NMR studies of the structure, kinetics and interactions of the ...

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NMR studies of the structure, kinetics and

interactions of the conserved RNA motifs

in the FMDV IRES

A thesis submitted to the University of Manchester

for the degree of PhD in the Faculty of Engineering

and Physical Sciences

2012

Usman Rasul

School of Chemistry

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Table of Contents

Title Page 1

Table of Contents 2

List of Figures 7

List of Tables 25

Symbols and Abbreviations 28

Abstract 32

Declaration 33

Copyright Statement 33

Acknowledgements 34

Chapter 1: Introduction 35

1.1 Significance of the project 35

1.1.1 The role of IRES in picornavirus translation 35

1.1.2 The IRES and antiviral therapy 37

1.1.3 The IRES in biotechnology 39

1.1.4 RNA structural biology 41

1.2 Picornaviruses and translation 43

1.2.1 Picornavirus classification and genome 43

1.2.2 mRNA and cap-dependent translation 44

1.2.3 FMDV and cap-independent translation 45

1.2.4 Internal Ribosome Entry Site (IRES) 46

1.2.5 FMDV IRES 47

1.3 Nucleic acid chemistry 50

1.3.1 Nucleic acids 50

1.3.2 RNA synthesis 52

1.3.2.1 Chemical synthesis 52

1.3.2.2 Enzymatic synthesis 54

1.3.3 RNA nucleotide structure 56

1.3.4 RNA base pairing and stacking 58

1.3.5 RNA structure 60

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1.4 RNA interactions 63

1.4.1 Intramolecular interactions 63

1.4.2 RNA and Mg2+

63

1.4.3 RNA-RNA interactions 65

1.5 Principles of NMR spectroscopy 67

1.5.1 Basic theory of NMR 67

1.5.2 Chemical shift, coupling constant and linewidth 68

1.5.3 Nuclear relaxation 69

1.5.3.1 Spin-lattice relaxation 70

1.5.3.2 Spin-spin relaxation 70

1.5.4 Nuclear Overhauser Effect (NOE) 72

1.5.5 NMR of rate processes 74

1.5.5.1 Base pair kinetics 74

1.5.5.2 Chemical exchange 75

1.5.6 1D 19

F-NMR and 31

P-NMR 77

1.5.7 Two-dimensional (2D) NMR spectroscopy 78

1.5.8 Three-dimensional (3D) NMR spectroscopy 79

1.6 Principles of molecular modelling 80

1.6.1 Molecular mechanics (MM) 80

1.6.2 Energy minimisation 81

1.6.3 Simulated annealing and molecular dynamics (MD) 82

1.7 Previous work 83

1.8 Aim of the project 84

Chapter 2: Materials and methods 86

2.1 RNA sample preparation for NMR studies 86

2.2 NMR spectroscopy 88

2.2.1 NMR spectrometers 88

2.2.2 NMR experimental parameters 88

2.2.3 Data processing and analysis 89

2.3 NMR techniques 91

2.3.1 Solvent suppression 91

2.3.2 1D NMR experiments with decoupling 93

2.3.3 Variable temperature (VT) experiments 94

2.3.4 T1 measurements 94

2.3.5 Water magnetisation transfer experiments 95

2.3.6 2D Double-Quantum Filtered Correlation Spectroscopy

(DQF-COSY) 96

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2.3.7 2D Total Correlation Spectroscopy (TOCSY) 97

2.3.8 2D Heteronuclear Single Quantum Coherence (HSQC) 98

2.3.9 2D Nuclear Overhauser Effect Spectroscopy (NOESY) 99

2.3.10 2D Heteronuclear Overhauser Effect Spectroscopy

(HOESY) 100

2.3.11 2D CPMG-HSQC-NOESY 101

2.3.12 3D NOESY/2Q-COSY 102

2.4 NMR assignment of RNA 103

2.4.1 Assignment strategy 103

2.4.2 Identification of base protons 106

2.4.2.1 Identification of exchangeable protons 106

2.4.2.2 Identification of non-exchangeable protons 107

2.4.3 Identification of sugar protons 108

2.4.4 Sequence-specific resonance assignment 109

2.5 Structure determination protocol of RNA 111

2.5.1 Restraints 111

2.5.1.1 Distance restraints 111

2.5.1.2 Dihedral angle restraints 112

2.5.1.3 Hydrogen bonds and planarity restraints 113

2.5.2 Structure calculation 114

2.5.3 Conformational analysis 117

2.5.4 Structure validation 120

2.6 Quantitative measurement of exchange rate constants 121

Chapter 3: NMR studies of the FMDV 16mer RNA and the effect

of Mg2+

123

3.1 Structure determination of the 16mer apo-RNA 123

3.1.1 NMR assignment 123

3.1.1.1 Exchangeable proton assignment 123

3.1.1.2 Non-exchangeable proton assignment 128


3.1.3 NMR solution structure 144

3.1.3.1 Ensemble and final structure 144

3.1.3.2 GNRA tetraloop 145

3.1.3.3 Intramolecular interactions 146


3.2 Effect of Mg2+

on 16mer RNA chemical shifts 152

3.2.1 Changes in proton chemical shift 152

3.2.2 Changes in phosphorus chemical shift 155

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3.3 Effect of Mg2+

on 16mer RNA stability 157

3.3.1 1H-NMR variable temperature (VT) series 157

3.3.2 31P-NMR variable temperature (VT) series 161

3.4 Imino proton exchange in the 16mer RNA 162

3.4.1 NOE exchange 162

3.4.2 T1 of imino protons 164

3.4.3 Exchange rate of imino protons 165

3.4.4 Effect of Mg2+

on imino proton exchange rates 168

3.5 Structure determination of the 16mer Mg2+

RNA complex 170







3.5.3.2 GNRA tetraloop 188



3.5.5 Comparison of the 16mer apo/Mg2+

RNA NMR structures 195

3.6 Mg2+

-induced structural changes to the 16mer apo-RNA 196

Chapter 4: : NMR studies of the FMDV 15mer RNA and its

complex with the 16mer RNA 201

4.1 Structure determination of the 15mer apo-RNA 201







4.1.3.2 Heptaloop 216



4.1.5 Comparison of the 15mer and 16mer apo-RNA NMR

structures 222

4.2 Effect of Mg2+

on the 15mer RNA 223

4.3 1GHz NMR studies of the 16mer apo-RNA 227

4.3.1 Effect of magnetic field strength on the 16mer apo-RNA 227

4.3.2 Sensitivity enhancement with 1GHz 229

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4.4 RNA-RNA interaction 230

4.4.1 Analysis of the RNA-RNA complex in 1H2O 230

4.4.2 Analysis of the RNA-RNA complex in 2H2O 234

4.4.3 Model of the RNA-RNA interaction 238

Chapter 5: 19

F-NMR studies of selectively fluorinated RNAs 240

5.1 19F-NMR studies of the 5-FU 16mer and 15mer RNAs 240

5.1.1 Identification of fluorination 240

5.1.2 Effect of the 19

F nucleus on the 5-FU 16mer RNA 244



5.1.2.3 The 2D 1H-

19F HOESY experiment 247

5.1.2.4 Effect of the 19

F nucleus on 31

P chemical shifts 248

5.1.3 Effect of magnetic field strength on the 5-FU 16mer RNA 250


F nucleus on the 5-FU 15mer RNA 251


on the 19

F signal 253

5.2 19F-NMR studies of the 5-FU 16mer/15mer complex 255

Chapter 6: Conclusion and Future work 257

6.1 Conclusion 257

6.1.1 Structure of the conserved RNA motifs 257

6.1.2 The role of Mg2+

259

6.1.3 RNA-RNA interaction 260

6.1.4 Errors and their implications 261

6.2 Future work 262

6.2.1 Binding of Mg2+

ions to RNA 262

6.2.2 Isotopically labelled RNA 263

6.2.3 RNA tertiary contacts 264

References 266

Appendices 277

Appendix I: NMRPipe script 277

Appendix II: XPLOR scripts 278

Word count: 63,403

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List of Figures

Figure 1.1.1 An IRES bicistronic expression vector. CAT and LUC are the first and

second reporter genes, respectively, surrounding the FMDV IRES. The arrow depicts the

direction of translation...………………….........................................................................40

Figure 1.2.1 The genomic structure of the FMDV viral RNA. The coding region or ORF

is divided into the L-region and three distinctive regions of P1, P2 and P3, coding for the

capsid and non-structural proteins. The 5’-UTR precedes the coding region and the 3’-

UTR is situated after the coding region…………………………………………………..43

Figure 1.2.2 The normal cap-dependent translation. The mRNA requires the 5’-end cap

(7-methylguanosine) structure along with complex interaction between the 40S ribosome

and several eukaryotic initiation factors (eIFs) to allow for translation initiation………..44

Figure 1.2.3 The IRES mediated cap-independent translation. Translation initiation does

not require the 5’-end cap structure and eIF4E in picornaviruses. Instead, the IRES

provides an entry/binding site for the 40S ribosome, so the ribosomal scanning can

start…………......................................................................................................................45

Figure 1.2.4 The FMDV IRES structure separated into five domains (1-5), from residues

1 (5’-C) to 462 (U-3’). Domain 3 is the central domain, from residues 86 (5’-G) to 299 (C-

3’). The apical region of domain 3 is found within the orange circle…………………….47

Figure 1.2.5 Illustration of the hammerhead region, a 79mer RNA (G150-U228) found in

domain 3 of the FMDV IRES. The 16mer RNA (U172-A187) shown in the red box and

36mer RNA (C159-G194) in the blue box, are displayed………………………………..48

Figure 1.2.6 Illustration of the apical region of domain 3 in the FMDV IRES, located

below the hammerhead region. The 15mer RNA (G229-C243) is indicated by the area

inside the red box…………………………………………………………………………49

Figure 1.3.1 Structures of purine and pyrimidine bases, and sugars, in DNA and RNA.

Atoms are numbered according to the IUPAC convention.35

……………………………50

Figure 1.3.2 A phosphodiester bond linking two nucleosides, adenosine and guanosine,

from the 5’ to 3’-end……………………………………………………………………...51

Figure 1.3.3 A phosphoramidite building block with four different sites required for

protection…………………………………………………………………………………53

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Figure 1.3.4 Seven dihedral angles, α, β, γ, δ, ε, ζ, χ, revealing the conformation of a

nucleotide and five dihedral angles, ν0, ν1, ν2, ν3, ν4, defining the conformation of the five-

membered sugar ring………………………………………………………………...........56

Figure 1.3.5 The two main sugar conformations (a) C3’-endo in RNA and (b) C2’-endo in

DNA………………………………………………………………………………………57

Figure 1.3.6 (a) the relationship between the syn/anti notations and the corresponding

dihedral angle values. (b) The pseudorotation phase cycle of the pentose sugar showing

the relationship between the pseudorotation phase angle (P), and the endo and exo

notations.35

………………………………………………………………………………...57

Figure 1.3.7 Illustration of canonical (a) G.C (b) A.U, and non-canonical (c) G.U, base

pairing…………………………………………………………………………………......58

Figure 1.3.8 Illustration of the base stacking interactions between adjacent bases in a base

paired helical stem. The rectangles represent the G, C, A, U bases, the unfilled circles

correspond to the pentose sugar, red circles represent the phosphorus atoms and green

triangles symbolise the base stacking interactions………………………………………..59

Figure 1.3.9 Illustration of the secondary structure of tRNA. The four-way junction

consists of the acceptor arm and the three hairpin loops, the D-loop, the T-loop and the

anticodon loop…………………………………………………………………………….61

Figure 1.3.10 G.A sheared base pairing found in GNRA tetraloop motifs, between the

first and fourth base. Two hydrogen bonds are formed between (guanine 2-amino and

adenine N7) and between (guanine N3 and adenine 6-amino); shown as blue dashed

lines……………………………………………………………………………………….62

Figure 1.4.1 Potential Mg2+

-ion interaction with the RNA phosphate backbone. The

diagram illustrates the Mg2+

ions (green circles), water molecules (red circles with

attached blue circles) and phosphate oxygens (lone red circles). Ion interactions can

involve specific interactions with RNA whereby the Mg2+

ions act as (a) chelated ions, (b)

water-positioned ions and (c) diffuse ions………………………………………………..64

Figure 1.4.2 Two main types of A-minor motifs (left) Type I and (right) Type II. The blue

broken lines represent hydrogen bonding within C.G base pairing and red broken lines

represent hydrogen bonding in A-minor interactions…………………………………….66

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Figure 1.5.1 Scheme illustrating (a) Spin-lattice (T1) relaxation where magnetisation

relaxes back to longitudinal axis; (b) Spin-spin (T2) relaxation where the spins precess in

the x,y plane and fan out as they lose coherence………………………………………….69

Figure 1.5.2 A plot of T1 (blue curve) and T2 (red curve) as a function of correlation time

(τc). Small molecules have a correlation time in the 10-12

to 10-10

second range while large

molecules are above 10-9

seconds………………………………………………………...71

Figure 1.5.3 Energy level diagram of a two spin system illustrating four energy levels

(N1>N2>N3>N4) with their corresponding α/β spin states. WA and WX represent single

quantum transition in HA and HX, respectively. W0 and W2 represent zero and double

quantum transitions, respectively…………………………………………………………72

Figure 1.5.4 Scheme illustrating the NOE effect in a two spin system; (a) spin populations

before saturation of nucleus HA, (b) spin populations after saturation of nucleus HA……73

Figure 1.5.5 Example of chemical exchange between two conformations of the same

nucleus. The nucleus can change magnetic environments between the two conformations

in fast, intermediate and slow exchange regimes, on the NMR timescale………………..76

Figure 1.5.6 The basic pulse sequence of a 2D NMR experiment. The evolution time (t1)

is the period between the two 90° pulses whereby T1 and T2 relaxation occurs. The

acquisition time (t2) begins immediately after the last 90° pulse…………………………78

Figure 1.5.7 The pulse sequence for a 3D NOESY-TOCSY experiment, constructed from

a combination of the NOESY and TOCSY pulse sequences……………………………..79

Figure 2.3.1 The presaturation pulse sequence. A selective, low power presaturation pulse

saturates the water frequency, which is followed by a non-selective, high power pulse to

excite the desired protons…………………………………………………………………91

Figure 2.3.2 The WATERGATE pulse sequence. The 90° (non-selective) and the 180°

(selective) pulses are shown in the top line. The τ delays inserted for gradient recovery.

The bottom line displays the pulsed magnetic gradients. Most protons experience gradient-

180°-gradient and are refocused, while the water protons experience gradient-0-gradient

and are dephased. Therefore, during signal acquisition (t2) the water signal is

suppressed………………………………………………………………………………...92

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Figure 2.3.3 1D X-{H} decoupled NMR experiment, where X represents an NMR active

nucleus apart from 1H. The decoupling pulse is activated at the same time as the FID is

acquired for nucleus X……………………………………………………………………93

Figure 2.3.4 The inversion recovery pulse sequence. A 180° pulse is followed by a

variable delay (τ) and then a 90° detection pulse…………………………………………94

Figure 2.3.5 The pulse sequence of the water magnetisation transfer experiment. The

DANTE sequence is followed by a variable delay (τm) and three 90° pulses. Gradient

pulses are represented by G1, G2 and G3.71

……………………………………………...95

Figure 2.3.6 The pulse sequence of a DQF-COSY experiment. The first 90° pulse is the

same as in a standard COSY experiment. The second 90° pulse is immediately followed

by a third 90° pulse; the third pulse acts in combination with second pulse as a double-

quantum filter to convert the double-quantum coherence created by the second pulse back

into single-quantum coherence.91

…………………………………………………………96

Figure 2.3.7 The pulse sequence of a TOCSY experiment. The first 90° pulse flips the

spin magnetisation onto the x,y plane. The evolution period is followed by the isotropic

mixing, which transfers magnetisation between spins connected via an unbroken network

of couplings.93

…………………………………………………………………………….97

Figure 2.3.8 1H-

13C HSQC sequence adapted from the INEPT pulse sequence. The first

pulses are derived from INEPT pulse sequence which transfers magnetisation from 1H to

13C nuclei. A 180° pulse on

1H nuclei forms a spin echo, so the evolution of coupling is

refocused. At mid-evolution the 13

C spin magnetisation then evolves during t1, at which

time it acquires a frequency label according to the offset of 13

C. The inverse INEPT step

transfers magnetisation back to 1H from

13C nuclei, yielding enhanced sensitivity of

13C

comparable to that of 1H.

95………………………………………………………………..98

Figure 2.3.9 The pulse sequence of a NOESY experiment. The first 90° pulse flips the

spins onto the x,y plane. The spins precess in the evolution period before a second 90°

pulse flips the spins onto longitudinal axis and a predetermined mixing period allows for

the exchange of magnetisation between dipolar spins. Finally the spins are flipped back

onto the x,y plane for detection.96

………….......................................................................99

Figure 2.3.10 The pulse sequence of the HOESY experiment, whereby X is the observed

nucleus. The X signal is recorded during t2 and the 1H signal is recorded as a function of

t1.98

……………………………………………………………………………………….100

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Figure 2.3.11 The pulse sequence of the 1H-

31P CPMG-HSQC-NOESY experiment. τ

represents the delays times around the 180° refocusing pulses, τm is the mixing time for

the NOESY component. The symbol G corresponds to the gradient pulses shown as black

filled shaped pulses.100

…………………………………………………………………..101

Figure 2.3.12 The 3D NOESY/2Q-COSY pulse sequence. The DANTE (Delays

Alternating with Nutation for Tailored Excitation) presaturation sequence is used to

suppress the residual water signal. The evolution period (t1) is followed by the NOE

mixing time (τm) and then the multiple quantum excitation step (τMQ). The t1 and t2 are the

first and second indirect dimensions, and t3 is the third (direct) dimension…………….102

Figure 2.4.1 Protocol for NMR assignment of RNA. The green boxes indicate the solvent

used. The red boxes contain information on the specific NMR experiments employed and

the cyan boxes represent the assignments that can be obtained from the corresponding

experiment(s)………………………………………………………………………….....103

Figure 2.4.2 Scheme representing intranucleotide H6/H8-H1’ connectivities (blue) and

internucleotide H6/H8-H1’ connectivities (red). These connectivities follow well

established NOE sequential assignment pathways………………………………………110

Figure 2.4.3 Scheme representing intranucleotide H1’-H2’ connectivities (blue) and

internucleotide H2’-H6/H8 connectivities (red)………………………………………...110

Figure 2.5.1 Scheme summarising the procedure for structure determination of RNA. The

restraints are added to the starting structure, which is followed by the simulated annealing

step. The lowest RMSD structure generated from the simulated annealing process is used

for the refinement step. The lowest RMSD structure from the refinement process is then

assessed for acceptance as the final NMR solution structure……………………………116

Figure 2.5.2 Definitions of the different parameters used in the 3DNA program.

Complementary base pair parameters (red), base pair step parameters (blue) and local

helical parameters (green) are clearly outlined. All images illustrate the positive values of

the corresponding parameters. Helical twist (Ω) is the same as twist (ω) and helical rise (h)

is the same as rise (Dz).107

…………………………………………………………..118-119

Figure 3.1.1 700 MHz 1H-NMR spectrum of the FMDV 16mer apo-RNA, at 2°C in

1H2O,

displaying the imino region. Six peaks were identified corresponding to the imino protons

of U175, U176, G185, G186, G177 and G178. The G178 loop imino proton peak can be

clearly observed highfield of the stem imino proton peaks……………………………..124

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Figure 3.1.2 700 MHz NOESY (250ms) spectrum of the FMDV 16mer apo-RNA, at 2°C

in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks

correspond to imino-imino connectivities. The sequential assignment starts from G178,

labelled in red, and finishes at G186, labelled in blue. Inset: Secondary structure of the

16mer RNA highlighting the imino-imino connectivities observed, represented by light

blue oval shapes…………………………………………………………………………126


in 1H2O, illustrating the imino-amino region of the spectrum. Connectivities from imino

protons to NH2*/NH2/H2/H5/H1’ protons can be observed (NH2* corresponds to the

proton involved in base pair hydrogen bonding); connectivities are marked by a black

circle……………………………………………………………………………………..127

Figure 3.1.4 Illustration of the identification of C5-H5 and C6-H6 peaks in the 1H-

13C

HSQC spectrum and the subsequent assignment of H5-H6 cross peaks in the NOESY

spectrum, of the FMDV 16mer apo-RNA. Bottom left panel: 600 MHz NOESY (400ms)

spectrum, at 25°C in 2H2O; blue circles indicate H5-H6 cross peaks. Top left panel: 600

MHz 1H-

13C HSQC spectrum at 25°C in

2H2O, displaying C6-H6 peaks. Bottom right

panel: 600 MHz 1H-


2H2O, displaying C5-H5 peaks…..129

Figure 3.1.5 Top/Bottom panels: 600 MHz NOESY (400ms) spectrum of the FMDV

16mer apo-RNA, at 25°C in 2H2O. The blue line represents C173 H6–H1’ to U175 H6–H1’

intra- and internucleotide connectivities, and the green line represents A180 H8-H1’ to

A183 H8-H1’ intra- and internucleotide connectivities; same colouring as in secondary

structure shown. The (i) corresponds to an intranucleotide connectivity. The red circles

correspond to H5-H6 connectivities……………………………………………………..131

Figure 3.1.6 Illustration of the identification of C6-H6, C8-H8 and C1’-H1’ peaks in the 1H-

13C HSQC spectrum and the subsequent assignment of H6/H8-H1’ cross peaks in the

NOESY spectrum, of the FMDV 16mer apo-RNA. (a) 600 MHz 1H-

13C HSQC spectrum

at 25°C in 2H2O, displaying the C6-H6 and C8-H8 peaks. (b,c) 600 MHz NOESY (400ms)

spectrum at 25°C in 2H2O. (d,e) 600 MHz

1H-


2H2O,

displaying the C1’-H1’ peaks……………………………………………………………132

Figure 3.1.7 Illustration of the assignment of intranucleotide H1’-H2’ and internucleotide

H2’-H6/H8 connectivities in the NOESY spectrum, with the aid of the 1H-

13C HSQC

spectrum, of the FMDV 16mer apo-RNA. Bottom panels: 600 MHz 1H-

13C HSQC spectra

at 25°C in 2H2O. Top panels: 600 MHz NOESY (400ms) spectra at 25°C in

2H2O…….133

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Figure 3.1.8 Illustration of the assignment of intranucleotide H1’-H2’ connectivities in the

DQF-COSY spectrum, with the aid of the 1H-

13C HSQC spectrum, of the FMDV 16mer

apo-RNA. Top right panel: 600 MHz 1H-


2H2O,

representing the C1’-H1’ resonances. Top left panel: 400 MHz DQF-COSY spectrum at

25°C in 2H2O, representing the H1’-H2’ cross peaks. Bottom left panel: 600 MHz

1H-

13C

HSQC spectrum at 25°C in 2H2O, representing the C2’-H2’ resonances……………….135

Figure 3.1.9 Illustration of the assignment of phosphorus to H6/H8/H1’ peaks in the 1H-

31P CPMG-HSQC-NOESY spectrum, with the aid of the

1H-

13C HSQC spectrum, of the

FMDV 16mer apo-RNA. Bottom left panel: 600 MHz 1H-

31P CPMG-HSQC-NOESY

(500ms) spectrum at 25°C in 2H2O, representing the phosphorus-H6/H8 peaks. Bottom

right panel: 600 MHz 1H-

31P CPMG-HSQC-NOESY (500ms) spectrum at 25°C in

2H2O,

representing the phosphorus-H1’ peaks. Top left panel: 600 MHz 1H-

13C HSQC spectrum

at 25°C in 2H2O, representing C6-H6 and C8-H8 peaks. Top right panel: 600 MHz

1H-

13C

HSQC spectrum at 25°C in 2H2O, representing C1’-H1’ peaks………………………...137

Figure 3.1.10 600 MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer

apo-RNA. Identification of base and sugar protons of the (a) U172 nucleotide and (b)

C173 nucleotide, illustrated by the F3/F1 NOE plane. Both positive (orange) and negative

(green) levels are shown…………………………………………………………………139

Figure 3.1.11 600 MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer

apo-RNA. Slices from the F3/F2 plane chosen at different F1 frequencies, identifying base

and sugar proton chemical shift of the C174 nucleotide. Only positive levels are shown. In

each slice strong inner NOE peaks correspond to coupling between two protons as well as

weaker outer NOEs to other base/sugar protons………………………………………...140

Figure 3.1.12 Illustration of the NMR solution structures of the FMDV 16mer apo-RNA

(a) Overlay of the 20 lowest RMSD structures, with an average RMSD of 0.18Å. (b)

Lowest RMSD solution structure (0.17Å); the red ribbon represents the RNA

backbone…………………………………………………………………………………144

Figure 3.1.13 The G178UAA181 tetraloop of the FMDV 16mer apo-RNA NMR structure,

shown in Figure 3.1.12b; the closing G177.C182 base pair is also illustrated. Colour coding

of nucleotides: guanosine (blue), uridine (cyan), adenosine (green) and cytosine

(red)……………………………………………………………………………………...145

Figure 3.1.14 The G.A sheared base pair in the FMDV 16mer apo-RNA NMR structure.

Hydrogen bonding distances between G178 NH2-A181 N7 (3.09Å) and G178 N3-A181

NH6 (5.05Å) are indicated by the red broken lines……………………………………..146

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Figure 3.2.1 A stack plot of 400 MHz 1H-NMR spectra (imino region) of the FMDV

16mer RNA in 1H2O, with increasing Mg

2+ concentration. Each

1H-NMR spectrum is

labelled 1-6; 1 (0eq - 5°C), 2 (0.5eq - 5°C), 3 (0.5eq - 2°C), 4 (1.0eq - 2°C), 5 (2.0eq -

2°C), 6 (5.0eq - 2°C). The U175, U176, G185, G186, G177 and G178 imino proton peaks

are labelled accordingly…………………………………………………………………153

Figure 3.2.2 700 MHz 1H-NMR spectra (imino region) of the FMDV 16mer RNA, in

1H2O at 2°C; (a) no Mg

2+ and (b) containing 5eq of Mg

2+. The large Mg

2+-induced

chemical shift change of 0.30ppm and 0.11ppm to G178 and G177, respectively, is clearly

identified…………………………………………………………………………………154

Figure 3.2.3 Histogram illustrating the imino proton chemical shift changes observed in 1H-NMR spectra, at different Mg

2+ concentrations; 0.5eq (red), 1.0eq (blue), 2.0eq (green)

and 5.0eq (orange)……………………………………………………………………….154

Figure 3.2.4 162 MHz stack plot of 31

P-NMR spectra of the FMDV 16mer RNA in 1H2O,

with increasing Mg2+

concentration. Each 31

P-NMR spectrum is labelled a-f; a (0eq - 5°C),

b (0.5eq - 5°C), c (0.5eq - 2°C), d (1.0eq - 2°C), e (2.0eq - 2°C), f (5.0eq - 2°C). Peaks are

labelled 1-9………………………………………………………………………………156

Figure 3.2.5 Histogram illustrating the phosphorus chemical shift changes observed in 31

P-NMR, at different Mg2+

concentrations; 0.5eq(red), 1.0eq (blue), 2.0eq (green), 5.0eq

(orange). The peak numbers 1-9 correspond to the labelling introduced in Figure 3.2.4.

Positive chemical shift changes represent a lowfield shift and negative chemical shift

changes represent a highfield shift………………………………………………………156

Figure 3.3.1 400 MHz 1H-NMR spectra of the FMDV 16mer RNA in

1H2O at two

different temperatures of 5°C and 35°C. (a) 16mer apo-RNA and (b) 16mer Mg2+

RNA

complex. The G186 and G177 imino peaks are clearly exchange retarded in the presence

of Mg2+

. The G178 loop imino proton peak could not be observed in the absence or

presence of Mg2+

………………………………………………………………………...158

Figure 3.3.2 700 MHz 1H-NMR spectra of the FMDV 16mer Mg

2+ RNA complex in

1H2O. A stack plot of the imino region is shown at variable temperatures; 2°C, 10°C, 15°C,

20°C, 25°C, 35°C, 38°C, 41°C, 45°C, 47°C, 50°C and 55°C. The U175, U176, G185,

G186, G177 and G178 imino proton peaks are labelled………………………………...160

15

Figure 3.4.1 400 MHz NOESY (150ms) spectrum of the FMDV 16mer apo-RNA at 5°C

in 1H2O (blue) and Mg

2+ RNA complex at 2°C in

1H2O (red). The imino-imino regions

(bottom panels) and imino-water regions (top panels) are displayed. The horizontal line in

the top panels represents the chemical shift of water; 4.995ppm at 5°C and 5.029ppm at

2°C……………………………………………………………………………………….162

Figure 3.4.2 700 MHz NOESY (150ms) spectrum of the FMDV 16mer Mg2+

RNA

complex in 1H2O at 2°C (red) and 27°C (orange). The imino-imino regions (bottom panels)

and imino-water regions (top panels) are displayed. The horizontal line in the top panels

represents the chemical shift of water; 5.029ppm at 2°C and 4.75ppm at 27°C………...163

Figure 3.4.3 T1 measurement of the U175, U176, G185, G186, G177 and G178 imino

protons for the FMDV 16mer apo-RNA, at 2°C (blue), 15°C (red) and 35°C (green).

Errors bars correspond to 5% of each T1 value………………………………………….165

Figure 3.4.4 Exchange rate constants (Kex) of the imino protons for the FMDV 16mer

apo-RNA at 2°C (blue), 15°C (red) and 35°C (green). Errors bars correspond to 10% of

each Kex value……………………………………………………………………………167

Figure 3.4.5 600 MHz 1H-NMR spectra of the FMDV 16mer apo-RNA in

1H2O at three

different temperatures of (a) 5°C, (b) 15°C and (c) 35°C. The U175, U176, G185, G186,

G177 and G178 imino proton peaks are labelled accordingly…………………………..167

Figure 3.4.6 Exchange rate constants (Kex) of the imino protons for the FMDV 16mer

Mg2+

RNA complex at 15°C (red) and 35°C (green). Errors bars correspond to 10% of

each Kex value……………………………………………………………………………169


RNA

complex, at 2°C in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal

peaks correspond to imino-imino connectivities. The sequential assignment starts from

U176, labelled in red, and finishes at G186, labelled in blue. Inset: Secondary structure of

the 16mer RNA highlighting the imino-imino connectivities observed, represented by light

blue oval shapes…………………………………………………………………………171


RNA

complex, at 2°C in 1H2O, illustrating the imino-amino region of the spectrum.

Connectivities from imino protons to NH2*/NH2/H2/H5/H1’ protons can be observed

(NH2* corresponds to the proton involved in base pair hydrogen bonding); connectivities

are marked by a black circle……………………………………………………………..172

16

Figure 3.5.3 Top and bottom panels: 600 MHz NOESY (400ms) spectrum of the FMDV

16mer Mg2+

RNA complex, at 25°C in 2H2O. The blue line represents U172 H6-H1’ to

G178 H1’-U179 H6 intra- and internucleotide connectivities, and the green line represents

A180 H8-H1’ to A187 H8-H1’ intra- and internucleotide connectivities; same colouring as

in secondary structure shown. The (i) corresponds to an intranucleotide connectivity and (s)

corresponds to a sequential connectivity. The red circles correspond to H5-H6

connectivities…………………………………………………………………………….174

Figure 3.5.4 Illustration of the identification of C6-H6, C8-H8 and C1’-H1’ peaks in the 1H-

13C HSQC spectrum and the subsequent assignment of H6/H8-H1’ cross peaks in the

NOESY spectrum, of the FMDV 16mer Mg2+

RNA complex. (a) and (d) 600 MHz 1H-

13C

HSQC spectrum at 25°C in 2H2O, displaying C6-H6 and C8-H8 peaks. (b), (c) and (e) 600

MHz NOESY (250ms) spectrum, at 25°C in 2H2O. (f) and (g) 600 MHz

1H-

13C HSQC

spectrum at 25°C in 2H2O, displaying C1’-H1’ peaks…………………………………..175



13C HSQC

spectrum, of the FMDV 16mer Mg2+

RNA complex. Bottom panels: 600 MHz 1H-

13C

HSQC spectra at 25°C in 2H2O. Top panels: 600 MHz NOESY (250ms) spectra at 25°C in

2H2O……………………………………………………………………………………..176




Mg2+

RNA complex. Top right panel: 600 MHz 1H-


2H2O,

representing the C1’-H1’ peaks. Top left panel: 600 MHz DQF-COSY spectrum at 25°C

in 2H2O, representing the H1’-H2’ cross peaks. Bottom left panel: 600 MHz

1H-

13C HSQC

spectrum at 25°C in 2H2O, representing the C2’-H2’ peaks…………………………….178




Mg2+

RNA complex. Top panel: 600 MHz DQF-COSY spectrum at 25°C in 2H2O,

representing the H3’-H4’ cross peaks. Middle panel: 600 MHz 1H-

13C HSQC spectrum at

25°C in 2H2O, representing the C3’-H3’ peaks. Bottom panel: 600 MHz

1H-

13C HSQC

spectrum at 25°C in 2H2O, representing the C4’-H4’ peaks…………………………….179

Figure 3.5.8 Illustration of the assignment of intranucleotide H5’-H5’’ connectivities in

the DQF-COSY spectrum, with the aid of the 1H-

13C HSQC spectrum, of the FMDV

16mer Mg2+

RNA complex. Top panel: 600 MHz DQF-COSY spectrum at 25°C in 2H2O,

representing the H5’-H5’’ cross peaks. Bottom panel: 600 MHz 1H-

13C HSQC spectrum at

25°C in 2H2O, representing the C5’-H5’ and C5’-H5’’ peaks…………………………..180

17

Figure 3.5.9 Illustration of the assignment of phosphorus to H6/H8/H1’ peaks in the 1H-

31P CPMG-HSQC-NOESY spectrum, with the aid of the

1H-


FMDV 16mer Mg2+

RNA complex. Top panels: 600 MHz 1H-

13C HSQC spectra at 25°C

in 2H2O. Bottom panels: 600 MHz

1H-

31P CPMG-HSQC-NOESY (500ms) spectra at 25°C

in 2H2O…………………………………………………………………………………..181

Figure 3.5.10 (a) 600 MHz 3D-NOESY/2Q-COSY (250ms) spectrum of the FMDV

16mer Mg2+

RNA complex. Identification of base and sugar protons of the U176

nucleotide. This is illustrated by the F3/F1 NOE plane at F2 = 9.05ppm. Both positive

(orange) and negative (green) levels are shown. The strong intensity of U176 H5 and

U176 H6 inner NOE cross peaks are associated with weaker outer NOE peaks to U175 H6,

U176 H3’, U175 H4’, U175 H2’ and U176 H5’’. (b) 600 MHz 3D-NOESY/2Q-COSY

(250ms) spectrum of the FMDV 16mer Mg2+

RNA complex. Slices from the F3/F2 plane

chosen at different F1 frequencies, identifying H2’(i)-H6/H8(i+1) internucleotide

connectivities. A sequential assignment is observed between U172 H2’-C174 H6. The

black circles represent the H2’(i)-H6/H8(i+1) peaks in each F1 slice…………………..183

Figure 3.5.11 Illustration of the NMR solution structures of the FMDV 16mer Mg2+

RNA

complex (a) Overlay of the 20 lowest RMSD structures with average RMSD of 0.17Å. (b)

Lowest RMSD solution structure (0.16Å). The red ribbon represents the RNA

backbone…………………………………………………………………………………187

Figure 3.5.12 The G178UAA181 tetraloop of the FMDV 16mer Mg2+

RNA complex NMR

structure shown in Figure 3.5.11b; the closing G177.C182 base pair is also illustrated.

Colour coding of nucleotides: guanosine (blue), uridine (cyan), adenosine (green) and

cytosine (red)…………………………………………………………………………….188

Figure 3.5.13 The G.A sheared base pair of the FMDV 16mer Mg2+

RNA NMR structure.

Hydrogen bonding distances between G178 NH2-A181 N7 (2.23Å) and G178 N3-A181

NH6 (3.45Å) are indicated by the red dotted lines……………………………………...189

Figure 3.6.1 Histogram illustrating the changes to the chemical shift of exchangeable and

non-exchangeable protons of the FMDV 16mer RNA, upon addition of 5eq of Mg2+

. Bars

shown in blue represent exchangeable protons. Bars shown in green and orange represent

non-exchangeable protons in the stem and loop, respectively. Negative values correspond

to a highfield chemical shift change and positive values correspond to a lowfield chemical

shift change……………………………………………………………………………...197

18

Figure 3.6.2 Illustration of the Mg2+

-induced structural changes to the GUAA tetraloop in

the FMDV 16mer RNA by comparison of the 16mer (a) apo-RNA structure and (b) Mg2+

RNA complex structure. The U179, A180 and A181 bases are stacked more tightly

together in the 16mer Mg2+

RNA complex, strengthening the base stacking

interactions………………………………………………………………………………197


-induced structural changes to the A181 nucleotide in


RNA complex structure. The G.A sheared base pair is formed by two hydrogen bonds

(black dashed lines); G178 NH2-A181 N7 is 3.09Å and 2.23Å in apo-RNA and Mg2+

complex, respectively and G178 N3-A181 NH6 is 5.05Å and 3.45Å in apo-RNA and

Mg2+

complex, respectively. Stronger base-phosphate intramolecular interactions were

formed (blue dashed line); A181 H8-A181 O5’ is 3.29Å and 2.65Å in the 16mer apo-

RNA and Mg2+

RNA complex, respectively…………………………………………….198


-induced structural changes to the A180 nucleotide in


RNA complex structure. A stronger base-phosphate intramolecular interaction is formed

(red dashed line); A180 H8-A180 O5’ is 3.08Å and 2.30Å in the 16mer apo-RNA and

Mg2+

RNA complex, respectively……………………………………………………….199

Figure 3.6.5 600 MHz 3D NOESY/2Q-COSY spectrum (250ms) of the FMDV 16mer (a)

apo-RNA and (b) Mg2+

RNA complex. The F2 chemical shift is labelled in each plane.

Both positive (orange) and negative (green) levels are shown. In the 16mer apo-RNA,

scalar coupling between A180 H1’ and A180 H2’ can be clearly observed by strong cross

peaks, characteristic of the C2’-endo sugar conformation. In the 16mer Mg2+

RNA

complex, coupling is still observed between A180 H1’-H2’, although with significantly

reduced intensity………………………………………………………………………...200


1H2O,

displaying the imino region. Four peaks were identified corresponding to the imino

protons of G229, U230, G231 and G240………………………………………………..202


in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks

correspond to imino-imino connectivities. The sequential assignment shown here is

between U230-G231. The G229 NH diagonal peak is shown with an arrow. Inset:

Secondary structure of the 15mer RNA highlighting the observed imino-imino

connectivities, represented by light blue oval shapes…………………...………………203

19


in 1H2O, illustrating the imino-amino region of the spectrum. Connectivities from imino

protons to NH2*/NH2/H2/H5 protons can be observed (NH2* corresponds to the proton

involved in base pair hydrogen bonding); connectivities are marked with a black

circle……………………………………………………………………………………..204


13C

HSQC spectrum and the subsequent assignment of H5-H6 cross peaks in the NOESY

spectrum, of the FMDV 15mer apo-RNA. Bottom left panel: 700 MHz NOESY (250ms)

spectrum, at 25°C in 2H2O; blue circles indicate H5-H6 cross peaks. Top left panel: 400

MHz 1H-

13C HSQC spectrum, at 25°C in

2H2O, displaying the C6-H6 peaks. Bottom right

panel: 400 MHz 1H-


2H2O, displaying the C5-H5

peaks…………………………………………………………………………………….206

Figure 4.1.5 700 MHz NOESY (250ms) spectrum of the FMDV 15mer apo-RNA, at

25°C in 2H2O. The blue line represents G229 H8-H1’ to A234 H1’-C235 H6 intra- and

internucleotide connectivities. The green line represents C238 H6-H1’ to C243 H6-H1’

intra- and internucleotide connectivities (same colouring in secondary structure shown).

H6/H8 and H2 chemical shifts are labelled in black and blue, respectively. The (i)

corresponds to an intranucleotide connectivity and (s) corresponds to a sequential

connectivity. The red circles correspond to H5-H6 connectivities……………………...207



13C HSQC

spectrum, of the FMDV 15mer apo-RNA. Bottom panels: 400 MHz 1H-

13C HSQC spectra

at 25°C in 2H2O. Top panels: 700 MHz NOESY (250ms) spectra at 25°C in

2H2O…….208

Figure 4.1.7 Illustration of the identification of H6/H8 and H1’ peaks in the NOESY

spectrum with the aid of both 1H-

13C HSQC and

1H-

31P CPMG-HSQC-NOESY spectra,

of the FMDV 15mer apo-RNA. (a) 600 MHz 1H-

31P CPMG-HSQC-NOESY spectrum, at

25°C in 2H2O, displaying the phosphorus-H6/H8 peaks. (b) 400 MHz

1H-

13C HSQC

spectrum, at 25°C in 2H2O, displaying the C6-H6 and C8-H8 peaks. (c) 700 MHz NOESY

(250ms) spectrum, at 25°C in 2H2O. (d) 400 MHz

1H-


2H2O, displaying the C1’-H1’ peaks. (e) 600 MHz

1H-

31P CPMG-HSQC-NOESY

spectrum at 25°C in 2H2O, displaying the phosphorus-H1’ peaks. The dashed lines in

panels (a) and (e) indicate phosphorus-H6/H8/H1’ peaks that could not be identified in the 1H-

31P CPMG-HSQC-NOESY spectrum………………………………………………..210

20

Figure 4.1.8 600 MHz 1H-

31P CPMG-HSQC-NOESY spectrum, at 25°C in

2H2O, of the

FMDV 15mer apo-RNA. Illustration of the phosphorus-H6/H8 (panel A), and

phosphorus-H1’ (panel B) correlations. Sequential assignment between G229-C232 is

represented by a blue line and between G240-C243 is indicated by a green line, in both

phosphorus-H6/H8 and phosphorus-H1’ regions. Peaks marked with a cross represent

phosphorus-proton correlations………………………………………………………….211

Figure 4.1.9 Illustration of the NMR solution structures of the FMDV 15mer apo-RNA. (a)

Overlay of the 20 lowest RMSD structures with average RMSD of 0.51Å. (b) Lowest

RMSD solution structure (0.35Å). The red ribbon represents the RNA backbone……..215

Figure 4.1.10 The heptaloop (A233ACCCCA239) of the FMDV 15mer apo-RNA NMR

structure, shown in Figure 4.1.9b. Colour coding of nucleotides: adenosine (green) and

cytosine (red)………………….…………………………………………………………216

Figure 4.2.1 600MHz 1H-NMR stack plot (imino region) of the 15mer RNA with (a) no

Mg2+

and (b) in the presence of 5.0eq Mg2+

, at 2°C in 1H2O……………………………223


RNA

complex, at 2°C in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal

peaks correspond to imino-imino connectivities. The sequential assignment shown here is

between U230-G231 and G231-G240. Inset: Secondary structure of the 15mer RNA

highlighting the observed imino-imino connectivities, represented by light blue oval

shapes……………………………………………………………………………………224


RNA

complex, at 2°C in 1H2O, illustrating the imino-amino region of the spectrum.

Connectivities from imino protons to NH2*/NH2/H2 protons can be observed (NH2*

corresponds to the proton involved in base pair hydrogen bonding); connectivities are

marked by a black circle…………………………………………………………………225


non-exchangeable protons of the FMDV 15mer RNA, upon addition of 5eq of Mg2+

. Bars

shown in blue represent exchangeable protons. Bars shown in green and orange represent

non-exchangeable protons in the stem and loop, respectively. Negative values correspond

to a highfield chemical shift change and positive values correspond to a lowfield chemical

shift change……………………………………………………………………………...226

21

Figure 4.3.1 1H-NMR stack plot of the 16mer apo-RNA (batch 2), at 2°C in

1H2O,

displaying the imino proton region with four different magnetic field strengths; (a) 400

MHz, (b) 600 MHz, (c) 800 MHz and (d) 1000 MHz. The intensity of the U176 and G178

imino proton peak is clearly reduced with increasing magnetic field strength, marked by

the red arrows……………………………………………………………………………228

Figure 4.3.2 Illustration of the NOESY (150ms) spectrum (imino-amino region) of the (a)

16mer apo-RNA (batch 1) at 700 MHz and (b) 16mer apo-RNA (batch 2) at 1GHz. The

G178 NH to C182 NH2* NOE cross peak can be clearly observed in the 1GHz NOESY

spectrum, but is absent in the 700 MHz NOESY spectrum……………………………..229

Figure 4.4.1 1H-NMR stack plot (imino region) of the (a) 16mer Mg

2+ RNA complex

(600MHz), (b) 15mer Mg2+

RNA complex (600MHz) and (c) RNA-RNA complex

(700MHz), at 2°C in 1H2O………………………………………………………………231

Figure 4.4.2 700 MHz NOESY (250ms) spectrum of the FMDV RNA-RNA complex, at

2°C in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks

correspond to imino-imino connectivities. The sequential assignment of the 16mer RNA

starts from G178 and finishes at G186 (black lines). The sequential assignment of the

15mer RNA starts from U230 and finishes at G240 (blue lines) Inset: Secondary structure

of the 16mer RNA (top) and 15mer RNA (bottom) highlighting the imino-imino

connectivities shown in the spectrum, represented by light blue oval shapes…………..232

Figure 4.4.3 700 MHz NOESY (250ms) spectrum of the FMDV RNA-RNA complex, at

2°C in 1H2O, illustrating the imino-amino region of the spectrum. Connectivities from

imino protons to NH2*/NH2/H2/H5/H1’ protons can be observed. (NH2* corresponds to

the proton involved in base pair hydrogen bonding); connectivities are marked by a black

and blue circle for the 16mer RNA and 15mer RNA, respectively. Assignments for both

the 16mer RNA (black) and 15mer RNA (blue) are shown……………………………..233

Figure 4.4.4 400 MHz 1H-NMR stack plot of the (a) 16mer Mg

2+ RNA complex, (b)

15mer Mg2+

RNA complex and (c) RNA-RNA complex, at 25°C in 2H2O The red asterisk

indicates the peaks of interest that show changes in chemical shift or linewidth……….235


non-exchangeable protons of the FMDV 16mer and 15mer Mg2+

RNAs, upon RNA-RNA

complex formation. Bars shown in blue and red represent proton chemical shift changes

identified in the NOESY spectrum in 1H2O and

2H2O, respectively. Negative values

correspond to a highfield chemical shift change and positive values correspond to a

lowfield chemical shift change…………………………………………………………..236

22


RNA-RNA complex, at 25°C in 2H2O. The black and blue lines represent the sequential

H6/H8-H1’ intra- and internucleotide connectivities, for the 16mer and 15mer Mg2+

RNAs,

respectively. The (i) corresponds to an intranucleotide connectivity and (s) corresponds to

a sequential connectivity. The red circles and light blue squares correspond to H5-H6

connectivities for the 16mer and 15mer Mg2+

RNAs, respectively. Assignments for both

the 16mer Mg2+

RNA (black) and 15mer Mg2+

RNA (blue) are shown………………...237

Figure 4.4.7 Scheme illustrating the possible RNA-RNA interaction between the FMDV

16mer RNA (left) and the 15mer RNA (right). The rectangles represent the bases, which

have been numbered. Base pairing is represented by three black lines for G.C base pairs

and two black lines for A.U base pairs. The unfilled circles correspond to the sugar ribose,

red circles represent the phosphorus atom and green triangles symbolise the base stacking

interactions. Chemical shift and linewidth changes to protons in nucleotides are

represented by the blue filled rectangles and circles. The broken black line between the

orange brackets indicates the possible area of RNA-RNA interaction………………….239

Figure 5.1.1 376 MHz 19

F-NMR stack plot of the 5-FU 16mer RNA, in 1H2O at (a) 2°C,

(b) 10°C and (c) 25°C. 19

F chemical shifts were referenced to CFCl3…………………..241

Figure 5.1.2 376 MHz 19

F-NMR stack plot of the 5-FU 15mer RNA, in 1H2O at (a) 2°C,

(b) 10°C and (c) 25°C. 19

F chemical shifts were referenced to CFCl3…………………..242

Figure 5.1.3 1GHz NOESY (150ms) spectrum of the 5-FU 16mer RNA at 2°C in 1H2O

(orange) overlaid on the 1GHz NOESY (250ms) spectrum of the unlabelled 16mer RNA

(batch 2) at 2°C in 1H2O (green). The overlay displays the U172, C173, C174, U175,

U176, U179 and C182 H5-H6 cross peaks found in the aromatic region of the spectra.

Cross peaks of the unlabelled 16mer RNA have been labelled by a cross……………...243

Figure 5.1.4 600 MHz NOESY (400ms) spectrum of the 5-FU 15mer RNA at 2°C in 1H2O (orange) overlaid on the 600 MHz NOESY (400ms) spectrum of the unlabelled

15mer RNA at 2°C in 1H2O (green). The overlay displays the U230, C232, C235, C236,

C237, C238, C241 and C243 H5-H6 cross peaks found in the aromatic region of the

spectra. Cross peaks of the unlabelled 15mer RNA have been labelled by a cross……..243

Figure 5.1.5 1GHz 1H-NMR stack plot (imino region) of the (a) 16mer apo-RNA (batch 2)

and (b) 5-FU 16mer RNA, at 2°C in 1H2O. The U175, U176, G185, G186, G177 and

G178 imino proton peaks are labelled…………………………………………………...244

23

Figure 5.1.6 1GHz NOESY (250ms) spectra of the FMDV 16mer apo-RNA (batch 2), left,

and 5-FU 16mer apo-RNA, right, at 2°C in 1H2O, illustrating the imino region of the

spectrum. The cross-diagonal peaks correspond to imino-imino connectivities. The

sequential assignment shown in both spectra is between G178-G186, although the G177-

U176 imino-imino connectivity is not observed for the 5-FU 16mer RNA. Insets:

Secondary structure of the 16mer RNA highlighting the observed imino-imino

connectivities, represented by light blue oval shapes…………………………………...245

Figure 5.1.7 1GHz NOESY (250ms) spectra of the FMDV 16mer apo-RNA (batch 2), left,

and 5-FU 16mer apo-RNA, right, at 2°C in 1H2O, illustrating the imino-amino region of

the spectrum. Connectivities from imino protons to NH2*/NH2/H2/H5/H1’ protons can be

observed (NH2* corresponds to the proton involved in base pair hydrogen bonding);

connectivities are marked by a black circle……………………………………………...246


5-FU 16mer RNA, at 25°C in 2H2O. The blue line represents intra- and internucleotide

connectivities from U172 H1’ to G177 H8 and the green line from U179 H1’ to A187 H8

connectivities. The (i) corresponds to an intranucleotide connectivity and (s) corresponds

to a sequential connectivity. The red circles correspond to H5-H6 connectivities……...247


19F HOESY (250ms) spectrum of the FMDV 5-FU 16mer RNA,

at 25°C in 2H2O.

19F chemical shifts were referenced to CF3COOH……………………248

Figure 5.1.10 162 MHz 31

P-NMR stack plot of the (a) 16mer apo-RNA (batch 1) and (b)

5-FU 16mer RNA, at 2°C in 1H2O. Peaks in the

31P-NMR spectrum of the 16mer apo-

RNA are labelled 1-9. Peaks 1, 2 and 9 correspond to U179, A180/C182 and A181

phosphorus (labelled in red), respectively……………………………………………….249

Figure 5.1.11 1H-NMR stack plot of the 5-FU 16mer RNA, at 2°C in

1H2O, displaying the

imino proton region with four different magnetic field strengths; (a) 400 MHz, (b) 600

MHz, (c) 800 MHz and (d) 1000 MHz. The intensity of the U176 and G178 imino proton

peak is clearly reduced with increasing magnetic field strength, marked by the red

arrows……………………………………………………………………………………250

Figure 5.1.12 A 1H-NMR stack plot (imino region) of the (a) 15mer apo-RNA (600 MHz)

and (b) 5-FU 15mer RNA (800 MHz), at 2°C in 1H2O. A lowfield shift of 0.78ppm is

observed for the U230 imino proton in the 5-FU 15mer RNA………………………….251

24

Figure 5.1.13 162 MHz 31

P-NMR stack lot of the (a) 15mer apo-RNA and (b) 5-FU

15mer RNA, at 25°C in 1H2O. A highfield shift of 0.52ppm is observed for the U230

phosphorus in the 5-FU 15mer RNA……………………………………………………252

Figure 5.1.14 376MHz 19

F-NMR stack plot of the 5FU 16mer RNA, with (a) no Mg2+

and

(b) with 5.0eq Mg2+

, at 25°C in 2H2O. A highfield shift of 0.27ppm was observed for the

U179 F5 peak in the 5-FU 16mer RNA. 19

F chemical shifts were referenced to

CFCl3…………………………………………………………………………………….253


19F HOESY (250ms) spectrum of the (a) FMDV 5-FU 16mer

apo-RNA and (b) FMDV 5-FU 16mer Mg2+

RNA complex, at 25°C in 2H2O.

19F chemical

shifts were referenced to CF3COOH…………………………………………………….254

Figure 5.1.16 376MHz 19


and

(b) with 5.0eq Mg2+

, at 25°C in 2H2O. A highfield shift of 0.05ppm was observed for the

U230 F5 peak in the 5-FU 15mer RNA. 19

F chemical shifts were referenced to

CFCl3…………………………………………………………………………………….254

Figure 5.2.1 376 MHz 19

F-NMR stack plot of the (a) 5-FU 16mer Mg2+

RNA complex, (b)

5-FU 15mer Mg2+

RNA complex, (c) 5-FU RNA-RNA complex, at 25°C in 2H2O. The red

asterisks represent the additional smaller peaks observed in the 5-FU RNA-RNA complex. 19

F chemical shifts were referenced to CFCl3.………………………………………………256


19F HOESY (250ms) spectrum of the (a) FMDV 5-FU 16mer

Mg2+

RNA complex and (b) FMDV 5-FU 15mer Mg2+


19F chemical shift referenced to CF3COOH. Assignments for both the 5-FU 16mer RNA

(black) and 5-FU 15mer RNA (blue) are shown………………………………………...256

25

List of Tables

Table 2.4.1 Summary of 1H and

13C chemical shifts observed in NOESY and

1H-

13C

HSQC spectra of RNA. H-bonded refers to ‘hydrogen’ bonded………………………..105

Table 2.5.1 Dihedral angle restraints used for defining nucleotide structure (α, β, γ, δ, ε, ζ,

χ) and the ribose sugar (ν1 and ν2) in the structure calculations.35

………………………113

Table 3.1.1 1H,

13C and

31P NMR chemical shifts of the FMDV 16mer apo-RNA, in

1H2O

and 2H2O…………………………………………………………………………………141

Table 3.1.2 A summary of the total number of restraints used for the structure

determination of the FMDV 16mer apo-RNA…………………………………………..143

Table 3.1.3 Ten intramolecular interactions in total were identified in the GUAA tetraloop

of the FMDV 16mer apo-RNA NMR structure. The interactions formed between donor

and acceptor atoms are given, the type of interaction, the specificity of the interaction and

the distances between the proton donor and acceptor atoms…………………………….147

Table 3.1.4 Local helical parameter values for the FMDV 16mer apo-RNA structure,

calculated by the 3DNA analysis program………………………………………………149

Table 3.1.5 Base pair step parameter values for the FMDV 16mer apo-RNA structure,

calculated by the 3DNA analysis program………………………………………………149

Table 3.1.6 Complementary base pair parameter values for the FMDV 16mer apo-RNA

structure, calculated by the 3DNA analysis program……………………………………149

Table 3.1.7 Dihedral angle values of nucleotides in the FMDV 16mer apo-RNA structure,

calculated by the 3DNA (black) and CURVES (red) analysis programs. Angles are all

measured in degrees……………………………………………………………………..150

Table 3.1.8 Dihedral angle values (ν1 and ν2), pseudorotation phase angle (Phase) and

amplitude (Amp) values that define the sugar ribose conformation for each nucleotide in

the FMDV 16mer apo-RNA structure. Values were calculated by the 3DNA (black) and

CURVES (red) analysis programs………………………………………………………151

Table 3.5.1 1H,

13C and

31P NMR chemical shifts of the FMDV 16mer Mg

2+ RNA

complex, in 1H2O and

2H2O……………………………………………………………..184

26


determination of the FMDV 16mer Mg2+

RNA complex……………………………….186

Table 3.5.3 Seven intramolecular interactions in total were identified in the GUAA

tetraloop of the FMDV 16mer Mg2+

RNA complex NMR structure. The interactions

formed between donor and acceptor atoms are given, the type of interaction, the

specificity of the interaction and the distances between the proton donor and acceptor

atoms…………………………………………………………………………………….190

Table 3.5.4 Local helical parameter values for the FMDV 16mer Mg2+

RNA complex


Table 3.5.5 Base pair step parameter values for the FMDV 16mer Mg2+

RNA complex


Table 3.5.6 Complementary base pair parameter values for the FMDV 16mer Mg2+

RNA

complex structure, calculated by the 3DNA analysis program………………………….192

Table 3.5.7 Dihedral angle values of nucleotides in the FMDV 16mer Mg2+

RNA

complex structure, calculated by the 3DNA (black) and CURVES (red) analysis programs.

Angles are all measured in degrees……………………………………………………...193



the FMDV 16mer Mg2+

RNA complex structure. Values were calculated by the 3DNA

(black) and CURVES (red) analysis programs………………………………………….194

Table 3.6.1 A comparison of dihedral angle and distance values obtained from the FMDV

16mer apo-RNA and Mg2+

RNA complex structures, highlighting the changes induced by

Mg2+

……………………………………………………………………………………..199

Table 4.1.1 1H,

13C and


1H2O

and 2H2O…………………………………………………………………………………212


determination of the FMDV 15mer apo-RNA…………………………………………..214

27

Table 4.1.3 Two specific and twelve non-specific intramolecular interactions identified in

the heptaloop of the FMDV 15mer apo-RNA NMR structure. The interactions formed

between donor and acceptor atoms are given, the type of interaction, the specificity of the

interaction and the distances between the proton donor and acceptor atoms……………217

Table 4.1.4 Local helical parameter values for the FMDV 15mer apo-RNA structure,

calculated by 3DNA analysis program…………………………………………………..218

Table 4.1.5 Base pair step parameter values for the FMDV 15mer apo-RNA structure,

calculated by 3DNA analysis program…………………………………………………..218

Table 4.1.6 Complementary base pair parameter values for the FMDV 15mer apo-RNA

structure, calculated by 3DNA analysis program………………………………………..219


calculated by the 3DNA (black) and CURVES (red) analysis programs. Angles are all

measured in degrees……………………………………………………………………..220



the FMDV 15mer apo-RNA structure. Values were calculated by the 3DNA (black) and

CURVES (red) programs………………………………………………………………..221

28

Symbols and Abbreviations

Symbols:

Å Ångström

A260 Absorbance at a wavelength of 260nm

B0 External magnetic field

°C Degrees Celsius

° Degree

Dx Shift

Dy Slide

Dz Rise

dx x-displacement

dy y-displacement

δ Chemical shift

Ep Potential energy

η Inclination

F1 First frequency dimension

F2 Second frequency dimension

F3 Third frequency dimension

h Helical rise

Hz Hertz

I Nuclear spin quantum number

J Coupling constant

K Kelvin

kex Exchange rate constant

κ Buckle

Mz Longitudinal magnetisation

Mxy Transverse magnetisation

φ Sugar pucker amplitude

P Pseudorotation phase angle

p0 Zero-order phase correction

p1 First-order phase correction

π Propeller

ρ Roll

R1 T1 relaxation rate

R1a Apparent T1 relaxation rate

S Svedburg

Sx Shear

Sy Stretch

Sz Stagger

σ Opening

29

θ Tip

T1 Spin-lattice relaxation time

T2 Spin-spin relaxation time

T2* Effective transverse relaxation time

t1, t2, t3 Evolution or detection period in pulse sequence

Tm Melting temperature

τ Tilt

τ Delay time

τ0 Base pair lifetime

τc Rotational correlation time

τm Mixing time

τm Magnetisation transfer delay

μ Nuclear magnetic moment

υ Resonance frequency

ω½ Linewidth

ω Twist

Ω Helical twist

W0 Zero quantum transition

W1 Single quantum transition

W2 Double quantum transition

30

Abbreviations:

1D One-dimensional

2D Two-dimensional

3D Three-dimensional

5-FU 5-fluorouridine

A Adenine

C Cytosine

CAT Chloramphenicol acetyltransferase

CHARMM Chemistry at Harvard macromolecular mechanics

COSY Correlated spectroscopy

CPG Controlled pore glass

CPMG Carr-Purcell-Meiboom-Gill

CrPV Cricket paralysis virus

DANTE Delays alternating with nutation for tailored excitation

DQF-COSY Double quantum filtered COSY

DMT 4,4-dimethoxytrityl

DNA Deoxyribonucleic acid

E.coli Escherichia coli

EFF Empirical force fields

eIF Eukaryotic initiation factor

eq Equivalents

EMCV Encephalomyocarditis virus

FID Free induction decay

G Guanine

GFP Green fluorescent protein

GHz Gigahertz

FMDV Foot-and-Mouth Disease virus

HCV Hepatitis C virus

HOESY Heteronuclear overhauser effect spectroscopy

HPLC High-performance liquid chromatography

HSQC Heteronuclear single quantum coherence

INEPT Insensitive nuclei enhancement by polarisation transfer

IRES Internal ribosome entry site

ITAF IRES trans-acting factor

IUPAC International union of pure and applied chemistry

LUC Luciferase

MD Molecular dynamics

Met Methionine

MHz Megahertz

miRNA microRNA

ml millilitre

MM Molecular mechanics

mM millimolar

31

mm millimetre

MQ Multiple quantum

mRNA Messenger RNA

ms milliseconds

NMR Nuclear magnetic resonance

NTPs Nucleotide triphosphates

nm nanometre

ns nanoseconds

NOE Nuclear overhauser effect

NOESY Nuclear overhauser effect spectroscopy

OD Optical density

ORF Open reading frame

PAGE Polyacrylamide gel electrophoresis

PES Potential energy surface

ppm parts per million

ps picoseconds

PTB Polypyrimidine tract binding protein

RDC Residual dipolar coupling

RF Radio-frequency

rMD Restrained molecular dynamics

RMSD Root Mean Square Deviation

RNA Ribonucleic acid

RNAi RNA interference

RNase Ribonuclease

siRNA Small interfering RNA

ssRNA Single-stranded RNA

TBDMS t-butyldimethylsilyl

TOCSY Total correlation spectroscopy

tRNA Transfer RNA

μg microgram

μl microlitre

μs microseconds

UTR Untranslated region

U Uracil

vdW van der Waals

VT Variable temperature

WATERGATE Water suppression by gradient-tailored excitation

32

Abstract

The structure, kinetics, and interactions of the conserved 16mer and 15mer RNA motifs of

the internal ribosome entry site (IRES) of the Foot-and-Mouth Disease virus (FMDV),

have been investigated by homonuclear and heteronuclear NMR techniques. The 16mer

RNA is endowed with a classic GNRA tetraloop motif, which is essential for IRES

activity and the 15mer RNA motif is a potential tetraloop receptor. We have determined

three high resolution NMR solution structures of the 16mer apo-RNA, the 16mer Mg2+

RNA complex and the 15mer apo-RNA with RMSDs of 0.17Å, 0.16Å and 0.35Å,

respectively. The high precision of these NMR structures was achieved by including a

large number of NMR experimental restraints, derived from NOEs and coupling constants,

and validating them using the MolProbity program. The 16mer RNA structure comprised

of six base pairs with a GUAA tetraloop and the 15mer RNA structure comprised of four

base pairs and a large heptaloop; this is the first heptaloop to be studied by NMR.

Addition of Mg2+

to the 16mer apo-RNA caused selective chemical shift changes to the

stem G177 and loop G178 imino proton resonances, suggesting Mg2+

-induced

conformational change to the GUAA tetraloop. This was supported by a significant

chemical shift change to the selectively 19

F-labelled loop U179 in the 5-FU 16mer RNA.

Furthermore, variable temperature experiments revealed retarded imino proton exchange

for the stem and loop imino protons, demonstrating the enhanced thermodynamic stability

conferred by Mg2+

. This enhancement in stability was confirmed by measuring the imino

proton exchange rates for the 16mer apo-RNA and the 16mer Mg2+

RNA complex.

Analysis of the 16mer apo-RNA and its Mg2+

RNA complex NMR solution structures

revealed that Mg2+

-induced structural changes to the GUAA tetraloop act to stabilise the

loop via stronger base stacking and intramolecular interactions. Fascinatingly, we

discovered that Mg2+

ions provide increased stability required for the formation of a G.A

sheared base pair in the GUAA tetraloop. RNA-RNA interactions between the 16mer and

15mer RNAs and their fluorinated analogues were studied by NMR spectroscopy. Small

changes to chemical shift and linewidth of proton peaks in the non-fluorinated RNA-RNA

complex provided evidence for a weak interaction between the loop of the 16mer RNA

and the stem of the 15mer RNA. 19

F-NMR experiments revealed additional peaks for the 19

F-labelled U179 of the fluorinated 16mer/15mer RNA complex providing further good

evidence of RNA-RNA interaction.

The NMR structures of the conserved RNA motifs and their interactions have yielded

important information in understanding the properties and behaviour of RNA. This will

provide the first stepping stone in understanding the IRES mechanism and its use in

antiviral therapy and biotechnology.

33

Declaration

No portion of the work referred to in this thesis has been submitted in support of an

application for another degree or qualification of this or any other university or other

institute of learning.

Copyright Statement

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certain copyright or related rights in it (the “Copyright”) and s/he has given The

University of Manchester certain rights to use such Copyright, including for

administrative purposes.

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may be made only in accordance with the Copyright, Designs and Patents Act 1988 (as

amended) and regulations issued under it or, where appropriate, in accordance with

licensing agreements which the University has from time to time. This page must form

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property (the “Intellectual Property”) and any reproductions of copyright works in the

thesis, for example graphs and tables (“Reproductions”), which may be described in this

thesis, may not be owned by the author and may be owned by third parties. Such

Intellectual Property and Reproductions cannot and must not be made available for use

without the prior written permission of the owner(s) of the relevant Intellectual Property

and/or Reproductions.

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University Library’s regulations (see

http://www.manchester.ac.uk/library/aboutus/regulations) and in The University’s policy

on presentation of Theses.

34

Acknowledgements

I would like to thank Allah the Almighty for giving me the opportunity to make a

contribution to scientific research and being able to finish this research project

successfully.

I would like to thank my supervisor Dr. Vasudevan Ramesh for his guidance and support

throughout my project and my colleagues (Dr Tony Cheung, Dr John King, Dr Misbah

Nareen, Misbah Ghafoor, Sadia Mohammed and Nick Chan) for helpful discussions.

For their assistance in use of NMR facilities, I would like to thank Tom Frankiel, Geoff

Kelly and Alain Oregioni of the National Institute of Medical Research (NIMR) in Mill

Hill, UK, Moreno Lelli of the Centre de RMN à Très Hauts Champs (CRMN) in Lyon,

France and Roger Speak at the School of Chemistry, University of Manchester.

Finally, I would like to express gratitude to my family for their continued support,

especially from my mother, Rehana Rasul, and my wife, Usma Rasul.

35

Chapter 1: Introduction

The introduction chapter consists of eight different sections. The first section highlights

the importance of the project and establishes the area of research. Sections 1.2 through to

1.6 provide relevant background, and finally the previous work and the aims of the project

are summarised in sections 1.7 and 1.8, respectively.

1.1 Significance of the project

1.1.1 The role of IRES in picornavirus translation

Protein synthesis involves the process of translation, in which eukaryotic messenger RNA

(mRNA) is decoded to produce proteins. Initiation of translation in mRNAs commences

with the 5ʹ-end cap-dependent recruitment of a 40S ribosomal unit, with the aid of specific

eukaryotic initiation factors (eIFs).1 In contrast, picornaviruses adopt a different method of

recruiting the ribosomal unit.

The picornaviruses are a family of animal viruses that contain positive-sense, single-

stranded, RNA genomes. They are able to take control of the host cell translation

machinery in a way that allows efficient translation of the viral coding region and

inhibition of host cell protein synthesis. An important feature of the picornaviral RNA

genome is the unusually long 5ʹ-untranslated region (5ʹ-UTR), preceding the viral coding

region. The 5ʹ-UTR exhibits a high degree of secondary structure elements, one of which

is a specialised regulatory element known as the internal ribosome entry site (IRES).2 The

IRES has attracted much attention as it is involved in a novel mechanism for the initiation

of protein synthesis, whereby translation is cap-independent.3 The IRES element is a

distinctive feature of the picornavirus genome, whose secondary structures are

phylogenetically conserved. However, different classes of the IRES element show no

similarity in primary sequence or secondary structure.4

This raises the question of how the

IRES elements can direct initiation of translation with dissimilar sequence and structure.

The most probable answer is that the tertiary structure of the IRES RNA plays a critical

36

role in the initiation of translation in picornaviruses. Both structural and functional studies

have shown a close relationship between IRES RNA structure and biological activity.5-6

In

addition, a second factor, which is also essential for IRES activity, is the RNA-binding

proteins such as eukaryotic initiation factors (eIFs) and IRES trans-acting factors

(ITAFs).7 Understanding the relationship between these two factors and IRES function

would improve our knowledge of cap-independent translation in viral RNAs.

The Picornaviridae family encompass many different species including the Foot-and-

Mouth disease virus (FMDV) and the Encephalomyocarditis virus (EMCV). The FMDV

IRES consists of multiple domains (domains 1-5); shown in Figure 1.2.4.8 Domain 3

contains a highly conserved hammerhead structure that includes a 4-way junction and 3

short stem-loop structures (Figure 1.2.5). At the apex of one of the short stem-loops is a

GNRA motif that has been found to be essential for IRES activity. The GNRA motif (N

represents any base and R represents a purine) is a tetraloop spanning residues

G178UAA181 in the FMDV IRES. Fascinatingly, mutation at any of the four bases in the

GUAA tetraloop, especially at the 4th

position, results in a considerable reduction in IRES

activity.9 It has been suggested that the GNRA motif plays a critical role in structural

organisation in the apical region of domain 3, possibly involving RNA-RNA interactions

to GNRA binding sites in domain 3.10

However, the exact role played by the GNRA motif

is still not well understood. Therefore, knowledge of the structure-function relationship of

the conserved GNRA motif is extremely important in determining the molecular basis of

its critical role in IRES function.

Understanding IRES biology is essential for providing insight into translational initiation

in viral RNAs. However, the basic mechanisms through which the IRES recruits the

ribosome have only recently come to light. The IRES mechanism in the FMDV is still

relatively unexplored and not well understood. A systematic study of the structure,

kinetics and interactions of conserved RNA motifs in the FMDV IRES will help to

decipher the mechanism of its function. Elucidating this mechanism should enable

scientists to develop new antiviral approaches and strategies for gene therapy, which has

been highlighted in the next two sub-sections.

37

1.1.2 The IRES and antiviral therapy

The IRES is essential in the FMDV life cycle and remains highly conserved within species.

Therefore, inhibition of the process of FMDV translation represents an attractive target in

the development of novel antiviral drugs. This sub-section will highlight the therapeutic

approaches being developed to target FMDV translation for the development of novel

antiviral therapy.

A number of different approaches have been investigated to target viral translation.

Strategies under investigation include targeting the cis-acting IRES and the trans-acting

host cell factors, which are essential for viral translation. The most common techniques for

targeting the IRES are oligonucleotide-based therapeutics such as antisense

oligonucleotides, RNA interference (RNAi), nucleic acid aptamers and ribozymes.

Additionally, small molecule inhibitors that target specific sites in the IRES are also

becoming a popular choice for therapy. These approaches intend to block translation

initiation in viral RNAs and can ultimately provide a cure for diseases such as Foot-And-

Mouth disease.

Antisense oligonucleotides are short nucleic acid molecules whose sequence is

complementary to the target RNA molecule. They can bind to the target RNA sequence, in

this case the IRES, and inhibit translation either through steric interference of the

translation machinery, changing the conformation of the RNA structure or cleavage of

RNA by RNase H. Antisense oligonucleotides have been directed against the FMDV

IRES resulting in significant reduction of FMDV translation initiation.11

RNAi controls

the post-transcriptional regulation of gene expression using microRNA (miRNA) and

small interfering RNA (siRNA). These RNA molecules can bind to complementary

sequences of the target RNA, which leads to the silencing of target gene expression. For

example, binding to mRNA can alter the level of protein expression. RNAi has been

investigated as a potential treatment against a variety of diseases including Hepatitis C.

Studies have used RNAi to target a conserved region of the Hepatitis C Virus (HCV)

IRES to inhibit translation, significantly reducing HCV infection in cell culture.12

Nucleic

38

acid aptamers are single stranded DNA or RNA molecules that can adopt complex three-

dimensional structures, which bind to a specific target molecule. An advantage of using

aptamers is that they can bind to highly structured targets such as the IRES RNA structure.

For example, RNA aptamers have been targeted against the HCV IRES, whereby two

aptamers were isolated that bound successfully to the HCV IRES, leading to inhibition of

IRES-dependent translation.13

Ribozymes, ribonucleic acid enzymes, are catalytic RNA

molecules, which are able to cleave their own phosphodiester bonds (self-cleaving) or of

other RNA molecules. There are many different types of ribozymes, including the

hammerhead ribozymes, group I/II introns and RNase P. The use of ribozymes to target

against the IRES in picornaviruses has been investigated as a therapeutic option. The

RNase P ribozyme has been shown to recognise the FMDV IRES element, leading to

strong inhibition of translation in cell culture.14

In addition to oligonucleotide-based strategies, small molecule inhibitors have also been

used to target the IRES in picornaviruses. Quinacrine is a well known compound with a

variety of different medical applications. It is a nucleic acid intercalator capable of

inhibiting replication, transcription and translation. Interestingly, it has also been shown to

inhibit cell-free IRES-mediated translation in EMCV, HCV and poliovirus.15

A new class

of benzimidazole inhibitors has also been found to show high affinity for domain IIa of the

HCV IRES, inhibiting HCV translation.16

Furthermore, NMR investigation revealed the

mechanism of action; it was determined that inhibitor binding induced a major

conformational change in domain IIa of HCV IRES.17

The results from these studies are very promising and provide an opportunity to develop

antiviral tools in the treatment of viral infections such as the Foot-And-Mouth disease.

However, these approaches will require a better understanding of the mechanism of

translation initiation in the FMDV and the role of cellular and viral trans-acting factors in

modulating IRES activity. This justifies the investigation of the FMDV IRES, which may

eventually have far-reaching implications for medicinal virology.

39

1.1.3 The IRES in biotechnology

Bicistronic vectors allow the efficient expression of two distinct coding sequences and are

important tools for molecular biology and gene therapy. One of the most promising

strategies to co-express multiple genes is to incorporate an IRES element into a gene

vector design.18

In IRES bicistronic expression vectors, two cistrons are linked together by

an IRES allowing co-translational expression of both genes. The advantage is that only

one promoter is needed for the expression of two genes. In biotechnology applications, the

expression of two proteins is often required, one being a marker or reporter gene such as

the green fluorescent protein (GFP), and the other being the gene of interest. This allows

the tracking and/or selection of transduced cells based on the detection of the selectable

marker protein. Recently, novel types of IRES-based retroviral vectors are being used for

expression and functional studies.19

The bicistronic vector design has been exploited to measure IRES activity by using

bicistronic reporter assays. Two reporter genes are placed either side of the IRES, which

code for two different proteins such as chloramphenicol acetyltransferase (CAT) and

luciferase (LUC). The first gene coding for CAT is translated by a cap-dependent manner

while translation initiation for the second gene coding for LUC is directed by the IRES,

which is cap-independent (Figure 1.1.1). The advantage of this setup is that the IRES can

independently drive translation of the downstream cistron. Subsequently, IRES activity is

measured by quantifying the expression of LUC, relative to the expression of CAT.

Bicistronic reporter assays have been invaluable to explore the link between IRES

structure and translation initiation in the FMDV. Several mutational studies have

investigated the sensitivity of the IRES to nucleotide substitutions. The most revealing

study showed that two highly conserved RNA motifs (GUAA and RAAA) in domain 3 of

the FMDV IRES, are essential for IRES function.9 This has prompted the recent escalation

of interest in the IRES element, and for the first time, includes the NMR investigation of

conserved RNA motifs in the FMDV IRES.

40

Figure 1.1.1 An IRES bicistronic expression vector. CAT and LUC are the first and second

reporter genes, respectively, surrounding the FMDV IRES. The arrow depicts the direction of

translation.

One of the advantages of IRES bicistronic expression, which makes it potentially of great

interest, is that simultaneous expression of two proteins may be required for novel gene

therapy. For the treatment of single gene defects, a selected marker gene is often necessary

to achieve sufficient expression of the therapeutic gene. For the treatment of disorders

with multiple gene defects such as cancer, a strategy for simultaneously targeting different

defective genes is required. The most popular IRES element used for gene therapy is from

the EMCV, owing to its high translation efficiency compared to IRES elements of other

picornaviruses. New bicistronic expression vectors are being developed to increase IRES

controlled expression.20

Thus, the importance of the IRES in molecular biology and gene therapy is apparent from

the above. However, since IRES-mediated translation is dependent on the availability of

host cell and viral proteins, adequate gene expression from the EMCV IRES element is

not always achieved. Therefore, in order to provide more effective IRES bicistronic

expression vectors, we must gain a deeper understanding of the underlying mechanisms

that control IRES activity. Ultimately, this will lay the foundation of future expression

vectors for biotechnology applications.

41

1.1.4 RNA structural biology

Currently, there has been great emphasis on studying the IRES. This is on a par with the

recent explosion in RNA research as RNA molecules have been shown to be involved in

wide variety of biological processes. Not only will scientists have a better comprehension

of the complexities of RNA structure, kinetics and interactions, but will also be able to

apply this knowledge into key technologies, eg. antiviral therapeutics and construction of

new expression vectors, as previously described.

However, how does one approach the large task of studying RNA in the form of the IRES.

Since the structure of biological macromolecules such as proteins and nucleic acids is

important for their function, structural biologists have endeavoured to unravel the three

dimensional structures of RNA in the hope that it will provide new information about their

function. X-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy are

two techniques that have been extensively used for the structure determination of proteins

and nucleic acid molecules.

The main advantage of using X-ray methods was that large RNA molecules could be

studied, eg. transfer RNA (tRNA). Recently, the 2009 Nobel Prize in Chemistry was given

for studies of the structure and function of the ribosome, in which the structure of a whole

ribosome was determined. Clearly, high-resolution X-ray structures of RNA have helped

structural biologists advance in the area of RNA biology. In contrast, NMR spectroscopy

methods are still limited to smaller RNA structures. However, it does provide several,

unique, advantages over X-ray methods, which have had a profound effect on our

understanding of RNA. Firstly, non-crystalline samples are used, which means that they

are studied in solution; commonly known as solution-state NMR. The advantage is that it

allows biomolecules to be studied under physiological conditions found in the cell.

Additionally, solution conditions can be varied such as temperature, pH and ionic strength,

which provide the possibility of studying biomolecules in more depth. Secondly, NMR

spectroscopy can be applied for molecules where single crystals cannot be produced. It

has generally been more difficult to obtain well-ordered RNA crystals for high-resolution

42

X-ray diffraction. Therefore, in these cases, NMR spectroscopy is the only available

option for high-resolution structure determination. Thirdly, the study of dynamics has

provided a new avenue for structural biologists as the focus has mainly been on structure

alone. NMR spectroscopy has been used to discover flexible regions in RNA structure,

identify local dynamical motions and study base pair kinetics. Evidently, NMR

spectroscopy provides the necessary capability and means to study the structure, kinetics

and interactions of the conserved RNA motifs in the FMDV IRES, and so is by far the

most appropriate biophysical technique to employ in this project.

Comparatively, structure determination of RNA by NMR spectroscopy has far lagged

behind that of proteins. As of March 2012, 87.4% of NMR structures published on the

PDB website are of proteins and only a small proportion of 4.4% are of RNA. In the past,

this has been due to the traditional view that RNA plays a minor role in the transfer of

genetic information and that proteins play a major role in biological processes such as

disease. The main challenge for NMR of RNA has always been the low dispersion, signal

overlap and rapid signal decay that occurs with increasing molecular weight. However,

with the advent of cheaper isotopic labelling of RNA and new labelling strategies, a new

era of RNA structural biology has arrived. This presents a great opportunity for scientists

to pave the way for detailed studies on RNA. Therefore, this project goes beyond trying to

understand the IRES mechanism, but using the IRES as a model for studying RNA

structure, kinetics and interactions.

43

1.2 Picornaviruses and translation

1.2.1 Picornavirus classification and genome

Picornaviruses are animal viruses composed of icosahedral capsid symmetry, in the

absence of an envelope. The viral capsid is composed of four viral proteins that

encapsulate a ‘positive-sense’ single strand ribonucleic acid (ssRNA). Positive-sense RNA

can be directly translated into desired viral proteins; the viral RNA genome is identical to

viral mRNA so it can be immediately translated by the host cell. The picornaviruses are

classified into genera based on their nucleotide sequences. Species within the same genus

will therefore contain similar nucleotide sequences and have a higher level of homology.

Currently, the family of picornaviruses have been divided into nine genera in the most

recent version of virus taxonomy.21

The Foot-And-Mouth Disease Virus (FMDV) is classified as an Aphthovirus. The FMDV

viral RNA is approximately 8,500 bases long consisting of a 5ʹ-untranslated region (5ʹ-

UTR), a coding region and a 3ʹ-untranslated region (3ʹ-UTR).22

The full FMDV genome is

illustrated in (Figure 1.2.1). The viral genome is translated as an open reading frame

(ORF), which starts downstream of the 5ʹ-UTR and ends upstream of the 3ʹ-UTR region.

Figure 1.2.1 The genomic structure of the FMDV viral RNA. The coding region or ORF is

divided into the L-region and three distinctive regions of P1, P2 and P3, coding for the capsid and

non-structural proteins. The 5ʹ-UTR precedes the coding region and the 3ʹ-UTR is situated after

the coding region.

44

1.2.2 mRNA and cap-dependent translation

Translation initiation in picornaviruses is different from normal eukaryotic mRNA

translation. mRNA translation is cap-dependent (Figure 1.2.2), but picornaviruses utilise a

unique mechanism of translation that is cap-independent (Figure 1.2.3). Normal mRNA

has a cap structure located at the 5ʹ-end of the mRNA molecule. It consists of a modified

guanine base (7-methylguanosine) that is attached to the mRNA via an unusual 5ʹ to 5ʹ

triphosphate linkage.

Figure 1.2.2 The normal cap-dependent translation. The mRNA requires the 5ʹ-end cap (7-

methylguanosine) structure along with complex interaction between the 40S ribosome and several

eukaryotic initiation factors (eIFs) to allow for translation initiation.

The initiation of translation in mRNA requires the 5ʹ-end cap structure, a 40S ribosome,

tRNA and several eukaryotic initiation factors (eIFs). The 40S ribosome can only gain

access to the initiation site via the 5ʹ-end of the mRNA. A cap-binding protein (eIF4E)

binds to the N-terminal domain of eIF4G, which is essential for cap-dependent translation

initiation. This initiation factor is part of the eIF4F complex that is also comprised of

eIF4G and eIF4A. The eIF4G acts as a bridge connecting the mRNA 5ʹ-end cap structure

to the 43S pre-initiation complex, which is composed of the 40S ribosomal unit and the

eIF2-GTP-tRNAMet

ternary complex.23

This interaction recruits the 43S complex, whereby

it binds to the C-terminal of the eIF4G bridge via eIF3, forming the 48S pre-initiation

complex. The ribosome scanning mechanism is aided by eIF1, eIF1A and eIF5, scanning

in the 5ʹ to 3ʹ direction until an authentic AUG initiation codon is found.

45

1.2.3 FMDV and cap-independent translation

Picornaviruses have two main characteristics, which prohibit their use of the conventional

cap-dependent mechanism for translation initiation. Firstly, picornaviral RNA does not

have a cap structure at its 5ʹ-end, unlike most mRNAs, which would prevent the cap-

binding complex from assembling. Secondly, the 5ʹ-UTR is very long and has several non-

authentic start codons, making canonical translation initiation very unlikely. Generally,

cap-independent translation of mRNA can be very inefficient, however, this is not the case

in picornaviral RNA translation. In picornaviral translation, initiation occurs by a cap-

independent mechanism that does not require the 5ʹ-end cap structure and whereby the

40S ribosome can recognise the authentic initiation codon.

Figure 1.2.3 The IRES mediated cap-independent translation. Translation initiation does not

require the 5ʹ-end cap structure and eIF4E in picornaviruses. Instead, the IRES provides an

entry/binding site for the 40S ribosome, so the ribosomal scanning can start.

Cap-dependent mRNA is inhibited in infected cells due to cleavage of the N-terminal of

eIF4G by viral L-proteases. This results in the separation of the cap-binding function

associated with N-terminus of eIF4G. Cleavage leaves one-third of the N-terminal

fragment and two-thirds of the C-terminal fragment.24

Initiation of translation in

picornaviruses only requires the C-terminal of eIF4G, which binds to the highly structured

picornaviral IRES and interacts with 40S ribosomal subunit bound eIF4A and eIF3.25

In

addition, the FMDV IRES also requires two IRES trans-acting factors known as

polypyrimidine tract binding protein (PTB) and ITAF45, for ribosomal binding.26

46

1.2.4 Internal Ribosome Entry Site (IRES)

Four major types of picornaviral IRES can be classified; type I of the Enterovirus group,

type II of the Cardiovirus and Aphthovirus groups and type III of the Hepatovirus

group.27

A fourth type has also been recently identified of the Teschovirus group. The type

I, II and III groups of picornaviral IRES elements have some similar structural features,

with a large central domain that has a characteristic four-way junction, which is essential

for IRES function. Besides the Picornavirdae family, IRES elements can also be found in

Flaviviridae and Dicistroviridae family of viruses.

IRESs are diverse RNA structures, which contain highly conserved secondary structures.

It is suggested that the IRES functions by replacing the cap structure and some eukaryotic

initiation factors (eIFs) with highly structured RNA. However, distinct differences can be

found between the IRES structures studied, based on their secondary/tertiary structure and

protein factor requirements.28

Three main types of IRES can be classified, the first are

compact folded IRES, the second are extended IRES with compact regions and the third

are extended and largely flexible IRES.29

The first class of IRES are the most highly

structured with tightly folded structures, eg. Cricket paralysis virus (CrPv) IRES. These

IRESs do not require any initiation factors and essentially operate as an RNA-based

ribosome recruitment apparatus. The second class of IRES are mostly extended but

maintain some tightly packed regions, eg. Hepatitis C virus (HCV) IRES. eIF2 and eIF3

are required to recruit the 80S ribosome and initiate translation. The third class of IRES do

not fold into globally compact structures, but retain some conformational flexibility, eg.

FMDV and EMCV IRES. This class of IRES do not fold into compact structures, and as

they are less structured, they require the most number of initiation factors as well as IRES

trans-acting factors (ITAFs). ITAFs are RNA-binding proteins that are able to enhance

IRES activity, but are not directly involved in the process of translation initiation. It has

been shown that ITAFs induce conformational changes in the IRES, which stabilises the

IRES conformation and forms a more compact RNA structure, allowing efficient

ribosomal recruitment.30

47

1.2.5 FMDV IRES

The FMDV IRES is an extended RNA region in the 5ʹ-UTR, spanning approximately 460

nucleotides, with a high degree of conserved secondary structure. The FMDV IRES is

modelled into five different domain structures, termed 1-5, from the 5ʹ to 3ʹ-end (Figure

1.2.4).

Figure 1.2.4 The FMDV IRES structure separated into five domains (1-5), from residues 1 (5ʹ-C)

to 462 (U-3ʹ). Domain 3 is the central domain, from residues 86 (5ʹ-G) to 299 (C-3ʹ). The apical

region of domain 3 is found within the orange circle.

440 450 460

U

C

AU

G

A

G

CA

U

A

U

G

C

G

C

A

U

U

U

U

C

CC

CC

C

C

CGG G

GG

GG

GAA

A

A

AA AA

A

CC

C

C

C

CC

C

C

C

G

GG

G

G

G

C

G

CA

G

C

C160

180

170

190

AUUAGC

C

GCA

200

210

220

C

C

G

A

U

G

CA

C

G

C

A

GC GCGGUAU CG

AC

CU

A

U

G

C

GG

G

C

UAG

230

240

150

140

GU

CA

UCACG

UUU

AC

GC

AA

AU

AAG

AU

CC

UG

CGAA

U

UC

CG

UGUG

AUGCUA

AA

AUUA

CC

GA

C

G U

AUGC

G

AG

C

UC

CGCGAU

CC

GAA

C

UC

UAC

A

UC

U

AG

CG

U C

5` 3`

130

120

110

100

90

250

260

270

280

290

86 299

CA

G

AC

C

AC

UG

G

C

UC

C

G

GA

GC

350

A

GC

G

A

C

UU

G

AG

UAGC

CA

G

U

AC

A

U

GA

360

340

330

GAGC

CAAAA

CG

UAUU

CGUG

GC

AA A

UUCGCGUAGCAGUGGU A A U UA UU

CGCG

U ACGGC

A

GCCGUG

AA

G

CCUUUA AC AUUAAUGACCCU

320

310

300

370

380

390

400

410

420

430A

C

U

CC

GG

U UUU

U AGCGCUG

GC

CU

AA

GGCGUGC

U AU A

AA

G

A UA U

CCAUC

GCUG

U ACG UCAAAACCAAG

GCA U

CGAG

U AUUC C

CC

A1

10

20

30

40

50

60

70

80

5` 3` 5` 3`

1

2

3

4

5

48

Interestingly, all five domains of the FMDV IRES appear to have different functions.31

Domain 3 is the central domain, which acts as a scaffold structure by forming long-range

RNA-RNA interactions with the other four domains.32

Situated in the apical region of

domain 3 is a highly conserved hammerhead structure. The hammerhead is constituted of

the 16mer RNA, which contains the essential GUAA tetraloop, the 36mer and 79mer

RNA motifs (Figure 1.2.5). Small insertions or deletions in this hammerhead region,

especially to the GNRA (G178UAA181) tetraloop in the 16mer RNA and the RAAA motif

(A199AAA202), can have drastic affects on IRES activity.9

The 15mer RNA, a potential

receptor for the GUAA tetraloop, is also found in domain 3 of the FMDV IRES, below the

hammerhead region (Figure 1.2.6). Mutational analysis has shown that substitution of

G240CACG244 can cause a significant decrease in IRES activity as severe as the

substitution to the GNRA tetraloop.10

Therefore, it has been proposed that long-range

tertiary contacts between the GNRA motif and the stem region of the 15mer RNA are

required for IRES activity.

Figure 1.2.5 Illustration of the hammerhead region, a 79mer RNA (G150-U228) found in domain

3 of the FMDV IRES. The 16mer RNA (U172-A187) shown in the red box and 36mer RNA

(C159-G194) in the blue box, are displayed.

A

U

G

A

G

C A

U

A

U

G

C

G

C

A

U

U

U

U

C

C

C

C C

C

C

CG G G

G G

G

G

G

A

A

A

A

A

A AA

A

C

C

C

C

C

C

C

C

C

C

G

G

G

G

G

G

C

G

C

A

G

C

C

5` 3`

150

160

180

170

190

AU

UA

GC

UG

C

GC

A

200

210

220

228

49

Figure 1.2.6 Illustration of the apical region of domain 3 in the FMDV IRES, located below the

hammerhead region. The 15mer RNA (G229-C243) is indicated by the area inside the red box.

Domains 2, 4 and 5 are involved in the interaction with RNA binding proteins, which

include the eukaryotic initiation factors (eIFs) and IRES trans-acting factors (ITAFs).

Domain 2 is a stem-loop structure, which consists of four helical sections separated by

three bulges. The loop contains five nucleotides, which forms a binding site for the ITAF

called polypyrimidine tract-binding protein (PTB). Two stem-loop structures are found in

domain 4, which are essential for the interaction between the FMDV IRES and eIF4G.33

Particularly, mutational analysis has shown that specific nucleotides are vital for this

RNA-protein interaction; nucleotides A312A313 in the distal part of domain 4 and two

highly conserved dinucleotides, A329C330 and G360A361. This RNA-protein interaction is

another important area for studying cap-independent translation.

Domain 5 is a stem-loop structure, which consists of two helices separated by one

conserved nucleotide (A424) and a tri-loop. The nucleotides in the loop and the bulged

A424 nucleotide are both critical for binding of eIF4B to the FMDV IRES.33

Domain 5 is

linked to a single stranded region approximately 25 nucleotides in length, known as the

polypyrimidine tract, which leads to the authentic AUG codon. There are two AUG

codons found in the ORF, of which both can function as authentic codons depending on

the genera of virus. In FMDV, initiation of translation can occur at either the first AUG

(termed Lab) or at the downstream AUG codon (termed Lb).11

AU

UA

GC

C

C

G

A

U

G

CA

C

G

C

AU

GC G

CG

GU

AU

AC

C

U

A

U

G

C

GG

G

C

U

A

G

230

240

150

140

5` 3`

248135

50

1.3 Nucleic acid chemistry

1.3.1 Nucleic acids

Nucleic acids are complex macromolecules composed of nucleotide sequences, commonly

found in the form of deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). They play

a central role in the transmission, expression and conservation of genetic information.

Nucleotides consist of three major components; a nitrogenous heterocyclic base, which

can be a monocyclic pyrimidine or bicyclic purine, a ribose or deoxyribose sugar and a

phosphate residue.34

Bases can be characterised into two heterocyclic types; purines,

adenine (A) and guanine (G) or pyrimidines, cytosine (C), thymine (T) and uracil (U).

Adenine, guanine and cytosine bases can be found in both DNA and RNA. However,

thymine is only found in DNA, and uracil only in RNA. The structures of these bases are

illustrated in (Figure 1.3.1).

Figure 1.3.1 Structures of purine and pyrimidine bases, and sugars, in DNA and RNA. Atoms are

numbered according to the IUPAC convention.35

51

In nucleosides the purine or pyrimidine base is attached to a five-membered sugar ring.

The ring nitrogen atom (N1) in cytosine, thymine and uracil is attached to the C1ʹ atom in

the sugar. Analogously, the ring nitrogen atom (N9) in adenine and guanine is also

attached to the C1ʹ atom in the sugar. The numbers corresponding to sugar atoms are

primed so that they can be distinguished from base atoms, according to the IUPAC

convention.35

Each sugar is connected to one of one of four possible heterocyclic bases in

DNA and RNA. There are two different configurations of the sugar moiety: 2-deoxyribose

found in DNA and ribose in RNA. The difference between the two is the 2ʹ-H group in

DNA being replaced by 2ʹ-hydroxyl (2ʹ-OH) group in RNA (Figure 1.3.1).

From nucleosides, nucleotides are formed by the addition of a phosphate group that forms

a phosphodiester bond. Two sugars are linked together by covalently bonding the

phosphate group to the 3ʹ carbon atom of one sugar and 5ʹ carbon atom of the other sugar

(Figure 1.3.2).

Figure 1.3.2 A phosphodiester bond linking two nucleosides, adenosine and guanosine, from the

5ʹ to 3ʹ-end.

52

1.3.2 RNA synthesis

1.3.2.1 Chemical synthesis

Chemical synthesis is normally the method of choice when synthesising small RNAs of

typically less than 30 nucleotides, due to low yields and high costs for larger RNAs. The

advantage of chemical synthesis is that modified nucleotides with site-specific labelling

can be introduced, eg. fluorinated nucleotides. However, the main disadvantage is that the

RNA cannot be isotopically labelled.

Solid phase synthesis is the method by which peptides and nucleic acids are synthesised

chemically. This method involves molecules being bound onto a solid support and

synthesised in a step-wise manner by a coupling reaction. The advantage is that a large

excess of the reactants can be used to force the reaction to high yield and the excess

reactants or by-products can be removed easily. In RNA synthesis, phosphoramidite

building blocks are used to synthesise the desired RNA sequence. Naturally occurring

nucleotides are insufficiently reactive to produce high yields. Therefore, nucleoside

phosphoramidites are used as they dramatically improve the selectivity and rate of

formation of internucleosidic linkages.

Phosphoramidites are protected at all reactive functional groups by attaching protecting

groups to prevent any undesired reactions. There are four different types of protecting

group, which are required for the 5ʹ-OH, 2ʹ-OH, 3ʹ-OH and the base amino groups (Figure

1.3.3). The 5ʹ-OH, 2ʹ-OH and 3ʹ-OH groups are protected by DMT (4,4-dimethoxytrityl),

TBDMS (t-butyldimethylsilyl) and 2-cyanoethyl groups, respectively. The base amino

groups in adenine, guanine and cytosine are protected by isobutyryl groups.

53

Figure 1.3.3 A phosphoramidite building block with four different sites required for protection.

Chemical synthesis of RNA is carried out from the 3ʹ to 5ʹ direction in a stepwise addition

of nucleotides. There are four essential chemical reactions of solid phase synthesis;

deprotection, coupling, capping and oxidation. Initially, the first phosphoramidite is

attached via its 3ʹ-OH group to a solid support; the most common solid support used is the

controlled pore glass (CPG). The deprotection step allows the addition of the next

monomer by removing the DMT group protecting the 5ʹ-OH. This results in a free 5ʹ-OH

group on the solid support bound nucleotide precursor. Since the 5ʹ-OH group is the only

reactive species on the precursor, it ensures that the next monomer will only bind to that

site. Any excess reactants and by-products are removed by washing the reaction column.

The second step is the coupling reaction, in which the phosphoramidite monomer is

activated by cleaving one of the groups protecting the phosphorus linkage. The 5ʹ-OH

group of the nucleotide precursor can then react with the activated phosphoramidite to

form a phosphite triester linkage. Any excess reactants, unbound base and by-products are

removed. The next step is capping, which is a safety step, designed to stop unreacted

nucleotide precursors from reacting with phosphoramidites later on in the reaction. This

can produce undesired oligonucleotides with deletions in their sequence. To prevent this

54

from happening, the unreacted 5ʹ-OH groups in the nucleotide precursors are capped with

a protective group. Again, any excess reactants, unbound base and by-products are

removed. The fourth step is the oxidation of the phosphite triester linkage produced in the

coupling reaction. This linkage is unstable under the conditions of synthesis and so it is

oxidised into a tetra-coordinated phosphate triester. This phosphate linkage is much more

stable and is a precursor of the naturally occurring phosphodiester bond found between

two nucleosides.

These four steps are repeated until the desired phosphoramidites are added to the

oligonucleotide. Subsequently, a final step is required, which involves cleavage,

deprotection and purification. The phosphoramidite attached to the solid support is

cleaved and all the protecting groups in the oligonucleotide chain are removed. The final

product consists of the desired oligonucleotide, cleaved protective groups and

oligonucleotides with deletions. To eliminate the undesired products, the oligonucleotides

are purified using polyacrylamide gel electrophoresis (PAGE) or chromatography

techniques such as HPLC (high-performance liquid chromatography).

1.3.2.2 Enzymatic synthesis

A powerful method of in vitro enzymatic transcription uses T7 RNA polymerase from the

T7 bacteriophage, to catalyse the formation of RNA from a synthetic DNA template. The

advantage of the enzymatic method is that larger RNAs can be synthesised (up to few

thousand nucleotides) and it allows simple incorporation of commercially available 2H,

13C and

15N isotopically labelled nucleotides. Enzymatic synthesis is much cheaper than

chemical synthesis, but it involves a great deal of labour intensive work.

T7 polymerase is extremely promoter specific and only transcribes DNA downstream of a

T7 promoter sequence. T7 RNA polymerase can either be obtained commercially or

produced in-house by overexpressing the T7 RNA polymerase gene in E.coli. High

transcriptional efficiency is obtained when the DNA template is prepared from chemically

synthesised double-stranded DNA, which consists of an 18 nucleotide T7 promoter

55

sequence at the 5ʹ-end. The first nucleotide that must be incorporated in the desired RNA

sequence must be a guanosine. In addition, the transcriptional efficiency is highly

dependent on the first six nucleotides of the desired RNA sequence; these six starting

sequences are generally purine-rich. This significantly restricts the RNA sequences that

can be synthesised efficiently. Another disadvantage of this technique is that by-products

with N±1 nucleotides are produced due to 5ʹ and 3ʹ inhomogeneity.

In enzymatic synthesis, the RNA is synthesised from the 5ʹ to 3ʹ direction. The T7 RNA

polymerase binds to the T7 promoter sequence and unwinds a section of DNA to produce

a single-stranded DNA template. The template strand (3ʹ-5ʹ) is used as a template for RNA

synthesis. As transcription proceeds, the RNA polymerase moves along the template

strand and uses base pair complementarity to create an RNA transcript. The main reaction

in this case is the formation of a phosphodiester bond between the 5ʹ-phosphate group of

one donor molecule and the 3ʹ-hydroxyl group of the acceptor molecule. The enzyme T4

RNA ligase from the T4 bacteriophage catalyses this reaction.

For the transcription reaction, the main reactants that are required include the T7 RNA

polymerase, nucleotide triphosphates (NTPs) and the DNA template. Magnesium ions are

also required to stabilise the DNA template. The conditions for the reaction are very

important, so every effort is made to optimise conditions such as the incubation time,

DNA template concentration, NTP concentration and type of T7 RNA polymerase. After

the transcription reaction, the RNA is purified using both PAGE and various

chromatography techniques.

56

1.3.3 RNA nucleotide structure

The conformation of nucleotides depends on the dihedral angles for rotation around each

bond. In total there are seven dihedral angles that are used to characterise the

conformation of a nucleotide; α, β, γ, δ, ε, ζ which are sugar-phosphate backbone angles,

and the glycosidic (χ) angle (Figure 1.3.4). The glycosidic angle describes the position of a

base with respect to the sugar. In nucleosides, the C1ʹ atom of the sugar is bonded to N1 in

pyrimidines and to N9 in purines, by a glycosidic bond. Two dihedral angle ranges

correspond to two conformations, syn and anti. The glycosidic angle is called anti when χ

= 180 ± 90° and syn when χ = 0 ± 90° (Figure 1.3.6a). The anti conformation is found in

A-DNA, A-RNA and B-DNA and is more stable than the syn conformation.

Ribose is a five-membered ring and so five dihedral angles can be specified for the sugar

moiety. The dihedral angle C4ʹ-O4ʹ-C1ʹ-C2ʹ is labelled ν0 and ν1, ν2, ν3, ν4 continue around

the ring in a clockwise direction (Figure 1.3.4). These dihedral angles can be used to

characterize the two main ribose conformations, C3ʹ-endo and C2ʹ-endo (Figure 1.3.5).

RNA will commonly adopt an A-form helical structure, whereby the sugar conformation

is predominantly C3ʹ-endo.

Figure 1.3.4 Seven dihedral angles, α, β, γ, δ, ε, ζ, χ, revealing the conformation of a nucleotide

and five dihedral angles, ν0, ν1, ν2, ν3, ν4, defining the conformation of the five-membered sugar

ring.

57

Figure 1.3.5 The two main sugar conformations (a) C3ʹ-endo in RNA and (b) C2ʹ-endo in DNA.

The pentose sugar conformation can also be specified in terms of the pseudorotation phase

angle (P).36

The pseudorotation phase angle defines the sugar pucker conformation and

identifies which two atoms of the other three are out of the plane. The relationship

between the phase angle and the endo/exo notations is illustrated in Figure 1.3.6b.

Figure 1.3.6 (a) the relationship between the syn/anti notations and the corresponding dihedral

angle values. (b) The pseudorotation phase cycle of the pentose sugar showing the relationship

between the pseudorotation phase angle (P), and the endo and exo notations.35

58

1.3.4 RNA base pairing and stacking

Two main types of base pairing can occur in RNA, either canonical or non-canonical.

Canonical base pairing is also known as Watson-Crick complementary base pairing,

which involves hydrogen bonding between G.C and A.U bases.37

G.C base pairing

involves three hydrogen bonds (Figure 1.3.7a), while A.U base pairing involves two

hydrogen bonds (Figure 1.3.7b). Watson-Crick base pairing allows the formation of a

right-handed helix of any sequence without any distortions. Conversely, non-canonical

base pairing includes any type of base pairing that is not Watson-Crick type. Wobble base

pairing is a common type of non-canonical base pairing that involves hydrogen bonding

between G.U bases, forming two hydrogen bonds (Figure 1.3.7c).

Figure 1.3.7 Illustration of canonical (a) G.C (b) A.U, and non-canonical (c) G.U, base pairing.

59

Base stacking interactions are caused by the non-covalent interaction of π-orbitals between

the aromatic rings of adjacent bases (Figure 1.3.8), which favour nucleic acid geometry

energetically and are an important contribution to stability in nucleic acids. Stacking

interactions between purine-purine bases are found to be the most stable, followed by

pyrimidine-purine and pyrimidine-pyrimidine bases.38

Therefore, stacking interactions are

highly influenced by the composition and sequence of stem-loop structures. Several non-

covalent forces have been considered that play a role in stabilisation including van der

Waals forces, electrostatic interactions and solvation effects.39

Figure 1.3.8 Illustration of the base stacking interactions between adjacent bases in a base paired

helical stem. The rectangles represent the G, C, A, U bases, the unfilled circles correspond to the

pentose sugar, red circles represent the phosphorus atoms and green triangles symbolise the base

stacking interactions.

60

1.3.5 RNA structure

RNA structure is described in terms of primary, secondary and tertiary structure. The

primary structure is the sequence of nucleotides, which creates the single-stranded RNA.

The secondary structure refers to the folding of the primary structure to form a duplex,

with hydrogen bonds allowing for canonical and non-canonical base pairing. It consists of

various regular and irregular secondary structural elements. The secondary structure

elements are associated via additional hydrogen bonds and van der Waals contacts to form

the tertiary structure. The tertiary structure portrays the complete 3D spatial arrangement

of the RNA.

The secondary structure is determined by the hydrogen bonding from the single-stranded

RNA sequence. This leads to the assembly of several recognisable domains of secondary

structure motifs including hairpin loops, bulges, and RNA junctions.40

Hairpin loops are

unpaired loops found at the end of a double helix stem and are among the most common

structural motifs found in RNA. The hairpin loop is not only a stable component of RNA

secondary structure but it is also an important functional element in many well

characterised RNAs. Hairpin loops can contain a number of unpaired bases, the most

abundant are tetraloops, which have four unpaired bases.41

Bulges are formed from an

excess of residues on one side of the duplex. A bulge can have three significant effects on

RNA tertiary structure; it distorts the stacking of bases next to the bulge and reduces helix

stability, induced a bend in the RNA and increases major groove accessibility. RNA

junctions are a common structural feature in RNA secondary structures that are formed by

three or more helices. They are dynamic structures capable of undergoing large

conformational changes. A classic example of a four-way junction is the secondary

structure of transfer RNA (tRNA) (Figure 1.3.9). The four-way junction is comprised of

the double-stranded acceptor stem and three hairpin loops; the D-loop, T-loop and the

anticodon loop. Four-way junctions are remarkably stable, forming coaxial stacking of

helices and long-range tertiary interactions.42

61

Figure 1.3.9 Illustration of the secondary structure of tRNA. The four-way junction consists of the

acceptor arm and the three hairpin loops, the D-loop, the T-loop and the anticodon loop.

RNA hairpin loops, which contain the GNRA motif, are the most commonly occurring

hairpins in various biologically active eukaryotic RNAs, the most abundant being GAAA

and GUAA. GNRA tetraloops exhibit an unusual stability; they are found to be

substantially more stable, with melting transition temperatures (Tm) more than 4°C higher

than other less frequently occurring sequences.43

The high thermodynamic stability can be

explained by the presence of intramolecular interactions and extensive base stacking.44

The main source of stability in GNRA tetraloops comes from hydrogen bonding. These

hydrogen bonds include the hydrogen bond base pairing between the first and fourth bases

forming a G.A sheared base pair. G.A mismatches are common non-canonical structural

motifs in RNA molecules.45

Hydrogen bonds are formed between (guanine 2-amino and

adenine N7) and between (guanine N3 and adenine 6-amino), as shown in Figure 1.3.10.46

G.A sheared mismatches serve an integral part of the U-turn folding motif observed in

GNRA tetraloops.47

62

Figure 1.3.10 G.A sheared base pairing found in GNRA tetraloop motifs, between the first and

fourth base. Two hydrogen bonds are formed between (guanine 2-amino and adenine N7) and

between (guanine N3 and adenine 6-amino); shown as blue dashed lines.

63

1.4 RNA interactions

1.4.1 Intramolecular interactions

Intramolecular interactions play a significant role in stabilising loop regions in RNA

tertiary structure and contribute to the high thermodynamic stability of hairpin loop

structures. These intramolecular interactions involve base-base, base-sugar, sugar-

phosphate and base-phosphate interactions. It is well known that an extensive network of

hydrogen bonding can exist within loop regions, further contributing to the stabilisation of

the loop tertiary structure. Base-phosphate interactions have been known to increase

stability in RNA structures by reducing intramolecular self-repulsion and promote

compact RNA folding.48

Base-phosphate interactions have been found to be conserved in

hairpin loops, internal loops, junctions and as part of tertiary interactions. A base-

phosphate interaction, which is conserved in most GNRA tetraloops, is found between the

imino and/or amino protons of the first base (G) and the phosphate oxygen atoms of the

fourth base (A). The 2ʹ-OH group is also able to hydrogen bond with highly

electronegative phosphate oxygen atoms.49

1.4.2 RNA and Mg2+

The role of metal ions is very significant in the biological function and activity of RNAs.

In the past decade several excellent articles have reviewed the role of Mg2+

.50-51

Mg2+

ions

are commonly used in ion-RNA interaction studies as they are the most relevant to in vivo

conditions. The ionic radius and the charge of a metal ion are two important properties

which are relevant to ion-RNA interactions. A smaller radius and bigger charge will

increase the charge density of the metal ion and so strengthen its electrostatic interactions

with the RNA structure. The radius and charge of metal ions are also related to hydration

energies. Mg2+

ions have a small radius (~0.65Å) and a charge of [+2], which allows a

tight packing of six water molecules in an octahedral arrangement. This forms the first

hydration layer and the ion can organise a second and possibly third hydration layer,

which contributes to the overall hydration energy.

64

Divalent ions such as Mg2+

in particular have been studied extensively. Mg2+

plays a

substantial role in RNA tertiary structure stabilisation; positive Mg2+

ions reduce the

repulsion of the negatively charged phosphates in RNA causing stabilisation.52

The

addition of Mg2+

can significantly enhance the thermodynamic stability of RNA motifs,

possibly by non-specific interactions with the phosphate backbone.53

As well as stabilising

the secondary structure they also promote RNA folding and alter RNA dynamics.54

The

folding of RNA tertiary structure increases the charge density and negative electrostatic

potential, which causes a dramatic increase in the number of cations pulled to the RNA

surface.55

This involves ion-RNA interaction whereby Mg2+

ions act as ‘diffuse ions’,

‘water-positioned ions’ or ‘chelated ions’ (Figure 1.4.1).56

Figure 1.4.1 Potential Mg2+

-ion interaction with the RNA phosphate backbone. The diagram

illustrates the Mg2+

ions (green circles), water molecules (red circles with attached blue circles)

and phosphate oxygens (lone red circles). Ion interactions can involve specific interactions with

RNA whereby the Mg2+

ions act as (a) chelated ions, (b) water-positioned ions and (c) diffuse ions.

Diffuse ions are largely hydrated forming a hexahydrate Mg(H2O)62+

complex, with first

and partial second hydration layers. Although there is no contact between the ion and the

RNA surface, ion-RNA interactions are mediated by electrostatic interactions between the

ionic charge of Mg2+

and the RNA electrostatic field. Water-positioned ions hold a single

hydration layer between themselves and the RNA surface. Consequently, the close

proximity of the ion to the RNA surface directly influences the positioning of the hydrated

65

ion. Electrostatic interactions between the ion and RNA surface and also the perturbation

of ion hydration layers have to be considered here in terms of energetic factors. Chelated

ions directly interact with specific sites on the RNA surface, held together by electrostatic

forces. However, the free energy required to partially dehydrate a fully hydrated ion

becomes a major energetic factor especially for metal ions such as Mg2+

with high

hydration energies.57

An essential ingredient for IRES function is the dependence on Mg2+

, which has been

proposed to enhance thermodynamic stability and promote RNA folding.53

Mg2+

ions are

suggested to play a significant role in the stabilisation and folding of RNA structures,

possibly by enhancing the electrostatic interactions with the phosphate backbone. UV

absorbance experiments confirmed the improved stability arising from Mg2+

binding with

a dramatic increase in melting temperature of the FMDV 16mer RNA (ΔTm 70°C –

75.5°C).58

It has also been established that Mg2+

ions play an important role in mediating

RNA-RNA interactions.59

1.4.3 RNA-RNA interactions

RNAs follow a hierarchical folding pathway whereby the secondary structure elements

such as helices, hairpin loops and bulges form first, followed by the final tertiary structure

of the RNA molecule. Long-range tertiary interactions in biologically active RNAs are

known to mediate RNA tertiary folding by the binding of GNRA tetraloops to specific

receptor sites.60

These RNA-RNA interactions include A-minor motifs, ribose zippers, and

A-platforms.61

A-minor motifs provide the majority of tertiary interactions used for

interhelical packing in complex RNAs studied to date.62

The tertiary contacts formed by

GNRA tetraloops, consist of A-minor interactions involving tandem adenosine docking

into the minor groove of an RNA helix.63

These A-minor interactions can involve adenosines, which are unpaired or part of non-

canonical G.A sheared base pairing; both can be found in GNRA tetraloops. The high

frequency and conservation of GNRA tetraloops emphasises the importance of long-range

66

interactions, which stabilise RNA tertiary structure.64

In GNRA tetraloops, the adenosine

nucleotide forms extensive hydrogen bond networks to both nucleotides in a Watson-

Crick base pair. Two types of A-minor interaction can be found: Type I and Type II

(Figure 1.4.2). Type I is the most common interaction, in which both the 2ʹ-OH, the N3

and N1 groups of the adenosine lie between the 2ʹ-OH groups of the corresponding base

pair receptor. GNRA tetraloops that have two adenosines in the 3rd

and 4th

position have

shown to form A-minor interactions, whereby Type I and Type II base triples form

interactions between the two adenosines and two C.G base pairs in the receptor helix.65

Figure 1.4.2 Two main types of A-minor motifs (left) Type I and (right) Type II. The blue broken

lines represent hydrogen bonding within C.G base pairing and red broken lines represent hydrogen

bonding in A-minor interactions.

A possible receptor site for the GUAA tetraloop in the FMDV IRES has been identified as

the 15mer RNA located in domain 3. Mutational and structural analysis has provided

significant evidence to propose that the 15mer RNA is a strong candidate to be a

functional receptor of the GNRA motif.10

This provides an ideal model for studying RNA-

RNA interactions in the FMDV IRES.

67

1.5 Principles of NMR spectroscopy66

1.5.1 Basic theory of NMR

Nuclear magnetic resonance (NMR) spectroscopy is a powerful analytical technique that

is used to solve complex structural problems in biological chemistry. NMR involves

perturbing magnetic nuclei and detecting perturbations in the spins of the nuclei in the

sample. This section will focus on the main principles of NMR spectroscopy.

The nuclear magnetic resonance phenomenon lies in the magnetic properties of the atomic

nucleus and forms the basis of NMR spectroscopy. The spin of a nucleus is a quantum

mechanical property, which gives rise to its magnetic properties. The spin of a nucleus is

characterised by the spin quantum number (I). The interaction of the nuclear magnetic

moment (µ) with the external magnetic field (B0), leads to a split in the energy of the spin

states. The total number of possible energy levels can be given by the 2I+1 rule.

Consequently, if we consider nuclei such as 1H,

13C or

31P with spin of ½, according to the

2I+1 rule there are two possible spin states.

Due to Boltzmann distribution, there is a slight excess of spin populations in the lower

energy state than the higher energy state. The population excess in the lower energy state

determines the probability of a transition of spins in the lower energy state to the higher

energy state. To stimulate the transition of spins from the lower energy state to the higher

energy state, a radiation of frequency which exactly matches the energy gap is required.

This is called the resonance condition. A radiofrequency (RF) pulse can be used to

stimulate the transition of spin populations between the two energy levels.

The absorption of energy can be detected as an emission signal, which is usually referred

to as the resonance signal. Since the energy gap between spin states of every nucleus will

be different, each nucleus will acquire a specific resonance frequency, which can be

observed as resonance peaks in an NMR spectrum.

68

1.5.2 Chemical shift, coupling constant and linewidth

The chemical shift describes the dependence of nuclear magnetic energy levels on the

electronic environment. Therefore, resonance signals observed in NMR spectra arise due

to nuclei in different chemical environments. Electrons within a molecule oppose the

external magnetic field (B0) and thereby shield the nuclei. Shielding of a nucleus increases

with the number of electrons. Consequently, nuclei with different electron densities will

be shielded to different strengths, therefore nuclei will precess at different frequencies and

give separate resonance signals in the NMR spectrum. The variations in resonance

frequencies of the same type of nucleus can be seen as chemical shifts in NMR spectra.

Chemical shift (δ) is given with respect to a reference frequency, measured in parts per

million (ppm).67

Scalar coupling (J) arises due to the interaction between spins of two non-equivalent

neighbouring nuclei. Coupling of two spins is influenced by bonded molecular electrons

on the localised magnetic field produced by the two nuclei. Scalar couplings are observed

through a small number of chemical bonds and so its value, measured in Hz, is highly

characteristic of these chemical bonds. The size of the scalar couplings is the coupling

constant, which can give information on the nature of the bond. Three bond couplings are

very useful in providing dihedral angle information of the bonds, using the Karplus

equation.68

The linewidth (ω½) of resonance signals is of paramount importance in NMR

spectroscopy as peaks have to be clearly resolved in order to assign them more accurately.

The separation between two peaks relative to their linewidth will determine the degree to

which they are resolved. The linewidth is the width of the peak measured at half the peak

height. In high-resolution NMR spectroscopy, the linewidth must be minimised to produce

well resolved peaks.

69

1.5.3 Nuclear relaxation

Nuclear relaxation is a process by which nuclei lose their excess energy so that they can

return to thermal equilibrium. The energy absorbed by nuclei from the RF pulse will

excite the spin populations in the lower energy state to a higher energy state. Over time

these excited spins will lose their excess thermal energy so that the magnetisation will

return to its equilibrium alignment with the external magnetic field (B0). The spins will

exchange energy with their surroundings by two relaxation processes, spin-lattice

relaxation and spin-spin relaxation.

The process by which longitudinal magnetisation (Mz) is restored to its Boltzmann

equilibrium along the z-axis is called spin-lattice relaxation or longitudinal relaxation

(Figure 1.5.1a). This relaxation process is time-dependent and is measured by the spin-

lattice relaxation time (T1), whereby the rate constant is the reciprocal of T1. The process

by which transverse magnetisation (Mxy) decays to the equilibrium value of zero is called

spin-spin relaxation or transverse relaxation (Figure 1.5.1b). This relaxation process is

time-dependent and is measured by the spin-spin relaxation time (T2). T2 is measured as

the time required for the transverse magnetisation (Mxy) to drop to 37% of its original

magnitude.

Figure 1.5.1 Scheme illustrating (a) Spin-lattice (T1) relaxation where magnetisation relaxes back

to longitudinal axis; (b) Spin-spin (T2) relaxation where the spins precess in the x,y plane and fan

out as they lose coherence.

70

1.5.3.1 Spin-lattice relaxation

During T1 relaxation, energy is transferred from the spins to the surrounding environment

called the lattice. As the surroundings are at thermal equilibrium, there is a higher

probability that energy will move into the surroundings than there is for the same amount

of energy to flow out of the surroundings. Energy is exchanged through dipolar interaction

of two nuclear magnetic dipoles, which causes fluctuations in local magnetic fields. This

provides an important means of relaxation through the dipolar mechanism. The

combination of local magnetic fields and molecular motion is what allows the nuclei to

exchange energy with the environment. The magnitude of the T1 relaxation time depends

upon several factors including the type of nuclei, size of the molecule and the temperature.

T1 relaxation is normally shorter for nuclei with high gyromagnetic ratios, so in liquids the

T1 for protons is in the order of 10-2

seconds while the T1 for carbons is much longer at

about 102 seconds. Apart from the type of nuclei, the other factors mentioned above are all

related to the motion of the molecule, which can be measured by the rotational correlation

time (τc). The rotational correlation time is the time required for a molecule to rotate by

one radian, which is normally in the range of picoseconds (ps) to milliseconds (ms). Since

T1 is dependent upon molecular motion, nuclei in solids have an extremely long T1 values

compared to the same nuclei in liquids. Smaller molecules that have short correlation

times will tumble faster and so will have a long T1 compared to larger molecules such as

DNA, RNA and proteins (Figure 1.5.2). Changing the temperature of the sample will also

change the correlation time of a molecule.

1.5.3.2 Spin-spin relaxation

T2 relaxation is based on the exchange of energy between spins and is also governed

largely by the dipolar mechanism. The transition between spin states of a nucleus changes

the local magnetic field at nearby nuclei, which allows the exchange of energy between

the spins of nuclei. The total energy exchanged between the spins in a spin system does

not change, but the lifetime of the spin states is shortened by this process and so provides

a means of relaxation. The major contributor to the decay of transverse magnetisation

71

(Mxy) is due to the magnetic field inhomogeneity. The spins of individual nuclei are

exposed to different local magnetic fields, which results in a spread of their individual

Larmor frequencies. Since transverse magnetisation exists due to coherence of the Larmor

frequency of individual spins, the decay of transverse magnetisation is largely governed

by this process. Although strictly speaking this is not a relaxation process, it is measured

by the effective transverse relaxation time, denoted (T2*).

T2* is of great practical importance as it determines the spectral resolution of the NMR

spectrum. The spectral resolution is measured by the linewidth of a resonance signal at

half-height. Shorter T2* values will cause excess line broadening, which can be described

by the following equation:

ω½ = 1/πT2* Equation 1.5.1

whereby (ω½) is the linewidth of the resonance signal at half-height; (ω½) is measured in

Hz if the T2* value is measured in seconds.

Figure 1.5.2 A plot of T1 (blue curve) and T2 (red curve) as a function of correlation time (τc).

Small molecules have a correlation time in the 10-12

to 10-10

second range while large molecules

are above 10-9

seconds.

72

1.5.4 Nuclear Overhauser Effect (NOE)

In contrast to scalar coupling, dipolar couplings are mediated through space rather than

through chemical bonds. Close proximity of nuclei can be detected through dipolar

couplings due to the Nuclear Overhauser Effect (NOE).69

In a two-spin system of nucleus

HA and HX, the irradiation of one proton (HA) can increase the intensity of the

neighbouring proton (HX).70

Normally, transfer of spin polarisation from one population to

another occurs via single quantum transitions (W1). However, dipole-dipole NOE

couplings arise from cross-relaxation via W0 (zero quantum) and W2 (double quantum)

transitions. Single quantum transitions can be excited by electromagnetic radiation such as

the radiofrequency (RF) pulse, but W0 or W2 transitions cannot. Both W0 and W2

transitions are only allowed in relaxation, therefore, the NOE is dependent on the process

of relaxation. In a two-spin system, four different energy states exist (Figure 1.5.3).

Figure 1.5.3 Energy level diagram of a two spin system illustrating four energy levels

(N1>N2>N3>N4) with their corresponding α/β spin states. WA and WX represent single quantum

transition in HA and HX, respectively. W0 and W2 represent zero and double quantum transitions,

respectively.

73

If we consider the two protons, HA and HX, irradiation of nucleus HA will excite the spins

of HA leading to WA transition. The population of spins in all four energy states can be

described as N1>N2>N3>N4 before the saturation of nucleus HA (Figure 1.5.4a). After

saturation, the population of spins in nucleus HA move from N1-N3 and N2-N4 until the

population of spins in the N1/N3 and N2/N4 energy states are evenly distributed (Figure

1.5.4b). The population difference for nucleus HX still remains the same. However, the

system must be restored to equilibrium by restoring population distribution, which can

only be achieved via WX, W0 and W2 transitions. This cross relaxation causes a change in

population difference of nucleus HX that leads to change in the intensity of nucleus HX.

Cross relaxation via W0 mechanism leads to a decrease in intensity of proton X.

Conversely, cross relaxation via W2 mechanism leads to an increase in intensity of proton

X. However, these two mechanisms function simultaneously, in addition to competing

with WX. Therefore, NOE dipolar coupling depends upon the relative rates at which these

three (WX, W0, W2) mechanisms occur.

Figure 1.5.4 Scheme illustrating the NOE effect in a two spin system; (a) spin populations before

saturation of nucleus HA, (b) spin populations after saturation of nucleus HA.

74

1.5.5 NMR of rate processes

1.5.5.1 Base pair kinetics

The kinetics of base pair opening and imino proton exchange represents a probe for the

dynamic motions of base pairs in nucleic acids. NMR spectroscopy can be used to

measure base pair kinetics and imino proton exchange rates, which can provide additional

information to complement the structural studies of nucleic acids and aid in the

investigation of RNA-Mg2+

and RNA-RNA interactions.

Base pair kinetics is interpreted by a two-state model (open/closed) whereby the base pair

opens and closes, but imino proton exchange only occurs in the open state; in the closed

state the imino protons are protected in Watson-Crick base pairing.71

Once it is in the open

state, the imino proton can exchange via an acid-base catalysed reaction with proton

acceptors. The imino proton exchange rate induced by a proton acceptor is:

Equation 1.5.2

where kcoll is the collision rate, [acc] is the proton acceptor concentration, and the ΔpK is

the pKa difference between the imino proton (pKG = 9.3, pKU = 9.2) and acceptor.72

The

collision rate is typically in the range of 108 to 10

9 s

-1. A successful collision is needed for

proton exchange. When the pK of the acceptor is larger than that of the donor, the rate of

exchange is efficient, although this is dependent on the proton acceptor concentration as

shown in the above equation. When the pK of the acceptor is much less than that of the

donor, as in imino proton exchange with water (pK of H3O+ is -1.7), the rate of exchange

is less efficient.

However, imino proton exchange in base pairs is a two-step process requiring base pair

opening, followed by a transfer to a proton acceptor such as water or a catalyst, eg. NH3.

The imino proton exchange time induced by a proton acceptor is:

75

Equation 1.5.3

where τ0 is the base pair lifetime, Kdiss is the base pair dissociation constant and kex,acc is

the imino proton exchange rate in the open state. The base pair lifetime is a measure of the

minimum time required for the base pair to open. If the imino proton exchange occurs at

every opening event, then the exchange time (τex) is equal to the base pair lifetime (τ0). In

RNA duplexes, base pair lifetimes can range from 2ms to 50ms and 0.1 to 2ms at 15°C for

d(G.C) and d(A.T) base pairs, respectively.72

Base pair lifetimes can vary for G.C and A.T

base pairs within an RNA sequence, depending on base pair stability and exposure to

solvent. Imino proton exchange rates can vary from 1.0s-1

to 40.0s-1

for guanine and uracil

imino protons at 35°C.73

1.5.5.2 Chemical exchange

Chemical exchange processes are significant in NMR spectroscopy as they can provide

information on dynamic processes. The linewidth and line intensity of NMR resonance

signals are sensitive to chemical exchange processes, so allow the means to study

dynamics in NMR. Here, the concept of exchange regimes will be introduced for

intramolecular exchange.

If a molecule is in equilibrium between two conformations, A and B, then the individual

nuclei concerned will experience two different chemical environments. Since the chemical

shift of nuclei depends upon their chemical environment, the effect on peak positions

depends on the rates of interconversion, kA and kB, and the magnitude of the difference in

chemical shifts. The rates of interconversion and the equilibrium are concentration

independent as the conformational changes occur within the same molecule.

If the interconversion rate is slow on the NMR timescale, whereby kA and kB << Δυ (Δυ is

the difference in chemical shift measured in Hz), then two separate peaks are observed.

This type of exchange regime is called slow exchange. The two peaks correspond to the

two different conformations of A and B and the relative intensities of the two peaks

76

depend on the molar concentrations of conformation A and B, at equilibrium. If the

interconversion rate increases, whereby kA and kB ~ Δυ, then the effects on the chemical

shifts are intermediate and this is called intermediate exchange. During this exchange

regime, there is a partial averaging of the two chemical shifts, which leads to a broadening

of both peaks. When the interconversion rate is fast on the NMR timescale, whereby kA

and kB >> Δυ, the effects of the chemical shifts are completely averaged and so only one

peak is observed. Since this chemical shift is an average of the A and B chemical shifts,

the single peak observed will appear between the positions corresponding to the A and B

peaks. The exact position of the peak will depend upon the ratio between the

concentrations of the two conformations. The three exchange regimes described here are

illustrated in Figure 1.5.5.

Figure 1.5.5 Example of chemical exchange between two conformations of the same nucleus. The

nucleus can change magnetic environments between the two conformations in fast, intermediate

and slow exchange regimes, on the NMR timescale.

77

1.5.6 1D 19

F-NMR and 31

P-NMR

The most commonly used 1D heteronuclear NMR experiments, apart from 13

C-NMR,

include 19

F-NMR and 31

P-NMR. The 19

F nucleus has a spin of ½, a similar gyromagnetic

ratio to 1H and a natural abundance of 100%, which gives it a relative sensitivity of 0.83

(when 1H is 1.00). One particular advantage of

19F-NMR spectroscopy is the large

chemical shift range (>1200ppm), which is much greater than for protons (~15ppm).74

This large range is attributed to the dominance of paramagnetic shielding, as opposed to

diamagnetic shielding in 1H-NMR, which is responsible for the wide variation of

19F

chemical shifts. The 19

F nucleus is easily accessible by NMR spectroscopy, but its

application has been less important because fluorine compounds are rarely found in nature.

However, 19

F-NMR has become increasingly more important in studying biological

systems, whereby 19

F analogues of biologically important molecules can be obtained.

Recently, 19

F-NMR has been exploited for structure analysis, binding studies and

metabolic studies.75 76

In nucleic acids, 5-fluorouridine (5-FU) substitutions are largely

used to incorporate 19

F labels in DNA or RNA sequences. Therefore, 19

F-NMR of

fluorinated DNA/RNA samples provides a valuable tool for probing nucleic acid

secondary structure, dynamics and interactions. 19

F chemical shifts of 5-FU pyrimidines

range between -90.0 to -85.0ppm and -169ppm to -164ppm when referenced to

Trifluoroacetic acid (CF3COOH) and Trichlorofluoromethane (CFCl3), respectively.

The 31

P nucleus also has a spin of ½ and a natural abundance of 100%. However, due to

its low gyromagnetic ratio, the relative sensitivity is only 6.6% when compared to 1H,

which can seriously limit the use of 31

P-NMR. The range of chemical shifts for

phosphorus is also very large (~4000ppm). However, in contrast to fluorine, phosphorus

has been studied more thoroughly by NMR spectroscopy as it is found in the phosphate

backbone of nucleic acids. 31

P chemical shifts are very sensitive to changes in the local

environment due to the large anisotropic distribution of electrons.77

Therefore, 31

P-NMR

spectroscopy is an excellent method to probe the backbone conformation of nucleic acid

structures. 31

P chemical shifts of nucleotides range between 4.0ppm and -6.0ppm, when

referenced to phosphoric acid.

78

1.5.7 Two-dimensional (2D) NMR spectroscopy

In two-dimensional (2D) NMR, scalar and/or dipolar couplings can be observed between

nuclei in a 2D spectrum, through a collection of several FID (free induction decay) signals.

The main difference between 1D and 2D experiments is that the FID signal detected is

recorded as a function of two time variables, t1 and t2.78

However, the FID signal is only

recorded during t2 and not t1. The advantage of 2D experiments over 1D experiments is

that heavily overlapped signals can be spread into the second dimension, thereby resolving

peaks and cross diagonal peaks that provide important coupling information. The basic

pulse sequence consists of two pulses or group of pulses that are separated by the

preparation time, evolution time (t1) and the acquisition time (t2) (Figure 1.5.6).

Figure 1.5.6 The basic pulse sequence of a 2D NMR experiment. The evolution time (t1) is the

period between the two 90° pulses whereby T1 and T2 relaxation occurs. The acquisition time (t2)

begins immediately after the last 90° pulse.

The evolution time (t1) is systematically increased in increments (t1 + Δt) after each new

pulse sequence. During this variable time period, the 90° pulse flips the magnetisation to

the transverse plane. This is directly followed by a second 90° pulse (mixing pulse) that

causes mixing of spin states and transfer of magnetisation between coupled signals. The

FID signal is produced and the experiment is repeated after the relaxation delay.

Consequently, several FIDs are collected during the acquisition time (t2), depending on the

number of increments completed in variable delay t1. Two Fourier transformations are

required for conversion to two frequency functions, F1 and F2.

79

1.5.8 Three-dimensional (3D) NMR spectroscopy

In three-dimensional (3D) NMR, a third frequency domain is introduced to the standard

2D NMR pulse sequence. The FID signal is recorded in the t3 time domain as a function of

two variable times, t2 and t1. The advantage over 2D NMR is that heavily overlapped

signals found in larger RNAs can be distributed into the third dimension, thereby

resolving peaks that provide important coupling information. The basic 3D pulse sequence

is similar to the 2D pulse sequence with an additional evolution period or mixing period

before detection (Figure 1.5.7). There are various examples of 3D NMR experiments but

here a homonuclear 3D NOESY-TOCSY pulse sequence is illustrated.79

Figure 1.5.7 The pulse sequence for a 3D NOESY-TOCSY experiment, constructed from a

combination of the NOESY and TOCSY pulse sequences.

The first t1 evolution period and τm mixing time coincides with the NOESY pulse

sequence. The second t2 evolution period and the isotropic mixing correspond to the

TOCSY part of the pulse sequence. The two evolution periods t1 and t2 are independently

incremented. Three Fourier transformations are required for conversion to three frequency

functions; F1, F2 and F3. In this experiment magnetisation is transferred from spin A to

spin B during the NOE mixing period and subsequently magnetisation from spin B is

transferred to spin C during the TOCSY mixing period. This is an example whereby both

dipolar and scalar couplings can be observed to yield information on both intranucleotide

and internucleotide connectivities.

80

1.6 Principles of molecular modelling

1.6.1 Molecular mechanics (MM)

Molecular mechanics (MM) is a method referring to the application of classical mechanics

in modelling molecular systems. It describes a molecule as a collection of atoms that

interact with each other based on equations of classical mechanics. Molecules are treated

as balls on springs to describe bonded interactions and individual atoms are treated as

single particles.80

The potential energy (Ep) of a molecular system is calculated using a force field, which are

functions and parameter sets derived from experimental data and quantum mechanical

calculations. The functional form of a force field can be described in (Equation 1.6.1),

although the functional form can vary in different force fields.

Equation 1.6.1

The force field will attempt to calculate the potential energy by defining the molecule in

terms of the energy between covalent bonds (Ebond), the energy between bond angles (Ebond

angle), planarity in aromatic rings (Eimproper), the energy of non-bonded interactions of atoms

separated by three bonds (Edihedral), the energy due to van der Waals interaction (EvdW) and

the energy due to electrostatic interactions (Eelectrostatic).

In addition to the functional form, force fields define a set of parameters for each type of

atom in the molecular system. These usually include values for atomic mass, atomic

charge, bond lengths, bond angles and bond dihedral angles. Force field calculations are

based on numerous approximations. These approximations will be derived from all

different types of experiments, hence they are called empirical force fields (EFF).

Different force fields are designed for different molecular systems, depending on the size

81

and the type of molecules involved. The most popular force fields include CHARMM81

(Chemistry at HARvard Macromolecular Mechanics) and AMBER82

(Assisted Model

Building and Energy Refinement), both developed for nucleic acids and proteins.

1.6.2 Energy minimisation

The geometry of a molecule determines many of its chemical and physical properties,

therefore calculating accurate geometries of a molecular system is very important. Stable

structures best define the accurate geometry of a molecule, therefore computational

methods are employed to explore and find the most stable structures.

Energy minimisation is a method which is used to find the best atomic arrangement of a

structure that leads to a more stable structure. The stability of a structure is measured by

its energy; the lower the energy the more stable the molecule. Therefore, energy

minimisation involves exploring various possibilities to find a structure with the lowest

energy value. This is achieved by creating a potential energy surface (PES). A PES

mathematically describes the relationship between the different molecular geometries and

their corresponding single point energies. Energy minimisation is used to find minima on

the potential energy surface, which involves adjusting the structural geometry of a

molecule in order to reduce the energy of the conformation.

Various algorithms have been designed to find the lowest energy conformation of a

molecular system. These include calculus based minimisation methods, genetic algorithms,

Monte Carlo methods, quantum mechanical and molecular mechanical optimisation

methods, molecular dynamics and simulated annealing methods. Several factors need to

be taken into account in order to identify the most appropriate algorithm(s) for the given

problem. In the case of modelling nucleic acids and proteins with respect to energy

minimisation, molecular mechanical methods are most commonly employed.

82

1.6.3 Simulated annealing and molecular dynamics (MD)

Simulated annealing heats the initial structure to a very high temperature. This provides

energy to atoms, which allows them to become unstuck from their original positions and

move randomly through conformational space. Subsequently, the temperature is reduced

slowly in order to increase the chance of finding the lowest energy conformations.

Molecular dynamics (MD) simulations can provide information about the time-dependent

motion of molecular systems. A molecular dynamics simulation allows molecules to

explore conformational space in the region defined by atomic positions and velocities.

Unlike energy minimisation calculations, the MD calculation accounts for thermal motion,

providing thermal energy to molecules so they can cross potential energy barriers.

Molecular dynamics is a very important tool used to study the thermodynamic properties

and dynamics of molecules. MD simulations require a force field, which calculates the

potential energy of the system and describes the terms by which the particles in the

simulation will interact. MD involves providing kinetic energy to the PES and the

subsequent motion of the molecular system over the PES. These motions follow the laws

of classical mechanics, but for more detailed simulations, quantum mechanics can also be

employed. MD simulations calculate the future positions and velocities of atoms based on

their current positions and velocities.

A variation of molecular dynamics is restrained molecular dynamics (rMD), which

incorporates experimental data to the molecular dynamics simulation. This type of

molecular dynamics has been used to generate the NMR solution structures reported in

this thesis. The experimental data in this case consists of distance restraints derived from

NOESY data.

One disadvantage of molecular dynamics simulations is that the timescale of simulations

are much shorter than chemical processes and most physical processes, which occur in

nanoseconds or longer. Therefore, long-term processes cannot be studied as they would

take too much computational time.

83

1.7 Previous work

As part of my MSc project (University of Manchester, 2007), I carried out a research

project. The structure and Mg2+

binding properties of the conserved 16mer RNA, located

in domain 3 of the FMDV IRES, were investigated by NMR spectroscopy and molecular

modelling. Various 1D and 2D NMR experiments were performed on the 16mer apo-RNA

in 1H2O. However, the full assignment and structure determination of the 16mer apo-RNA

are now fully reported in chapter 3 of this thesis.

Molecular dynamics simulations were used to investigate the effect of Mg2+

on the FMDV

16mer RNA structure, using HyperChem software. The presence of Mg2+

alone created a

compact structure, which did not conform to the standard helical structure of A-form RNA.

Interestingly, in the presence of both water and Mg2+

, a good quality structure was

produced. The addition of Mg2+

to the 16mer RNA in water resulted in the stabilisation of

the entire 16mer RNA structure. Furthermore, the addition of excess Mg2+

around the

GNRA tetraloop revealed shorter hydrogen bond distances between the G.A sheared base

pair. These results provided preliminary evidence for the effect of Mg2+

on the 16mer

RNA structure.

Molecular modelling methods were also applied to investigate the possible RNA-RNA

interaction between the FMDV 16mer RNA and the 15mer RNA. It was hypothesised that

the two adenosine nucleotides in the GUAA tetraloop of the 16mer RNA, form A-minor

interactions with the two target G.C base pairs (G231.C241 and C232.G240) found in the

15mer RNA. It was found that the A181 nucleotide in the tetraloop could form four

possible hydrogen bonds with its target G.C base pair (G231.C241), while the A180

nucleotide could only form two possible hydrogen bonds with its target G.C base pair

(G232.C240).

84

1.8 Aim of the project

The principal aim of the project is to elucidate the three dimensional structure of the

conserved RNA motifs in the FMDV IRES and study their kinetics and interactions. For

NMR investigation, two highly conserved RNA motifs have been chosen for NMR

structure determination: the 16mer RNA and the 15mer RNA, located in the apical region

of domain 3 of the FMDV IRES. The results obtained will help us to gain a better

comprehension on the three-dimensional nature of RNA structures, which are complex

and not understood at this stage.

Mg2+

plays a vital role in the stabilisation of RNA structures and the 16mer RNA was

selected to investigate the effect of Mg2+

on RNA structure and kinetics, using NMR

spectroscopy. The aim is to solve the three-dimensional structure of the 16mer RNA in the

absence of Mg2+

and then to titrate Mg2+

to study effect on 16mer RNA tertiary structure

conformation. Subsequently, the three-dimensional structure of the 16mer RNA in the

presence of Mg2+

will be solved. Additionally, the imino proton exchange rates of the


RNA complex will be measured to quantify the effect of Mg2+

.

To allow the folding of RNA structures into complex three-dimensional structures, long-

range RNA-RNA interactions are essential for folding and function of biologically active

RNAs. However, the molecular details of these interactions are largely unknown. We

believe that the GNRA tetraloop of the 16mer RNA is involved in long-range tertiary

contacts with a conserved 15mer RNA. The objective here is to firstly solve the three-

dimensional structure of the 15mer RNA. Subsequently, the main goal is to merge the

16mer and 15mer RNAs and study the RNA-RNA complex using NMR methods. This

will provide us with any evidence of an interaction involving these two RNAs and in the

larger context it will enhance our knowledge of interactions between RNAs.

Fluorine substituted RNAs can be effectively used to study the effect of Mg2+

on RNA

structure as well as the interaction between two RNAs. The 5-FU 16mer RNA was used to

investigate the effect of Mg2+

. Subsequently, the 5-FU 15mer RNA was used to make an

85

RNA-RNA complex of two fluorinated RNA samples. The purpose was to provide any

additional evidence that could support the results found in chapters 3 and 4. Additionally,

this was a great opportunity to study fluorine substituted RNAs by NMR spectroscopy and

develop the scope for 19

F-NMR studies.

86

Chapter 2: Materials and Methods

Chapter 2 highlights all the relevant materials required to repeat the study as well as

methods used to prepare NMR samples, implement NMR experiments, process and

analyse NMR data, and finally employ structure determination protocols. Additionally, a

section has been devoted to the quantification of imino proton exchange rates.

Assistance in setting up the NMR experiments was provided by the scientific staff at

NIMR (London, U.K.) and CRMN (Lyon, France) and technical staff at the University of

Manchester. However, all the data processing, analysis, interpretation and structure

calculations were carried out by myself.

2.1 RNA sample preparation for NMR studies

Five main samples were prepared for NMR investigation. This included the FMDV

16mer-gGUAAc-tetraloop RNA sample (batches 1 and 2) and the 15mer-cAACCCCAg-

heptaloop RNA sample, with sequence 5ʹ-UCCUUG(GUAA)CAAGGA-3ʹ and 5ʹ-

GUGC(AACCCCA)GCAC-3ʹ, respectively. In addition, two fluorinated samples were

also prepared; 5-fluorouracil (5-FU) 16mer and 15mer RNAs with fluorination at the H5

position of U179 and U230 bases, respectively. These five samples were purchased from

Metabion international AG (Martinsried, Germany), synthesised chemically with HPLC

purification and used without further purification.

RNA samples from Metabion were prepared for NMR by dissolving the RNA sample in

filtered Q-water. After lyophilisation of the sample, 400μl of 20mM NaCl and 10mM

H3PO4 was added. The sample was annealed by gently heating to 80C followed by slow

cooling. NMR samples were prepared by addition of 90% filtered Q-water (1H2O) and

10% deuterated water (2H2O) or ~100% deuterated water, to a 5mm NMR tube, with a

total volume of 600μl. Samples that were reconstituted from 1H2O to

2H2O were

lyophilised extensively to remove any trace of water from the sample. The final

concentrations of the 16mer (batch 1), 16mer (batch 2), 15mer, 5-FU 16mer and 5-FU

87

15mer RNA samples were 1.21mM, 0.45mM, 0.75mM, 0.2mM and 0.16mM, respectively,

in 600μl solution.

Calculation of RNA concentrations was based on UV absorbance measurements at 260nm

(A260). For UV absorbance measurements, the diluted RNA samples were placed in a

clean sterile cuvette with another cuvette containing 1000μl filtered Q-water prepared as a

‘blank’. The baseline was zeroed using another blank in the sample chamber before the

A260 measurement was made. RNA absorbs at 260nm producing a sharp peak observable

in the spectrum. The absorbance at this wavelength was measured and used to calculate

the concentration of the sample using the molecular weight of the RNA and also the

calculated (µg/OD260) value. These values were obtained using an oligonucleotide

calculator found online at www.ribotask.com.

A 64mM Mg2+

stock solution (1.0ml) was made in 1H2O, based on the concentration of the

16mer RNA (batch 1). The 16mer RNA sample (batch 1) was titrated with Mg2+

.

Titrations were performed by four separate additions of Mg2+

, with concentrations of

0.605mM (0.5eq), 1.21mM (1.0eq), 2.42mM (2.0eq) and 6.05mM (5.0eq). A total of

100μl of Mg2+

was added to the sample, which was subsequently lyophilised and

constituted to 600μl of 1H2O. The 16mer apo-RNA and Mg

2+ RNA complex (batch 1) was

used for structure determination and monitoring the effect of Mg2+

. For the 16mer RNA

sample (batch 2), a 35μl addition of Mg2+

was made corresponding to 5.0eq. This sample

was used for measuring the imino proton exchange rates and also for producing the

16mer/15mer RNA-RNA complex. An addition of 59μl of Mg2+

was made to the 15mer

RNA sample, corresponding to 5.0eq. Subsequently, the 16mer/15mer RNA-RNA

complex was formed by addition of 437μl of the 15mer RNA to the 16mer RNA (batch 2),

producing a 0.45mM RNA-RNA complex. The remaining 15mer Mg2+

RNA sample was

made up to 600μl producing a concentration of 0.23mM.

A second 76.7mM Mg2+

stock solution (1.0ml) was made in 2H2O. A volume of 13μl and

10.5μl of this stock solution was added to the 5-FU 16mer and 5-FU 15mer RNA samples,

respectively, corresponding to 5.0eq of Mg2+

. The fluorinated 16mer/15mer RNA-RNA

http://www.ribotask.com/

88

complex was formed by addition of 225μl of the 5-FU 15mer to a 0.06mM 5-FU 16mer

RNA sample. The sample was lyophilised and constituted into 600μl of 2H2O, producing a

0.06mM RNA-RNA complex of the two fluorinated RNAs.

2.2 NMR spectroscopy

2.2.1 NMR spectrometers

NMR experiments were acquired using Bruker Avance 600/700 MHz spectrometers and

Varian Inova 600/800 MHz spectrometers at NIMR (National Institute for Medical

Research) in Mill Hill, London. Access was also given to the latest Bruker Avance 1GHz

spectrometer at the Centre de RMN à Très Hauts Champs (CRMN) in Lyon, France. Our

research group was one of the first groups in the world to gain access to the 1GHz

spectrometer. Furthermore, a Bruker 400 MHz spectrometer was also used at the

University of Manchester, School of Chemistry.

Only the Bruker 600 MHz, 700 MHz and 1GHz spectrometers were equipped with

cryogenically cooled probes. All the spectrometers, except for the Bruker 400 MHz, were

equipped with triple resonance (1H,

13C,

15N) probes. The Varian 600 MHz spectrometer

was equipped with probes capable of detecting 19

F and 31

P nuclei. The Bruker 400 MHz

spectrometer was equipped with a broadband probe capable of detecting 19

F and 31

P nuclei.

All spectrometers were equipped with variable temperature units.

2.2.2 NMR experimental parameters

Several NMR experimental parameters were taken into consideration when performing

multidimensional NMR experiments. The most important parameters required to set up

NMR experiments included the 90º pulse width, number of scans, spectral width, carrier

frequency, total number of data points, number of experiments (for 2D and 3D only) and

water suppression techniques.

89

NMR spectra were acquired using the software supplied by Bruker and Varian

spectrometers. The 90º pulses were calibrated separately for each sample on the

spectrometers used in the NIMR facility and EU-NMR. Generally, for small RNAs the

pulse width is between 6-10μs. However, the spectrometers used in the University of

Manchester had default settings and so the pulse width was set to 10.00μs. The number of

scans used for NMR experiments varies greatly and depends on the spectrometer

frequency, the concentration of the RNA sample and the type of experiment. For 1H-NMR

experiments, the number of scans generally ranged between 128-1024 scans. A larger

number of scans were used for 31

P-NMR and 19

F-NMR experiments for better sensitivity.

For 2D experiments, the number of scans normally ranged between 16-64 scans. The

spectral width used in 1H-NMR experiments was dependent on the type of solvent in the

sample. Samples in 1H2O had a larger spectral width range between 20-26ppm, which

allowed the observation of the lowfield imino proton region. Samples in 2H2O had smaller

spectral width of typically 10ppm. The carrier frequency was always set to the water

signal frequency for the 1H dimension to minimise any artefacts. The total number of data

points acquired for 1D NMR experiments usually ranged between 8192 and 65536. In 2D

experiments the number of data points in the first dimension was normally 4096, but the

number of data points in the second dimension was the same as the number of experiments.

The number of experiments refers to the number of increments in the evolution period(s),

which is only applicable for 2D and 3D NMR experiments. This typically ranged between

400-1200 in 2D NMR experiments. The WATERGATE water suppression technique was

used to reduce the sharp water signal of samples in 1H2O.

83 Conversely, the presaturation

technique was used to suppress any residual water peak of samples in 2H2O.

84 Details for

these two water suppression techniques are given in sub-section 2.3.1.

2.2.3 Data processing and analysis

All 1D NMR data were processed using the Spinworks program. The NMRPipe program

was used to process both 2D and 3D raw NMR data.85

NMRPipe is a program designed

for multidimensional spectral processing and analysis. The NMRPipe script that was used

consisted of three parts (Appendix I). The first part converts the .fid file from

90

Bruker/Varian format to the NMRPipe format. This allows NMRPipe to read acquisition

parameters for the direct dimension and indirect dimension(s). These parameters were

checked and altered if necessary such as the acquisition mode and carrier frequency. Since

the carrier frequency was positioned on the water peak, it was calibrated according to the

temperature of the NMR experiment.86

The second part of the script involves various

methods of data processing, which includes using a solvent filter, baseline correction,

applying a window function, zero filling, Fourier transform and phase correction. The

solvent filter was applied for spectra in both 1H2O and

2H2O by using the SOL or POLY

functions. Baseline correction was also performed using different POLY functions.

Gaussian (GMB) and sine bell (SP) window functions were used in all processing scripts.

Zero filling was applied to increase the number of data points to the nearest power of two.

Fourier transformation was applied to the direct dimension and the indirect dimension(s)

to create two frequency dimensions. Phase correction was applied using the PS command

for all dimensions; both zero-order (p0) and first-order (p1) phase correction was used.

Peaks were phased in both the horizontal and vertical dimensions to reduce negative peaks

and intensify the positive peaks. The final part of the script converts the processed data

file into a .ucsf format, which can be visualised by using the Sparky87

program.

The Sparky program was used to display and assign 2D and 3D NMR spectra. The

CCPNMR Analysis88

program was also used to assign NOESY spectra. The Analysis

program is similar to the Sparky program as it serves the purpose of visualisation and

assignment of NMR data. However, there are some advantages of using Analysis over

Sparky. The main advantage of using the Analysis program is that it allows an efficient

generation of distance restraints based on the intensity of assigned peaks in the NOESY

spectra. Therefore, the Analysis program was used to generate distance restraints for

structure calculations.

91

2.3 NMR techniques

2.3.1 Solvent suppression

The analysis of 1H-NMR spectra is complicated in the presence of a large water peak,

which is much more intense than the peaks corresponding to the solute of interest.

Suppression of the water peak greatly increases the sample sensitivity. This is especially

important for biological macromolecules, since the concentration of the solute is generally

1mM or less.

Various methods have been developed to suppress the large water resonance in NMR of

non-deuterated solvents. The most common water suppression techniques include

presaturation and water suppression by gradient-tailored excitation (WATERGATE).

Presaturation is a simple two pulse experiment that utilises a relatively, long, low power

radiofrequency (RF) pulse to selectively saturate a specific frequency, and a non-selective

90° pulse to excite the desired resonance (Figure 2.3.1). This method will significantly

reduce the intensity of the water peak, but with two disadvantages. Firstly, the peaks close

to the water frequency experience a loss in intensity and secondly the exchangeable proton

peaks are suppressed by the process of saturation transfer.

Figure 2.3.1 The presaturation pulse sequence. A selective, low power presaturation pulse

saturates the water frequency, which is followed by a non-selective, high power pulse to excite the

desired protons.84

92

The WATERGATE pulse sequence was created to provide highly selective and effective

water suppression by using a combination of tailored excitation with pulsed magnetic field

gradients. Magnetic field gradients disturb the magnetic field homogeneity of the sample

and since the gradient is gated on and off it is called a ‘pulsed’ magnetic field gradient.

The WATERGATE pulse sequence (Figure 2.3.2) consists of a non-selective 90° pulse

that uniformly excites all the protons regardless of their chemical shift. A subsequent echo

section of the pulse sequence is formed by two short magnetic field gradient pulses with a

centrally placed 180° selective pulse. The 90° pulse is followed by the first magnetic field

gradient, which dephases all the proton resonances. The 180° selective pulse acts on all

protons except for the water protons. The second magnetic field gradient then refocuses all

the coherences dephased by the first magnetic field gradient, provided that they

experienced the 180° selective pulse. Conversely, the water proton resonance further

dephases, as it was unaffected by the 180° selective pulse. Therefore, the magnetic field

gradients and the 180° selective pulse both act as a gradient spin echo for all protons

except for water. The FID is acquired with the water significantly suppressed.

Figure 2.3.2 The WATERGATE pulse sequence. The 90° (non-selective) and the 180° (selective)

pulses are shown in the top line. The τ delays inserted for gradient recovery. The bottom line

displays the pulsed magnetic gradients. Most protons experience gradient-180°-gradient and are

refocused, while the water protons experience gradient-0-gradient and are dephased. Therefore,

during signal acquisition (t2) the water signal is suppressed.83

93

2.3.2 1D NMR experiments with decoupling

19F-NMR and

31P-NMR experiments are performed with

1H decoupling in order to

simplify NMR spectra, by eliminating unwanted couplings. These experiments are

heteronuclear decoupling experiments because the observed and decoupled nuclei are

different. In 1D 19

F-NMR and 31

P-NMR decoupling experiments, the 19

F or 31

P nucleus is

observed with normal RF field, while at the same time irradiating the 1H nucleus with a

second, stronger RF field. Broadband decoupling techniques are used to irradiate all

protons within the proton spectral range, resulting in effective decoupling of all protons. It

is normally applied using a set of multiple pulses.

Figure 2.3.3 displays the pulse sequence of an ‘inverse gated decoupling’ technique. Here

the observed X nucleus is perturbed by a 90° pulse and the FID signal recorded. The

decoupling is applied only during the acquisition time to decouple all the protons.

Figure 2.3.3 1D X-{H} decoupled NMR experiment, where X represents an NMR active nucleus

apart from 1H. The decoupling pulse is activated at the same time as the FID is acquired for

nucleus X.

94

2.3.3 Variable temperature (VT) experiments

Variable temperature (VT) experiments involve performing 1D NMR experiments at

several different temperatures. VT experiments can allow the observation temperature-

dependent effects in NMR spectra, which can provide valuable information on RNA

structure and kinetics. A stack plot of the 1D NMR spectra, at different temperature points,

can be used to monitor changes in chemical shift or linewidth. Typically, 1H-NMR VT

experiments are used to monitor the extent of imino proton exchange for RNA samples in

1H2O. This can give useful information on the relative stability of base pairs.

2.3.4 T1 measurements

The inversion recovery experiment is used for the measurement of T1 values. A 180° pulse

is applied to invert the magnetisation to the -z-axis, and after a variable delay of τ seconds,

a second 90° pulse is applied which flips the magnetisation to the -y-axis where it can be

detected. The experiment is repeated with several different values of τ. Typically, a

relaxation delay of 5T1 must be given to allow the magnetisation to reach equilibrium,

although this is usually not practical. Once the experiments are complete, the spectra can

be plotted as a function of τ. Figure 2.3.4 represents the pulse sequence used for inversion

recovery experiments.

Figure 2.3.4 The inversion recovery pulse sequence. A 180° pulse is followed by a variable delay

(τ) and then a 90° detection pulse.

95

2.3.5 Water magnetisation transfer experiments

Imino proton exchange rates were obtained by experiments involving magnetisation

transfer from water, using the pulse sequence shown in Figure 2.3.5.72

This experiment

uses a selective 180° pulse sequence that selectively inverts the water magnetisation.89

The inversion is followed by a magnetisation transfer delay (τm), which is variable;

typically 20 different delays times are used from 5-100ms. During the variable delay,

weak gradients (G1 and G2) are applied to reduce radiation damping of the water signal.

Subsequently, the acquisition pulses are used to suppress the water signal and detect the

FID signal.

Figure 2.3.5 The pulse sequence of the water magnetisation transfer experiment. The DANTE

sequence is followed by a variable delay (τm) and three 90° pulses. Gradient pulses are represented

by G1 and G2.72

During the variable delay (τm), imino protons will exchange with water and the inverted

magnetisation is transferred from water to the imino protons. This causes the imino proton

peak intensity to decrease. By using different delay times, the rate of change of the imino

proton peak intensity can be calculated, which corresponds to the rate of imino proton

exchange.

96

2.3.6 2D Double-Quantum Filtered Correlation Spectroscopy (DQF-COSY)

The most basic 2D NMR experiment is a homonuclear COSY (COrrelated SpectroscopY)

experiment, in which scalar couplings can be observed.90

The COSY experiment is used

for determining basic connectivity between protons within a spin system. Scalar couplings

separated by three chemical bonds can be observed between non-equivalent protons.

However, cross peaks are usually difficult to observe when they are found very close to

the intense diagonal peaks.

A COSY spectrum can be purged of undesirable features in order to focus on more

relevant peaks, such as the cross peaks found close to the broad diagonal. This is achieved

by suppressing the single quantum transitions found by selecting coherence transfer

between the evolution and detection periods.91

The Double-Quantum Filtered (DQF)

COSY (Figure 2.3.6) experiment is employed to significantly reduce the intensity of the

diagonal peaks so to resolve any cross peaks that would originally be obscured by the

broad diagonal in a standard COSY spectrum. Consequently, important spectral regions

close to the diagonal can be analysed. The signals observed during t2 all derive from

double-quantum coherence present between the second and third 90° pulses. The

advantage of this is that only cross peaks between coupled nuclei are observed and single

resonances are removed.

Figure 2.3.6 The pulse sequence of a DQF-COSY experiment. The first 90° pulse is the same as

in a standard COSY experiment. The second 90° pulse is immediately followed by a third 90°

pulse; the third pulse acts in combination with second pulse as a double-quantum filter to convert

the double-quantum coherence created by the second pulse back into single-quantum coherence.92

97

2.3.7 2D Total Correlation Spectroscopy (TOCSY)

The homonuclear 2D TOCSY (TOtal Correlation SpectroscopY) experiment is very

similar to COSY, except that the cross peaks found in TOCSY spectra correspond to

coupling of protons within an extended spin system. Therefore, scalar couplings can be

observed for protons separated by more than three chemical bonds, within the same spin

system. TOCSY experiments are useful for identifying protons belonging to a network of

couplings and for identifying different spin systems within a molecule.

The TOCSY pulse sequence (Figure 2.3.7) differs from the COSY pulse sequence by the

addition of an isotropic mixing time (τm). In practice isotropic mixing is achieved by the

use of specially designed multi-pulse sequences (a spin-lock field) for net magnetisation

transfer among scalar coupled homonuclear spins.93

During the spin-lock period

magnetisation transfer is possible between all coupled nuclei within a spin system, even if

they are not directly coupled. Consequently, correlation of each spin to other spins within

the same spin system can be observed as cross peaks. The extent of the correlation

depends upon the length of the mixing period, which can be adjusted experimentally.

Shorter mixing times allow for correlations between adjacent nuclei to be observed, while

correlations between more distant nuclei can be observed with longer mixing times as the

time of magnetisation transfer transpires for a longer period.

Figure 2.3.7 The pulse sequence of a TOCSY experiment. The first 90° pulse flips the spin

magnetisation onto the x,y plane. The evolution period is followed by the isotropic mixing, which

transfers magnetisation between spins connected via an unbroken network of couplings.94

98

2.3.8 2D Heteronuclear Single Quantum Coherence (HSQC)

The HSQC (Heteronuclear Single Quantum Correlation) experiment can be used to reveal

couplings between 1H-

13C nuclei that are directly coupled. In heteronuclear 2D NMR

experiments, problems can arise with the lower gyromagnetic ratio of 13

C nuclei compared

with 1H, which reduces the sensitivity of

13C nuclei. To solve this problem the HSQC

pulse sequence uses an adapted version of an INEPT (Insensitive Nuclei Enhancement by

Polarisation Transfer) sequence.95

The INEPT sequence was developed to take advantage

of the high gyromagnetic ratio of 1H and enhance the sensitivity of nuclei such as

13C.

The HSQC experiment incorporates this INEPT sequence at the start of the pulse sequence,

improving the sensitivity of those nuclei with low gyromagnetic ratio. This is followed by

a 180° proton pulse at mid-evolution that removes the effect of proton coupling. The final

part of the pulse sequence involves an inverse INEPT step, which enhances the sensitivity

of insensitive nuclei to a sensitivity comparable to 1H (Figure 2.3.8).

Figure 2.3.8 1H-

13C HSQC sequence adapted from the INEPT pulse sequence. The first pulses are

derived from INEPT pulse sequence which transfers magnetisation from 1H to

13C nuclei. A 180°

pulse on 1H nuclei forms a spin echo, so the evolution of coupling is refocused. At mid-evolution

the 13

C spin magnetisation then evolves during t1, at which time it acquires a frequency label

according to the offset of 13

C. The inverse INEPT step transfers magnetisation back to 1H from

13C

nuclei, yielding enhanced sensitivity of 13

C comparable to that of 1H.

96

99

2.3.9 2D Nuclear Overhauser Effect Spectroscopy (NOESY)

Dipolar 2D NMR experiments are used to observe proximity of nuclei through space,

when approximately <5.0Å apart. Although the scalar 2D NMR experiments are

extremely useful in structure determination, by far the most important technique used for

structure determination of large molecules, such as proteins and nucleic acids, is the

homonuclear NOESY (Nuclear Overhauser Enhancement SpectroscopY) experiment.

The pulse sequence of the NOESY experiment consists of three 90° pulses, with t1 as

evolution period (between 1st and 2nd pulses) and τm as the mixing time (between 2nd and

3rd pulses), followed by t2 as FID detection (Figure 2.3.9).97

The mixing time (τm) can be

varied in NOESY experiments, which can be extremely useful in structure determination.

A longer mixing time can produce more intense NOE peaks, however in larger molecules

shorter mixing times are usually employed to avoid problems of ‘spin diffusion’. Spin

diffusion is when the magnetisation is transferred from one spin to another via multiple

other spins. The result is that the cross-relaxation rate is no longer proportional to NOE

intensity leading to generation of inaccurate distance restraints.

Figure 2.3.9 The pulse sequence of a NOESY experiment. The first 90° pulse flips the spins onto

the x,y plane. The spins precess in the evolution period before a second 90° pulse flips the spins

onto longitudinal axis and a predetermined mixing period allows for the exchange of

magnetisation between dipolar spins. Finally the spins are flipped back onto the x,y plane for

detection.97

100

2.3.10 2D Heteronuclear Overhauser Effect Spectroscopy (HOESY)

Heteronuclear NOEs can also be observed in 2D HOESY (Heteronuclear Overhauser

Effect Spectroscopy) experiments. 1H-

19F NOE’s are readily accessible due to the 100%

abundance and high gyromagnetic ratio of the 19

F nucleus. Therefore, 1H-

19F HOESY

experiments can provide additional distance restraints for structure determination and have

wider applications in studying macromolecular complexes.98

The pulse sequence of the HOESY experiment is shown in Figure 2.3.10. In a two spin

system of 1H and X nuclei, X is the observed nucleus. The first 90° (

1H) pulse flips the

magnetisation onto the x,y plane. The two vectors of 1H magnetisation (α and β),

corresponding to the two X spin states, precess for a period of t1/2. After a 180° pulse on

X, the two spin state labels interchange and precess for a further period of t1/2. This

refocuses the two vectors of 1H magnetisation removing any scalar interaction between

1H

and X nuclei and allowing only dipolar interactions to give rise to signals. The second 90°

(1H) pulse flips the magnetisation onto the z-axis. During the mixing time (τm)

magnetisation is transferred between 1H and X spins through dipolar interactions. The FID

signal is recorded following the 90° (X) observed pulse.

Figure 2.3.10 The pulse sequence of the HOESY experiment, whereby X is the observed nucleus.

The X signal is recorded during t2 and the 1H signal is recorded as a function of t1.

99

101

2.3.11 2D CPMG-HSQC-NOESY

2D 31

P-NMR experiments can provide important information about the structure and

dynamics of nucleic acids. The 2D 1H-

31P CPMG-HSQC-NOESY experiment allows the

observation of proton-phosphorus correlations. The effective application of the HSQC

component requires the use of a Carr-Purcell-Meiboom-Gill (CPMG) pulse, which is a set

of closely spaced 180° refocusing pulses, during magnetisation transfer between scalar

coupled phosphorus and proton nuclei.100

This optimises the scalar coupling between

proton and phosphorus, which results in enhanced sensitivity. Additionally, the CPMG

pulse reduces phosphorus line broadening in the presence of conformational exchange,

which is experienced by the phosphate backbone in nucleic acids. The CPMG-HSQC can

then be applied in combination with a NOESY, which contains both scalar and dipolar

coupling information. Scalar coupling is observed between phosphorus and H3ʹ/H5ʹ/H5ʹʹ

protons. Dipolar coupling is observed between these protons and other protons within

5.0Å. The pulse sequence of the CPMG-HSQC-NOESY experiment is shown in Figure

2.3.11.

Figure 2.3.11 The pulse sequence of the 1H-

31P CPMG-HSQC-NOESY experiment. τ represents

the delays times around the 180° refocusing pulses, τm is the mixing time for the NOESY

component. The symbol G corresponds to the gradient pulses shown as black filled shaped

pulses.101

102

2.3.12 3D NOESY/2Q-COSY

In 2D NMR experiments such as 2D COSY, scalar couplings are restricted to providing

information on intranucleotide connectivities. Conversely, dipolar couplings provide

information on internucleotide connectivities, such as in 2D NOESY experiments.

Therefore, NOESY and COSY experiments can be combined into one experiment to take

advantage of both scalar and dipolar couplings. A combination of a NOESY and 2Q-

COSY pulse sequence is employed to produce a homonuclear 3D NOESY/2Q-COSY

experiment, which uses a multiple quantum (MQ) excitation step (Figure 2.3.12). Three

frequency dimensions (F1, F2 and F3) produce the 3D NMR spectrum consisting of a

NOESY, 2Q-COSY and a back-transfer plane.

If we consider two coupled protons, HA and HX, scalar coupling can be observed between

HA and HX as well as strong NOEs. These NOEs are known as ‘inner’ NOEs, since they

contribute to MQ coherence between the two coupled protons. Furthermore, weaker NOEs

from HA and HX can be observed to other protons within a distance of 5.0Å. These NOEs

are known as ‘outer’ NOEs. In addition to the connectivities observed, the 3JH3ʹ-H4ʹ

coupling constants can be measured for C3ʹ-endo sugar puckers and 3JH1ʹ-H2ʹ for C2ʹ-endo

sugar puckers.

Figure 2.3.12 The 3D NOESY/2Q-COSY pulse sequence. The DANTE (Delays Alternating with

Nutation for Tailored Excitation) presaturation sequence is used to suppress the residual water

signal. The evolution period (t1) is followed by the NOE mixing time (τm) and then the multiple

quantum excitation step (τMQ). The t1 and t2 are the first and second indirect dimensions, and t3 is

the third (direct) dimension.102

103

2.4 NMR assignment of RNA

2.4.1 Assignment strategy

NMR spectroscopy is a powerful technique that is used to solve complex structural

problems. It has been used for many years to study the structure and dynamics of RNA.

However, within the current limitations of NMR spectroscopy, approximately 50

nucleotides can be analysed at high resolution with complete assignments. To assign the

structure of RNA, various NMR experiments are employed, which provide information on

base pairing, local conformation, secondary and tertiary structure of RNA. The NMR

assignment strategy of RNA is well established and usually follows a standard

methodology.103

In the past decade, advances in NMR spectroscopy have promoted

experiments for both unlabelled and labelled RNA.104-105

This assignment strategy was

applied when performing 2D and 3D NMR experiments (Figure 2.4.1).

Figure 2.4.1 Protocol for NMR assignment of RNA. The green boxes indicate the solvent used.

The red boxes contain information on the specific NMR experiments employed and the cyan boxes

represent the assignments that can be obtained from the corresponding experiment(s).

104

The NOESY experiment (2°C/1H2O) was mainly used for the assignment of imino-imino

connectivities for local secondary structure and imino-amino connectivities to establish

base pairing involved in the RNA helix. Cross peaks corresponding to imino to

H2/H5/H6/H8 connectivities were also assigned. The NOESY experiment (25°C/ 2H2O)

was used for H5-H6 assignment and sequential assignment of H6/H8-H1ʹ intra- and

internucleotide connectivities. The 1H-

13C HSQC experiment allows the observation of

1H-

13C one bond couplings, which means that proton chemical shifts can be clearly

identified due to the large dispersion of 13

C chemical shifts. Consequently, 1H-

13C HSQC

experiments were extremely useful for the assignment of cross peaks in NOESY spectra.

The DQF-COSY experiment (25°C/2H2O) was used for identification of sugar proton spin

systems. The main use of the DQF-COSY was to identify H1ʹ-H2ʹ and H3ʹ-H4ʹ cross

peaks, which provided information on the sugar pucker conformation of nucleotides. The

1H-

31P CPMG-HSQC-NOESY experiment (25°C/

2H2O) was used for phosphate backbone

assignment and confirmation of H3ʹ, H4ʹ, H5ʹ, H5ʹʹ chemical shifts, provided by 1H-

31P

couplings. The homonuclear 3D NOESY/2Q-COSY experiment allowed the identification

of sugar proton chemical shifts.

To aid in the assignment of NOESY and 1H-

13C HSQC spectra, the standard observable

chemical shifts of various 1H and

13C nuclei present in RNA were utilised. Chemical shifts

are characterised by specific ranges, which correspond to particular atoms within the RNA

structure. Table 2.4.1 presents the 1H and

13C chemical shifts of peaks observed in

NOESY and 1H-

13C HSQC spectra.

The assignment of cross peaks in NOESY spectra is the most important aspect of the

assignment procedure. There are three main steps to be taken which would allow for

complete assignment of RNA. The first step is the identification of base protons (NH, NH2,

H2, H5, H6 and H8). The second step is to identify the sugar proton spin system (H1ʹ, H2ʹ,

H3ʹ, H4ʹ, H5ʹ and H5ʹʹ). The third step is to correlate the base protons to sugar protons,

with intra- and internucleotide connectivities. These three steps are discussed in the

following sub-sections.

105

Atoms 1H

(ppm)

13

C

(ppm)

C1ʹH1ʹ 5.0-6.0 89-95

C2ʹH2ʹ 4.0-5.0 70-80

C3ʹH3ʹ 4.0-5.0 70-80

C4ʹH4ʹ 4.0-5.0 81-86

C5ʹH5ʹ/H5ʹʹ 3.5-5.0 62-70

C2H2 6.5-8.5 145-155

C5H5 5.0-6.5 95-105

C6H6 7.0-8.0 137-140

C8H8 7.0-8.5 133-140

NH2 G (non H-bonded) 5.5-6.5 -

NH2 C (non H-bonded) 7.0-7.5 -

NH2 A (non H-bonded) 6.0-6.5

NH2 G (H-bonded in GC) 8.0-9.0 -

NH2 C (H-bonded in GC) 8.0-9.0 -

NH2 A (H-bonded in UA) 7.5-8.5 -

NH G (in GC) 12.0-13.5 -

NH U (in UA) 13.0-15.0 -

NH G (in GA) 10.0-11.0 -

Table 2.4.1 Summary of 1H and

13C chemical shifts observed in NOESY and

1H-

13C HSQC

spectra of RNA. H-bonded refers to ‘hydrogen’ bonded.

106

2.4.2 Identification of base protons

2.4.2.1 Identification of exchangeable protons

Exchangeable proton correlations provide information on base stacking and base pairing.

Therefore, the assignment of exchangeable protons is crucial in determining the secondary

and tertiary structure of RNA. There are two different protons that fall in the exchangeable

category, imino and amino protons. Imino protons correspond to an –NH group on uracil

and guanine bases. Amino protons correspond to an –NH2 group on adenine, guanine and

cytosine bases.

Imino protons can be unambiguously assigned by observing NOE cross peaks between

imino-imino protons of guanine and uracil. Imino-imino connectivities between

neighbouring base pairs are generally at least 3.5-4.0Å apart, while cross-strand imino-

imino connectivities can be up to 5.0Å apart, in A-form RNA. Uracil and guanine imino

proton chemical shifts can be distinguished in Watson-Crick base pairing. Uracil and

guanine imino protons resonate between 13.0-15.0ppm and 12.0-14.0ppm, respectively,

which means that uracil imino protons are generally found to be lowfield of guanine imino

protons. The weaker hydrogen bonding of A.U base pairs compared to G.C base pairs

results in uracil imino protons being more exposed to the external magnetic field than

guanine imino protons. Therefore, uracil imino protons are more deshielded than guanine

imino protons, causing uracil imino protons to be shifted lowfield of guanine imino

protons. However, it can be difficult to distinguish between guanine and uracil imino

protons as they can be affected by ring current shifts and hydrogen bonding.

Imino protons from unpaired bases or non-canonical base pairing have distinct highfield

chemical shifts in the range of 10-12ppm. This is because guanine and uracil bases, which

are base paired and located in the helix, are involved in base stacking interactions. Base

stacking interactions are caused by the non-covalent interaction of π-orbitals between the

aromatic rings of adjacent bases. When an external magnetic field is directed

perpendicular to the plane of the base aromatic rings, a magnetic field is induced called

107

the ring current effect. Imino protons experience a deshielding effect because the induced

magnetic field is in the same direction as the external magnetic field, which results in a

lowfield shift. Therefore, if we apply the same principle to bases that are partially base

stacked or are not involved in base stacking, the imino proton chemical shifts would

appear more highfield.

Amino protons are found highfield of imino protons due to the increased diamagnetic

shielding provided by the second electron from hydrogen. However, the two protons in

NH2 have distinct chemical shifts as one is involved in hydrogen bonding in Watson-Crick

base pairing. The hydrogen bonded proton is found lowfield of its non-hydrogen bonded

analogue, due to deshielding from the electronegative oxygen hydrogen bond acceptor.

Cytosine NH2 resonances can be observed easily, in comparison with adenine and guanine

NH2 resonances, due to slow exchange. In Watson-Crick base pairing, two strong NOE

cross peaks can be observed from guanine NH to cytosine NH2. Adenine and guanine

amino protons are difficult to observe unless under low temperature or pH. However,

adenine amino protons can be identified from intense NOE cross peaks from uracil NH to

adenine NH2, in A.U base pairs.

2.4.2.2 Identification of non-exchangeable protons

Pyrimidine H5 and H6 resonances can be readily identified through strong NOE cross

peaks in NOESY (1H2O and

2H2O) spectra, which are observable due to the covalently

fixed 2.4Å distance between the two protons. Generally, H5 chemical shifts are identified

by examining NOE cross peaks between guanine NH to cytosine H5 and uracil NH to its

own H5. Subsequently, the identified H5 chemical shifts can be used to identify strong

H5-H6 NOE cross peaks, in NOESY (1H2O/

2H2O) spectra. Uracil and cytosine H5

resonances can be distinguished from C5 chemical shifts in 1H-

13C HSQC spectra. Uracil

C5 chemical shifts are typically found lowfield (100-105ppm) of cytosine C5 chemical

shifts (95-100ppm). From the assignment of H5-H6 cross peaks in the NOESY (2H2O)

spectrum, the H6 chemical shifts are identified, which allows the subsequent assignment

of C6-H6 correlations in 1H-

13C HSQC spectra. C6-H6 and C8-H8 peaks are found in the

108

same region in 1H-

13C HSQC spectra, due to similar

1H and

13C chemical shifts. Therefore,

the assignment of C6-H6 correlations allows for easier assignment of C8-H8 correlations

through process of elimination and the identification of H8 chemical shifts.

In Watson-Crick A.U base pairing, the adenine H2 proton gives rise to a strong NOE cross

peak to the base paired uracil imino proton, which is observed in the NOESY (1H2O)

spectrum. H2 proton chemical shifts are in a similar range to that of H6 and H8, but they

can be easily identified from 1H-

13C HSQC spectra, since C2 chemical shifts are found

lowfield of C6 and C8. Both intra- and internucleotide connectivities between H2-H1ʹ can

be observed in the NOESY (2H2O) spectrum, corresponding to a distance of 4.5 Å or

longer.

2.4.3 Identification of sugar protons

Anomeric H1ʹ chemical shifts are generally found in the 5.0-6.0ppm range; unusually

shifted H1ʹ resonances can be observed further highfield at 3.5-4.5ppm. In 1H-

13C HSQC

spectra, C1ʹ resonances are observed at 90-95ppm and provide the best way to assign H1ʹ

resonances. NOESY and TOCSY spectra can also be used to distinguish between H1ʹ and

H5 resonances within the same spectral region, through correlations between H5 and H6

protons.

In the DQF-COSY spectrum, the H1ʹ-H2ʹ coupling constant is small (<2Hz),

corresponding to the C3ʹ-endo conformation found in A-form RNA, so cross peaks

between H1ʹ and H2ʹ are not always evident. The C3ʹ-endo conformation is predominant in

regular RNA secondary structure so only a small number of cross peaks are expected in

the anomeric-sugar region. Conversely, the coupling constant between H3ʹ-H4ʹ is

approximately 7Hz for C3ʹ-endo sugars, so cross peaks between H3ʹ-H4ʹ can be more

easily observed and used to indicate a C3ʹ-endo conformation.

Sugar proton chemical shifts from H2ʹ, H3ʹ, H4ʹ, H5ʹ and H5ʹʹ can be quite difficult to

identify. These resonances all appear within the 3.0-5.0ppm range and most of them lie in

109

a narrower range of 4.0-5.0ppm. There are two different methods used to identify sugar

proton chemical shifts. The first method involves the observation of NOE cross peaks to

H1ʹ protons, in the NOESY (2H2O) spectrum. Strong NOE cross peaks are observed for

H1ʹ-H2ʹ connectivities due to the covalently fixed distance of 2.8-3.0Å, so they can be

easily assigned. Similarly, H1ʹ-H3ʹ, H1ʹ-H4ʹ, H1ʹ-H5ʹ and H1ʹ-H5ʹʹ cross peaks can also be

observed and assigned. The second method used for observation of sugar proton

resonances is by involving correlation to 31

P nuclei in the phosphodiester backbone, using

1H-

31P CPMG-HSQC-NOESY experiments. Correlations between phosphorus and H3ʹ,

H5ʹ, H5ʹʹ can be observed through scalar coupling.

2.4.4 Sequence-specific resonance assignment

In clearly defined A-form duplex regions of RNA, sequence-specific sequential

assignment is possible, which allows the identification of each nucleotide in the RNA

sequence. H6/H8-H1ʹ NOE cross peaks can be easily identified; each aromatic H6/H8

proton shows two NOE cross peaks to H1ʹ. The first corresponds to an intranucleotide

connectivity between the H6/H8 and H1ʹ protons. The second corresponds to an

internucleotide connectivity between the H1ʹ proton and its 3ʹ-neighbouring H6/H8 proton.

These connectivities are shown in Figure 2.4.2. The sequential distance between H6/H8-

H1ʹ is greater than 4.0Å in A-form RNA helices. The combination of both intranucleotide

and internucleotide connectivities can lead to full sequential assignment.

Sequence-specific sequential assignment can also be achieved by identifying

intranucleotide H1ʹ-H2ʹ connectivities and then internucleotide H2ʹ-H6/H8 connectivities

(Figure 2.4.3). The internucleotide connectivity corresponds to a correlation between the

H2ʹ proton and its 3ʹ-neighbouring H6/H8 proton. In the C3ʹ-endo conformation, the

sequential H2ʹ to H6/H8 protons are in close proximity (~3.0Å) and so can be observed as

strong NOE cross peaks.

110

Figure 2.4.2 Scheme representing intranucleotide H6/H8-H1ʹ connectivities (blue) and

internucleotide H6/H8-H1ʹ connectivities (red). These connectivities follow well established NOE

sequential assignment pathways.

Figure 2.4.3 Scheme representing intranucleotide H1ʹ-H2ʹ connectivities (blue) and

internucleotide H2ʹ-H6/H8 connectivities (red).

111

2.5 Structure determination protocol of RNA

Structure determination involves several steps to generate a final solution structure. Firstly,

a range of NMR experiments are performed that provide the necessary information to

produce a complete assignment of proton-proton connectivities in NOESY (1H2O/

2H2O)

spectra. Subsequently, NOE distance restraints are generated which provide crucial

structural information. Secondly, information is extracted from NMR data that allows

dihedral angle and hydrogen bond restraints to be produced. Finally, these restraints are

inputted into a structure determination program, which will calculate a number of

structures. These structures are then refined to create a single viable structure.

2.5.1 Restraints

2.5.1.1 Distance restraints

A distance restraint is the most important parameter used for structure determination.

Distance restraints are derived from the NOE cross peak volume. The theory of NOE

states that the dipolar cross relaxation rate is proportional to the inverse sixth power of the

distance between two interacting 1H spins. Since the cross relaxation rate is also

proportional to the intensity of the NOE cross peak, the intensity can be directly related to

the interproton distance. This relationship is shown in Equation 2.5.1, where (i) and (j)

represent two 1H spins.

NOEij ~ 1/rij6 Equation 2.5.1

Interproton distances were calculated from the NOE cross peak volume in NOESY

(1H2O/

2H2O) spectra, using the CCPNMR Analysis program. The NOE cross peak

volumes were converted to distances and automatically calibrated. A reference was chosen

as the distance between H5-H6 protons (2.43Å). Distance restraints were generated with

high and low error bounds of approximately 20%. NOE cross peak intensities were

112

categorised into strong (1.8–2.5 Å), medium (2.6–3.3 Å), weak (3.4–5.0 Å) and very weak

(5.1–7.0 Å) ranges. These ranges were based on the intensities of known H5-H6 distances.

2.5.1.2 Dihedral angle restraints

Dihedral angle restraints are used with distance restraints to improve the tertiary RNA

structure, specifically the RNA backbone conformation. They can be experimentally

generated from 3J coupling constants using the Karplus relationship. There are six

backbone dihedral angles and a seventh glycosidic angle that were incorporated into

structure calculations. Additionally, the ν1 and ν2 dihedral angles were used to restrain the

sugar pucker conformation.

The NOE cross peak distances of H8-H1ʹ (purine) and H6-H1ʹ (pyrimidine) were used to

define the glycosidic angle (χ). Strong intranucleotide H6/H8-H1ʹ cross peaks (2.0-2.5Å)

indicate syn conformation of glycosidic angle. Conversely, weak intranucleotide H6/H8-

H1ʹ cross peaks (3.5-4.5Å) indicate anti conformation of the glycosidic angle.

The sugar ribose conformation was also restrained for each nucleotide by defining the δ

angle. The DQF-COSY experiment was used to observe H1ʹ-H2ʹ cross peaks. 3JH1ʹ-H2ʹ

couplings less than 2Hz are characteristic of the C3ʹ-endo conformation. Couplings less

than 2Hz are usually not observed in a DQF-COSY experiment as the negative and

positive lines will cancel out. Therefore, the absence of H1ʹ-H2ʹ cross peaks are generally

assumed to indicate C3ʹ-endo conformation, while the observation of clear cross peaks

indicates C2ʹ-endo conformation.

Coupling constant information was unavailable to obtain the (α), (β), (γ), (ε) and (ζ)

dihedral angle restraints used to define nucleotide structure. Therefore, dihedral angles

were loosely restrained to the range within A-form parameters.35

All dihedral angles used

in the structure determination of the 16mer apo-RNA, 16mer Mg2+

RNA complex and

15mer apo-RNA are summarised in Table 2.5.1.

113

Dihedral angle Restraint value (°)

α -60 ± 30

β 165 ± 15

γ 60 ± 30

δ (C3ʹ-endo) 60 ± 30

δ (C2ʹ-endo) 165 ± 15

ε -165 ± 15

ζ -60 ± 30

χ (C3ʹ-endo) -165 ± 15

χ (C2ʹ-endo) -120 ± 30

ν1 (C3ʹ-endo) -25.0 ± 30

ν2 (C3ʹ-endo) 37.3 ± 30

ν1 (C2ʹ-endo) 25.0 ± 30

ν2 (C2ʹ-endo) -35.0 ± 30

Table 2.5.1 Dihedral angle restraints used for defining nucleotide structure (α, β, γ, δ, ε, ζ, χ) and

the ribose sugar (ν1 and ν2) in the structure calculations.35

2.5.1.3 Hydrogen bond and planarity restraints

Hydrogen bond restraints are crucial for RNA secondary/tertiary structure determination.

They provide information on base pairing, with distance restraints between the donor and

acceptor of typically 3.1±0.4Å. Chemical shifts of exchangeable imino protons can be

shifted to lowfield arising from base stacking interactions. This can be invaluable in

distinguishing between Watson-Crick base pairing and non-canonical base pairing or

unpaired guanine/uracil bases. The imino proton chemical shifts resonate to lowfield due

to the ring current effect caused by base stacking interactions of adjacent base pairs in the

helical stem. Therefore, uracil and guanine imino protons resonating between 12ppm and

15ppm were restrained as canonical base pairs. Loop imino proton chemical shifts

resonate highfield of the stem imino proton resonances between 10ppm and 11ppm.

Restraints between base pairs were validated by the observation of imino-amino and

imino-H2/H5 base pair connectivities. If these observation were not made then hydrogen

bonds and planarity restraints were loosely restrained for base pairing.

114

2.5.2 Structure calculation

Structure determination is the process by which experimental data are utilised to generate

three-dimensional structures with the aid of computational structure determination

programs. The most popular biomolecular structure determination program used for

macromolecules such as DNA, RNA and proteins is XPLOR-NIH.106

Structure determination calculations require a starting structure, to which experimental

restraints can be added. The starting structure is created by using two separate files, a

structure file and a topology file. The structure file contains all the atomic coordinates of

the atoms found in the molecule. The topology file adds additional information such as

atom types, charges and bond lengths etc., to complete the construction of the molecule.

The structure determination program can extract information from both files to define the

starting structure.

Once the starting structure is generated, experimental restraints can be input into the

structure determination calculation. NOE distance restraints, dihedral angle restraints,

hydrogen bond restraints and also planarity restraints can be included. However, the

inclusion of experimental restraints is not always possible depending on the quality of

NMR data. Subsequently, restraints can be added to the structure calculation that will

avoid the production of very random structures, improving the viability of structures

generated. Distance restraints are derived directly from NOE data. Dihedral angle

restraints and hydrogen bond restraints can be approximated with certain assumptions and

refined, accordingly. When the selected restraints are included with the starting structure,

the structure calculation can proceed.

The next step in the structure determination process is to take the randomised starting

structure and run the structure determination calculations, which is outlined in Figure 2.5.1.

Firstly, the process of ‘simulated annealing’ is carried out. The initial temperature was set

to 3500K and reduced by 12.5K every time step, until the final temperature of 25K was

reached. High temperature dihedral angle dynamics were run for 800ps. This was

115

followed by torsion angle minimisation and Cartesian-space energy minimisation. The

simulated annealing process generated 200 structures, of which the 20 lowest RMSD (root

mean square deviation) structures were accepted.

The lowest RMSD structure generated by the simulated annealing step was then placed

into the refinement step. The refinement step takes the violations produced in the

simulated annealing step into consideration. Violations are defined as parameters that have

a value which has fallen outside the error bounds. The refinement step aims to reduce the

number of violations in order to generate improved structures. The 200 structures that are

produced from the refinement step are analysed and checked to see whether there are large

violations to the distance restraints and dihedral angle restraints. In the refinement step, an

initial energy minimisation step was followed by a high temperature step (10ps). The

temperature for the dihedral angle simulated annealing calculation was set to 2000K and

reduced by 25K every time step, until the final temperature of 25K was reached. A torsion

angle energy minimisation and a Cartesian-space energy minimisation were run to

produce the refined structures. Structures were rejected based on the number of violations

they produced. If this was consistent throughout most of the refined structures generated

then restraints would have to be examined and modified accordingly. Modifications would

be placed into the starting structure and the processes discussed above would be repeated.

When the top 20 average structures have been collected, a final structure can be acquired

on the basis of lowest energy values, low root mean square deviation (RMSD) and least

violations. The VMD-XPLOR program was used to visualise the ensemble of structures

and calculate the RMSD values.107

The final structure can also be represented as an

ensemble of aligned structures, to show the extent of convergence of lowest energy

structures.

116

Figure 2.5.1 Scheme summarising the procedure for structure determination of RNA. The

restraints are added to the starting structure, which is followed by the simulated annealing step.

The lowest RMSD structure generated from the simulated annealing process is used for the

refinement step. The lowest RMSD structure from the refinement process is then assessed for

acceptance as the final NMR solution structure.

117

2.5.3 Conformational analysis

The 3DNA108

program allows one to analyse the three-dimensional nucleic acid structures

by characterising all base interactions and double helical character of base pair steps.109

A

standard nucleic acid base reference frame is utilised for the description of nucleic acid

base pair geometry.110

Several parameters can be used to describe the geometry of base

pairs and sequential base pair steps, which are shown in Figure 2.5.2.

The first parameters are the complementary base pair parameters; shear (Sx), stretch (Sy),

stagger (Sz), buckle (κ), propeller (π) and opening (σ). These six base pair parameters

define the relative position and orientation of two complementary bases. The 3DNA

program generates values for all complementary base pairs.

Six rigid body parameters are required to describe the position and orientation of one base

pair to another. These six parameters consist of three rotations and three translations. In

3DNA two sets of such parameters are used, local helical parameters; x-displacement (dx),

y-displacement (dy), inclination (η), tip (θ), helical twist (Ω) and helical rise (h), and base

pair step parameters; shift (Dx), slide (Dy), rise (Dz), tilt (τ), roll (ρ) and twist (ω). Local

helical parameters describe the regularity of the helix and the base pair step parameters

describe the stacking geometry from a local perspective.

Dihedral angles that define nucleotide structure can be calculated by 3DNA. Dihedral

angles (ν1 and ν2) that define the sugar pucker conformation of a nucleotide are also given,

including the phase pseudorotation angle (P) and the amplitude (φ).

119

Figure 2.5.2 Definitions of the different parameters used in the 3DNA program. Complementary

base pair parameters (red), base pair step parameters (blue) and local helical parameters (green)

are clearly outlined. All images illustrate the positive values of the corresponding parameters.

Helical twist (Ω) is the same as twist (ω) and helical rise (h) is the same as rise (Dz).108

120

2.5.4 Structure validation

MolProbity is a structure validation program that evaluates the quality of protein and

nucleic acid structures.111

It uses the all-atom analysis method to assess the quality of

structures by combining information from atom positions, optimisation of explicit

hydrogen atoms and the van der Waals radii of atoms. This all-atom analysis method

operates by rolling a virtual 0.5Å diameter ball around the van der Waal surface of atoms.

It then calculates the number of non-bonded atom pairs that overlap by more than 0.4Å.

This is reported as a ‘clash score’, which is the number of serious clashes per 1000 atoms.

The clashes do not represent strained conformations but indicate the inconsistencies in the

structure.

MolProbity also runs a geometrical analysis of the structure, which includes an evaluation

of bond lengths, bond angles, sugar pucker conformation and backbone conformation. The

bond length and angles are assessed by comparing them with nucleic acid parameter sets

that contain information on equilibrium bond lengths and angles. The value given

corresponds to the percentage of nucleotides that contain deviated bond length(s) or

angle(s). The ribose sugar pucker is assessed by measuring the perpendicular distance

between the C1ʹ-N1/N9 glycosidic bond vector and the subsequent 3ʹ-end phosphate group;

>2.9Å for C3ʹ-endo and <2.9Å for C2ʹ-endo. Recently a consensus classification and

nomenclature have been defined for the RNA backbone, using all of the backbone

dihedral angles.112

Fifty-four conformers have been identified, which are used to identify

the conformers present in an RNA structure.

The final structures of the 16mer apo-RNA, 16mer Mg2+

RNA complex and 15mer RNA,

were all validated using the MolProbity program. Values for the five different parameters

mentioned were obtained.

121

2.6 Quantitative measurement of exchange rate constants

Imino proton exchange rates were measured using a 600 MHz Varian INOVA

spectrometer. Imino proton exchange theory has been previously described in sub-section

1.5.5.

The T1 rate constants of the imino protons of the 16mer apo-RNA and 16mer Mg2+

RNA

complex (batch 2) were determined by semi-selective inversion recovery experiments,

whereby the inversion pulse was applied to the imino proton region. In total, 13 delays

were used in the inversion recovery experiments; 0ms, 50ms, 100ms, 150ms, 200ms,

250ms, 300ms, 350ms, 400ms, 450ms, 500ms, 750ms and 1000ms. A relaxation delay of

8.0s was used between each pulse sequence to allow magnetisation to reach equilibrium.

The apparent relaxation rates of the imino protons (R1a) were determined by measuring

the integral of each imino proton peak in the stack plot and fitting the data to a three

parameter exponential function using the MestReNova program. The R1a value for each

imino proton was calculated ten times in order to acquire an average value. T1 values were

subsequently calculated by using the T1=1/R1a relationship. T1 measurements for water

were not made, so a value of 0.3s-1

was used for the apparent relaxation rate of water

(R1w).71

Imino proton exchange rates were measured by water magnetisation transfer experiments

at three different temperatures of 2°C, 15°C, 35°C for the 16mer apo-RNA and two

temperatures of 15°C and 35°C for the 16mer Mg2+

RNA complex. Since the imino proton

exchange rates are temperature dependent, a temperature of 15°C was chosen to measure

the exchange rate of the rapidly exchanging G178 loop imino proton and 35°C to measure

exchange rate of the exchange retarded stem imino protons. In total, 20 different delay

times were used ranging from 5ms to 100ms. A relaxation delay of 17.5s was allowed for

complete relaxation of the water longitudinal magnetisation. The exchange rates were

calculated by fitting relative peak intensities to the equation:

122

Equation 2.6.1

whereby Kex is the exchange rate constant of the imino proton, It and I0 are the imino

proton peak intensities at times (t) and (0), respectively.73

Peak intensities were measured

using the MestReNova program; the peak intensity at time (0) corresponds to a time of

5ms. Using the equation, exchange rates were calculated for each time period after 5ms

providing a maximum of nineteen values for Kex. These values were averaged to give a

final value of Kex. The exchange rate of the imino protons is related to their T1 values

according to the equation:

R1a = R1 + Kex

Equation 2.6.2

whereby R1a is the apparent T1 relaxation rate that is measured and R1 is the actual T1

relaxation rate, which is independent of the exchange rate.

The T1 values and subsequent imino proton exchange rates were also measured for the

16mer/15mer RNA-RNA complex at 15°C, using the same methods described above.

123

Chapter 3: NMR studies of the FMDV 16mer RNA and the effect

of Mg2+

Chapter 3 will firstly describe the NMR assignment and structure determination of the

16mer apo-RNA, and also include a conformational analysis of the NMR solution

structure. Subsequently, the focus of this chapter shifts to the effect of Mg2+

on the 16mer

RNA, which includes the effect on RNA chemical shifts, RNA stability and imino proton

exchange rates. Finally, the structure determination of the 16mer Mg2+

RNA complex is

detailed in a similar manner to the 16mer apo-RNA.

3.1 Structure determination of the 16mer apo-RNA

3.1.1 NMR assignment

The chemical shift table for the FMDV 16mer apo-RNA is displayed at the end of this

sub-section in Table 3.1.1. Chemical shifts were provided for all 1H,

13C and

31P nuclei

that were identified.

3.1.1.1 Exchangeable proton assignment

Imino proton identification and assignment

Six peaks were observed in the imino region of the 1H-NMR spectrum in

1H2O at 2°C,

between 10-15ppm (Figure 3.1.1). The initial assignment of the imino proton peaks was

based on knowledge of chemical shifts. Chemical shifts can be used to ambiguously

assign uracil and guanine imino proton peaks and most importantly distinguish between

stem and loop imino proton peaks. The imino proton peak at 10.54ppm was clearly shifted

to highfield of the stem imino proton peaks, meaning that it could only correspond to the

loop G178 or U179 loop imino protons. The presence of stem imino proton peaks found

between 12.0-15.0ppm, clearly showed that these bases were stacked in a double-helical

124

conformation. At this stage, the 1D 1H-NMR spectrum did not provide unambiguous

assignment of the uracil and guanine imino protons, so the NOESY (1H2O) spectrum was

analysed for accurate assignment. Therefore, the identification of imino proton peaks

shown in Figure 3.1.1 were made according to the assignment of the imino-imino region

of the NOESY (1H2O) spectrum.

The 16mer apo-RNA contains eight guanine and uracil bases, but only six imino proton

peaks were observed. The missing imino proton peaks were from the U172 and U179

bases. The most likely cause of the missing U172 imino proton peak is that the U172 base

has a higher exposure to water, possibly due to fraying of its terminal base pair.

Consequently, the imino proton peak intensity had reduced due to rapid exchange with

water. Similarly, as the U179 base is part of the GNRA tetraloop, there is a greater

probability that it will have higher exposure to water.


1H2O,

displaying the imino region. Six peaks were identified corresponding to the imino protons of U175,

U176, G185, G186, G177 and G178. The G178 loop imino proton peak can be clearly observed

highfield of the stem imino proton peaks.

125

Imino proton identification in the 1D 1H-NMR spectrum was aided by the interpretation of

the imino region in the 2D NOESY (1H2O) spectrum. Six diagonal peaks were found in

the imino region (Figure 3.1.2), corresponding to the six imino proton peaks found in the

1H-NMR spectrum. To start the identification, using a method of sequential assignment, at

least one peak must first be identified by making certain assumptions. Firstly, it was

assumed that the highfield loop imino proton peak corresponded to G178. This

assumption was based on the fact that G178 it is more likely to be stable than U179 due to

its ability to form a G.A sheared base pair. Secondly, uracil imino proton peaks are

generally found downfield from guanine imino proton peaks. Since there are two uracil’s

in the 16mer RNA (U175 and U176), it was assumed that the two most downfield peaks at

14.05ppm and 13.38ppm corresponded to the U175 and U176 imino protons, respectively.

Using this information the imino region was assigned by means of imino-imino sequential

assignment.

The imino-imino connectivities that were observed in the 16mer apo-RNA are shown in

Figure 3.1.2. An imino-imino sequential assignment was attained from G178 to G186

(G178-G177-U176-U175-G185-G186). The cross-diagonal peaks observed in the NOESY

(1H2O) spectrum correspond to NOE connectivities between two imino protons that were

close in space. NOE connectivities could not be observed to either the U172 or U179

imino protons. Interestingly, a highfield diagonal peak was found at 11.20ppm, which may

correspond to the U179 imino proton.

The observation of imino-imino connectivities confirmed two important features of the

16mer apo-RNA. Firstly, the presence of imino-imino connectivities indicated that imino-

imino distances were less than 5.0Å, which is characteristic of a stable, double-helical,

tertiary RNA structure. Secondly, the imino-imino sequential assignment observed in

Figure 3.1.2 partially confirmed that the 16mer RNA sequence is correct.

126

Figure 3.1.2 700 MHz NOESY (250ms) spectrum of the FMDV 16mer apo-RNA, at 2°C in 1H2O,

illustrating the imino region of the spectrum. The cross-diagonal peaks correspond to imino-imino

connectivities. The sequential assignment starts from G178, labelled in red, and finishes at G186,

labelled in blue. Inset: Secondary structure of the 16mer RNA highlighting the imino-imino

connectivities observed, represented by light blue oval shapes.

Imino-amino assignment

Once the imino proton chemical shifts were identified, the cross peaks in the imino-amino

region were assigned. In the imino-amino region, connectivities can be observed from

imino to the NH2*/NH2 amino protons (NH2* corresponds to the proton involved in base

pair hydrogen bonding), H2, H5, and H1ʹ protons. The previously identified imino proton

127

chemical shifts were used to assign cross peaks in the imino-amino region. The full

assignment of connectivities in the imino-amino region is shown in Figure 3.1.3. These

connectivities provided strong evidence of base pairing between C173-G186, C174-G185,

U175-A184, U176-A183 and G177-C182.


illustrating the imino-amino region of the spectrum. Connectivities from imino protons to

NH2*/NH2/H2/H5/H1ʹ protons can be observed (NH2* corresponds to the proton involved in base

pair hydrogen bonding); connectivities are marked with a black circle.

128

3.1.1.2 Non-exchangeable proton assignment

H5-H6 assignment

In the FMDV 16mer RNA, there are four uracil nucleotides and three cytosine nucleotides,

so seven H5-H6 cross peaks were expected to be observed in total. H5-H6 cross peaks are

usually very strong as the distance between H5-H6 protons is approximately 2.4Å.

The H5 and H6 proton chemical shifts of the 16mer apo-RNA, were identified from three

different spectra: NOESY in 1H2O, NOESY in

2H2O and

1H-

13C HSQC in

2H2O. Figure

3.1.4 illustrates the assignment of H5-H6 cross peaks observed in the NOESY (2H2O)

spectrum, with the aid of the 1H-

13C HSQC spectrum. The

1H-

13C HSQC spectrum allows

the identification of H5 and H6 proton chemical shifts due to the C5 and C6 chemical

shifts being distinctly different. Six H5 and H6 proton chemical shifts were identified in

the 1H-

13C HSQC spectrum, corresponding to U172, C173, C174, U175, U179 and C182.

Subsequently, six H5-H6 cross peaks were assigned in the NOESY (2H2O) spectrum

(Figure 3.1.4).

129


13C HSQC

spectrum and the subsequent assignment of H5-H6 cross peaks in the NOESY spectrum, of the

FMDV 16mer apo-RNA. Bottom left panel: 600 MHz NOESY (400ms) spectrum, at 25°C in 2H2O;

blue circles indicate H5-H6 cross peaks. Top left panel: 600 MHz 1H-

13C HSQC spectrum at 25°C

in 2H2O, displaying C6-H6 peaks. Bottom right panel: 600 MHz

1H-

13C HSQC spectrum at 25°C

in 2H2O, displaying C5-H5 peaks.

130

H6/H8-H1ʹ sequential assignment

The next step in the assignment strategy was to sequentially assign aromatic H6/H8

protons to H1ʹ sugar protons. The 1H-

13C HSQC spectrum was used to identify C8-H8,

C6-H6 and C1ʹ-H1ʹ peaks to aid the assignment of H6/H8-H1ʹ cross peaks in the NOESY

(2H2O) spectrum. C6-H6 peaks were first identified from the

1H-

13C HSQC spectrum, as

shown in Figure 3.1.4. C8-H8 peaks are located within a similar 1H and

13C chemical shift

range of C6-H6 peaks in 1H-

13C HSQC spectra. Therefore, H8 proton chemical shifts were

identified only when the C6-H6 peaks were labelled. Finally, C1ʹ-H1ʹ peaks were

identified from the 1H-

13C HSQC spectrum. With the combination of H6, H8 and H1ʹ

chemical shift values a sequential assignment was achieved by considering intranucleotide

and internucleotide H6/H8-H1ʹ connectivities observed in the NOESY (2H2O) spectrum. A

sequential assignment was attempted from the first nucleotide (U172) to the last

nucleotide (A187), in the 16mer RNA.

To start the sequential assignment an intranucleotide U172 H6-H1ʹ connectivity was

needed to be found, which was observed in the spectrum. No internucleotide connectivity

was observed between U172 H1ʹ and C173 H6. A sequential assignment was acquired

from C173 H6-H1ʹ to U175 H6-H1ʹ. No connectivity was observed between U175 H1ʹ

and A180 H8. Subsequently, A180 H8-H1ʹ to A183 H8-H1ʹ connectivities were added to

the partially acquired sequential assignment of the 16mer apo-RNA. Figure 3.1.5 displays

the sequential assignment discussed, and Figure 3.1.6 illustrates the use of the 1H-

13C

HSQC spectrum for identification and the NOESY (2H2O) spectrum for assignment.

131

.

Figure 3.1.5 Top/Bottom panels: 600 MHz NOESY (400ms) spectrum of the FMDV 16mer apo-

RNA, at 25°C in 2H2O. The blue line represents C173 H6–H1ʹ to U175 H6–H1ʹ intra- and

internucleotide connectivities, and the green line represents A180 H8-H1ʹ to A183 H8-H1ʹ intra-

and internucleotide connectivities; same colouring as in secondary structure shown. The (i)

corresponds to an intranucleotide connectivity. The red circles correspond to H5-H6 connectivities.

132

Figure 3.1.6 Illustration of the identification of C6-H6, C8-H8 and C1ʹ-H1ʹ peaks in the 1H-

13C

HSQC spectrum and the subsequent assignment of H6/H8-H1ʹ cross peaks in the NOESY

spectrum, of the FMDV 16mer apo-RNA. (a) 600 MHz 1H-


2H2O,

displaying the C6-H6 and C8-H8 peaks. (b,c) 600 MHz NOESY (400ms) spectrum at 25°C in

2H2O. (d,e) 600 MHz

1H-


2H2O, displaying the C1ʹ-H1ʹ peaks.

133

H1ʹ-H2ʹ and H2ʹ-H6/H8 sequential assignment

Strong intranucleotide NOE cross peaks were observed between H1ʹ-H2ʹ protons, which

are covalently fixed to 2.8-3.0 Å. The assignment of H1ʹ-H2ʹ cross peaks in the NOESY

(2H2O) spectrum was supported by the

1H-

13C HSQC spectrum (Figure 3.1.7). Known

chemical shift values of H1ʹ protons were used to identify correlations to H2ʹ protons,

observed in the sugar region of the NOESY (2H2O) spectrum. A strong internucleotide

NOE cross peak from the H2ʹ proton to the H6/H8 proton of the next nucleotide was

observed. Therefore, H2ʹ protons can be correlated to the sequential H6/H8 protons, and in

doing so they were used to confirm the H6/H8-H1ʹ sequential assignment.

Figure 3.1.7 Illustration of the assignment of intranucleotide H1ʹ-H2ʹ and internucleotide H2ʹ-

H6/H8 connectivities in the NOESY spectrum, with the aid of the 1H-


FMDV 16mer apo-RNA. Bottom panels: 600 MHz 1H-

13C HSQC spectra at 25°C in

2H2O. Top

panels: 600 MHz NOESY (400ms) spectra at 25°C in 2H2O.

134

Sugar proton assignment

Intranucleotide and internucleotide connectivities from H1ʹ to H2ʹ, H3ʹ, H4ʹ and H5ʹ/H5ʹʹ

protons were also assigned in the NOESY (2H2O) spectrum. However, as the H6/H8-H1ʹ

sequential assignment was incomplete, these assignments were difficult to obtain

unambiguously. The 1H-

13C HSQC spectrum was used to assist in the identification of the

sugar protons; C2ʹ/C3ʹ, C4ʹ and C5ʹ chemical shifts all appear in different regions of the

1H-

13C HSQC spectrum.

The DQF-COSY spectrum was used to identify intranucleotide H1ʹ-H2ʹ cross peaks. In

RNA, the sugar pucker conformation is normally C3ʹ-endo, which produces H1ʹ-H2ʹ

coupling constants of approximately 2Hz or less. The positive and negative parts of the

multiplet structure of the cross peaks cancel out as the coupling constant is less than the

linewidth. This means that H1ʹ-H2ʹ cross peaks that are not observed in the DQF-COSY

spectrum, are characteristic of the C3ʹ-endo sugar conformation. Observed H1ʹ-H2ʹ cross

peaks are assumed to be characteristic of the C2ʹ-endo sugar conformation. Figure 3.1.8

displays the H1ʹ-H2ʹ cross peaks observed in the DQF-COSY spectrum, which were

confirmed by the 1H-

13C HSQC spectrum. The U172, A180 and A187 H1ʹ-H2ʹ cross peaks

were clearly observed in the DQF-COSY spectrum of the 16mer apo-RNA. The U172 and

A187 nucleotides are found at the terminus of the 16mer RNA, and so are more likely to

adopt sugar conformations that conform to C2ʹ-endo. Similarly, it was expected that the

loop nucleotides may also conform to the C2ʹ-endo sugar conformation. The DQF-COSY

spectrum provided evidence that the loop A180 nucleotide did not conform to the C3ʹ-endo

sugar conformation, but more likely had the C2ʹ-endo sugar conformation in the 16mer

apo-RNA.

135

Figure 3.1.8 Illustration of the assignment of intranucleotide H1ʹ-H2ʹ connectivities in the DQF-

COSY spectrum, with the aid of the 1H-

13C HSQC spectrum, of the FMDV 16mer apo-RNA. Top

right panel: 600 MHz 1H-


2H2O, representing the C1ʹ-H1ʹ

resonances. Top left panel: 400 MHz DQF-COSY spectrum at 25°C in 2H2O, representing the H1ʹ-

H2ʹ cross peaks. Bottom left panel: 600 MHz 1H-


2H2O,

representing the C2ʹ-H2ʹ resonances.

136

Phosphorus identification and assignment

The 1H-

31P CPMG-HSQC-NOESY experiment was used to observe connectivities

between phosphorus and proton nuclei. This experiment involved both scalar and dipolar

coupling; scalar coupling between the phosphorus nuclei to H3ʹ/H5ʹ/H5ʹʹ protons, and

dipolar coupling between the H3ʹ/H5ʹ/H5ʹʹ protons to other sugar and base protons. The

H6/H8 protons and the H1ʹ protons were identified relative to the phosphorus chemical

shift, which aided in confirming the H6/H8-H1ʹ sequential assignment as well as

identifying proton chemical shifts that had not been previously identified. The most

intense peaks corresponded to the scalar correlation between phosphorus nuclei and the

H3ʹ, H5ʹ, H5ʹʹ protons. These protons gave strong intranucleotide NOEs to nearby sugar

protons in the same nucleotide. Both intra- and internucleotide NOEs were observed to

H6/H8 and H1ʹprotons, but these NOEs were much weaker in intensity.

Since most of the H6/H8 and H1ʹ chemical shifts had been identified in the 16mer apo-

RNA, this information was used to identify the corresponding correlations to phosphorus

chemical shifts. In the 1H-

31P CPMG-HSQC-NOESY spectrum, four H6/H8 and two H1ʹ

correlations to phosphorus were assigned, using the 1H-

13C HSQC spectrum (Figure 3.1.9).

Subsequently, three phosphorus chemical shifts were identified for the C173, C174 and

C182 nucleotides. Due to the lack of clearly observed peaks in the spectrum, it was not

possible to obtain a sequential assignment between phosphorus and H6/H8/H1ʹprotons.

137

Figure 3.1.9 Illustration of the assignment of phosphorus to H6/H8/H1ʹ peaks in the 1H-

31P

CPMG-HSQC-NOESY spectrum, with the aid of the 1H-


16mer apo-RNA. Bottom left panel: 600 MHz 1H-

31P CPMG-HSQC-NOESY (500ms) spectrum at

25°C in 2H2O, representing the phosphorus-H6/H8 peaks. Bottom right panel: 600 MHz

1H-

31P

CPMG-HSQC-NOESY (500ms) spectrum at 25°C in 2H2O, representing the phosphorus-H1ʹ

peaks. Top left panel: 600 MHz 1H-


2H2O, representing C6-H6 and

C8-H8 peaks. Top right panel: 600 MHz 1H-


2H2O, representing

C1ʹ-H1ʹ peaks.

138

Identification of proton resonances using 3D NMR

A 3D NOESY/2Q-COSY experiment was performed to reduce the complication of

overlapping peaks in the 2D NOESY (2H2O) spectrum. Two different illustrations of the

3D NMR spectrum are shown (Figures 3.1.10 and 3.1.11) for the identification of proton

chemical shifts, corresponding to two different planes; the F3/F1 and F3/F2 planes. The

F3/F1 plane was used to identify the base and sugar protons of the U172 and C173

nucleotide in the 16mer apo-RNA, shown in Figures 3.1.10a and 3.1.10b, respectively.

Figure 3.1.10a displays the F3/F1 NOE plane chosen at a specific F2 frequency (9.422

ppm) that allows the observation of coupling between the U172 H5 and H6 protons.

Strong inner NOE cross peaks were observed between the U172 H5 and H6 protons.

Weaker outer NOEs were observed to other sugar/base protons, which included U172 H1ʹ,

U172 H3ʹ, U172 H4ʹ, U172 H5ʹ and U172 H5ʹʹ. Figure 3.1.10b displays the F3/F1 NOE

plane chosen at a specific F2 frequency (9.36ppm) that allows the observation of coupling

between the C173 H5 and H6 protons. Strong inner NOE cross peaks were observed

between the C173 H5 and H6 protons. Weaker outer NOEs were observed to other

sugar/base protons, which included C173 H1ʹ, C173 H5, C173 H2ʹ, C173 H3ʹ, C173 H5ʹ

and C173 H5ʹʹ. In both cases, the non-exchangeable H5/H6 base protons and sugar protons

in the U172 and C173 nucleotides were unambiguously identified.

The F3/F2 plane was used to identify the base and sugar protons of the C174 nucleotide in

the 16mer apo-RNA. Figure 3.1.11 displays the different F1 slices in the F3/F2 plane of

the 3D NMR spectrum. F1 frequencies were chosen depending on the chemical shifts of

C174 H5ʹ, H5ʹʹ, H4ʹ, H3ʹ, H5 and H6 protons. In this case, intra- and internucleotide base

and sugar protons in the C174 nucleotide were clearly identified.

This 3D NMR experiment has allowed the successful identification of several sugar

proton resonances in the 16mer apo-RNA. It clearly demonstrates the powerful advantage

of using 3D NMR techniques, without the necessity for labelled RNA samples.

139

Figure 3.1.10 600 MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer apo-RNA.

Identification of base and sugar protons of the (a) U172 nucleotide and (b) C173 nucleotide,

illustrated by the F3/F1 NOE plane. Both positive (orange) and negative (green) levels are shown.

140

Figure 3.1.11 600 MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer apo-RNA.

Slices from the F3/F2 plane chosen at different F1 frequencies, identifying base and sugar proton

chemical shift of the C174 nucleotide. Only positive levels are shown. In each slice strong inner

NOE peaks correspond to coupling between two protons as well as weaker outer NOEs to other

base/sugar protons.

141

Table 3.1.1 1H,

13C and


1H2O and

2H2O.

NH NH2* NH2 H2 H5 H6 H8 H1ʹ H2ʹ H3ʹ H4ʹ H5ʹ H5" C2 C5 C6 C8 C1ʹ C2ʹ C3ʹ C4ʹ C5ʹ 31P

U172 13.79 5.935 8.156 5.721 4.642 4.599 4.399 3.949 4.074 104.4 144.0 93.23 75.37 74.25 85.29 62.15 C173 8.606 7.228 5.943 8.132 5.732 4.504 4.619 4.540 4.228 4.565 97.96 142.5 94.34 75.56 72.62 86.76 65.23 -4.255

C174 8.471 7.014 5.651 7.923 5.523 4.421 4.562 4.468 4.601 4.145 97.64 141.8 94.34 75.39 75.70 81.88 64.57 -4.579

U175 14.05 5.458 7.950 5.545 4.459 4.750 4.417 3.901 3.844 103.3 142.3 94.08 72.67 87.92 63.89

U176 13.38 5.806 7.899 5.648 93.21

G177 12.28 8.113 5.760 5.783 92.64

G178 10.53 5.983 5.150 91.58 80.20

U179 11.20 5.499 7.794 6.131 4.285 4.498 4.064 104.0 143.3 91.75 65.62

A180 7.846 8.051 5.671 4.376 4.592 4.251 3.896 3.821 154.4 140.6 92.02 76.81 75.65 83.58 65.87

A181 8.135 8.165 6.060 4.528 5.129 5.428 155.0 141.3 92.06 76.86 74.09 83.1

C182 8.522 7.056 6.066 7.771 3.773 4.364 4.173 4.303 4.222 99.42 141.9 94.79 74.47 82.71 68.93 -3.038

A183 7.633 6.545 6.704 8.044 5.864 4.513 4.787 4.365 4.109 4.498 152.2 139.8 92.91 75.65 72.53 86.24 64.62

A184 7.876 6.641 7.405 5.888 4.568 4.560 152.9 92.35 72.34 72.17

G185 12.90 8.310 6.109 5.528 4.484 4.424 4.031 92.37 75.42 65.64

G186 12.63 8.049 5.685 4.484 4.434 4.067 93.21 75.42 72.93 62.15 -4.330

A187 8.086 7.802 6.082 4.125 4.293 4.299 4.128 154.7 140.1 91.75 78.0 83.94 70.36

142


Distance restraints were generated from NOESY spectra of the 16mer apo-RNA.

Exchangeable NOE restraints were obtained from the NOESY spectrum of the 16mer apo-

RNA in 1H2O, with 150ms mixing time. The exchangeable NOE restraints consisted of

imino-imino connectivities, imino-amino connectivities and imino-H2/H5 connectivities.

Non-exchangeable NOE restraints were obtained from the NOESY spectrum of the 16mer

apo-RNA in 2H2O, with 250ms mixing time. The non-exchangeable NOE restraints

consisted of H5-H6 connectivities, H6/H8-H1ʹ intra- and internucleotide connectivities,

H1ʹ-H5 connectivities, aromatic to aromatic connectivities, H1ʹ to sugar and sugar to

H6/H8 connectivities.

The absence of strong intranucleotide H6/H8-H1ʹ cross peaks for stem nucleotides,

allowed the glycosidic angle of all stem nucleotides to be restrained to the anti

conformation. No intranucleotide connectivities were observed for G178 and U179, so

their glycosidic angles were not defined. A180 and A181 were also not defined to allow

some conformational flexibility in the loop region. In the DQF-COSY experiment, the

U172 and A187 H1ʹ-H2ʹ cross peaks were clearly observed. Therefore, the δ angle was

restrained to the C2ʹ-endo conformation for U172 and A187 and the other stem nucleotides

were restrained to the C3ʹ-endo conformation. The δ angle for the loop nucleotides was not

restrained due to sugar pucker dynamics found in GNRA tetraloops.44

The α, β, γ, ε and ζ

dihedral angles were solely defined for the stem nucleotides, except for U172 and A187.

The imino proton chemical shifts resonate lowfield due to the ring current effect caused by

base stacking interactions of adjacent base pairs in the helical stem. The only exception

was the G178 loop imino proton. Therefore, the bases corresponding to the stem uracil

and guanine imino proton peaks were restrained as canonical base pairs. Hydrogen bond

restraints were added for all six base pairs. However, since the U172 imino proton peak

was not observed and no connectivities from it were found in the NOESY (1H2O)

spectrum, the hydrogen bond restraints between U172-A187 were loosely restrained. For

143

the same reason, planarity restraints were added for all base pairs in the stem, except for

the U172-A187 base pair.

Distance, dihedral angle, hydrogen bond and planarity restraints were added to the

structure calculation. From the 20 lowest RMSD structures generated from the refinement

step, no violations were found in the distance restraints and hydrogen bond restraints. The

restraints that were used for these structure calculations are summarised in Table 3.1.2.

Restraints

NOEs 125

Strong 1.8 – 2.5 Å 9

Medium 2.6 – 3.3 Å 39

Weak 3.4 – 5.0 Å 77

Very weak 5.1 – 7.0 Å 0

Internucleotide NOEs 71

Intranucleotide NOEs 54

Hydrogen bonds 36

Planarity 10

Dihedral angles 98

Helix (, , , ε, ζ) 50

Ribose pucker (, ν1, ν2) 36

Glycosidic (χ) 12

NOE per residue 7.8

Restraints per

residue

16.8

Total restraints 269

Table 3.1.2 A summary of the total number of restraints used for the structure determination of the

FMDV 16mer apo-RNA.

144

3.1.3 NMR solution structure

3.1.3.1 Ensemble and final structure

The 20 lowest RMSD structures generated by the refinement step were aligned by

considering all atoms (Figure 3.1.12a). The average RMSD of all 20 structures was

calculated to be 0.18Å.

Figure 3.1.12 Illustration of the NMR solution structures of the FMDV 16mer apo-RNA (a)

Overlay of the 20 lowest RMSD structures, with an average RMSD of 0.18Å. (b) Lowest RMSD

solution structure (0.17Å); the red ribbon represents the RNA backbone.

The overlay of the 20 lowest RMSD structures displays the overall conformational

homogeneity of the 16mer apo-RNA tertiary structure. An important observation was that

the helical stem region and tetraloop region were aligned very well, despite the dynamic

nature of the tetraloop region. The tetraloop region has been significantly restrained by

internucleotide connectivities, which has the effect of reducing the RMSD of the loop

region. When considering the helical stem and the tetraloop separately, there was no

145

significant difference in RMSD. The average RMSD of the helical stem alone was 0.178Å

and of the tetraloop alone was 0.183Å. From the ensemble of 20 structures, the structure

with the lowest RMSD (0.17Å) was selected for structure and conformational analysis

(Figure 3.1.12b).

3.1.3.2 GNRA tetraloop

The tertiary conformation of the G178UAA181 tetraloop is displayed in Figure 3.1.13. The

overall geometry was similar to that found in previously studied GNRA tetraloops.43-44

GNRA tetraloops adopt an asymmetric loop structure, where only G of GNRA is stacked

on the 5ʹ side and NRA is stacked on the 3ʹ side of the stem. A sharp turn was found

between the G178 and U179 nucleotides, which allows U179 to stack on the 3ʹ side of the

stem. Hence the U179, A180 and A181 nucleotides in the GNRA tetraloop were in a loose

stacking formation. The term ‘loose’ is used here because the U179, A180 and A181 bases

do not stack directly on top of each other, unlike the bases in the helical stem. This base

stacking formation would significantly contribute to the stability of the tetraloop.

Figure 3.1.13 The G178UAA181 tetraloop of the FMDV 16mer apo-RNA NMR structure, shown in

Figure 3.1.12b; the closing G177.C182 base pair is also illustrated. Colour coding of nucleotides:

guanosine (blue), uridine (cyan), adenosine (green) and cytosine (red).

146

3.1.3.3 Intramolecular interactions

In total, ten intramolecular interactions were found in the GUAA tetraloop of the 16mer

apo-RNA NMR structure. Two specific, base-base interactions were found between G178

and A181, corresponding to the two hydrogen bonds found in the G.A sheared base pair;

G178 NH2-A181 N7 and G178 N3-A181 NH6 were found to be 3.09Å and 5.05Å,

respectively (Figure 3.1.14). The G178 and A181 bases do not adopt the correct

orientation for G.A sheared base pairing, which is why one hydrogen bond distance is too

large for hydrogen bonding.

Figure 3.1.14 The G.A sheared base pair in the FMDV 16mer apo-RNA NMR structure.

Hydrogen bonding distances between G178 NH2-A181 N7 (3.09Å) and G178 N3-A181 NH6

(5.05Å) are indicated by the red broken lines.

One specific, base-phosphate interaction was found in the GUAA tetraloop; G178 NH2-

A181 OP at 2.11Å (Table 3.1.3). In particular, this base-phosphate interaction is

conserved in most GNRA tetraloops. Base-phosphate interactions that involve the N-H

donor group, almost always have the anionic OP oxygen as the acceptor. This type of

hydrogen bond forms the strongest of all base-phosphate interactions and so will

significantly contribute to the stabilisation of the GNRA tetraloop. This explains why this

specific base-phosphate interaction is so highly conserved in GNRA tetraloops.

147

Furthermore, two specific, base-base interactions were found between U179 O4-A180

NH6. Here the O4 oxygen of U179 is able to interact with both the amines in A180 with

distances of 2.30Å and 2.94Å (Table 3.1.3). One specific, base-sugar interaction was

identified in the GUAA tetraloop; G178 NH2-A180 O2ʹ at 2.49Å (Table 3.1.3).

Although specific interactions will play a large role in stabilising the GUAA tetraloop,

non-specific interactions will also play a significant part in stabilisation. Four non-specific,

intranucleotide base-phosphate interactions were found for G178, U179, A180 and A181

(Table 3.1.3).

Donor/acceptor atoms Type of

interaction

Specificity Distance (Å)

G178 NH2 – A181 N7 base-base specific 3.09

G178 N3 – A181 NH6 base-base specific 5.05

G178 NH2 – A181 OP base-phosphate specific 2.11

G178 NH2 – A180 O2ʹ base-sugar specific 2.49

G178 H8 – G178 O5ʹ base-phosphate non-specific 3.05

U179 O4 – A180 NH6 base-base specific 2.30

U179 O4 – A180 NH6 base-base specific 2.94

U179 H6 – U179 O5ʹ base-phosphate non-specific 2.84

A180 H8 – A180 O5ʹ base-phosphate non-specific 3.08


Table 3.1.3 Ten intramolecular interactions in total were identified in the GUAA tetraloop of the

FMDV 16mer apo-RNA NMR structure. The interactions formed between donor and acceptor

atoms are given, the type of interaction, the specificity of the interaction and the distances between

the proton donor and acceptor atoms.

148


The 3DNA program was used to perform a conformational analysis on the final 16mer

apo-RNA NMR structure. The local helical parameters, base pair step parameters and

complementary base pair parameters were calculated and are displayed in Tables 3.1.4,

3.1.5 and 3.1.6, respectively.

When analysing the 16mer apo-RNA structure, it was first determined whether the RNA

helical structure did indeed exhibit an A-form conformation. The two best parameters that

can be used to distinguish between A-form and B-form conformations are the ‘x-

displacement’ (local helical parameter) and ‘slide’ (base pair step parameter). Values

obtained for the 16mer apo-RNA were compared with average values obtained for A-

DNA and B-DNA crystal structures.110

Both the ‘x-displacement’ value of -3.19Å (Table

3.1.4) and ‘slide’ value of -1.40Å (Table 3.1.5), suggested that the 16mer apo-RNA

structure is characteristic of an A-form helical conformation.

One other parameter that may be useful for analysing the conformation of the tetraloop in

the 16mer apo-RNA structure is the complementary base pair parameter, ‘shear’. Since the

G.A base pair in the tetraloop is a sheared base pair, the ‘shear’ parameter value of the

G.A base pair can be compared with the other canonical base pairs. The ‘shear’ parameter

value of the G.A base pair was found to be 7.67Å (Table 3.1.6). In contrast to the average

canonical base pair values (-0.05Å), the value of 7.67Å confirms the sheared conformation

of the G.A base pair.

The base pairing between U172-A187 could not be confirmed by NMR and the final

NMR solution structure clearly displayed a frayed base pair. This is shown by the

deviation from average values, consistent with standard base pairing, of the

complementary base pair parameters ‘propeller’ and ‘opening’ (Table 3.1.6).

149

Base

Pair Nucleotides

Xdisp

(dx)

Ydisp

(dy)

Inclination

(η)

Tip

(θ)

Helical Twist

(Ω)

Helical

Rise (h)

1-2 U-A / C-G -0.48 -2.90 -12.93 20.83 36.28 3.72

2-3 C-G / C-G -5.90 -1.23 18.75 -0.66 26.49 2.15

3-4 C-G / U-A -2.90 0.80 6.50 -0.92 33.56 2.97

4-5 U-A / U-A -1.40 -1.76 -7.51 -2.07 32.25 4.01

5-6 U-A / G-C -7.71 0.55 32.54 -2.63 25.79 1.56

6-7 G-C / G-A -0.75 1.88 4.02 2.62 63.99 3.05

Ave. - -3.19 -0.44 6.79 2.86 36.39 2.91

Table 3.1.4 Local helical parameter values for the FMDV 16mer apo-RNA structure, calculated

by the 3DNA analysis program.

Base

Pair Nucleotides

Shift

(Dx)

Slide

(Dy)

Rise

(Dz)

Tilt

(τ)

Roll

(ρ)

Twist

(ω)

1-2 U-A / C-G 0.45 -1.14 3.96 -12.61 -7.83 33.19

2-3 C-G / C-G 0.59 -1.92 2.87 0.30 8.18 25.22

3-4 C-G / U-A -0.41 -1.34 3.14 0.53 3.75 33.36

4-5 U-A / U-A 1.12 -1.27 3.84 1.15 -4.16 31.97

5-6 U-A / G-C -0.14 -2.10 3.15 1.11 13.78 21.82

6-7 G-C / G-A -2.11 -0.62 3.01 -2.78 4.26 63.81

Ave. - -0.08 -1.40 3.33 -2.05 3.00 34.89

Table 3.1.5 Base pair step parameter values for the FMDV 16mer apo-RNA structure, calculated

by the 3DNA analysis program.

Base

Pair Nucleotides

Shear

(Sx)

Stretch

(Sy)

Stagger

(Sz)

Buckle

(κ)

Propeller

(π)

Opening

(σ)

1 U-A -0.65 -0.26 -1.55 -11.52 37.14 -23.50

2 C-G 0.04 -0.25 -0.04 -11.16 -13.07 -4.48

3 C-G -0.15 -0.12 0.08 -3.72 -13.78 0.80

4 U-A 0.06 -0.09 0.28 1.22 -25.87 -2.57

5 U-A 0.08 -0.10 -0.26 2.13 2.75 -1.62

6 G-C 0.02 -0.14 0.13 4.46 5.45 -2.23

7 G-A 7.67 -6.79 1.27 28.09 12.93 -35.78

Table 3.1.6 Complementary base pair parameter values for the FMDV 16mer apo-RNA structure,

calculated by the 3DNA analysis program.

150

Dihedral angles that define nucleotide structure were calculated using both the 3DNA and

CURVES conformational analysis programs (Table 3.1.7). The 3DNA program was used

to generate dihedral angle values for nucleotides of the six stem base pairs, and for the

loop G178 and A181 nucleotides. Fourteen dihedral angle values were acquired by the

3DNA program. To obtain the dihedral angle values of U179 and A180 loop nucleotides,

the CURVES program was used. Notably, the δ angle for the loop nucleotides was not

restrained, but revealed a C3ʹ-endo conformation for G178, U179 and A181, and a C2ʹ-

endo conformation for A180. Evidence for the C2ʹ-endo conformation was found for A180

in the DQF-COSY and 3D NOESY/2Q-COSY spectra. The most significant deviation in

dihedral angles were found for the χ angle of A180, the ζ angle of U172 and the α angle of

U179 nucleotide. The deviation in the α angle of U179 is due to the rotation around the P-

O5ʹ bond that forms the U-turn motif found in GNRA tetraloops.

No. Nucleotide C1ʹ-N

(χ)

C5ʹ-C4ʹ

(γ)

C4ʹ-C3ʹ

(δ)

C3ʹ-O3ʹ

(ε)

O3ʹ-P

(ζ)

P-O5ʹ

(α)

O5ʹ-C5ʹ

(β)

1 U -130.7 54.3 149.6 -178.0 -102.2 - -

2 C -161.5 53.8 82.2 -156.3 -63.1 -66.7 172.4

3 C -158.0 57.0 81.7 -161.6 -76.8 -64.4 169.0

4 U -151.9 54.1 81.3 -155.5 -65.6 -66.9 170.9

5 U -152.9 55.4 81.8 -158.2 -69.7 -64.5 171.0

6 G -154.4 53.1 82.1 -152.8 -61.0 -64.5 174.9

7 G -146.3 53.1 81.9 -155.5 -70.1 -69.2 172.8

8 U -159.6 55.2 84.4 -146.0 -65.9 148.3 164.8

9 A -119.8 49.6 140.1 -162.7 -91.0 -68.0 -174.8

10 A -154.4 54.4 83.8 -153.1 -52.5 -69.0 166.4

11 C -157.3 67.5 88.4 -156.9 -61.1 -70.1 156.7

12 A -168.3 59.4 84.4 -161.1 -89.1 -65.2 169.7

13 A -156.6 56.1 84.4 -153.4 -65.0 -64.2 170.4

14 G -155.3 54.3 82.6 -156.0 -67.7 -65.5 171.2

15 G -154.1 55.4 82.8 -160.7 -72.0 -65.1 171.5

16 A -112.6 52.8 148.4 - - -63.4 -173.5


calculated by the 3DNA (black) and CURVES (red) analysis programs. Angles are all measured in

degrees.

151

Dihedral angles (ν1 and ν2) that define the sugar ribose conformations were also calculated

by the 3DNA and CURVES programs, including the parameters pseudorotation phase

angle and the amplitude (Table 3.1.8). Based on these values, the calculated sugar ribose

conformation is also given. These values were very useful in cross-checking the dihedral

angle restraints, which were used in the 16mer apo-RNA structure calculations.

Six out of the sixteen nucleotides shown in Table 3.1.8, were calculated to have a C3ʹ-endo

sugar ribose conformation. The C4ʹ-exo conformation was revealed for six nucleotides,

three C2ʹ-endo and one C2ʹ-exo conformations were found. The six nucleotides whose

sugar ribose adopted a C4ʹ-exo conformation, did not deviate significantly from the C3ʹ-

endo conformation. The C2ʹ-endo conformation was found for U172, A180 and A187. The

C2ʹ-exo conformation was found for the C182 nucleotide, which is very close to the phase

angle of the C3ʹ-endo conformation.

No. Nucleotide v1 v2 Amp Phase Conformation

1 U 33.4 -36.3 36.7 172.5 C2ʹ-endo

2 C -20.4 36.2 40.0 25.4 C3ʹ-endo

3 C -9.4 29.8 39.9 41.7 C4ʹ-exo

4 U -20.7 37.5 41.8 26.2 C3ʹ-endo

5 U -16.9 34.9 40.7 31.1 C3ʹ-endo

6 G -7.1 28.4 40.8 46.0 C4ʹ-exo

7 G -14.8 31.5 37.8 33.5 C3ʹ-endo

8 U -14.4 30.3 36.9 32.3 C4ʹ-exo

9 A 33.8 -30.9 34.4 157.4 C2ʹ-endo

10 A -12.5 30.5 38.0 36.6 C4ʹ-exo

11 C -34.3 41.0 41.0 359.3 C2ʹ-exo

12 A -11.0 28.6 36.5 38.3 C4ʹ-exo

13 A -32.2 41.0 41.1 5.2 C3ʹ-endo

14 G -25.0 37.8 39.6 17.2 C3ʹ-endo

15 G -11.5 29.6 37.5 38.0 C4ʹ-exo

16 A 38.4 -37.8 39.5 163.0 C2ʹ-endo

Table 3.1.8 Dihedral angle values (ν1 and ν2), pseudorotation phase angle (Phase) and amplitude

(Amp) values that define the sugar ribose conformation for each nucleotide in the FMDV 16mer

apo-RNA structure. Values were calculated by the 3DNA (black) and CURVES (red) analysis

programs.

152

3.2 Effect of Mg2+

on 16mer RNA chemical shifts

In this section, the results obtained from the Mg2+

titration of the FMDV 16mer RNA

(batch 1) will be discussed. 400 MHz 1H-NMR and 162 MHz

31P-NMR spectra were

measured in the absence of Mg2+

and in the presence of varying Mg2+

concentrations, in

1H2O. Both

1H-NMR and

31P-NMR experiments were initially performed at 5ºC in the

absence of Mg2+

and with 0.5eq of Mg2+

. Subsequently, 1H-NMR and

31P-NMR

experiments were performed at 2ºC, including the repetition of the 0.5eq titration point, in

order to reduce imino proton exchange and increase signal intensity. Both 400 MHz and

700 MHz NOESY experiments were also performed on the 16mer RNA, in the absence

and presence of 5.0eq (6.05mM) Mg2+

. The imino-imino, imino-amino and aromatic

regions were analysed to discover any differences in NOE patterns and intensity. Changes

in these two parameters would be indicative of conformational change in the tertiary RNA

structure.

3.2.1 Changes in proton chemical shift

A 1H-NMR stack plot of the imino region, at different titration points, is shown in Figure

3.2.1. The imino proton region is shown illustrating the stem (U175, U176, G185, G186,

G177) and the loop (G178), imino proton peaks.

Interestingly, upon addition of 5.0eq of Mg2+

, a large lowfield chemical shift change was

observed for the G178 (Δδ=0.30ppm) and G177 (Δδ =0.11ppm) imino proton peaks

(Figure 3.2.2). Both the G178 and G177 chemical shift changes were significant and

clearly provided evidence that Mg2+

was having an effect in the loop region. These

chemical shift changes could be attributed to changes in base stacking or to direct binding

of Mg2+

. No significant chemical shift changes were observed for the U175, U176, G185

and G186 imino proton peaks. Figure 3.2.3 plots the changes in imino proton chemical

shifts observed in 1H-NMR, at different Mg

2+ concentrations. Changes in chemical shift

observed for U175, U176, G185 and G186 imino proton peaks are not shown as the values

are below the detectable range of ≥0.05ppm.

153

Another interesting observation was found when comparing the imino region with

increasing Mg2+

concentration, which is the slight increase in line broadening. Line

broadening from addition of Mg2+

may be caused by RNA aggregation or intermediate

chemical exchange from non-specific and diffuse Mg2+

ions.

Figure 3.2.1 A stack plot of 400 MHz 1H-NMR spectra (imino region) of the FMDV 16mer RNA

in 1H2O, with increasing Mg

2+ concentration. Each

1H-NMR spectrum is labelled 1-6; 1 (0eq -

5°C), 2 (0.5eq - 5°C), 3 (0.5eq - 2°C), 4 (1.0eq - 2°C), 5 (2.0eq - 2°C), 6 (5.0eq - 2°C). The U175,

U176, G185, G186, G177 and G178 imino proton peaks are labelled accordingly.

154

Figure 3.2.2 700 MHz 1H-NMR spectra (imino region) of the FMDV 16mer RNA, in

1H2O at 2°C;

(a) no Mg2+

and (b) containing 5eq of Mg2+

. The large Mg2+

-induced chemical shift change of

0.30ppm and 0.11ppm to G178 and G177, respectively, is clearly identified.

Figure 3.2.3 Histogram illustrating the imino proton chemical shift changes observed in 1H-NMR

spectra, at different Mg2+

concentrations; 0.5eq (red), 1.0eq (blue), 2.0eq (green) and 5.0eq

(orange).

155

3.2.2 Changes in phosphorus chemical shift

The 31

P-NMR spectra were also analysed to study the effect of Mg2+

on the 16mer RNA

structure. Phosphorus chemical shifts are very sensitive to the conformation of RNA, since

the phosphorus is located in the RNA backbone. Therefore, the changes in phosphorus

chemical shifts will signify a change in the RNA phosphate backbone. Phosphorus peaks

were assigned based on 1H-

31P CPMG-HSQC-NOESY data of both the 16mer apo-RNA

and Mg2+

RNA complex.

A 31

P-NMR stack plot, at different titration points, is shown in Figure 3.2.4. Nine

phosphorus peaks were identified in the absence of Mg2+

, excluding the large phosphorus

solvent peak. Peaks 3-9 were observed between -3.1ppm and -4.2ppm, which largely

consist of stem phosphorus peaks. Peaks 1 and 2 were observed significantly more

lowfield at -1.21ppm and -1.88ppm, respectively. Peaks 1, 2 and 9 were identified to be

from U179, A180/C182 and A181, respectively. Interestingly, the U179 and A180/C182

phosphorus peaks were observed significantly more lowfield of the stem phosphorus

peaks, while the A181 phosphorus peak was found to be the most highfield.

Upon addition of Mg2+

, the most significant chemical shift changes were observed for

U179 (lowfield Δδ=0.33ppm) and A181 (highfield Δδ=0.31ppm) (Figure 3.2.5). Mg2+

-

phosphate interactions generally produce a lowfield 31

P chemical shift change due to

deshielding of the phosphorus nucleus.113

However, changes in structure conformation,

such as increased base stacking interactions, can lead to a simultaneous highfield shift.

Therefore, the lowfield shift of the U179 phosphorus suggests that Mg2+

is directly

coordinating with the U179 phosphate group, possibly in the form of chelated ions.

Conversely, the highfield shift observed for A181 suggests conformational change in the

A181 phosphate group. It is possible that the highfield shift may also be caused by

intramolecular interactions with the phosphate oxygen of A181. Highfield chemical shift

changes were also observed for all other phosphorus peaks, which indicated Mg2+

-induced

structural changes to the entire 16mer RNA phosphate backbone.

156

Figure 3.2.4 162 MHz stack plot of 31

P-NMR spectra of the FMDV 16mer RNA in 1H2O, with

increasing Mg2+

concentration. Each 31

P-NMR spectrum is labelled a-f; a (0eq - 5°C), b (0.5eq -

5°C), c (0.5eq - 2°C), d (1.0eq - 2°C), e (2.0eq - 2°C), f (5.0eq - 2°C). Peaks are labelled 1-9.

Figure 3.2.5 Histogram illustrating the phosphorus chemical shift changes observed in 31

P-NMR,

at different Mg2+

concentrations; 0.5eq(red), 1.0eq (blue), 2.0eq (green), 5.0eq (orange). The peak

numbers 1-9 correspond to the labelling introduced in Figure 3.2.4. Positive chemical shift

changes represent a lowfield shift and negative chemical shift changes represent a highfield shift.

157

3.3 Effect of Mg2+

on 16mer RNA stability

In this section, the results obtained from the variable temperature (VT) series of the

FMDV 16mer apo-RNA and Mg2+

RNA complex will be discussed. 400 MHz 1H-NMR

and 162 MHz 31

P-NMR VT spectra of the 16mer apo-RNA and Mg2+

RNA complex were

analysed. Additionally, 700 MHz 1H-NMR VT spectra were analysed for the 16mer Mg

2+

RNA complex only.

3.3.1 1H-NMR variable temperature (VT) series

The imino region of the 1H-NMR VT series was investigated for both the 16mer apo-RNA

and Mg2+

RNA complex, at two different temperatures of 5°C and 35°C (Figure 3.3.1).

The loop G178 imino proton peak could not be observed clearly in the 16mer apo-RNA or

Mg2+

RNA complex and so the effect of Mg2+

on the G178 imino proton could not be

assessed. The most significant difference in peak intensity was found for the G177 and

G186 imino proton peaks. The G186 and G177 imino proton peaks were clearly exchange

retarded in the presence of Mg2+

demonstrating the enhanced thermodynamic stability

conferred by Mg2+

. Fascinatingly, at a higher temperature of 45°C, all the imino protons of

the apo-RNA were absent but were clearly observable in the presence of Mg2+

(data not

shown). These results strongly suggest a role for Mg2+

in increasing the stability of base

pairing in the stem of the 16mer RNA structure. This increase in stability is mainly

apparent in the G177 and G186 base pairs, which are highly susceptible to temperature-

induced destabilisation. These findings prompted the measurement of imino proton

exchange rates in order to quantify the effect of Mg2+

(section 3.4).

The change in imino proton chemical shifts with temperature was analysed, for both the


RNA complex. Most notably, for the 16mer apo-RNA, the

largest chemical shift changes were found for U176 (highfield Δδ=0.24ppm) and U175

(highfield Δδ=0.10ppm), between 5°C and 35°C (data not shown). This suggests that the

two A.U base pairs, involving U175 and U176, are more susceptible to temperature-

induced conformational changes. It was also observed that the U175, U176 and G185

158

imino proton chemical shift changes were not as large in the Mg2+

RNA complex when

compared to the apo-RNA (data not shown). This suggests a Mg2+

-induced stabilisation of

the tertiary conformation involving the stem U175, U176 and G185 base pairs.

Figure 3.3.1 400 MHz 1H-NMR spectra of the FMDV 16mer RNA in

1H2O at two different

temperatures of 5°C and 35°C. (a) 16mer apo-RNA and (b) 16mer Mg2+

RNA complex. The G186

and G177 imino peaks are clearly exchange retarded in the presence of Mg2+

. The G178 loop

imino proton peak could not be observed in the absence or presence of Mg2+

.

159

A second 700 MHz 1H-NMR VT series was performed on the 16mer Mg

2+ RNA complex

in order to observe the G178 loop imino proton more clearly and monitor changes in its

intensity and chemical shift. Additional temperature points between 35°C-45°C allowed

more accurate monitoring of the decrease in intensity of G177 and G186 imino proton

peaks. Figure 3.3.2 displays a stack plot of the imino region between 2°C-55°C.

Interestingly, the G178 imino proton peak could be observed between 2°C-15°C but

disappeared completely at 20°C. The U175, U176 and G185 imino proton peaks could be

observed up to the highest temperature point of 55°C. The G177 and G186 imino proton

peaks could still be observed at 50°C, but were rendered absent at 55°C. Again, this

clearly illustrates the enhanced stability produced by Mg2+

.

160

Figure 3.3.2 700 MHz 1H-NMR spectra of the FMDV

16mer Mg2+

RNA complex in 1H2O. A stack plot of the

imino region is shown at variable temperatures; 2°C,

10°C, 15°C, 20°C, 25°C, 35°C, 38°C, 41°C, 45°C, 47°C,

50°C and 55°C. The U175, U176, G185, G186, G177

and G178 imino proton peaks are labelled.

161

3.3.2 31P-NMR variable temperature (VT) series

The 31

P-NMR VT series was analysed for both the 16mer apo-RNA and Mg2+

RNA

complex. For the 16mer apo-RNA, the largest chemical shift changes were observed for

the loop U179 (highfield Δδ=0.66ppm) and A181 (lowfield Δδ=0.59ppm) phosphate

peaks, between 5°C-35°C. This strongly suggests that the U179 and A181 elements of the

phosphate backbone are susceptible to changes in conformation induced by an increase in

temperature. Interestingly, the largest Mg2+

-induced phosphorus chemical shift changes

were also observed for U179 and A181, as previously described in sub-section 3.2.2.

Therefore, the VT series data was used to explain why Mg2+

ions are able to specifically

induce conformation changes in the U179 and A181 phosphate backbones.

When compared to the 16mer Mg2+

RNA complex data, it was observed that the change in

chemical shift, between 5°C-35°C, was significantly reduced for the U179 phosphorus

peak. This would suggest that the addition of Mg2+

confers stability for the U179

phosphate backbone, possibly due to specific RNA-Mg2+

interactions.

162

3.4 Imino proton exchange in the 16mer RNA

3.4.1 NOE exchange

In the NOESY (1H2O) spectra of the 16mer apo-RNA and Mg

2+ RNA complex,

differences were observed in the NOE cross peaks corresponding to connectivity between

the imino proton and water. These NOE cross peaks represent the imino proton

exchanging with water. Cross peaks between imino proton and water were observed in the

16mer apo-RNA for U175, U176, G185, G186 and G177 (Figure 3.4.1). However, in the

Mg2+

RNA complex these cross peaks had significantly reduced in intensity (U175, U176,

G185, G186) or were undetectable (G177). The G186 cross peak was retained the most in

the presence of Mg2+

. This is possibly due to its higher exposure to water as it is adjacent

to the terminal base pair. This evidence strongly suggested that Mg2+

was having the effect

of reducing imino proton exchange.

Figure 3.4.1 400 MHz NOESY (150ms) spectrum of the FMDV 16mer apo-RNA at 5°C in 1H2O

(blue) and Mg2+

RNA complex at 2°C in 1H2O (red). The imino-imino regions (bottom panels) and

imino-water regions (top panels) are displayed. The horizontal line in the top panels represents the

chemical shift of water; 4.995ppm at 5°C and 5.029ppm at 2°C.

163

For the 16mer Mg2+

RNA complex, differences were also observed to the imino-water

NOE cross peaks at two different temperatures, 2°C and 27°C (Figure 3.4.2). At 2°C the

imino-water cross peaks were absent, indicating the lack of imino proton exchange with

water. However, when the temperature was raised to 27°C, imino-water cross peaks were

clearly observed for U175, U176, G185, G186 and G177. This evidence suggests that the

Mg2+

-induced effect observed in Figure 3.4.1 may be reversible with increasing

temperature.


RNA complex in

1H2O at 2°C (red) and 27°C (orange). The imino-imino regions (bottom panels) and imino-water

regions (top panels) are displayed. The horizontal line in the top panels represents the chemical

shift of water; 5.029ppm at 2°C and 4.75ppm at 27°C.

164

3.4.2 T1 of imino protons

The imino proton exchange rates of the 16mer apo-RNA (batch 2) were measured by

using a water magnetisation transfer experiment. However, to calculate the exchange rates

of the imino protons, T1 measurement of the individual imino protons was also necessary.

For the 16mer apo-RNA, T1 experiments were performed at three different temperatures;

2°C, 15°C and 35°C. In these experiments the apparent T1 (T1a) value was measured, so

henceforth the T1 values stated will refer to T1a.

The T1 measurement of the imino protons at these three different temperatures revealed

some interesting results (Figure 3.4.3). At 2°C, the T1 value of the G178 loop imino proton

was significantly smaller compared to the T1 value of the stem imino protons. This is due

to the faster exchange rate of the G178 loop imino proton; the rate of T1 relaxation

increases as the value of Kex increases according to Equation 2.6.2 (Methods: section 2.6).

We also observed that the T1 values of the stem guanine imino protons were very similar

and that they were smaller than the uracil imino proton T1 values. At 15°C, the T1 values

of G178 and U175 decrease owing to the faster exchange rate of these imino protons.

Conversely, an increase in the T1 values of the stem U176, G185, G186, G177 imino

protons is observed, since the T1 of selectively inverted protons is inversely proportional

to the correlation time. However, as the temperature increases to 35°C, the T1 relaxation

rate increases for all the imino protons, again due to faster exchange of the imino protons.

Since the T1 values measured for the imino protons are dependent on the exchange rate,

they can give an indication of how imino proton exchange rates change with temperature.

For example, when comparing the T1 values of imino protons at 2°C and 35°C, it is clear

that the G185 imino proton exchange rate has been affected the least by the increase in

temperature. Nevertheless, the exchange rate constants themselves give a much more

accurate measurement of imino proton exchange.

165

Figure 3.4.3 T1 measurement of the U175, U176, G185, G186, G177 and G178 imino protons for

the FMDV 16mer apo-RNA, at 2°C (blue), 15°C (red) and 35°C (green). Errors bars correspond to

5% of each T1 value.

3.4.3 Exchange rate of imino protons

The imino proton exchange rates of the 16mer apo-RNA, were also measured at 2°C,

15°C and 35°C (Figure 3.4.4). At 2°C, the exchange rates of the stem imino protons were

negligible (between 0-1 s-1

), which suggests that the base pairing is very stable at this

temperature. The exchange rate of the G178 loop imino proton was found to be

approximately 5.0s-1

, which shows that the exchange with water is still relatively slow.

Fascinatingly, at 15°C the exchange rate of G178 increases dramatically to approximately

24.0s-1

. This illustrates that the exchange rate of the G178 imino proton is heavily

dependent on temperature, most likely due to its higher exposure to water. The exchange

rates of the stem imino protons were still negligible at 15°C, except for the U175 imino

proton exchange rate, which increases to approximately 3.0s-1

. This implies that the base

pair involving U175 may be the first to destabilise with increasing temperature.

166

Since the exchange rates of the stem imino protons were negligible at 15°C, the affect of

temperature on exchange rates could not be accurately determined as the values were

within the error bounds. However, at 35°C, the exchange rate of the stem imino protons

increased considerably. The highest exchange rate was found for U175 (23.0s-1

) followed

by G186 (13.0s-1

), U176 (12.0s-1

), G177 (8.0s-1

) and G185 (4.0s-1

). Higher exchange rates

were expected for uracil imino protons as A.U base pairs are less stable than G.C base

pairs. This is supported by the results as the imino proton exchange rates of U175 and

U176 were significantly higher than for G185 and G177. The reason for the higher

exchange rate of the G186 imino proton is most likely due to its proximity to the terminal

U172-A187 base pair, which is shown to be frayed in the 16mer apo-RNA NMR structure.

The G178 imino proton exchange rate could not be calculated accurately at 35°C, even

though the imino proton peak could be observed in the 1H-NMR spectrum, albeit with low

intensity.

These exchange rates were also compared to the 1H-NMR spectra of the 16mer apo-RNA

(batch 2) at 2°C, 15°C and 35°C (Figure 3.4.5). The 1H-NMR data supported the

calculated exchange rate values. The largest increase in exchange rate was observed for

the U175 imino proton, which is clearly reflected in the 1H-NMR spectra; the U175 imino

proton peak has the largest intensity at 2°C, but at 35°C it has the smallest intensity due to

faster exchange with water. Conversely, the G185 imino proton has the lowest exchange

rate at 35°C and in the 1H-NMR spectra the peak has the largest intensity. These results

show that 1H-NMR VT series experiments can indeed be used to estimate the relative

exchange of imino protons.

Interestingly, a comparison of the 1H-NMR VT series of the 16mer apo-RNA (batches 1

and 2) revealed differences in imino proton exchange. As previously discussed, in the

16mer apo-RNA (batch 1), the G186 and G177 imino proton peaks disappear at 35°C.

However, this is not the case with the 16mer RNA (batch 2). The evidence suggests that

the imino proton exchange rates and possibly base pair kinetics are different between these

two 16mer RNA batches even though the tertiary structure is the same. This clearly

highlights the importance of studying both structure and kinetics of RNA.

167

Figure 3.4.4 Exchange rate constants (Kex) of the imino protons for the FMDV 16mer apo-RNA at

2°C (blue), 15°C (red) and 35°C (green). Errors bars correspond to 10% of each Kex value.

Figure 3.4.5 600 MHz 1H-NMR spectra of the FMDV 16mer apo-RNA in

1H2O at three different

temperatures of (a) 5°C, (b) 15°C and (c) 35°C. The U175, U176, G185, G186, G177 and G178

imino proton peaks are labelled accordingly.

168


on imino proton exchange rates

The imino proton exchange rates were measured for the 16mer Mg2+

RNA complex at two

temperatures of 15°C and 35°C (Figure 3.4.6). Subsequently, a comparison was made

between the imino proton exchange rates of the 16mer apo-RNA and Mg2+

RNA complex.

At 15°C, the exchange rate of the G178 imino proton increased from 24.0s-1

to 37.0s-1

,

which is an increase of approximately 55%. A possible explanation for this is that the

Mg2+

ions are positioned close to the imino proton of the G178 base, which is easily

accessible due to the large major groove of the GUAA tetraloop. Since these Mg2+

ions

are fully or partially hydrated, it is likely that it would be the equivalent of exposing the

G178 imino proton to more water.

The changes in exchange rates of the stem imino protons were observed at 35°C. In

particular, the exchange rate of the U175 and U176 imino protons reduced significantly by

approximately 75% and 55% respectively, upon addition of Mg2+

. This suggests that Mg2+

is able to increase the stability of the A.U base pairs involving the U175 and U176 bases.

Consequently, this may have the effect of further stabilising the stem region of the 16mer

RNA and possibly the global tertiary structure. A small, but detectable, decrease in

exchange rate was found for the G177 imino proton, possibly caused by enhanced stability

conferred in the GUAA tetraloop. No significant change in exchange rate was found for

the G185 and G186 imino protons.

These imino proton exchange results give further evidence of Mg2+

-induced stabilisation

and support the 1H-NMR data. Importantly, the exchange rate constants of the imino

protons have been calculated, which provided a more accurate picture of imino proton

exchange. Therefore, together with the results revealed in the VT series experiments, the

measurement of imino proton exchange rates was justified in order to quantify the imino

proton exchange phenomenon and understand the influence of temperature and Mg2+

on

RNA base pair kinetics.

169

Figure 3.4.6 Exchange rate constants (Kex) of the imino protons for the FMDV 16mer Mg2+

RNA

complex at 15°C (red) and 35°C (green). Errors bars correspond to 10% of each Kex value.

170

3.5 Structure determination of the 16mer Mg2+

RNA complex


The chemical shift table for the FMDV 16mer Mg2+

RNA complex is displayed at the end

of this sub-section in Table 3.5.1. Chemical shifts were provided for all 1H,

13C and

31P

nuclei that were identified.


The assignment of imino-imino and imino-amino/H2/H5/H1ʹ connectivities was achieved

by using the same methodology employed for the 16mer apo-RNA, described in sub-

section 3.1.1.

The 700 MHz NOESY (1H2O) spectra of the 16mer apo-RNA and Mg

2+ RNA complex

were analysed. Figure 3.5.1 displays the imino region of the 16mer Mg2+

RNA complex.

The loop G178 imino proton diagonal peak could be clearly observed in the 16mer apo-

RNA, but it was absent in the Mg2+

RNA complex. This was likely due to the faster

exchange rate of the G178 imino proton, which was discovered in the imino proton

exchange experiments. Imino-imino NOE cross peaks between U176-U175, G185-U175

and G186-G185 could be observed in both the 16mer apo-RNA and Mg2+

RNA complex.

The weak G177-U176 cross peak observed in the 16mer apo-RNA could not be found in

the Mg2+

RNA complex.

Figure 3.5.2 displays the imino-amino region of the 16mer Mg2+

RNA complex. No

connectivities from the G178 imino proton were found in this region. The most significant

changes in chemical shift (>0.1ppm) were found for C173 NH2*, G177 NH, A184 NH2

and G185 NH2. No significant chemical shift changes were observed for H5-H6 NOE

cross peaks in the aromatic region. It was concluded that chemical shift changes observed

in the NOESY (1H2O) spectrum, upon addition of Mg

2+, suggests that small structural

changes had occurred in the stem region.

171


RNA complex, at

2°C in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks correspond to

imino-imino connectivities. The sequential assignment starts from U176, labelled in red, and

finishes at G186, labelled in blue. Inset: Secondary structure of the 16mer RNA highlighting the

imino-imino connectivities observed, represented by light blue oval shapes.

172


RNA complex, at

2°C in 1H2O, illustrating the imino-amino region of the spectrum. Connectivities from imino

protons to NH2*/NH2/H2/H5/H1ʹ protons can be observed (NH2* corresponds to the proton

involved in base pair hydrogen bonding); connectivities are marked by a black circle.

173



Sequential assignment was achieved by considering intranucleotide and internucleotide

H6/H8-H1ʹ connectivities, as observed in the 16mer apo-RNA. Firstly, the H5-H6 cross

peaks were assigned using chemical shift values obtained from NOESY and TOCSY

spectra in 1H2O. The U172 H6-H1ʹ intranucleotide connectivity was first identified, to

begin the sequential assignment. A sequential assignment was attained from U172 H6-H1ʹ

to G178 H1ʹ-U179 H6. No connectivity was observed between U179 H6-H1ʹ and U179

H1ʹ-A180 H8. The sequential assignment was commenced from A180 H8-H1ʹ to A187

H8-H1ʹ. Figure 3.5.3 illustrates the sequential assignment discussed here. Figure 3.5.4

illustrates the identification of C6-H6, C8-H8 and C1ʹ-H1ʹ peaks in the 1H-

13C HSQC

spectrum and the assignment of H6/H8-H1ʹ NOE cross peaks in the NOESY (2H2O)

spectrum.

In comparison to the sequential assignment attained for the 16mer apo-RNA it must be

emphasised here that a near full sequential assignment was produced for the 16mer Mg2+

RNA complex. This was achieved as a consequence of the newly observed NOE cross

peaks, which were found in the NOESY (2H2O) spectrum. An example would be the

assignment of the U176 H5-H6 cross peak, which was not observed for the 16mer apo-

RNA. The identification of this NOE cross peak provided the possibility of assigning the

U175 H1ʹ-U176 H6 internucleotide connectivity. Subsequently, this allowed the

sequential assignment to proceed, which highlights the importance of assigning

internucleotide connectivities. The broad peaks observed in the 16mer apo-RNA spectrum

indicate an intermediate exchange in conformation. Therefore, it is likely that the

enhanced stability induced by Mg2+

has reduced conformational exchange, producing

sharper and clearly resolved cross peaks.

174

Figure 3.5.3 Top and bottom panels: 600 MHz NOESY (400ms) spectrum of the FMDV 16mer

Mg2+

RNA complex, at 25°C in 2H2O. The blue line represents U172 H6-H1ʹ to G178 H1ʹ-U179

H6 intra- and internucleotide connectivities, and the green line represents A180 H8-H1ʹ to A187

H8-H1ʹ intra- and internucleotide connectivities; same colouring as in secondary structure shown.

The (i) corresponds to an intranucleotide connectivity and (s) corresponds to a sequential

connectivity. The red circles correspond to H5-H6 connectivities.

175

Figure 3.5.4 Illustration of the identification of C6-H6, C8-H8 and C1ʹ-H1ʹ peaks in the 1H-

13C

HSQC spectrum and the subsequent assignment of H6/H8-H1ʹ cross peaks in the NOESY

spectrum, of the FMDV 16mer Mg2+

RNA complex. (a) and (d) 600 MHz 1H-

13C HSQC spectrum

at 25°C in 2H2O, displaying C6-H6 and C8-H8 peaks. (b), (c) and (e) 600 MHz NOESY (250ms)

spectrum, at 25°C in 2H2O. (f) and (g) 600 MHz

1H-


2H2O,

displaying C1ʹ-H1ʹ peaks.

176


The assignment of H1ʹ-H2ʹ and H2ʹ-H6-H8 NOE cross peaks in the NOESY (2H2O)

spectrum was correlated with the 1H-

13C HSQC spectrum (Figure 3.5.5). A sequential

assignment was achieved between all sixteen nucleotides of the 16mer RNA, which was

used to confirm the previous H6/H8-H1ʹ sequential assignment. This demonstrates the

advantage of using different types of sequential assignment in the NOESY (2H2O)

spectrum to unambiguously assign cross peaks.




FMDV 16mer Mg2+

RNA complex. Bottom panels: 600 MHz 1H-


2H2O. Top panels: 600 MHz NOESY (250ms) spectra at 25°C in

2H2O.

177


A DQF-COSY experiment was used to identify H1ʹ-H2ʹ cross peaks, and in this case,

identify correlations between other sugar protons within three chemical bonds. Figure

3.5.6 displays the H1ʹ-H2ʹ cross peaks observed in the DQF-COSY spectrum, which were

confirmed by the 1H-

13C HSQC spectrum. The A180, U179 and A187 H1ʹ-H2ʹ cross peaks

were clearly observed in the DQF-COSY spectrum, indicating a possible C2ʹ-endo sugar

conformation for these nucleotides. Intranucleotide H3ʹ-H4ʹ and H5ʹ-H5ʹʹ connectivities

were identified using the DQF-COSY and 1H-

13C HSQC spectra. Eight H3ʹ-H4ʹ (Figure

3.5.7) and thirteen H5ʹ-H5ʹʹ (Figure 3.5.8) cross peaks were observed in the DQF-COSY

spectrum. These were used to confirm the sugar-sugar assignments made in the NOESY

(2H2O) spectrum of the 16mer Mg

2+ RNA complex.

178



13C HSQC spectrum, of the FMDV 16mer Mg

2+ RNA

complex. Top right panel: 600 MHz 1H-


2H2O, representing the

C1ʹ-H1ʹ peaks. Top left panel: 600 MHz DQF-COSY spectrum at 25°C in 2H2O, representing the

H1ʹ-H2ʹ cross peaks. Bottom left panel: 600 MHz 1H-


2H2O,

representing the C2ʹ-H2ʹ peaks.

179




2+ RNA

complex. Top panel: 600 MHz DQF-COSY spectrum at 25°C in 2H2O, representing the H3ʹ-H4ʹ

cross peaks. Middle panel: 600 MHz 1H-



C3ʹ-H3ʹ peaks. Bottom panel: 600 MHz 1H-



C4ʹ-H4ʹ peaks.

180

Figure 3.5.8 Illustration of the assignment of intranucleotide H5ʹ-H5ʹʹ connectivities in the DQF-



2+ RNA

complex. Top panel: 600 MHz DQF-COSY spectrum at 25°C in 2H2O, representing the H5ʹ-H5ʹʹ

cross peaks. Bottom panel: 600 MHz 1H-



C5ʹ-H5ʹ and C5ʹ-H5ʹʹ peaks.

181


In Figure 3.5.9, ten peaks were observed corresponding to phosphorus-H6/H8 correlations

and twelve peaks corresponding to phosphorus-H1ʹ correlations. The assignment was

aided by using the 1H-

13C HSQC spectrum. Consequently, twelve phosphorus chemical

shifts were identified from the 1H-

31P CPMG-HSQC-NOESY spectrum.

Figure 3.5.9 Illustration of the assignment of phosphorus to H6/H8/H1ʹ peaks in the 1H-

31P

CPMG-HSQC-NOESY spectrum, with the aid of the 1H-


16mer Mg2+

RNA complex. Top panels: 600 MHz 1H-


2H2O. Bottom

panels: 600 MHz 1H-

31P CPMG-HSQC-NOESY (500ms) spectra at 25°C in

2H2O.

182

Identification of proton resonances using 3D NMR

For the 16mer Mg2+

RNA complex, the 3D NOESY/2Q-COSY spectrum was used to

identify the base and sugar protons of the U176 nucleotide. This was because in the 16mer

apo-RNA it was difficult to identify U176 proton chemical shifts. Figure 3.5.10a displays

the F3/F1 plane chosen at a specific F2 frequency of 9.05ppm, which allows the

observation of scalar coupling between the U176 H5 and H6 protons. Strong inner NOE

cross peaks were observed between the U176 H5 and H6 protons. Weaker outer NOEs

were observed to other sugar/base protons, including U175 H6, U176 H3ʹ, U175 H4ʹ,

U175 H2ʹ and U176 H5ʹʹ.

Internucleotide connectivities were observed in the 3D NMR spectrum, which provided a

potential means of sequential assignment. The shortest internucleotide distance is between

H2ʹ(i)- H6/H8(i+1). Therefore, these internucleotide connectivities were observed more

easily due to stronger NOE intensities. Figure 3.5.10b displays different F1 slices in the

F3/F2 plane of the 3D NMR spectrum. The F1 frequencies correspond to either the H2ʹ or

H6/H8 chemical shifts. In each F1 slice, the internucleotide connectivity between H2ʹ(i)-

H6/H8(i+1) was observed; the black circles represent the H2ʹ(i)-H6/H8(i+1) peaks in each

F1 slice. For example, the first slice was chosen at an F1 frequency corresponding to C173

H6 (8.145ppm); the U172 H2ʹ-C173 H6 peak was clearly observed at 4.594ppm. In

addition, an intranucleotide connectivity between C173 H2ʹ-C173 H6 was observed.

Therefore, a sequential assignment was achieved from U172 H2ʹ-C174 H6, shown by the

arrow in Figure 3.5.10b.

183

Figure 3.5.10 (a) 600 MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer Mg2+

RNA complex. Identification of base and sugar protons of the U176 nucleotide. This is illustrated

by the F3/F1 NOE plane at F2 = 9.05ppm. Both positive (orange) and negative (green) levels are

shown. The strong intensity of U176 H5 and U176 H6 inner NOE cross peaks are associated with

weaker outer NOE peaks to U175 H6, U176 H3ʹ, U175 H4ʹ, U175 H2ʹ and U176 H5ʹʹ. (b) 600

MHz 3D NOESY/2Q-COSY (250ms) spectrum of the FMDV 16mer Mg2+

RNA complex. Slices

from the F3/F2 plane chosen at different F1 frequencies, identifying H2ʹ(i)-H6/H8(i+1)

internucleotide connectivities. A sequential assignment is observed between U172 H2ʹ-C174 H6.

The black circles represent the H2ʹ(i)-H6/H8(i+1) peaks in each F1 slice.

184

Table 3.5.1 1H,

13C and

31P NMR chemical shifts of the FMDV 16mer Mg

2+ RNA complex, in

1H2O and

2H2O.


U172 5.898 8.114 5.543 4.594 4.560 4.357 3.922 4.053 104.2 140.2 93.91 75.30 73.93 85.05 61.89 C173 8.510 7.167 5.888 8.150 5.715 4.449 4.631 4.519 4.206 4.576 97.7 142.6 94.28 75.52 72.43 82.15 64.91 -4.471

C174 8.415 6.954 5.604 7.966 5.518 4.400 4.575 4.476 4.621 4.145 97.5 141.9 94.17 72.58 72.03 81.79 64.44 -4.794

U175 14.04 5.453 7.981 5.532 4.439 4.582 4.478 4.139 103.2 142.3 94.19 75.54 72.01 81.79 -4.812

U176 13.44 5.662 8.060 5.64 4.421 4.662 4.473 4.602 4.151 103.8 141.9 93.39 72.71 72.13 81.80 64.47 -4.753

G177 12.41 8.153 5.830 7.655 5.839 4.454 4.683 4.583 4.543 4.158 136.1 92.38 76.74 72.03 81.99 64.60 -4.423

G178 7.289 5.632 4.377 4.579 4.328 4.444 4.080 136.5 93.39 75.74 72.03 82.11 64.38

U179 11.27 5.380 7.724 5.38 4.379 4.148 4.116 4.188 3.835 103.5 143.1 92.76 75.74 72.84 82.99 64.34 -2.105

A180 7.765 8.085 5.69 4.390 4.605 4.293 4.080 3.939 154.8 140.2 92.62 75.50 74.15 82.94 65.17 -3.134

A181 8.168 8.122 6.026 4.456 4.948 4.445 4.280 4.574 155.6 143.9 91.85 75.54 73.15 81.95 65.48 -5.222

C182 8.585 7.059 6.062 7.848 3.676 4.330 4.453 4.172 4.270 4.200 99.6 141.8 95.01 74.41 74.29 82.58 68.72 -2.942

A183 7.724 6.548 6.705 8.1 5.859 4.481 4.806 4.464 4.498 4.101 152.1 139.9 92.74 75.71 72.28 81.80 64.36

A184 7.940 6.478 7.376 7.884 5.899 4.486 4.691 4.409 4.604 4.144 153.4 139.0 92.22 75.67 72.11 81.73 64.44 -4.747

G185 12.93 8.369 6.251 7.069 5.562 4.454 4.038 4.438 135.5 92.29 76.74 65.35 -4.381

G186 12.61 8.091 7.06 5.659 4.467 4.423 4.272 4.023 4.431 135.8 93.06 75.55 75.39 83.80 65.35

A187 8.054 7.773 6.062 4.112 4.274 4.061 4.493 155.1 140.0 91.89 77.96 75.42 65.39 -4.417

185


Distance restraints were generated from the NOESY spectra of the 16mer Mg2+

RNA

complex. Exchangeable NOE restraints were obtained from the NOESY spectrum in 1H2O,

with 150ms mixing time. Non-exchangeable NOE restraints were obtained from the

NOESY spectrum in 2H2O, with 250ms mixing time. The non-exchangeable NOE distance

restraints consisted of base and sugar ribose connectivities.

The absence of strong intranucleotide H6/H8-H1ʹ cross peaks for stem nucleotides,

allowed the glycosidic angle of all stem nucleotides to be restrained to the anti

conformation. The glycosidic angles were not defined for loop nucleotides to allow some

conformational flexibility in the loop region. All stem nucleotides were restrained to the

C3ʹ-endo conformation, except for A187. The δ angle for the loop nucleotides was not

restrained, even though 3JH3-ʹH4ʹ couplings larger than 2Hz for the A180 and A181 loop

nucleotides were clearly observed in the DQF-COSY spectrum.

Hydrogen bond restraints were added for all six base pairs. However, since the U172

imino proton peak was not observed and no connectivities were found in the NOESY

(1H2O) spectrum, the hydrogen bond restraints between U172-A187 were loosely

restrained. For the same reason, planarity restraints were added for all base pairs except

for U172-A187.



step, one violation was found in the distance restraints. The restraints that were used for

these structure calculations are summarised in Table 3.5.2.

186

Restraints

NOEs 260

Strong 1.8 – 2.5 Å 13

Medium 2.6 – 3.3 Å 73

Weak 3.4 – 5.0 Å 173

Very weak 5.1 – 7.0 Å 1

Internucleotide NOEs 125

Intranucleotide NOEs 135

Hydrogen bonds 36

Planarity 10

Dihedral angles 98

Helix (, , , ε, ζ) 50


Glycosidic (χ) 12

NOE per residue 16.3

Restraints per

residue

25.3



FMDV 16mer Mg2+

RNA complex.

187





calculated to be 0.17Å.

Figure 3.5.11 Illustration of the NMR solution structures of the FMDV 16mer Mg2+

RNA

complex (a) Overlay of the 20 lowest RMSD structures with average RMSD of 0.17Å. (b) Lowest

RMSD solution structure (0.16Å). The red ribbon represents the RNA backbone.


homogeneity of the 16mer Mg2+

RNA complex tertiary structure. The helical stem region

and tetraloop region are aligned very well, despite the dynamic nature of the tetraloop

region. When considering the helical stem and the tetraloop separately, there was no

188

significant difference in RMSD. The average RMSD of the helical stem alone was 0.164Å

and of the tetraloop alone was 0.186Å. This is likely due to the high number of intra- and

internucleotide connectivities in the loop region. From the ensemble of 20 structures, the

structure with the lowest RMSD (0.16Å) was selected for structure analysis and

conformational analysis (Figure 3.5.11b).

3.5.3.2 GNRA tetraloop

Analogous to the 16mer apo-RNA NMR structure, a sharp turn was found between the

G178 and U179 nucleotides (Figure 3.5.12). Fascinatingly, the U179, A180 and A181

bases stack directly on top of each other, unlike in the 16mer apo-RNA NMR structure.

Therefore, it is likely that upon addition of Mg2+

, the base stacking interactions between

U179, A180 and A181 bases are strengthened, increasing the stability of the GNRA

tetraloop.

Figure 3.5.12 The G178UAA181 tetraloop of the FMDV 16mer Mg2+

RNA complex NMR structure

shown in Figure 3.5.11b; the closing G177.C182 base pair is also illustrated. Colour coding of

nucleotides: guanosine (blue), uridine (cyan), adenosine (green) and cytosine (red).

189


In total, seven intramolecular interactions were found in the GUAA tetraloop of the 16mer

Mg2+

RNA complex NMR structure. Two specific, base-base interactions were found

between G178 and A181, which correspond to the two hydrogen bonds involved in G.A

sheared base pairing; G178 NH2–A181 N7 and G178 N3–A181 NH6 were found to be

2.23Å and 3.45Å, respectively (Figure 3.5.13). The G178 and A181 bases were in the

correct orientation for G.A sheared base pairing and the hydrogen bond distances were

shorter compared with the 16mer apo-RNA NMR structure. Therefore, these results

provide good evidence that Mg2+

may be required to form a G.A sheared base pair in

GNRA tetraloops.

Figure 3.5.13 The G.A sheared base pair of the FMDV 16mer Mg2+

RNA NMR structure.

Hydrogen bonding distances between G178 NH2-A181 N7 (2.23Å) and G178 N3-A181 NH6

(3.45Å) are indicated by the red dotted lines.

One specific, base-phosphate interaction was found in the GUAA tetraloop, which is

found to be conserved in the GNRA tetraloop (Table 3.5.3); G178 NH2-A181 OP at

1.88Å. Interestingly, this distance is longer in the 16mer apo-RNA NMR structure at

2.11Å. This indicates that the base-phosphate interaction is stronger in the 16mer Mg2+

190

RNA complex, which would enhance stability of the GUAA tetraloop. In addition, four

intranucleotide, base-phosphate interactions were found for G178, U179, A180 and A181.


interaction


G178 NH2 – A181 N7 base-base specific 2.23

G178 N3 – A181 NH6 base-base specific 3.45

G178 NH2 – A181 OP base-phosphate specific 1.88

G178 H8 – G178 O5ʹ base-phosphate non-specific 3.10

U179 H6 – U179 O5ʹ base-phosphate non-specific 2.73



Table 3.5.3 Seven intramolecular interactions in total were identified in the GUAA tetraloop of

the FMDV 16mer Mg2+

RNA complex NMR structure. The interactions formed between donor

and acceptor atoms are given, the type of interaction, the specificity of the interaction and the

distances between the proton donor and acceptor atoms.

191


The 3DNA program was used to perform a conformational analysis on the final 16mer

Mg2+

RNA complex NMR structure. The local helical parameters, base pair step

parameters and complementary base pair parameters are displayed in Tables 3.5.4, 3.5.5

and 3.5.6, respectively.

For the 16mer Mg2+

RNA complex structure, both the ‘x-displacement’ (-3.84Å) and

‘slide’ (-1.46Å) parameter values suggest that the 16mer Mg2+

RNA complex structure is

characteristic of an A-form helical conformation. The ‘shear’ parameter value of the G.A

sheared base pair was found to be 6.72Å (Table 3.5.6), which confirms the sheared

conformation of the G.A base pair. In contrast, a value of 7.67Å was found for the 16mer

apo-RNA, in which the G178 and A181 bases were not in the right orientation to form the

sheared base pair.

Base

Pair Nucleotides

Xdisp

(dx)

Ydisp

(dy)

Inclination

(η)

Tip

(θ)

Helical Twist

(Ω)

Helical

Rise (h)

1-2 U-A / C-G -1.66 -0.99 0.43 11.32 36.11 3.85

2-3 C-G / C-G -4.44 0.95 16.84 -4.92 34.15 2.48

3-4 C-G / U-A -5.12 -0.07 12.30 1.33 29.45 2.57

4-5 U-A / U-A -4.86 -0.67 21.50 -5.10 32.87 2.41

5-6 U-A / G-C -6.00 -0.35 32.03 4.54 29.40 1.73

6-7 G-C / G-A -0.94 1.30 7.72 8.44 57.88 3.35

Ave. - -3.84 0.03 15.14 2.60 36.65 2.73

Table 3.5.4 Local helical parameter values for the FMDV 16mer Mg2+

RNA complex structure,


192

Base

Pair Nucleotides

Shift

(Dx)

Slide

(Dy)

Rise

(Dz)

Tilt

(τ)

Roll

(ρ)

Twist

(ω)

1-2 U-A / C-G -0.12 -1.00 3.91 -6.98 0.27 35.45

2-3 C-G / C-G -0.33 -1.81 3.14 2.85 9.75 32.65

3-4 C-G / U-A -0.03 -2.02 3.05 -0.67 6.21 28.80

4-5 U-A / U-A 0.62 -1.73 3.18 2.82 11.89 30.57

5-6 U-A / G-C -0.01 -1.71 3.06 -2.19 15.45 24.99

6-7 G-C / G-A -1.69 -0.52 3.24 -8.13 7.43 56.91

Ave. - -0.26 -1.46 3.26 -2.05 8.50 34.90

Table 3.5.5 Base pair step parameter values for the FMDV 16mer Mg2+

RNA complex structure,


Base

Pair Nucleotides

Shear

(Sx)

Stretch

(Sy)

Stagger

(Sz)

Buckle

(κ)

Propeller

(π)

Opening

(σ)

1 U-A -0.69 -0.28 -1.93 1.82 6.77 -4.46

2 C-G -0.17 -0.22 0.10 5.67 -21.18 -0.80

3 C-G 0.14 -0.24 -0.06 -0.00 -18.62 -6.90

4 U-A -0.14 -0.22 0.40 0.02 -16.57 -9.67

5 U-A -0.05 0.04 0.21 -2.50 -12.48 0.50

6 G-C -0.11 -0.17 -0.25 -0.03 5.96 0.44

7 G-A 6.72 -5.83 1.53 17.01 5.37 -18.58

Table 3.5.6 Complementary base pair parameter values for the FMDV 16mer Mg2+

RNA complex

structure, calculated by the 3DNA analysis program.


CURVES conformational analysis programs (Table 3.5.7). The 3DNA program was used

to generate dihedral angles values for nucleotides of the stem base pairs and the G.A

sheared base pair in the 16mer Mg2+

RNA complex structure. Fourteen dihedral angles

values were acquired by the 3DNA program. The CURVES program was used to obtain

the dihedral angle values for U179 and A180 nucleotides.

All four nucleotides in the loop revealed δ angle values that conform to the C3ʹ-endo

conformation. In contrast, values conforming to the C2ʹ-endo conformation for the A180

nucleotide were found in the 16mer apo-RNA structure. The only significant deviation in

193

the dihedral angles were found in the α angle of the U179 and γ angle of U172. Analogous

to the 16mer apo-RNA structure, the deviation in the α angle U179 is caused by the

rotation around the P-O5ʹ bond that forms the U-turn motif found in GNRA tetraloops.

Base Nucleotides C1ʹ-N

(χ)

C5ʹ-C4ʹ

(γ)

C4ʹ-C3ʹ

(δ)

C3ʹ-O3ʹ

(ε)

O3ʹ-P

(ζ)

P-O5ʹ

(α)

O5ʹ-C5ʹ

(β)

1 U -163.9 -175.8 84.5 -152.7 -88.9 - -

2 C -156.4 55.9 84.8 -153.6 -65.1 -63.6 167.8

3 C -156.9 52.7 81.9 -158.7 -67.6 -68.5 174.1

4 U -158.4 54.2 81.1 -154.8 -63.4 -64.1 174.0

5 U -153.7 51.9 80.9 -153.5 -65.7 -69.9 178.2

6 G -157.7 53.2 81.8 -152.3 -61.0 -69.1 172.2

7 G -153.7 53.5 81.3 -156.8 -61.3 -68.4 171.8

8 U -143.9 55.8 85.6 -153.9 -55.7 158.9 153.6

9 A -138.9 57.5 86.3 -154.3 -63.9 -68.5 165.9

10 A -145.9 55.3 84.2 -151.1 -55.9 -65.0 158.7

11 C -151.6 66.2 89.1 -157.7 -65.8 -66.4 158.3

12 A -155.0 54.1 80.6 -155.3 -69.0 -64.3 172.8

13 A -161.3 53.5 81.0 157.4 -67.9 -66.2 172.1

14 G -161.3 55.0 80.7 -154.5 -63.9 -63.9 170.7

15 G -162.6 54.0 81.3 -158.9 -69.6 -66.8 173.2

16 A -121.4 64.3 149.1 - - -64.3 -178.9

Table 3.5.7 Dihedral angle values of nucleotides in the FMDV 16mer Mg2+

RNA complex

structure, calculated by the 3DNA (black) and CURVES (red) analysis programs. Angles are all

measured in degrees.

Dihedral angles (ν1 and ν2) that define the sugar ribose conformation were also calculated

by the 3DNA and CURVES programs, including the pseudorotation phase angle and the

amplitude (Table 3.5.8). Eleven out of the sixteen nucleotides were calculated to have a

C3ʹ-endo sugar ribose conformation. The C4ʹ-exo conformation was revealed for three

nucleotides and one C2ʹ-exo conformation was found for C182. In contrast, only six C3ʹ-

endo conformations were found in stem and loop nucleotides of the 16mer apo-RNA. All

four loop nucleotides adopted sugar pucker conformations within or close to the C3ʹ-endo

conformation.

194

Base Nucleotides v1 v2 Amp Phase Conformation

1 U -10.4 27.9 36.0 39.9 C4ʹ-exo

2 C -31.4 40.3 40.5 6.0 C3ʹ-endo

3 C -22.8 37.3 40.0 21.3 C3ʹ-endo

4 U -23.8 39.5 42.5 21.6 C3ʹ-endo

5 U -22.1 38.6 42.4 24.5 C3ʹ-endo

6 G -8.8 28.8 39.5 43.2 C4ʹ-exo

7 G -17.1 34.2 39.7 30.6 C3ʹ-endo

8 U -11.6 28.0 35.6 36.2 C4ʹ-exo

9 A -28.4 37.2 38.3 7.24 C3ʹ-endo

10 A -23.0 36.5 39.0 20.5 C3ʹ-endo

11 C -34.0 40.4 40.4 358.9 C2ʹ-exo

12 A -27.7 42.1 43.9 16.6 C3ʹ-endo

13 A -25.1 40.1 42.7 20.1 C3ʹ-endo

14 G -25.5 41.4 44.2 20.7 C3ʹ-endo

15 G -21.8 37.1 40.3 23.2 C3ʹ-endo

16 A 39.4 -39.0 40.6 163.7 C2ʹ-endo



Mg2+

RNA complex structure. Values were calculated by the 3DNA (black) and CURVES (red)

analysis programs.

195

3.5.5 Comparison of the 16mer apo/Mg2+

RNA NMR structures

The successful determination of the 16mer apo and Mg2+

complex RNA structures

allowed a comparison to be made between the two RNAs. In this sub-section the

similarities and differences will be highlighted.

The assignment of the 16mer apo and Mg2+

complex RNAs proceeded using the NMR

assignment strategy. The assignments produced from the NOESY spectra in both 1H2O

and 2H2O were extremely important as they provided the distance restraints required for

structure determination. The number of NOE restraints per residue generated for the

16mer apo and Mg2+

complex RNA structures was 7.8 and 16.3, respectively. Relatively,

52% fewer NOEs per residue were used in the 16mer apo-RNA structure. The reason for

this is because the NOESY (2H2O) spectrum of the 16mer apo-RNA, possessed broad

lines and reduced sensitivity when compared to the 16mer Mg2+

RNA complex. This

hindered efforts to produce a H6/H8-H1ʹ sequence-specific sequential assignment, which

made further assignment difficult. Regardless of the limitations faced with the 16mer apo-

RNA, the 16mer apo-RNA structures generated were of very good quality with low

RMSDs. This may be due to the larger number of internucleotide connectivities obtained

compared to intranucleotide connectivities; 71 and 54, respectively.

In contrast, the H6/H8-H1ʹ assignment of the 16mer Mg2+

RNA complex was more

straightforward. A greater number of intra- and internucleotide NOE cross peaks could be

observed and assigned, most likely due to the stabilisation effect produced by the presence

of Mg2+

. Consequently, the final twenty 16mer Mg2+

RNA complex structures generated

had low RMSDs, with no significant difference in RMSD found between the stem and

loop region. The final solution structures of both the 16mer apo and Mg2+

complex RNAs

were of a high standard. Structure validation using the Molprobity program, revealed no

bad bonds, angles, sugar pucker and backbone conformations for the NMR structures of

both the 16mer apo-RNA and Mg2+

RNA complex. Clash scores were found to be low;

31.2 and 21.4 for the 16mer apo-RNA and Mg2+

RNA complex, respectively.

196

3.6 Mg2+

-induced structural changes to the 16mer apo-RNA

The only major structural difference in the stem region of the 16mer apo-RNA structure

was found for the U172 nucleotide, upon addition of Mg2+

. The DQF-COSY experiment

provided clear evidence of the change from C2ʹ-endo to C3ʹ-endo sugar conformation for

the U172 nucleotide, which was also observed in the NMR structures. Consequently, the

U172 base changes its orientation to stack with C173, which was not the case in the 16mer

apo-RNA structure. Therefore, this would have the effect of increasing stability of the

terminal stem region. This is another example of the stabilising effect of Mg2+

on RNA.

Significant chemical shift changes to exchangeable and non-exchangeable protons were

also observed in the stem region of the 16mer RNA, upon addition of Mg2+

(Figure 3.6.1).

It was observed that the conformation of the GUAA tetraloop was significantly different

between the 16mer apo and Mg2+

RNA structures. Mg2+

is known to interact with the

negatively charged phosphate backbone, so was likely that Mg2+

has caused the

conformational change in the phosphate backbone of the GUAA tetraloop. The

consequence of these structural changes in the loop was that the U179, A180 and A181

bases were tightly stacked together on the 3ʹ-end, creating a more compact GUAA

tetraloop (Figure 3.6.2) allowing for stronger base stacking interactions. These structural

changes allowed the formation of stronger intramolecular interactions due to shorter

hydrogen bond distances. Crucially, the two hydrogen bonds involved in forming a G.A

sheared base pair were shorter (Figure 3.6.3). This suggests that Mg2+

is required to form a

stable G.A sheared base pair, which was not possible in the 16mer apo-RNA structure.

Base-phosphate interactions between A180 H8-A180 O5ʹ and A181 H8-A181 O5ʹ were

also stronger in the Mg2+

RNA complex structure, adding to the stability of the loop

(Figures 3.6.3 and 3.6.4). Both the intramolecular and base stacking interactions are

improved following the Mg2+

-induced structural changes to the GUAA tetraloop, which

significantly contribute towards its stability. These findings demonstrate the ability of

Mg2+

to enhance stability in the GNRA tetraloop and the mechanism by which Mg2+

is

able to do that. Table 3.6.1 summaries the changes highlighted here.

197

Figure 3.6.1 Histogram illustrating the changes to the chemical shift of exchangeable and non-

exchangeable protons of the FMDV 16mer RNA, upon addition of 5eq of Mg2+

. Bars shown in

blue represent exchangeable protons. Bars shown in green and orange represent non-exchangeable

protons in the stem and loop, respectively. Negative values correspond to a highfield chemical

shift change and positive values correspond to a lowfield chemical shift change.


-induced structural changes to the GUAA tetraloop in the

FMDV 16mer RNA by comparison of the 16mer (a) apo-RNA structure and (b) Mg2+

RNA

complex structure. The U179, A180 and A181 bases are stacked more tightly together in the

16mer Mg2+

RNA complex, strengthening the base stacking interactions.

198


-induced structural changes to the A181 nucleotide in the


RNA

complex structure. The G.A sheared base pair is formed by two hydrogen bonds (black dashed

lines); G178 NH2-A181 N7 is 3.09Å and 2.23Å in apo-RNA and Mg2+

complex, respectively and

G178 N3-A181 NH6 is 5.05Å and 3.45Å in apo-RNA and Mg2+

complex, respectively. Stronger

base-phosphate intramolecular interactions were formed (blue dashed line); A181 H8-A181 O5ʹ is

3.29Å and 2.65Å in the 16mer apo-RNA and Mg2+

RNA complex, respectively.

199


-induced structural changes to the A180 nucleotide in the


RNA

complex structure. A stronger base-phosphate intramolecular interaction is formed (red dashed

line); A180 H8-A180 O5ʹ is 3.08Å and 2.30Å in the 16mer apo-RNA and Mg2+

RNA complex,

respectively.

Parameters 16mer

apo-RNA

16mer Mg2+

RNA

complex

Difference

Δ

A180 Delta (º) 140.1 84.2 55.9

G178 NH2 – A181 N7 (Å) 3.09 2.23 0.86

G178 N3 – A181 NH6 (Å) 5.05 3.45 1.60

G178 NH – A181 OP (Å) 3.35 3.31 0.04

G178 NH2 – A181 OP (Å) 2.11 1.88 0.23

G178 H8 – G178 O5ʹ (Å) 3.05 3.10 0.05

U179 H6 – U179 O5ʹ (Å) 2.84 2.73 0.11

A180 H8 – A180 O5ʹ (Å) 3.08 2.30 0.78

A181 H8 – A181 O5ʹ (Å) 3.29 2.65 0.64

Table 3.6.1 A comparison of dihedral angle and distance values obtained from the FMDV 16mer

apo-RNA and Mg2+

RNA complex structures, highlighting the changes induced by Mg2+

.

200

Experimental evidence has also been found to support the changes to the loop

conformation that are described here. The DQF-COSY experiment of the 16mer Mg2+

RNA complex revealed 3JH3ʹ-H4ʹ couplings greater than 2 Hz for A180 and A181

nucleotides, characteristic of the C3ʹ-endo conformation. This supports the transition from

C2ʹ-endo to C3ʹ-endo sugar conformation of the A180 nucleotide, which is observed in the

NMR structures. A comparison of the F3/F1 planes of the 3D NOESY/2Q-COSY spectra

also supports the change in sugar puckering of the A180 nucleotide (Figure 3.6.5).

Significant chemical shift changes to the non-exchangeable protons of loop nucleotides

can also be observed in the 16mer RNA, upon addition of Mg2+

(Figure 3.6.1). This could

be as a result of changes in electronic environment caused by changes in base stacking

observed in the NMR structures.

Figure 3.6.5 600 MHz 3D NOESY/2Q-COSY spectrum (250ms) of the FMDV 16mer (a) apo-

RNA and (b) Mg2+

RNA complex. The F2 chemical shift is labelled in each plane. Both positive

(orange) and negative (green) levels are shown. In the 16mer apo-RNA, scalar coupling between

A180 H1ʹ and A180 H2ʹ can be clearly observed by strong cross peaks, characteristic of the C2ʹ-

endo sugar conformation. In the 16mer Mg2+

RNA complex, coupling is still observed between

A180 H1ʹ-H2ʹ, although with significantly reduced intensity.

201

Chapter 4: NMR studies of the FMDV 15mer RNA and its complex

with the 16mer RNA

In this chapter the NMR assignment and structure determination of the 15mer apo-RNA

will be described, which will include a conformational analysis of the NMR solution

structure. Secondly, the effect of Mg2+

on the 15mer RNA will be briefly discussed.

Finally, the results and analysis of the 16mer/15mer RNA-RNA interaction will be

presented.

4.1 Structure determination of the 15mer apo-RNA


The chemical shift table for the FMDV 15mer apo-RNA is displayed at the end of this

sub-section in Table 4.1.1. Chemical shifts were provided for all 1H,

13C and

31P nuclei

that were identified.


Imino proton identification and assignment

The 15mer RNA contains four guanine and uracil bases so a maximum of four imino

proton peaks could be observed in the imino region of the 1H-NMR spectrum. Four imino

proton peaks were observed in the 1H-NMR spectrum at 2°C (Figure 4.1.1). The

1H-NMR

spectrum clearly illustrates three distinct peaks in the imino region corresponding to G229

(13.17ppm), U230 (13.90ppm) and G231 (12.72ppm). However, a fourth peak was

observed with much reduced intensity, close to the chemical shift of the G231 imino

proton peak. This peak was identified to be G240 (12.78ppm). It is likely that the G240

imino proton is in fast exchange with water due to its proximity to the loop, causing a

significant reduction in signal intensity and line broadening.

202


1H2O,

displaying the imino region. Four peaks were identified corresponding to the imino protons of

G229, U230, G231 and G240.

As the 1H-NMR spectrum does not provide unambiguous assignment of the guanine and

uracil bases, the NOESY (1H2O) spectrum was analysed to provide a more accurate

assignment. Two strong imino proton peaks were found in the NOESY (1H2O) spectrum,

corresponding to the U230 and G231 peaks found in the 1H-NMR spectrum. The G229

imino proton peak could also be observed in the NOESY (1H2O) spectrum, but it was very

weak. The G240 imino proton peak could not be clearly identified in the NOESY (1H2O)

spectrum due to line broadening and overlapping with the G231 imino proton peak. One,

imino-imino connectivity was observed in the NOESY (1H2O) spectrum between U230-

G231 as shown in Figure 4.1.2. No imino-imino connectivities were observed between

G229-U230 and G231-G240.

203


illustrating the imino region of the spectrum. The cross-diagonal peaks correspond to imino-imino

connectivities. The sequential assignment shown here is between U230-G231. The G229 NH

diagonal peak is shown with an arrow. Inset: Secondary structure of the 15mer RNA highlighting

the observed imino-imino connectivities, represented by light blue oval shapes.

Imino-amino assignment

The imino-amino region of the NOESY (1H2O) spectrum was assigned using the identified

chemical shift values of U230, G229 and G231 imino protons. The full assignment of

connectivities in the imino-amino region is shown in Figure 4.1.3. These results provided

strong evidence of base pairing between U230-A242 and G231-C241 base pairs.

204


illustrating the imino-amino region of the spectrum. Connectivities from imino protons to

NH2*/NH2/H2/H5 protons can be observed (NH2* corresponds to the proton involved in base

pair hydrogen bonding); connectivities are marked with a black circle.

205


H5-H6 assignment

In the FMDV 15mer RNA, there is one uracil and three cytosine nucleotides in the stem

and four cytosine nucleotides in the loop. Accordingly, eight H5-H6 cross peaks are

expected to be observed in the NOESY (2H2O) spectrum. The U230, C232, C235, C236,

C237, C238, C241 and C243 H5-H6 cross peaks were assigned successfully. Figure 4.1.4

illustrates the assignment of H5-H6 cross peaks in the NOESY (2H2O) spectrum, with the

aid of the 1H-

13C HSQC spectrum. Interestingly, the H5 chemical shifts of the loop C236,

C237 and C238 nucleotides were found to be lowfield to those of the stem cytosine

nucleotides, except for the C235 nucleotide.


A sequential assignment was carried out from G229 H8-H1ʹ to C243 H6-H1ʹ. The

intranucleotide G229 H8-H1ʹ connectivity was found to start the sequential assignment. A

sequential assignment was attained from G229 H8-H1ʹ to A234 H1ʹ-C235 H6. In this part

of the sequential assignment, connectivities to A234 H2 were used instead of A234 H8.

No intranucleotide connectivity was observed between C235 H6-H1ʹ and C237 H1ʹ-C238

H8. Subsequently, the sequential assignment commenced from C238 H6-H1ʹ to C243 H6-

H1ʹ; connectivities to A239 H2 were used instead of A239 H8. The reason for using

connectivities to A234 H2 and A239 H2 is because the cross peaks corresponding to these

connectivities could be clearly observed. This is because the A234 and A239 nucleotides

are found in the 15mer RNA loop and so distances from intra- and internucleotide H2-H1ʹ

were shorter than H8-H1ʹ. The use of H2-H1ʹ connectivities highlighted the importance of

using alternative methods for sequential assignment depending on the RNA tertiary

structure. Figure 4.1.5 illustrates the sequential assignment discussed.

206


13C HSQC

spectrum and the subsequent assignment of H5-H6 cross peaks in the NOESY spectrum, of the

FMDV 15mer apo-RNA. Bottom left panel: 700 MHz NOESY (250ms) spectrum, at 25°C in 2H2O;

blue circles indicate H5-H6 cross peaks. Top left panel: 400 MHz 1H-

13C HSQC spectrum, at 25°C

in 2H2O, displaying the C6-H6 peaks. Bottom right panel: 400 MHz

1H-

13C HSQC spectrum, at

25°C in 2H2O, displaying the C5-H5 peaks.

207

Figure 4.1.5 700 MHz NOESY (250ms) spectrum of the FMDV 15mer apo-RNA, at 25°C in

2H2O. The blue line represents G229 H8-H1ʹ to A234 H1ʹ-C235 H6 intra- and internucleotide

connectivities. The green line represents C238 H6-H1ʹ to C243 H6-H1ʹ intra- and internucleotide

connectivities (same colouring in secondary structure shown). H6/H8 and H2 chemical shifts are

labelled in black and blue, respectively. The (i) corresponds to an intranucleotide connectivity and

(s) corresponds to a sequential connectivity. The red circles correspond to H5-H6 connectivities.

208


Six sequential connectivities of H1ʹ-H2ʹ and H2ʹ-H6/H8 NOE cross peaks were assigned

in the NOESY (2H2O) spectrum, with the aid of the

1H-

13C HSQC spectrum (Figure 4.1.6).

Four of the sequential connectivities were found between stem nucleotides. The sequential

assignments produced here were used to confirm the H6/H8-H1ʹ sequential assignments.




FMDV 15mer apo-RNA. Bottom panels: 400 MHz 1H-


2H2O. Top

panels: 700 MHz NOESY (250ms) spectra at 25°C in 2H2O.

209


Intranucleotide and internucleotide cross peaks from H1ʹ to H2ʹ, H3ʹ, H4ʹ, H5ʹ and H5ʹʹ

protons were assigned. Analogously, intranucleotide and internucleotide cross peaks from

H2ʹ, H3ʹ, H4ʹ, H5ʹ and H5ʹʹ protons to H2/H6/H8 protons were also assigned.

The DQF-COSY spectrum was analysed to find H1ʹ-H2ʹ cross peaks. The C243 H1ʹ-H2ʹ

cross peak was clearly observed in the DQF-COSY spectrum; no other H1ʹ-H2ʹ cross

peaks were found. The C243 nucleotide is the last nucleotide located at the 3ʹ-end of the

FMDV 15mer RNA, which means that it is more susceptible to changes in sugar pucker

conformation.


The chemical shifts of H6/H8 and H1ʹ protons were identified relative to the 31

P chemical

shifts in the 1H-

31P CPMG-HSQC-NOESY spectrum, which assisted in confirming the

H6/H8-H1ʹ sequential assignment. Figure 4.1.7 illustrates the identification of H6/H8 and

H1ʹ chemical shifts in the NOESY (2H2O) spectrum, using both the

1H-

13C HSQC

spectrum and the 1H-


Using the 1H-

31P CPMG-HSQC-NOESY spectrum, phosphorus correlations to H6/H8 and

H1ʹ protons in both the 5ʹ and 3ʹ directions can be observed, resulting in a phosphorus

driven pathway for sequential assignment of the 15mer apo-RNA. Therefore, a sequential

assignment was attempted from phosphorus to H6/H8 (Figure 4.1.8: panel A) and

phosphorus to H1ʹ (Figure 4.1.8: panel B). Panel A in Figure 4.1.8 illustrates a sequential

assignment from U230 phosphorus to C232 H6 and C241 phosphorus to C243 H6. Panel

B in Figure 4.1.8 illustrates a sequential assignment from G229 H1ʹ to G231 H1ʹ and C241

H1ʹ to C243 H1ʹ. A sequential assignment from phosphorus to H6/H8/H1ʹ could not be

found for either the loop nucleotides or the G240 nucleotide.

210

Figure 4.1.7 Illustration of the identification of H6/H8 and H1ʹ peaks in the NOESY spectrum

with the aid of both 1H-

13C HSQC and

1H-

31P CPMG-HSQC-NOESY spectra, of the FMDV

15mer apo-RNA. (a) 600 MHz 1H-


2H2O,

displaying the phosphorus-H6/H8 peaks. (b) 400 MHz 1H-


2H2O,

displaying the C6-H6 and C8-H8 peaks. (c) 700 MHz NOESY (250ms) spectrum, at 25°C in 2H2O.

(d) 400 MHz 1H-


2H2O, displaying the C1ʹ-H1ʹ peaks. (e) 600 MHz

1H-

31P CPMG-HSQC-NOESY spectrum at 25°C in

2H2O, displaying the phosphorus-H1ʹ peaks.

The dashed lines in panels (a) and (e) indicate phosphorus-H6/H8/H1ʹ peaks that could not be

identified in the 1H-


211



2H2O, of the FMDV

15mer apo-RNA. Illustration of the phosphorus-H6/H8 (panel A), and phosphorus-H1ʹ (panel B)

correlations. Sequential assignment between G229-C232 is represented by a blue line and between

G240-C243 is indicated by a green line, in both phosphorus-H6/H8 and phosphorus-H1ʹ regions.

Peaks marked with a cross represent phosphorus-proton correlations.

212

Table 4.1.1 1H,

13C and


1H2O and

2H2O.


G229 13.18 7.743 5.722 8.106 5.781 4.822 4.629 4.415 4.054 3.936 139.1 92.16 75.03 74.91 82.28 62.86

G229

U230 13.90 5.244 7.963 5.745 4.772 4.666 4.537 4.274 103.4 142.2 93.45 72.88 73.03 82.40 65.36 -4.431 U230

G231 12.72 7.984 5.779 7.764 5.807 4.512 4.550 4.216 141.0 92.94 75.56 82.48 -4.075 G231

C232 5.087 7.491 5.397 4.416 4.134 97.27 136.3 94.15 73.79 64.82 -4.606 C232

A233 8.195 5.809 4.684 4.803 141.0 93.25 72.72 74.89 -4.627 A233

A234 7.849 7.763 5.569 4.057 4.335 154.2 141.0 93.55 76.08 64.84 -4.658 A234

C235 5.392 7.551 3.913 4.310 97.27 142.1 64.98 -4.640 C235

C236 5.716 7.892 5.702 98.91 141.7 92.71 C236

C237 5.581 7.790 5.579 4.486 4.406 98.44 141.7 93.72 76.10 82.14 -4.392 C237

C238 5.706 7.895 5.679 4.537 4.383 4.171 98.05 141.7 93.21 76.08 82.74 65.13 -4.179 C238

A239 8.101 8.193 5.955 4.896 4.618 4.295 154.2 141.0 93.02 75.48 83.62 65.44 -3.754 A239

G240 12.78 7.627 5.557 4.567 4.452 4.507 4.136 140.5 93.41 76.10 64.83 G240

C241 8.571 6.994 5.313 7.765 5.491 4.442 97.23 141.0 93.98 75.32 -4.680 C241

A242 8.157 6.342 7.535 8.054 5.967 4.408 4.711 4.456 4.132 4.584 153.6 139.6 93.25 75.91 72.67 82.33 -4.280 A242

C243 8.298 7.072 5.283 7.408 5.701 3.908 4.144 4.010 97.70 141.4 92.57 77.61 83.48 -4.373 C243

213


Distance restraints were generated from the NOESY spectra of the 15mer apo-RNA.

Exchangeable NOE restraints were obtained from the NOESY spectrum of the 15mer apo-

RNA in 1H2O, with 400ms mixing time. Non-exchangeable NOE restraints were obtained

from the NOESY spectrum of the 15mer apo-RNA in 2H2O, with both 150ms and 250ms

mixing times.

All nucleotides except for A234, C235, C236 and C237 were restrained to the anti

conformation. No H6/H8-H1ʹ intranucleotide connectivities were observed for the four

loop nucleotides mentioned, so their glycosidic angles were not defined. All stem

nucleotides were restrained to the C3ʹ-endo conformation except for the C243 nucleotide,

which was restrained to the C2ʹ-endo conformation. The δ angle was not restrained for the

loop nucleotides.

Hydrogen bond restraints were added for all four base pairs. However, no connectivities

could be observed from the G240 imino proton in the NOESY (1H2O) spectrum so the

hydrogen bond restraints between C232-G240 were loosely restrained. The G229-C243

base pair was also loosely restrained as the terminal base pairs are subject to motion due to

the proximity to solution. Planarity restraints were added for three base pairs with the

exception of the C232-G240 base pair.



step, one violation was found in the distance restraints and hydrogen bond restraints. The

restraints that were used for these structure calculations are summarised in Table 4.1.2.

214

Restraints

NOEs 178

Strong 1.8 – 2.5 Å 9

Medium 2.6 – 3.3 Å 49

Weak 3.4 – 5.0 Å 115

Very weak 5.1 – 7.0 Å 5

Inter-nucleotide NOEs 87

Intra-nucleotide NOEs 91

Hydrogen bonds 24

Planarity 6

Dihedral angles 71

Helix (, , , ε, ζ) 36


Glycosidic (χ) 11

NOE per residue 11.9

Restraints per

residue

18.6



FMDV 15mer apo-RNA.

215





calculated to be 0.51 Å.

Figure 4.1.9 Illustration of the NMR solution structures of the FMDV 15mer apo-RNA. (a)

Overlay of the 20 lowest RMSD structures with average RMSD of 0.51Å. (b) Lowest RMSD

solution structure (0.35Å). The red ribbon represents the RNA backbone.


homogeneity of the 15mer apo-RNA tertiary structure. The RMSD of the 15mer apo-RNA

is good considering the dynamic nature of the heptaloop, which considerably increases the

RMSD of the whole 15mer apo-RNA structure. When considering the helical stem and the

216

tetraloop separately, a significant difference in RMSD was found. The average RMSD of

the helical stem alone was 0.33Å and of the tetraloop alone was 0.66Å. From the

ensemble of 20 structures, the structure with the lowest RMSD (0.35Å) was selected for

solution structure analysis and conformational analysis (Figure 4.1.9b).

4.1.3.2 Heptaloop

A heptaloop was found for the 15mer apo-RNA structure (Figure 4.1.10). Since the loop

contains seven nucleotides, it was assumed that the loop structure will be less stable

compared to the helical stem region. Loop regions in RNA structures are stabilised by

base stacking interactions and intramolecular interactions. In Figure 4.1.10, the A233 and

A234 bases were stacked together on the 5ʹ-end, while bases C237, C238 and A239 were

stacked on the 3ʹ-end. The C235 and C236 bases look as though they are not stacked with

either the 5ʹ-end or 3ʹ-end bases. These base stacking interactions will significantly

contribute to the stabilisation of the entire heptaloop region.

Figure 4.1.10 The heptaloop (A233ACCCCA239) of the FMDV 15mer apo-RNA NMR structure,

shown in Figure 4.1.9b. Colour coding of nucleotides: adenosine (green) and cytosine (red).

217


Fourteen intramolecular interactions were identified in total of which twelve were found

to be non-specific (Table 4.1.3). Six base-phosphate interactions were identified, all

involving the O5ʹ phosphate oxygen and the aromatic H6/H8 protons. Three sugar-

phosphate interactions were identified, which involved the O3ʹ phosphate oxygen and the

2ʹ-hydroxyl group of the sugar ribose. One base-sugar interaction was identified between

the 2ʹ-OH group of the sugar ribose and the O5ʹ phosphate oxygen atom. Two sugar-sugar

interactions were found between the 2ʹ-OH and O4ʹ atoms, in the sugar ribose. Only two

specific interactions were identified in the heptaloop, both involving base-base

interactions. Despite the dynamic nature of such a large heptaloop, these intramolecular

interactions will increase the stability of the loop and therefore the entire 15mer RNA

structure.


interaction


A233 N1 – A239 NH6 base-base specific 3.01

C236 NH4 – C235 N3 base-base specific 2.01


A233 (2ʹ-OH) – A234 O4ʹ sugar-sugar non-specific 3.07


A234 (2ʹ-OH) – C235 O5ʹ base-sugar non-specific 2.57

C235 (2ʹ-OH) – C236 O3ʹ sugar-phosphate non-specific 2.73

C236 H6 – C236 O5ʹ base-phosphate non-specific 2.86


C237 H6– C237 O5ʹ base-phosphate non-specific 3.49


C238 H6 – C238 O5ʹ base-phosphate non-specific 2.91


A239 (2ʹ-OH) – G240 O4ʹ sugar-sugar non-specific 3.27

Table 4.1.3 Two specific and twelve non-specific intramolecular interactions were identified in

the heptaloop of the FMDV 15mer apo-RNA NMR structure. The interactions formed between

donor and acceptor atoms are given, the type of interaction, the specificity of the interaction and

the distances between the proton donor and acceptor atoms.

218


The 3DNA program was used to perform a conformational analysis on the FMDV 15mer

apo-RNA NMR structure. The local helical parameters, base pair step parameters and

complementary base pair parameters for the 15mer apo-RNA structure are displayed in

Tables 4.1.4, 4.1.5 and 4.1.6, respectively.

The x-displacement and slide parameter values of -4.23Å and -1.62Å, respectively, were

found for the 15mer apo-RNA structure, which is characteristic of an A-form helical

conformation. This was good considering that the 15mer RNA consisted of a large

heptaloop, which could have altered the A-form conformation of the stem region.

Base

Pair Nucleotides

Xdisp

(dx)

Ydisp

(dy)

Inclination

(η) Tip (θ)

Helical

Twist (Ω)

Helical

Rise (h)

1-2 G-C / U-A -3.97 0.22 15.73 1.94 34.61 2.45

2-3 U-A / G-C -3.93 0.81 11.45 -1.60 33.15 3.21

3-4 G-C / C-G -4.80 0.25 18.44 2.70 33.48 3.28

Ave. - -4.23 0.43 15.21 1.01 33.75 2.98

Table 4.1.4 Local helical parameter values for the FMDV 15mer apo-RNA structure, calculated

by 3DNA analysis program.

Base

Pair Nucleotides

Shift

(Dx)

Slide

(Dy) Rise (Dz) Tilt (τ) Roll (ρ) Twist (ω)

1-2 G-C / U-A -0.22 -1.64 2.97 -1.14 9.25 33.37

2-3 U-A / G-C -0.37 -1.59 3.59 0.91 6.49 32.52

3-4 G-C / C-G -0.31 -1.64 3.96 -1.53 10.45 31.81

Ave. - -0.30 -1.62 3.51 -0.59 8.73 32.57

Table 4.1.5 Base pair step parameter values for the FMDV 15mer apo-RNA structure, calculated

by 3DNA analysis program.

219

Base

Pair Nucleotides

Shear

(Sx)

Stretch

(Sy)

Stagger

(Sz)

Buckle

(κ)

Propeller

(π)

Opening

(σ)

1 G-C -0.86 0.03 0.01 2.51 2.99 3.84

2 U-A -0.07 -0.10 -0.24 10.43 -11.22 -4.65

3 G-C 0.27 -0.04 0.08 -0.77 -22.01 -2.87

4 C-G 0.38 -0.08 -0.12 -11.14 -2.83 -11.52

Table 4.1.6 Complementary base pair parameter values for the FMDV 15mer apo-RNA structure,

calculated by 3DNA analysis program.


CURVES programs (Table 4.1.7). The 3DNA program was used to generate dihedral

angles values for the stem base pair nucleotides. Therefore, eight dihedral angles values

were calculated by the 3DNA program. The CURVES program was used to obtain the

dihedral angle values of the seven loop nucleotides in the heptaloop.

The dihedral angle values calculated for the stem nucleotides were all found to be within

the ranges used in the structure calculations. However, a few significant deviations from

standard dihedral angles were found for the loop nucleotides; C236 (χ) angle, C238 (ε)

angle and C235 (α) angle. The deviation in the C235 (α) angle is due to the rotation

around the P-O5ʹ bond that forms the turn in the heptaloop. Interestingly, this deviation in

the (α) angle is also observed for the U179 nucleotide in the 16mer RNA NMR structures,

which marks the turning point of the loop from the 5ʹ-end to the 3ʹ-end.

220

Base Nucleotides C1ʹ-N

(χ)

C5ʹ-C4ʹ

(γ)

C4ʹ-C3ʹ

(δ)

C3ʹ-O3ʹ

(ε)

O3ʹ-P

(ζ)

P-O5ʹ

(α)

O5ʹ-C5ʹ

(β)

1 G -157.1 54.4 85.4 -156.1 -62.0 - -

2 U -158.4 53.9 80.5 -156.1 -70.9 -65.9 172.8

3 G -155.4 54.4 83.0 -152.0 -61.1 -62.1 173.8

4 C -155.9 52.9 82.4 -156.8 -88.8 -69.9 178.1

5 A -165.6 52.7 82.4 -156.1 -67.4 -70.2 -168.2

6 A -158.9 53.4 82.8 -153.6 -56.8 -66.1 -179.3

7 C -146.5 55.5 82.2 -155.9 -84.8 179.6 175.0

8 C -129.0 52.0 86.5 -157.2 -59.5 -90.6 179.3

9 C -158.9 53.7 83.0 -165.8 -95.7 -79.0 -175.5

10 C -159.5 53.1 82.4 -133.5 -53.1 -67.5 174.4

11 A -168.2 63.9 85.8 -153.9 -62.9 -66.3 171.7

12 G -158.2 57.0 87.1 -153.8 -66.0 -65.0 172.9

13 C -157.4 54.2 83.2 -153.3 -60.1 -67.3 177.7

14 A -169.1 54.8 81.1 -156.8 -68.8 -66.3 171.0

15 C -132.3 66.6 149.0 - - -63.7 -175.9


calculated by the 3DNA (black) and CURVES (red) analysis programs. Angles are all measured in

degrees.

Dihedral angle values (ν1 and ν2) that define the sugar ribose conformation were also

calculated by the 3DNA and CURVES programs, including the pseudorotation phase

angle and the amplitude (Table 4.1.8). Eight out of the fifteen nucleotides were calculated

to have a C3ʹ-endo sugar ribose conformation and one C2ʹ-endo sugar ribose conformation.

The C4ʹ-exo conformation was revealed for four nucleotides and two C2ʹ-exo

conformations were found. The stem C232 nucleotide, whose sugar ribose adopted a C4ʹ-

exo conformation, did not deviate significantly from the C3ʹ-endo conformation.

Conversely, the loop A234 and C237 nucleotides also adopted C4ʹ-exo conformations, but

the deviation from the C3ʹ-endo conformation was found to be greater. This is most likely

due to the dynamic nature of the loop nucleotides. The C2ʹ-exo conformation was found

for the G229 and G240 nucleotides with phase angles of very close to the phase angle

corresponding to the C3ʹ-endo conformation.

221

Base Nucleotides ν1 v2 Amp Phase Conformation

1 G -36.0 41.7 41.7 357.7 C2ʹ-exo

2 U -26.7 40.0 41.7 16.3 C3ʹ-endo

3 G -28.3 39.3 40.2 12.0 C3ʹ-endo

4 C -11.8 29.1 36.5 37.2 C4ʹ-exo

5 A -18.8 33.6 38.5 26.0 C3ʹ-endo

6 A -4.5 24.6 38.1 48.5 C4ʹ-exo

7 C -15.6 32.5 39.1 31.6 C3ʹ-endo

8 C -10.4 25.7 33.0 37.0 C4ʹ-exo

9 C 3.4 19.0 39.5 60.3 C4ʹ-exo

10 C -15.9 32.8 39.6 31.5 C3ʹ-endo

11 A -22.8 34.2 36.9 17.2 C3ʹ-endo

12 G -35.5 40.2 40.4 355.6 C2ʹ-exo

13 C -34.4 43.2 43.3 4.2 C3ʹ-endo

14 A -29.0 42.6 44.2 15.3 C3ʹ-endo

15 C 36.2 -37.3 38.2 167.3 C2ʹ-endo



apo-RNA structure. Values were calculated by the 3DNA (black) and CURVES (red) programs.

222

4.1.5 Comparison of the 15mer and 16mer apo-RNA NMR structures

Analogous to the 16mer apo-RNA and Mg2+

RNA complex structures, the successful

determination of the 15mer and 16mer apo-RNA structures allowed for a review of their

similarities and differences.

The number of NOE restraints per residue generated for the 15mer and 16mer apo-RNA

structures was 11.9 and 7.8, respectively. Relatively, 35% fewer NOEs per residue were

used for the 16mer apo-RNA structure, which was attributed to the presence of broad lines

in the NOESY (1H2O/

2H2O) spectra of the 16mer apo-RNA. By contrast, the sensitivity

observed in the NOESY (1H2O/

2H2O) spectra of the 15mer apo-RNA was much better,

which allowed easier assignment of NOE cross peaks. However, as the 15mer RNA

contained a heptaloop, the assignment of this region proved much more difficult.

Consequently, the assignment of the 15mer apo-RNA involved a greater amount of effort

in order to assign more internucleotide connectivities, which were required to sufficiently

restrain the heptaloop region. The final twenty 15mer apo-RNA NMR structures generated

had low RMSDs, when considering the large heptaloop.

Structure validation using the Molprobity program revealed no bad bonds, angles, sugar

pucker and backbone conformations for the 15mer and 16mer apo-RNA NMR structures.

Clash scores were found to be low; 31.2 and 12.5 for the 15mer and 16mer apo-RNA

structures, respectively. The final solution structures of both the 16mer and 15mer apo-

RNAs were of a high standard. This justified the investigation of possible RNA-RNA

interactions between the 16mer and 15mer RNAs.

223

4.2 Effect of Mg2+

on the 15mer RNA

A comparison was made between the imino regions of the 15mer apo-RNA and Mg2+

-

RNA complex (Figure 4.2.1). A large, lowfield chemical shift change was observed for

the U230 imino proton peak (Δδ=0.19ppm). This lowfield shift suggests a conformational

change possibly caused by improved stacking of the U230 base with the G229 base. A

second lowfield shift was also observed for the G240 imino proton (Δδ=0.07ppm).

Another interesting observation was that the G229 imino proton intensity was

significantly reduced in the 15mer Mg2+

RNA complex. This was likely to be caused by

faster exchange with water as G229 is part of the terminal base pair and the hydrated Mg2+

ions are more likely to affect the terminal base pairs. These findings provided the first

evidence of Mg2+

-induced effects in the 15mer RNA.

Figure 4.2.1 600MHz 1H-NMR stack plot (imino region) of the 15mer RNA with (a) no Mg

2+ and

(b) in the presence of 5.0eq Mg2+

, at 2°C in 1H2O.

In the imino region of the NOESY (1H2O) spectrum, imino-imino connectivities were

observed between U230-G231 (Figure 4.2.2); the same was observed for the 15mer apo-

224

RNA. Interestingly, in the 15mer Mg2+

RNA complex, an additional imino-imino

connectivity was found between G231-G240, which was not observed in the 15mer apo-

RNA. This cross-strand internucleotide connectivity was observed possibly due to the

stabilising affect conferred by the Mg2+

ions or due to a shorter distance between the

imino protons of G231 and G240.


RNA complex, at

2°C in 1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks correspond to

imino-imino connectivities. The sequential assignment shown here is between U230-G231 and

G231-G240. Inset: Secondary structure of the 15mer RNA highlighting the observed imino-imino

connectivities, represented by light blue oval shapes.

225

In the imino-amino region of the NOESY (1H2O) spectrum (Figure 4.2.3), connectivities

were observed from the G240 imino proton. In contrast, connectivities from G240 NH

could not be observed in the 15mer apo-RNA, due to the overlapping of the G231 NH.

Consequently, the C232 NH2* and NH proton chemical shifts were identified in the

15mer Mg2+

RNA complex.


RNA complex, at

2°C in 1H2O, illustrating the imino-amino region of the spectrum. Connectivities from imino

protons to NH2*/NH2/H2 protons can be observed (NH2* corresponds to the proton involved in

base pair hydrogen bonding); connectivities are marked by a black circle.

226

Chemical shift changes were found to both the imino and amino exchangeable protons in

the NOESY (1H2O) spectrum. Additionally, chemical shift changes were also observed to

the non-exchangeable protons in the NOESY (2H2O) spectrum. Figure 4.2.4 summarises

the most significant chemical shift changes observed in the NOESY spectra. These results

provided further evidence of Mg2+

-induced structural changes to the RNA structure.

Figure 4.2.4 Histogram illustrating the changes to the chemical shifts of exchangeable and non-

exchangeable protons of the FMDV 15mer RNA, upon addition of 5eq of Mg2+

. Bars shown in

blue represent exchangeable protons. Bars shown in green and orange represent non-exchangeable

protons in the stem and loop, respectively. Negative values correspond to a highfield chemical

shift change and positive values correspond to a lowfield chemical shift change.

227

4.3 1GHz NMR studies of the 16mer apo-RNA

4.3.1 Effect of magnetic field strength on the 16mer apo-RNA

Access to the 1GHz spectrometer provided a great opportunity to acquire NMR spectra at

the highest possible magnetic field strength. This was very important has higher magnetic

field strengths generally lead to higher sensitivity in NMR experiments. The 16mer apo-

RNA (batch 2) was chosen for NMR study at 1GHz, in which a 1H-NMR and NOESY

experiment was performed in 1H2O.

The 1GHz 1H-NMR spectrum revealed an interesting result, in which the U176 imino

proton peak appeared significantly reduced in intensity compared to spectra acquired

previously at lower magnetic field strengths. To monitor the effect of magnetic field

strength on the intensity of the imino proton peaks, 1H-NMR experiments were performed

at four different magnetic field strengths (Figure 4.3.1). The stack plot in Figure 4.3.1

clearly showed that the intensity of the U176 imino proton peak decreased dramatically

with increasing magnetic field strength. Similarly, a decrease in intensity was also found

for the G178 imino proton peak; this decrease was not as large and could only be clearly

observed when comparing the G178 and G177 imino proton peak intensities in the 400

MHz and 1GHz 1H-NMR spectra.

A possible explanation for the change in intensity found for the U176 imino proton peak

could be attributed to conformational exchange of the U176 imino proton. If there are two

environments for the U176 imino proton, it is likely that they are in fast exchange since

only one imino proton peak is observed for U176. As the magnetic field strength is

increased, so is the separation in Hz of the two electronic environments. Therefore, the

exchange between the two species is observed as intermediate exchange on the NMR

timescale, which leads to broadening of the peak and reduced intensity.

By measuring the linewidth of individual imino proton peaks, it was possible to measure

the contribution of exchange broadening to the linewidth. For U175, G177, G185 and

228

G186 imino proton peaks, the linewidth was unaffected with increasing magnetic field

strength. Conversely, the linewidth increased for the U176 and G178 imino proton peaks,

with increasing magnetic field strength. This increase in linewidth is attributed to

exchange broadening.

Figure 4.3.1 1H-NMR stack plot of the 16mer apo-RNA (batch 2), at 2°C in

1H2O, displaying the

imino proton region with four different magnetic field strengths; (a) 400 MHz, (b) 600 MHz, (c)

800 MHz and (d) 1000 MHz. The intensity of the U176 and G178 imino proton peak is clearly

reduced with increasing magnetic field strength, marked by the red arrows.

229

4.3.2 Sensitivity enhancement with 1GHz

The NOESY (1H2O) spectrum of the 16mer apo-RNA (batch 2) was also analysed.

Fascinatingly, an additional cross peak was observed from the G178 imino proton in the

imino-amino region (Figure 4.3.2). NOE cross peaks were difficult to observe from the

G178 imino proton, largely due to the fast exchange with water. These NOE cross peaks

were crucial for generating distance restraints for structure determination, in order to

sufficiently restrain the GNRA tetraloop region. The 1GHz spectrometer offered greater

sensitivity, which allowed the assignment of a new internucleotide connectivity (G178

NH – C182 NH2*). This demonstrates the advantage of using the highest magnetic field

strength possible, especially for RNA structures with dynamic regions.

Figure 4.3.2 Illustration of the NOESY (150ms) spectrum (imino-amino region) of the (a) 16mer

apo-RNA (batch 1) at 700 MHz and (b) 16mer apo-RNA (batch 2) at 1GHz. The G178 NH to

C182 NH2* NOE cross peak can be clearly observed in the 1GHz NOESY spectrum, but is absent

in the 700 MHz NOESY spectrum.

230

4.4 RNA-RNA interaction

4.4.1 Analysis of the RNA-RNA complex in 1H2O

A comparison was made between the imino proton regions of the 16mer Mg2+

RNA

complex (batch 2), the 15mer Mg2+

RNA complex and the RNA-RNA complex (Figure

4.4.1). In the RNA-RNA complex, the G178 loop imino proton peak was clearly observed

highfield at 10.86ppm. Interestingly, a broad peak was also observed highfield of the

G178 imino proton peak in the 16mer Mg2+

RNA complex. This peak may correspond to

the G178 imino proton in a minor conformation, as the chemical shift matches that

observed in the 16mer apo-RNA. The U176, G177 and G185 imino proton peaks were

also observed without any overlapping of the 15mer Mg2+

RNA peaks. However, the

U175 imino proton peak was overlapped with U230 and G186 was overlapped with G231.

Both G229 and G240 imino proton peaks of the 15mer Mg2+

RNA could not be clearly

identified in the RNA-RNA complex.

No significant chemical shift changes were observed for the imino proton peaks of the

16mer and 15mer Mg2+

RNAs. However, changes to the intensity and linewidth were

observed for the G178 imino proton peak. In the RNA-RNA complex, the line intensity of

the G178 imino proton peak was notably reduced and the linewidth had increased. This

may be caused by an increase in the imino proton exchange with water. Therefore, to

provide evidence of this increase in exchange, the imino proton exchange rate was

calculated for the G178 imino proton. Fascinatingly, a dramatic increase in exchange rate

was found for the G178 imino proton, from 37.0s-1

to 145.0s-1

. This significant change in

exchange rate provided the first evidence of a possible interaction between the 16mer and

15mer Mg2+

RNAs. The only explanation was that there may be a change in dynamics of

the 16mer loop region, allowing more solvent exposure to the major groove of the G178

base. Furthermore, the unknown peak found at 10.56ppm was not observed in the RNA-

RNA complex. In light of the increased exchange rate found for G178 imino proton, it is

possible that this peak is no longer visible due to fast exchange.

231

Figure 4.4.1 1H-NMR stack plot (imino region) of the (a) 16mer Mg

2+ RNA complex (600MHz),

(b) 15mer Mg2+

RNA complex (600MHz) and (c) RNA-RNA complex (700MHz), at 2°C in 1H2O.

In the NOESY (1H2O) spectrum of the RNA-RNA complex, the connectivities in the

imino-imino and imino-amino region were assigned. Figure 4.4.2 displays the imino-

imino connectivities observed for the 16mer Mg2+

RNA (G178-G186) and also for the

15mer Mg2+

RNA (U230-G240). The pattern of imino-imino cross peaks was found to be

similar to that of both RNAs alone. These results suggest that it is unlikely that any major

conformational changes have occurred in the stem region of the 16mer or 15mer Mg2+

RNA tertiary structures. Analogously, connectivities for both RNAs were assigned in the

imino-amino (Figure 4.4.3) and aromatic regions of the NOESY (1H2O) spectrum.

Subsequently, chemical shift changes for both RNAs were identified in the NOESY (1H2O)

spectrum (Figure 4.4.5). For the 16mer Mg2+

RNA, large chemical shift changes were

found to the U176 H5 and H6 protons; 0.12ppm and 0.07ppm, respectively. Interestingly,

232

these chemical shifts match those of the U176 H5/H6 protons in the 16mer apo-RNA,

suggesting that the Mg2+

-induced effect on these protons had been reversed. A chemical

shift change was also observed for the C173 H5 proton of the 16mer RNA (Δδ=0.06ppm).

For the 15mer Mg2+

RNA, four significant chemical shift changes were found in the stem

region; U230 H5 (Δδ=0.05ppm), A242 NH2* (Δδ=0.07ppm), C243 NH2* (Δδ=0.1ppm)

and C243 H1ʹ (Δδ=0.05ppm). These results suggest a possible RNA-RNA interaction

involving the stem region of the 15mer Mg2+

RNA.

Figure 4.4.2 700 MHz NOESY (250ms) spectrum of the FMDV RNA-RNA complex, at 2°C in

1H2O, illustrating the imino region of the spectrum. The cross-diagonal peaks correspond to imino-

imino connectivities. The sequential assignment of the 16mer RNA starts from G178 and finishes

at G186 (black lines). The sequential assignment of the 15mer RNA starts from U230 and finishes

at G240 (blue lines) Inset: Secondary structure of the 16mer RNA (top) and 15mer RNA (bottom)

highlighting the imino-imino connectivities shown in the spectrum, represented by light blue oval

shapes.

233

Figure 4.4.3 700 MHz NOESY (250ms) spectrum of the FMDV RNA-RNA complex, at 2°C in

1H2O, illustrating the imino-amino region of the spectrum. Connectivities from imino protons to

NH2*/NH2/H2/H5/H1ʹ protons can be observed. (NH2* corresponds to the proton involved in

base pair hydrogen bonding); connectivities are marked by a black and blue circle for the 16mer

RNA and 15mer RNA, respectively. Assignments for both the 16mer RNA (black) and 15mer

RNA (blue) are shown.

234

4.4.2 Analysis of the RNA-RNA complex in 2H2O

Subsequently, the RNA-RNA complex was constituted in 2H2O. The

1H-NMR spectrum

of the RNA-RNA complex was compared with the 1H-NMR spectra of the 16mer and

15mer Mg2+

RNAs (Figure 4.4.4). The H6/H8 aromatic protons were observed between

6.6-8.5ppm and H5/H1ʹ protons were observed between 5.1-6.2ppm, in all three spectra.

In order to detect any changes to the chemical shift or linewidth of peaks in the RNA-

RNA complex, specific peaks were selected within the 1H-NMR spectra of the 16mer and

15mer Mg2+

RNAs. These peaks were selected on the basis that they could be clearly

observed in the RNA-RNA complex and that the peaks did not overlap between the 16mer

and 15mer Mg2+

RNAs.

For the 16mer Mg2+

RNA, six peaks were identified corresponding to A183 H2,

G185/G186 H8, A181 H8, A187 H2, A181 H1ʹ and C182 H5. The A183 H2 and

G185/G186 H8 proton peaks can be found in a very distinct region of the spectrum and

can be easily identified in the RNA-RNA complex. Interestingly, these two peaks are

broader in the RNA-RNA complex. The line broadening could be caused by shorter T2

relaxation due to a longer correlation time, which would indicate RNA-RNA complex

formation. Conversely, the A181 H8 peak was found to be more intense in the RNA-RNA

complex. For the 15mer Mg2+

RNA, three peaks were identified corresponding to

A233/A239 H8, A242 H1ʹ and A239 H1ʹ. Interestingly, small chemical shift changes were

observed for the A242 H1ʹ and A239 H1ʹ peaks.

On the whole, small changes in chemical shift, line intensity and linewidth, indicated a

possible weak RNA-RNA interaction. These results did not conclusively point towards a

specific binding site, but there was enough evidence to suggest changes, structural and/or

dynamic, to the 16mer and 15mer Mg2+

RNAs within the RNA-RNA complex.

235

Figure 4.4.4 400 MHz 1H-NMR stack plot of the (a) 16mer Mg

2+ RNA complex, (b) 15mer Mg

2+

RNA complex and (c) RNA-RNA complex, at 25°C in 2H2O The red asterisk indicates the peaks

of interest that show changes in chemical shift or linewidth.

Analysis of the NOESY (2H2O) spectrum of the RNA-RNA complex did not reveal any

significant chemical shift changes except for U230 H5 (Δδ=0.06ppm) in the 15mer Mg2+

RNA. Interestingly, the number of chemical shift changes observed in the NOESY (2H2O)

spectrum was much less when compared to the number observed in the NOESY (1H2O)

spectrum (Figure 4.4.5). This may be attributed to the increase in the temperature, 2°C to

25°C, when observing NOESY spectra in 1H2O and

2H2O, respectively. A higher

temperature could reduce the RNA-RNA interactions between the 16mer and 15mer Mg2+

RNAs and obscure any observable changes in chemical shift.

236

Figure 4.4.5 Histogram illustrating the changes to the chemical shifts of exchangeable and non-

exchangeable protons of the FMDV 16mer and 15mer Mg2+

RNAs, upon RNA-RNA complex

formation. Bars shown in blue and red represent proton chemical shift changes identified in the

NOESY spectra in 1H2O and

2H2O, respectively. Negative values correspond to a highfield

chemical shift change and positive values correspond to a lowfield chemical shift change.

In the NOESY (2H2O) spectrum of the RNA-RNA complex, the H6/H8-H1ʹ sequential

assignment was attempted for both the 16mer and 15mer Mg2+

RNAs. Figure 4.4.6

displays the sequential assignment observed for the 16mer and 15mer Mg2+

RNAs in the

NOESY (2H2O) spectrum of the RNA-RNA complex. A near full assignment was

achieved for the 16mer Mg2+

RNA from C173 H6-H1ʹ to G177 H8-H1ʹ and U179 H1ʹ-

A180 H8 to A187 H8-H1ʹ. By contrast, the same sequential assignment produced for the

16mer Mg2+

RNA alone had a very similar sequential pattern indicating that the initial

16mer Mg2+

RNA structure had not changed conformation. Two short sequential

assignments were achieved for the 15mer Mg2+

RNA from U230 H1ʹ-G231 H8 and C241

H6-H1ʹ to C243 H6-H1ʹ. Again this sequential pattern was found to be very similar to that

observed for the 15mer Mg2+

RNA alone. No intra- or internucleotide connectivities were

observed for nucleotides found in the 15mer loop region.

237

Figure 4.4.6 Top and bottom panels: 600 MHz NOESY (250ms) spectrum of the FMDV RNA-

RNA complex, at 25°C in 2H2O. The black and blue lines represent the sequential H6/H8-H1ʹ

intra- and internucleotide connectivities, for the 16mer and 15mer Mg2+

RNAs, respectively. The

(i) corresponds to an intranucleotide connectivity and (s) corresponds to a sequential connectivity.

The red circles and light blue squares correspond to H5-H6 connectivities for the 16mer and

15mer Mg2+

RNAs, respectively. Assignments for both the 16mer Mg2+

RNA (black) and 15mer

Mg2+

RNA (blue) are shown.

238

4.4.3 Model of the RNA-RNA interaction

With the information gathered from the analysis of the RNA-RNA complex, a model of

the possible RNA-RNA interaction was proposed between the 16mer and the 15mer

RNAs. Firstly, the potential regions involved in the RNA-RNA interaction were identified

for the 16mer RNA and the 15mer RNA.

In the 16mer RNA, the G178 imino proton was found to have an increased exchange rate.

As previously stated, this may have been caused by a change in dynamics of the GNRA

tetraloop, prompted by the close proximity of the 15mer RNA. Changes were also

observed to the line intensity of the A181 H8 peak. Additionally, chemical shift changes

were found to the U176 H5 and H6 protons, and linewidth changes to A183 H2. Therefore,

the G178-A181 base pair in the 16mer loop and the U176-A183 base pair in the 16mer

stem were found to be possible sites of interaction. In the 15mer RNA, chemical shift

changes to protons in the U230 and A242 nucleotides pointed to towards a possible site of

interaction around the U230-A242 base pair.

Figure 4.4.7 is a scheme that illustrates the possible RNA-RNA interaction between the

16mer and 15mer RNAs. The possible sites of interaction have been highlighted along

with the specific nucleotides in which chemical shift and linewidth changes were observed.

From this model it was hypothesised that the 16mer RNA and 15mer RNA may have

formed a heterodimer complex, whereby the 16mer RNA loop interacts with the stem

region of the 15mer RNA and the 15mer RNA loop interacts with stem region of the

16mer RNA. However, this may be one of many interactions occurring between the two

RNAs since the interaction was found to be weak.

239

Figure 4.4.7 Scheme illustrating the possible RNA-RNA interaction between the FMDV 16mer

RNA (left) and the 15mer RNA (right). The rectangles represent the bases, which have been

numbered. Base pairing is represented by three black lines for G.C base pairs and two black lines

for A.U base pairs. The unfilled circles correspond to the sugar ribose, red circles represent the

phosphorus atom and green triangles symbolise the base stacking interactions. Chemical shift and

linewidth changes to protons in nucleotides are represented by the blue filled rectangles and circles.

The broken black line between the orange brackets indicates the possible area of RNA-RNA

interaction.

240

Chapter 5: 19

F-NMR studies of selectively fluorinated RNAs

This chapter will focus on studying the two fluorinated RNAs, the 19

F labelled 16mer and

15mer RNA motifs of the FMDV IRES, by NMR spectroscopy. Firstly, the fluorination of

the RNAs will be confirmed by NMR, followed by assessing the influence of the 19

F

nucleus on proton and phosphorus chemical shifts. The chapter will conclude by studying

the effects of Mg2+

and investigating RNA-RNA interactions.

5.1 19F-NMR studies of the 5-FU 16mer and 15mer RNAs

5.1.1 Identification of fluorination

19F-NMR experiments were performed on both the 5-FU 16mer and 15mer RNAs in order

to identify and confirm that a 19

F label had been incorporated into the two RNA samples.

For the 5-FU 16mer RNA, a broad fluorine peak was identified corresponding to U179 F5

(-165.7ppm) at a temperature of 2°C (Figure 5.1.1). To increase the intensity of the

fluorine peak the 19

F-NMR experiments were performed at higher temperatures of 10°C

and 25°C whereby distinct chemical shifts for the fluorine peak were observed at three

different temperatures (Figure 5.1.1). A change in chemical shift of 0.27ppm was

observed between 2°C and 10°C, and 0.44ppm between 10°C and 25°C. These results

strongly suggest that different conformations of the U179 base are present at different

temperatures, most likely attributed to differences in base stacking.

Analogously, distinct fluorine chemical shifts were found for the 5-FU 15mer RNA

corresponding to U230 F5, at the same three temperatures of 2°C, 10°C and 25°C (Figure

5.1.2). A change in chemical shift of 0.22ppm was observed between 2°C and 10°C, and

0.41ppm between 10°C and 25°C. Interestingly, these chemical shift differences were very

similar to that found for the 5-FU 16mer RNA. This implies that the 19

F nucleus is

sensitive to base stacking for both the stem and loop fluorinated bases. In contrast to the 5-

FU 16mer RNA, the chemical shift of U230 F5 was found highfield (-168.3ppm) at a

241

temperature of 2°C. Generally, fluorine nuclei are shielded by neighbouring stacked bases

in A-form RNA, but are deshielded in unpaired regions of RNA. This variation in

chemical shift has been previously observed between Watson-Crick and unpaired 5-FU

nucleotides.114

More importantly, it demonstrates that the 19

F label is a sensitive marker of

RNA secondary structure, as one can distinguish between F5 chemical shifts of stem and

loop 5-FU nucleotides.

Figure 5.1.1 376 MHz 19

F-NMR stack plot of the 5-FU 16mer RNA, in 1H2O at (a) 2°C, (b) 10°C

and (c) 25°C. 19

F chemical shifts were referenced to CFCl3.

242

Figure 5.1.2 376 MHz 19

F-NMR stack plot of the 5-FU 15mer RNA, in 1H2O at (a) 2°C, (b) 10°C

and (c) 25°C. 19


The 19

F chemical shifts alone do not provide any information to allow the identification of

the fluorine peak to a specific nucleotide. Therefore, the next logical step was to identify

the nucleotide in which the fluorination had taken place. The best approach was to observe

the strong H5-H6 cross peaks in the NOESY spectra. The H5-H6 cross peak of the

fluorinated base should be absent due to fluorination at the H5 position. Figure 5.1.3 is an

overlay of the aromatic region of the NOESY (1H2O) spectrum, of the 16mer RNA and the

5-FU 16mer RNA (both apo-RNAs). The overlay clearly displays the seven H5-H6 peaks

from the 16mer RNA in green and six H5-H6 peaks from the 5-FU 16mer RNA in orange.

The missing cross peak from the 5-FU 16mer RNA is U179 H5-H6, which was to be

expected if the H5 position of U179 was fluorinated. Analogously, the U230 H5-H6 peak

was not observed in the 5-FU 15mer RNA, confirming the fluorination within U230

(Figure 5.1.4).

243

Figure 5.1.3 1GHz NOESY (150ms) spectrum of the 5-FU 16mer RNA at 2°C in 1H2O (orange)

overlaid on the 1GHz NOESY (250ms) spectrum of the unlabelled 16mer RNA (batch 2) at 2°C in

1H2O (green). The overlay displays the U172, C173, C174, U175, U176, U179 and C182 H5-H6

cross peaks found in the aromatic region of the spectra. Cross peaks of the unlabelled 16mer RNA

have been labelled by a cross.

Figure 5.1.4 600 MHz NOESY (400ms) spectrum of the 5-FU 15mer RNA at 2°C in 1H2O

(orange) overlaid on the 600 MHz NOESY (400ms) spectrum of the unlabelled 15mer RNA at

2°C in 1H2O (green). The overlay displays the U230, C232, C235, C236, C237, C238, C241 and

C243 H5-H6 cross peaks found in the aromatic region of the spectra. Cross peaks of the unlabelled

15mer RNA have been labelled by a cross.

244


F nucleus on the 5-FU 16mer RNA


The shielding or deshielding effects caused by the highly electronegative fluorine nucleus

can be studied by comparing proton chemical shifts in the 5-FU substituted RNAs. The

chemical shift of the 5-FU H6 proton will be influenced significantly due to the close

proximity to the fluorine nucleus, as will the imino proton within the same aromatic ring.

Generally, the H6 proton is deshielded by approximately 0.2ppm, while the imino NH

proton is deshielded by approximately 0.6-0.7ppm.114

For the 5-FU 16mer RNA, no

significant chemical shift changes were found to the stem or loop imino proton peaks

(Figure 5.1.5). The U179 imino proton could not be identified in either the 1H-NMR

spectrum of the 16mer apo-RNA or 5-FU 16mer RNA.

Figure 5.1.5 1GHz 1H-NMR stack plot (imino region) of the (a) 16mer apo-RNA (batch 2) and (b)

5-FU 16mer RNA, at 2°C in 1H2O. The U175, U176, G185, G186, G177 and G178 imino proton

peaks are labelled.

245

The NOESY spectra of the 16mer apo-RNA and 5-FU 16mer RNA were analysed in both

1H2O and

2H2O. Figure 5.1.6 displays the imino region of the NOESY (

1H2O) spectrum of

these two RNAs. The same imino-imino connectivities were observed in the fluorinated

16mer RNA when compared to the non-fluorinated 16mer RNA, indicating a similar local

helical conformation of the 5-FU 16mer RNA. This was further confirmed by the imino-

amino region of the NOESY (1H2O) spectrum (Figure 5.1.7), whereby the same NOE

patterns were observed as in the unlabelled 16mer RNA.

Figure 5.1.6 1GHz NOESY (250ms) spectra of the FMDV 16mer apo-RNA (batch 2), left, and 5-

FU 16mer apo-RNA, right, at 2°C in 1H2O, illustrating the imino region of the spectrum. The

cross-diagonal peaks correspond to imino-imino connectivities. The sequential assignment shown

in both spectra is between G178-G186, although the G177-U176 imino-imino connectivity is not

observed for the 5-FU 16mer RNA. Insets: Secondary structure of the 16mer RNA highlighting

the observed imino-imino connectivities, represented by light blue oval shapes.

246

Figure 5.1.7 1GHz NOESY (250ms) spectra of the FMDV 16mer apo-RNA (batch 2), left, and 5-

FU 16mer apo-RNA, right, at 2°C in 1H2O, illustrating the imino-amino region of the spectrum.

Connectivities from imino protons to NH2*/NH2/H2/H5/H1ʹ protons can be observed (NH2*

corresponds to the proton involved in base pair hydrogen bonding); connectivities are marked by a

black circle.


The H6/H8-H1ʹ region of the NOESY (2H2O) spectrum of 5-FU 16mer RNA was assigned

and a sequential assignment was produced (Figure 5.1.8). The sequential assignment

started from the U172 H6-H1ʹ intranucleotide connectivity and ended with G177 H8-H1ʹ

intranucleotide connectivity. The G177 H1ʹ-G178 H8, G178 H8-H1ʹ, G178 H1ʹ-U179 H6

and U179 H6-H1ʹ connectivities could not be observed. The sequential assignment was

247

commenced from U179 H1ʹ-A180 H8 and ended at A187 H8-H1ʹ. Subsequently, the

comparison of the NOESY (1H2O/

2H2O) spectra did not reveal any changes to proton

chemical shifts or NOE patterns. Evidently, this suggested that the presence of a 5-FU

nucleotide did not alter the conformation of the 16mer RNA.

Figure 5.1.8 Top and bottom panels: 600 MHz NOESY (400ms) spectrum of the FMDV 5-FU

16mer RNA, at 25°C in 2H2O. The blue line represents intra- and internucleotide connectivities

from U172 H1ʹ to G177 H8 and the green line from U179 H1ʹ to A187 H8 connectivities. The (i)

corresponds to an intranucleotide connectivity and (s) corresponds to a sequential connectivity.

The red circles correspond to H5-H6 connectivities.

5.1.2.3 The 2D 1H-

19F HOESY experiment

The chemical shift of the U179 H6 proton could not be identified in the NOESY (2H2O)

spectrum. However, the U179 H6 proton could be clearly observed in the 1H-

19F HOESY

spectrum (Figure 5.1.9). A lowfield chemical shift change of 0.18ppm was observed,

which is consistent with studies on 5-FU substituted RNA. Furthermore, one additional

248

cross peak was observed corresponding to a correlation between U179 F5 and the U179

H2ʹ proton. This was the first 1H-

19F HOESY experiment that was performed and since it

was successful it justified the continuation of studying the effect of Mg2+

and investigating

RNA-RNA interactions using this technique.


19F HOESY (250ms) spectrum of the FMDV 5-FU 16mer RNA, at

25°C in 2H2O.

19F chemical shifts were referenced to CF3COOH.

5.1.2.4 Effect of the 19

F nucleus on 31

P chemical shifts

A 31

P-NMR experiment was performed to observe any changes to the phosphorus

chemical shifts with the addition of a 5-FU nucleotide. These chemical shift changes can

be caused by the influence of fluorine on the phosphorus nucleus or changes in

conformation to the phosphate backbone. A comparison was made between the 31

P-NMR

spectrum of the 16mer apo-RNA (batch 1) and 5-FU 16mer RNA (Figure 5.1.10). Nine

peaks were originally identified in the 16mer apo-RNA; peaks 1, 2 and 9 were identified

as U179, A180/C182 and A181, respectively. The U179 phosphorus peak could not be

observed in the 5-FU 16mer RNA, possibly due to the lower sample concentration.

However, it was also possible that U179 phosphorus has shifted highfield into the large

phosphate peak. Additionally, peaks 2-9 exhibited the same chemical shifts in the 5-FU

16mer RNA when compared to 16mer apo-RNA. These results strongly suggest that the

phosphate backbone of the 16mer RNA has not been perturbed with the substitution of the

5-FU nucleotide.

249

Figure 5.1.10 162 MHz 31

P-NMR stack plot of the (a) 16mer apo-RNA (batch 1) and (b) 5-FU

16mer RNA, at 2°C in 1H2O. Peaks in the

31P-NMR spectrum of the 16mer apo-RNA are labelled

1-9. Peaks 1, 2 and 9 correspond to U179, A180/C182 and A181 phosphorus (labelled in red),

respectively.

250

5.1.3 Effect of magnetic field strength on the 5-FU 16mer RNA

Analogous to the 16mer RNA (batch 2), 1H-NMR experiments for the 5-FU 16mer RNA,

at four different magnetic field strengths, revealed a significant decrease in intensity of the

U176 and G178 imino proton peaks (Figure 5.1.11). This provided additional

confirmation that this effect was specific for the 16mer RNA sequence, regardless of the

actual sample.

Figure 5.1.11 1H-NMR stack plot of the 5-FU 16mer RNA, at 2°C in

1H2O, displaying the imino

proton region with four different magnetic field strengths; (a) 400 MHz, (b) 600 MHz, (c) 800

MHz and (d) 1000 MHz. The intensity of the U176 and G178 imino proton peak is clearly reduced

with increasing magnetic field strength, marked by the red arrows.

251


F nucleus on the 5-FU 15mer RNA

The 1D 1H-NMR and 2D NOESY (

1H2O) spectra of the 15mer apo-RNA and 5-FU 15mer

RNA were compared. The imino region of the 1H-NMR spectrum in

1H2O was analysed

(Figure 5.1.12). Two fascinating observations were found. Firstly, the U230 imino proton

chemical shift of the 5-FU 15mer RNA shifted lowfield (Δδ=0.78ppm), which is

consistent with published values.114

The deshielding affect is caused by the highly

electronegative fluorine nucleus reducing the electron density around the imino proton.

Secondly, the intensity of the U230 imino proton peak had significantly reduced and the

linewidth increased. This observation was explained by an increase in exchange of the

U230 imino proton with water; the electronegativity of fluorine reduces the pKa value of

the N3 nitrogen resulting in the increase in imino proton exchange.

Figure 5.1.12 A 1H-NMR stack plot (imino region) of the (a) 15mer apo-RNA (600 MHz) and (b)

5-FU 15mer RNA (800 MHz), at 2°C in 1H2O. A lowfield shift of 0.78ppm is observed for the

U230 imino proton of the 5-FU 15mer RNA.

252

The U230 H6 chemical shift was identified using the NOESY (1H2O) spectrum by

observing NOE patterns corresponding to intranucleotide connectivities from U230 H6 to

its sugar protons. A lowfield chemical shift change of 0.33ppm was observed for the U230

H6 proton. No significant chemical shift changes could be found for the U230 sugar

protons. Interestingly, lowfield chemical shift changes were also found for protons in the

base paired A242 base; A242 NH2* (Δδ=0.29ppm) and A242 NH2 (Δδ=0.12ppm). These

results highlighted the local influence of the fluorine nucleus on proton chemical shifts.

Interestingly, 31

P-NMR experiments revealed that the U230 phosphorus peak shifted

highfield by 0.52ppm in the 5-FU 15mer RNA (Figure 5.1.13), although this was not

confirmed. In addition, some phosphorus chemical shifts appeared to be different between

the two samples. This would suggest that the RNA backbone of the two structures may be

slightly different, but this effect could also arise if the temperature effects are different for

the two samples.

Figure 5.1.13 162 MHz 31

P-NMR stack lot of the (a) 15mer apo-RNA and (b) 5-FU 15mer RNA,

at 25°C in 1H2O. A highfield shift of 0.52ppm is observed for the U230 phosphorus in the 5-FU

15mer RNA.

253


on the 19

F signal

19F-NMR experiments were performed on the 5-FU 15mer and 16mer RNA samples, with

and without Mg2+

. The objective was to investigate whether any chemical shift changes to

the 19

F signal could be observed with the addition of Mg2+

. Fascinatingly, for the 5-FU

16mer RNA, a highfield chemical shift change of 0.27ppm was observed (Figure 5.1.14).

The highfield shift of the U179 F5 peak suggests better base stacking of U179 since

fluorine signals in stem nucleotides are found further highfield. This supports the

observation found in the NMR solution structures, whereby the U179 base is stacked

better with the A180 base, in the 16mer Mg2+

RNA complex structure. 1H-

19F HOESY

experiments confirmed the change in chemical shift of the U179 H6 proton, which was

observed in the NOESY spectra (Figure 5.1.15). Interestingly, two more 1H-

19F NOE

peaks were observed in the 5-FU 16mer Mg2+

RNA complex, corresponding to U179 H1ʹ

and U179 H3ʹ. This could be a result of the GNRA tetraloop developing a more compact

structure as observed in the NMR solution structures.

Figure 5.1.14 376MHz 19


and (b) with

5.0eq Mg2+

, at 25°C in 2H2O. A highfield shift of 0.27ppm was observed for the U179 F5 peak in

the 5-FU 16mer RNA. 19


254


19F HOESY (250ms) spectrum of the (a) FMDV 5-FU 16mer apo-

RNA and (b) FMDV 5-FU 16mer Mg2+


19F chemical shifts were

referenced to CF3COOH.

For the 5-FU 15mer RNA, a chemical shift change of 0.05ppm was observed (Figure

5.1.16). This chemical shift change is within the detectable limit so did not correspond to a

significant change in chemical shift.

Figure 5.1.16 376MHz 19


and (b) with

5.0eq Mg2+

, at 25°C in 2H2O. A highfield shift of 0.05ppm was observed for the U230 F5 peak in

the 5-FU 15mer RNA. 19


255

5.2 19F-NMR studies of the 5-FU 16mer/15mer complex

The 19

F-NMR (2H2O) spectrum of the 5-FU 16mer/15mer Mg

2+ RNA complex revealed

some very interesting data (Figure 5.2.1). Two clear peaks could be observed at -

165.5ppm and -167.73ppm corresponding to the U179 F5 peak of the 5-FU 16mer RNA

and the U230 F5 peak of the 5-FU 15mer RNA, respectively. Three important

observations were made when comparing with the 19

F-NMR spectra of the fluorinated

Mg2+

-RNAs alone. Firstly, the U179 F5 peak had reduced in intensity and slightly

broadened compared to that of the 5-FU 16mer Mg2+

RNA alone. Secondly, two smaller

peaks were observed at -164.8ppm and -166.0ppm, flanking the U179 F5 peak. It is

possible that these peaks could represent the U179 F5 peak in two different conformations

and would explain why the U179 F5 had reduced in intensity and broadened. What is

more fascinating is that one of these peaks could represent the 5-FU 16mer Mg2+

RNA in

its bound form of the RNA-RNA complex. Thirdly, the U230 F5 peak had shifted back to

the chemical shift found in the 5-FU 15mer apo-RNA, indicating a possible RNA-RNA

interaction involving the stem region of the 15mer RNA. These findings suggest an RNA-

RNA interaction between the 16mer and 15mer Mg2+

RNAs and demonstrated how

fluorinated RNAs could be used to uniquely probe RNA-RNA interactions.

A 1H-

19F HOESY experiment of the 5-FU 16mer/15mer Mg

2+ RNA complex may have

provided additional information, specifically intermolecular 1H-

19F NOEs between the

fluorinated 16mer and 15mer RNAs. Since it was hypothesised that the 16mer RNA loop

region interacts with the 15mer RNA stem region, intermolecular NOEs would have

provided good evidence of an RNA-RNA interaction. Due to time constraints and the

unavailability of spectrometer time, the 1H-

19F NOESY experiment was not performed.

Figure 5.2.2 displays the 1H-

19F HOESY (

2H2O) spectrum of the 5-FU 16mer and 15mer

Mg2+

RNAs. These spectra would have been used as a control to assess whether any

additional peaks could be observed in the 1H-

19F HOESY spectrum of the RNA-RNA

complex. If these additional peaks had been discovered then they may have corresponded

to intermolecular NOE connectivities.

256

Figure 5.2.1 376 MHz 19

F-NMR stack plot of the (a) 5-FU 16mer Mg2+

RNA complex, (b) 5-FU

15mer Mg2+

RNA complex, (c) 5-FU RNA-RNA complex, at 25°C in 2H2O. The red asterisks

represent the additional smaller peaks observed in the 5-FU RNA-RNA complex. 19

F chemical

shifts were referenced to CFCl3.


19F HOESY (250ms) spectrum of the (a) FMDV 5-FU 16mer Mg

2+

RNA complex and (b) FMDV 5-FU 15mer Mg2+


19F chemical

shift referenced to CF3COOH. Assignments for both the 5-FU 16mer RNA (black) and 5-FU

15mer RNA (blue) are shown.

257

Chapter 6: Conclusion and Future work

6.1 Conclusion

The main aim of the project was to investigate the three dimensional structure, kinetics

and interactions of the conserved RNA motifs in the FMDV IRES. Specifically, there

were three main areas in this thesis that were of prime interest: the NMR structures of the

16mer and 15mer apo-RNAs, the effect of Mg2+

on these conserved RNAs and the RNA-

RNA interaction. In this chapter, the findings of these three main areas will be

summarised. Finally, suggestions for future work are indicated.

6.1.1 Structure of the conserved RNA motifs

The NMR structure of the 16mer and 15mer apo-RNAs provided deep insight into the

tertiary structure of these RNA motifs. Firstly, the NMR structures confirmed the

predicted secondary structure of the 16mer and 15mer RNAs. The 16mer apo-RNA NMR

structure was found to have six base pairs and four unpaired nucleotides in the tetraloop.

The 15mer apo-RNA NMR structure was found to have four base pairs with a large

heptaloop. This demonstrates that NMR structures can be used to ascertain and verify the

secondary structure of RNA motifs, which have been previously determined by other

experimental methods.

Secondly, detailed analysis of the NMR structures provided interesting features of the

tertiary structures. For the 16mer apo-RNA NMR structure, the six base pairs were found

to conform to the canonical A-form helical conformation. However, the terminal base pair

(U172-A187) was found to be frayed, caused by more exposure to the solvent. This

suggested that terminal A.U base pairing may be susceptible to destabilisation and fraying.

Additionally, the NMR data clearly showed a C2ʹ-endo sugar conformation for the U172

and A187 nucleotides. Four unpaired nucleotides were found in the GNRA tetraloop

(G178UAA181), with the G178 base stacked on the 5ʹ-end and the three succeeding bases

stacked on the 3ʹ-end. The GUAA tetraloop structure also showed that the two main

258

factors involved in stabilising the GUAA tetraloop are base stacking and intramolecular

interactions. Base stacking would have the largest contribution to the stabilisation of the

GUAA tetraloop. However, both specific and non-specific base-phosphate interactions

were observed in the GUAA tetraloop, which would play a significant role in stabilising

the GUAA tetraloop. Furthermore, the NMR data confirmed that the A180 nucleotide, in

the GUA180A tetraloop, possessed a C2ʹ-endo sugar conformation. Interestingly, the NMR

data also confirmed the existence of conformational exchange of the sugar pucker for the

loop nucleotides of the 16mer apo-RNA.

For the 15mer apo-RNA NMR structure, the four base pairs were found to conform to the

typical A-form helical conformation, despite the presence of a large heptaloop. The

terminal base pair (G229-C243) formed a standard base pairing conformation and was not

frayed like the terminal A.U base pair in the 16mer apo-RNA NMR structure. This

suggests that terminal G.C base pairs are much more stable than terminal A.U base pairs

in hairpin loops. Seven unpaired nucleotides formed the heptaloop in the 15mer apo-RNA

NMR structure. Since this was the first heptaloop RNA structure to be studied by any

structure determination method to the best of our knowledge, it was an exciting prospect

to observe the tertiary structure of the heptaloop (A233ACCCCA239). The base stacking

formation was not as simple as observed for the GUAA tetraloop in the 16mer apo-RNA

NMR structure. The first two bases (A233 and A234) stacked at an angle on the 5ʹ-end,

while the last three bases (C237, C238 and A239) were found to be stacked. Similar to the

GUAA tetraloop of the 16mer apo-RNA NMR structure, the largest contribution to

stabilisation would come from the base stacking interactions. However, a large number of

non-specific base-phosphate and sugar-phosphate interactions were also observed in the

heptaloop structure, playing a significant part in loop stability.

Overall, the 16mer and 15mer apo-RNA NMR structures have provided detailed

information about the tertiary conformation of RNA hairpin loops, which will impart a

greater understanding of RNA structure. This also presented a good opportunity to study

the role of Mg2+

and investigate the RNA-RNA interaction between the 16mer and 15mer

RNAs.

259

6.1.2 The role of Mg2+

Mg2+

plays an essential role in RNA stability, RNA folding and RNA-RNA interactions.

The 16mer RNA was used as a model to study the effect of Mg2+

on RNA structure and

stability using NMR methods. The results obtained provided new knowledge about the

role of Mg2+

. There are two main effects of Mg2+

that will be discussed. The first describes

the effect of Mg2+

on RNA structure and the second will describe the effect of Mg2+

on

RNA stability.

It was hypothesised that Mg2+

can interact with 16mer RNA, thereby changing the tertiary

conformation in order to increase the stability of the entire 16mer RNA. 1H-NMR

experiments immediately confirmed that Mg2+

was having an effect on the 16mer

tetraloop, as large chemical shift changes were observed for the loop G178 and G177

imino protons. Significant chemical shift changes to the exchangeable and non-

exchangeable protons in the 16mer stem and loop regions, strongly suggested a Mg2+

-

induced change in tertiary conformation. This was supported by analogous chemical shift

changes observed in the 15mer RNA. 31

P-NMR experiments were able to establish that

Mg2+

induced changes to the phosphate backbone of the entire length of the 16mer RNA,

specifically to the tetraloop region. A lowfield chemical shift change found for the U179

phosphorus indicated possible specific interaction of chelated Mg2+

ions in proximity to

the G178 and U179 nucleotides. 19

F-NMR experiments revealed a significant 19

F chemical

shift change to U179 F5, which further supported the effect of Mg2+

on the tetraloop.

For the first time, the NMR structure of a hairpin loop was generated, with and without

Mg2+

. This provided a novel understanding into the Mg2+

-induced structural changes and

its link to RNA stability. The NMR structures revealed enhanced stability for the terminal

base pair (U172-A187), since the base pair was not frayed in the 16mer Mg2+

RNA

complex and the U172 base was able to stack with the adjacent C173 base. This structural

change was accompanied by a change in sugar conformation for U172 from C2ʹ-endo to

C3ʹ-endo. The most significant structural changes were found for the 16mer GUAA

tetraloop. The tetraloop was altered by the presence of Mg2+

to become more compact,

260

allowing for better base stacking and stronger intramolecular interactions. These changes

would directly lead to a significant enhancement in loop stability, contributing to the

stability of the whole 16mer RNA structure.

Evidence of Mg2+

increasing thermodynamic stability of the 16mer RNA was clearly

apparent in the VT series and imino proton exchange experiments. The G186 and G177

imino proton peaks were clearly exchange retarded in the presence of Mg2+

at 35°C. In

addition, Mg2+

was able to significantly stabilise the base pairing at a higher temperature

of 45°C. Imino proton exchange rates revealed that the stem base pairing was very stable

up to 15°C, due to low exchange rates of the stem imino protons. Conversely, the loop

G178 imino exchange rate was higher compared to the stem imino protons due to

increased exposure to the solvent. From the imino proton exchange experiments, it was

discovered that Mg2+

was able to significantly increase the base pair stability of A.U base

pairs, which would have the consequence of stabilising the stem region even further.

Mg2+

has long been known to increase stability of RNA structure, but the mechanism by

which Mg2+

is able to induce structural and dynamical changes to RNA structure has still

eluded researchers. This investigation has provided a greater understanding for the role of

Mg2+

with the conclusion that Mg2+

is able to alter the RNA structure to improve stability

through maximising base stacking and intramolecular interactions.

6.1.3 RNA-RNA interaction

It was hypothesised, by E. Martínez-Salas and co-workers10

that the FMDV 16mer and

15mer RNA motifs interact with each other, forming long-range tertiary contacts.

Although it was not possible to study the interaction between these two RNAs as part of

the whole native hammerhead region, it presented an important opportunity to investigate

the interaction between two small hairpin loop structures.

The most significant observation that was made in the RNA-RNA complex is that the loop

G178 imino proton peak of the 16mer RNA was considerably reduced in intensity caused

261

by faster exchange with water; confirmed by the calculated imino proton exchange rates.

This suggested the possibility that the 16mer tetraloop was involved in forming RNA-

RNA interactions. Changes in linewidth and intensity of peaks in the 1H-NMR spectra and

chemical shift changes in the NOESY spectra provided further evidence of RNA-RNA

interactions. In addition, a lower number of chemical shift changes were observed at a

higher temperature, indicating a temperature-dependent interaction between the two RNAs.

19F-NMR experiments revealed two extra fluorine peaks for U179 F5 of the 5-FU 16mer

RNA indicating a possible equilibrium between two different RNA-RNA interactions.

These results indicated that there is a weak interaction between the two RNAs. However,

it is entirely possible that this interaction is much stronger when the two RNAs are part of

the whole apical region of domain 3 in the FMDV IRES. Therefore, further work must be

performed to understand the critical role of the 16mer GNRA tetraloop in IRES function.

6.1.4 Errors and their implications

The results described in the thesis were all subject to experimental, systematic and random

errors. Experimental errors were mainly related to errors in the measurement of chemical

shifts and the assignment of resonances in NMR spectra. These were very few and

minimised further by carefully checking the chemical shift data and assignments to make

sure that they were all correct. Examples of systematic errors can include instrumental

error, changes in environment during the running of long NMR experiments such as

variation in temperature and imperfect calibration of NMR data. Since these errors are

inherent to any data acquisition they were difficult to minimise by any later action.

Random errors can be related to the measurement of chemical shifts, coupling constants,

linewidth, NOE intensities, T1 and Kex from NMR data. These errors were minimised by

either setting a detectable limit for changes in values or by making repeated measurements.

The errors mentioned above were not significant enough to affect either the results or their

interpretation. Therefore, the conclusions drawn from the results are reliable and valid.

262

6.2 Future work

The purpose of studying the FMDV IRES was to use it as a model to gain better

understanding in the area of RNA structural biology and also to unravel the mechanism of

translation initiation. The studies conducted in this thesis have provided valuable insight

into the structure, kinetics and interactions of RNA and are the first steps to advancing our

knowledge on IRES translation initiation. The relationship between the IRES structure and

its function is still unknown at this time. To provide a greater understanding of the

complex nature of the conserved RNA motifs in the FMDV IRES further experiments

must be performed in order to achieve this outcome.

6.2.1 Binding of Mg2+

ions to RNA

NMR spectroscopy has been used to investigate the effect of Mg2+

binding with RNA,

whereby chemical shift mapping and imino proton exchange experiments were utilised.

However, other experimental methods must be imposed to provide direct evidence for the

interaction between Mg2+

and RNA structure. Two important techniques used in NMR for

the investigation of Mg2+

interaction are paramagnetic line broadening and the use of

cobalt hexamine (Co(NH3)63+

).115

116

Line broadening experiments use paramagnetic ions

such as Mn2+

to substitute Mg2+

. The proximity of Mn2+

ions to RNA structure induces

line broadening due to faster relaxation of specific proton peaks, which represent nuclei

within 5Å from Mn2+

. These peaks can be identified and indicate the possible site of Mg2+

binding. Cobalt hexamine (Co(NH3)63+

) is an analogue of Mg2+

that is used in NMR

spectroscopy to mimic Mg2+

ions. The advantage of using cobalt hexamine is that it

provides direct NOE contacts between the NH3 protons of (Co(NH3)63+

) and RNA protons.

2J-[

1H,

15N]-HSQC NMR experiments can also be used to probe the direct coordination of

Mg2+

to N7 of purines.117

Changes in chemical shift to N7 and H8 nuclei can both be

monitored upon addition of Mg2+

, which may give an indication of the binding mode of

Mg2+

.

263

6.2.2 Isotopically labelled RNA

Uniformly, 13

C- and 15

N-isotopically labelled RNA samples would provide a great

opportunity to perform a wide variety of NMR experiments. Not only will this

substantially aid in more reliable assignments for structure determination, but will also

provide new prospects of studying RNA dynamics and interactions. A new strategy can

then be employed to study the larger conserved RNA motifs in domain 3 of the FMDV

IRES. The most promising would be the 36mer RNA found in the apical region of domain

3, which constitutes the 16mer RNA. The knowledge gained could bring us closer to

understanding the relationship between RNA structure/dynamics and function.

Various 2D NMR experiments can be performed for the identification and assignment of

protons in 13

C/15

N-labelled RNAs. These experiments include the 2D 1H-

15N HSQC

118,

the 2D HCNCH119

and the HNN-COSY120

. The combination of uniformly labelled RNA

and 3D NMR experiments can provide a means of unambiguous assignment and

significantly reduce the problem of overlapping peaks. These experiments include the 3D

1H-

13C HSQC-NOESY

121, the 3D HCCH-TOCSY

122 and the 3D HCP

123.

In order to study larger RNAs by NMR spectroscopy, such as the 79mer RNA in domain 3

of the FMDV IRES, different labelling strategies will have to be employed. This is

because there are two major problems that come with larger RNAs, spectral overlap and

increased linewidth.124

Signal overlapping problems become increasingly substantial with

larger RNAs as there are a greater number of protons in the same given spectra width.

Furthermore, the slower tumbling of larger RNAs results in more efficient T2 relaxation

and increasing linewidths. Therefore, to overcome these limitations, alternate labelling

strategies must be sought, mainly involving selective and segmental labelling of RNA.125

An example of selective labelling involves the use of deuterium (2H) isotopes, which can

produce >90% enriched RNA samples. The great advantage of this labelling strategy is

that it solves the problem of spectral overlap and increases sensitivity due to reduced

linewidths. Segmental labelling involves ligating an isotopically labelled RNA to a larger

unlabelled RNA. This method may be applied to the 79mer RNA, whereby the labelled

264

36mer RNA is ligated with unlabelled RNA. It is believed that the 79mer RNA produces a

stable four-way junction, which has yet to be studied by NMR, and will be crucial to

understanding its role in the IRES mechanism. Subsequently, 2D and 3D filtered/edited

experiments can then be employed to remove the signals from protons attached to labelled

nuclei while retaining signals from protons attached to unlabelled nuclei, and vice

versa.126

This would facilitate the NMR assignment and structure determination of the

labelled and unlabelled regions of the 79mer RNA.

6.2.3 RNA tertiary contacts

IRES activity depends upon the formation of tertiary RNA contacts within domain 3 of the

FMDV IRES. With the advances in labelling strategies and NMR techniques, it is possible

to study RNA-RNA interactions within large RNAs instead of studying two separate RNA

molecules. Therefore, in order to investigate the possible RNA-RNA interaction between

the GUAA tetraloop of the 16mer RNA with the 15mer RNA, one would need an RNA

that would encompass both the 16mer and 15mer RNA motifs. This would have to involve

segmental labelling, whereby the 16mer and 15mer RNAs are isotopically labelled as part

of the larger RNA. Filtered-edited NMR experiments are extremely useful in this case as

long-range tertiary interactions can be identified.

265

Papers to be published

The results of the project described in chapters 3 to 5 will be published in peer reviewed

journals and they are listed below.

1. Usman Rasul, Vasudevan Ramesh (2012) NMR studies of the structure, kinetics

and interactions of the conserved RNA motifs in the FMDV IRES. RNA, (To be

submitted).

2. Usman Rasul, Vasudevan Ramesh (2012) Elucidation of RNA-RNA interactions

in the FMDV IRES using NMR spectroscopy. Nucleic Acids Res., (To be

submitted).

266

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277

Appendices

Appendix I: NMRPipe script

csh

if (-e /opt/NMRPipe/com/nmrInit.linux9.com) then

source /opt/NMRPipe/com/nmrInit.linux9.com

endif

if (-e /opt/NMRPipe/dynamo/com/dynInit.com) then

source /opt/NMRPipe/dynamo/com/dynInit.com

endif

#!/bin/csh

bruk2pipe -in /home/chemistry/Desktop/Usman/URdata_22_09_11/3/ser \

-bad 0.0 -noaswap -DMX -decim 12 -dspfvs 12 -grpdly -1 \

-xN 4096 -yN 984 \

-xT 2048 -yT 492 \

-xMODE DQD -yMODE States-TPPI \

-xSW 13227.513 -ySW 12019.231 \

-xOBS 599.927 -yOBS 599.927 \

-xCAR 5.029 -yCAR 5.029 \

-xLAB 1Hx -yLAB 1H \

-ndim 2 -aq2D States \

-out /home/chemistry/Desktop/Usman/URdata_22_09_11/3/test.fid -verb -

ov

sleep 5

nmrPipe -verb -in

/home/chemistry/Desktop/Usman/URdata_22_09_11/3/test.fid \

| nmrPipe -fn SOL -fl 32 \

| nmrPipe -fn GMB -lb -3.0 -gb 0.05 -c 0.5 \

| nmrPipe -fn ZF -auto \

| nmrPipe -fn FT -auto \

| nmrPipe -fn PS -p0 155.0 -p1 -91.0 -di \

| nmrPipe -fn POLY -auto -ord 1 \

| nmrPipe -fn TP \

| nmrPipe -fn SP -off 0.5 -end 0.98 -c 0.5 \

| nmrPipe -fn ZF -auto \

| nmrPipe -fn FT -auto \

| nmrPipe -fn PS -p0 -95.0 -p1 176.0 -di \

| nmrPipe -fn POLY -auto -ord 1 \

| nmrPipe -fn TP \

| nmrPipe -ov -verb -out

/home/chemistry/Desktop/Usman/URdata_22_09_11/3/15merMg2+_H2O_150ms.ft2

/opt/sparky/bin/./pipe2ucsf

/home/chemistry/Desktop/Usman/URdata_22_09_11/3/15merMg2+_H2O_150ms.ft2

/home/chemistry/Desktop/Usman/URdata_22_09_11/3/15merMg2+_H2O_150ms.ucsf

278

Appendix II: XPLOR scripts

Simulated Annealing Script:

from pdbTool import PDBTool

from xplorPot import XplorPot

from rdcPotTools import create_RDCPot

from varTensorTools import create_VarTensor

import varTensorTools

from ivm import IVM

from potList import PotList

import protocol

from protocol import initMinimize

from ivm import IVM

from xplor import command

import random

from atomAction import SetProperty

from simulationTools import StructureLoop

from vec3 import Vec3

from psfGen import seqToPSF




from ivm import IVM


import protocol

from avePot import AvePot

from simulationTools import MultRamp, StaticRamp, InitialParams, StructureLoop,

AnnealIVM, FinalParams

from simulationTools import AnnealIVM

from monteCarlo import randomizeTorsions

from noePotTools import create_NOEPot

xplor.parseArguments()

# this checks for typos on the command-line. User-customized arguments can

# also be specified

command = xplor.command

protocol.initParams("nucleic")

protocol.initTopology("nucleic")

# parameters to ramp up during the simulated annealing protocol

#

rampedParams=[]

highTempParams=[]

init_t = 3500. # Need high temp and slow annealing to converge

final_t=25

bathTemp=2000

seqToPSF(open('16mer.seq').read(), seqType='rna')

#seqToPSF(open('15mer.seq').read(), seqType='rna', startResid=21)

279

command("write psf output=FMDV16mer.psf end")

for atom in AtomSel("all"):

atom.setPos( Vec3(float(atom.index())/10,

random.uniform(-0.5,0.5),

random.uniform(-0.5,0.5)) )

pass

protocol.fixupCovalentGeom(useVDW=1,maxIters=100)

pots = PotList()

noex = create_NOEPot("noex",

"FMDV16Mgex.tbl")

noex.setPotType("soft") #if incorrect noes suspected set soft if not set hard

rampedParams.append( MultRamp(0.2,30.,"noex.setScale( VALUE )") )

noen = create_NOEPot("noen",

"FMDV16MgL9b.tbl")

noen.setPotType("soft") #if incorrect noes suspected set soft if not set hard

rampedParams.append( MultRamp(0.2,30.,"noen.setScale( VALUE )") )

hbon = create_NOEPot("hbon",

"hbon16.tbl")

hbon.setPotType("hard")

rampedParams.append( MultRamp(0.2,30.,"hbon.setScale( VALUE )") )

hbs = create_NOEPot("hbs",

"hbonsoft16.tbl")

hbs.setPotType("soft")

rampedParams.append( MultRamp(0.2,30.,"hbs.setScale( VALUE )") )

protocol.initDihedrals("tor16.tbl",

scale=5, #initial force constant

useDefaults=0)

highTempParams.append( StaticRamp("pots['CDIH'].setScale(10)") )

rampedParams.append( StaticRamp("pots['CDIH'].setScale(200)") )

# set custom values of threshold values for violation calculation

#

pots.add( XplorPot('CDIH') )

pots['CDIH'].setThreshold( 5 )

xplor.command("@plane16.tbl")

## radius of gyration term

##

#protocol.initCollapse(Rtarget=10.16)

#pots.append( XplorPot('COLL') )

pots.add( XplorPot("BOND") )

pots.add( XplorPot("DIHE") )

pots.add( XplorPot("ANGL") )

pots.add( XplorPot("IMPR") )

rampedParams.append( MultRamp(0.4,1.0,"pots['ANGL'].setScale(VALUE)"))

rampedParams.append( MultRamp(0.1,1.0,"pots['IMPR'].setScale(VALUE)"))

pots.add( XplorPot("VDW") )

rampedParams.append( StaticRamp("protocol.initNBond(nbxmod=5)") )

rampedParams.append( MultRamp(0.95,0.85,

"xplor.command('param nbonds repel VALUE end

end')") )

280

rampedParams.append( MultRamp(.004,4,

"xplor.command('param nbonds rcon VALUE end

end')") )

pots.add(noex)

pots.add(noen)

pots.add(hbon)

pots.add(hbs)

#pots.append(AvePot(XplorPot("plan",xplor.simulation)) )

# IVM setup

# the IVM is used for performing dynamics and minimization in torsion-angle

# space, and in Cartesian space.

#

from selectTools import IVM_groupRigidSidechain

from selectTools import IVM_breakRiboses

dyn = IVM()

protocol.initDynamics(dyn,potList=pots)

#IVM_groupRigidSidechain(dyn)

#IVM_breakRiboses(dyn, sel=0, breakSelStr="name O4' or name C1'")

protocol.torsionTopology(dyn)

minc = IVM()

protocol.initMinimize(minc,potList=pots)

IVM_groupRigidSidechain(minc)

#IVM_breakRiboses(minc, sel=0, breakSelStr="name O4' or name C1'")

protocol.cartesianTopology(minc,"not resname ANI")

# object which performs simulated annealing

#


cool = AnnealIVM(initTemp =init_t,

finalTemp=final_t,

tempStep =12.5,

ivm=dyn,

rampedParams = rampedParams)

#cart_cool is for optional cartesian-space cooling

cart_cool = AnnealIVM(initTemp =init_t,

finalTemp=25,

tempStep =12.5,

ivm=minc,


def calcOneStructure( structData ):

randomizeTorsions(dyn)

# initialize parameters for high temp dynamics.

InitialParams( rampedParams )

# high-temp dynamics setup - only need to specify parameters which

# differfrom initial values in rampedParams

InitialParams( highTempParams )

# high temperature bit - using only P-P nonbonded terms

highTempParams.append( StaticRamp("""protocol.initNBond(repel=1.2,

cutnb=100,

tolerance=45,

selStr="name P")""") )

281

protocol.initDynamics(dyn,

potList=pots, # potential terms to use

bathTemp=init_t,

initVelocities=1,

finalTime=800, # stops at 800ps or 8000 steps

numSteps=8000, # whichever comes first

printInterval=100)

dyn.setETolerance( init_t/100 ) #used to det. stepsize. default: t/1000

dyn.run()

protocol.initNBond() #reset to include all atoms

# initialize parameters for cooling loop


# initialize integrator for simulated annealing

#


potList=pots,

numSteps=100, #at each temp: 100 steps or

finalTime=.2 , # .2ps, whichever is less

printInterval=100)

# perform simulated annealing

#

cool.run()

# final torsion angle minimization

#

protocol.initMinimize(dyn,

printInterval=50)

dyn.run()

protocol.initDynamics(minc,

potList=pots,

numSteps=100, #at each temp: 100 steps or

finalTime=.4 , # .2ps, whichever is less

printInterval=100)

cart_cool.run()

# final all- atom minimization

#

protocol.initMinimize(minc,

potList=pots,

dEPred=10)

minc.run()

structData.writeStructure(pots)

simWorld.setRandomSeed( 785 )

outPDBFilename = 'SCRIPT_STRUCTURE.pdb'

StructureLoop(numStructures=200,

pdbTemplate=outPDBFilename,

structLoopAction=calcOneStructure,

genViolationStats=1,

averageTopFraction=0.3, #report stats on best 30% of structs

averageContext=FinalParams(rampedParams),

averageSortPots=[pots['BOND'],pots['ANGL'],pots['IMPR'],

noex,noen,pots['CDIH'],hbon,hbs],

averageFilename="SCRIPT_ave.pdb", #generate regularized ave

structure

averageFitSel="name P",

averagePotList=pots).run()

282

Refinement Script:

from pdbTool import PDBTool


from rdcPotTools import create_RDCPot



from ivm import IVM


import protocol

from protocol import initMinimize

from ivm import IVM

from xplor import command

import random

from atomAction import SetProperty

from simulationTools import StructureLoop

from vec3 import Vec3

from psfGen import seqToPSF


from ivm import IVM


import protocol

from avePot import AvePot

from simulationTools import MultRamp, StaticRamp, InitialParams, StructureLoop,

AnnealIVM


xplor.parseArguments()

# this checks for typos on the command-line. User-customized arguments can

# also be specified.

#

command = xplor.command

from noePotTools import create_NOEPot

protocol.initParams("nucleic")

protocol.initTopology("nucleic")

seed=10

numberOfStructures=200

startStructure=0

outFilename = "SCRIPT_STRUCTURE.pdb"

rampedParams=[]

highTempParams=[]

init_t=2000

final_t=25

bathTemp=2000

startFile="annealFMDV16meru_85.pdb"

simWorld.setRandomSeed(seed)

seqToPSF(open('16mer.seq').read(), seqType='rna')

#seqToPSF(open('15mer.seq').read(), seqType='rna', startResid=21)

#command("write psf output=29mer.psf end")

#

# starting coords

283

#

protocol.initCoords(startFile)

protocol.covalentMinimize()

# list of potential terms used in refinement

pots = PotList()

crossTerms=PotList('cross terms') # can add some pot terms which are not

# refined against- but included in analysis

noex = create_NOEPot("noex",

"FMDV16Mgex.tbl")

noex.setPotType("soft") #if incorrect noes suspected

rampedParams.append( MultRamp(0.2,30.,"noex.setScale( VALUE )") )

noen = create_NOEPot("noen",

"FMDV16MgL9b.tbl")

noen.setPotType("soft") #if incorrect noes suspected

rampedParams.append( MultRamp(0.2,30.,"noen.setScale( VALUE )") )

hbon = create_NOEPot("hbon",

"hbon16.tbl")

hbon.setPotType("hard")

hbon.setScale(1000)

rampedParams.append( MultRamp(0.2,30.,"hbon.setScale( VALUE )") )

hbs = create_NOEPot("hbs",

"hbonsoft16.tbl")

hbs.setPotType("soft")

hbon.setScale(1000)

rampedParams.append( MultRamp(0.2,30.,"hbs.setScale( VALUE )") )

protocol.initDihedrals("tor16.tbl",

scale=5) #initial force constant

pots.append(AvePot(XplorPot,"cdih") )

rampedParams.append( StaticRamp("pots['CDIH'].setScale(200)") )

protocol.initRamaDatabase('nucleic')

pots.append(AvePot(XplorPot,"rama") )

rampedParams.append( MultRamp(1,1,"xplor.command('rama scale VALUE end')"))

xplor.command("@rna_orient1.setup")

pots.append(AvePot(XplorPot,"orie") )

rampedParams.append( StaticRamp("pots['ORIE'].setScale(0.2)") )

rampedParams.append( MultRamp(0.002,0.3,"xplor.command('orie scale VALUE end')"))

xplor.command("@plane16.tbl")

pots.add( XplorPot("BOND") )

pots.add( XplorPot("DIHE") )

pots.add( XplorPot("ANGL") )

pots.add( XplorPot("IMPR") )

rampedParams.append( MultRamp(0.4,1.0,"pots['ANGL'].setScale(VALUE)"))

rampedParams.append( MultRamp(0.1,1.0,"pots['IMPR'].setScale(VALUE)"))

protocol.initNBond(cutnb=4.5)

pots.add( XplorPot("VDW") )

rampedParams.append( StaticRamp("protocol.initNBond(nbxmod=5)") )

rampedParams.append( MultRamp(0.95,0.85,

"xplor.command('param nbonds repel VALUE end

end')") )

rampedParams.append( MultRamp(.004,4,

284

"xplor.command('param nbonds rcon VALUE end

end')") )

pots.add(noen)

pots.add(noex)

pots.add(hbon)

pots.add(hbs)

pots.append(AvePot(XplorPot("plan",xplor.simulation)) )

mini = IVM() #initial alignment of orientation tensor axes


from selectTools import IVM_breakRiboses

IVM_groupRigidSidechain(mini)

#IVM_breakRiboses(mini, sel=0, breakSelStr="name O4' or name C1'")

protocol.cartesianTopology(mini,"not resname ANI")

protocol.initMinimize(mini,

numSteps=20)

mini.fix("not resname ANI")

mini.run() #this initial minimization is not strictly necessary

#uncomment to allow Da, Rh to vary

#for medium in ('bic1','phg1'): media[medium].setFreedom("varyDa, varyRh")

#for medium in ('bic2',):

# media[medium].setFreedom("varyDa, varyRh, fixAxisTo bic1")

#for medium in ('phg2','phg3',):

# media[medium].setFreedom("varyDa, fixAxisTo phg1, fixRhTo phg1")

dyn = IVM()

protocol.initDynamics(dyn,potList=pots)

IVM_groupRigidSidechain(dyn)

#IVM_breakRiboses(dyn, sel=0, breakSelStr="name O4' or name C1'")

#protocol.cartesianTopology(dyn,"not resname ANI")

protocol.torsionTopology(dyn)


minc = IVM()

protocol.initMinimize(minc,potList=pots)

IVM_groupRigidSidechain(minc)

#IVM_breakRiboses(minc, sel=0, breakSelStr="name O4' or name C1'")

protocol.cartesianTopology(minc,"not resname ANI")

anneal= AnnealIVM(initTemp =init_t,

finalTemp=25,

tempStep =25,

ivm=dyn,


# initialize parameters for initial minimization.


# initial minimization

protocol.initMinimize(dyn,

numSteps=1000)

dyn.run()

from simulationTools import testGradient

#testGradient(potList,eachTerm=1)

def calcOneStructure( structData ):

285

# initialize parameters for high temp dynamics.


# high temperature bit - using only P-P nonbonded terms

highTempParams.append( StaticRamp("""protocol.initNBond(repel=1.2,

cutnb=100,

tolerance=45,

selStr="name P")""") )


initVelocities=1,

bathTemp=init_t,

potList=pots,

finalTime=10)

dyn.run()

protocol.initNBond() #reset to include all atoms

# perform simulated annealing

#


finalTime=0.2, #time to integrate at a given temp.

numSteps=0, # take as many steps as necessary

eTol_minimum=0.001 # cutoff for auto-TS det.

)

anneal.run()

#

# torsion angle minimization

#

protocol.initMinimize(dyn)

dyn.run()

##

##all atom minimization

##

minc.run()

structData.writeStructure(pots,crossTerms)

from simulationTools import StructureLoop, FinalParams

StructureLoop(numStructures=numberOfStructures,

startStructure=startStructure,

structLoopAction=calcOneStructure,

pdbTemplate=outFilename,

genViolationStats=1,

averageTopFraction=0.3,

averagePotList=pots,

averageSortPots=[pots['BOND'],pots['ANGL'],pots['IMPR'],

noex,noen,pots['CDIH'],hbon,hbs],

#averageAccept=accept, #only use structures which pass accept()

averageContext=FinalParams(rampedParams),

averageFilename="SCRIPT_ave.pdb", #generate regularized ave

structure

averageFitSel="name P",

averageCompSel="not resname ANI and not name H*" ).run()

NMR studies of the structure, kinetics and interactions of the ...

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