to overcome. Solubility of the reactants in the synthetic routes for the capping intermediates and the capped products have been problematic for previously reported syntheses. Also, separation is another issue during the purification of intermediates and for the cap structure and the capped
RNA products.
The dimethoxytrityl (DMT) protecting group has been used in DNA and RNA solid-phase synthesis for blocking the 5’-hydroxyl group on the growing chain of an oligomer attached to the solid support. One advantage is that each nucleotide coupling efficiency can be monitored via on-line UV/Vis detector at 436 nm, which tracks the absorbance of the DMT-cation, and consequently yields can be estimated from the peak area of each synthetic cycle. Another advantage is the lipophilicity of the DMT group which contains three aromatic rings. When the DMT group is left on the 5’ end of a final oligomer, the overall lipophilicity of the DMT oligomer is much greater as compared to the shorter, acylated failure sequences. This is very advantageous during reverse-phase purification, whereby the DMT-product elutes much later being well separated from failure sequences.
In our approach to a new synthetic method for capped RNA, we initially encountered problems in the synthesis of an intermediate for the capping reagent, namely
GDP. The reaction involved treating an activated GMP imidazolide with an excess of triethylammonium phosphate to form a new pyrophosphate bond for the GDP product.
150
During reverse-phase preparatory chromatography, the GDP product was not successfully
separated from the excess phosphate reagent. If the original GMP were chemically
modified with the DMT group, the purification of the subsequent intermediates should be
facilitated due to the different retention time on reverse-phase chromatography, and their
solubility in polar aprotic solvents should also be improved.
Kamimura reported the use of a trimethoxytrityl (TMT) group on both guanosine
amino groups for the synthesis of the cap structure, m7GpppG.1 The DMT group is ten
times more stable having one fewer methoxy group and is commonly used in nucleoside
chemistry, therefore we opted to use this group on the exocyclic amino group of the
guanosine capping reagent. We also intended to synthesize longer substrates than
previously reported in literature.
In the retrosynthetic analysis for capped RNA, we chose the final bond
connection to be the pyrophosphate bond between the α:β phosphates (Scheme 3-12).
This can allow for the separate synthesis and purification of the 5’-PO4-oligomer or
nucleotide, which would simplify the final coupling reaction mixture. This would also
focus all remaining modifications, namely N7-methylation, N2-tritylation, and
imidazolide-activation on the terminal phosphate of the capping reagent, m7GDPDMT imidazolide (Figure 3-4, 104).
151
Scheme 3-12: Retrosynthetic analysis for final coupling of capped RNA.
3.2 Synthesis of Capping Reagent m7GDPDMT imidazolide
The activated intermediate m7GDPDMT imidazolide 104 (Figure 3-4) is the key
intermediate for coupling to a 5’-PO4 nucleotide or oligonucleotide to form either the cap structure or capped RNA respectively. As such, the decision was made to start from GMP as the starting material for the route toward this key intermediate.
Figure 3-4: Key capping reagent intermediate, m7GDPDMT imidazolide 104.
3.2.1 Synthesis of GMPDMT: Transient Protection and Tritylation
The first step in the synthetic route toward m7GDPDMT imidazolide was the installation of the DMT group onto the guanosine amino group. As mentioned in the introduction, The TMT group was previously used in the synthesis of the cap structure
152
m7GpppG by Kamimura et al. on both of the coupling partners (see Scheme 3-7). In our
work, we were only interested in having the capping reagent guanosine possessing a
DMT group for greater flexibility during a planned synthesis particularly with oligomer
substrates. In an additional contrast, the DMT group would be the only “protecting”
group/ purification handle that would be carried throughout the entire synthesis until the very last deprotection step after coupling.
Transient silylation of the amino group was found to be necessary for its facile subsequent acylation in a previous report from the Jones lab.2. The use of TMSCl in
amino acylation provided a smoother overall reaction that avoided the production of
13 unwanted, dark-colored side products. It was shown that when labeled [2- C-1,7,NH2-
15 15 N3]-guanosine was treated with an excess of TMSCl, the signals from the N1 and
15N2 amino shifted toward that of the silylated form. This was indicated by a large
change in the 15N chemical shift of the N1 from a pyrrole-type nitrogen (153 ppm) to an
pyridine-type signal (217 ppm) for the O6 silylation. Also present was a shift for the
aniline-type nitrogen of the amino group (75 ppm → 86 ppm). This confirmed the
reaction of the TMS to these sites. In light of the advantageous effect of using TMSCl in
the transient silylation prior to acylation, it was decided that amino tritylation would
require the 5 eq to cover reaction at the 2’, 3’ hydroxyls, the O6 and the amino groups.
Commercially available GMP monohydrate (Na+ form) first underwent transient
protection using 5 eq excess TMSCl and Et3N in DMSO/pyridine to temporarily silylate
the ribose hydroxyls, as well as the guanosine O6 and amino group (Scheme 3-13, 105
→ 106). After silylation was complete, DMTCl and additional Et3N was added to
tritylate the amino group (106 → 107). The presence of the pyridine in the mixed solvent
153
system was sufficient as a nucleophilic catalyst to not require the use of DMAP in the
reaction mixture. After the tritylation was complete, a quick extraction with methylene
chloride and cold triethylammonium bicarbonate partitioned the fully protected product
into the organic layer.
The cleavage of the TMS groups was achieved with aqueous ammonia overnight at 4 °C (107 → 108). After evaporation of any residual organic solvent, the mixture was partitioned between water and ether where the product was isolated from any excess dimethoxytritonol by moving the product into the aqueous layer. By isolating the product by a series of extractions, no chromatography was required. The final isolation was via precipitation in ethyl acetate and ether which resulted in 75% isolated yield of GMPDMT
108 in the triethylammonium form.
Scheme 3-13: Transient silylation and amino tritylation of GMP.
154
3.2.2 Synthesis of GMPDMT imidazolide: Activation
Activation of the terminal phosphate by a suitable leaving group, like imidazole,
is an effective way to form a pyrophosphate bond when reacted with an incoming
phosphate. There have been numerous reports of using this group for the pyrophosphate bridge in the cap structure and related analogs.3-8 We selected the imidazolide as the
activated form in our planned synthesis of capped RNA due to the ease of handling of the
reagents, and their availability.
GMPDMT 108 was reacted using a system containing imidazole, 2,2’-
dithiopyridine, PPh3, Et3N under anhydrous conditions in DMF at room temperature
(Scheme 3-14). The Mitsunobu-type reaction was initiated by PPh3 attacking the
disulfide bond on dithiopyridine (Scheme 3-15, a), causing a reduction to the free thiol.
The phosphenium cation was subsequently attacked by a phosphate oxygen of GMPDMT b, that released 2-thiopyridine. Imidazole then displaced the phosphenium leaving group via an SN2 mechanism at the phosphate center that released POPh3 c. The resultant
product, GMPDMT imidazolide 109 was isolated in 96% yield by precipitation as the Na+ form in a solution of NaClO4 in acetone:ether (1:1).
155
Scheme 3-14: Imidazolide activation of GMPDMT forming GMPDMT imidazolide, 109.
Scheme 3-15: Mechanism of phosphorimidazolide formation for GMPDMT imidazolide, 109.
3.2.3 Synthesis of GDPDMT: Pyrophosphate Bond Formation
The pyrophosphate bond forming reaction for the intermediate GDPDMT involved
DMT reaction of the GMP imidazolide with an excess of both (Et3NH)3PO4 and ZnCl2 in anhydrous DMF at room temperature (Scheme 3-16). The ZnCl2 was used as a Lewis
acid to minimize the coulombic repulsions from highly anionic substrates, and to
156
facilitate the solubility of the reactants. When this reaction was performed without the use of ZnCl2, and it was found to be very slow in product formation, only ~20% conversion
DMT over 3-4 hours. The addition of ZnCl2 increased the conversion of GDP to 92% over
3-4 hours. After addition of EDTA to sequester the Zn2+ and reverse-phase purification,
GDPDMT 110 was isolated in 57% yield as the ammonium form.
Scheme 3-16: Pyrophosphate bond formation resulting in GDPDMT, 110.
3.2.4 Synthesis of m7GDPDMT imidazolide: N7-Methylation and Activation
The N7 of guanosine is the most basic site on the guanine moiety, having a pKa of
2.4, and therefore is the preferential site of methylation using a variety reagents.9 10
Previous papers reported using MeI to methylate the guanosine intermediate in the synthesis of cap analogs.5, 11 Other reagents such as dimethylsulfate and methyl-p-
toluenesulfonate have been reported for the methylation of purines.9 Since methylation
reactions can be non-specific, reacting at multiple nucleophilic sites, and since the N7-
methylated guanosine product may not be fully stable for chromatography, care had to be
taken to optimize the reaction conditions to favor the N7-methyl product.
DMT + Initial trials of methylation of GDP (NH4 form) using MeI with DIPEA in
anhydrous DMF showed facile over-methylation, with starting material still present. In
switching to the slightly less reactive Me2SO4, it was found that the identity of the
157
reactant counterion, the temperature during addition, the number of equivalents of
reagent and whether the Me2SO4 was added as a dilution in the appropriate solvent or as a
neat solution were all contributing factors for reaction optimization. The triethylammonium form of GDPDMT increased the solubility of the reactant GDPDMT, and led to rapid over-methylation. It was found the optimal counterion was the ammonium form of GDPDMT, which possessed the solubility for more optimal product formation, as
well as being convenient coming from the previous reverse-phase chromatographic
purification. The addition of a solution of Me2SO4 (1.38 M) in DMF reduced the
formation of di- and tri-methylation; however, when a neat solution of Me2SO4 was
added, rapid over-methylation was observed. This was counteracted by chilling the
reaction flask to 0 °C during neat reagent addition. Lastly, the equivalents of Me2SO4 were optimized to maximize the singularly methylated product (2.5 eq), while minimizing any di- and tri methylated by-products. The optimal reaction conditions are shown in Scheme 3-17, where after 21 hours, m7GDPDMT 111 was isolated as the crude
precipitate as the Na+ form from sieved-dried 0.08 M sodium perchlorate in acetone:ether
(1:1) in ~80% yield as determined by HPLC integration.
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Scheme 3-17: Optimized reaction conditions for N7-methylation of GDPDMT forming m7GDPDMT, 111.
The remaining synthetic step toward the capping reagent was the imidazolide activation at the terminal, β phosphate using the 2,2’-dithiodipyridine/PPh3/imidazole
system in an analogous manner for synthesizing GMPDMT imidazolide 109. The number
of equivalents of all reagents had to be increased to push the completion of the reaction
compared to conditions with the monophosphate. Overall, the optimized conditions are shown in Scheme 3-18 whereby m7GDPDMT imidazolide 104 was isolated as the crude precipitate as the Na+ form in ~80% yield as determined by HPLC integration. Attempts
at purifying the material via reverse-phase chromatography resulted in early detritylation
of the product, hydrolysis of the imidazole group, and overall very poor yields.
Therefore, the intermediate had to be used in the final coupling as the crude precipitate.
159
Scheme 3-18: Imidazolide activation of m7GDPDMT forming m7GDPDMT imidazolide, 104, as key capping reagent intermediate.
3.2.5 Conclusions
The inclusion of the DMT group on the guanosine amino group altered the solubility of the capping intermediates that allowed modification and optimization of reported literature procedures. Overall, the DMT group facilitated the intermediate’s solubility in a positive manner. Additionally, the positive charge resulting from the N7 methyation made the intermediates very prone to detritylation in aqueous solvents, even those that have been pH buffered. Therefore, the majority of the steps in the route to m7GDPDMT imidazolide were used as crude precipitates, with carefully chosen reaction
conditions that optimized the formation of the desired product while suppressing
undesired side-products. The summary of the synthetic scheme for m7GDPDMT imidazolide 104 is shown in Scheme 3-19.
160
O O TMS N NH N N
O N N NH N 2 TMSCl (5 eq), O N NH DMTCl (1.4 eq), -O P O -O P O TMS O Et3N(5eq) O Et3N(1.4eq) O- O- OH OH DMSO/pyridine DMSO/pyridine TMSO OTMS
105TMS 106 O O N N N NH N DMT DMT 2, 2'-dithiodipyridine (2 eq) O N N O N N N - Imidazole(5eq), O P O TMS - H O 3M aq. NH3 O P O PPh (2 eq) - O 3 O O- 4°C overnight Et N(1eq),DMF TMSO OTMS 3 OH OH 75% 96%
107 108
O O
N NH N NH DMT DMT O N N N O O N N N N H - H N P O O O P O P O O Me SO (2.5 eq) (Et3NH)3PO4 (5 eq), 2 4 O- O- O- DIPEA (1.0 eq) ZnCl (6 eq), DMF OH OH 2 OH OH 57% DMF
109 110
O H C 3 H C O N N 3 N DMT N O O N N N 2, 2'-dithiodipyridine (4 eq) H Imidazole (10 eq), O O N N N DMT O P O P O N O PPh3 (4 eq) H O O N P O P O O Et N(1eq),DMF O O OH OH 3 OH OH
111 (isolated as crude ppt) 104 (isolated as crude ppt)
Scheme 3-19: Synthetic route for key capping reagent m7GDPDMT imidazolide, 104.
3.3 Synthesis of the Cap Structure and 5’-Capped Oligoribonucleotides
7 7 3.3.1 Synthesis of m GpppG and m Gppp(s)G
The unmodified cap structure was synthesized using GMP (TEA+ form) as the incoming nucleophile to react with the capping reagent m7GDPDMT imidazolide (Scheme
3-20). The reaction required both 2 eq of the capping reagent 104 and ZnCl2 in DMSO at room temperature under a nitrogen atmosphere. After reacting 21 hours and quenching
161 the reaction with EDTA (TEA+ form), the DMT-cap structure 112a was purified using reverse-phase chromatography in 42% isolated yield. The DMT group on the N7-methyl guanosine moiety of the cap structure was sufficiently labile to be cleanly detritylated upon dissolving in purified water (pH ~5.5) and standing at room temperature over 18 hours. After a final HPLC purification, m7GpppG 113a was formed in 72% isolated yield, and 30% overall yield from GMP (Scheme 3-20).
O O H3C N N N NH OH OH DMT N O O O N N N O N NH2 104 (2 eq), ZnCl2 (2 eq), H - O O P O P O P O O P O O O - O O O O DMSO, RT H2N N N OH OH 42 % OH OH HN N 105 O 112a
H2O, RT 72 %
O H3C N N OH OH N O O O N NH2 O O P O P O P O O H2N N N O O O OH OH HN N O 113a
Scheme 3-20: Synthesis of the unmodified cap structure, m7GpppG, 113a.
7 The α-thiophosphate cap structure, m Gppp(s)G, was synthesized using guanosine monothiophosphate (GMP(s)) as the nucleophile in the capping reaction. GMP(s) was synthesized according to a procedure based on Yoshikawa’s selective 5’-phosphorylation on unprotected nucleosides12, that was later modified to include thiophosphorylation by
Whitesides (Scheme 3-21, 114→116).13
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Scheme 3-21: 5’-thiophosphorylation of guanosine to form GMP(s), 116.
7 The modified cap structure, m Gppp(s)G, was synthesized by reacting GMP(s) 116
with 2 eq of capping reagent 104 and 3 eq of ZnCl2 in DMSO at room temperature for 24 hours (Scheme 3-22). After quenching with EDTA (TEA+ form) and HPLC purification,
the DMT-thiocap structure 112b was isolated in 38% yield. After mild water detritylation
7 and final purification, the α-thiophosphate cap structure, m Gppp(s)G 113b was isolated
in 58% yield, with 22% overall yield from GMP(s). These yields represent both
diastereomers, D1 and D2, since the newly created α-thiophosphate contains a chiral
phosphorus center. Each were successfully separated on reverse-phase and characterized
separately (See Experimental section and Appendix).
163
O O H3C N N N NH OH OH DMT N O O O N N N S N NH2 104 (2 eq), ZnCl2 (3 eq), H - O O P O P O P O O P O O O - S O O O DMSO, RT H2N N N OH OH 38 % HN OH OH N 116 O 112b H2O, RT 58 %
O H3C N N OH OH N O O O N NH2 O O P O P O P O O S O O H2N N N OH OH HN N
O 113b
7 Scheme 3-22: Synthesis of α-thiophosphate cap structure, m Gppp(s)G, 113b.
3.3.2 Synthesis of 5’-Capped Oligonucleotides
Initial trials of the capping reaction required the synthesis of a variety of 5’- phosphorylated oligonucleotides. Solid-phase synthesis of the oligonucleotides was performed using standard phosphoramidite chemistry for the internucleotide linkages using pyridinium-trifluoroacetate/N-methylimidazole as the activator reagent. The coupling cycle and the reaction conditions are shown in Scheme 3-23. The different 2’ groups on the phosphoramidites dictated different recycling times on the AKTA
Oligopilot during the coupling step due to different steric influences which resulted in different reaction kinetics. The different times are as follows: 3 minutes for deoxyribonucleotide couplings, 10 minutes for 2’-O-methyl nucleotides, and 15 minutes for 2’-O-TBS protected ribonucleotides. The length of the recycling time allowed for
164
Scheme 3-23: Synthetic cycle for solid-phase synthesis of oligonucleotides on the AKTA Oligopilot. Recycling times vary depending on the nature of the 2’ substituent: 3 minutes for DNA, 10 minutes for 2’- O-methyl, and 15 minutes for 2’-O-TBS RNA.
165
sufficient contact time for the incoming 3’-phosphoramidite to react with the CPG-bound
5’-OH of the growing chain.
The 5’ terminal was phosphorylated using a non-nucleoside phosphoramidite that
allowed for DMT-on reverse-phase purification. The final phosphorylation was achieved
using 2-cyanoethyl-3-(4,4’-dimethoxy)-2,2-di(ethoxycarbonyl)propyl-1-N,N-diisopropyl
phosphoramidite (Scheme 3-24, 117→c).14 The purification and deprotection steps are
shown in Scheme 3-25 (a → c) which was the general sequence used for all the 5’-PO4-
oligonucleotide substrates used in the capping reaction.
iPr EtOOC COOEt EtOOC COOEt pyr·TFA, NMI O O N O O O-Oligo DMT P iPr DMT P MeCN O O CN CN 117 b HO-Oligo a
EtOOC COOEt 1. I ,pyr/H O (99:1) O 2 2 O O DMT P O-Oligo-3' 2. Base deprotection, O cleavage from CPG c
Scheme 3-24: 5’-phosphorylation with a non-nucleoside phosphoramidite 117 for the final coupling on the CPG-bound oligomer.
166
Scheme 3-25: Purification and deprotection sequence for 5’-PO4 oligonucleotides.
3.3.2.1 Capping Reaction Model Systems: pT8mer and 2’-O-Me-pGAUGC
As a model for longer oligonucleotides and to test the chemistry of installing the
5’-phosphate group, a 5’-PO4-T8mer was synthesized and used as a substrate in the capping reaction. Initial solubility tests of pT8mer showed that it was not soluble in
+ DMSO as the Na form, only when H2O was added did the oligomer solubilize and was detected via LCMS. To increase the solubility in an anhydrous organic solvent, like
DMF, the oligomer was converted to the TEA+ form, where it was found to be soluble.
Initial capping reaction trials showed 50% conversion to product at room temperature with 40 eq of both 104 and ZnCl2 over the course of 1 week.
To increase its efficiency, the capping reaction on pT8mer was run at both 30 °C and 45 °C, and it was shown that either level of elevated temperature was successful in increasing the conversion to ~65%. Increasing the amount of 104 and ZnCl2 to 100 eq also proved to improve conversion of DMT–capped T8mer to 84%. It was later discovered that conversion yields could be pushed even further if ZnCl2 was removed from the reaction system.
167
In a paper from Piccarilli for the solid-phase synthesis of AppDNA, which is an
intermediate in DNA ligation reactions, it was reported that elevated reaction temperature
was the most significant variable for increasing the yields of the pyrophosphate bond
formation. The reaction was between a solution-phase adenosine phosphorimidazolide
15 and CPG-bound 5’-PO4-DNA oligomer for forming the pyrophosphate bond in DMF .
The addition of ZnCl2, 6-(trifluoromethyl)-1-hydroxyl-benzotriazole, or tetrazole did not
increase the coupling efficiency; nor did the use of pyridine or DMSO as alternative
solvents.
Despite the fact that the above system is rather different in terms of reaction
phase, it was informative in probing different factors for increasing the yields of the
solution-phase reaction of the present capping reaction study. By eliminating ZnCl2 from the reaction mixture, it was found that using 100 eq of 104 in DMF at 30 °C over 4 days pushed the conversion of m7GDMTpppT8mer to 97%, based on HPLC UV integration.
Initial trials using a mixed sequence 2’-O-Me-p(GAUGC)2 oligonucleotide for the
capping reaction using the same conditions as the pT8mer were unsuccessful, both with
and without ZnCl2. As a result, a shorter mixed sequence 2’-O-Me-pGUAUC was
synthesized, and tested in the capping reaction. Using 50 eq of 104 over 6 days at 30 °C
resulted in 80% conversion to capped product, m7GDMTppp-2’-O-Me-GUAUC, and using
75 eq under similar conditions resulted in 95% conversion over the same time. Clearly,
the need for the ZnCl2 for capping oligonucleotides was not supported by these results.
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3.3.2.2 Capping Reaction of p(2’-O-Me-GAUGC)2, p(2’-O-Me-GUAUC)4 and p(2’-
OH- GUAUC)4
In the effort to synthesize longer capped RNA, a mixed sequence 2’-O-Me-
+ p20mer was prepared, p(2’-O-Me-GUAUC)4 (TEA form) for use as a substrate. The initial trials were unsuccessful using conditions successful for pT8mer and p(2’-O-Me-
GAUGC). There was no evidence of DMT-capped-20mer, m7GDMTppp-(2’-O-Me-
GUAUC)4, formation at 30 °C over 5 days with 40 eq of capping reagent 104. Increasing the temperature to 45-50 °C also showed no product formation over 20 hours. Increasing the temperature even further to 60 °C, product began to appear over 3 days, however, the majority of the capped product was detritylated and coeluted with the 2’-O-Me-p20mer starting material (Figure 3-5 a, b).
169
2.50 p20mer & m7Gppp20mer a) 2.00 DMTm7ppp20mer 1.50 AU 1.00 0.50
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes
7 -4 1776.3 m ppp20mer (M-4) 100 b) p20mer (M-4)-4
1666.2 %
0 m/z 1600 1650 1700 1750 1800 1850 1900 1950 2000
Figure 3-5: LCMS of partially detriylated capped 2’-O-Me-20mer ( reaction mixture a) HPLC of the crude capping reaction of p20mer showing minor portion of DMT capped 20mer at 8.3 minutes, with the majority of the product coeluting with the p20mer starting material at 5.6 minutes; b) ESI-MS of showing both M-4 ions of starting material 1666 m/z and capped 20mer 1776 m/z coeluting.
Temperature clearly played an important role in facilitating the kinetics of the
reaction, but more likely, acted to disrupt any secondary structure the lyophilized
oligonucleotide may have adopted while in aqueous solution or upon being introduced to
a polar, aprotic solvent like DMF. The next experiment was to preheat the starting
oligomer in DMF to disrupt any structure before adding capping reagent at a moderate
temperature (45 °C) for the reaction.
Preheating the 2’-O-Me-p20mer at 70 °C for 30 minutes and quick-quenching the
reaction tube in room temperature oil before adding capping reagent at 45 °C was the
most effective procedure at increasing yields, achieving 30% conversion after 12 days. A
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slow-cool method with periodic intervals of 70 °C heat application resulted in some product formation, but at a cost of a significant amount of detritylated capped 20mer.
7 DMT Optimized conditions for m G ppp-2’-O-Me-20mer preparation in DMF was
established as follows: preheating treatment at 70 °C for 2 hours, followed by a quench at
45 °C reaction temperature, the addition of 104 in portions (50 eq) every 2 days over the
course of 13 days. This procedure blended a balance between keeping the reaction as anhydrous as possible while pushing the reaction toward product formation (Figure 3-6).
After DMT-on HPLC purification, mild water detritylation, and a final column, m7Gppp-
(2’-O-Me-GUAUC)4 113f was isolated in 18% yield (Table 3-1, entry 6),, with recovery
of the p20mer starting material as 73% of the original oligomer. These taken together
account for 91 % of the initial amount of p20mer oligomer.
1.20 DMTm7ppp20mer 1.00 0.80 p20mer
AU 0.60 0.40 0.20
0.00
2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes
Figure 3-6: Crude HPLC of optimized DMTcap-2’-O-Me-20mer reaction, showing the clear separation of the DMT-m7ppp20mer from both the starting material and the excess capping reagent as the later eluting collection of peaks.
In applying this method to the p10mer, p(2’-O-Me-GAUGC)2, the reaction
7 DMT resulted in a conversion of 29% over 7 days to m G ppp(2’-O-Me-GAUGC)2. After
DMT-on HPLC purification, mild water detritylation, and a final column, the capped
7 10mer m Gppp(2’-O-Me-GAUGC)2 113e was isolated in 15% yield (Table 3-1, entry 5),
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with recovery of the p10mer starting material as 45% of the original oligomer. This taken
together account for 60 % of the p10mer oligomer.
The attention was focused on using this method in DMF to RNA 20mer, 2’-OH-
+ p(GUAUC)4 in the TEA form. Initial trials using DMF as the solvent were unsuccessful
in forming any product, despite raising the temperature to 60 °C. In switching to the more polar solvent DMSO, initial trials were successful in product formation. Optimized
7 DMT conditions resulted in 56% conversion to m G ppp(GUAUC)4 in 8 days. After DMT-
on HPLC purification, detritylation and a final column, the capped-RNA-20mer,
7 m Gppp(GUAUC)4 113g, was isolated in 40% yield (Table 3-1, entry 8), with recovery of the RNA p20mer as 18 % of the starting material oligomer. Taken together, both yields represent 58% of the starting material RNA p20mer.
Since DMSO was such a superior solvent over DMF in achieving product conversion with the RNA 20mer, the capping reaction was revisited on a previously used
2’-O-Me p20mer in DMSO. Over the course of 8 days at 45 °C, m7GDMTppp(2’-O-Me-
GUAUC)4 was formed in 72% conversion. After DMT-on purification, detritylation and
7 a final column, the capped-2’-O-Me 20mer, m Gppp-(2’-O-Me-GUAUC)4 113f, was isolated in 63 % yield as the ammonium salt (Table 3-1, entry 7).
172
The general capping reaction and summary of the reaction conditions are shown
in below in Table 3-1 for all substrates.
O H3C 1. 5'-PO -GMP or oligonucleotide, N N 4 O O O conditions below 7 DMT R O P O P O P O N N NH m GDP imidazolide, 104 O 2 2. RP HPLC X O O 3. Water OH OH
113a-g
Entry Product R R 2' group X eq. 104 eq. ZnCl Solvent T C % conversion % isolated yield 2 1 113a G OH O 2 2 DMSO RT >99 30 2 113b G OH S 2 2 DMSO RT >99 22 3 113c T8 H O 100 na DMF 30 97 - 4 113d GAUGC OMe O 75 na DMF 30 94 - 5 113e (GAUGC)2 OMe O 100 na DMF 45 29 15 6 113f (GUAUC)4 OMe O 300 na DMF 45 32 18 7 113f (GUAUC)4 OMe O 300 na DMSO 45 72 63 8 113g (GUAUC) OH O 300 na DMSO 45 56 40 4
Table 3-1: Summary of conversion yields for cap structures (entries 1-2) and capped oligonucleotides (entries 3-8).
3.4 Conclusions
A general synthetic method for the preparation of 5'-capped oligoribonucleotides
was established that utilizes the DMT group as a purification handle during reverse-phase
HPLC. The method is suitable for the synthesis of a wide range of substrates from the dinucleotide cap structure to 2’-O-Me and 2’-OH mixed sequence 20mers.
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3.5 Experimental
Preparation of N2-dimethoxytrityl guanosine monophosphate (GMPDMT, 108).
Guanosine monophosphate hydrate (2.03g, 4.8 mmol, Na+ form) was dried via
coevaporation with pyridine three times, leaving 10 mL on the last evaporation. The flask
was put under a nitrogen atmosphere and 30 mL of DMSO was added and the mixture
was stirred to form a suspension. The flask was set in a room temperature water bath, and
then triethylamine was added (3.3 mL, 24.0 mmol, 5.0 eq) followed by TMSCl (3.0 mL,
24.0 mmol, 5 eq). After 5 minutes, the water bath was removed. After 1 hour, additional
triethylamine was added (0.9 mL, 6.7 mmol, 1.4 eq) followed by DMTCl (2.28g, 6.7
mmol, 1.4 eq). After 1½ hours, the reaction was diluted with 50 mL of methylene
chloride, and then poured into a separatory funnel containing 120 mL of cold aqueous
TEAB and 40 mL methylene chloride. The aqueous layer was extracted with three 30 mL
portions of methylene chloride. The organic layers were pooled and reduced to an oil. To
this, 20 mL of acetonitrile, 20 mL of water, and 9 mL of concentrated ammonia was
added, and set at 4°C. After 18 hours, an additional 30 mL water was added and then the
solid was concentrated to remove the remaining organic solvent. The aqueous mixture
was extracted with three 30 mL portions of ether, and then the aqueous layer was reduced
to an oil. The oil was evaporated with triethylamine and MeOH, then dissolved in 8 mL
of MeOH, which was then precipitated in 400 mL of ethyl acetate:ether (1:1). The
precipitate was filtered and dried in a vacuum dessicator over fresh P2O5 overnight, resulting in 3.06g of 108 (3.53 mmol, TEA+ form, 74%) and was used without further purification. UV λmax 236 nm. The mass was confirmed via ESI-LCMS in negative
- mode as m/z (M-1) 664.2 (calculated for C31H31N5O10P : 664.6)
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Preparation of N2-dimethoxytrityl guanosine phosphorimidazolide (GMPDMT imid,
109). To a flask containing 2.37g (2.73 mmol) of 108 was added 25 mL of DMF, which was dried via coevaporation with acetonitrile three times, and placed under a nitrogen atmosphere. Imidazole (0.93g, 13.65 mmol, 5.0 eq), triethylamine (0.38 mL, 2.73 mmol,
1.0 eq), dithiodipyridine (1.20g, 5.46 mmol, 2.0 eq), and PPh3 (1.43g, 5.46 mmol, 2.0 eq)
were added directly, and the reaction was stirred at room temperature. After 3 hours, the reaction mixture was poured into 150 mL of a sieve-dried 0.08 M NaClO4 solution in
acetone:ether (1:1), then chilled at 4°C for 1 hour to precipitate the product as the sodium
salt. The resulting solid was collected by filtration and washed with two 20 mL portions
of cold acetone:ether (1:1), dried in a vacuum dessicator over fresh P2O5 overnight, and
resulted in 1.92g of 109 (2.61 mmol, 95%), which was then used without further
purification. UV λmax 233 nm. The mass was confirmed via ESI-LCMS in negative
- mode as m/z (M-1) 714.6 (calculated for C34H33N7O9P : 714.6)
Preparation of N2-dimethoxytrityl guanosine diphosphate (GDPDMT, 110). To a flask containing 1.70g (2.00 mmol) of 109 was added 15 mL of DMF, which was dried via coevaporation with acetonitrile three times, then placed under a nitrogen atmosphere. A stock solution of (Et3NH)3PO4 was prepared by combining 1.36 mL of 85% H3PO4, 9 mL
of Et3N, and 10 mL of DMF, which was dried via coevaporation with acetonitrile three
times, then placed under a nitrogen atmosphere. In a separate pear-shaped flask, ZnCl2
(1.63 g, 12.0 mmol, 6.0 eq) was dissolved in 10 mL DMF and dried via coevaporation with acetonitrile three times, then placed under a nitrogen atmosphere. To the guanosine
175
flask, a portion of the (Et3NH)3PO4 solution (10.2 mL, 11.0 mmol, 5.5 eq), and the ZnCl2 solution were added at room temperature. After 4 hours, the reaction mixture was poured into a 0.07 M EDTA (TEA+ form) solution, stirred until clear, and then extracted with
three 30 mL portions of ether. The aqueous layer was reduced to an oil, and then purified
by reverse-phase HPLC using a gradient of acetonitrile in 0.1M NH4HCO3 buffer (pH
1 7.3) to give 0.91g of 110 (1.14 mmol, 57%) in the ammonium form. UV λmax 236 nm; H
NMR (DMSO-d6, 500 MHz): δ 7.91 (s, 1H), 7.57 (br s), 7.29-7.28 (m, 4H), 7.21-7.17
(m, 5H), 6.86-6.84 (m, 4H), 5.12 (d, J = 3.5 Hz, 1H), 4.05-4.04 (m, 1H), 3.91-3.85 (m,
31 4H), 3.71 (s, 3H), 3.70 (s, 3H); P NMR (DMSO-d6, 500 MHz): δ -9.31 (d, J = 17.2
Hz), -10.31 (d, J = 17.2 Hz) The mass was confirmed via ESI-LCMS in negative mode
- as m/z (M-1) 744.5 (calculated for C31H32N5O13P2 : 744.6)
Preparation of N7-methyl-N2-dimethoxytrityl guanosine diphosphate (m7GDPDMT,
111). To a flask containing 0.76g (0.96 mmol) of 110 was added 10 mL of DMF and 1 mL of MeOH to dissolve the material. This was dried via coevaporation with acetonitrile three times, then placed under a nitrogen atmosphere, and chilled to 0 °C using an ice bath. To this, 167 uL of DIPEA (0.96 mmol, 1.0 eq) was added and allowed to stir 5 minutes, followed by the dropwise addition of 0.25 mL of Me2SO4 (2.69 mmol, 2.8 eq)
before removing the ice bath. After 18 hours, the reaction was poured into a stirred
solution of sieve-dried 0.08 M NaClO4 in acetone:ether (1:1), and chilled at 4°C for 10
minutes. The solid was collected and washed with ice-cold acetone:ether, dried in a
vacuum dessicator over fresh KOH, and resulted in 1.05g of 111 in the sodium form,
which was used without further purification. UV λmax 232 nm; The mass was confirmed
176
- via ESI-LCMS in negative mode as m/z (M-1) 758.1 (calculated for C32H34N5O13P2 :
758.6)
Preparation of N7-methyl-N2-dimethoxytrityl guanosine diphosphate imidazolide
(m7GDPDMT imid, 104). To a flask containing 0.99g (1.0 mmol) of 111 was added 10 mL of DMF, which was dried via coevaporation with acetonitrile three times, and placed under a nitrogen atmosphere. Imidazole (0.68 g, 10.0 mmol, 10.0 eq), triethylamine (0.14 mL, 1.0 mmol, 1.0 eq), dithiodipyridine (0.88 g, 4.0 mmol, 4.0 eq), and PPh3 (1.05 g, 4.0 mmol, 4.0 eq) were added directly, and the reaction was stirred at room temperature.
After 18 hours, the reaction mixture was poured into 200 mL of a stirred solution of sieve-dried 0.08 M NaClO4 in acetone:ether (1:1), then chilled at 4 °C for 1 hour to precipitate the product as the sodium salt. The resulting solid was collected by filtration, washed with two 20 mL portions of cold acetone:ether (1:1), dried in a vacuum dessicator over fresh KOH overnight, and resulted in 0.99 g of 104, and was used without further purification. UV λmax 232 nm; The mass was confirmed via ESI-LCMS in negative
- mode as m/z (M-1) 808.6 (calculated for C35H38N7O12P2 810.7)
Preparation of Guanosine monothiophosphate (GMP(s), 116). Guanosine monohydrate (0.68 g, 2.25 mmol) was dried via coevaporation with pyridine three times, placed under a nitrogen atmosphere, and then had 20 mL of triethylphosphate added and stirred to suspend the solid. The flask was heated to 130 °C for 5 minutes to dissolve the solid, then was removed and chilled to 0 °C in an ice bath. To this, 1.57 mL of 2,6- lutidine (13.50 mmol, 6.0 eq) and 0.47 mL of thiophosphoryl chloride (4.50 mmol, 2.0
177
eq) was added at 0 °C. After 3 hours, the reaction mixture was poured onto 150 mL of pet
ether to precipitate the crude product, and then was triturated with additional pet ether.
After decanting the solvent, the precipitate was dissolved in 0.1M NH4HCO3 buffer and
allowed to stir for 15 minutes. The aqueous layer was concentrated to an oil, and then
purified by reverse-phase HPLC using a gradient of acetonitrile in 0.1M NH4HCO3 buffer (pH 7.3) to give 0.50 g of 116 (1.20 mmol, 53%) in the ammonium form. UV λmax
1 251 nm; H NMR (D2O, 400 MHz): δ 8.15 (s, 1H), 5.87 (d, J = 6.0 Hz, 1H), 4.73-4.69
31 (m, 1H), 4.46-4.44 (m, 1H), 4.31-4.29 (m, 1H), 4.09-4.06 (m, 2H); P NMR (D2O, 400
MHz): δ 49.14. The mass was confirmed via ESI-LCMS in negative mode as m/z (M-1)
- 378.2 (calculated for C10H13N5O7PS : 378.3)
Preparation of N7-methyl-N2-dimethoxytrityl-GpppG (m7GDMTpppG, 112a). To a
flask containing 0.063 g of GMP (0.11 mmol, TEA+ form) was added 1 mL of DMSO, dried via coevaporation with acetonitrile three times, and then placed under a nitrogen atmosphere. In a separate flask, 0.030g of ZnCl2 (0.223 mmol, 2.0 eq) had added 2 mL of
DMSO, and was dried via coevaporation with acetonitrile in a similar manner, was added
to the GMP at room temperature. In a separate flask, 1 mL of DMSO was added to a flask
containing 0.26 g of 104 (0.22 mmol, 2.0 eq), which was dried via coevaporation with
acetonitrile in a similar manner, was added to the GMP-ZnCl2 flask. After 21 hours, the
reaction was poured onto 25 mL of a 0.015 M EDTA solution (TEA+ form), stirred until the solution cleared, and was extracted with three 15 mL portions of ether. The aqueous layer was concentrated to an oil, and purified by reverse-phase HPLC using a gradient of acetonitrile in 0.1M NH4HCO3 buffer (pH 7.3) to give 0.054 g of 112a (0.046 mmol,
178
42%) in the ammonium form. UV λmax 239.4 nm; The mass was confirmed via ESI-
- LCMS in negative mode as m/z (M-1) 1104.1 (calculated for C42H46N10O20P3 : 1103.8)
Preparation of N7-methyl-GpppG (m7GpppG, 113a). To a flask containing 0.054 g of
+ 112a (0.046 mmol, NH4 form), was added 20 mL of water. The solution was allowed to
stand sealed at room temperature for 18 hours, lyophilized, and the residue purified by
reverse-phase HPLC using a gradient of acetonitrile in 0.1M NH4HCO3 buffer (pH 7.3)
to give 0.028g of 113a (0.033 mmol, 72%) in the ammonium form. UV λmax 253.6 nm;
1 H NMR (D2O, 500 MHz): δ 8.98 (solvent exchange reduces intensity), 8.04 (s, 1H),
5.85 (d, J = 3.0 Hz, 1H), 5.77 (d, J = 6.0 Hz, 1H), 4.60 (t, J = 5.5 Hz, 1H), 4.51-4.50 (m,
1H), 4.44-4.42 (m, 1H), 4.41-4.39 (m, 1H), 4.37-4.29 (m, 3H), 4.27-4.18 (m, 3H), 3.98
13 (s, 3H); C NMR (D2O, 500 MHz): δ 160.7, 158.0, 157.1, 156.5, 151.8, 139.8, 139.1,
117.9, 110.5, 92.3, 89.5, 86.3 (d, JCP = 9.3 Hz), 86.3 (d, JCP = 8.3 Hz), 77.4, 76.6, 72.9,
31 71.6, 68.0 (d, JCP = 6.0 Hz), 66.9 (d, JCP = 5.2 Hz), 38.7; P NMR (D2O, 500 MHz): δ -
10.48 (d, J = 19.2 Hz, 2P), -22.08 (app t, J = 19.1 Hz, 1P) The mass was confirmed via
- HRESI in negative mode as m/z (M-1) 801.0743 (calculated for C21H28N10O18P3 :
801.0801)
7 DMT Preparation of N7-methyl-N2-dimethoxytrityl-Gppp(s)G (m G ppp(s)G 112b, D1
+ + and D2). GMP(s) 116 (0.067 g, 0.16 mmol, NH4 form) was converted to the TEA form by evaporation with triethylamine and acetonitrile, then was dissolved in 2 mL of DMSO and placed under a nitrogen atmosphere. ZnCl2 (0.066g, 0.49 mmol, 3.0 eq) was
dissolved in 2 mL of DMSO, dried via coevaporation with acetonitrile three times, and
179
then placed under a nitrogen atmosphere. Compound 104 (0.33 g, 0.31 mmol, 2.0 eq) was
dissolved in 2 mL of DMSO, dried via coevaporation with acetonitrile three times, and
placed under a nitrogen atmosphere stirring. To this solution, 116 and ZnCl2 were added.
After 24 hours at room temperature, the reaction was poured onto 25 mL of a 0.015 M
EDTA solution (TEA+ form), stirred until the solution cleared, and then extracted with
three 15 mL portions of ethyl acetate. The aqueous layer was reduced to an oil, and then
purified by reverse-phase HPLC using a gradient of acetonitrile in 0.1M NH4HCO3 buffer (pH 7.3) to give 0.072 g of 112b (0.061 mmol, 38%) in the ammonium form. UV
λmax 239 nm; The mass was confirmed via ESI-LCMS in negative mode as m/z (M-1)
- 1119.5 (calculated for C42H46N10O19P3S 1119.8)
7 Preparation of N7-methyl-Gppp(s)G (m Gppp(s)G 113b, D1 and D2). To a flask
+ containing 0.072 g of 112b (0.061 mmol, NH4 form) was added 20 mL of water. The
solution was allowed to stand sealed at room temperature for 18 hours, lyophilized, and
the residue was purified by reverse-phase HPLC using a gradient of acetonitrile in 0.1M
NH4HCO3 buffer (pH 7.3) to give 0.031 g of j (0.035 mmol, 58%) in the ammonium
1 form. UV λmax 252 nm; D1: H NMR (D2O, 500 MHz): δ 8.93 (s, solvent exchange
reduces intensity), 8.01 (s, 1H), 5.79 (d, J = 3.5 Hz, 1H), 5.70 (d, J = 6.0 Hz, 1H), 4.58-
4.56 (m, 1H), 4.49-4.47 (m, 1H), 4.40-4.38 (m, 2H), 4.32-4.26 (m, 3H), 4.22-4.15 (m,
13 3H), 3.95 (s, 3H); C NMR (D2O, 500 MHz): δ161.0, 158.6, 158.0, 156.4, 154.0, 151.8,
140.1, 118.5, 110.6. 92.2, 89.3, 86.4 (d, JCP = 9.6 Hz), 86. 2 (d, JCP = 9.6 Hz), 77.4, 76.5,
31 73.0, 71.7, 68.1 (d, JCP = 6.3 Hz), 66.9 (d, JCP = 5.3 Hz), 38.7; P NMR (D2O, 500
MHz): δ 43.33 (d, J = 26.1 Hz, 1P), -11.67 (d, J = 19.6 Hz, 1P), -24.21 (app t, J = 22.9
180
1 Hz). D2: H NMR (D2O, 500 MHz): δ 8.95 (solvent exchange reduces intensity), 7.97 (s,
1H), 5.89 (d, J = 3.0 Hz, 1H), 5.70 (d, J = 6.0 Hz, 1H), 4.56-4.54 (m, 1H), 4.46-4.44 (m,
1H), 4.39-4.34 (m, 3H), 4.28-4.23 (m, 3H), 4.20-4.16 (m, 2H), 3.94 (s, 3H); 13C NMR
(D2O, 500 MHz): δ 160.95, 158.44, 157.74, 156.33, 153.89, 151.66, 139.94, 118.35,
110.51, 92.32, 89.37, 86.21 (app t, J = 9.4 Hz), 77.41, 76.56, 73.02, 71.46, 68.32 (d, JCP =
31 5.8 Hz), 66.84 (d, JCP = 4.8 Hz), 38.70; P NMR (D2O, 500 MHz): δ 43.42 (d, J = 27.1
Hz, 1P), -11.68 (d, J = 19.6 Hz, 1P), -24.15 (dd, J = 19.6, J = 26.7 Hz, 1P). The mass was confirmed via HRESI in negative mode as m/z (M-1) 817.0573 (calculated for
- C21H28N10O17P3S 817.0573)
General Method for 5’-Capped Oligoribonucleotide Synthesis:
All 2’-O-methyl and 2’-OH oligoribonucleotides were prepared by solid-phase
synthesis using standard phosphoramidite chemistry for the coupling steps. For each
coupling step, the activator used was 0.22 M pyridinium trifluoroacetate/ 0.11 M N-
methylimidazole in acetonitrile. The final 5’ phosphorylation utilized a dimethyoxytrityl
(DMT) -protected phosphorylation reagent, 2-cyanoethyl-3-(4,4’-dimethoxy)-2,2-
di(ethoxycarbonyl)propyl-1-N,N-diisopropyl phosphoramidite, following the referenced
method for synthesis and deprotection.7 Each batch of CPG-bound crude 2’-O-methyl
oligonucleotide was base-deprotected and cleaved from the solid support by suspension
in concentrated ammonia for 2 days. The 2’-O-TBS oligoribonucleotide was deprotected
and cleaved using 40% aqueous methylamine at 65 °C for 10 min. The CPG was removed by filtration, and the filtrate concentrated on a speed vac to remove most of the
181
ammonia, and lyophilized. The 2’-O-TBS oligoribonucleotides were desilylated using
Et3N-HF in N-methylpyrrolidinone at 65 °C, then had the excess fluoride scavenged
using trimethylsilyl-isopropyl ether, and finally the oligomer was precipitated in ether.
Semi-preparative reverse phase HPLC purification was performed on the crude DMT-on oligonucleotides on a Waters Novapak C18 19 x 300 mm column using gradients of
acetonitrile in 0.1M triethylammonium acetate (TEAA) buffer (pH 6.8). The product fractions were lyophilized. The DMT group was removed by dissolving the residue in 10 mL of 0.6 M acetic acid, monitored by analytical HPLC. After ~5 hours, the mixture was
neutralized with ammonia and then lyophilized. Deprotection of the 5’ terminal
phosphate group was achieved by dissolving the residue in 15 mL of concentrated
ammonia for 2 hours for the 2’-O-methyl oligomer and 5 mM aqueous ammonia for 2
days for the 2’-OH oligomer. The sample was concentrated on a speed vac and then
lyophilized. Purification of the DMT-off oligonucleotides was performed using a solvent
system of acetonitrile and TEAA buffer (see above). Desalting of pure samples was
performed on a Waters Novapak C18 19 x 300 mm column using gradients of acetonitrile
in degassed Millipore water to yield the triethylammonium form of the oligonucleotide.
The starting oligonucleotide (TEA+ form) was dissolved in water, transferred to a
small epindorf tube (2 mL) or small glass flask (5 mL) and lyophilized. A micro stir bar
was added, then the material was dried in a vacuum dessicator over fresh P2O5 overnight.
The dessicator was opened under nitrogen, and then DMF or DMSO was added with stirring. The flask was then heated at 75 °C for 2 hours in an oil bath, then cooled to the reaction temperature in an oil bath under nitrogen for 5 minutes prior to the addition of
182
the capping reagent 104. After the indicated reaction time, the reaction mixture was diluted with 5 mL of 0.1 M triethylammonium acetate (TEAA) buffer (pH 6.8), extracted with three 4 mL portions of ethyl acetate, and then the aqueous layer was lyophilized.
Semi-preparative reverse phase HPLC purification of the crude DMT-on-capped RNA was performed on a Waters Novapak C18 19 x 300 mm column using gradients of acetonitrile in 0.1M NH4HCO3 buffer (pH 7.3). Detritylation was performed by
dissolving the pure DMT-on capped RNA fraction in 10 mL of water for 6 hours, then
lyophilizing the sample. Purification of the DMT-off capped RNA was performed on a
Waters Novapak C18 7.8 x 300 mm column using gradients of acetonitrile in 0.1M
NH4HCO3 buffer (pH 7.3).
All analytical reverse phase HPLC was performed on a Waters 2960 system, with
an Atlantis C18 column, 100 Ǻ, 4.6 mm x 50 mm, 3μm using gradients of acetonitrile in
0.1M TEAA (pH 6.8) with a flow rate of 1.0 mL/min. ESI-MS was acquired using a
Waters Micromass single quadrupole LCZ system. Optical density of purified samples
was performed on a Cary Varian spectrometer at 260 nm at 25 °C.
7 Preparation of m Gppp-(2’-O-Me-GAUGC)2 (113e) To 20 ODs (0.203 umol, ε = 98.7
OD/umol) of the lyophilized p(GAUGCGAUGC) (TEA+ form) was added 0.3 mL of
DMF and a microstir bar. Following the above procedure, 0.0241 g of e was added and
reacted at 45 °C for 7 days prior to workup. DMT-on purification, detritylation and
DMT-off purification yielded 3.1 ODs of 113e (0.029 umol, 15%, ε = 107.2 OD/umol),
183
and 8.9 ODs of recovered starting material p10mer (0.0903 umol). UV λmax = 258 nm;
The mass was confirmed via ESI-LCMS in negative mode as m/z (M-3) 1285.9
(calculated 1285.1)
7 Preparation of m Gppp-(2’-O-Me-GUAUC)4 (113f) To 50 ODs (0.245 umol, ε = 204
+ OD/umol), of the lyophilized p(GUAUC)4 (TEA form) was added 0.4 mL of DMSO and
a microstir bar. Following the above procedure, 0.0292 g (24.51 umol, 100 eq) of e was
initially added at 45 °C, and then was added every 2 days for a total of 3 additions (73.5
umol, 300 eq total). After 7 days, the reaction was worked up, the reaction mixture was
precipitated into 20 mL of ether, and the supernatant was decanted. The pellet was
washed with 3 mL of ether, then dried under a stream of nitrogen. This was followed by
DMT-on purification, detritylation and DMT-off purification to yield 33.1 ODs of 113f (
0.156 umol, 63%, ε = 212.5 OD/umol). UV λmax = 259 nm; The mass was confirmed via
ESI-LCMS in negative mode as m/z (M-4) 1775.9 (calculated 1775.1)
7 Preparation of m Gppp-(GUAUC)4 (113g) To 36 ODs (0.178 umol, ε = 204 OD/umol),
+ of the lyophilized p(GUAUC)4 (TEA form) was added 0.5 mL of DMSO and a microstir bar. Following the above procedure, 0.0212 g (17.84 umol, 100 eq) of e was initially added at 45 °C, and then was added every 2 days for a total of 3 additions (53.52 umol,
300 eq total). After 8 days, the reaction mixture was precipitated into 20 mL of ether, and
the supernatant was decanted. The pellet was washed with 3 mL of ether, then dried
184
under a stream of nitrogen. This was followed by DMT-on purification, detritylation and
DMT-off purification to yield 15.2 ODs of 113g ( 0.0717 umol, 40%, ε = 212.5
OD/umol), and 6.6 ODs of recovered starting material RNA p20mer (0.0325 umol). UV
λmax = 259 nm; The mass was confirmed via ESI-LCMS in negative mode as m/z (M-4)
1706.1 (calculated 1705.1)
185
3.6 References
1. Kamimura, T., Osaki, Y., Sekine, M., Hata, T., "An Effective Method for the Synthesis of the Cap Structure of Eukaryotic Messenger Ribonucleic Acids". Tetrahedron Letters, 1984, 25(25), 2683-2686.
2. Fan, Y., Gaffney, B., Jones, R., "Transient Silyation of the Guanosine O6 and Amino Group Facilitates N-Acylation". Organic Letters, 2004, 6(15), 2555-2557.
3. Stepinski, J., Bretner, M., Jankowska, M., Felczak, K., Stolarski, R., Wieczorek, Z., Cai, A-L., Rhoads, R., Temeriuz, Haber, D., Darzynkiewicz, E., "Synthesis and Properties of P1, P2-, P1, P3- and P1, P4- Dinucleotide Di-, Tri- and Tetraphosphate mRNA 5-Cap Analogues". Nucleosides and Nucleotides, 1995, 14(3-5), 717-721.
4. Sawai, H.W., H., Nakamura-Ozaki, A., "Synthesis and Reactions of Nucleoside 5'-Diphosphate Imidazolide. A Nonenzymatic Capping Agent for 5'- Monophosphorylated Oligoribonucleotides in Aqueous Solution." Journal of Organic Chemistry, 1999, 64, 5836-5840.
5. Jemielity, J., Stepinski, J., Jaremko, M., Haber, D., Stolarski, R., Rhoads, R., Darzynkiewicz, E., "Synthesis of Novel mRNA 5' Cap-Analogues: Dinucleoside P1, P3-Tri, P1, P4-Tetra, and P1, P5-Pentaphosphates". Nucleosides, Nucleotides, and Nucleic Acids, 2003, 22(5-8), 691-694.
6. Stepinski, J., Zuberek, J., Jemielty, J., Kalek, M., Stolarski, R., Darzynkiewicz, E., "Novel Dinucleoside 5',5'-Triphosphate Cap Analogues. Synthesis and Affinity for Murine Translation Factor eIF4E." Nucleosides, Nucleotides, and Nucleic Acids, 2005, 24, 629-633.
7. Worch, R., Niedzwiecka, A., Stepinski, J., Mazza, C., Jankowska-Anyszka, M., Darzynkiewicz, E., Cusack, S., Stolarski, R., "Specificity of Recognition of mRNA 5' Cap by Human Nuclear Cap-Binding Complex". RNA, 2005, 11, 1355- 1363.
8. Kore, A.R., Shanmugasundaram, M., "Synthesis and Biological Evaluation of Trimethyl-Substituted Cap Analogs". Bioorganic & Medicinal Chemistry, 2008, 18, 880-884.
9. Jones, J.W., Robins, R.K., "Purine Nucleosides. III. Methylation Studies of Certain Naturally Occurring Purine Nucleosides." Journal of the American Chemical Society, 1963, 85, 193-201.
10. . Current Protocols in Nucleic Acid Chemistry, ed. Egli, M., Herdewijin, P., Matsuda, A., Sanghvi, Y. 2010: John Wiley & Sons, Inc.
186
11. Stepinski, J., Waddell, C., Stolarski, R., Darzynkiewicz, E., Rhoads, R., "Synthesis and Properties of mRNA Containing the Novel "Anti-Reverse" Cap Analogs 7-Methyl(3'-O-methyl)GpppG and 7-Methyl(3'-deoxy)GpppG". RNA, 2001, 7, 1486-1495.
12. Yoshikawa, M., Kato, T., Takenishi, T.,, "A Novel Method For Phosphorylation of Nucleosides to 5'-Nucleotides". Tetrahedron Letters, 1967, 50, 5065-5068.
13. Moran, J., Whitesides, G., "A Practical Enzymatic Synthesis of (Sp)-Adenosine 5'-O-(1-Thiophosphate) ((Sp)-ATP-a-S)". Journal of Organic Chemistry, 1984, 49, 704-706.
14. Guzaev, A., Salo, H., Azhayev, A, Lonnberg, H., "A New Approach for Chemical Phosphorylation of Oligonucleotides at the 5'-Terminus". Tetrahedron, 1995, 51(34), 9357-9384.
15. Dai, Q., Saikia, M., Li, N., Pan, T., Piccirilli, J.A., "Efficient Chemical Synthesis of AppDNA by Adenylation of Immobilized DNA-5'-Monophosphate". Organic Letters, 2009, 11(5), 1067-1070.
187
3.7 Appendix
188
m7GpppG, 113a
1H NMR
13C NMR
31P NMR
189
HPLC 2-20% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
0.50
0.40 0.30
AU 0.20 0.10
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 Minutes
LRMS
400.3 100
%
801.4
266.6 534.1
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200
UV
190
7 m Gppp(s)G, D1, 113b
1H NMR
13C NMR
31P NMR
191
HPLC 2-20% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
0.60
0.40
AU 0.20
0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 Minutes
LRMS
408.0 100
%
271.7 544.4 817.1
0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200
UV
192
7 m Gppp(s)G, D2, 113b
1H NMR
13C NMR
31P NMR
193
HPLC 2-20% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
1.00
0.80
0.60
AU 0.40 0.20 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00 12.00 Minutes
LRMS
408.0 100
%
271.7 816.9 544.4 0 m/z 200 300 400 500 600 700 800 900 1000 1100 1200
UV
194
7 m Gppp(2’-O-methyl-GAUGC)2, 113e
HPLC 2-40% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
0.80
0.60
AU 0.40
0.20
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes LRMS 1285.9 100
%
0 m/z 700 800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900
UV
195
7 m Gppp(2’-O-methyl-GUAUC)4, 113f
HPLC 2-40% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
0.80
0.60
AU 0.40
0.20 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes
LRMS
1775.9 100
%
0 m/z 600 800 1000 1200 1400 1600 1800 2000 2200 2400
UV
196
7 m Gppp(GUAUC)4, 113g
HPLC 2-40% acetonitrile in 0.1 M triethylammonium acetate buffer (pH 6.8), 280 nm
1.00
AU 0.50
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 Minutes LRMS
1706.1 100
%
0 m/z 400 600 800 1000 1200 1400 1600 1800 2000 2200 2400
UV
197
ELIZABETH VELIATH-HOUSTON
EDUCATION:
RUTGERS UNIVERSITY, Piscataway, New Jersey 2005-2010 PhD in Organic Chemistry, January 2011
Awards: GAANN Fellowship 2009-2010, Excellence in Leadership Award 2009, Rutgers Excellence Fellowship 2006-2007
MARYMOUNT at FORDHAM UNIVERSITY, Tarrytown, New York 1998-2001 BS, May 2001, Summa Cum Laude Majors: Chemistry, Foods and Nutrition (double major); GPA 4.0/4.0
Honors: President’s Award for Academic Excellence, Department Gold Medal for Nutrition, Department Gold Medal for Chemistry, Marymount Trustee Scholarship, Dean’s List
RELEVENT EXPERIENCE:
MERCK, Rahway, NJ 2010- Associate Chemist, Department of RNA Process Chemistry
RUTGERS UNIVERSITY, Piscataway, NJ 2005-2010 Graduate Student, Department of Chemistry and Chemical Biology
KRAFTFOODS, Technical Center, Tarrytown, New York 2002-2005 Associate Research Scientist II, Coffee Division, Global Technology & Quality, Specialty Coffee Development Associate Research Scientist I, Technical Guidance Research 2001-2002
AMERICAN HEALTH FOUNDATION, Valhalla, New York 1999 Researcher Department of Biochemical Pharmacology
PUBLICATIONS:
Gaffney, B.A., Veliath, E., Zhao, J., Jones, R.A. (2010) “One-Flask Syntheses of c-di- GMP and the [Rp,Rp] and [Rp,Sp] Dithiophosphate Analogs.” Organic Letters 12(14): 3269-3271
Zhang, Z., Dai, J., Veliath, E., Jones, R.A., Yang, D. (2010). "Structure of a Two-G- Tetrad Intramolecular G-Quadruplex Formed by a Variant Human Telomeric Sequence in K+ Solution: Insights into the Interconversion of Human Telomeric G-Quadruplex Structures." Nucleic Acids Research 38(3): 1009-1021
Zhao, J., Veliath, E., Kim, S., Gaffney, B., Jones, R.A. (2009). "Thiophosphate Analogs of c-di-GMP: Impact on Polymorphism." Nucleosides, Nucleotides, and Nucleic Acids 28(5-7): 352-378
Weisburger, J. H., Veliath, E., Larios, E., Pittman, B., Zang, E., Hara, Y. (2002). "Tea Polyphenols Inhibit the Formation of Mutagens During the Cooking of Meat." Mutation Research, Genetic Toxicology and Environmental Mutagenesis 516(1-2): 19-22
198
TEACHING:
RUTGERS UNIVERSITY, Piscataway, New Jersey 2005-2008 Teaching Assistant, Department of Chemistry and Chemical Biology Chem 310: second semester Organic Chemistry lab for chemistry majors Chem 309: first semester Organic Chemistry lab for chemistry majors Chem 171: General Chemistry lab requirement