Bifacial Peptide Nucleic Acid (Bpna) As a Regulator of Nucleic Acid Function

Home , Aptamer, Coding strand, Peptide nucleic acid, RNTP

Bifacial Peptide Nucleic Acid (bPNA) as a Regulator of Nucleic Acid Function

Dissertation

Presented in Partial Fulfillment of the Requirement for the Degree Doctor of Philosophy

in the Graduate School of the Ohio State University

Xin Xia, B.S.

Graduate Program in Chemistry

The Ohio State University

2015

Dissertation Committee:

Dr. Dennis Bong, Advisor

Dr. Karin Musier-Forsyth

Dr. Kotaro Nakanishi

Xin Xia

2015

Abstract

This dissertation contains research summaries regarding the characterizations and applications of a novel bifacial peptide nucleic acid (bPNA) triplex system. The main content is divided into two parts: part one (chapter 2) presents systematic studies on effective in vitro inhibition of transcription, reverse-transcription and exonuclease function via the formation of synthetic bPNA-nucleic acid triplex structures; part two (chapters 3 and 4) demonstrates that bPNA triplex hybrid functionally substitutes for native duplex structures that are crucial for proper functions of aptamers and ribozyme, further a two- way communication system describing RNA-templated oxidative coupling of bPNA fragments leads to the emergence of ribozyme cleavage is discussed in details.

In part one, we speculate that the thermodynamically stable synthetic bPNA-nucleic acid triplex can be utilized to generate template distortions that are inhibitory to nucleic acid based enzymatic reactions. Three enzymatic systems were investigated: T7 RNA polymerase, Exonuclease T, and AMV reverse transcriptase. bPNA hybridization kinetics and inhibitory efficacies on each system will be discussed in detail in chapter 2.

In part two, study on bPNA-nucleic acid triplex system was expanded to investigate the structure-function relation of nucleic acid. In chapter 3, three biologically active nucleic acids folds were selected: IgE DNA aptamer, spinach RNA aptamer and minimal type I hammerhead ribozyme. Replacement of a duplex stem with unstructured oligo-T/U strands, which are bPNA binding sites, imposed structure-function loss in all three nucleic

ii acids folds. Functional rescue was observed upon bPNA-driven refolding of oligo-T/U strands into triplex hybrid system. Further, in chapter 4, we demonstrated a 2-way communication between the abiotic bPNA hybridization site and the native ribozyme cleavage site, where the ribozyme-templated bPNA ligation in turn restore the ribozyme self-cleavage activity.

In summation, we will demonstrate to you that bPNA triplex stem structure is compatible with biological processes and presented to be competitive inhibitor for DNA/RNA specific enzymes; further, bPNA triplex stem is biologically similar to native stem structures, not only the bPNA hybridization but also the nucleic acid templated bPNA ligation can restore native nucleic acid activities, demonstrate readout and transformation of non-native macromolecules through an abiotic template interface in DNA/RNA template topologies that are not accessible via native base-pairing.

iii

To my parents and my fiancé

Acknowledgement

I hold great appreciation to Dr. Bong, as he is an intelligent, patient and responsible mentor to me. He took me to into his lab on my third year, and since then he has been constantly helping me to not only build up knowledge, but also confidence. His persistence in research originality is highly admirable, and it has inspired me and will keep on inspiring me far beyond the graduate school.

I also want to express my appreciation to Dr. Karin Musier-Forsyth and Dr. Kotaro

Nakanishi for serving on my graduation committee, providing guidance and constructive input.

Thank Dr. Thomas Magliery for his generosity in sharing his lab equipment constantly throughout my graduate study. Also, thank Dr. Kurt Fredrick for a productive collaboration.

Finally, I want to thank my father Ming Xia and mother Ningning Zhang for their endless support and unconditional love throughout these years; thank my fiancé Zhun Zhou for holding my hands during all the ups and downs, studying and researching together; last but not the least, thank to my two fluffy friends Pangpang and Niuniu for necessary distractions.

Vita

2001-2004 ...... Jinling High School, Nanjing, China

2004-2008 ...... B.S., China Pharmaceutical University, Nanjing, China

2008-2009 ...... Q.A., Simcere Pharmaceutical Group, Nanjing, China

2009-2015 ...... Graduate Research/Teaching Assistant, The Ohio State University

Publications during Ph.D. program at the Ohio State University

 Zhou, Z., Xia, X. and Bong, D.* (2015) “Synthetic Polymer Hybridization with DNA and RNA Directs Nanoparticle Loading, Silencing Delivery, and Aptamer Function.” J. Am. Chem. Soc., DOI: 10.1021/jacs.5b05481

 Piao, X., Xia, X., Mao, J. and Bong, D.* (2015) “Peptide ligation and RNA cleavage via an abiotic interface.” J. Am. Chem. Soc., 137, 3751-3754.

 Xia, X., Piao, X., and Bong, D.* (2014) "Bifacial PNA as an allosteric switch for aptamer and ribozyme function." J. Am. Chem. Soc., 136, 7265-7268.

 Xia, X., Piao, X., Fredrick, K, and Bong, D., (2014) “Bifacial PNA Complexation Inhibits Enzymatic Access to DNA and RNA.” ChemBioChem. 15, 31-36.

 Piao, X., Xia, X., and Bong, D.* (2013) “Bifacial Peptide Nucleic Acid Directs Cooperative Folding and Assembly of Binary, Ternary, and Quaternary DNA Complexes.” Biochemistry, 52, 6313-6323.

 Saibal Bandyopadhyay, Xin Xia, Andre Matseiyeu, Georgeta Mihai, Sanjay Rajagopalan and Dennis Bong.* (2012) “Z-Group Ketone Chain Transfer Agents for RAFT Polymer Nanoparticle Modification via Hydrazone Conjugation.” Macromolecules, 45, 6766-6773.

Field of Study

Major field: Chemistry vi

Table of Contents

Abstract ...... ii

Acknowledgement ...... v

Vita ...... vi

Table of Contents ...... vii

List of Figures ...... xvii

List of Tables ...... xxi

CHAPTER 1

Nucleic Acids Structures and Targeting Strategies ...... 1

1.1 Native Nucleic Acids Structures ...... 2

1.1.1 Duplexes ...... 2

1.1.2 G-Quadruplex ...... 4

1.1.3 RNA Folds ...... 6

1.2 Nucleic Acids Triplex ...... 7

1.2.1 Structural Characters of Triplex ...... 7

1.3.2 Nucleobase Modifications ...... 10

1.3.3 Sugar Modifications ...... 12

1.3.4 Backbone Modifications ...... 13

1.4 Peptide Nucleic Acid (PNA) as DNA Mimic ...... 14

1.4.1 PNA Structure and Recognition Mechanism ...... 14

vii

1.4.2 Antigene Therapeutics of PNA ...... 17

1.4.3 Antisense Regulation of PNA ...... 20

1.4.4 PNAs as Detection Probes and Biosensors ...... 21

1.4.5 Chemical Modifications of PNA ...... 21

1.5 Bifacial Nucleobase Analogues ...... 23

1.5.1 Bifacial Nucleobase Displaying PNA for Triplex Formation ...... 24

1.5.2 Melamine Derivatives Recognize T-T/U-U Mismatch ...... 25

1.6 PNA With Native Peptide Backbone ...... 26

1.7 Melamine Displayed Bifacial Peptide Nucleic Acid (bPNA) ...... 27

1.8 References for Chapter 1 ...... 31

CHAPTER 2 bPNA Triplex Inhibits Enzymatic Access to DNA and RNA ...... 46

2.1 Introduction ...... 47

2.2 Results and Discussion ...... 50

2.2.1 Triplexes Inhibit in vitro T7 RNA Transcription...... 52

2.2.2 Triplexes are Resistant Towards Exonuclease T Digestion ...... 56

2.2.3 Triplexes Inhibit AMV Reverse Transcription ...... 59

2.3 Conclusion ...... 62

2.4 Experimental ...... 62

2.4.1 Material ...... 62

2.4.2 Nucleic Acid Sequences ...... 64

2.4.3 Equipment ...... 65

2.4.4 Procedures ...... 66

2.4.4.1 Amino Acid Derivative Synthesis ...... 66

viii

2.4.4.2 Solid Phase Peptide Synthesis and Characterization ...... 66

2.4.4.3 bPNA Binding Analysis ...... 66

2.4.4.4 Transcriptional Repression ...... 67

2.4.4.5 T7 Run-off Transcription Control ...... 67

2.4.4.6 Exonuclease T Resistance ...... 68

2.4.4.7 Exonuclease T digestion control ...... 68

2.4.4.8 Inhibition of reverse transcription ...... 68

2.4.4.9 Reverse transcription control ...... 69

2.5 Additional Data ...... 70

2.5.1 Amino Acid Derivative ...... 70

2.7.2 HPLC and MALDI-TOF Mass Spectra of the Synthetic Peptides ...... 72

2.6 Acknowledgements ...... 74

2.7 References for Chapter 2 ...... 75

CHAPTER 3 bPNA as Allosteric Switch for Aptamer and Ribozyme Function ...... 81

3.1 Overview...... 82

3.2 Introduction ...... 83

3.3 Results and Discussion ...... 85

3.3.1 Rescue of Aptamer-IgE Binding via Triplex Formation ...... 85

3.3.2 Rescue RNA Aptamer-Small Molecule Binding and Fluorogenics...... 88

3.3.3 Rescue Hammerhead Ribozyme Cleavage...... 92

3.3 Conclusion ...... 99

3.4 Experimental ...... 100

3.4.1 Material ...... 100

3.4.2 DNA Sequences ...... 100

3.4.3 Experimental Procedures ...... 102

3.4.3.1 Synthesis of DFHBI ...... 102

3.4.3.2 Radiolabeling of oligonucleotides ...... 102

3.4.3.3 RNA Transcription and Purification...... 103

3.4.3.4 Electrophoretic Mobility Shift Assays (EMSA) ...... 103

3.4.3.5 IgE-Aptamer Filter Binding Assays ...... 104

3.4.3.6 Fluorescence Activation Assay ...... 104

3.4.3.7 Apparent Kd Measurement for Fluorescence Activation ...... 105

3.4.3.8 U-Ribozyme Rescue Experiment ...... 105

3.4.3.9 In situ Splicing ...... 106

3.5 Equation Table ...... 107

3.6 Additional Data ...... 108

3.6.1 DFHBI Synthesis...... 108

3.6.2 Fluorescence Activation of tRNALysU-Spinach ...... 110

3.6.3 Mg2+ Dependent Cleavage of U-3 ...... 111

3.6.4 Mg2+ Dependent Cleavage of U-(2,3) ...... 113

3.7 References for Chapter 3 ...... 115

CHAPTER 4 bPNA Ligation and Ribozyme Cleavage via An Abiotic Interface ...... 121

4.1 Overview...... 122

4.2 Introduction ...... 122

4.1.2 Structure Defines Function ...... 122

4.2.2 Abiotic Template Interface for bPNA Ligations ...... 123

4.3 Results and Discussion ...... 124

4.4 Experimental procedures ...... 129

4.3.1 RNA Transcription and Purification ...... 129

4.3.2 Ribozyme Cleavage with Pre-oxidized Peptide ...... 130

4.3.3 Ribozyme Facilitated bPNA Oxidation ...... 130

4.4.4 In situ Oxidation and Cleavage ...... 130

4.5 Additional Data ...... 131

4.5.1 bPNA Initiated U-3 Cleavage Under 1 mM Mg2+ ...... 131

4.5.2 U-3 Cleavage with Pre-oxidized bPNAs ...... 132

4.5.3 In situ bPNA Oxidation and U-3 Cleavage ...... 133

4.5.4 bPNA Initiated U-(2,3) Cleavage Under 1 mM Mg2+ ...... 135

4.5.5 U-(2,3) Cleavage with Pre-oxidized bPNAs ...... 136

4.6 References for Chapter 4 ...... 137

APPENDIX A

Peptide Ligation and RNA Cleavage via An Abiotic Template Interface...... 145

A.1 Overview ...... 146

A.2 Introduction: ...... 146

A.3 Results and Discussion ...... 148

A.4 Conclusion ...... 156

A.5 Acknowledgement ...... 156

A.6 References for Appendix A ...... 157

APPENDIX B

General Guidelines for Bacterial Cell Culture ...... 164

B.1 Sterilization ...... 165

B.2 Stock solution preparation ...... 165

B.3 Pouring Agar plates ...... 166

B.4 Streak bacteria cells onto agar plate ...... 166

APPENDIX C

Preparation of Competent E. Coli Cells ...... 168

C.1 Introduction ...... 169

C.2 Protocol 1: Preparing Chemically Competent E. Coli Cells ...... 170

C.2.1 Material ...... 170

C.2.2 Procedure ...... 170

C.3 Protocol 2: Preparing Electrocompetent E. Coli Cells ...... 173

C.3.1 Material ...... 173

C.3.2 Procedure ...... 173

C.4 References for Appendix C ...... 176

APPENDIX D

Transformation of Competent E. Coli Cells ...... 177

D.1 Introduction ...... 178

D.2 Protocol 1: Transforming Chemically Competent E. Coli Cells ...... 178

D.3 Protocol 2: Transforming Electrocompetent E. Coli Cells ...... 179

D.4 Protocol 3: Determine Transformation Efficiency ...... 180

D.5 Reference for Appendix D ...... 181

xii

APPENDIX E

T7 RNA Polymerase Purification ...... 182

E.1 Introduction ...... 183

E.2 Protocol 1: Purification of His-Tag T7 RNA Polymerase ...... 183

E.2.1 Material ...... 183

E.2.2 Equipment and Consumables ...... 184

E.2.3 Solutions ...... 184

E.2.4 Procedures ...... 186

E.3 Protocol 2: SDS-PAGE for Purity Analysis ...... 188

E.3.1 Material ...... 188

E.3.2 Apparatus ...... 189

E.3.3 Procedure ...... 189

E.4 Protocol 3: T7 RNA Polymerase Activity Test ...... 193

E.4.1 Material ...... 193

E.4.2 Equipment ...... 193

E.4.3 DNA Template Design for T7 Transcription ...... 194

E.5 References for Appendix E ...... 197

APPENDIX F

Drosophila Schneider 2 Cells Culture and Protein Expression ...... 199

F.1 Introduction ...... 200

F.2 Materials ...... 201

F.2.1 Cell Culture Reagents ...... 201

F.2.2 Buffers and Stock Solutions ...... 201

F.2.3 Consumables and Apparatus ...... 201

xiii

F.3 Protocol 1: General Culture of Drosophila melanogaster S2 Cells ...... 202

F.3.1 Thawing Cells From Frozen Stock: ...... 202

F.3.2 Subculturing Drosophila melanogaster S2 Cells ...... 203

F.3.3 Generating Frozen Stocks of Drosophila melanogaster S2 Cells ...... 203

F.4 Protocol 2: Hygromycin Kill Curve for Hygromycin B Selection ...... 205

F.4.1 Overview ...... 205

F.4.2 Procedures ...... 205

F.5 Protocol 3: Calcium Phosphate Transfection ...... 207

F.5.1 Overview ...... 207

F.5.2 Procedure ...... 208

F.6 Protocol 4: Stable Transfection of S2 Cells ...... 210

F.7 Protocol 5: Adapting Stably Transfected S2 Cells ...... 211

F.8 Protocol 6: HER2ECD(1-4) Expression and Purification via S2 Cells ...... 211

F.9 References for Appendix F ...... 213

APPENDIX G

Protein Expression via Baculovirus System ...... 215

G.1 Introduction ...... 216

G.1.1 Overview ...... 216

G.1.2 Bac-to-Bac HBM TOPO Expression System ...... 217

G.1.3 TOPO Cloning ...... 219

G.1.4 Tn7 Transposition ...... 219

G.1.5 Honey Bee Melitin Secretion Signal Peptide ...... 220

G.2 Protocol 1: General Methods in Sf9 Cell Culture ...... 221

G.2.1 Material ...... 221

xiv

G.2.2 Recover Frozen Sf9 Cells in SF-900 II SFM ...... 221

G.2.3 Maintain Sf9 Cell Culture ...... 222

G.2.4 Cryopreservation of Sf9 Cells ...... 222

G.3 Protocol 2: Acquire HER2ECD(1-3)-encoding Sequence via PCR ...... 223

G.3.1 Material ...... 223

G.3.2 Procedure ...... 223

G.4 Protocol 3: TOPO Cloning HER2ECD(1-3) into pFastBac/HBM ...... 227

G.4.1 Overview ...... 227

G.4.2 Material ...... 227

G.4.3 Procedure ...... 228

G.5 Protocol 4 - Generating Recombinant Bacmid via Tn7 Transposition...... 232

G.5.1 Material ...... 232

G.5.1 Procedure ...... 232

G.6 Protocol 5: Analyzing Recombinant Bacmid DNA by PCR ...... 238

G.6.1 Material ...... 239

G.6.2 Procedure ...... 240

G.7 References for Appendix G ...... 242

References ...... 246

Chapter 1 ...... 246

Chapter 2 ...... 260

Chapter 3 ...... 265

Chapter 4 ...... 271

Appendix A ...... 279

Appendix C ...... 285

Appendix D ...... 285

Appendix E ...... 285

Appendix F ...... 286

Appendix G ...... 288

xvi

List of Figures

Figure 1.1. Watson-Crick base pair ...... 3

Figure 1.2. Helical structures of DNAs and RNAs ...... 5

Figure 1.3. G-quadruplex structure ...... 6

Figure 1.4. RNA folds with Hoogsteen pair and wobble pair ...... 7

Figure 1.5. 2D and 3D triplex structures ...... 8

Figure 1.6. Triplex strand orientations...... 9

Figure 1.7. Nucleobase modifications ...... 11

Figure 1.8. Sugar conformations...... 13

Figure 1.9. Sugar modifications ...... 14

Figure 1.10. Backbone modifications for charge neutralization ...... 15

Figure 1.11. PNA structure ...... 16

Figure 1.12. Schematic illustration of possible PNA binding modes ...... 17

Figure 1.13. PNA antigene functions ...... 19

Figure 1.14. PNA antisense functions ...... 20

Figure 1.15. 4-step PCR for mutation analysis ...... 22

Figure 1.16. Janus-Wedge nucleobase for bifacial targeting ...... 24

Figure 1.17. Triplex formation via bifacial nucleobase displayed PNA ...... 25

Figure 1.18. Melamine-acridine conjugate for bifacial targeting ...... 26

Figure 1.19. Melamine tagged PNA with native peptide backbone ...... 27

Figure 1.20. Melamine displaying bifacial peptide nucleic acid ...... 29

xvii

Figure 1.21. Binding studies on trimer and dimer formation ...... 30

Figure 2.1. bPNA and DNA constructs...... 51

Figure 2.2. bPNA-DNA binding profiles...... 53

Figure 2.3. Transcriptional repression by bPNA-DNA triplex ...... 55

Figure 2.4. Control transcriptions ...... 56

Figure 2.5. ExoT resistance of bPNA-DNA and RNA triplexes ...... 58

Figure 2.6. Exonuclease T resistance control ...... 59

Figure 2.7. Reverse transcription inhibition via bPNA-RNA triplex ...... 61

Figure 2.8. AMV reverse transcription control ...... 61

Figure 2.9. 1H NMR spectra of Fmoc-Lys-Melamine monomer ...... 70

Figure 2.10. 13C NMR spectrum of Fmoc-Lys-Melamine monomer ...... 71

Figure 2.11. HPLC and MALDI of H2N-(EM*)6G ...... 72

Figure 2.12. HPLC and MALDI of H2N-(EM*)8G ...... 72

Figure 2.13. HPLC and MALDI of H2N-(EM*)10G ...... 73

Figure 2.14. HPLC and MALDI of Cbf-(β-A)-(EM*)6G (Cbf-6) ...... 73

Figure 2.15. HPLC and MALDI of Cbf-(β-A)-(EM*)8G (Cbf-8) ...... 74

Figure 2.16. HPLC and MALDI of Cbf-(β-A)-(EM*)10G (Cbf-10 ...... 74

Figure 3.1. bPNA and nucleic acid designs ...... 84

Figure 3.2. Rescue of Aptamer−Protein Binding ...... 86

Figure 3.3. Binding profiles of IgE-T2 ...... 86

Figure 3.4. Binding onto IgE by D17.4 and IgE-T2...... 87

Figure 3.5. Rescue of RNA aptamer-DFHBI binding ...... 88

Figure 3.6. Fluorescence activation via bPNA-U-Spinach complexation ...... 89

Figure 3.7. Binding of bPNAs to U-Spinach ...... 90

Figure 3.8. bPNA dependent fluorescence activation ...... 91 xviii

Figure 3.9. DFHBI dependent fluorescence activation ...... 92

Figure 3.10. U-ribozymes rescue by bPNA ...... 93

Figure 3.11. U-3 ribozyme activity rescue via bPNAs ...... 94

Figure 3.12. Kinetics of U-3 ribozyme cleavage rescue...... 95

Figure 3.13. Mg2+ dependence of U-3 cleavage...... 96

Figure 3.14. in situ splicing of U-3 via bPNA 10...... 97

Figure 3.15. U-(2,3) activity restoration via bPNA triplexation ...... 98

Figure 3.16. Kinetic of U-(2,3) ribozyme cleavage rescue...... 99

Figure 3.17. 1H NMR of DFHBI precursor ...... 108

Figure 3.18. 1H NMR of DFHBI ...... 109

Figure 3.19. Fluorescence activation of tRNALysU-Spinach ...... 110

Figure 3.20. U-3 cleavage at higher Mg2+ concentration ...... 111

Figure 3.21. U-3 cleavage at lower Mg2+ concentration ...... 112

Figure 3.22. Gel images of Mg2+ dependent U-(2,3) cleavage ...... 113

Figure 3.23. Kinetics of Mg2+ dependent U-(2,3) cleavage ...... 114

Figure 4.1. bPNA structure and experimental designs ...... 125

Figure 4.2. bPNA structures of N-ternimal thiol (6) and C-terminal thiol (7) ...... 126

Figure 4.3. U-3 ribozyme-templated bPNA thiol oxidation ...... 127

Figure 4.4. Thiol oxidation coupled U-3 self-cleavage ...... 128

Figure 4.5. Two-way communication scheme ...... 129

Figure 4.6. bPNA length dependent U-3 cleavage ...... 131

Figure 4.7. U-3 cleavage restored by pre-oxidized bPNAs...... 132

Figure 4.8. in siu bPNA oxidation and U-3 cleavage ...... 133

Figure 4.9. Emergence of U-3 cleavage via templated bPNA oxidation ...... 134

Figure 4.10. bPNA length dependence of U-(2,3) cleavage ...... 135 xix

Figure 4.11. U-(2,3) cleavage restored by pre-oxidized bPNAs ...... 136

Figure A.1. bPNA structure and experimental design ...... 147

Figure A.2. DNA-template bPNA native ligation ...... 149

Figure A.3. DNA-templated bPNA oligomerization ...... 151

Figure A.4. Template effect on bPNA native ligation ...... 152

Figure A.5. DNA- and RNA-templated thiol oxidation ...... 153

Figure A.6. Ribozyme-templated bPNA thiol oxidation leads to ribozyme cleavage ..... 155

Figure E.1: Purity evaluation through SDS-PAGE ...... 192

Figure E.2. T7 RNA polymerase activity estimation ...... 196

Figure F.1. Hygromycin B kill curve result ...... 207

Figure G.1. Baculovirus life cycle in vivo and in vitro ...... 218

Figure G.2. PCR primer set design scheme ...... 224

Figure G.3. PCR product yield examination with 1.0% agarose gel in 1× TBE ...... 226

Figure G.4. Double digestion for inserted fragment orientation determination ...... 231

Figure G.5. Gel image of double digested plasmid minipreps ...... 232

Figure G.6. lac operon tightly regulates the expression of β-galacosidase ...... 236

Figure G.7. Blue-white screen basics ...... 237

Figure G.8. Transposed area is primed under pUC/M13 forward primer and reverse primers ...... 238

Figure G.9. PCR analysis of recombinant bacmid ...... 241

List of Tables

Table B.1. Antibiotic Solutions Recipes ...... 165

Table C.1. Equilibrium Buffer ...... 171

Table C.2. Storage Buffer ...... 172

Table E.1. Lysis Buffer (500 mL) ...... 184

Table E.2. Washing Buffer (200 mL) ...... 184

Table E.3. Elution Buffer 1 (100 mM Imidazole, 50 mL) ...... 184

Table E.4. Elution Buffer 2 (200 mM Imidazole, 50 mL) ...... 185

Table E.5. Elution Buffer 3 (300 mM Imidazole, 50 mL) ...... 185

Table E.6. Elution Buffer 4 (400 mM Imidazole, 50 mL) ...... 185

Table E.7. Elution Buffer 5 (500 mM Imidazole, 50 mL) ...... 185

Table E.8. Storage Buffer (2000 mL) ...... 185

Table E.9. Resolving Gel Recipes ...... 190

Table E.10. Stacking Gel Recipes ...... 190

Table E.11. 10x Tris-Glycine-SDS Buffer ...... 191

Table E.12. 10x High Yield Transcription Buffer ...... 195

Table E.13. in vitro T7 Transcription Protocol...... 195

Table F.1. Freezing Medium Recipe ...... 204

Table F.2. Ca2+-DNA Mixture ...... 208

Table F.3. 2x HEPES-Buffered Saline ...... 209

Table F.4. Lysis Buffer Recipe ...... 212

xxi

Table G.1. Freezing Medium Recipe ...... 222

Table G.2. HER2ECD(1-3) PCR Cloning Protocol ...... 224

Table G.3. PCR Thermocycle ...... 225

Table G.4. TOPO Ligation Protocol ...... 228

Table G.5. Double Digestion Recipe for Orientation Check ...... 230

Table G.6. Selection Agar Plate Recipe ...... 233

Table G.7. DH10Bac Growing Medium ...... 237

Table G.8. Estimation on Product Length from Bacmid PCR ...... 239

Table G.9. Bacmid PCR Protocol...... 240

Table G.10. Bacmid PCR Thermocycle...... 240

xxii

CHAPTER 1

Nucleic Acids Structures and Targeting Strategies

This thesis describes studies on synthetic hybrid nucleic acid structures and their biological functions derived from bifacial peptide nucleic acid (bPNA). The bPNA hybrid structures exhibit both similarities and differences from native folds. In this chapter, we will first describe native nucleic acid folds; following the overview on synthetic strategies inspired from the native structures for improved nucleic acid recognition and hybridization; then the discussion on rational design and preliminary studies regarding our bPNA system will be presented.

1.1 Native Nucleic Acids Structures

DNA and RNA molecules are biopolymers of deoxyribose and ribose 1’-conjugated purine and pyrimidine nucleobases linked through 3’-5’ phosphodiester backbones. The four bases A, T/U, G and Care displayed, enabling A-T/U and G-C Watson-Crick base pairing.

(Figure 1.1) Both the genetic inheritances as well as the phenotypic presentations are determined by accurate biological processes involving DNA and RNA molecules.

Nucleobase complementarity, together with the structural and electrostatic characters of their phosphate backbone is the origin of nucleic acid structures and biological functions.

1.1.1 Duplexes

Both DNA and RNA molecules are competent in Watson-Crick pairing, (Figure 1.1) which leads to folding that is driven by base-stacking interactions. Generally, DNAs adopts a right-handed double helical structure termed B-form helix exhibiting a 10.5 bp pitch and a helical diameter of 20 Å; two unequally sized grooves are presented along the helical 2 structure with major groove being 22 Å wide and minor groove being 12 Å wide. According to crystal structures, the width of the major groove provides sufficient accessibility for enzymatic readout on the Hoogsteen edge of paired nucleobases, meanwhile the narrowness of the minor groove suggests more closed chemical environments and less intermolecular interactions. (Figure 1.2 b) RNA helices, on the contrary, preferably adopt the A-form helical structure with an 11 bp pitch and a wider helical diameter around 23 Å; the differences in helical topology are attributed to the presentation of 2’-hydroxyl group on the ribose ring leading to 3’-endo sugar conformations other than 2’-endo that is presented in B-form helices.

Figure 1.1. Watson-Crick base pair. Two hydrogen bonds between A-T/U pair, and three hydrogen bonds between G-C pair. 3

The difference in ribose conformation also results in a deeper and narrower major groove and wider and shallower minor groove. (Figure 1.2 c). Noticeably, B-form DNA double helices can interchanged into A-form (Figure 1.2 a) during enzymatic processing, for example DNA replication and RNA transcription. In addition to A- and B-form helices, DNA can adopt Z-form left-handed double helices (Figure 1.2 d) when unique sequences such as poly-d(GC)poly-d(GC) are presented1.

1.1.2 G-Quadruplex

Besides native duplex folding topologies, both DNA and RNA can fold into G-quadruplexes via Hoogsteen pairing. G-quadruplex formation is generally initiated under high concentration of monovalent metals for guanine rich nucleic acid sequences; it is a multi- layer planer structure with 4 Hoogsteen paired guanine positioned in one layer. (Figure

1.3) It can be an intramolecular, bimolecular or tetramolecular topology, adopting either parallel or antiparallel orientations.2

The most physiologically significant G-quadruplex is telomeric quadruplex consisting of repeating d(GGTTAG) sequence resides at the end of each chromatid, and functions as chromosome protecting segment from gene deterioration3 due to the lagging strand

“replication problem”.4-6 Telomere length is tightly associated with aging and mortality, and it is speculated that cancer cell immortality is presumably related to the up-regulation of telomerase that extensively elongates telomeric repeats.7 Also, putative G-quadruplex sequences have been detected in both prokaryotic and eukaryotic genome, majorly located near promoter regions, their possible roles in gene expression regulation thus have been suggested.8

Figure 1.2. Helical structures of DNAs and RNAs. (a) A-DNA; (b) B-DNA; (c) Z-DNA; (d) A- RNA.1

Figure 1.3. G-quadruplex structure. (Left) Schematic illustration of G-quadruplex hydrogen bonding network via Hoogsteen face interactions. (Right) Bimolecular anti-parallel DNA G- 9 quadruplex structure of d(G4T3G4).

1.1.3 RNA Folds

RNA molecules extensively employ Wobble pairing10, 11 and Hoogsteen edge interactions12, 13 to enhance and stabilize critical tertiary conformations. While native DNA function primarily as informational molecules, RNAs exhibit a wider range of biological function as catalytic ribozymes,14 regulatory riboswitches,15 and essential structural and functional parts in protein synthesis.16 (Figure 1.4)

Figure 1.4. RNA folds with Hoogsteen pair and wobble pair. (Left) Group I intron structure.17 (Right) HDV ribozyme that utilizes wobble pair at the active site.18

1.2 Nucleic Acids Triplex

1.2.1 Structural Characters of Triplex

The discovery of triplex nucleic acid19-21 demonstrated the functional relevance of an additional recognition interface within the major groove formed by the Hoogsteen edge of purine bases, this new interface presents both base pair and sequence specific information from a pre-formed duplex. The triple-helix formation is generally guided through the major groove Hoogsteen base pair between a Watson-Crick base-paired purine and an incoming purine or pyrimidine, and the interactions are believed to be exclusively on the purine side of the duplex.22 (Figure 1.5) 7

A B

Figure 1.5. 2D and 3D triplex structures. (A) Triple-helix structure with right hand two strands forming base-pair the left hand one strand docking on the major groove.22 (B) Schematic illustrations of purine exclusive Hoogsteen recognition.23

Triple-helical structures can be categorized into 3 classes based on the orientation of the third docking strand: TC-triplex, GT-triplex and GA-triplex. (Figure 1.6) TC-triplexes include T-AT and C-GC+ triple-helical structures, in which the poly-T and poly-C strands are in the same orientation as the poly-A and poly-G; GT-triplexes contain C-GG and T-

AT triplexes, whose third strand can be parallel to poly-purine or anti-parallel to poly- purine via reverse Hoogsteen; GA-triplexes contain C-GG and T-AA triplexes that only form through reverse Hoogsteen interactions.24 It has been reported by Singleton and co- workers25 that within excellent agreement with the nearest-neighbor analysis the triplex structures are less thermodynamically stable than duplexes, the enthalpy contribution for a base-triplet formation is about 2.0 ± 0.1 kcal/mole, which is less than 1/3 of that from a base-pair formation at 6.3 ± 0,3 kcal/mole. The formation conditions for C-GC+ triplex requires acidic conditions with pH < 6.0 for efficient cytosine protonation;26 furthermore, high divalent metal (Mg2+) concentrations and cationic additives (Spermidine or other polyamines) are required to overcome the electrostatic repulsion between the three phosphate backbones in a triplex structure.27-32 8

Figure 1.6. Triplex strand orientations.

Even though the formation of stable triplex requires non-physiological conditions and there are specific sequence limitations, triple-helical nucleic acids still have great potential as artificial gene targeting and regulation agents. Genomic mapping indicates abundant sites that are potential triplex formation sites within human genome, especially concentrated near promoter regions, indicating the possible physiological relevance of triple-helical structures in regulatory roles.33-39

Triplex formation under physiological conditions requires extensive modifications to the nucleobase, sugar, and the phosphate backbone. Below, we briefly discuss each category of modifications.

1.3.2 Nucleobase Modifications

Nucleobase modifications that favor triplex formation have been primarily focused on the pyrimidine base that docks on the Hoogsteen face of the pre-formed duplex. Methylation

Me on 5-cytosine ( C) elevates the pKa of the protonated N3 site and thus ameliorates the pH restrictions of forming C-GC+ triplex (Figure 1.7 a), MeC modification enhances thermal stability of DNA triplexes,40, 41 however the stabilizing effect on RNA triplex presented to be minimal.42

Strobel and Dervan demonstrated that C-methylated strands can tightly associate with duplex DNA under physiological salt and pH conditions, and more intriguingly, can regulate enzymatic cleavage.43, 44 Because of thymine containing oligonucleotides can adopt both parallel triplexation (TC) and anti-parallel triplexation (GT) orientations, thymine modifications also facilitate enhanced recognition capability. Improved triplex stability was observed from modification of 2’-deoxyuridine.45 Conjugation of amino group derivatives to the 5th position on the pyrimidine ring further promotes triplex binding and lowers the required concentration of divalent metals,46-49 presumably by decreasing electrostatic repulsion between strands by installation of cationic amine groups (Figure 1.7 b). In addition to amino groups, intercalators can as well be conjugated to oligonucleotides to enhance triplex stability.50, 51 (Figure 1.7 c) With the conjugation of an intercalator, triplex formation can occur under neutral conditions with un-modified cytosine.50

There are fewer studies on purine for triplex formation. One of the most studied modifications is 6-thioguanine. 52 (Figure 1.7 d) Given the decreased basicity of S vs. O, this O−>S modification effectively results in the loss of a hydrogen bond acceptor and prevents the formation of competing structures under high salt concentrations, such as G- quadruplex53 and GA homodimer.30 In contrast, 8-amino guanine and adenine derivatives 10

a b

c d

f e

C C

Figure 1.7. Nucleobase modifications. (a) 5-methylcytosine. (b) 2’-deoxyuridine and its amino group derivatives.48 (c) Intercalator conjugation.51 (d) 6-thioguanine.52 (e) 8-amino derivatives of adenine, guanine and their triplexation patterns.54 (f) Synthetic bases for full major groove targeting during strand disruption.55, 56 11

provide addition hydrogen bonding sites on the purine ring, and promote GA-triplex

(antiparallel) formation under near physiological conditions. 52, 54 (Figure 1.7 e)

As mentioned above, triplex formation stems from purine exclusive Hoogsteen recognition, thus expand the recognition interface beyond purine base, besides enhancing Hoogsteen hydrogen binding, can also facilitate in triplex formation. Nucleobases contain novel hydrogen binding patterns and enable to simultaneously target purine-pyrimidine from duplex major groove are synthesized and characterized. 55-58 (Figure 1.7 f) Those synthetic bases prove to restore triplex stability disruption when oligopurine-oligopyrimidine continuity is interrupted by a reverse base pair such as T-A and C-G.59, 60

1.3.3 Sugar Modifications

Thermal stability of nucleic acid triplexes have been reported to be higher for RNA triplexes than DNA triplexes,19, 61 which prompts synthetic investigations into construct ribose analogues that exhibits similar topology.24 The intrinsic problem associated with RNA triplex is the short lifetime of ribonucleotides under neutral conditions due to 2’-OH associated degradation and RNase A digestion. Hence, a degradation resistant RNA backbone mimic can further broadens the utility of RNA therapeutics. The general principle for sugar modification is to maintain sugar conformation as 3’-endo but diminish the reactivity of 2’-hydroxyl group. (Figure 1.8)

Figure 1.8. Sugar conformations. (Left) C2’-endo sugar pucker of deoxyribose. (Right) C3’- endo sugar pucker of ribose.

Examples of sugar modifications include: 1) 2’-Methyl (2’-OMe) modification proves to effectively maintain the sugar conformation and the backbone flexibility while blocking the

2’-OH assisted hydrolysis, thus favors stable triplex formation;62 2) 2’-aminoethyl (2’-AE) modification further incorporates positive charge into the backbone, thus decreases electrostatic repulsion;63 3) conformational restricted ribose analogues such like Locked nucleic acid (LNA, O2’, O4’-methylene-linked nucleic acid) and ENA (O2’, O4’-ethylene- linked nucleic acid) stabilize nucleic acid folds through covalent linkage of the ribose 2’ and 4’ position. 64-68 (Figure 1.9)

1.3.4 Backbone Modifications

Strong electrostatic repulsion, originated from the poly-anionic phosphate backbone, is a destabilizing to triplex formation. Beside previously mentioned cationic group incorporation to partially neutralize local repulsion, chemical modifications on the backbone that neutralize the repulsive can elicit more efficient triplexation. Such modifications can be achieved via cationic phosphoramidate during oligonucleotide solid phase synthesis.69, 70 Other than changing electrostatic properties, backbone has also been modified to phosphorothioate24 or N3’-P5’ phosphoramidate71 linkage that displays 13 higher resistance against nuclease for the expectation of future in vivo applications.

(Figure 1.10)

a b

c d

Figure 1.9. Sugar modifications. (a) O2’-Methyl modification (2’-OMe). (b) O2’-Aminoethyl modification (2’-AE). (c) O2’, O4’-methylene-linked nucleic acid (LNA). (d) O2’, O4’-ethylene- linked nucleic acid (ENA).

1.4 Peptide Nucleic Acid (PNA) as DNA Mimic

1.4.1 PNA Structure and Recognition Mechanism

Apart from extensive modification on nucleic acid scaffold, Nielsen and co-workers established a polymeric form of oligonucleotide mimic termed peptide nucleic acid (PNA), which displays native nucleobases on a synthetic neutral pseudo-peptide backbone.72-75

Each nucleobase is conjugated to the glycine nitrogen on the N-(2-aminoethyl) glycine backbone via a methylene carbonyl linker. The pseudo-peptide backbone resembles the backbone bond arrangement on native DNA oligonucleotides, which presents as 6 bonds 14

along the backbone between nucleoside and 3 bonds linking the nucleobase to the phosphate backbone, but without the destabilizing electrostatic repulsions. (Figure 1.11)

Figure 1.10. Backbone modification for charge neutralization. Modifications on phosphate backbone by substituting O with S (PS), or NH (PN). These modifications still maintain the phosphate diester nucleic acid backbone signature.

Figure 1.11. PNA structure. (Top) PNA forms duplex with complimentary DNA strand.72 (Bottom) Backbone bond arrangement comparison between PNA and DNA, PNA design follows “6+3” rule, which represents 6 bonds (blue) on the backbone for monomeric unit, and 3 bonds (red) linking base to the backbone.74

As anticipated, PNAs have been reported to exhibit higher thermal stability when complexed with complimentary DNA and RNA, compare to the corresponding native duplex systems, presumably the significant thermal stabilization stems from the abolished backbone repulsions.75 However, the original attempts of forming more stable PNA- 16

DNA2/RNA2 triplex system turned out to be unsuccessful due to the preferable formations of PNA2-DNA/RNA triplexes or PNA-DNA/RNA duplex through duplex strand displacement.74, 76(Figure 1.12) It has been indicated that the strand displacement can be initiated through inherent DNA breathing that displays temporary structural disruption, which allows the quick association of complementary PNA strand.76

Figure 1.12. Schematic illustration of possible PNA binding modes.74

1.4.2 Antigene Therapeutics of PNA

The strand displacement process upon PNA association although differs from conventional nucleic acid triplex formation, presents a novel strategy for DNA targeting.

The invasive nature of PNA targeting ameliorates the sequence selection restriction on poly-purine or poly-pyrimidine sequences for Hoogsteen recognition, PNA can carry mixed-base sequences for a broader range of targeting.77 One of the intriguing aspects will be antigene therapeutics.

The possible mechanism of PNA hybridization through DNA breathing permits PNA facilitated genome structure probing. Unique structural features that are thermodynamically less stable have been reported to be frequently presented at promoter regions or replication origins, such as inverted repeats,78 H-form DNA79 and cruciform DNA,

Corey and co-workers have demonstrated that PNAs are able to target those structures and further regulate the associated biological functions.80

It has been demonstrated that PNA hybridization to regular duplex DNA can be used to down-regulate gene transcription and translation at the genomic level, and the regulatory functions are based on the two structural features arisen from strand displacement: the D- loop formation on the displaced strand, and the hetero-duplex or hetero-triplex formation on the complementary strand. (Figure 1.13 a) For example, Nielsen and co-workers demonstrated that D-loop is prone to single strand specific endonuclease digestion, hence

PNA hybridization can facilitate sequence specific genome cleavage.81 (Figure 1.13 b)

Further, hetero-duplex and hetero-triplex structures from PNA hybridization are lack of electrostatic signature in addition to high thermal stability and possible increase in strand bulkiness, thus impede enzymatic recognition and access. (Figure 1.13 c) For instance, triplex has been indicated to block transcription factor access to promoter region thus down-regulate the gene transcription;82 Allfrey presented highly specific PNA invasion into

CAG repeats and subsequently down-regulated gene expression under semi- physiological conditions in permeabilized cells;83 moreover, an in situ negative feedback regulatory loop has also been explored, in which the transient duplex disruption caused by transcription initiates the PNA binding on the template that eventually turn down the transcription efficiency.84

Figure 1.13. PNA antigene functions. (a) Formation of D-loop and PNA-DNA hybrid. (b) D- loop mediated double strand cleavage. (c) Inhibition of enzymatic access via PNA-DNA hybrid.

Corey and co-workers further exploited the capability of PNA targeting on telomeric G- quadruplex.85 Surprisingly, PNA with complementary sequence can efficiently hybrid with human telomeric repeat and consequently repress telomerase elongation activity.

1.4.3 Antisense Regulation of PNA

PNA can hybrid with mRNA in either triplex form (PNA2-mRNA) or duplex form (PNA- mRNA) with similar affinity and specificity, hence PNA applications in antisense regulation have been proposed and exploited.

Nielsen and co-workers and others presented translational inhibition via employment of

PNA-mRNA hybrid duplex and triplex.77, 86 (Figure 1.14 a) Moreover, in contrast to down- regulation of protein translation, PNA has been utilized in modulating mRNA splicing pattern.87 (Figure 1.14 b) In addition, one would expect the stable hybrid of PNA-RNA is as well inhibitory to many binding proteins, and presumably can be applied to intercept signal transduction pathways.

Figure 1.14. PNA antisense functions. (a) Ribosomal rest during translation due to PNA hybridization. (b) Alternative mRNA splicing pattern from mRNA masking via PNA hybridization.

1.4.4 PNAs as Detection Probes and Biosensors

The high specificity of sequence recognition and the low tolerance of sequence mismatch75 prompted development of PNAs as high sensitivity probes and biosensors.

Stanley first described mutation analysis utilizing Tm difference between PNA and DNA primers for specific mutant sequence PCR amplification88. (Figure 1.15) Later on, chemical modifications on PNAs, which are complementary to targeting sequences, for high sensitivity detection have been developed, modifications include but not limit to PNA-chip anchoring,89, 90 and fluorescence labeling.91-93 Recently, structure formation dependent detection approaches have been developed. Researchers employed homologous rather than complementary PNAs in dimeric heteroquadruplex formation for probing DNA and

RNA G-quadruplex displaying sequences.94-98

1.4.5 Chemical Modifications of PNA

Inherent obstacles of PNA that prevents its broad biological applications are the inefficient cellular uptake and the endosome trapping of non-modified PNA, which can be overcome via chemical modifications of PNAs with proper internalization signals.99 Generally, modifications can be categorized into two classes: one for attaching cell penetrating molecules via amine, carboxyl or thiol specific reaction to the PNA terminus or reactive side chains;100 the other for incorporating the attachment of signal molecules via step-wise solid phase synthesis along with the PNA generation.101

Figure 1.15. 4-step PCR for mutation analysis. Additional PNA annealing step enables the amplification of mutation containing DNA.

Cell penetrating peptide (CPP) displaying positively charged amino acids such as lysine and arginine is the most commonly used internalization signal in antisense PNA delivery, it can be fused to PNA via disulfide or maleimide chemistry,102-105 or coupled with PNA solid phase synthesis.106, 107 It has been reported that PNA-CPP conjugates can exhibit comparable gene inhibitory effect as commercial siRNA knock down assays.103 Similar chemistry also applies to ligand conjugation for receptor targeting leading to positive endocytosis pathway.99, 102

The ease of chemical modifications on PNA overcomes numerous application obstacles, in addition to those mentioned above, PNAs have been conjugated with intercalator and cationic molecules hence are able to hybridize with nucleic acids under physiological conditions, hugely improved from initial conditions under which low salt concentration that destabilize duplex is required for kinetically trapping PNA-DNA/RNA complexes before applied to experiments.72, 74-76

1.5 Bifacial Nucleobase Analogues

Although PNA hybridization system differs from triplex nucleic system in both binding mechanism and thermal stability, they share the similarity in only being able to target one strand of the nucleic acid duplex. Hence, a hybridization system that can evenly readout information from both strands would be expected to exhibit more proficient and unbiased targeting.

Lehn and co-workers108 first established a Janus-Wedge nucleobase analogue displaying two hydrogen bonding arrays that are able to simultaneously bind two native nucleobase via their Watson-Crick faces. (Figure 1.16) It is postulated that Janus Wedge nucleobases can be employed for generalized duplex targeting, and applied in base mismatch probing.

Tor and co-worker also established a Janus-like nucleobase that displays self- complementary hydrogen bonding faces and can engage in Watson-Crick interaction with both T and A,109 however only duplex formation is detected.

Figure 1.16. Janus-Wedge nucleobase for bifacial targeting.108 (a) Schematic illustration of Janus-Wedge nucleobase design, hydrogen bonding patterns are indicated. (b) C-U mismatch targeting heterocycle.

1.5.1 Bifacial Nucleobase Displaying PNA for Triplex Formation

McLaughlin further exploited the Janus-Wedge heterocycle initiated triad formation by displaying multiple of the nucleobase analogue on the PNA backbone.110 Kinetic studies indicated that Janus-Wedge displaying PNA can readily associate with pre-installed triplex

24 binding site with picomolar affinity and similar thermodynamic profile as duplex. (Figure

1.17)

Figure 1.17. Triplex formation via bifacial nucleobase displayed PNA. 110 (a) Hydrogen bonding scheme of a triad layer. (b) Design of pre-installed binding site.

Following the initial study on Janus-Wedge PNA, additional nucleobase analogues have been designed and tested.111 Different Janus-Wedge nucleobases can be mixed and displayed on one PNA backbone, thus allows for targeting various sequences. Further, the Janus-Wedge displaying PNA exhibits thermodynamically low tolerance for sub- optimal hydrogen bonding.

1.5.2 Melamine Derivatives Recognize T-T/U-U Mismatch

Janus-Wedge nucleobases that target natural base pairs are generally self- complementary, and as depicted by Lehn108 and Tor109 targeting process needs to compete with both base pair and self-aggregation. Targeting mismatch interface with

Janus-Wedge nucleobases eliminates the aggregation problem, and has been presented to be potential approaches in diagnostics and therapeutics.

Baranger and Zimmerman demonstrated that melamine-acridine conjugate functions as a

U-U mismatch ligand, exhibits sub-micromolar affinity towards CUG repeats in Myotonic dystrophy type I. Multiple binding events of the ligand subsequently prevent the binding of muscleblind-like proteins to the CUG repeat.112-114 (Figure 1.18)

Figure 1.18. Melamine-acridine conjugate for bifacial targeting.

1.6 PNA With Native Peptide Backbone

Nielsen termed the nucleobases installed polyamide molecule as peptide nucleic acid

(PNA), however it is constructed via pseudopeptide backbone for the purpose of achieving optimal bond arrangement similar to native nucleic acid. The development in PNA related antigene and antisense approaches prompted investigations into possible nucleic acid targeting PNAs endow native peptide backbone. Eschenmoser and co-workers115 demonstrated the melamine tagged peptide for proficient poly-T targeting and PNA-DNA duplex formation. Noticeably, the alternative placement of anionic aspartic acid side chain was not obstructive to the peptide-oligoT recognition; on the contrary, negatively charge

26 melamine peptide exhibits comparable binding profile to many of the neutral PNAs. (Figure

1.19)

Figure 1.19. Melamine tagged PNA with native peptide backbone.115

1.7 Melamine Displayed Bifacial Peptide Nucleic Acid (bPNA)

Inspired by the work of Zimmerman,113 Eschenmoser115 and others, we designed and synthesized a melamine-displaying 21-mer oligo-peptide, and its oligo-T targeting function was comprehensively studied. The general architecture mimics native nucleic acid via alternating the placements of glutamic acids and melamine-derived lysines along the native peptide backbone, and we term this peptide as bifacial peptide nucleic acid (bPNA) attributing to the Janus-Wedge character of melamine molecules. (Figure 1.20)

Preliminary binding studies reveal that polymeric display of bifacial melamine moieties presents two multivalent arrays of hydrogen bonding sites, to which two strands of oligo-

T can associate simultaneously. In addition to the heterotrimeric stem of bPNA-(dT10)2, 27

a b

bPNA can also cooperatively fold linear dT10C10T10 into a heterodimeric stem-loop structure. The binding kinetics of heterotrimer and heterodimer formations along the bPNA have been systematically investigated, which render to be 4000 nM2 for trimer system and

2.7 nM for dimeric stem-loop system. (Figure 1.21)

However similar to many nucleic acid targeting strategies that utilizing hydrogen bonding network as the recognition interface, bPNA system presented in this thesis is drastically different in binding mechanism, kinetics and topology.

Melamine bPNA fully exhibits its Janus-Wedge property of unbiased recognition on T-rich sequences, which is distinctly different from both triplex system and PNA hybridization system. Compare to the established Janus-Wedge system from McLaughlin, the bifacial recognition doesn’t require pre-installed binding site, yet no destabilizing effects were observed from the negative charges. And the most intriguing character of melamine bPNA recognition is the presentation of associative binding. Unlike conventional PNA hybridization that is presented to be invasive and result in strand displacement, melamine bPNA associatively brings unstructured nucleic acid strand into well-defined and thermal stable folds. Further, melamine bPNA hybridization occurs at physiological condition without chemical modification such as intercalator or cationic group, which vacates modification sites for prosthetic group or specific targeting ligands, thus broadens the targeting range and the potential applications towards.

Figure 1.20. Melamine displaying bifacial peptide nucleic acid.116 Schematic illustrations of its structure and nucleic acid binding topologies.

In the following chapters, applications of melamine bPNA in multiple biological systems will be presented in details. In chapter 2, we first investigated the inhibitory regulation of bPNA modulated triplex system in transcription, reverse transcription and exonuclease digestion systems; as the conventional triplex system and PNA system exhibit repression in transcription,117-119 DNA replication120 after proper base, sugar and backbone modifications, this novel bPNA triplex system can as well be efficiently inhibitory to multiples enzymatic processes however with no additional modifications. In Chapter 3, we will demonstrate that bPNA triplex can also function as allosteric activity switch for the onset of ribozyme catalysis and aptamer target binding. In Chapter 4, a two way communication study will be presented, between an abiotic interface for bPNA ligation and a biological ribozyme cleavage site. Overall, this thesis will provide systematic description of this novel bPNA triplex system, through which design-based nucleic acid bio-techniques can be crafted. It is anticipated that the bPNA-based methodology presented herein will be a broadly useful tool for study and control of nucleic acid function.

Journalofthe A m erican Chem icalSociety Com m unication

trichlorotriazine17 w ith am m onia and/or dimethylam ine and Boc-lysine yie ld ed the dimeth ylated and tetram ethylated m elam inolysine derivatives, w hich w ere used in SPPS to provide peptides 3 and 4,respectively. N anoparticle assem bly from m elam ine and cyanuric acid derivatives results from noncovalentpolym erization ofthe two- fold sym m etric heterocycle recognition faces;thym ine hasonly one recognition face com plem entary to m elam ine, and thus discrete assem bly w as anticipated. G ratifyingly, no large peptide−DNA aggregates w ere detectable by dynam ic light scattering, consis te nt with the model of discrete triplex form ation. Peptide-triggered base-stacking sig natu res w ere observed by UV absorbance changes,w ith a 1:2 stoichiom etry between 1 and dT 10, consistent w ith bivalent m elam ine− thym ine recognition and formation oftriplex 5;peptide alone did not induce such a signal (Figures 1 and 2, Supporting a b

Figure 3. C ircular dichroism spectra in D PBS, pH 7.4, of (A) 1 com plexed with dT 10 (□) vs dT 10 alone (--) and (B) 1 com plexed with dT 10C 10T 10 (○) vs dT 10C 10T 10 alone (--).Peptide 1 in both is at 5 μM concentration ( ), while dT 10 and dT 10C 10T 10 are m aintained at 10 and 5 μM,respectively.Electrophoretic m obility shiftassaysimaged by C y5 fluorescence for(C) C y5-dT 10 (D N A 1) and (D ) C y5-dT 10C 10T 10 (D N A 2)at 20 nM in eachlane, w ithincreasing peptide 1 concentration from leftto right.(E) Relative electrophoretic m obilities of the free DNA oligos and their peptide com plexes, a m ixture of Figure 2. Peptide 1 titrated into (A) dT and (B) Flc-dT C T - 10 10 10 10 com plex 5 and DNA shown in the central lane.See SI for further D abcyl, followed by UV absorbance (260 nm ) and fluorescein 1 c d details. em ission (521 nm ),respectively.Saturation is observed at 33 and 50 m ol% peptide, in dicating 1:2 and 1:1 peptide:D NA bindin g stoichiom etries in (A) and (B),respectively. C ooperative m elting w as observed fortriplex 5 and hairpin 6 by both UV and fluorescence dequenching and differential Information (SI)). U nlike DNA and peptide−DNA 4 triplex scanning calorim etry (D SC ) (Figure 4, SI). T he hairpin structures w hich optimally form at high saltw ith divalentm etal ions (1−2 M N aCl, 50 mM M gC l2), robust assem bly w as observed understandard saltconditions(D ulbecco’sphosphate buffered saline, DPBS), akin to th e conditions used by Eschenm oser and K rishnam urthy9c for PNA−DNA duplex formation.Similarly,UV absorbance signatures indicated a 1:1 Figure 1.21. Binding studies on trimer and dimer formation.116 (a) UV titration of bPNA 10 intobinding dT10. (b)stoichiom Fluorescence etry quenchingbetween titiration1 ofand bPNAdT 1010 intoC 10 FlcT-10dT,10asC10Tw10 ould-Dabcyl.be (c) Fluorescence titration of dT10 into Cbf-bPNA 10. (d) Fluorescence anisotropy titration of dTexpected10C10T10 into Cbfif -anbPNAintram 10 olecular peptide−DNA triplex structure formed from the dT 10 terminiofdT 10C 10T 10.Indeed,binding of1 to Flc-dT 10C 10T 10-Dabcylresulted in m aximalfluorescein quenching at a 1:1 peptide−DNA ratio, supportive of the formation of heterodimeric hairpin structure 6,w hich w ould Figure 4.(A) First-derivative plot of m elting transitions of triplex 5 9c (--) and hairpin 6 ( ) followed by UV absorbance (260 nm ). bring the 3′and 5′ends of the oligo in close proximity, N ormalized absorbance change is shown inset.(B) D SC upscan traces resulting in efficientdabcylquenching offluorescein (Figure 2). oftriplex 5 ( ) and hairpin 6 ( ), with downscan traces shown as T riplex and hairpin formation30 w as further corroborated by dashed regular and bold lines.Peptide−DNA ratios used in triplex 5 circular dichroism , w hich indicated significant structuring of and hairpin 6 experiments were 1:2 and 1:1,respectively.D SC and UV DNA upon addition ofpeptide,signified by the developm entof experim ents w ere perform ed in DPBS, pH 7.4 at peptide a negative CD signalat 260 nm at the expense ofa positive CD concentrations of 25 μM (D SC ) and in UV experiments, 1 μM(5) and 2.5 μM(6). signalat 280 nm ,w hich weassign to the peptide com plex and free DNA,respectively (Figure 3).N otably,w hile the peak at heterodimer structure 6 w as m ore thermally stable (T = 54 280 nm is com pletely ablated in the triplex,there is a residual m °C) than the heterotrimeric triplex 5 (T m = 43 °C ). For peak in the hairpin; this is consistent w ith the presence of an com parison, a dA 10-T 10 duplex has T m = 35 °C,and a dA 10- unstructured C loop found in hairpin 6 but not triplex 5. 10 (T 10)2 triplex has T m = 17 °C in the presence of 50 mM 18 C lean transformation ofDNA to peptide−DNA com plex bands M gC l2. It is calculated thata dA 10-T 10 duplex w illhave T m = on native polyacrylam ide electrophoresis indicated discrete 22.5 °C undersimilarsaltconditions.Reversible peptide−DNA peptide−DNA recognition in both triplex and hairpin contexts com plexation w as supported by observation of endothermic (Figure 3). m elting and exothermic cooling peaksby D SC ;though triplex 5

833 dx.doi.org/10.1021/ja2099326 |J. Am .Chem .Soc.2012,134,832−835 1.8 References for Chapter 1

1. Arnott, S., Historical article: DNA polymorphism and the early history of the double

helix. Trends Biochem. Sci. 2006, 31, 349-354.

2. Bochman, M. L.; Paeschke, K.; Zakian, V. A., DNA secondary structures: stability

and function of G-quadruplex structures. Nature Reviews Genetics 2012.

3. Tsai, Y. C.; Qi, H.; Liu, L. F., Protection of DNA ends by telomeric 3′ G-tail

sequences. J. Biol. Chem. 2007.

4. Lingner, J.; Cooper, J. P.; Cech, T. R., Telomerase and DNA end replication: no

longer a lagging strand problem? Science 1995, 269, 1533-4.

5. Paeschke, K.; Simonsson, T.; Postberg, J.; Rhodes, D.; Lipps, H. J., Telomere

end-binding proteins control the formation of G-quadruplex DNA structures in vivo.

Nat. Struct. Mol. Biol. 2005, 12, 847-54.

6. Gilson, E.; Géli, V., How telomeres are replicated. Nature Reviews Molecular Cell

Biology 2007.

7. Neidle, S.; Parkinson, G., Telomere maintenance as a target for anticancer drug

discovery. Nature Reviews Drug Discovery 2002.

8. Huppert, J. L.; Balasubramanian, S., G-quadruplexes in promoters throughout the

human genome. Nucleic Acids Res. 2007.

9. Burge, S.; Parkinson, G. N.; Hazel, P.; Todd, A. K.; Neidle, S., Quadruplex DNA:

sequence, topology and structure. Nucleic Acids Res. 2006, 34, 5402-15.

10. Agris, P. F.; Vendeix, F. A. P.; Graham, W. D., tRNA's wobble decoding of the

genome: 40 years of modification. J. Mol. Biol. 2007.

11. Varani, G.; McClain, W. H., The G· U wobble base pair. EMBO reports 2000.

12. Nissen, P.; Ippolito, J. A.; Ban, N.; Moore, P. B.; Steitz, T. A., RNA tertiary

interactions in the large ribosomal subunit: the A-minor motif. Proc. Natl. Acad. Sci.

U. S. A. 2001, 98, 4899-903.

13. Theimer, C. A.; Blois, C. A.; Feigon, J., Structure of the human telomerase RNA

pseudoknot reveals conserved tertiary interactions essential for function. Mol. Cell

2005, 17, 671-82.

14. Fedor, M. J.; Williamson, J. R., The catalytic diversity of RNAs. Nat. Rev. Mol. Cell

Biol. 2005, 6, 399-412.

15. Smith, A. M.; Fuchs, R. T.; Grundy, F. J.; Henkin, T. M., Riboswitch RNAs:

regulation of gene expression by direct monitoring of a physiological signal. RNA

Biol. 2010, 7, 104-10.

16. Stern, S.; Powers, T.; Changchien, L. M.; Noller, H. F., RNA-protein interactions in

30S ribosomal subunits: folding and function of 16S rRNA. Science 1989, 244,

783-90.

17. Adams, P. L.; Stahley, M. R.; Gill, M. L.; Kosek, A. B., Crystal structure of a group

I intron splicing intermediate. RNA 2004.

18. Sripathi, K. N.; Banas, P.; Reblova, K.; Sponer, J.; Otyepka, M.; Walter, N. G.,

Wobble pairs of the HDV ribozyme play specific roles in stabilization of active site

dynamics. Phys. Chem. Chem. Phys. 2015, 17, 5887-900.

19. Felsenfeld, G.; Davies, D. R.; Rich, A., Formation of a Three-Stranded

Polynucleotide Molecule. J. Am. Chem. Soc. 1957, 79, 2023-2024.

20. Morgan, A. R.; Wells, R. D., Specificity of the three-stranded complex formation

between double-stranded DNA and single-stranded RNA containing repeating

nucleotide sequences. J. Mol. Biol. 1968.

21. Beal, P. A.; Dervan, P. B., Second structural motif for recognition of DNA by

oligonucleotide-directed triple-helix formation. Science 1991, 251, 1360-3.

22. Radhakrishnan, I.; Patel, D. J., Solution structure of a purine· purine· pyrimidine

DNA triplex containing G· GC and T· AT triples. Structure 1993.

23. Ghosh, I.; Stains, C. I.; Ooi, A. T.; Segal, D. J., Direct detection of double-stranded

DNA: Molecular methods and applications for DNA diagnostics. Mol Biosyst 2006,

2, 551-60.

24. Oussedik, K.; Halby, L.; Arimondo, P. B., The triple helix: 50 years later, the

outcome. Nucleic acids … 2008.

25. Plum, G. E.; Park, Y. W.; Singleton, S. F.; Dervan, P. B.; Breslauer, K. J.,

Thermodynamic characterization of the stability and the melting behavior of a DNA

triplex: a spectroscopic and calorimetric study. Proc. Natl. Acad. Sci. U. S. A. 1990,

87, 9436-40.

26. Lyamichev, V. I.; Mirkin, S. M.; Frank-Kamenetskii, M. D.; Cantor, C. R., A stable

complex between homopyrimidine oligomers and the homologous regions of

duplex DNAs. Nucleic Acids Res. 1988, 16, 2165-78.

27. Lyamichev, V. I.; Voloshin, O. N.; Frank-Kamenetskii, M. D.; Soyfer, V. N.,

Photofootprinting of DNA triplexes. Nucleic Acids Res. 1991, 19, 1633-8.

28. Thomas, T.; Thomas, T. J., Selectivity of polyamines in triplex DNA stabilization.

Biochemistry 1993, 32, 14068-74.

29. Thomas, T. J.; Kulkarni, G. D.; Greenfield, N. J.; Shirahata, A.; Thomas, T.,

Structural specificity effects of trivalent polyamine analogues on the stabilization

and conformational plasticity of triplex DNA. Biochem. J. 1996, 319 ( Pt 2), 591-9.

30. Arimondo, P. B.; Barcelo, F.; Sun, J. S.; Maurizot, J. C.; Garestier, T.; Helene, C.,

Triple helix formation by (G,A)-containing oligonucleotides: asymmetric sequence

effect. Biochemistry 1998, 37, 16627-35.

31. Blume, S. W.; Lebowitz, J.; Zacharias, W.; Guarcello, V.; Mayfield, C. A.;

Ebbinghaus, S. W.; Bates, P.; Jones, D. E., Jr.; Trent, J.; Vigneswaran, N.; Miller,

D. M., The integral divalent cation within the intermolecular purine*purine.

pyrimidine structure: a variable determinant of the potential for and characteristics

of the triple helical association. Nucleic Acids Res. 1999, 27, 695-702.

32. Floris, R.; Scaggiante, B.; Manzini, G.; Quadrifoglio, F.; Xodo, L. E., Effect of

cations on purine.purine.pyrimidine triple helix formation in mixed-valence salt

solutions. Eur. J. Biochem. 1999, 260, 801-9.

33. Goni, J. R.; de la Cruz, X.; Orozco, M., Triplex-forming oligonucleotide target

sequences in the human genome. Nucleic Acids Res. 2004, 32, 354-60.

34. Goni, J. R.; Vaquerizas, J. M.; Dopazo, J.; Orozco, M., Exploring the reasons for

the large density of triplex-forming oligonucleotide target sequences in the human

regulatory regions. BMC Genomics 2006, 7, 63.

35. Durland, R. H.; Kessler, D. J.; Gunnell, S.; Duvic, M.; Pettitt, B. M.; Hogan, M. E.,

Binding of triple helix forming oligonucleotides to sites in gene promoters.

Biochemistry 1991, 30, 9246-55.

36. Duval-Valentin, G.; Thuong, N. T.; Helene, C., Specific inhibition of transcription

by triple helix-forming oligonucleotides. Proc. Natl. Acad. Sci. U. S. A. 1992, 89,

504-8.

37. Maher, L. J., 3rd, Inhibition of T7 RNA polymerase initiation by triple-helical DNA

complexes: a model for artificial gene repression. Biochemistry 1992, 31, 7587-94.

38. Maher, L. J., 3rd; Dervan, P. B.; Wold, B., Analysis of promoter-specific repression

by triple-helical DNA complexes in a eukaryotic cell-free transcription system.

Biochemistry 1992, 31, 70-81.

39. Young, S. L.; Krawczyk, S. H.; Matteucci, M. D.; Toole, J. J., Triple helix formation

inhibits transcription elongation in vitro. Proc. Natl. Acad. Sci. U. S. A. 1991, 88,

10023-6.

40. Lee, J. S.; Woodsworth, M. L.; Latimer, L. J. P.; Morgan, A. R., Poly(pyrimidine)

poly(purine) synthetic DNAs containing 5-methylcytosine form stable triplexes at

neutral pH. Nucleic Acids Res. 1984, 12, 6603-6614.

41. Xodo, L. E.; Manzini, G.; Quadrifoglio, F.; van der Marel, G. A.; van Boom, J. H.,

Effect pf 5-methylcytosine on the stability of triple-stranded DNA—a

thermodynamic study. Nucleic Acids Res. 1991, 19, 5625-5631.

42. Wang, S.; Xu, Y.; Kool, E. T., Recognition of RNA by triplex formation: divergent

effects of pyrimidine C-5 methylation. Bioorg. Med. Chem. 1997.

43. Strobel, S. A.; Dervan, P. B., Single-site enzymatic cleavage of yeast genomic

DNA mediated by triple helix formation. Nature 1991, 350, 172-4.

44. Strobel, S. A.; Doucette-Stamm, L. A.; Riba, L.; Housman, D. E.; Dervan, P. B.,

Site-specific cleavage of human chromosome 4 mediated by triple-helix formation.

Science 1991, 254, 1639-42.

45. Mills, M.; Arimondo, P. B.; Lacroix, L.; Garestier, T.; Klump, H.; Mergny, J. L.,

Chemical modification of the third strand: differential effects on purine and

pyrimidine triple helix formation. Biochemistry 2002, 41, 357-66.

46. Sollogoub, M.; Darby, R. A. J.; Cuenoud, B.; Brown, T.; Fox, K. R., Stable DNA

Triple Helix Formation Using Oligonucleotides Containing 2‘-Aminoethoxy,5-

propargylamino-U†. Biochemistry 2002, 41, 7224-7231. 35

47. Brazier, J. A.; Shibata, T.; Townsley, J.; Taylor, B. F.; Frary, E.; Williams, N. H.;

Williams, D. M., Amino-functionalized DNA: the properties of C5-amino-alkyl

substituted 2'-deoxyuridines and their application in DNA triplex formation. Nucleic

Acids Res. 2005, 33, 1362-71.

48. Bijapur, J.; Keppler, M. D.; Bergqvist, S.; Brown, T.; Fox, K. R., 5-(1-

propargylamino)-2'-deoxyuridine (UP): a novel thymidine analogue for generating

DNA triplexes with increased stability. Nucleic Acids Res. 1999, 27, 1802-9.

49. Froehler, B. C.; Wadwani, S.; Terhorst, T. J.; Gerrard, S. R.,

Oligodeoxynucleotides containing C-5 propyne analogs of 2′-deoxyuridine and

2′-deoxycytidine. Tetrahedron Lett. 1992.

50. Sun, J. S.; Francois, J. C.; Montenay-Garestier, T.; Saison-Behmoaras, T.; Roig,

V.; Thuong, N. T.; Helene, C., Sequence-specific intercalating agents: intercalation

at specific sequences on duplex DNA via major groove recognition by

oligonucleotide-intercalator conjugates. Proceedings of the National Academy of

Sciences 1989, 86, 9198-9202.

51. Mouscadet, J. F.; Ketterle, C.; Goulaouic, H.; Carteau, S.; Subra, F.; Le Bret, M.;

Auclair, C., Triple helix formation with short oligonucleotide-intercalator conjugates

matching the HIV-1 U3 LTR end sequence. Biochemistry 1994, 33, 4187-96.

52. Spackova, N.; Cubero, E.; Sponer, J.; Orozco, M., Theoretical study of the guanine

--> 6-thioguanine substitution in duplexes, triplexes, and tetraplexes. J. Am. Chem.

Soc. 2004, 126, 14642-50.

53. Davis, J. T., G-quartets 40 years later: from 5'-GMP to molecular biology and

supramolecular chemistry. Angew. Chem. Int. Ed. Engl. 2004, 43, 668-98.

54. Avino, A.; Cubero, E.; Gonzalez, C.; Eritja, R.; Orozco, M., Antiparallel triple

helices. Structural characteristics and stabilization by 8-amino derivatives. J. Am.

Chem. Soc. 2003, 125, 16127-38.

55. Guianvarc’h, D.; Benhida, R.; Fourrey, J.-L.; Maurisse, R.; Sun, J.-S., Incorporation

of a novel nucleobase allows stable oligonucleotide-directed triple helix formation

at the target sequence containing a purine·pyrimidine interruption. Chemical

Communications 2001, 1814-1815.

56. Guianvarc'h, D.; Fourrey, J. L.; Maurisse, R.; Sun, J. S.; Benhida, R., Synthesis,

incorporation into triplex-forming oligonucleotide, and binding properties of a novel

2'-deoxy-C-nucleoside featuring a 6-(thiazolyl-5)benzimidazole nucleobase. Org

Lett 2002, 4, 4209-12.

57. Guianvarc'h, D.; Fourrey, J. L.; Maurisse, R.; Sun, J. S., Design of artificial

nucleobases for the recognition of the AT inversion by triple-helix forming

oligonucleotides: A structure–stability relationship study and neighbour bases

effect. Bioorg. Med. Chem. 2003.

58. Wang, Y.; Rusling, D. A.; Powers, V. E.; Lack, O.; Osborne, S. D.; Fox, K. R.;

Brown, T., Stable recognition of TA interruptions by triplex forming oligonucleotides

containing a novel nucleoside. Biochemistry 2005, 44, 5884-92.

59. Best, G. C.; Dervan, P. B., Energetics of Formation of Sixteen Triple Helical

Complexes Which Vary at a Single Position within a Pyrimidine Motif. J. Am. Chem.

Soc. 1995, 117, 1187-1193.

60. Greenberg, W. A.; Dervan, P. B., Energetics of Formation of Sixteen Triple Helical

Complexes Which Vary at a Single Position Within a Purine Motif. J. Am. Chem.

Soc. 1995, 117, 5016-5022.

61. Roberts, R. W.; Crothers, D. M., Stability and properties of double and triple helices:

dramatic effects of RNA or DNA backbone composition. Science 1992, 258, 1463-

62. Shimizu, M.; Konishi, A.; Shimada, Y.; Inoue, H.; Ohtsuka, E., Oligo(2'-O-

methyl)ribonucleotides. Effective probes for duplex DNA. FEBS Lett. 1992, 302,

155-8.

63. Seidman, M. M.; Puri, N.; Majumdar, A.; Cuenoud, B.; Miller, P. S.; Alam, R., The

development of bioactive triple helix-forming oligonucleotides. Ann. N. Y. Acad.

Sci. 2005, 1058, 119-27.

64. Torigoe, H.; Hari, Y.; Sekiguchi, M.; Obika, S.; Imanishi, T., 2'-O,4'-C-methylene

bridged nucleic acid modification promotes pyrimidine motif triplex DNA formation

at physiological pH: thermodynamic and kinetic studies. J. Biol. Chem. 2001, 276,

2354-60.

65. Petersen, M.; Wengel, J., LNA: a versatile tool for therapeutics and genomics.

Trends Biotechnol. 2003, 21, 74-81.

66. Brunet, E.; Alberti, P.; Perrouault, L.; Babu, R.; Wengel, J.; Giovannangeli, C.,

Exploring cellular activity of locked nucleic acid-modified triplex-forming

oligonucleotides and defining its molecular basis. J. Biol. Chem. 2005, 280, 20076-

85.

67. Sun, B. W.; Babu, B. R.; Sorensen, M. D.; Zakrzewska, K.; Wengel, J.; Sun, J. S.,

Sequence and pH effects of LNA-containing triple helix-forming oligonucleotides:

physical chemistry, biochemistry, and modeling studies. Biochemistry 2004, 43,

4160-9.

68. Hojland, T.; Kumar, S.; Babu, B. R.; Umemoto, T.; Albaek, N.; Sharma, P. K.;

Nielsen, P.; Wengel, J., LNA (locked nucleic acid) and analogs as triplex-forming

oligonucleotides. Org Biomol Chem 2007, 5, 2375-9.

69. Michel, T.; Martinand-Mari, C.; Debart, F.; Lebleu, B.; Robbins, I.; Vasseur, J. J.,

Cationic phosphoramidate alpha-oligonucleotides efficiently target single-stranded

DNA and RNA and inhibit hepatitis C virus IRES-mediated translation. Nucleic

Acids Res. 2003, 31, 5282-90.

70. Michel, T.; Debart, F.; Heitz, F.; Vasseur, J. J., Highly stable DNA triplexes formed

with cationic phosphoramidate pyrimidine alpha-oligonucleotides. Chembiochem

2005, 6, 1254-62.

71. Gryaznov, S. M.; Lloyd, D. H.; Chen, J. K.; Schultz, R. G.; DeDionisio, L. A.;

Ratmeyer, L.; Wilson, W. D., Oligonucleotide N3'-->P5' phosphoramidates.

Proceedings of the National Academy of Sciences 1995, 92, 5798-5802.

72. Nielsen, P. E.; Haaima, G., Peptide nucleic acid (PNA). A DNA mimic with a

pseudopeptide backbone. Chemical Society Reviews 1997, 26, 73.

73. Nielsen, P. E., Sequence-selective DNA recognition by synthetic ligands.

Bioconjug. Chem. 1991, 2, 1-12.

74. Nielsen, P. E., Peptide Nucleic Acid. A Molecule with Two Identities. Accounts of

Chemical Research 1999, 32, 624-630.

75. Nielsen, P. E.; Egholm, M.; Buchardt, O., Peptide nucleic acid (PNA). A DNA mimic

with a peptide backbone. Bioconjug. Chem. 1994, 5, 3-7.

76. Nielsen, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O., Sequence-selective

recognition of DNA by strand displacement with a thymine-substituted polyamide.

Science 1991, 254, 1497-500.

77. Hanvey, J. C.; Peffer, N. J.; Bisi, J. E.; Thomson, S. A.; Cadilla, R.; Josey, J. A.;

Ricca, D. J.; Hassman, C. F.; Bonham, M. A.; Au, K. G.; et al., Antisense and

antigene properties of peptide nucleic acids. Science 1992, 258, 1481-5.

78. Lilley, D. M., The inverted repeat as a recognizable structural feature in supercoiled

DNA molecules. Proc. Natl. Acad. Sci. U. S. A. 1980, 77, 6468-6472.

79. Mirkin, S. M.; Lyamichev, V. I.; Drushlyak, K. N.; Dobrynin, V. N.; Filippov, S. A.;

Frank-Kamenetskii, M. D., DNA H form requires a homopurine-homopyrimidine

mirror repeat. Nature 1987, 330, 495-497.

80. Norton, J. C.; Waggenspack, J. H.; Varnum, E.; Corey, D. R., Targeting peptide

nucleic acid-protein conjugates to structural features within duplex DNA. Bioorg.

Med. Chem. 1995, 3, 437-445.

81. Demidov, V.; Frank-Kamenetskii, M. D.; Egholm, M.; Buchardt, O.; Nielsen, P. E.,

Sequence selective double strand DNA cleavage by peptide nucleic acid (PNA)

targeting using nuclease S1. Nucleic Acids Res. 1993, 21, 2103-2107.

82. Vickers, i. A.; Griffith, M. C.; Ramasamy, K.; Risen, L. M.; Freier, S. M., Inhibition

of NF-κB specific transcriptional activation by PNA strand invasion. Nucleic Acids

Res. 1995, 23, 3003-3008.

83. Morris, P. L., Invasion of the CAG Triplet Repeats by a Complementary Peptide

Nucleic Acid Inhibits Transcription of the Androgen Receptor and TATA-binding

Protein Genes and Correlates with Refolding of an Active Nucleosome Containing

a Unique AR Gene Sequence. J. Biol. Chem. 1996, 271, 13228-13233.

84. Larsen, H. J.; Nielsen, P. E., Transcription-mediated binding of peptide nucleic acid

(PNA) to double-stranded DNA: sequence-specific suicide transcription. Nucleic

Acids Res. 1996.

85. Norton, J. C.; Piatyszek, M. A.; Wright, W. E.; Shay, J. W.; Corey, D. R., Inhibition

of human telomerase activity by peptide nucleic acids. Nat. Biotechnol. 1996, 14,

615-619.

86. Knudsen, H.; Nielsen, P. E., Antisense properties of duplex-and triplex-forming

PNAs. Nucleic Acids Res. 1996.

87. Abes, S.; Williams, D.; Prevot, P.; Thierry, A.; Gait, M. J.; Lebleu, B., Endosome

trapping limits the efficiency of splicing correction by PNA-oligolysine conjugates.

J. Control. Release 2006, 110, 595-604.

88. Orum, H.; Nielsen, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O.; Stanley, C., Single

base pair mutation analysis by PNA directed PCR clamping. Nucleic Acids Res.

1993, 21, 5332-5336.

89. Wang, J.; Palecek, E.; Nielsen, P. E.; Rivas, G.; Cai, X.; Shiraishi, H.; Dontha, N.;

Luo, D.; Farias, P. A. M., Peptide Nucleic Acid Probes for Sequence-Specific DNA

Biosensors. J. Am. Chem. Soc. 1996, 118, 7667-7670.

90. D'Agata, R.; Corradini, R.; Grasso, G.; Marchelli, R.; Spoto, G., Ultrasensitive

detection of DNA by PNA and nanoparticle-enhanced surface plasmon resonance

imaging. Chembiochem 2008, 9, 2067-70.

91. Kuhn, H.; Demidov, V. V.; Coull, J. M.; Fiandaca, M. J.; Gildea, B. D.; Frank-

Kamenetskii, M. D., Hybridization of DNA and PNA molecular beacons to single-

stranded and double-stranded DNA targets. J. Am. Chem. Soc. 2002, 124, 1097-

103.

92. Molenaar, C.; Wiesmeijer, K.; Verwoerd, N. P.; Khazen, S.; Eils, R.; Tanke, H. J.;

Dirks, R. W., Visualizing telomere dynamics in living mammalian cells using PNA

probes. EMBO J. 2003, 22, 6631-41.

93. Stender, H.; Williams, B.; Coull, J., PNA fluorescent in situ hybridization (FISH) for

rapid microbiology and cytogenetic analysis. Peptide Nucleic Acids 2014.

94. Datta, B.; Schmitt, C.; Armitage, B. A., Formation of a PNA2-DNA2 hybrid

quadruplex. J. Am. Chem. Soc. 2003, 125, 4111-4118.

95. Marin, V. L.; Armitage, B. A., RNA guanine quadruplex invasion by complementary

and homologous PNA probes. J. Am. Chem. Soc. 2005, 127, 8032-8033.

96. Marin, V. L.; Armitage, B. A., Hybridization of complementary and homologous

peptide nucleic acid oligomers to a guanine quadruplex-forming RNA.

Biochemistry 2006, 45, 1745-1754.

97. Roy, S.; Tanious, F. A.; Wilson, W. D.; Ly, D. H.; Armitage, B. A., High-affinity

homologous peptide nucleic acid probes for targeting a quadruplex-forming

sequence from a MYC promoter element. Biochemistry 2007, 46, 10433-10443.

98. Lusvarghi, S.; Murphy, C. T.; Roy, S.; Tanious, F. A.; Sacui, I.; Wilson, W. D.; Ly,

D. H.; Armitage, B. A., Loop and backbone modifications of peptide nucleic acid

improve g-quadruplex binding selectivity. J. Am. Chem. Soc. 2009, 131, 18415-

18424.

99. Koppelhus, U.; Nielsen, P. E., Cellular delivery of peptide nucleic acid (PNA).

Advanced drug delivery reviews 2003.

100. Koppelhus, U.; Awasthi, S. K.; Zachar, V., Cell-dependent differential cellular

uptake of PNA, peptides, and PNA-peptide conjugates. … and Nucleic Acid …

2002.

101. Cutrona, G.; Carpaneto, E. M.; Ulivi, M.; Roncella, S.; Landt, O.; Ferrarini, M.; Boffa,

L. C., Effects in live cells of a c-myc anti-gene PNA linked to a nuclear localization

signal. Nat. Biotechnol. 2000, 18, 300-3.

102. Pooga, M.; Soomets, U.; Hallbrink, M.; Valkna, A.; Saar, K., Cell penetrating PNA

constructs regulate galanin receptor levels and modify pain. Nat. Biotechnol. 1998.

103. Turner, J. J.; Jones, S.; Fabani, M. M.; Ivanova, G., RNA targeting with peptide

conjugates of oligonucleotides, siRNA and PNA. Blood Cells 2007.

104. Abes, S.; Turner, J. J.; Ivanova, G. D.; Owen, D.; Williams, D.; Arzumanov, A.;

Clair, P.; Gait, M. J.; Lebleu, B., Efficient splicing correction by PNA conjugation to

an R6-Penetratin delivery peptide. Nucleic Acids Res. 2007, 35, 4495-502.

105. Ivanova, G. D.; Arzumanov, A.; Abes, R.; Yin, H., Improved cell-penetrating

peptide–PNA conjugates for splicing redirection in HeLa cells and exon skipping

in mdx mouse muscle. Nucleic acids … 2008.

106. Good, L.; Awasthi, S. K.; Dryselius, R.; Larsson, O.; Nielsen, P. E., Bactericidal

antisense effects of peptide-PNA conjugates. Nat. Biotechnol. 2001, 19, 360-4.

107. Bendifallah, N.; Rasmussen, F. W.; Zachar, V.; Ebbesen, P.; Nielsen, P. E.;

Koppelhus, U., Evaluation of cell-penetrating peptides (CPPs) as vehicles for

intracellular delivery of antisense peptide nucleic acid (PNA). Bioconjug. Chem.

2006, 17, 750-8.

108. Branda, N.; Kurz, G.; Lehn, J.-M., JANUS WEDGES: a new approach towards

nucleobase-pair recognition. Chemical Communications 1996, 2443.

109. Shin, D.; Tor, Y., Bifacial Nucleoside as a Surrogate for Both T and A in Duplex

DNA. J. Am. Chem. Soc. 2011.

110. Chen, D.; Meena; Sharma, S. K.; McLaughlin, L. W., Formation and stability of a

Janus-Wedge type of DNA triplex. J. Am. Chem. Soc. 2004, 126, 70-71.

111. Chen, H.; Meena; McLaughlin, L. W., A Janus-Wedge DNA triplex with A-W1-T

and G-W2-C base triplets. J. Am. Chem. Soc. 2008, 130, 13190-1.

112. Arambula, J. F.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C., A simple

ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein

binding. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 16068-73.

113. Wong, C. H.; Fu, Y.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C.,

Selective inhibition of MBNL1-CCUG interaction by small molecules toward

potential therapeutic agents for myotonic dystrophy type 2 (DM2). Nucleic Acids

Res. 2011, 39, 8881-90.

114. Jahromi, A. H.; Nguyen, L.; Fu, Y.; Miller, K. A.; Baranger, A. M.; Zimmerman, S.

C., A novel CUG(exp).MBNL1 inhibitor with therapeutic potential for myotonic

dystrophy type 1. ACS Chem. Biol. 2013, 8, 1037-43.

115. Mittapalli, G. K.; Reddy, K. R.; Xiong, H., Mapping the landscape of potentially

primordial informational oligomers: Oligodipeptides and oligodipeptoids tagged

with triazines as recognition elements. Angewandte Chemie International Edition

2007.

116. Zeng, Y.; Pratumyot, Y.; Piao, X.; Bong, D., Discrete assembly of synthetic

peptide-DNA triplex structures from polyvalent melamine-thymine bifacial

recognition. J. Am. Chem. Soc. 2012, 134, 832-5.

117. Hewett, P. W.; Daft, E. L.; Laughton, C. A.; Ahmad, S.; Ahmed, A.; Murray, J. C.,

Selective inhibition of the human tie-1 promoter with triplex-forming

oligonucleotides targeted to Ets binding sites. Mol. Med. 2006, 12, 8-16.

118. Karympalis, V.; Kalopita, K.; Zarros, A.; Carageorgiou, H., Regulation of gene

expression via triple helical formations. Biochemistry (Mosc.) 2004, 69, 855-60.

119. Svinarchuk, F.; Nagibneva, I.; Cherny, D.; Ait-Si-Ali, S.; Pritchard, L. L.; Robin, P.;

Malvy, C.; Harel-Bellan, A.; Chern, D., Recruitment of transcription factors to the

target site by triplex-forming oligonucleotides. Nucleic Acids Res. 1997, 25, 3459-

64.

120. Guieysse, A. L.; Praseuth, D.; Francois, J. C.; Helene, C., Inhibition of replication

initiation by triple helix-forming oligonucleotides. Biochem. Biophys. Res. Commun.

1995, 217, 186-94.

CHAPTER 2

bPNA Triplex Inhibits Enzymatic Access

to DNA and RNA

This chapter is reproduced with permission from ChemBioChem, 2014, 15, 31-36.

2.1 Introduction

We have previously reported that synthetic melamine-displaying α-peptides, termed bifacial peptide nucleic acid (bPNA), can selectively bind thymine-rich DNA into synthetic heterotriplex structures composed of two T-rich DNA strands assembled onto a bPNA strand.1, 2 When T-rich domains are separated by an intervening oligonucleotide sequence of 4-10 nt, a hairpin triplex stem-loop structure is formed with high affinity (Kd ~ 2 nM). We hypothesized that these high melting complexes (Tm=57°C) would be competitive with protein enzymes whose native substrates are DNA or RNA. Herein we report the effective in vitro inhibition of transcription, reverse-transcription and exonuclease function via the formation of synthetic bPNA-nucleic acid triplex structures formed by folding of DNA or

RNA around a single bPNA template strand. These studies demonstrate versatile and selective bPNA targeting of both DNA and RNA substrates with native-like oligonucleotide composition, suggesting possible application of bPNAs as synthetic regulators of nucleic acid function.

Many native nucleic acid processes are controlled via induction of structure in DNA or

RNA substrates, by protein complexation,3 small molecule metabolite binding4 or both.

This mechanism for repression of enzymatic “read-through” has been observed in transcriptional, reverse-transcriptional and translational regulation; further, the increased stability of folded nucleic acid structures towards chemical and enzymatic degradation is well-known. Moderate substrate affinity appears to be sufficient to modulate nucleic acid function, as observed in native riboswitches with nanomolar to micromolar range affinity

47 to metabolite targets.5, 6 Metabolite binding toggles native riboswitch sequences between folded conformations with distinct function, thus regulating transcription, translation initiation, mRNA degradation and splicing. A similar strategy of sequence- encoded conformational control is found natively in prokaryotic transcriptional regulation wherein palindromic transcripts signal transcriptional termination by folding into RNA hairpin structures.6, 7 We hypothesized that bifacial peptide nucleic acid (bPNA), which binds T- rich DNA and induces the formation of synthetic bPNA-DNA triplex-stem loop (hairpin) structures, could be used in a similar fashion as an artificial repressor of nucleic acid function.

Peptide nucleic acids from α-amino acids presenting nucleobases on derivatized sidechains are synthetically more convenient relative to the well-studied PNA backbone reported by Nielsen and Buchardt,8 which features a non-native peptide backbone N- amide acylated with a nucleobase moiety. A neutral PNA backbone9 results in enhanced stability of PNA hybrids with DNA and RNA relative to native hybrids, due to reduced interstrand electrostatic repulsion.8 Increased hybrid stability permits strand invasion of native duplexes to form PNA hybrid duplex and triplex structures, which suggested potential utility as antigene and antisense therapeutic agents. Notably, Nielsen demonstrated inhibition of enzymatic restriction upon complexation of an A10 tract proximal to restriction sites by oligo-T PNA;10 PNA-DNA triplex formation via Watson-Crick and

Hoogsteen base-pairing thus blocked enzymatic access to restriction sites. Corey11 and

Nielsen12 independently reported the design of bis-PNA “tail-clamps” which target pyrimidine-rich duplex sites for triplex invasion; this strategy has been shown to be effective in transcriptional repression.11, 13

In this chapter, the inhibitory effects of bPNA triplex were tested in three different 48 enzymatic systems: T7 RNA transcription, Exonuclease T digestion and AMV reverse transcription. For T7 RNA transcription system, DNA templates bearing T10-spacer-T10 bPNA triplexation sequences at various locations were designed, it is postulated that upon association a hairpin structure consisting the bPNA-nucleic acid triplex stem should be presented. Kinetic studies concerning bPNA association affinity and specificity on different DNA templates were conducted. bPNA showed robust cooperative association

toward all DNA templates despite the sequence complexity, and the measured Kds are uniformly around 20 nM. Transcriptional repression was observed for all template constructs with 50%-90% repression efficiency. For the exonuclease T system, efficient and structure specific exonuclease T resistance was observed for both DNA and RNA molecules, and the level of resistance was proportional to the length of bPNA. In the AMV reverse transcriptase system, reverse transcription was assayed on a 142 nt RNA template with bPNA binding site embedded in the center. Early aborted, short cDNA product was produced in reactions with bPNA structured RNA templates, and the production amount of short aborted cDNA was also proportional to the length of bPNA.

Overall, we find that bPNAs presenting 6, 8 and 10 bifacial nucleobases can selectively target T- and U-rich sites on long DNA and RNA substrates of otherwise heterogeneous composition. The mode of bPNA recognition drives a significant structural re-organization in the nucleic acid, transforming unstructured single-stranded nucleic acids into well folded triplex hairpin loop. Nanomolar range bPNA binding affinity provides sufficient hairpin stability to compete with enzymes for their DNA and RNA substrates, thus greatly limiting substrate access in transcription, exonuclease digestion and reverse transcription.

2.2 Results and Discussion

Bifacial peptide nucleic acid (bPNA) presents an alternative approach to block enzymatic access via non-native structuring of nucleic acid substrates. Conventional PNA triplex hybrids mimic DNA triplexes14, 15 by replacing two DNA strands with PNA: two PNA strands bind to a native oligopyrimidine via both Watson- Crick and Hoogsteen base-pairing, which can also lead to intramolecular folding, as in G-quartets.16-18 In contrast, melamine- based bPNA strands do not fold intramolecularly and serve as the template strand, binding two native pyrimidine nucleic acid (NA) strands on their Watson-Crick faces to form a bPNA-

NA triplex structure. (Figure 2.1) This approach is similar to “Janus-wedge” nucleobase- pairing introduced by Lehn,19-21 but addresses mismatch sites in which the two nucleobases are identical. It has been previously demonstrated that 2-fold symmetric triazine bases can base-pair with DNA when displayed on a PNA backbone22 and recognize T-T and U-U mismatch sites when coupled to an intercalator.23 Eschenmoser and Krishnamurthy have demonstrated that synthetic triazine bases displayed at alternate residues on α-peptide backbone and peptoid backbones can effectively form duplex structures with DNA and RNA partners.24, 25 A single-stranded oligonucleotide with two separated oligothymidine or oligouracil tracts is folded upon bPNA binding into a hairpin secondary structure with a bPNA-nucleic acid triplex stem. The recognition nucleobase mimic on bPNA is 1,3,5- triazine-2,4,6-triamine (melamine) which is attached to the bPNA backbone via lysine sidechain.

Figure 2.1. bPNA and DNA constructs. (Top) Structure of bPNAs 6, 8 and 10 used in this study, corresponding to repeat units n of the same number, and the melamine- thymine/uracil interaction driving bPNA triplex formation. Folding of a T/U-rich sequence in DNA or RNA in a bPNA triplex hairpin blocks enzyme (yellow) processing. (Bottom) Design of DNA constructs used in transcriptional repression and exonuclease resistance. A control construct features a sequence encoding tRNA-Lys flanked by an unstructured 5’ sequence and 3’ T7 RNA polymerase promoter (red). Binding sites for bPNAs are inserted as shown (blue) at locations 5’ of tRNA-Lys, mid-sequence, post-promoter (pp) and 3’ of the promoter. Decathymidine tracts are separated by the 21 nt promoter sequence in the hairpin promoter loop (HPPL) design.

Small molecules, peptides, lipids and polymers26 displaying melamine derivatives have been previously shown to avidly bind to synthetic hydrogen bonding complements in organic27-30 and aqueous solution.1, 2, 31-36 A bPNA sequence was designed with alternating residues of melamine- modified lysine (M*) and glutamic acid (E) to yield peptides of the general form (EM*)n, where repeat units (n) of 6, 8 and 10 were studied. This polyanionic design was expected to impart greater water solubility and ease of purification relative to conventional PNA, and minimize non-specific interactions with nucleic acids while binding and folding thymine-rich DNA sequences into hairpin structures. The biophysical underpinnings of this interaction are similar to DNA triplex formation,37, 38 with assembly driven by exothermic base-stacking. Prior bPNA binding studies revealed low nanomolar affinity to dT10 tracts with non-interacting C10 linkers, and related studies on triazine polymers suggested uracil nucleobase binding should be similarly robust.

2.2.1 Triplexes Inhibit in vitro T7 RNA Transcription.

We tested bPNA 10 affinity to longer DNA sequences (~127 nt) encoding a T7 RNA polymerase promoter region, tRNA-Lys and a dT10C10T10 domain. Fluorescence anisotropy binding isotherms generated with fluorescein-labeled 10 indicated low nanomolar (~21 nM) affinity to the large, heterogeneous DNA strand while no binding was observed to a similar DNA construct with a random unstructured sequence in place of the bPNA binding site (Figure 2.2). Similar 127 nt DNA templates were designed with 5’-

T10(CA)2T10-3’ domains placed at 4 different positions throughout the template: directly 3’ and 5’ of the T7 promoter site, in the middle of the template, at the 5’ terminus. These templates were all designed to host bPNA strands with 10 triazine rings (10) targeting the

10 T-T pairs presented in a DNA hairpin conformation.

Figure 2.2. bPNA-DNA binding profiles. (Top) Binding curves describing association of bPNA 10 with the 5’- hairpin DNA transcription template (Figure 2.1, T10C10T10) (—) or control sequence (---). Binding to DNA with double stranded () and single stranded () promoter regions is shown. (Center) Binding curves obtained by EMSA are shown of 10 to DNA transcription templates HPPL (, hairpin promoter loop binding site) and 3’HP (, 3’ hairpin binding site). (Bottom) SYBR gold stained EMSA indicating binding saturation and full gel shift of the 3’HP DNA to complex upon treatment with bPNA 10.

An additional DNA template was designed in which two T10 tracts flanked the 21 nt T7 promoter sequence, with the notion that a triplex stem-loop would be formed that constrained the promoter into the loop region. All templates exhibited a clean gel-shift upon bPNA binding and cooperative thermal transitions (~55 °C) by UV spectroscopy, supporting discrete and specific 1:1 recognition, in line with prior studies on smaller DNA strands (Figure 2.2).

Quantitation of native electrophoretic mobility shift assays (EMSA) of DNA treated with unlabeled bPNA 10 yielded binding curves that fit well to 1:1 binding models.39

As with the dC10 loop, bPNA 10 exhibited ~20 nM binding affinity to the DNA template with

4 nucleotide dCACA spacer in between T-tracts, in the context of a t-RNA-Lys encoding

DNA sequence. Interestingly, despite the potential for higher entropic penalty on association, the DNA template with the 21 nt T7 promoter sequence flanked by T10 tracts exhibited a tighter binding to 10 (Kd = 4 nM), suggesting strain in the shorter loops or the existence of stabilizing interactions within the stem- loop folded promoter sequence.

Affinity for 10 in all template designs, including the promoter stem-loop design, was unaffected by whether or not the promoter region was single or double- stranded, though it is unclear whether or not bPNA binding resulted in duplex dissociation.

Having established that longer native-like DNA templates exhibited similar bPNA binding affinity and specificity as previously found with shorter substrates, we considered bPNA

10 as an artificial transcriptional repressor. The effect of bPNA targeting on transcription was studied using the same DNA substrate templates and a T7 RNA polymerase run-off transcription assay. In the absence of 10, run-off transcription yielded full-length RNA transcripts as expected. Prior incubation with bPNA 10 resulted in significant, though wide- ranging, transcriptional repression, as judged by total transcript production (Figure 2.3).

Weakest repression (50%) was observed with the 5’ hairpin triplex, which resulted in 54 production of truncated transcript. Electrophoretic mobility of the truncated product corresponded to aborted transcription at the 5’ hairpin site, indicating that T7 disengages

Figure 2.3. Transcriptional repression by bPNA-DNA triplex. (Top) Illustration of aborted T7 RNA polymerase transcription and initiation upon encountering a bPNA hairpin structure in the middle of the template or sequestering the promoter sequence (black) as a loop. Transcriptional efficiency of T7 as labelled, with 100% efficiency defined as product produced in the bPNA-free control run-off at 4 hours. Corresponding SYBR gold stained gels of run-off transcription product (*) produced from template (<) over time are shown at right with gels A-E representing no bPNA (all templates similar), 5’ hairpin, 3’ hairpin, promoter-loop hairpin and post- promoter hairpin structured templates, respectively, as defined in Figure 2.1. with the template upon encountering the artificial folded structure. Up to 90% repression was observed when the bPNA hairpin site was placed proximal to the promoter sequence and when the promoter sequence was folded into the loop domain of the hairpin, suggesting that transcriptional initiation is especially sensitive to competing bPNA-DNA

55 folded structures. Effective repression from placement of the 21 nt T7 promoter into the loop of the triplex hairpin structure directly demonstrates that the synthetic hairpin structure can prevent T7 binding and transcriptional initiation, and underscores the stability of DNA hairpin triplexes with 4 to 21 nt loop domains.

Transcriptions on control DNA templates with bPNA addition showed no inhibitory effect,

(Figure 2.4) further proves the observed transcription inhibition on T10 bearing template is originated from the bPNA-DNA triplex formation.

Figure 2.4. Control transcriptions. Transcriptional Repression Control with T7 Run-off Control Oligo 1 (Top) and T7 Run-off Control Oligo 2 (Bottom).

2.2.2 Triplexes are Resistant Towards Exonuclease T Digestion

The ability of bPNA folded DNA and RNA structures to limit enzymatic access suggested resistance to nuclease degradation. We tested the stability of both RNA-bPNA and DNA- bPNA hairpin complexes towards exonuclease degradation. It was not possible to eliminate background degradation of long RNA substrates, so our RNA studies were

56 limited to bPNA complexes of rU10C10U10. As expected, discrete gel-shifts on binding bPNAs 8 and 10 were observed with rU10C10U10 (Figure 2.5). While the free RNA substrate was rapidly consumed by exonuclease T (exo T) in less than one minute, the hairpin complexes remained detectable for over 4 hours, as judged by the quantity of free bPNA observed under saturating conditions. Gel quantification revealed that while ~40% of bPNA 8 was released over 4 hours, negligible quantities (<3%) of bPNA 10 from their respective RNA complexes upon exo T treatment. This difference is likely reflective of the greater stability afforded by a longer bPNA-RNA binding interface. Longer DNA substrates were studied with bPNA 10 and dT10C4T10 targeting sites placed in middle, 3’ and 5’ termini.

Following formation of triplex stem-loop structures, these complexes were exposed to exonuclease T. Exonuclease T operates in the 3’ to 5’ direction, and thus hairpin structures placed at the 3’ end of a single-stranded RNA or DNA were largely resistant to exo T digestion while 5’ hairpin modified oligonucleotides were degraded down to a structure with the electrophoretic mobility of the hairpin structure alone. DNA or RNA substrates with hairpin triplexes in the middle of the sequence showed digestion was halted at the position of the synthetic secondary structure as well. (Figure 2.5)

Figure 2.5. ExoT resistance of bPNA-DNA and RNA triplexes. (Left, Top) Complete gel shift of carboxyfluorescein labelled bPNA 10 (>) upon treatment with rU10C10U10 to form bPNA-RNA complex (*) that remains detectable over 4 hours of exoT exposure. (Left, Bottom) Gel quantitation of bPNAs 8 and 10 released from their respective complexes with rU10C10U10 over time. (Top, Right) Complete gel shift of carboxyfluorescein labelled bPNA 10 (>) upon treatment with DNA featuring a T10CACAT10 bPNA binding site in the middle of the sequence, to form bPNA-DNA complex (*) indicated above that remains detectable over 4 hours of exoT exposure, following digestion to the hairpin structure indicated. (Bottom, Right) Gel quantitation of exoT degradation of bPNA 10 complexes with DNA featuring 3’ and 5’ bPNA binding sites, as described in Figure 2.1.

The triplex structure dependent digestion resistance was further proved via control experiments on DNA substrate grants no bPNA recognition site. Digestion results indicate no difference between sample with bPNA addition and sample without bPNA addition.

(Figure 2.6)

Figure 2.6. Exonuclease T resistance control.

2.2.3 Triplexes Inhibit AMV Reverse Transcription

Inhibition of RNA-substrate based processes such as reverse transcription was similarly effective. Initial attempts to study ribosome displacement using a toe-print assay on a well- studied RNA sequence40 were unsuccessful as 70S bacterial ribosome binding was sensitive to U-rich bPNA binding sites41 even in the absence of bPNA. We instead utilized the 142 nt mRNA containing a mid-sequence bPNA binding site (rU10CAAAU10) as a substrate for reverse-transcription inhibition. This system could accept bPNA with lengths varying from 10, 8 and 6 melamine- displaying repeat units (Figure 2.1), which yield bPNA-

RNA complexes of decreasing stability as the size of the recognition interface is decreased,

59 similar to what was observed with exo T resistance. The RNA substrate was produced by run-off transcription, purified, annealed with 5’ 32P end-labeled DNA primer, and subjected to reverse-transcription conditions with AMV reverse transcriptase. Each system exhibited a bPNA concentration-dependent suppression of full-length cDNA product. As expected, the most effective suppression of reverse- transcription was shown by the most tightly binding bPNA 10 and weaker, though significant, suppression was found with bPNAs 8 and 6 (Figure 2.7). As the yield of full length cDNA product decreased, increased yield of a truncated cDNA was observed; this truncation product corresponded to halted reverse- transcription at the rU10CAAAU10 site. As the same bPNA binding site was being targeted, the identical truncated DNA reverse- transcript product was observed with bPNAs 6, 8, and 10, suggestive of aborted enzymatic processing upon encountering the bPNA triplex.

Interestingly, full-length cDNA production was sharply decreased to ~10% on treatment with 10, and exhibited no further change at an approximate 1:1 mole ratio to the RNA, as expected based on the molecular recognition interface (Figures 2.1 & 2.7). Similar, but less pronounced, saturation effects were observed with bPNAs 6 and 8, likely due to weaker binding. Further, control reverse transcription showed no inhibitory effect of bPNA towards AMV reverse transcriptase when no bPNA binding sites were displayed on mRNA template. (Figure 2.8)

Figure 2.7. Reverse transcription inhibition via bPNA-RNA triplex. Production of cDNA from reverse transcription of RNA template bearing a bPNA U10CAAAU10 binding site, showing repression as function of bPNA length and mole ratio to RNA. Data obtained by EMSA quantitation, with a representative gel shown at right, where full length cDNA product (*) and truncated cDNA product (<) are indicated. Assays are run at constant RNA substrate concentration (50 nM).

Figure 2.8. AMV reverse transcription control. Control Oligo 2 was used.

2.3 Conclusion

Overall, we find that bPNAs presenting 6, 8 and 10 bifacial nucleobases can selectively target T and U-rich sites on long DNA and RNA substrates of heterogeneous composition.

Nanomolar range bPNA binding affinity provides sufficient hairpin stability to compete with enzymes for their native DNA and RNA substrates, greatly limiting substrate access in the context of transcription, exonuclease digestion and reverse- transcription. These results are remarkable considering the non- native nucleobase employed in bPNA. Though bPNA forms triplex hybrid structures with polypyrimidine nucleic acid much like conventional

PNA, the bifacial triamino triazine bPNA recognition element generates a unique hybrid fold that is the inverse of the PNA triplex; two native nucleic acid strands are templated along a synthetic bPNA strand to form a triplex has no native or artificial cognate. This mode of recognition drives a significant structural reorganization in the nucleic acid, transforming unstructured single-stranded nucleic acids into well-folded triplex hairpin loops. Our studies demonstrate that structure nucleation42-46 upon formation of bPNA- nucleic acid hybrids may be used to limit substrate access and thereby modulate the function of enzymes involved in nucleic acid transcription, reverse transcription and degradation, and suggests the utility of bPNA as a tool for biotechnology.

2.4 Experimental

2.4.1 Material

Chemicals for amino acid derivative synthesis, solid phase peptide synthesis, purification and characterization were purchased from Sigma-Aldrich, Chem-Impex and AAPPTec without further purification unless otherwise specified. Rink Resin LS (100-200 mesh, 0.28 mmol/g) was purchased from Advanced ChemTech

All DNA oligonucleotides were purchased from Integrated DNA Technology (IDT). DNA concentrations were measured at 260 nm using extinction coefficient provided by IDT. For

DNA oligonucleotides with significant secondary structure at room temperature, the concentrations were measured at 50 °C in order to make sure there was no interference from the secondary structure.

T7 RNA polymerase, Exonuclease T, AMV reverse transcriptase and SYBR gold stain were purchased from Invitrogen. Binding experiments and transcriptional repression experiments were performed in 1x RNA polymerase reaction buffer (NEB), supplemented with 10 mM DTT and 15mM Mg2+.

Peptides were synthesized as previously reported using standard Fmoc solid phase peptide synthesis.1 Fluorescently-labeled bPNAs were prepared by N-termination with carboxyfluorescein on a β-alanine residue. All bPNAs were purified to homogeneity on reversed phase HPLC and their identity confirmed by mass spectroscopy.

2.4.2 Nucleic Acid Sequences

The DNA sequence encoding tRNA-Lys was used in transcriptional repression and exonuclease resistance assays, T7 RNA polymerase promoter sequence underlined and hairpin sequence shown in bold red:

5’ hairpin 5’ - TTT TTT TTT TCA CAT TTT TTT TTT CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GGT TAA TGC TAA GTT AGC TTT ACA GTG GGC CCC TAT AGT GAG TCG TAT TAA TTT C - 3’ mid hairpin 5’ - CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GTT TTT TTT TTC ACA TTT TTT TTT TGT TAA TGC TAA GTT AGC TTT ACA GTG GGC CCC TAT AGT GAG TCG TAT TAA TTT C - 3’ pp hairpin 5’-CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GGT TAA TGC TAA GTT AGC TTT ACA GTG GGG CCC TTT TTT TTT TCA CAT TTT TTT TTT TAT AGT GAG TCG TAT TAA TTT C-3’

3’ hairpin 5’-CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GGT TAA TGC TAA GTT AGC TTT ACA GTG GGC CCC TAT AGT GAG TCG TAT TAA TTT CTT TTT TTT TTC ACA TTT TTT TTT T-3’

Hairpin promoter loop 5’-CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GGT TAA TGC TAA GTT AGC TTT ACA GTG GGC CCT TTT TTT TTT CCT ATA GTG AGT CGT ATT AAT TTT TTT TTT-3’

T7 Run-off control oligo 1: 5’- GAC GCG ACT AGT TAC GGA GCT CAC ACT CTA CTC AAC AGC TGC ACT GCC GAA GCA GCC ACA CCT GGA CCC GTC CTT CAC CAT TTC ATT CAG TTG CGT CTA TAG TGA GTC GTA TTA ATT TC -3’

T7 Run-off control oligo 2 5’- AGT TCT GGT ACT CGC ACT CTC GTA AAC GAT CAC TGT AAA GAG GTG TTG GTT CTC TTA ATC TTT AAC TTA AAA GGT TAA TGC TAA GTT AGC TTT ACA GTG GGC CCC TAT AGT GAG TCG TAT TAA TTT C -3’

AMV reverse transcription primer 5’- AGC TCT GGT ACT CGC ACT CTC GTA AAC G-3’ mRNA sequence used in RT repression studies, primer binding site underlined: 5’-GGU AAA GUG UCA UAG CAC CAA CUG UUA AUU AAA UUA AAU UAA AAA GGA AAU AAA AAU GUU UUU UUU UUC AAA UUU UUU UUU UAA CUC GCU GCA CAA AUG GCU AAA CUG AAU GGG AAU AAA GGU UUU UCU UUC UGA AGA UAA A-3’.

Anisotropy binding studies were carried out with a 5’ bPNA binding site featuring a 10 base loop instead (T10(CA)5T10), while a control sequence replaced this T-rich sequence with AGC TCT GGT ACT CGC ACT CTC GTA AAC GAT.

2.4.3 Equipment

MALDI-TOF Mass spectra were acquired on Bruker Microflex MALDI-TOF instrument.

Electrospray mass spectroscopy was acquired on Bruker MicroTOF equipped with an electrospray ionization source. Mass spectrometry instruments were provided by a grant from the Ohio BioProducts Innovation Center. NMR spectra were acquired on Bruker

Advance DPX 400 instrument. Solid phase peptide synthesis was performed on AATTPec

Apex 396 peptide synthesizer. All peptides were purified on RP-C18 preparative HPLC column and the purity was confirmed on RP-C18 analytical column. UV melting was performed on Varian Cary-100 UV-Vis Spectrophotometer equipped with Cary

Temperature Controller. Fluorescence anisotropy was performed on Molecular Devices

SpectraMax M5. Circular Dichroism spectra were obtained on Jasco J815 Circular

Dichroism Spectrometer. Fluorescence gel images were acquired using a Typhoon Trio

Variable Mode Imager (Amersham Biosciences).

2.4.4 Procedures

2.4.4.1 Amino Acid Derivative Synthesis

Fmoc-Lys(Melamine)-OH was synthesized as previously reported1.

2.4.4.2 Solid Phase Peptide Synthesis and Characterization

Standard Fmoc chemistry was employed. DIC/HOBt were used as the coupling reagents.

And 20% piperidine in NMP was used for Fmoc deprotection. 95% TFA was used to cleave peptide from resin and remove the protecting groups. Resin was removed by filtration through cotton and the peptide was precipitated and washed by cold diethyl ether. Crude peptides were then dissolved in solvent A and purified by HPLC on a C18 reversed phase column using a gradient of 10–50% solvent B in 50 min (solvent A= 0.1% TFA in water, solvent B = 0.01% TFA in 45% acetonitrile, 45% isopropanol, 10% water). The UV detector was set at 238 nm. Purified peptides were lyophilized to dryness.

2.4.4.3 bPNA Binding Analysis

Native electrophoretic mobility shift assays (EMSA) and fluorescence anisotropy assays were performed to determine bPNA-NA binding affinity. DNA concentration was held constant at 100 nM in the EMSA experiments while bPNA concentration was varied from

0 nM to 1600 nM. All native PAGE samples were equilibrated at room temperature for 1hr before analysis. Gels were stained by using SYBR® gold for 20 minutes and scanned with a Typhoon gel scanner (excitation/emission = 488/526 nm) followed by quantification using Image Quant software. For anisotropy experiments, carboxyfluorescein-labeled bPNA concentration was maintained at 60nM while DNA concentration was varied from 0 66 nM to 1440 nM. Samples were equilibrated at room temperature shielded from light for

1hr before analysis with a SpectraMax® M5 plate reader (excitation/emission 492/525 nm).

EMSA and anisotropy data were fit to a 1:1 binding model39 with the general form of

Equation 2.1

Equation 2.1 1 1 [푏푃푁퐴푁퐴] = (퐾 + 푏푃푁퐴 + 푁퐴 ) − √(퐾 + 푏푃푁퐴 + 푁퐴 )2 − 4푏푃푁퐴 푁퐴 2 푑 0 0 2 푑 0 0 0 0

Where Kd, bPNA0 and NA0 represent the dissociation constant, initial bPNA concentration and initial concentration of nucleic acid (NA), respectively. For anisotropy data sets, As replaces NA0 as anisotropy of saturated binding.

2.4.4.4 Transcriptional Repression

Transcription run-off assays were performed using 2μM promoter-duplexed DNA templates with and without 8 equivalents of bPNA, annealed in a 1:1.1 ratio of DNA/T7 promoter (5 minutes at 95 °C, cooled slowly to room temperature in 1mM EDTA, 10mM

Tris-Cl and 50mM NaCl). After 1hr equilibration, 3.75 unit/μL T7 RNA polymerase was added, followed by addition of 4mM of each NTP and incubation at 37 °C. Reaction aliquots were quenched in 200mM EDTA at 3, 8, 15, 30, 60 and 90 minute time points, analyzed on 3% EtBr/agarose gel and visualized by UV.

2.4.4.5 T7 Run-off Transcription Control

Control Oligos (see Materials section for sequences) were mixed with T7 Promoter at 1:1.1 ratio, the mixtures were heated up to 95°C for 5min, then gradually cooled down to room temperature. Samples with and without 8 equivalents of bPNA 10 were then mixed with

3.75unit/μL T7 RNA polymerase, followed by addition of 4mM of each NTP and incubation at 37°C. Reaction aliquots were quenched in 200mM EDTA at 3, 8, 15, 30, 60 and 90 67 minute time points, analyzed on 3% EtBr/agarose gel and visualized by UV.

2.4.4.6 Exonuclease T Resistance

All experiments were conducted in 1x NEBuffer 4 (50mM Potassium Acetate, 20mM Tris-

Acetate, 10mM Magnesium Acetate, 1mM DTT, pH7.9), supplemented with 10% glycerol.

Saturated binding conditions of bPNA to DNA and RNA substrates were determined by an EMSA, titrating 100 nM carboxyfluorescein (Cbf) labeled bPNA against DNA or RNA.

Exonuclease T (20 units, NEB) was added into a 150uL reaction solution under conditions in which complete bPNA complexation was observed (100nM bPNA to 200 nM DNA/RNA) and incubated at 37°C for 4 hours. Reaction aliquots were quenched with 250mM EDTA at various time points and on native PAGE gel (RNA) or native gradient PAGE (DNA), and quantified using SYBR gold stain.

2.4.4.7 Exonuclease T digestion control

All experiments were conducted in 1x NEBuffer 4 (50mM Potassium Acetate, 20mM Tris-

Acetate, 10mM Magnesium Acetate, 1mM DTT, pH7.9), supplemented with 10% glycerol.

8 equivalents of bPNA 10 were added to control DNA, followed by the addition of

Exonuclease T (20 units, NEB) into a 150uL reaction solution. Reaction aliquots were quenched with 250mM EDTA at various time points and on native PAGE gel (RNA) or native gradient PAGE (DNA), and quantified using SYBR gold stain.

2.4.4.8 Inhibition of reverse transcription

Reverse transcription primers (100 pmole) were end-labeled by mixing with T4 polynucleotide kinase (50 units) and ATP γ-32P in 1x T4 PNK buffer (70mM Tris-Cl, 10mM

MgCl2, 5mM DTT, pH 7.6) at 25 °C, followed by incubation at 37 °C for 30 minutes and enzyme inactivation at 65 °C for 20 minutes. Labeled DNA primer was purified using a

Sephadex G15 column and radioactivity was diluted to 200,000 CPM. RNA substrate was prepared using standard T7 run-off transcription and purification methods

(MEGAshortscript T7 and MEGAclear kits, Invitrogen) from the appropriate DNA template annealed to the 17 nt sense promoter sequence. Purified RNA (50 nM) was incubated with excess radiolabeled RT primer and varying concentrations of bPNAs 6, 8 or 10.

Following heat denaturation (65 °C, 10 minutes) and ice-cooling, reaction was initiated by addition of AMV reverse transcriptase (RT) to a final concentration of 0.4 unit/μL.

Reactions were incubated at 37 °C for 30 minutes before being quenched with EDTA- formamide dye. Reactions were analyzed using denaturing PAGE (6%, 7 M urea) after drying, overnight exposure to a phosphor image plate and Typhoon gel scanner.

2.4.4.9 Reverse transcription control

RNA template was transcribed from T7 Run-off Control Oligo 2 and further purified.

Purified RNA (1 μM) was incubated with excess RT primer and 8 equivalents of bPNA 10.

Following heat denaturation (65 °C, 10 minutes) and ice-cooling, reaction was initiated by addition of AMV reverse transcriptase (RT) to a final concentration of 0.4 unit/μL.

Reactions were incubated at 37 °C and aliquots at different time points were quenched with EDTA-Urea dye. Reactions were analyzed using denaturing gradient PAGE (4% -

20%, 7 M urea). Gels were stained with SYBR®Gold.

2.5 Additional Data

2.5.1 Amino Acid Derivative

Figure 2.9. 1H NMR spectra of Fmoc-Lys-Melamine monomer. 1H NMR (400 MHz, d6-DMSO) δ (ppm) 1.35 (d, J=2.8Hz, 2H), 1.50 (t, J=5.4Hz, 2H), 1.62-1.71 (m, 2H), 3.25 (d, J=2.4Hz, 2H), 3.93 (m, 1H), 4.22 (t, J=5.4Hz, 1H), 4.27-4.30 (m, 2H), 7.31 (t, J=5.8Hz, 2H), 7.40 (t, J=6Hz, 2H), 7.63 (d, J = 8Hz, 1H), 7.72 (m, 2H), 7.88 (m, 6H), 8.09 (s, 1H).

Figure 2.10. 13C NMR spectrum of Fmoc-Lys-Melamine monomer.13C NMR (100 MHz, d6- DMSO) δ (ppm) 23.48, 28.80, 30.95, 47.18, 54.30, 66.10, 120.61, 125.78, 127.56, 128.15, 141.23, 144.34, 156.69, 158.96, 174.45. Calculated Mass [M+H]: 478.2205. Found: 478.2382.

2.7.2 HPLC and MALDI-TOF Mass Spectra of the Synthetic Peptides

xp1196-1 0:H11 MS Raw a 1000

Intens. [a.u.] Intens. b

3 10 5 800

2.5 10 5

5 2 10 600 2275.583

1.5 10 5

400 1 10 5

5 10 4

200 0

-5 10 4 0 10 20 30 40 50 0 1500 2000 2500 3000 3500 m /z Time (min)

Figure 2.11. HPLC and MALDI of H2N-(EM*)6G (a) HPLC trace of H2N-(EM*)6G (6) on a RP- C18 analytical column using a gradient of 10-50% solvent B over 50 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of 6: Calculated [M+H]: 2273.353, Found: 2275.583.

EM 8 0:C5 MS Raw b a [a.u.] Intens.

1500 2.5 10 5

2 10 5

3006.812 1.5 10 5 1000

1 10 5

5 10 4 500

-5 10 4 0 10 20 30 40 50 0 1500 2000 2500 3000 3500 Time (min) m /z

Figure 2.12. HPLC and MALDI of H2N-(EM*)8G (a) HPLC trace of H2N-(EM*)8G (8) on a RP- C18 analytical column using a gradient of 10-50% solvent B over 46 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of (8): Calculated [M+H]: 3006.107, Found: 3006.812.

a b Intens. [a.u.] Intens. 5 7 10 2500

6 10 5

2000 5 10 5

4 10 5 1500 3739.541

3 10 5

1000 2 10 5

1 10 5 500 1869.836 0

5 0 -1 10 1500 2000 2500 3000 3500 4000 4500 m /z 0 10 20 30 40 50 Time (min)

Figure 2.13. HPLC and MALDI of H2N-(EM*)10G (a) HPLC trace of H2N-(EM*)10G (10) on a RP- C18 analytical column using a gradient of 10-50% solvent B over 50 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of (10): Calculated [M+H]: 3738.862, Found: 3739.541.

E M 6\0_C 6\2\1sref a b [a.u.] Intens. b

6000

4 10 5

3.5 10 5 4000

3 10 5

5 2.5 10 2702.209

2 10 5

5 1.5 10 2000

1 10 5

5 10 4

0 0 10 20 30 40 50 1000 1500 2000 2500 3000 3500 4000 4500 m /z Time (min)

Figure 2.14. HPLC and MALDI of Cbf-(β-A)-(EM*)6G (Cbf-6) (a) HPLC trace of Cbf-(β-A)- (EM*)6G (Cbf-6) on a RP-C18 analytical column using a gradient of 10-50% solvent B over 47 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of (Cbf-6): Calculated [M+H]: 2702.728, Found: 2702.209.

* EM 8\0_C 8\2\1sref 5000 a b [a.u.] Intens.

4000 2 10 5

1.5 10 5 3000

3436.586 1 10 5 2000

5 10 4

1000

0 10 20 30 40 50 0 1000 1500 2000 2500 3000 3500 4000 4500 Time (min) m /z

Figure 2.15. HPLC and MALDI of Cbf-(β-A)-(EM*)8G (Cbf-8) (a) HPLC trace of Cbf-(β-A)- (EM*)8G (Cbf-8) on a RP-C18 analytical column using a gradient of 10-50% solvent B over 50 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of (Cbf-8): Calculated [M+H]: 3435.480, Found: 3436.586.

a b * EM 10\0_C 10\4\1SR ef

Intens. [a.u.] 5000Intens.

4000

4 10 5

5 3.5 10 3000

5 3 10 4167.691

2.5 10 5 2000 2 10 5

1.5 10 5

5 1 10 1000

5 10 4

0 0 1000 1500 2000 2500 3000 3500 4000 4500 0 10 20 30 40 50 m /z Time (min) Figure 2.16. HPLC and MALDI of Cbf-(β-A)-(EM*)10G (Cbf-10) (a) HPLC trace of Cbf-(β-A)- (EM*)10G (Cbf-10) on a RP-C18 analytical column using a gradient of 10-50% solvent B over 50 min, monitored by a UV-Vis detector at 238 nm. (b) MALDI-TOF Mass of (Cbf-10): Calculated [M+H]: 4168.232 Found: 4167.691.

2.6 Acknowledgements

We thank H. Zhu for technical assistance. This work was supported in part by NIH grant GM072528 (K.F.). 74

2.7 References for Chapter 2

1. Zeng, Y.; Pratumyot, Y.; Piao, X.; Bong, D., Discrete assembly of synthetic

peptide-DNA triplex structures from polyvalent melamine-thymine bifacial

recognition. J. Am. Chem. Soc. 2012, 134, 832-5.

2. Piao, X.; Xia, X.; Bong, D., Bifacial Peptide Nucleic Acid Directs Cooperative

Folding and Assembly of Binary, Ternary, and Quaternary DNA Complexes.

Biochemistry 2013, 52, 6313-6323.

3. Højfeldt, J. W.; Van Dyke, A. R.; Mapp, A. K., Transforming ligands into

transcriptional regulators: building blocks for bifunctional molecules. Chemical

Society Reviews 2011, 40, 4286-4294.

4. Zhang, J.; Lau, M. W.; Ferre-D'Amare, A. R., Ribozymes and riboswitches:

modulation of RNA function by small molecules. Biochemistry 2010, 49, 9123-31.

5. Henkin, T. M., Riboswitch RNAs: using RNA to sense cellular metabolism. Genes

& Development 2008, 22, 3383-3390.

6. Wilson, K. S.; von Hippel, P. H., Transcription termination at intrinsic terminators:

the role of the RNA hairpin. Proceedings of the National Academy of Sciences

1995, 92, 8793-8797.

7. Greive, S. J.; von Hippel, P. H., Thinking quantitatively about transcriptional

regulation. Nat Rev Mol Cell Biol 2005, 6, 221-32.

8. Nielsen, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O., Sequence-selective

recognition of DNA by strand displacement with a thymine-substituted polyamide.

Science 1991, 254, 1497-1500.

9. Kumar, V. A.; Ganesh, K. N., Conformationally constrained PNA analogues:

structural evolution toward DNA/RNA binding selectivity. Acc Chem Res 2005, 38,

404-12.

10. Nielsen, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O., Sequence specific inhibition

of DNA restriction enzyme cleavage by PNA. Nucleic Acids Research 1993, 21,

197-200.

11. Kaihatsu, K.; Shah, R. H.; Zhao, X.; Corey, D. R., Extending Recognition by

Peptide Nucleic Acids (PNAs): Binding to Duplex DNA and Inhibition of

Transcription by Tail-Clamp PNA-Peptide Conjugates. Biochemistry 2003, 42,

13996-14003.

12. Bentin, T.; Larsen, H. J.; Nielsen, P. E., Combined Triplex/Duplex Invasion of

Double-Stranded DNA by "Tail-Clamp" Peptide Nucleic Acid. Biochemistry 2003,

42, 13987-13995.

13. Belotserkovskii, B. P.; Liu, R.; Hanawalt, P. C., Peptide nucleic acid (PNA) binding

and its effect on in vitro transcription in friedreich's ataxia triplet repeats. Molecular

carcinogenesis 2009, 48, 299-308.

14. Rougee, M.; Faucon, B.; Mergny, J. L.; Barcelo, F.; Giovannangeli, C.; Garestier,

T.; Hélène, C., Kinetics and thermodynamics of triple-helix formation: effects of

ionic strength and mismatches. Biochemistry 1992, 31, 9269-78.

15. Felsenfeld, G.; Davies, D. R.; Rich, A., Formation of a three-stranded

polynucleotide molecule. J. Am. Chem. Soc. 1957, 79, 2023-2024.

16. Datta, B.; Schmitt, C.; Armitage, B. A., Formation of a PNA2-DNA2 hybrid

quadruplex. Journal of the American Chemical Society 2003, 125, 4111-4118.

17. Datta, B.; Bier, M. E.; Roy, S.; Armitage, B. A., Quadruplex Formation by a

Guanine-Rich PNA Oligomer. Journal of the American Chemical Society 2005, 127,

4199-4207.

18. Gupta, A.; Lee, L.-L.; Roy, S.; Tanious, F. A.; Wilson, W. D.; Ly, D. H.; Armitage,

B. A., Strand Invasion of DNA Quadruplexes by PNA: Comparison of Homologous

and Complementary Hybridization. Chembiochem 2013, 14, 1476-1484.

19. Branda, N.; Kurz, G.; Lehn, J.-M., JANUS WEDGES: a new approach towards

nucleobase-pair recognition. Chem. Commun. 1996, 1996, 2443-2444.

20. Chen, H.; Meena; McLaughlin, L. W., A Janus-Wedge DNA Triplex with A-W1-T

and G-W2-C Base Triplets. J. Am. Chem. Soc. 2008, 130, 13190-13191.

21. Shin, D.; Tor, Y., Bifacial Nucleoside as a Surrogate for Both T and A in Duplex

DNA. J. Am. Chem. Soc. 2011, 133, 6926-6929.

22. Vysabhattar, R.; Ganesh, K. N., Cyanuryl peptide nucleic acid: synthesis and DNA

complexation properties. Tetrahedron Lett. 2008, 49, 1314-1318.

23. Arambula, J. F.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C., A simple

ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein

binding. Proc. Natl. Acad. Sci. USA 2009, 106, 16068-16073.

24. Mittapalli, G. K.; Osornio, Y. M.; Guerrero, M. A.; Reddy, K. R.; Krishnamurthy, R.;

Eschenmoser, A., Mapping the landscape of potentially primordial informational

oligomers: oligodipeptides tagged with 2,4-disubstituted 5-aminopyrimidines as

recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2478-84.

25. Mittapalli, G. K.; Reddy, K. R.; Xiong, H.; Munoz, O.; Han, B.; De Riccardis, F.;

Krishnamurthy, R.; Eschenmoser, A., Mapping the landscape of potentially

primordial informational oligomers: oligodipeptides and oligodipeptoids tagged

with triazines as recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2470-7. 77

26. De Greef, T. F. A.; Smulders, M. M. J.; Wolffs, M.; Schenning, A.; Sijbesma, R. P.;

Meijer, E. W., Supramolecular Polymerization. Chemical Reviews 2009, 109,

5687-5754.

27. Whitesides, G. M.; Simanek, E. E.; Mathias, J. P.; Seto, C. T.; Chin, D.; Mammen,

M.; Gordon, D. M., Noncovalent Synthesis: Using Physical-Organic Chemistry To

Make Aggregates. Acc. Chem. Res. 1995, 28, 37-44.

28. Prins, L. J.; Reinhoudt, D. N.; Timmerman, P., Noncovalent synthesis using

hydrogen bonding. Angew. Chem. Int. Ed. 2001, 40, 2382-2426.

29. ten Cate, M. G. J.; Huskens, J.; Crego-Calama, M.; Reinhoudt, D. N.,

Thermodynamic stability of hydrogen-bonded nanostructures: A calorimetric study.

Chem.--Eur. J. 2004, 10, 3632-3639.

30. Oshovsky, G. V.; Reinhoudt, D. N.; Verboom, W., Supramolecular chemistry in

water. Angew. Chem. Int. Ed. 2007, 46, 2366-2393.

31. Ma, M.; Paredes, A.; Bong, D., Intra- and intermembrane pairwise molecular

recognition between synthetic hydrogen-bonding phospholipids. J. Am. Chem. Soc.

2008, 130, 14456-8.

32. Ma, M.; Gong, Y.; Bong, D., Lipid Membrane Adhesion and Fusion Driven by

Designed, Minimally Multivalent Hydrogen-Bonding Lipids. J. Am. Chem. Soc.

2009, 131, 16919-16926.

33. Ma, M.; Bong, D., Protein assembly directed by synthetic molecular recognition

motifs. Org. & Biomolec. Chem. 2011, 9, 7296-9.

34. Ma, M.; Bong, D., Determinants of cyanuric acid and melamine assembly in water.

Langmuir 2011, 27, 8841-53.

35. Ma, M.; Bong, D., Directed peptide assembly at the lipid-water interface

cooperatively enhances membrane binding and activity. Langmuir 2011, 27, 1480-

36. Zhou, Z.; Bong, D., Small-Molecule/Polymer Recognition Triggers Aqueous-Phase

Assembly and Encapsulation. Langmuir 2013, 29, 144-150.

37. Soto, A. M.; Loo, J.; Marky, L. A., Energetic contributions for the formation of

TAT/TAT, TAT/CGC(+), and CGC(+)/CGC(+) base triplet stacks. J. Am. Chem.

Soc. 2002, 124, 14355-14363.

38. SantaLucia, J., Jr.; Hicks, D., The thermodynamics of DNA structural motifs. Annu.

Rev. Biophys. Biomol. Struct. 2004, 33, 415-40.

39. Agou, F.; Raveh, S.; Mesnildrey, S.; Veron, M., Single strand DNA specificity

analysis of human nucleoside diphosphate kinase B. J Biol Chem 1999, 274,

19630-8.

40. Shoji, S.; Walker, S. E.; Fredrick, K., Reverse translocation of tRNA in the ribosome.

Molecular cell 2006, 24, 931-942.

41. Pestka, S., Studies on the Formation of Transfer Ribonucleic Acid-Ribosome

Complexes VI. OLIGOPEPTIDE SYNTHESIS AND TRANSLOCATION ON

RIBOSOMES IN THE PRESENCE AND ABSENCE OF SOLUBLE TRANSFER

FACTORS. Journal of Biological Chemistry 1969, 244, 1533-1539.

42. Ghadiri, M. R.; Choi, C., Secondary structure nucleation in peptides. Transition

metal ion stabilized .alpha.-helices. J. Am. Chem. Soc. 1990, 112, 1630-2.

43. Ghadiri, M. R.; Fernholz, A. K., Peptide architecture. Design of stable .alpha.-

helical metallopeptides via a novel exchange-inert ruthenium(III) complex. Journal

of the American Chemical Society 1990, 112, 9633-5.

44. Torres, O.; Yuksel, D.; Bernardina, M.; Kumar, K.; Bong, D., Peptide tertiary

structure nucleation by side-chain crosslinking with metal complexation and double

"click" cycloaddition. ChemBioChem 2008, 9, 1701-5.

45. Schafmeister, C. E.; Po, J.; Verdine, G. L., An All-Hydrocarbon Cross-Linking

System for Enhancing the Helicity and Metabolic Stability of Peptides. Journal of

the American Chemical Society 2000, 122, 5891-5892.

46. Chapman, R. N.; Dimartino, G.; Arora, P. S., A Highly Stable Short alpha -Helix

Constrained by a Main-Chain Hydrogen-Bond Surrogate. Journal of the American

Chemical Society 2004, 126, 12252-12253.

CHAPTER 3

bPNA as Allosteric Switch for

Aptamer and Ribozyme Function

This chapter is reproduced with permission from J. Am. Chem. Soc., 2014, 136, 7265-7268.

3.1 Overview

We demonstrate herein that bifacial peptide nucleic acid (bPNA) hybrid triplexes functionally substitute for duplex DNA or RNA. Structure−function loss in three non-coding nucleic acids was inflicted by replacement of a duplex stem with unstructured oligo-T/U strands, which are bPNA binding sites. Functional rescue was observed on refolding of the oligo-T/U strands into bPNA triplex hybrid stems. Bifacial PNA binding was thus used to allosterically switch-on protein and small- molecule binding in DNA and RNA aptamers, as well as catalytic bond scission in a ribozyme. IgE aptamer showed aborted protein binding activity after the stem replacement, upon triplexation via bPNA IgE binding was recovered with no observable activity difference compare to the native system.

Fluorogenic activation of DFHBI small molecule via spinach RNA aptamer folding was lost via P-II stem oligo-U substitution, bPNA triggered triplexation was able to re-fold aptamer structure and restore 50% of its fluorogenic activity under GFP filter. Duplex stems that support the catalytic site of minimal type I hammerhead ribozyme were replaced with oligo-

U loops, severely crippling or ablating the native RNA cleavage function. Refolding of the

U-loop into bPNA triplex stems completely restored the cleavage function in the hybrid system. These studies indicate that bPNA may have general utility as an allosteric trigger for a wide range of functions in non-coding nucleic acids.

3.2 Introduction

We recently reported a new class of bifacial1-3 α-peptide nucleic acids (bPNAs),4-6 derived from studies on artificial recognition.7, 8 Bifacial PNA utilizes a synthetic triazine9-12 base with two identical hydrogen-bonding faces available. This enables bPNA to simultaneously dock13-17 two oligo-thymine (Figure 3.1) with low nanomolar affinity.4, 5 Two T/U-rich tracts separated by 4−25 nucleotides of random sequence may thus be folded into triplex stem−loop (hairpin) structures upon binding to a single bPNA strand. These unique bPNA hybrid structures are stable enough to block enzymatic access to DNA and RNA substrates.6 Bifacial PNA is distinct from conventional PNAs18, 19 in terms of its α-peptide backbone10, 20-23 as well as its use of a non-native triazine recognition interface.9, 10, 13, 24, 25

While PNA triplex26, 27 hybrids are formed from sequestration of native oligonucleotides with two strands of PNA, bPNA serves as a template to assemble two native oligonucleotides. Generally, PNA targeting has been applied as a dissociative operation, wherein native duplex28-32 or quadruplex33-35 folds are disrupted in favor of PNA hybrid structures. In contrast, bPNA targeting is associative, uniting two non-interacting strands on a bPNA bridge. We hypothesized that bPNA-driven association could be used as a synthetic allosteric switch36 for non-coding nucleic acid function.37, 38 To test this notion, known aptamer and ribozyme sequences were functionally crippled by replacement of a critical duplex element with T/U tracts. We predicted that triplex hybridization of the T/U tracts with bPNA would mimic the native duplex and tighten the overall fold to restore function in an allosterically coupled domain. Three systems were chosen to represent a breadth of function: molecular recognition of protein, molecular recognition of small molecule, and chemical catalysis.

Figure 3.1. bPNA and nucleic acid designs. (Top) Structure of bPNAs studied, with n = 6, 8, and 10 corresponding to 6, 8, and 10, showing the recognition interface between bPNA nucleobase melamine (M) and thymine/uracil. (Bottom) Aptamer and ribozyme sequences engineered with bPNA binding sites (red) are indicated with predicted fold based on the original sequence. The conserved catalytic site of the hammerhead ribozyme is shown, and the scissile bond is indicated with a red arrow; t-RNA Lys attached to the 3′ end is not shown.

3.3 Results and Discussion

3.3.1 Rescue of Aptamer-IgE Binding via Triplex Formation

A protein-binding DNA aptamer (D17.4) has been derived via SELEX that binds immunoglobulin E (IgE) with 7 nM affinity.39 This compact 60-nt stem−loop aptamer is an ideal framework for bPNA structure−function manipulation: the 25-nt loop forms the primary molecular recognition module, which is held in place by a sequence-mutable duplex stem. The stem structure was ablated by replacement with two T10 tracts to yield

IgE-T1 (Figure 3.1). As expected, this mutant did not exhibit a thermal melting transition and was retained on nitrocellulose filters, indicative of an unfolded DNA structure. Binding to IgE was evaluated using a nitrocellulose filter-binding assay after DNA radiolabeling.39

As reported, analysis of D17.4 binding to IgE yielded a Kd value of 7 nM, while a null binding result was observed for the unstructured IgE-T1. Stem-ablated DNA IgE-T1 was treated with bPNA 10, which was anticipated to serve as a folding “splint” to structure the terminal T-tracts into a triplex hybrid stem4, 5 and restore the IgE recognition interface

(Figure 3.2). Indeed, addition of bPNA 10 to IgE-T1 resulted in clean complexation to a single product, as judged by electrophoretic mobility shift assay (EMSA) and nominal retention on nitrocellulose filters. However, IgE association by the complex 10·IgE-T1 was extremely weak. Inspection of the original D17.4 sequence suggested that predicted wobble base- pair interactions within the recognition loop may have been disrupted due to register-shifted bPNA complexation resulting from asymmetric T-tract lengths at the 3′

(T10) and 5′ (T13) termini (Figure 3.1). Stem replacement was redesigned as sequence IgE-

T2, which punctuated the 5′ T13 tract with two CG/GC base-pairs found in the original

D17.4 sequence (Figure 3.1).

Figure 3.2. Rescue of aptamer−protein binding. (Left) Binding of known IgE (blue) DNA aptamer D17.4 is shown with predicted fold and wobble base-pairs shown in purple. (Right) Illustration of bPNA refolding of IgE-T2 into a functionally identical bPNA-DNA complex that binds IgE.

This new DNA sequence, IgE-T2, was itself also unfolded and ineffective in binding IgE.

Complexation with bPNA 10 indicated sub-nanomolar (Kd = 0.2 nM) affinity (Figure 3.3), an order of magnitude tighter than previously found T-tract hairpin loop complexes.4, 5

Figure 3.3. Binding profiles of IgE-T2. Binding isotherms of (left) IgE-T2 binding to 10 and [IgE- T2·10] complex binding to IgE (right) obtained by gel shift (top, left) and filter-binding assays. Free IgE-T2 and the [IgE-T2·10] complex are indicated on the gel by * and >, respectively. Binding data were obtained in triplicate and fitted to a 1:1 binding model to yield the dissociation constants indicated.

Importantly, the redesigned 10·IgE-T2 triplex stem−loop structure exhibited low nanomolar affinity (5 nM) to IgE, similar to that reported for D17.4 (7 nM), indicating full recovery of molecular recognition via bPNA triplex stem replacement. Thus, bPNA rescue of DNA aptamer function revealed good tolerance of duplex−triplex replacement, as well as intolerance of a two base-pair shift in triplex site. (Figure 3.4) These findings demonstrate the general utility of duplex replacement as well the precision of bPNA T- tract targeting.

Figure 3.4. Binding onto IgE by D17.4 and IgE-T2. Protran filter images are shown. With the increase in IgE concentration, more radiolabeled aptamer-bPNA 10 complexes were retained on the top filter. Original aptamer showed a 7.4 nM Kd, while bPNA 10 meditated IgE-T2 showed a comparable Kd of 5.6nM.

3.3.2 Rescue RNA Aptamer-Small Molecule Binding and Fluorogenics.

Successful rescue of protein recognition in the relatively simple D17.4/IgE-T2 “stem−loop”

DNA fold prompted further investigation of aptamers with multiple secondary structural elements. Of particular interest was the “Spinach” aptamer, an RNA mimic of green fluorescent protein (GFP) recently disclosed in an elegant series of studies by Jaffrey and co-workers.40 This RNA aptamer binds to a family of small- molecule fluorophores that closely mimic the luminescent nucleus of GFP. (Figure 3.5)

Figure 3.5. Rescue of RNA aptamer−DFHBI binding. Stem II of the Spinach RNA aptamer is subject to replacement with rU10GAGAU10 (red dash). Fluorescence (green) and apparent DFHBI binding are abolished on stem replacement and restored by bPNA triplex stem refolding.

The compound 3,5-difluoro-4-hydroxybenzylidene imidazolinone (DFHBI) is weakly emissive in solution but becomes strongly luminescent when bound by Spinach.

Fluorophore binding (and fluorescence) may be disrupted by insertion of unstructured domains into the Spinach framework, with stem II particularly sensitive to modification.

Jaffrey41 and Hammond42 independently demonstrated how a Spinach fluorescence readout could be coupled to recognition of other substrates by insertion of a second aptamer domain into stem II of Spinach. Conformational tightening of the inserted aptamer fold upon target binding; restores Spinach structure and turns on DFHBI binding and emission.

This design was used to couple bPNA binding to Spinach fluorescence by replacement of stem II with U10 tracts to yield U-Spinach. As expected, loss of stem II completely abolished 88

Spinach fluorescence. Gratifyingly, treatment of U-Spinach with bPNAs 6, 8 or 10 restored

DFHBI binding to approximately 50% of fluorescence intensity. Notably, the spectral features of DFHBI on binding the bPNA-U-Spinach complex were identical to those with unmodified Spinach complex (Figure 3.5).

Figure 3.6. Fluorescence activation via bPNA-U-Spinach complexation. (Left) Excitation (black) and emission (green) of the original Spinach sequence (---) and the U-Spinach·6 complex ( ) bound to DFHBI. (Right) Fluorescence activation curve of DFHBI by the U-Spinach·6 complex, with apparent DFHBI dissociation constant indicated.

Fluorescence activation of DFHBI by U-Spinach as a function of bPNA 6 concentration was measured, yielding an apparent Kd of bPNA-RNA binding of 0.3 μM, though the actual affinity of bPNA to U-Spinach is likely much tighter, given the thermal stability of analogous bPNA-RNA hairpin complexes (Figure 3.6). Similar fluorescence activation profiles were observed for bPNA 8 and bPNA 10, and we speculate them as combined outcomes of bPNA hybridization and DFHBI binding. (Figure 3.7) Fluorescence activation of DFHBI by the RNA·bPNA complexes was comparable to that reported for the original Spinach aptamer (0.7 μM, Figure 3.8), underscoring the functional similarity of a bPNA-triplex to a duplex stem structure. 89

Figure 3.7. Binding of bPNAs to U-Spinach. Electrophoretic mobility shift assays of U-Spinach aptamer complexation with bPNA 6 (Top), 8 (Middle), 10 (Bottom). bPNAs was carboxyfluorescein labeled at N-terminus. Multiple bands of complex may due to different topologies under folding conditions without DFHBI. The apparent Kds of bPNA 6, 8, 10 hybrid onto U-Spinach are 70 nM, 196 nM and 240 nM individually.

Figure 3.8. bPNA dependent fluorescence activation. bPNA 6 (Figure 3.5), bPNA 10 (Top), bPNA 8 (Bottom) was titrated into solution mixtures containing U-Spinach and excess of DFHBI. Fluorescence activation is more responsive to bPNA addition, yielding apparent activation Kd of bPNA 0.3 μM for bPNA 6, 0.49 μM for bPNA 10 and 0.34 μM for bPNA 8, compare to 0.7μM of DFHBI activation affinity for all bPNAs.

Figure 3.9. DFHBI dependent fluorescence activation. All three types of bPNA mediated U- Spinach complexes (Top left: bPNA 6; Top right: bPNA 8; Bottom left: bPNA 10) showed similar affinity towards the DFHBI, and the binding/fluorescence activation affinity is comparable to reported value of 537 nM.

3.3.3 Rescue Hammerhead Ribozyme Cleavage

Aptamer stem-replacement studies indicated the ability of bPNA triplex hybrid stems to support recognition of both protein and small-molecule targets by DNA and RNA aptamer folds. We hypothesized that chemical catalysis, another non- coding nucleic acid function,

92 could be similarly placed under bPNA control, much like an artificial riboswitch.43-46 To this end, a minimal type I hammerhead ribozyme fold47 was sequence- engineered to install bPNA-sensitive catalytic function. The self-splicing ribozyme features a conserved catalytic nucleus supported by three duplex stems (Figure 3.9).

Figure 3.10. U-ribozyme rescue by bPNA. Mutants of the hammerhead ribozyme fold, with replacement of stem III with rU10GAGAU10 to yield U-3 and replacement of stems II and III with rU6GAGAU6 and rU10GAGAU10, respectively, to yield U-(2,3). Splicing function is greatly diminished or ablated on stem replacement (red dash) and may be rescued by triplex refolding with bPNA (blue dash).

Structure−function studies indicate that the duplex stems may be varied in sequence without loss of splicing activity. Stems II and III of the minimal type I hammerhead fold were subject to replacement with rU6CACAU6 and rU10CACAU10 sequences, with single- and double-stem replacements yielding U-3 and U-(2,3), respectively. Stem knock-out “U- mutants” U-3 and U-(2,3) of this ribozyme system (Figure 3.1 & 3.9) were produced as tRNA-Lys fusions by run-off transcription from the appropriate DNA template. While the structurally intact ribozyme was quantitatively spliced during run-off transcription

93 conditions, it was possible to isolate each of the U-ribozyme-tRNA fusions as full-length transcripts in high yield. On splicing, well-folded ribozyme and t-RNA products were readily detectable with minimal degradation by gel was not detectably active in self-splicing at any

Mg2+ concentration. Triplex refolding of the U-loop by addition of bPNA to U-3 resulted in allosteric turn-on of splicing activity, significantly above background (Figure 3.10).

Figure 3.11. U-3 ribozyme activity rescue via bPNAs. Rescue U-3 Ribozyme splicing by bPNA 6, 8, 10. Under 10mM Mg2+ standard condition, U-3 presented self-splicing activity (Top right); however, upon the addition of bPNA 6 (Top right), 8 (Bottom left) and 10 (Bottom, right), slicing rate has been significantly enhanced.

Interestingly, bPNAs 6, 8, and 10 all gave similar rate enhancements for U-3 splicing, restoring cleavage rates to the reported 1 min-1 rate, (Figure 3.11) despite the previously reported length-dependent bPNA affinity.5 This finding supports the notion that bPNA targeting can benefit from cooperative refolding, increasing affinity beyond that observed with unstructured nucleic acids.

Figure 3.12. Kinetics of U-3 ribozyme cleavage rescue. Rescue of self-splicing function by bPNA 6 (), 8 () and 10 (●) in U-3 hammerhead ribozyme. Association of U-3 ribozyme to bPNA results in faster splicing rate comparable to native mini hammerhead ribozyme, which is 1 minute-1 or 0.016 s-1. Splicing rate is 0.019 s-1 with bPNA 6; 0.019 s-1 with bPNA 8, 0.016 s-1 with bPNA 10 and 0.0005 s-1 for U-3 alone.

Further, triplex re-folding of U-3 ribozyme exhibited less Mg2+ dependence required for cleavage. Compare to conventional conditions for cleavage of mini hammerhead type I ribozyme that requires 10 mM Mg2+, bPNAs re-constructed U-3 ribozymes readily

95 exhibited self-cleavage activity even Mg2+ is lower than physiological concentration of 1 mM. (Figure 3.12 and Additional Data)

0.25 mM Mg2+ 0.5 mM Mg2+

with bPNA 6,8,10

U-3 alone U-3 alone

Figure 3.13. Mg2+ dependence of U-3 cleavage. Rescue of self-splicing function by bPNAs 6 (◆), 8 (■), and 10 (●) in U-3 hammerhead ribozymes. U-3 splicing activity observed with bPNA

as indicated from 0.1 mM to 0.8 mM Mg2+, as well as U-3 alone (---).

Targeting and ribozyme activation by bPNA could be accomplished even under the heterogeneous conditions of the run-off transcription reaction, with quantitative in situ splicing of U-3 observed upon addition of bPNA 10 (Figure 3.13).

Though a triple-stem replacement mutant was prepared, it did not exhibit splicing on bPNA complexation. In contrast, splicing of the double-stem knockout U-(2,3) could be completely restored by bPNA to native activity (Figure 3.14).

Figure 3.14. in situ splicing of U-3 via bPNA 10. T7 run-off transcription was performed. Transcription on U-3 only didn’t show noticeable splicing product, while with the addition of bPNA 10, self-cleavage products were observed via gel electrophoresis.

Figure 3.15. U-(2,3) activity restoration via bPNA triplexation. U-(2,3) splicing can be triggered by bPNAs 6, 8, and 10 and also exhibits Mg2+ dependence, with representative data from bPNA 6 shown (left). U-(2,3) splicing was analyzed by urea-denaturing PAGE for bPNAs 6, 8, and 10 at 10 mM MgCl2 (right). Background cleavage of U-(2,3) alone (*) was undetectable, while the tRNA (<) and ribozyme (≪) cleavage products could be clearly observed.

Surprisingly, all bPNAs again elicited identical splicing rates (Figure 3.15) and identical

Mg2+ dependence, (Figure 3.14 and Additional Data) despite the two different sizes of the

U-loop bPNA binding sites in U-(2,3). The smaller stem II site (U6CACAU6) was designed to be addressed exclusively by 6, while the larger stem III site (U10CACAU10) was expected to preferentially bind 10. The identical U-(2,3) splicing rates when complexed to bPNAs 6,

8, and 10 suggest high tolerance for bPNA length mismatching5 at these sites, leading to catalysis with both over-saturation and sub-saturation of the U-U sites by bPNA.

Figure 3.16. Kinetics of U-(2,3) Ribozyme cleavage rescue. Rescue of self-splicing function by bPNA 6 (), 8 () and 10 (●) in U-(2,3) hammerhead ribozyme. Association of U-(2,3) ribozyme to bPNA results in restoration of splicing function. Observed splicing rate is 0.001 s-1 with bPNA 6; 0.0016 s-1 with bPNA 8, 0.001 s-1 with bPNA 10. U-(2,3) only didn’t show any splicing product, the decrease in full length RNA is due to background RNA degradation

3.3 Conclusion

In summary, efficient allosteric control48, 49 of non-coding nucleic acid function by bPNA has been demonstrated in three distinct systems. Protein and small-molecule recognition by DNA and RNA aptamers may be turned on by bPNA triplex stem refolding. Catalysis of self-splicing by a hammerhead ribozyme may also be triggered by bPNA triplex refolding. The breadth of function and fold topology represented by these three nucleic acid systems suggest the possibility that bPNA may provide a general strategy to 99 synthetically switch on nucleic acid function, provided there is a structurally important and mutable duplex stem that could be subject to duplex−triplex hybrid replacement. Nucleic acids designed with a bPNA binding site structurally coupled to a functional nucleus may thus be turned on by a bPNA allosteric switch. It is anticipated that further development of the bPNA-based methodology presented herein will yield broadly useful tools for study and control of nucleic acid function.

3.4 Experimental

3.4.1 Material

MEGAshortscript T7 kit and MEGAclear purification kit for RNA purification were purchased from Invitrogen (Carlsbad, CA). T4 polynucleotide kinase was purchased from

New England Biolabs (Ipswich, MA). Immunoglobulin E human myeloma plasma (IgE) was purchased from Athens Research and Technology (Athens, GA). Oligonucleotides were purchased from Integrated DNA technologies (Coralville, IA). Synthesis reagents for

DFHBI were ordered from Oakwood Products (West Columbia, SC). Bio-Spin 30 columns were purchased from Bio-Rad. Protran nitrocellulose filter was from Whatman, and

Hybond-N+ filters were purchased from GE HealthCare.

3.4.2 DNA Sequences

IgE aptamer 17.4 5’-GGGGCACGTTTATCCGTCCCTCCTAGTGGCGTGCCCC-3’

IgE-T1 5’-TTTTTTTTTTTTCATCCGTACCTTCTAGTGGTTTTTTTTTT-3’

5'-TTTTTTTTTTACGTTTATCCGTCCCTCCTAGTGGCGATTTT IgE-T2 TTTTTT-3’ 5’-GAAATTAATACGACTCACTATAGACGCAACTGAATGAAATG GTGAAGGACGGGTCCAGGTGTGGCTGCTTCGGCAGTGCAGC Spinach GTTGAGTAGAGTGTGAGCTCCGTAACTAGTCGCGTC-3’

100

5’-GAAATTAATACGACTCACTATAGACGCAACTGAATGAAATG GTGAAGGACGGGTCCAGGTTTTTTTTTTCTGCTTCGGCAGTTT U-Spinach TTTTTTTGCTGTTGAGTAGAGTGTGAGCTCCGTAACTAGTCGC GTC-3’

5’-GAAATTAATACGACTCACTATAGGCCCGGATAGCTCAGTCG GTAGAGCAGCGGACGCAACTGAATGAAATGGTGAAGGACGG tRNALys U- GTCCAGGTTTTTTTTTTCTGCTTCGGCAGTTTTTTTTTTGCTGT Spinach TGAGTAGAGTGTGAGCTCCGTAACTAGTCGCGTCCGCGGGTC CAGGGTTCAAGTCCCTGTTCGGGCGCCA-3’

5’-GAAATTAATACGACTCACTATAGGCAGTGCCTGATGAGGCC AATGGGCCGAAACTTTTTTTTTTGAGATTTTTTTTTTGTCGCAC U-3 Ribozyme TGTAAAGCTAACTTAGCATTAACCTTTTAAGTTAAAGATTAAGA GAACCAACACCTCTTTACAGTGA-3’

5’-GAAATTAATACGACTCACTATAGGCAGTGCCTGATGAGTTT TTTGAGATTTTTTCGAAACTTTTTTTTTTGAGATTTTTTTTTTGT U-(2,3) Ribozyme CGCACTGTAAAGCTAACTTAGCATTAACCTTTTAAGTTAAAGA TTAAGAGAACCAACACCTCTTTACAGTGA-3’

5’-GAAATTAATACGACTCACTATAGGTTTTTTTTCCTGATGAGT U-(1,2,3) TTTTTTTGAGATTTTTTTTCGAAACTTTTTTTTGAGATTTTTTTT Ribozyme GTCGTTTTTTTTTAAAGCTAACTTAGCATTAACCTTTTAAGTTA AAGATTAAGAGAACCAACACCTCTTTACAGTGA-3’

5’-GAAATTAATACGACTCACTATAGGCAGTGCCTGATGAGGCC AATGGGCCGAAACTCGTAAGAGTCGCACTGTAAAGCTAACTT Original A Ribozyme GCATTAACCTTTTAAGTTAAAGATTAAGAGAACCAACACCTCTT TACAGTGA-3’

T7 Promoter 5’-TAATACGACTCACTATA-3’

Promoter sequence is underlined and will not appear in final RNA transcripts

101

3.4.3 Experimental Procedures

3.4.3.1 Synthesis of DFHBI

DFHBI was synthesized using detailed procedure from Paige et al.50

(Z)-2,6-difluoro-4-((2-methyl-5-oxooxazol-4(5H)-ylidene)methyl)phenyl acetate (2).

N-Acetylglycine (74.1 mg, 0.63 mmol), anhydrous sodium acetate (52 mg, 0.63 mmol), 4- hydroxy-3,5- difluorobenzaldehyde (100 mg, 0.63 mmol), and acetic anhydride (0.25 ml) were stirred at 110 °C for 2 h. After allowing the reaction to cool to room temperature, cold ethanol (2.0 ml) was added while stirring and the reaction was left stirring overnight at 4 °C.

The resulting crystalline solid was then washed with a small amount of cold ethanol, hot water, hexanes and dried to afford 120 mg (yield 68%) of 2 as a pale yellow solid: 1H NMR

(500 MHz, CDCl3) δ 7.76 (d, J = 13.6 Hz, 2H), 6.95 (s, 1H), 2.42 (s, 3H), 2.39 (s, 3H)

(Z)-4-(3,5-difluoro-4-hydroxybenzylidene)-1,2-dimethyl-1H-imidazol-5(4H)-one(3,DFHBI).

Modifications were applied to the work-up of final product 3. Briefly, compound 2 (100mg,

0.36mmol) was refluxed with 0.12mL 33% CH3NH2 (in EtOH), 1.5mL EtOH and 70mg

K2CO3 for 3 hours. Reaction mixture was removed from heat and upon cooling formed an orange precipitate. Solvent was removed under vacuum, and solid was dissolved and extracted with 1:1 ethyl acetate and 500mM NaOAc (pH 3) three times. Organic layers were combined and concentrated under vacuum, following flash column purification. 1H

NMR (400MHz, CD3OD) δ7.77 (qd, J=8.0,1.6 Hz, 2H), 6.89 (s, 1H), 3.17 (s, 3H), 2.39 (s,

3H).

3.4.3.2 Radiolabeling of oligonucleotides

90 pmole of oligonucleotide was mixed with [γ-32P]-ATP and T4 polynucleotide kinase in

1x polynucleotide kinase buffer, incubated at 37 °C for 1 hour. Reaction was quenched by

102 heat denaturation at 65 °C for 20 minutes. Radiolabeled oligonucleotide was purified using

Bio-Spin 30 column.

3.4.3.3 RNA Transcription and Purification

DNA oligonucleotides containing the desired mRNA sequences were constructed to imbed

T7 promoter sequence at 3’ end. Transcription assays were performed using

MEGAshortscript T7 kit, and sequentially purified using MEGAclear purification kit. The purity of transcribed mRNAs was analyzed through denaturing electrophoresis.

3.4.3.4 Electrophoretic Mobility Shift Assays (EMSA)

IgE targeting aptamer and aptamer derivatives: 0.05 nM of radio-labeled oligonucleotide was mixed with bPNA 10 at various concentrations (0 nM to 50 nM) in 1x PBSM buffer

(138 mM NaCl, 2.7 mM KCl, 1.1 mM KH2PO4, 1 mM MgCl2, pH 7.4) supplemented with

0.05% BSA, 1mM DTT and 10% glycerol. The binding mixtures were allowed for equilibrium at room temperature for 30 minutes before applying to electrophoresis. Gels were dried and exposed to imaging plate overnight, gel images were scanned using

Typhoon scanner and qualified using ImageQuant software. Equilibrium dissociation

51, 52 constant Kd was obtained through curve fitting with Equation 3.1

U-Spinach RNA derivative: 100 nM of carboxyfluorescein labeled bPNA 10 was incubated with various concentrations of U-Spinach (0 nM to 400 nM) in 1x binding buffer (40 mM

HEPES, 125 mM KCl, 5 mM MgCl2, pH 7.4), samples were allowed for equilibrium at room temperature for 30 minutes before applied to electrophoresis. Fluorescent gels were scanned using Typhoon scanner with excitation wavelength at 488 nm and emission wavelength at 526 nm.

103

U-3 and U-(2,3) ribozyme derivatives: 100 nM of carboxyfluorescein labeled bPNAs 6,8,10 were incubated with various concentrations of ribozyme derivatives (0nM to 1600nM) in

1x Tris-Cl buffer (50mM Tris, pH 7.4), samples were allowed for equilibrium at room temperature for 30 minutes before applied to electrophoresis. Fluorescent gels were scanned using Typhoon scanner with excitation wavelength at 488 nm and emission wavelength at 526 nm. Gels were quantified using ImagQuant software, and apparent equilibrium dissociation constant Kd,app was obtained through curve fitting with Equation

3.2.51

3.4.3.5 IgE-Aptamer Filter Binding Assays

Filter binding assays were conducted using 96-well dot-blot system, pre-wetted filter sandwich was applied with Protran filter at the top, Hybond-N+ filter at the bottom. Samples were prepared as follows: 0.05 nM of radio labeled aptamer was incubated with 6.0nM of bPNA 10 in 1x PBSM buffer supplemented with 1mM DTT, 0.05% BSA and 10% glycerol.

Mixture was equilibrated at room temperature for 30 minutes for aptamer-bPNA complex formation. Then the equilibrated complex was titrated with IgE from 0nM until 2μM and incubated at 37 °C for 10 minutes. Equilibrated samples were applied to filtration system, followed immediately by 200μL buffer wash twice. After filtering all the binding mixtures, filters were air dried and exposed to imaging plate overnight. Filter images later on were scanned by Typhoon scanner, quantified using ImagQuant and fitted using Equation

3.3.52

3.4.3.6 Fluorescence Activation Assay

Two types of fluorescence activation experiments were performed. For fluorescence activation as a function of DFHBI concentration, 2 μM 1:1 ratio of U-Spinach and bPNA complex was first equilibrated at room temperature for 30 minutes, followed by the addition 104 of DFHBI from 0 nM to 2.8 μM. Samples were allowed to equilibrate for another 15 minutes before applying to spectra measurement. For each concentration of fluorophore measured, a background signal for fluorophore alone was also measured and subtracted from the signal measured for RNA and fluorophore together. For fluorescence activation as a function of DFHBI concentration 2 μM U-Spinach RNA was equilibrated with 0 μM to 10

μM of bPNA 6, 8,10 and 10 μM of DFHBI in 1x binding buffer, equilibrated at room temperature for 30 minutes before applying to spectra measurement. The background signal for 10 μM DFHBI was also subtracted from the fluorescence reading of the ternary complex.

3.4.3.7 Apparent Kd Measurement for Fluorescence Activation

For apparent Kd of DFHBI, 200nM of U-Spinach RNA was equilibrated with 500nM of bPNA 8, 10 or 1μM of bPNA 6 in 1x binding buffer, followed by titration of DFHBI from

0nM to 2.8μM. Samples were heated to 65 °C and gradually cooled down to room temperature. Fluorescence intensity was measured using plate reader with 465nM as excitation wavelength and 505 as emission wavelength. Intensity data was quantified

52 using Equation 3.4 . For apparent Kd of bPNA, measurements described in 3.6.6 were quantified.

3.4.3.8 U-Ribozyme Rescue Experiment

500 nM U-Ribozyme construct was mixed with 4 μM of bPNA 6, 8, 10 in 1x Tris-Cl buffer without Mg2+, heated up to 75 °C and gradually cooled to room temperature in order to achieve specific complexation. Cleavage reaction was initiated by adding different concentrations of Mg2+ into complex solutions. Aliquots of reaction mixtures were taken out at 3, 8, 15, 45, 90 minutes and quenched with 500 mM EDTA. Cleavage products were analyzed through denaturing electrophoresis, gels were stained with SYBR® Gold 105

(Invitrogen). Data was quantified using Equation 3.553.

3.4.3.9 In situ Splicing

Transcription assays were performed in 1x RNA polymerase reaction buffer provided by

New England Biolabs (NEB), supplemented with 10 mM DTT and 15mM Mg2+. 200 nM annealed DNA with/without 8 equivalent of bPNA 10 was mixed together. Then 4 unit/μL

T7 RNA polymerase was added followed by addition of ribonucleotides (4 mM NTP each). Reaction solution was incubated at 37ºC, and fractions of reaction solution were taken out and quench into 250 mM EDTA at different time points. The results were resolved on 14% denaturing PAGE containing 8 M Urea in 1x TBE buffer, and stained with SYBR® gold (Invitrogen) nucleic acid stain. Gels were scanned using Typhoon

Image Scanner.

106

3.5 Equation Table

1 1 Equation 3.1 [퐶표푚푝푙푒푥] = (퐾 + 퐷푁퐴 + 푏푃푁퐴 ) − 2√(퐾 + 퐷푁퐴 + 푏푃푁퐴 )2 − 4퐷푁퐴 푏푃푁퐴 2 푑 0 0 2 푑 0 0 0 0

푏푃푁퐴0 × 푅푁퐴0 Equation 3.2 [퐶표푚푝푙푒푥] = 퐾푑,푎푝푝 + 푅푁퐴0

1 [퐶표푚푝푙푒푥] = (퐾 + 퐼𝑔퐸 + 푎푝푡푎푚푒푟 푏푃푁퐴 ) 2 푑 0 0 Equation 3.3 1 − 2√(퐾 + 퐼𝑔퐸 + 푎푝푡푎푚푒푟 푏푃푁퐴 )2 − 4퐼𝑔퐸 푎푝푡푎푚푒푟 푏푃푁퐴 2 푑 0 0 0 0 1 [퐶표푚푝푙푒푥] = (퐾 + 퐷퐹퐻퐵퐼 + 푈푆푝𝑖푛푎푐ℎ 푏푃푁퐴 ) 107 푑,푎푝푝 0 0 Equation 3.4 2 1 2

2 − √(퐾 + 퐷퐹퐻퐵퐼 + 푈푆푝𝑖푛푎푐ℎ 푏푃푁퐴 ) − 4퐷퐹퐻퐵퐼 푈푆푝𝑖푛푎푐ℎ 푏푃푁퐴 2 푑,푎푝푝 0 0 0 0

Equation 3.5 [푃푟표푑푢푐푡] = 푈푅𝑖푏표푧푦푚푒 × (1 − 푒푥푝−푘푡)

3.6 Additional Data

3.6.1 DFHBI Synthesis

O F O N AcO F

108

Figure 3.17. 1H NMR of DFHBI precursor. (Z)-2,6-difluoro-4-((2-methyl-5-oxooxazol-4(5H)-ylidene)methyl)phenyl acetate.

109

Figure 3.18. 1H NMR of DFHBI. (Z)-4-(3,5-difluoro-4-hydroxybenzylidene)-1,2-dimethyl-1H-imidazol-5(4H)-one.

3.6.2 Fluorescence Activation of tRNALysU-Spinach

Figure 3.19. Fluorescence activation of tRNALysU-Spinach. U-Spinach sequence was designed to replace the anti-codon stem of tRNALys to achieve higher stability due to the compact folding of tRNA (Left). Fluorescence activation upon bPNA 10 addition was measured (Top right). Fluorescence intensities at 505nm with different bPNA 10 concentrations were plotted, yielding apparent Kd of 338nM, comparable to original U-Spinach constructs (Bottom right).

110

3.6.3 Mg2+ Dependent Cleavage of U-3

Figure 3.20. U-3 cleavage at higher Mg2+ concentration. Rescue of self-splicing function of U- 3 RNA ribozyme is Mg2+ dependent. Under 0.8 mM and 0.5 mM Mg2+ concentration, U-3 presented minimal self-splicing activity, while upon complexation with bPNA 6, 8 and 10, splicing activity is restored, and three bPNAs presented similar rescue behavior (Figure 3.12).

111

Figure 3.21. U-3 Ribozyme cleavage at lower Mg2+ concentration. Rescue of self-splicing function of U-3 RNA ribozyme at lower Mg2+ concentrations. At 0.25mM Mg2+ concentration, difference about splicing rates can be seen among 3 bPNAs with bPNA 10 showing the highest splicing rate, and bPNA 6 the lowest. When Mg2+ concentration reached 0.1mM, only bPNA 10 showed noticeable self-splicing rescue (Figure 3.12).

112

3.6.4 Mg2+ Dependent Cleavage of U-(2,3)

Figure 3.22. Gel images of Mg2+ dependent U-(2,3) cleavage. (a). Rescue of self-splicing function of U-(2,3) ribozyme is Mg2+ dependent. U-(2,3) self-splicing rescue stopped at 1 mM Mg2+

113

Figure 3.23. Kinetics of Mg2+ dependent U-(2,3) cleavage. Kinetic plots of bond scission activity modulated by bPNA 8 (Top) and 10 (Bottom) at different Mg2+ concentrations.

114

3.7 References for Chapter 3

1. Shin, D.; Tor, Y., Bifacial Nucleoside as a Surrogate for Both T and A in Duplex

DNA. J. Am. Chem. Soc. 2011, 133, 6926-6929.

2. Branda, N.; Kurz, G.; Lehn, J.-M., JANUS WEDGES: a new approach towards

nucleobase-pair recognition. Chem. Commun. 1996, 1996, 2443-2444.

3. Chen, H.; Meena; McLaughlin, L. W., A Janus-Wedge DNA Triplex with A-W1-T

and G-W2-C Base Triplets. J. Am. Chem. Soc. 2008, 130, 13190-13191.

4. Zeng, Y.; Pratumyot, Y.; Piao, X.; Bong, D., Discrete assembly of synthetic

peptide-DNA triplex structures from polyvalent melamine-thymine bifacial

recognition. J. Am. Chem. Soc. 2012, 134, 832-5.

5. Piao, X.; Xia, X.; Bong, D., Bifacial Peptide Nucleic Acid Directs Cooperative

Folding and Assembly of Binary, Ternary, and Quaternary DNA Complexes.

Biochemistry 2013, 52, 6313-6323.

6. Xia, X.; Piao, X.; Fredrick, K.; Bong, D., Bifacial PNA complexation inhibits

enzymatic access to DNA and RNA. Chembiochem 2013, 15, 31-36.

7. Arambula, J. F.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C., A simple

ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein

binding. Proc. Natl. Acad. Sci. USA 2009, 106, 16068-16073.

8. Zhou, Z.; Bong, D., Small-Molecule/Polymer Recognition Triggers Aqueous-Phase

Assembly and Encapsulation. Langmuir 2013, 29, 144-150.

9. Vysabhattar, R.; Ganesh, K. N., Cyanuryl peptide nucleic acid: synthesis and DNA

complexation properties. Tetrahedron Lett. 2008, 49, 1314-1318.

10. Mittapalli, G. K.; Reddy, K. R.; Xiong, H.; Munoz, O.; Han, B.; De Riccardis, F.;

Krishnamurthy, R.; Eschenmoser, A., Mapping the landscape of potentially

115

primordial informational oligomers: oligodipeptides and oligodipeptoids tagged

with triazines as recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2470-7.

11. Hysell, M.; Siegel, J. S.; Tor, Y., Synthesis and stability of exocyclic triazine

nucleosides. Organic & Biomolecular Chemistry 2005, 3, 2946-2952.

12. Cafferty, B. J.; Gállego, I.; Chen, M. C.; Farley, K. I.; Eritja, R.; Hud, N. V., Efficient

Self-Assembly in Water of Long Noncovalent Polymers by Nucleobase Analogues.

J. Am. Chem. Soc. 2013, 135, 2447-2450.

13. Whitesides, G. M.; Simanek, E. E.; Mathias, J. P.; Seto, C. T.; Chin, D.; Mammen,

M.; Gordon, D. M., Noncovalent Synthesis: Using Physical-Organic Chemistry To

Make Aggregates. Acc. Chem. Res. 1995, 28, 37-44.

14. Ma, M.; Paredes, A.; Bong, D., Intra- and intermembrane pairwise molecular

recognition between synthetic hydrogen-bonding phospholipids. J. Am. Chem. Soc.

2008, 130, 14456-8.

15. Ma, M.; Gong, Y.; Bong, D., Lipid Membrane Adhesion and Fusion Driven by

Designed, Minimally Multivalent Hydrogen-Bonding Lipids. J. Am. Chem. Soc.

2009, 131, 16919-16926.

16. Ma, M.; Bong, D., Determinants of cyanuric acid and melamine assembly in water.

Langmuir 2011, 27, 8841-53.

17. Ma, M.; Bong, D., Directed peptide assembly at the lipid-water interface

cooperatively enhances membrane binding and activity. Langmuir 2011, 27, 1480-

18. Porcheddu, A.; Giacomelli, G., Peptide Nucleic Acids (PNAs), A Chemical

Overview. Curr. Med. Chem. 2005, 12, 2561-2599.

19. Nielsen, P. E., Peptide nucleic acids: protocols and applications. Taylor & Francis:

2004. 116

20. Mittapalli, G. K.; Osornio, Y. M.; Guerrero, M. A.; Reddy, K. R.; Krishnamurthy, R.;

Eschenmoser, A., Mapping the landscape of potentially primordial informational

oligomers: oligodipeptides tagged with 2,4-disubstituted 5-aminopyrimidines as

recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2478-84.

21. Ura, Y.; Beierle, J. M.; Leman, L. J.; Orgel, L. E.; Ghadiri, M. R., Self-assembling

sequence-adaptive peptide nucleic acids. Science 2009, 325, 73-7.

22. Garner, P.; Dey, S.; Huang, Y., α-Helical Peptide Nucleic Acids (αPNAs): A New

Paradigm for DNA-Binding Molecules. Journal of the American Chemical Society

2000, 122, 2405-2406.

23. Huang, Y.; Dey, S.; Zhang, X.; Sönnichsen, F.; Garner, P., The α-Helical Peptide

Nucleic Acid Concept: Merger of Peptide Secondary Structure and Codified

Nucleic Acid Recognition. Journal of the American Chemical Society 2004, 126,

4626-4640.

24. Ariga, K.; Kunitake, T., Molecular Recognition at Air-Water and Related Interfaces:

Complementary Hydrogen Bonding and Multisite Interaction. Acc. Chem. Res.

1998, 31, 371-378.

25. De Greef, T. F. A.; Smulders, M. M. J.; Wolffs, M.; Schenning, A.; Sijbesma, R. P.;

Meijer, E. W., Supramolecular Polymerization. Chemical Reviews 2009, 109,

5687-5754.

26. Kool, E. T., Preorganization of DNA: Design Principles for Improving Nucleic Acid

Recognition by Synthetic Oligonucleotides. Chemical Reviews 1997, 97, 1473-

1488.

27. Betts, L.; Josey, J. A.; Veal, J. M.; Jordan, S. R., A nucleic acid triple helix formed

by a peptide nucleic acid-DNA complex. Science 1995, 270, 1838-41.

117

28. Nielsen, P. E.; Egholm, M.; Berg, R. H.; Buchardt, O., Sequence-selective

recognition of DNA by strand displacement with a thymine-substituted polyamide.

Science 1991, 254, 1497-1500.

29. Bentin, T.; Nielsen, P. E., Superior duplex DNA strand invasion by acridine

conjugated peptide nucleic acids. J. Am. Chem. Soc. 2003, 125, 6378-9.

30. Bentin, T.; Larsen, H. J.; Nielsen, P. E., Combined Triplex/Duplex Invasion of

Double-Stranded DNA by "Tail-Clamp" Peptide Nucleic Acid. Biochemistry 2003,

42, 13987-13995.

31. Kaihatsu, K.; Shah, R. H.; Zhao, X.; Corey, D. R., Extending Recognition by

Peptide Nucleic Acids (PNAs): Binding to Duplex DNA and Inhibition of

Transcription by Tail-Clamp PNA-Peptide Conjugates. Biochemistry 2003, 42,

13996-14003.

32. Rapireddy, S.; He, G.; Roy, S.; Armitage, B. A.; Ly, D. H., Strand invasion of mixed-

sequence B-DNA by acridine-linked, gamma-peptide nucleic acid (gamma-PNA).

J. Am. Chem. Soc. 2007, 129, 15596-600.

33. Datta, B.; Schmitt, C.; Armitage, B. A., Formation of a PNA2-DNA2 hybrid

quadruplex. Journal of the American Chemical Society 2003, 125, 4111-4118.

34. Datta, B.; Bier, M. E.; Roy, S.; Armitage, B. A., Quadruplex Formation by a

Guanine-Rich PNA Oligomer. Journal of the American Chemical Society 2005, 127,

4199-4207.

35. Gupta, A.; Lee, L.-L.; Roy, S.; Tanious, F. A.; Wilson, W. D.; Ly, D. H.; Armitage,

B. A., Strand Invasion of DNA Quadruplexes by PNA: Comparison of Homologous

and Complementary Hybridization. Chembiochem 2013, 14, 1476-1484.

36. Perutz, M. F., Mechanisms of cooperativity and allosteric regulation in proteins.

Quarterly reviews of biophysics 1989, 22, 139-237. 118

37. Breaker, R. R., Engineered allosteric ribozymes as biosensor components.

Current opinion in biotechnology 2002, 13, 31-39.

38. Fechter, E. J.; Dervan, P. B., Allosteric Inhibition of Protein-DNA Complexes by

Polyamide-Intercalator Conjugates. J. Am. Chem. Soc. 2003, 125, 8476-8485.

39. Wiegand, T. W.; Williams, P. B.; Dreskin, S. C.; Jouvin, M.-H.; Kinet, J.-P.; Tasset,

D., High-affinity oligonucleotide ligands to human IgE inhibit binding to Fc epsilon

receptor I. The Journal of Immunology 1996, 157, 221-230.

40. Paige, J. S.; Wu, K. Y.; Jaffrey, S. R., RNA Mimics of Green Fluorescent Protein.

Science 2011, 333, 642-646.

41. Paige, J. S.; Nguyen-Duc, T.; Song, W.; Jaffrey, S. R., Fluorescence Imaging of

Cellular Metabolites with RNA. Science 2012, 335, 1194.

42. Kellenberger, C. A.; Wilson, S. C.; Sales-Lee, J.; Hammond, M. C., RNA-Based

Fluorescent Biosensors for Live Cell Imaging of Second Messengers Cyclic di-

GMP and Cyclic AMP-GMP. Journal of the American Chemical Society 2013, 135,

4906-4909.

43. Suess, B.; Weigand, J. E., Engineered riboswitches: overview, problems and

trends. RNA Biol. 2008, 5, 24-9.

44. Roth, A.; Breaker, R. R., The structural and functional diversity of metabolite-

binding riboswitches. Annual review of biochemistry 2009, 78, 305-334.

45. Topp, S.; Gallivan, J. P., Emerging Applications of Riboswitches in Chemical

Biology. ACS Chemical Biology 2010, 5, 139-148.

46. Ogawa, A., Rational design of artificial riboswitches based on ligand-dependent

modulation of internal ribosome entry in wheat germ extract and their applications

as label-free biosensors. RNA 2011, 17, 1-11.

119

47. Murray, J. B.; Terwey, D. P.; Maloney, L.; Karpeisky, A.; Usman, N.; Beigelman,

L.; Scott, W. G., The structural basis of hammerhead ribozyme self-cleavage. Cell

1998, 92, 665-673.

48. Buskirk, A. R.; Liu, D. R., Creating small-molecule-dependent switches to

modulate biological functions. Chem Biol 2005, 12, 151-61.

49. Link, K. H.; Breaker, R. R., Engineering ligand-responsive gene-control elements:

lessons learned from natural riboswitches. Gene therapy 2009, 16, 1189-201.

50. Paige, J. S.; Wu, K. Y.; Jaffrey, S. R., RNA mimics of green fluorescent protein.

Science 2011, 333, 642-6.

51. Lundblad, J. R.; Laurance, M.; Goodman, R. H., Fluorescence polarization

analysis of protein-DNA and protein-protein interactions. Molecular endocrinology

1996, 10, 607-12.

52. Agou, F.; Raveh, S.; Mesnildrey, S.; Veron, M., Single strand DNA specificity

analysis of human nucleoside diphosphate kinase B. J Biol Chem 1999, 274,

19630-8.

53. Burgin, A. B., Jr.; Gonzalez, C.; Matulic-Adamic, J.; Karpeisky, A. M.; Usman, N.;

McSwiggen, J. A.; Beigelman, L., Chemically modified hammerhead ribozymes

with improved catalytic rates. Biochemistry 1996, 35, 14090-7.

120

CHAPTER 4

bPNA Ligation and Ribozyme Cleavage

via An Abiotic Interface

This chapter is reproduced with permission from J. Am. Chem. Soc., 2015, 137, 3751-3754.

121

4.1 Overview

We report herein DNA- and RNA- templated chemical transformation of bifacial peptide nucleic acid (bPNA) fragments directed by an abiotic triplex hybrid interface. Assembly of one bPNA strand with two unstructured oligo T/U strands enables facile insertion of DNA and RNA template sites within partially folded nucleic acids; this template topology is not easily accessed through native base-pairing. RNA-templated oxidative coupling of bPNA fragments is found to result in the emergence of ribozyme cleavage function, thus establishing a connection between engineered and native reaction sites. These data demonstrate the use of new topologies in nucleic acid- templated chemistry that could serve as chemically sensitive DNA and RNA switches.

4.2 Introduction

4.1.2 Structure Defines Function

For the sustenance of life, highly efficient and specific chemical reactions, which produce essential building blocks, convert metabolites, respond to extracellular stimuli and more, are constantly catalyzed in biological organisms via enzymes. After dominating the enzymatic research area for decades, the statement “enzymes are catalytic proteins” was proven to be incomprehensive due to the discovery of catalytic RNA molecules,1-4 which are now termed as ribozymes. The “RNA world” hypothesis5, 6 surfaced after numerous laboratory studies indicated that RNA is capable to catalyze a large variety of reactions that are biologically significant;7-10 further with in-lab evolution, new ribozymes can be

122

generated to fit new catalytic needs.11-15 Compare to the large reservoir of protein building blocks, the fact that RNA with as few as 4 building blocks can present similar biological functions is astounding. One possible explanation draw from available crystal structures indicates ribozymes, though simpler in primary sequences, are capable of adopting compact and complicated tertiary structures;16, 17 flexible backbone enables contact of primarily distant nucleotide sequences through globular folding driven by Watson-Crick interactions, base stacking and metabolite binding. This explanation has been widely supported by the observations of sustained ribozyme activity disregard extensive sequence alterations: Group I intron catalyzes accurate mRNA splicing via recognition of a substrate stem-loop despite the minimal sequence homology of the mRNA;18 hammerhead ribozyme can be elongated, shortened or engineered from cis-cleaving into trans multi-turnover catalytic unit as long as the integrity of active conformation is well maintained.19, 20

This “structure determines functions” point of view inspired new horizons of RNA engineering that utilizes ribozymes to carry out more complicated reactions,21, 22 instead of simply cleave and ligate a phosphate backbone, ribozyme activities were employed into gene regulation and modification,14, 23 or further evolved to self-replicating,24 or as a structural switch for metabolite sensor.25

4.2.2 Abiotic Template Interface for bPNA Ligations

DNA recognition has been exploited to direct the chemistry of native and artificial26-30 macromolecules. Watson−Crick base-pairing can be taken out of context to enable group transfer,31-33 nucleic acid detection,34 chemical library selection,35 and self-replication.36 In addition, base-pairing templates can code for synthesis of non-native scaffolds37-39 and

123

multisite macromolecular modification.40-42 The abiotic base triple interface between bifacial peptide nucleic acid (bPNA)43-46 and nucleic acids provides an opportunity to explore alternative template topologies. Triazine47-55 bases in bPNA simultaneously dock two oligo T/U strands to form an obligate triplex hybrid.

Unlike conventional PNA, which dissociatively invades native structures, bPNA recognition is an associative process that unites non-interacting native domains. Though symmetric56-60 two-strand61-64 recognition of this type has no cognate in extant biology, amino and oxo 2,4,6-substituted triazines65, 66 recapitulate the Watson−Crick hydrogen- bonding patterns, fueling the speculative notion of triazine-derived precursors67 to DNA and RNA. We have previously demonstrated that bPNA hybridization can trigger DNA and

RNA chemistry.50

My specific contributions to this project was to establish a two-way communication that links the abiotic interface with a native catalytic cleavage site. Integration of a template site into a ribozyme fold renders RNA splicing dependent on oxidative ligation of bPNA; this could serve as a blueprint for chemically sensitive nucleic acid switches68, 69 and gates70, 71with applications in DNA/RNA nanotechnology.72 Overall, these data demonstrate readout and transformation of non-native macromolecules through an abiotic template interface in DNA/RNA template topologies that are not accessible via native base-pairing.

4.3 Results and Discussion

In Appendix A, details regarding facts that hybridization can likewise trigger bPNA chemistry will be discussed. Briefly here, single-stranded and partially structured

124

DNA/RNA topologies were found to serve as templates to catalyze bPNA coupling and controlled chain extension (oligomerization) of bi-reactive bPNAs. (Figure 4.1)

Figure 4.1. bPNA structure and experimental designs. (Top) T/U-rich tracts can bind bPNA via melamine- derivatized lysine (M*). (Bottom) The four template topologies shown catalyze thioester exchange, fragment ligation, chain extension, and oxidative coupling with activation of ribozyme splicing function with bPNA substrates (dark lines).

Observations showed successful catalysis with partially folded DNA templates, and these data collectively demonstrate that effective molarity increases on DNA and RNA templates 125

can catalyze acyl transfer and oxidative coupling as well as chain extension of bPNA fragments. (Appendix A) Hence, it has prompted the investigation of Un-template loops imbedded within RNA folds. This notion was tested using a minimal type I hammerhead ribozyme in which stem III was replaced with a rU10CACAU10 loop (U-3-ribozyme). It was initially thought that the constrained U-loop would exhibit orientational bias with respect the RNA template; thus, two 4mer bPNAs bearing N-terminal (6) and C-terminal (7) thiols were prepared and studied. (Figure 4.2)

N-terminal thiol (6) C-terminal thiol (7)

Figure 4.2. bPNA structures of N-terminal thiol (6) and C-terminal thiol (7).

However, both thiols and their mixture, gave identical template-enhanced oxidation profiles with U-3-ribozyme (Figure 4.3). Interestingly, the U-3-ribozyme was able to achieve a similar yield of oxidation at lower catalyst loading (12%) compared to the DNA

(50%), suggesting a higher exchange rate off the RNA loop template, though more quantitative measurements are needed to pinpoint the origin of this difference.

126

Figure 4.3. U-3 ribozyme-templated bPNA thiol oxidation. Oxidation of bPNAs 6 (Mpa- (EM*)4G) and 7 (EM*)4C alone (○) and with 12 mol% U-3-ribozyme (●), followed by Ellman’s test. Mpa = mercaptopropionamide. Lines are drawn to guide the eye.

This notion was tested using oxidative thiol coupling since amide bond ligation occurs on a time scale similar to RNA degradation. While 4mer bPNA (EM*)4 and bPNAs 4, 6, and

7 only weakly activate cleavage of the U-3-ribozyme (Figure 4.4), oxidative coupling 127

Figure 4.4. Thiol oxidation coupled U-3 self-cleavage. (Top) U-3-ribozyme cleavage upon oxidation of bPNA thiols 6 and 7 and addition of Mg2+. (Bottom) Denaturing PAGE of RNA cleavage triggered by 4mer bPNA (EM*)4 and 10% oxidized 6 and 7, with full-length RNA (*), tRNA (<), and ribozyme (≪) cleavage products indicated, along with gel quantification, with the bPNA additives indicated. Lines are drawn to guide the eye. produces an ∼8mer bPNA disulfide product that binds more tightly to the template and strongly activates function. Oxidation conditions are more concentrated than those for ribozyme cleavage; thus, reactions were studied by dilution of partially oxidized samples into ribozyme cleavage conditions with 1 mM Mg2+. PAGE analysis of the reaction indicated the formation of two RNA products upon 10% bPNA oxidation, which were identified as the tRNA fusion and the hammerhead ribozyme components (Figure 4.4).

128

Positive control experiments using 8mer bPNA (EM*)8 and fully oxidized and purified mixed disulfides of 6 and 7 gave higher yields of splicing. Ribozyme catalytic activity therefore may be used to report on fragment oxidative coupling, indicating two-way communication between an engineered abiotic template site and a native RNA splicing site (Figure 4.5). This connection makes possible functional selection and optimization of template and redox-switchable ribozymes.

Figure 4.5. Two-way communication between an engineered abiotic template site and a native RNA splicing site.

4.4 Experimental procedures

4.3.1 RNA Transcription and Purification

DNA oligonucleotides containing the desired mRNA sequences were constructed to imbed

T7 promoter sequence at 3’ end. Transcription assays were performed using

MEGAshortscript T7 kit, and sequentially purified using MEGAclear purification kit. The purity of transcribed mRNAs was analyzed through denaturing electrophoresis.

129

4.3.2 Ribozyme Cleavage with Pre-oxidized Peptide

500 nM U-Ribozyme constructs were mixed with 4 μM of pre-oxidized disulfide bPNA 6, 7 in 1x Tris-Cl (pH 7.4, 50mM) buffer, Cleavage reaction was initiated by adding 1mM Mg2+ for U-3 cleavage or 10mM Mg2+ for U23. Aliquots of reaction mixtures were taken out at

15, 30, 60, 120, 180 minutes and quenched with 500 mM EDTA. Cleavage products were analyzed through denaturing electrophoresis, gels were stained with SYBR® Gold

(Invitrogen). Data was quantified using Equation 4.1.

Equation 4.1: 푃푟표푑푢푐푡 = 푈푅𝑖푏표푧푦푚푒× (1 – 푒푥푝-kt)

4.3.3 Ribozyme Facilitated bPNA Oxidation

25 μM U-3-ribozyme constructs were mixed with 200 μM reduced bPNA mixture containing 6 and 7 in 1x Tris-Cl (pH7.4, 50mM), the mixture was incubated at 37 °C.

Oxidation progress was monitored and quantified through Ellman’s Test. Control experiment with only 200 μM bPNA mixture was performed simultaneously and quantified using the same method.

4.4.4 In situ Oxidation and Cleavage

Oxidation of bPNA mixture containing 6 and 7 with the presence of U-ribozyme was performed as described above. When oxidation percentage reached 10%, 30% and 50%, portions of oxidation mixture was taken out and diluted 50-fold and cleavage reaction was initiated with the addition of 1mM Mg2+, aliquots of reaction mixtures were pipetted out at

15, 30, 60, 120, 180 minutes and quenched with 500 mM EDTA. Cleavage products were analyzed through denaturing electrophoresis, gels were stained with SYBR® Gold

(Invitrogen). Data was quantified using Equation 4.1.

130

Data sum m ary: U3 cleavage 1mM Mg2+ 4.5 Additional Data

4.5.1 bPNA Initiated U-3 Cleavage Under 1 mM Mg2+ Data sum m ary: U3 cleavage 1mM Mg2+

0 15 30 60 120 180 0 15 30 60 120 180 0 15 30 60 120 180 U3 control U3 with bPNA 8 U3 with bPNA 4

2+ Under 1mM Mg , no cleavage is observed for U3-bPNA 4 mixture. bPNA 8 showed robust cleavage with an apparent r0at e co15ns tant30 as 600.0027 120 s -1.180 0 15 30 60 120 180 0 15 30 60 120 180 U3 control U3 with 8mer bPNA U3 with 4mer bPNA 400 Under 1mM Mg2+, no U3 cleavage is observed when mixed with 4mer bPNA. Meanwhile complexation with 8 mer bPNA showed robust cleavage with an apparent rate constant as 0.0027 s-1. 350 400 300

) 350

M 250

( 300

]

)

M c 200250 y = m1*(1-exp(-m2*m0))

( Value Error

]

t o 150 m1 361.32 0.96262

c r 200 m2y = m10.0026829*(1-exp(-m2*m06)).4803e-5

P Value Error

[ d Chisq 8.2143 NA

o 150 m1 361.32 0.96262 r 100 R 0.99996 NA m2 0.0026829 6.4803e-5

[ Chisq 8.2143 NA 50100 R 0.99996 NA

50 0 0 2000 4000 6000 8000 1 104 1.2 104 0 4 4 0 2000 T4000ime (se6000co8000nd) 1 10 1.2 10 Time (second)

Figure 4.6. bPNA length dependent U-3 cleavage. Under 1mM Mg2+, no U-3 cleavage is observed when mixed with 4mer bPNA. Meanwhile complexation with 8 mer bPNA showed robust cleavage with an apparent rate constant as 0.0027 s-1.

131

4.5.2 U-3 Cleavage with Pre-oxidized bPNAs U3 cleavage 1mM Mg2+ with pre-oxidized bPNAs

0 15 30 60 120 180 0 15 30 60 120 180 0 15 30 60 120 180 U3 with bPNA 6 U3 with bPNA 6 + 7 U3 with bPNA 7

When pre-oxidized bPNAs was added, U3 restored cleavage activity with the apparent rate constant as 0.0027 s-1. No orientation bias was presented.

Pre-oxidation 500

400

) M

n 300

(

]

c u

d 200

o r

P y = m1*(1-exp(-m2*m0)) [ Value Error 100 m1 396.93 5.0213 m2 0.0027394 0.00035838 Chisq 370.77 NA R 0.99855 NA 0 0 2000 4000 6000 8000 1 104 1.2 104 Time (second)

Figure 4.7. U-3 cleavage restored by pre-oxidized bPNAs. When pre-oxidized bPNAs was added, U-3 restored cleavage activity with the apparent rate constant as 0.0027 s-1. No orientation bias was presented.

132

4.5.3 In situ bPNA Oxidation and U-3 Cleavage In s itu U3 cleavage under 1mM Mg2+ with bPNA 6 + 7

0 15 30 60 120 180 0 15 30 60 120 180 0 15 30 60 120 180 10% Oxidation 30% Oxidation 50% Oxidation

In s itu oxidation and cleavage showed similar rate constant disregard the difference in oxidation percentage. 250 250

200 200

)

M n

150 n 150 (

( y = m1*(1-exp(-m2*m0))

]

] t

t Value Error c

c m1 194.36 3.8856 u

u m2 0.0014389 0.00013032 d

100 d 100 o

y = m1*(1-exp(-m2*m0)) o Chisq 92.834 NA r

Value Error r R 0.99829 NA

P [ m1 230.19 5.1051 [ m2 0.001452 0.0001672 50 Chisq 318.79 NA 50 R 0.99617 NA

0 0 0 2000 4000 6000 8000 1 104 1.2 104 0 2000 4000 6000 8000 1 104 1.2 104

Time (second) Time (second) 10% Oxidation k = 0.0015 s-1 30% Oxidation k = 0.0014 s-1 250

200

) M

n 150 (

] t

c u y = m1*(1-exp(-m2*m0)) d 100 o Value Error r m1 212.25 7.5891

P [ m2 0.0018487 0.00042002 Chisq 764.73 NA 50 R 0.9893 NA

0 0 2000 4000 6000 8000 1 104 1.2 104

Time (second) 50% Oxidation k = 0.0018 s-1

Figure 4.8. in situ bPNA oxidation and U-3 cleavage. In situ oxidation and cleavage showed similar rate constant disregard the difference in oxidation percentage.

133

Figure 4.9. Emergence of U-3 cleavage via templated bPNA oxidation. Comparison of in situ cleavage at 10% oxidation to bPNA 6+7 backgound oxidation without the presence of U- 3-ribozyme.

134

4.5.4 bPNA Initiated U-(2,3) Cleavage Under 1 mM Mg2+

U23-ribozyme cleavage 10mM Mg2+

0 15 30 60 120 180 0 15 30 60 120 180 0 15 30 60 120 180 U23 control U23 with 8mer bPNA U23 with 4mer bPNA Under 1mM Mg2+, no cleavage is observed when mixe d with 4mer bPNA. Complexation with 8mer bPNA showed robust cleavage with an apparent rate constant as 0.0012 s-1.

350

300

250

)

(

200

]

d 150 y = m1*(1-exp(-m2*m0))

r Value Error

P m1 300.1 7.6987 [ 100 m2 0.0012361 0.00014791 Chisq 680.57 NA 50 R 0.99506 NA

0 0 2000 4000 6000 8000 1 104 1.2 104 Time (second)

2+ Figure 4.10. bPNA length dependence of U-(2,3 ) cleavage. Under 1mM Mg , no cleavage is observed when mixed with 4mer bPNA. Complexation with 8mer bPNA showed robust cleavage with an apparent rate constant as 0.0012 s-1.

135

4.5.5 U-(2,3) Cleavage with Pre-oxidized bPNAs

In s itu U23 cleavage under 10mM Mg2+ with bPNA 6 + 7

0 15 30 60 120 180 0 15 30 60 120 180 0 15 30 60 120 180 U23 with bPNA 6 U23 with bPNA 6 + 7 U23 with bPNA 7)

When pre-oxidized bPNA 4 was added, U23 restored cleavage activity with the apparent rate constant as 0.0011 s-1. No orientation bias was observed.

U23 Preoxidation 350

300

250

)

(

200

]

t y = m1*(1-exp(-m2*m0))

c Value Error u m1 293.9 12.396

d 150

o m2 0.0011473 0.00021598 r Chisq 1709.1 NA

P [ 100 R 0.98718 NA

0 0 2000 4000 6000 8000 1 104 1.2 104 Time (second)

Figure 4.11. U-(2,3) cleavage restored by pre-oxidized bPNAs. When pre-oxidized bPNA 4 -1 was added, U23 restored cleavage activity with the apparent rate constant as 0.0011 s . No orientation bias was observed.

136

4.6 References for Chapter 4

1. Cech, T. R.; Zaug, A. J.; Grabowski, P. J., In vitro splicing of the ribosomal RNA

precursor of Tetrahymena: involvement of a guanosine nucleotide in the excision

of the intervening sequence. Cell 1981, 27, 487-96.

2. Grabowski, P. J.; Zaug, A. J.; Cech, T. R., The intervening sequence of the

ribosomal RNA precursor is converted to a circular RNA in isolated nuclei of

Tetrahymena. Cell 1981, 23, 467-76.

3. Kruger, K.; Grabowski, P. J.; Zaug, A. J.; Sands, J.; Gottschling, D. E.; Cech, T.

R., Self-splicing RNA: autoexcision and autocyclization of the ribosomal RNA

intervening sequence of Tetrahymena. Cell 1982, 31, 147-57.

4. Zaug, A. J.; Grabowski, P. J.; Cech, T. R., Autocatalytic cyclization of an excised

intervening sequence RNA is a cleavage-ligation reaction. Nature 1983, 301, 578-

83.

5. Gilbert, W., Evolution of antibodies. The road not taken. Nature 1986, 320, 485-6.

6. Cech, T. R., The efficiency and versatility of catalytic RNA: implications for an RNA

world. Gene 1993, 135, 33-6.

7. Mandal, M.; Breaker, R. R., Gene regulation by riboswitches. Nature reviews.

Molecular cell biology 2004, 5, 451-63.

8. Doudna, J. A.; Cech, T. R., The chemical repertoire of natural ribozymes. Nature

2002, 418, 222-8.

9. Sarver, N.; Cantin, E. M.; Chang, P. S.; Zaia, J. A.; Ladne, P. A.; Stephens, D. A.;

Rossi, J. J., Ribozymes as potential anti-HIV-1 therapeutic agents. Science 1990,

247, 1222-5.

137

10. Pyle, A. M., Ribozymes: a distinct class of metalloenzymes. Science 1993, 261,

709-14.

11. Seelig, B.; Jaschke, A., A small catalytic RNA motif with Diels-Alderase activity.

Chemistry & biology 1999, 6, 167-76.

12. Tarasow, T. M.; Tarasow, S. L.; Eaton, B. E., RNA-catalysed carbon-carbon bond

formation. Nature 1997, 389, 54-7.

13. Serganov, A.; Keiper, S.; Malinina, L.; Tereshko, V.; Skripkin, E.; Hobartner, C.;

Polonskaia, A.; Phan, A. T.; Wombacher, R.; Micura, R.; Dauter, Z.; Jaschke, A.;

Patel, D. J., Structural basis for Diels-Alder ribozyme-catalyzed carbon-carbon

bond formation. Nature structural & molecular biology 2005, 12, 218-24.

14. Mikheeva, S.; Jarrell, K. A., Use of engineered ribozymes to catalyze chimeric

gene assembly. Proceedings of the National Academy of Sciences of the United

States of America 1996, 93, 7486-90.

15. Wilson, D. S.; Szostak, J. W., In vitro selection of functional nucleic acids. Annual

review of biochemistry 1999, 68, 611-47.

16. Celander, D. W.; Cech, T. R., Visualizing the higher order folding of a catalytic

RNA molecule. Science 1991, 251, 401-7.

17. Gesteland, R. F.; Atkins, J. F., The RNA world : the nature of modern RNA

suggests a prebiotic RNA world. p xxiii, 630 pages.

18. Doudna, J. A.; Cormack, B. P.; Szostak, J. W., RNA structure, not sequence,

determines the 5' splice-site specificity of a group I intron. Proceedings of the

National Academy of Sciences of the United States of America 1989, 86, 7402-6.

19. Tuschl, T.; Eckstein, F., Hammerhead ribozymes: importance of stem-loop II for

activity. Proceedings of the National Academy of Sciences of the United States of

America 1993, 90, 6991-4. 138

20. Martick, M.; Scott, W. G., Tertiary contacts distant from the active site prime a

ribozyme for catalysis. Cell 2006, 126, 309-20.

21. Breaker, R. R., Engineered allosteric ribozymes as biosensor components.

Current opinion in biotechnology 2002, 13, 31-9.

22. Breaker, R. R., Natural and engineered nucleic acids as tools to explore biology.

Nature 2004, 432, 838-45.

23. Muller, S., Engineered ribozymes as molecular tools for site-specific alteration of

RNA sequence. Chembiochem : a European journal of chemical biology 2003, 4,

991-7.

24. Wochner, A.; Attwater, J.; Coulson, A.; Holliger, P., Ribozyme-catalyzed

transcription of an active ribozyme. Science 2011, 332, 209-12.

25. Kellenberger, C. A.; Wilson, S. C.; Sales-Lee, J.; Hammond, M. C., RNA-based

fluorescent biosensors for live cell imaging of second messengers cyclic di-GMP

and cyclic AMP-GMP. Journal of the American Chemical Society 2013, 135, 4906-

26. Schmidt, J. G.; Nielsen, P. E.; Orgel, L. E., Information transfer from peptide

nucleic acids to RNA by template-directed syntheses. Nucleic Acids Research

1997, 25, 4797-4802.

27. Schmidt, J. G.; Christensen, L.; Nielsen, P. E.; Orgel, L. E., Information transfer

from DNA to peptide nucleic acids by template-directed syntheses. Nucleic Acids

Research 1997, 25, 4792-4796.

28. Zhan, Z. Y. J.; Lynn, D. G., Chemical amplification through template-directed

synthesis. Journal of the American Chemical Society 1997, 119, 12420-12421.

29. Goodwin, J. T.; Mehta, A. K.; Lynn, D. G., Digital and analog chemical evolution.

Accounts of chemical research 2012, 45, 2189-99. 139

30. Sharma, A. K.; Heemstra, J. M., Small-Molecule-Dependent Split Aptamer Ligation.

Journal of the American Chemical Society 2011, 133, 12426-12429.

31. Grossmann, T. N.; Seitz, O., DNA-catalyzed transfer of a reporter group. Journal

of the American Chemical Society 2006, 128, 15596-15597.

32. McKee, M. L.; Milnes, P. J.; Bath, J.; Stulz, E.; Turberfield, A. J.; O'Reilly, R. K.,

Multistep DNA-Templated Reactions for the Synthesis of Functional Sequence

Controlled Oligomers. Angewandte Chemie-International Edition 2010, 49, 7948-

7951.

33. McKee, M. L.; Evans, A. C.; Gerrard, S. R.; O'Reilly, R. K.; Turberfield, A. J.; Stulz,

E., Peptidomimetic bond formation by DNA-templated acyl transfer. Organic &

Biomolecular Chemistry 2011, 9, 1661-1666.

34. Silverman, A. P.; Kool, E. T., Detecting RNA and DNA with templated chemical

reactions. Chemical Reviews 2006, 106, 3775-3789.

35. Kleiner, R. E.; Dumelin, C. E.; Liu, D. R., Small-molecule discovery from DNA-

encoded chemical libraries. Chemical Society Reviews 2011, 40, 5707-5717.

36. Von Kiedrowski, G., A SELF-REPLICATING HEXADEOXYNUCLEOTIDE.

Angewandte Chemie International Edition in English 1986, 25, 932-935.

37. Li, Z. Y.; Zhang, Z. Y. J.; Knipe, R.; Lynn, D. G., DNA-catalyzed polymerization.

Journal of the American Chemical Society 2002, 124, 746-747.

38. Rosenbaum, D. M.; Liu, D. R., Efficient and sequence-specific DNA-templated

polymerization of peptide nucleic acid aldehydes. Journal of the American

Chemical Society 2003, 125, 13924-13925.

39. Lo, P. K.; Sleiman, H. F., Nucleobase-Templated Polymerization: Copying the

Chain Length and Polydispersity of Living Polymers into Conjugated Polymers. J.

Am. Chem. Soc. 2009, 131, 4182-4183. 140

40. Ura, Y.; Beierle, J. M.; Leman, L. J.; Orgel, L. E.; Ghadiri, M. R., Self-assembling

sequence-adaptive peptide nucleic acids. Science 2009, 325, 73-7.

41. Heemstra, J. M.; Liu, D. R., Templated synthesis of peptide nucleic acids via

sequence-selective base-filling reactions. J. Am. Chem. Soc. 2009, 131, 11347-9.

42. McHale, R.; O'Reilly, R. K., Nucleobase Containing Synthetic Polymers:

Advancing Biomimicry via Controlled Synthesis and Self-Assembly.

Macromolecules 2012.

43. Zeng, Y.; Pratumyot, Y.; Piao, X.; Bong, D., Discrete assembly of synthetic

peptide-DNA triplex structures from polyvalent melamine-thymine bifacial

recognition. J. Am. Chem. Soc. 2012, 134, 832-5.

44. Piao, X.; Xia, X.; Bong, D., Bifacial Peptide Nucleic Acid Directs Cooperative

Folding and Assembly of Binary, Ternary, and Quaternary DNA Complexes.

Biochemistry 2013, 52, 6313-6323.

45. Xia, X.; Piao, X.; Fredrick, K.; Bong, D., Bifacial PNA complexation inhibits

enzymatic access to DNA and RNA. Chembiochem : a European journal of

chemical biology 2013, 15, 31-36.

46. Xia, X.; Piao, X.; Bong, D., Bifacial peptide nucleic acid as an allosteric switch for

aptamer and ribozyme function. Journal of the American Chemical Society 2014,

136, 7265-8.

47. Arambula, J. F.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C., A simple

ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein

binding. Proc. Natl. Acad. Sci. USA 2009, 106, 16068-16073.

48. Vysabhattar, R.; Ganesh, K. N., Cyanuryl peptide nucleic acid: synthesis and DNA

complexation properties. Tetrahedron Lett. 2008, 49, 1314-1318.

141

49. Zhou, Z.; Bong, D., Small-Molecule/Polymer Recognition Triggers Aqueous-Phase

Assembly and Encapsulation. Langmuir 2013, 29, 144-150.

50. Ma, M.; Paredes, A.; Bong, D., Intra- and intermembrane pairwise molecular

recognition between synthetic hydrogen-bonding phospholipids. J. Am. Chem. Soc.

2008, 130, 14456-8.

51. Ma, M.; Gong, Y.; Bong, D., Lipid Membrane Adhesion and Fusion Driven by

Designed, Minimally Multivalent Hydrogen-Bonding Lipids. J. Am. Chem. Soc.

2009, 131, 16919-16926.

52. Ma, M.; Bong, D., Protein assembly directed by synthetic molecular recognition

motifs. Org. & Biomolec. Chem. 2011, 9, 7296-9.

53. Ma, M.; Bong, D., Determinants of cyanuric acid and melamine assembly in water.

Langmuir 2011, 27, 8841-53.

54. Ma, M.; Bong, D., Directed peptide assembly at the lipid-water interface

cooperatively enhances membrane binding and activity. Langmuir 2011, 27, 1480-

55. Ma, M.; Bong, D., Controlled fusion of synthetic lipid membrane vesicles. Accounts

of chemical research 2013, 46, 2988-97.

56. Felsenfeld, G.; Davies, D. R.; Rich, A., Formation of a three-stranded

polynucleotide molecule. J. Am. Chem. Soc. 1957, 79, 2023-2024.

57. Pilch, D. S.; Levenson, C.; Shafer, R. H., Structural analysis of the (dA)10.2(dT)10

triple helix. Proc Natl Acad Sci U S A 1990, 87, 1942-6.

58. Betts, L.; Josey, J. A.; Veal, J. M.; Jordan, S. R., A nucleic acid triple helix formed

by a peptide nucleic acid-DNA complex. Science 1995, 270, 1838-41.

142

59. Soto, A. M.; Loo, J.; Marky, L. A., Energetic contributions for the formation of

TAT/TAT, TAT/CGC(+), and CGC(+)/CGC(+) base triplet stacks. J. Am. Chem.

Soc. 2002, 124, 14355-14363.

60. Duca, M.; Vekhoff, P.; Oussedik, K.; Halby, L.; Arimondo, P. B., The triple helix: 50

years later, the outcome. Nucleic Acids Res. 2008, 36, 5123-38.

61. Shin, D.; Tor, Y., Bifacial Nucleoside as a Surrogate for Both T and A in Duplex

DNA. J. Am. Chem. Soc. 2011, 133, 6926-6929.

62. Branda, N.; Kurz, G.; Lehn, J.-M., JANUS WEDGES: a new approach towards

nucleobase-pair recognition. Chem. Commun. 1996, 1996, 2443-2444.

63. Chen, H.; Meena; McLaughlin, L. W., A Janus-Wedge DNA Triplex with A-W1-T

and G-W2-C Base Triplets. J. Am. Chem. Soc. 2008, 130, 13190-13191.

64. Largy, E.; Liu, W.; Hasan, A.; Perrin, D. M., Base-pairing behavior of a carbocyclic

Janus-AT nucleoside analogue capable of recognizing A and T within a DNA

duplex. Chembiochem : a European journal of chemical biology 2013, 14, 2199-

208.

65. Mittapalli, G. K.; Reddy, K. R.; Xiong, H.; Munoz, O.; Han, B.; De Riccardis, F.;

Krishnamurthy, R.; Eschenmoser, A., Mapping the landscape of potentially

primordial informational oligomers: oligodipeptides and oligodipeptoids tagged

with triazines as recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2470-7.

66. Hysell, M.; Siegel, J. S.; Tor, Y., Synthesis and stability of exocyclic triazine

nucleosides. Organic & Biomolecular Chemistry 2005, 3, 2946-2952.

67. Menor ‑ Salván, C.; Ruiz ‑ Bermejo, D.; Guzmán, M. I.; Osuna ‑ Esteban, S.;

Veintemillas ‑ Verdaguer, S., Synthesis of pyrimidines and triazines in ice:

143

implications for the prebiotic chemistry of nucleobases. Chem.-Eur. J. 2009, 15,

4411-4418.

68. Liedl, T.; Olapinski, M.; Simmel, F. C., A surface-bound DNA switch driven by a

chemical oscillator. Angewandte Chemie 2006, 45, 5007-10.

69. Krishnan, Y.; Simmel, F. C., Nucleic acid based molecular devices. Angewandte

Chemie 2011, 50, 3124-56.

70. Yoshida, W.; Yokobayashi, Y., Photonic Boolean logic gates based on DNA

aptamers. Chemical communications 2007, 195-7.

71. Saghatelian, A.; Volcker, N. H.; Guckian, K. M.; Lin, V. S.; Ghadiri, M. R., DNA-

based photonic logic gates: AND, NAND, and INHIBIT. Journal of the American

Chemical Society 2003, 125, 346-7.

72. Chakraborty, S.; Mehtab, S.; Krishnan, Y., The predictive power of synthetic

nucleic acid technologies in RNA biology. Accounts of chemical research 2014, 47,

1710-9.

144

APPENDIX A

Peptide Ligation and RNA Cleavage

via an Abiotic Template Interface

This chapter is reproduced with permission from J. Am. Chem. Soc., 2015, 137, 3751-3754.

145

A.1 Overview

We report herein DNA- and RNA- templated chemical transformation of bifacial peptide nucleic acid (bPNA) fragments directed by an abiotic triplex hybrid interface. Assembly of one bPNA strand with two unstructured oligo T/U strands enables facile insertion of DNA and RNA template sites within partially folded nucleic acids; this template topology is not easily accessed through native base-pairing. Triplex hybridization of reactive bPNA fragments on DNA and RNA templates is shown to catalyze amide bond ligation and controlled bPNA chain extension. RNA-templated oxidative coupling of bPNA fragments is found to result in the emergence of ribozyme cleavage function, thus establishing a connection between engineered and native reaction sites. These data demonstrate the use of new topologies in nucleic acid- templated chemistry that could serve as chemically sensitive DNA and RNA switches.

A.2 Introduction:

DNA recognition has been exploited to direct the chemistry of native and artificial1-5 macromolecules. Watson−Crick base-pairing can be taken out of context to enable group transfer,6-8 nucleic acid detection,9 chemical library selection,10 and self-replication.11 In addition, base-pairing templates can code for synthesis of non-native scaffolds12-14 and multisite macromolecular modification.15-17 The abiotic base triple interface between bifacial peptide nucleic acid (bPNA)18-21 and nucleic acids provides an opportunity to explore alternative template topologies. Triazine22-30 bases in bPNA simultaneously dock two oligo T/U strands to form an obligate triplex hybrid (Figure A.1). 146

Figure A.1. bPNA structure and experimental design. (Top) T/U-rich tracts can bind bPNA via melamine- derivatized lysine (M*). (Bottom) The four template topologies shown catalyze thioester exchange, fragment ligation, chain extension, and oxidative coupling with activation of ribozyme splicing function with bPNA substrates (dark lines).

147

Unlike conventional PNA, which dissociatively invades native structures, bPNA recognition is an associative process that unites non-interacting native domains. Though symmetric31-35 two-strand36-39 recognition of this type has no cognate in extant biology, amino and oxo 2,4,6-substituted triazines40, 41 recapitulate the Watson−Crick hydrogen- bonding patterns, fueling the speculative notion of triazine-derived precursors42 to DNA and RNA. We have previously demonstrated that bPNA hybridization can trigger DNA and

RNA chemistry.25 We report herein that hybridization can likewise trigger bPNA chemistry.

Single-stranded and partially structured DNA/RNA topologies were found to serve as templates to catalyze bPNA coupling and controlled chain extension (oligomerization) of bi-reactive bPNAs. Furthermore, integration of a template site into a ribozyme fold renders

RNA splicing dependent on oxidative ligation of bPNA; this could serve as a blueprint for chemically sensitive nucleic acid switches43, 44 and gates45, 46with applications in DNA/RNA nanotechnology.47 Overall, these data demonstrate readout and transformation of non- native macromolecules through an abiotic template interface in DNA/RNA template topologies that are not accessible via native base-pairing.

A.3 Results and Discussion

An n-mer of bPNA has the general form (EM*)n, wherein M* = melamine-modified lysine and E = glutamic acid (Figure 1). Hybridization of bPNA with dTnC4Tn DNA results in triplex stem loop (hairpin) structures with n thymine− melamine−thymine (TMT) base triples.

Binary, ternary, and quaternary bPNA−DNA complexes can be formed with one, two, or

19 three bPNAs bound to a single dTnC4Tn DNA strand. Successful preorganization of DNA into hairpin configurations with weakly binding 4mer bPNAs prompted investigation of

DNA hairpin templates as catalysts for native chemical ligation48, 49 of two bPNA 4mer fragments. Nominal background ligation of 4mer bPNAs 1 (C-terminal thioester) and 2 (N-

148

terminal cysteine) was observed over several days at 200 nM fragment concentration

(Figure A.2). Remarkably, a strong template effect was observed with the T10 hairpin DNA template, despite the modest binding affinity of 4mer bPNA. Consistent with the key role of the TMT interface, ligation yield dropped sharply with T→C substitutions in the DNA template. The reaction profile is consistent with product inhibition,50 further underscored by competitive inhibition of ligation with 10mer bPNA (Figure A.2). As the reaction progresses, a well-defined UV transition emerges at ∼50 °C, similar to the thermal stability observed with an 8-mer bPNA−DNA complex.

Figure A.2. DNA-template bPNA native ligation. (A) Native chemical ligation of 1 (Cbf-β- (EM*)4G-COSR2) and 2 (CM*(EM*)3G) (200 nM each) with dT10C4T10 (Cbf = carboxyfluorescein; R2 = (CH2)2SO3Na); β = β-alanine. (B) Ligation yield with template (left) and (right) inhibitor concentration indicated. 10mer (EM*)10 bPNA was used as inhibitor at 1000 nM fixed DNA template. Lines are drawn to guide the eye.

149

Notably, optimum reaction temperature is at the Tm of the 4mer−DNA complex (25 °C), with decreased rate at higher and lower temperatures, suggestive of the importance of dynamic complexation. Despite the modest DNA affinity of the fragments, the T10 hairpin template increased ligation rates by 2500 fold over background; increased effective molarity51 in the bPNA−DNA ternary complex can readily account for this rate acceleration.

DNA-templated native chemical ligation suggested the possibility of chain extension

(oligomerization) through multiple on-template couplings of bi-reactive bPNAs. This appeared reasonable as the thermal stability of bPNA−DNA complexes increases from binary to ternary to quaternary.19 Rapid cyclization52, 53 of bPNAs bearing N-terminal cysteine and C-terminal thioester functionality prompted investigation of cysteine-free amide coupling of bPNAs. Though direct peptide aminoacylation with thioesters is low yielding reaction in aqueous milieu, amino acid side chains can greatly influence reaction rate and yield.54 Accordingly, a 4mer bPNA (3) fitted with N-terminal glycine and C-terminal histidine thioester was prepared for on-template chain extension. The reaction mixture was spiked with 5 mol% thioester 1 (Figure A.3) to fluorescently label the products for

PAGE analysis. While background coupling was insignificant, ligation was observed on incubation with dTnC4Tn hairpin templates (n = 8, 10, 15, 18), with 25− 50% overall conversion wherein higher yields corresponded to the longer templates (SI, Figures S5 and S6). Furthermore, longer ligation products were observed with longer templates.

(Figure A.3) For n = 8 and 10, dimer was the dominant outcome, consistent with the notion that two 4mer bPNAs could fit on the template at once. Longer trimer and tetramer bPNA products from two and three on-template couplings were clearly detected as major products with the dT15C4T15 and dT18C4T18 templates, commensurate with higher order complex formation. The identity of the oligomers was confirmed by band isolation and

150

MALDI-MS. Though fluorescence labeling was used to image the gel, isolated bands yielded masses corresponding to the unlabeled, thioester hydrolyzed population. Thus, on-template, direct aminolysis of thioester fragments from ternary, quaternary, and apparent quaternary bPNA−DNA complexes leads to bPNA chain extension by virtue of the length-matching abiotic TMT interface.

Figure A.3. DNA-templated bPNA oligomerization. (Top) Peptide chain extension of 3 (GM*(EM*)3H-COSR2; R2 = (CH2)2SO3Na) with dTnC4Tn DNA. (Lower) Fluorescence-stained denaturing PAGE of extension reactions with hairpin template T-tract indicated. Product bands were identified as (a) hydrolyzed 3, (b) 3, (c) dimer, (d) trimer, and (e) tetramer by MALDI-MS (c−e are C-terminal acids).

151

Figure A.4. Template effect on bPNA native ligation. Native chemical ligation of bPNAs 1 and 2 using T10-hairpin, T10-duplex and T10-(duplex)2 templates. Rate increases with longer hairpin templates (left) and with duplex-organized templates (right). Lines are drawn to guide the eye.

Just as chain extension was higher yielding with longer templates, native chemical ligation of bPNA cysteine and thioester fragments was also significantly faster with T15 and T18 hairpin templates relative to T10 (Figure A.4). Two factors likely contribute to this effect: (1) length-enhanced binding of 4mers and (2) template preorganization by first coupling product, leading to enhanced binding and catalysis of subsequent fragments. To probe the effect of pre-structuring, duplex- organized DNA templates were tested in bPNA native chemical ligation. Indeed, ligation rates of bPNAs 1 and 2 increased as the T10 tracts were buttressed by one (T10-duplex) and two duplexes (T10 - (duplex)2). Duplex presentation of 152

the T-tracts likely increases coupling efficiency by decreasing the entropic cost of bPNA triplex hybridization (Figure A.4). Constraining both ends of the unstructured T-tracts in

T10-(duplex)2 results in further enhancement of ligation.

In addition to acceleration of amide bond coupling, oxidation of bPNA dithiol 4 with T10-

(duplex)2 was also significantly accelerated over background. (Figure A.5)

Figure A.5. DNA- and RNA-templated thiol oxidation. (Left) Oxidation of bPNA 4 (Ac- CM*(EM*)3C) alone (○) and with 50 mol% T10-(duplex)2 DNA (●), followed by PAGE. (Right) Oxidation of bPNAs 6 (Mpa-(EM*)4G) and 7 (EM*)4C alone (○) and with 12 mol% U-3-ribozyme (●), followed by Ellman’s test. Mpa = mercaptopropionamide. Lines are drawn to guide the eye.

While thiol oxidation is more facile than amide bond formation under these conditions, the duplex-constrained template limited products formed to dimeric and trimeric extension; in contrast, a wide range of oxidation products were formed off template (SI, Figures S10 and S11). Unlike amide bond chain extension (Figure A.3), three bPNA fragments may be oxidatively coupled on a T10 template; this is perhaps due to the increased flexibility of the

153

disulfide linkage. Successful catalysis with partially folded DNA templates prompted investigation of Un-template loops imbedded within RNA folds. This notion was tested using a minimal type I hammerhead ribozyme55 in which stem III was replaced with an

25 rU10CACAU10 loop (U-3-ribozyme). It was initially thought that the constrained U-loop would exhibit orientational bias with respect the RNA template; thus, two 4mer bPNAs bearing N-terminal (6) and C-terminal (7) thiols were prepared and studied. However, both thiols and their mixture, gave identical template-enhanced oxidation profiles with U-3- ribozyme (Figure A.5). Interestingly, the U-3-ribozyme was able to achieve a similar yield of oxidation at lower catalyst loading (12%) compared to the DNA (50%), suggesting a higher exchange rate off the RNA loop template, though more quantitative measurements are needed to pinpoint the origin of this difference.

The U-3-ribozyme sequence has ablated self-cleavage activity due to the loss of stem III structure. We have previously demonstrated that duplex stems in aptamers and ribozyme folds can be functionally replaced with bPNA triplex hybrid stems when base-pairing sequences are replaced with T/U tracts. This allows bPNA to be used as an allosteric switch for both aptamer affinity and ribozyme catalysis. We hypothesized that native nucleic acid function could report on the coupling of short bPNA fragments. This notion was tested using oxidative thiol coupling since amide bond ligation occurs on a time scale similar to RNA degradation. While 4mer bPNA (EM*)4 and bPNAs 4, 6, and 7 only weakly activate cleavage of the U-3- ribozyme (Figure A.6), oxidative coupling produces an ∼8mer bPNA disulfide product that binds more tightly to the template and strongly activates function.25 Oxidation conditions are more concentrated than those for ribozyme cleavage; thus, reactions were studied by dilution of partially oxidized samples into ribozyme cleavage conditions with 1 mM Mg2+. PAGE analysis of the reaction indicated the

154

formation of two RNA products upon 10% bPNA oxidation, which were identified as the tRNA fusion and the hammerhead ribozyme components (Figure A.6).

Figure A.6. Ribozyme-templated bPNA thiol oxidation leads to the emergence of ribozyme self-cleavage. (Top) U-3-ribozyme cleavage upon oxidation of bPNA thiols 6 and 7 and addition 2+ of Mg . (Bottom) Denaturing PAGE of RNA cleavage triggered by 4mer bPNA (EM*)4 and 10% oxidized 6 and 7, with full-length RNA (*), tRNA (<), and ribozyme (≪) cleavage products indicated, along with gel quantification, with the bPNA additives indicated. Lines are drawn to guide the eye.

Positive control experiments using 8mer bPNA (EM*)8 and fully oxidized and purified mixed disulfides of 6 and 7 gave higher yields of splicing. Ribozyme catalytic activity therefore may be used to report on fragment oxidative coupling, indicating two-way communication between an engineered abiotic template site and a native RNA splicing site. This connection makes possible functional selection56, 57 and optimization of template and redox-switchable ribozymes. 155

A.4 Conclusion

Overall, these data collectively demonstrate that effective molarity increases on DNA and

RNA templates can catalyze acyl transfer and oxidative coupling as well as chain extension of bPNA fragments. Insertion of template sites into folded nucleic acids is uniquely achieved through the thymine−melamine− thymine triplex interface with bPNA.

Native nucleic acid function can thus be linked with engineered reactivity through allostery and template effects using the abiotic TMT interface. It is anticipated that facile inclusion of partially folded template topologies in nucleic acid directed chemistry will have use in

DNA/RNA nanotechnology.47

A.5 Acknowledgement

Research was supported in part by NSF-DMR. Facilities were provided by The Ohio State

University.

156

A.6 References for Appendix A

1. Schmidt, J. G.; Nielsen, P. E.; Orgel, L. E., Information transfer from peptide

nucleic acids to RNA by template-directed syntheses. Nucleic Acids Research

1997, 25, 4797-4802.

2. Schmidt, J. G.; Christensen, L.; Nielsen, P. E.; Orgel, L. E., Information transfer

from DNA to peptide nucleic acids by template-directed syntheses. Nucleic Acids

Research 1997, 25, 4792-4796.

3. Zhan, Z. Y. J.; Lynn, D. G., Chemical amplification through template-directed

synthesis. Journal of the American Chemical Society 1997, 119, 12420-12421.

4. Goodwin, J. T.; Mehta, A. K.; Lynn, D. G., Digital and analog chemical evolution.

Accounts of chemical research 2012, 45, 2189-99.

5. Sharma, A. K.; Heemstra, J. M., Small-Molecule-Dependent Split Aptamer Ligation.

Journal of the American Chemical Society 2011, 133, 12426-12429.

6. Grossmann, T. N.; Seitz, O., DNA-catalyzed transfer of a reporter group. Journal

of the American Chemical Society 2006, 128, 15596-15597.

7. McKee, M. L.; Milnes, P. J.; Bath, J.; Stulz, E.; Turberfield, A. J.; O'Reilly, R. K.,

Multistep DNA-Templated Reactions for the Synthesis of Functional Sequence

Controlled Oligomers. Angewandte Chemie-International Edition 2010, 49, 7948-

7951.

8. McKee, M. L.; Evans, A. C.; Gerrard, S. R.; O'Reilly, R. K.; Turberfield, A. J.; Stulz,

E., Peptidomimetic bond formation by DNA-templated acyl transfer. Organic &

Biomolecular Chemistry 2011, 9, 1661-1666.

9. Silverman, A. P.; Kool, E. T., Detecting RNA and DNA with templated chemical

reactions. Chemical Reviews 2006, 106, 3775-3789.

157

10. Kleiner, R. E.; Dumelin, C. E.; Liu, D. R., Small-molecule discovery from DNA-

encoded chemical libraries. Chemical Society Reviews 2011, 40, 5707-5717.

11. Von Kiedrowski, G., A SELF-REPLICATING HEXADEOXYNUCLEOTIDE.

Angewandte Chemie International Edition in English 1986, 25, 932-935.

12. Li, Z. Y.; Zhang, Z. Y. J.; Knipe, R.; Lynn, D. G., DNA-catalyzed polymerization.

Journal of the American Chemical Society 2002, 124, 746-747.

13. Rosenbaum, D. M.; Liu, D. R., Efficient and sequence-specific DNA-templated

polymerization of peptide nucleic acid aldehydes. Journal of the American

Chemical Society 2003, 125, 13924-13925.

14. Lo, P. K.; Sleiman, H. F., Nucleobase-Templated Polymerization: Copying the

Chain Length and Polydispersity of Living Polymers into Conjugated Polymers. J.

Am. Chem. Soc. 2009, 131, 4182-4183.

15. Ura, Y.; Beierle, J. M.; Leman, L. J.; Orgel, L. E.; Ghadiri, M. R., Self-assembling

sequence-adaptive peptide nucleic acids. Science 2009, 325, 73-7.

16. Heemstra, J. M.; Liu, D. R., Templated synthesis of peptide nucleic acids via

sequence-selective base-filling reactions. J. Am. Chem. Soc. 2009, 131, 11347-9.

17. McHale, R.; O'Reilly, R. K., Nucleobase Containing Synthetic Polymers:

Advancing Biomimicry via Controlled Synthesis and Self-Assembly.

Macromolecules 2012.

18. Zeng, Y.; Pratumyot, Y.; Piao, X.; Bong, D., Discrete assembly of synthetic

peptide-DNA triplex structures from polyvalent melamine-thymine bifacial

recognition. J. Am. Chem. Soc. 2012, 134, 832-5.

19. Piao, X.; Xia, X.; Bong, D., Bifacial Peptide Nucleic Acid Directs Cooperative

Folding and Assembly of Binary, Ternary, and Quaternary DNA Complexes.

Biochemistry 2013, 52, 6313-6323. 158

20. Xia, X.; Piao, X.; Fredrick, K.; Bong, D., Bifacial PNA complexation inhibits

enzymatic access to DNA and RNA. Chembiochem : a European journal of

chemical biology 2013, 15, 31-36.

21. Xia, X.; Piao, X.; Bong, D., Bifacial peptide nucleic acid as an allosteric switch for

aptamer and ribozyme function. Journal of the American Chemical Society 2014,

136, 7265-8.

22. Arambula, J. F.; Ramisetty, S. R.; Baranger, A. M.; Zimmerman, S. C., A simple

ligand that selectively targets CUG trinucleotide repeats and inhibits MBNL protein

binding. Proc. Natl. Acad. Sci. USA 2009, 106, 16068-16073.

23. Vysabhattar, R.; Ganesh, K. N., Cyanuryl peptide nucleic acid: synthesis and DNA

complexation properties. Tetrahedron Lett. 2008, 49, 1314-1318.

24. Zhou, Z.; Bong, D., Small-Molecule/Polymer Recognition Triggers Aqueous-Phase

Assembly and Encapsulation. Langmuir 2013, 29, 144-150.

25. Ma, M.; Paredes, A.; Bong, D., Intra- and intermembrane pairwise molecular

recognition between synthetic hydrogen-bonding phospholipids. J. Am. Chem. Soc.

2008, 130, 14456-8.

26. Ma, M.; Gong, Y.; Bong, D., Lipid Membrane Adhesion and Fusion Driven by

Designed, Minimally Multivalent Hydrogen-Bonding Lipids. J. Am. Chem. Soc.

2009, 131, 16919-16926.

27. Ma, M.; Bong, D., Protein assembly directed by synthetic molecular recognition

motifs. Org. & Biomolec. Chem. 2011, 9, 7296-9.

28. Ma, M.; Bong, D., Determinants of cyanuric acid and melamine assembly in water.

Langmuir 2011, 27, 8841-53.

159

29. Ma, M.; Bong, D., Directed peptide assembly at the lipid-water interface

cooperatively enhances membrane binding and activity. Langmuir 2011, 27, 1480-

30. Ma, M.; Bong, D., Controlled fusion of synthetic lipid membrane vesicles. Accounts

of chemical research 2013, 46, 2988-97.

31. Felsenfeld, G.; Davies, D. R.; Rich, A., Formation of a three-stranded

polynucleotide molecule. J. Am. Chem. Soc. 1957, 79, 2023-2024.

32. Pilch, D. S.; Levenson, C.; Shafer, R. H., Structural analysis of the (dA)10.2(dT)10

triple helix. Proc Natl Acad Sci U S A 1990, 87, 1942-6.

33. Betts, L.; Josey, J. A.; Veal, J. M.; Jordan, S. R., A nucleic acid triple helix formed

by a peptide nucleic acid-DNA complex. Science 1995, 270, 1838-41.

34. Soto, A. M.; Loo, J.; Marky, L. A., Energetic contributions for the formation of

TAT/TAT, TAT/CGC(+), and CGC(+)/CGC(+) base triplet stacks. J. Am. Chem.

Soc. 2002, 124, 14355-14363.

35. Duca, M.; Vekhoff, P.; Oussedik, K.; Halby, L.; Arimondo, P. B., The triple helix: 50

years later, the outcome. Nucleic Acids Res. 2008, 36, 5123-38.

36. Shin, D.; Tor, Y., Bifacial Nucleoside as a Surrogate for Both T and A in Duplex

DNA. J. Am. Chem. Soc. 2011, 133, 6926-6929.

37. Branda, N.; Kurz, G.; Lehn, J.-M., JANUS WEDGES: a new approach towards

nucleobase-pair recognition. Chem. Commun. 1996, 1996, 2443-2444.

38. Chen, H.; Meena; McLaughlin, L. W., A Janus-Wedge DNA Triplex with A-W1-T

and G-W2-C Base Triplets. J. Am. Chem. Soc. 2008, 130, 13190-13191.

39. Largy, E.; Liu, W.; Hasan, A.; Perrin, D. M., Base-pairing behavior of a carbocyclic

Janus-AT nucleoside analogue capable of recognizing A and T within a DNA

160

duplex. Chembiochem : a European journal of chemical biology 2013, 14, 2199-

208.

40. Mittapalli, G. K.; Reddy, K. R.; Xiong, H.; Munoz, O.; Han, B.; De Riccardis, F.;

Krishnamurthy, R.; Eschenmoser, A., Mapping the landscape of potentially

primordial informational oligomers: oligodipeptides and oligodipeptoids tagged

with triazines as recognition elements. Angew. Chem. Int. Ed. 2007, 46, 2470-7.

41. Hysell, M.; Siegel, J. S.; Tor, Y., Synthesis and stability of exocyclic triazine

nucleosides. Organic & Biomolecular Chemistry 2005, 3, 2946-2952.

42. Menor‑ Salván, C.; Ruiz‑ Bermejo, D.; Guzmán, M. I.; Osuna‑ Esteban, S.;

Veintemillas‑ Verdaguer, S., Synthesis of pyrimidines and triazines in ice:

implications for the prebiotic chemistry of nucleobases. Chem.-Eur. J. 2009, 15,

4411-4418.

43. Liedl, T.; Olapinski, M.; Simmel, F. C., A surface-bound DNA switch driven by a

chemical oscillator. Angewandte Chemie 2006, 45, 5007-10.

44. Krishnan, Y.; Simmel, F. C., Nucleic acid based molecular devices. Angewandte

Chemie 2011, 50, 3124-56.

45. Yoshida, W.; Yokobayashi, Y., Photonic Boolean logic gates based on DNA

aptamers. Chemical communications 2007, 195-7.

46. Saghatelian, A.; Volcker, N. H.; Guckian, K. M.; Lin, V. S.; Ghadiri, M. R., DNA-

based photonic logic gates: AND, NAND, and INHIBIT. Journal of the American

Chemical Society 2003, 125, 346-7.

47. Chakraborty, S.; Mehtab, S.; Krishnan, Y., The predictive power of synthetic

nucleic acid technologies in RNA biology. Accounts of chemical research 2014, 47,

1710-9.

161

48. Roloff, A.; Seitz, O., Bioorthogonal reactions challenged: DNA templated native

chemical ligation during PCR. Chemical Science 2013, 4, 432-436.

49. Dawson, P. E.; Muir, T. W.; Clarklewis, I.; Kent, S. B. H., Synthesis of proteins by

native chemical ligation. Science 1994, 266, 776-779.

50. Dose, C.; Ficht, S.; Seitz, O., Reducing product inhibition in DNA-template-

controlled ligation reactions. Angewandte Chemie-International Edition 2006, 45,

5369-5373.

51. Li, X.; Liu, D. R., DNA-templated organic synthesis: nature's strategy for controlling

chemical reactivity applied to synthetic molecules. Angewandte Chemie 2004, 43,

4848-70.

52. Li, Y.; Yongye, A.; Giulianotti, M.; Martinez-Mayorga, K.; Yu, Y.; Houghten, R. A.,

Synthesis of cyclic peptides through direct aminolysis of peptide thioesters

catalyzed by imidazole in aqueous organic solutions. Journal of combinatorial

chemistry 2009, 11, 1066-72.

53. White, C. J.; Yudin, A. K., Contemporary strategies for peptide macrocyclization.

Nature chemistry 2011, 3, 509-24.

54. Payne, R. J.; Ficht, S.; Greenberg, W. A.; Wong, C.-H., Cysteine-Free Peptide and

Glycopeptide Ligation by Direct Aminolysis. Angewandte Chemie 2008, 120, 4483-

4487.

55. Murray, J. B.; Terwey, D. P.; Maloney, L.; Karpeisky, A.; Usman, N.; Beigelman,

L.; Scott, W. G., The structural basis of hammerhead ribozyme self-cleavage. Cell

1998, 92, 665-673.

56. Tuerk, C.; Gold, L., Systematic evolution of ligands by exponential enrichment:

RNA ligands to bacteriophage T4 DNA polymerase. Science 1990, 249, 505-510.

162

57. Ellington, A. D.; Szostak, J. W., In vitro selection of RNA molecules that bind

specific ligands. Nature 1990, 346, 818-822.

163

APPENDIX B

General Guidelines for Bacterial Cell Culture

164

B.1 Sterilization

 All glassware should be cleaned and rinsed with plenty amount of ddH2O, make

sure there is no residual detergent.

 All liquid needs to be either autoclave or sterile filtration.

 Autoclave requires 30 minutes sterilization time. For large volume liquid 40

minutes sterilization is recommended.

B.2 Stock solution preparation

Luria Broth:

25 g of Luria Broth powder in 1 L of distilled H2O.

Luria Broth Agar:

40 g of Luria Broth Agar powder in 1 L of distilled H2O

Table B.1 Antibiotic Solutions:

Antibiotic Stock concentration Storage condition Final concentration Ampicillin 50 mg/ml in water –20 °C 100 µg/mL (1/500) Chloramphenicol 34 mg/ml in ethanol –20 °C 170 µg/mL (1/200) Kanamycin 10 mg/ml in water –20 °C 50 µg/mL (1/200) Streptomycin 10 mg/ml in water –20 °C 50 µg/mL (1/200) Tetracycline HCl 5 mg/ml in ethanol –20 °C 50 µg/mL (1/100)

165

B.3 Pouring Agar plates

Always prepare agar plate next to the flame

 Let autoclaved LB-agar cool to ~50 °C, add proper amount of antibiotic solution

referring to the table above.

 Slowly pour ~25 mL of warm agar into 100 mm petri dish, avoid generating

bubbles. If bubbles form, quickly run flame over the agar surface.

 Place 5 dishes in one stack, and sit a 250 mL flask filled with hot water on top of

the plates to prevent water condensation on the lids.

 Once solidified, put dishes into the packing sleeve, and store upside down in cold

room. Plates with antibiotics have an average shelf life about 2 weeks.

B.4 Streak bacteria cells onto agar plate

 Pre-warm agar plates in 37 °C incubator for 20 minutes.

 Sterilize inoculation loop with flame, remove a small amount of bacteria culture

(single colony, glycerol stock or liquid culture) with the inoculation loop, and

immediately streak the cells onto the agar plate following the pattern below.

3 4

166

 Seal the plate, and place it upside down in the 37 °C incubator overnight to

develop colonies.

167

APPENDIX C

Preparation of Competent E. Coli Cells

168

C.1 Introduction

For laboratory protein expression, E. Coli expression system is the most frequently used due to the advantages of easy selection, short induction and comparative high expression yields. Foreign genes that embed unique selection markers, most commonly antibiotic resistance markers, are introduced into E. Coli cells, this process of horizontal gene transfer is termed transformation1. The precondition for E. Coli transformation is competence, which is the capability of the bacterial cells to uptake external genetic materials. There are two categories of competent cells: i) chemically competent cells uptake foreign genes with the facilitation of calcium chloride2, which enhances plasmid adherence through the electrostatic interactions between divalent calcium cations and anionic phosphate backbones; ii) electrocompetent cells work through electroporation3, which utilizes electrical pulses to generate pores on bacterial membrane for genetic material permeation.

For routine protein expression and purification, transformation efficiency of chemically competent cells is adequate; for ligation and SELEX library screening that requires optimal transformation efficiency, electrocompetent cells are recommended. Occasionally, in order to be able to select single colony, chemically competent cells can be used instead of electrocompetent cells.

Described in details below are protocols developed in Bong lab for preparing both chemically competent cells and electrocompetent cells.

169

C.2 Protocol 1: Preparing Chemically Competent E. Coli Cells

C.2.1 Material

Glycerol stock of E. Coli cells

500mL sterilized Luria Broth

LB-agar plate

*No antibiotic for BL21 cells, 100μg/mL streptomycin for DH10B cells

1000 mL shaker flask

500 mL centrifuge bottle

15 mL Falcon tube

Sterilized inoculation loop

1.5 mL microcentrifuge tubes

50 mL and 5 mL serological pipettes

37 °C incubator and 37 °C shaker

2M MgCl2, 1M CaCl2, 50% Glycerol and 200mL distilled H2O, all sterilized.

C.2.2 Procedure

Day 1

 Pre-warm LB-agar plate at 37 °C for 20 minutes.

 Gently scrape the E. Coli glycerol stock with inoculation loop, then streak cells on

to the pre-warmed agar plate.

 Keep the plate upside down (agar side up), seal the plate with Para-film and

incubate in the 37 °C incubator overnight.

Day 2

170

 Pick one single colony from the overnight agar plate with inoculation loop, seed

the cells into 5 mL sterilized Luria Broth in a 15mL Falcon tube.

 Slightly loosen the cap to allow sufficient air exchange, then inoculate the 5 ml

culture overnight in a bench-top shaker at 37 °C 225 rpm.

Day 3

 Take 2.5 mL sterilized Luria Broth in a separate tube as blank.

 Add 4 mL overnight culture into 500 mL sterilized Luria Broth in a 500 mL shaker

flask, loosely cover the flask with aluminum foil, then shake the cell culture in a

37 °C shaker at 225 rpm. Monitor the culture growth by measuring the optical

density at 600nm.

 During the inoculation, chill serological pipettes, centrifuge tubes in the cold room.

Prepare the following solutions and store in 4 °C. Turn on centrifuge and set

temperature at 4 °C

Table C.1 Equilibrium Buffer:

Reagent Volume (mL) [Concentration] (mM)

2M MgCl2 4.00 80.00

1M CaCl2 2.00 20.00

ddH2O 94.00 100.00

171

Table C.2 Storage Buffer

Reagent Volume (mL) [Concentration] (mM) 50% glycerol 2.00 20%

1M CaCl2 0.50 100.00

ddH2O 2.50

5.00

 Once the OD600 reaches 0.6, which indicates the cells are in mid-log growth phase,

remove the cells from shaker and immediately chill on ice for 10 minutes in the

cold room. From this point forward, keep cells cold.

 Transfer culture to pre-chilled 50mL centrifuge tubes. Pellet cells at 4000rpm for

10 minutes at 4°C.

 Carefully pour out the supernatant, invert tubes on a paper towel for 1 minute to

remove residual liquid.

 Re-suspend cell pellet in 50mL ice-cold Equilibrium buffer and incubate on ice for

5 minutes. Centrifuge at 4000rpm for 10min at 4 °C. Repeat this step one more

time.

 Remove the supernatant, invert the tube on paper towel for 1min to remove

residual equilibrium buffer.

 Re-suspend each pellet in 5mL ice cold Storage buffer, and make 100 μL

aliquots in 1.5 mL microcentrifuge tubes.

 Flash freeze all the cells in liquid nitrogen, then store in -80 °C freezer.

172

C.3 Protocol 2: Preparing Electrocompetent E. Coli Cells

C.3.1 Material

Glycerol stock of E. Coli cells

500mL sterilized Luria Broth

LB-agar plate

*No antibiotic for BL21 cells, 100μg/mL streptomycin for DH10B cells

1000 mL shaker flask

15 mL Falcon tube

500 mL centrifuge bottle

Sterilized inoculation loop

1.5 mL microcentrifuge tubes

50 mL and 5 mL serological pipettes

37 °C incubator and 37 °C shaker

1 L Sterilized 10% glycerol

C.3.2 Procedure

Day 1

 Pre-warm LB-agar plate at 37 °C for 20 minutes.

 Gently scrape the E. Coli glycerol stock with inoculation loop, then streak cells on

to the pre-warmed agar plate.

 Keep the plate upside down (agar side up), seal the plate with Para-film and

incubate in the 37 °C incubator overnight.

173

Day 2

 Pick one single colony from the overnight agar plate with inoculation loop, seed

the cells into 5 mL sterilized Luria Broth in a 15mL Falcon tube.

 Slightly loosen the cap to allow sufficient air exchange, then inoculate the 5 ml

culture overnight in a bench-top shaker at 37 °C 225 rpm.

Day 3

 Take 2.5 mL sterilized Luria Broth in a separate tube as blank.

 Add 4 mL overnight culture into 500 mL sterilized Luria Broth in a 500 mL shaker

flask, loosely cover the flask with aluminum foil, then shake the cell culture in a

37 °C shaker at 225 rpm. Monitor the culture growth by measuring the optical

density at 600nm.

 During the inoculation, chill serological pipettes, centrifuge tubes and 10% glycerol

in the cold room. Turn on centrifuge and set temperature at 4 °C.

 Once the OD600 reaches 0.6, which indicates the cells are in mid-log growth phase,

remove the cells from shaker and immediately chill on ice for 10 minutes in the

cold room. From this point forward, keep cells cold.

 Transfer culture to pre-chilled 500mL centrifuge tubes. Pellet cells at 4000rpm for

10 minutes at 4°C.

 Carefully pour out the supernatant, invert tubes on a paper towel for 1 minute to

remove residual liquid.

 Re-suspend cell pellet in 250mL ice-cold 10% glycerol and incubate on ice for 5

minutes. Centrifuge at 4000rpm for 10min at 4 °C. Repeat this step two more

times. Large volume and additional washing step is to ensure the removal of salts.

174

 Remove the supernatant, invert the tube on paper towel for 1min to remove

residual equilibrium buffer.

 Re-suspend each pellet in 50 mL ice cold 10% glycerol, and make 100 μL aliquots

in 1.5 mL microcentrifuge tubes.

 Flash freeze all the cells in liquid nitrogen, then store in -80 °C freezer.

175

C.4 References for Appendix C

1. Avery, O. T.; Macleod, C. M.; McCarty, M., Studies on the Chemical Nature of the

Substance Inducing Transformation of Pneumococcal Types : Induction of

Transformation by a Desoxyribonucleic Acid Fraction Isolated from

Pneumococcus Type Iii. The Journal of experimental medicine 1944, 79, 137-58.

2. Cohen, S. N.; Chang, A. C.; Hsu, L., Nonchromosomal antibiotic resistance in

bacteria: genetic transformation of Escherichia coli by R-factor DNA. Proceedings

of the National Academy of Sciences of the United States of America 1972, 69,

2110-4.

3. Dower, W. J.; Miller, J. F.; Ragsdale, C. W., High efficiency transformation of E.

coli by high voltage electroporation. Nucleic acids research 1988, 16, 6127-45.

176

APPENDIX D

Transformation of Competent E. Coli Cells

177

D.1 Introduction

As explained above, the capability of cells to uptake extracellular genetic material is called competence, and the whole process of importing foreign gene is termed transformation1.

In laboratory, we utilize the concept of competence and conduct transformation with the proper E. Coli strains to fit our needs of protein expression.

Transformation of chemically competent cells includes two steps: i) plasmid DNA forming complexes with Ca2+, and adhere to the cell surface; ii) heat shock of the cells to perturb the membrane integrity, thus plasmid is uptaken into the cell.

Transformation of electrocompetent cells is distinctly different, and requires less time. In brief, a pulse of electricity is applied through cell suspension mixed with plasmid DNA, due to the absence of salt, electrical pulse forces negatively charged plasmid DNA penetrate through the cell membrane.

Described below are the detailed protocols regarding transformation of both chemically competent cells and electrocompetent cells.

D.2 Protocol 1: Transforming Chemically Competent E. Coli Cells

 Slowly thaw cells on ice, add 1~2 μL of plasmid DNA (purified and sequence

confirmed) into 100μL cell suspension, gently tap the tube to mix, and incubate the

mixture on ice for 30 minutes.

 Heat shock the cell-plasmid mixture without any agitation at 42 ºC for 45 seconds

then quickly place the tube on ice for 2 minutes.

178

 Add 900 μL Luria Broth at room temperature to the mixture and inoculate the cells

at 37 ºC in a bench-top shaker at 225 rpm for 1 hour. This step is to recover the

cells from heat shock and allow them to generate antibiotic resistance.

 Spread 100 ~ 300 μl of cell culture with a sterile glass spreader onto a pre-warmed

agar plate made with appropriate antibiotic. You can also make a “low” plate (using

50μl of culture) if you are worried about getting too many colonies.

 Incubate the plate upside down overnight at 37 °C.

 Save the rest of the transformed cells in liquid culture at 4 °C. If nothing appears

on the plate, try again and plate a larger volume of cells.

D.3 Protocol 2: Transforming Electrocompetent E. Coli Cells

 Slowly thaw cells on ice, add 1~2 μL of plasmid DNA (purified and sequence

confirmed) into 100μL cell suspension, gently tap the tube to mix, and place tube

on ice.

 Clean and chill an electroporation cuvette on ice. For bacterial electroporation,

cuvettes with 0.1 cm and 0.2 cm gap width are most appropriate.

 Quickly transfer cell-plasmid mixture into the cuvette, clean and dry the cuvette

surface, and place the cuvette to the holder, push the holder to the right and apply

the pulse. When electroporation is done, a reading reflecting the conductivity will

appear, if reading is lower than 5.00, repeat the electroporation with less amount

of plasmid DNA.

 Quickly add 900 μL Luria Broth at room temperature to the mixture and inoculate

the cells at 37 ºC in a bench-top shaker at 225 rpm for 1 hour. This is to recover

the cells from electric shock and allow them to generate antibiotic resistance.

179

 Make 10-fold, 100-fold and 1000-fold cell culture dilutions, and plate 100 μL of both

original culture and dilutions onto 4 agar plates prepared with suited antibiotics.

 Seal the plates and incubate the plates upside down overnight at 37 °C.

 Save the rest of the transformed cells in liquid culture at 4 °C. If nothing appears

on the plate, try again and plate a larger volume of cells.

D.4 Protocol 3: Determine Transformation Efficiency

Transformation efficiency is evaluated by the number of colony forming units per μg of plasmid DNA. Generally:

 Pick a plate with proper density of single colonies; count the number of single

colonies.

 Calculate the transformation efficiency using the Equation 4.1

cfu Number of colonies × Dilution factor Equation 4.1 Transformation efficiency = = μg Mass of plasmid in μg

180

D.5 Reference for Appendix D

1. Chen, I.; Dubnau, D., DNA uptake during bacterial transformation. Nature reviews.

Microbiology 2004, 2, 241-9.

181

APPENDIX E

T7 RNA Polymerase Purification

182

E.1 Introduction

T7 RNA polymerase from Bacteriophage is a robust enzymatic machinery in making oligoribonucleotide with high processivity1 and fidelity.2 It overwhelms the native E. Coli RNA polymerase catalysis in vivo and has been integrated into E. Coli genome to enable large- scale mRNA and the downstream protein productions;3 besides its in vivo application, in vitro RNA synthesis using synthetic DNA template via T7 RNA polymerase has been widely applied to facilitate research in the aspect of RNA biology.4

E.2 Protocol 1: Purification of His-Tag T7 RNA Polymerase

E.2.1 Material

1 LB-agar plate with 100 μg/mL 50% Glycerol

Ampicillin 14.3 M β-Mercaptoethanol

1 L LB medium with 100 μg/mL 0.5 M EDTA

Ampicillin 1 M K2HPO4

Ni-NTA agarose beads (Qiagen #30210) 1 M KH2PO4

1 M Tris-HCl, pH 7.9 1 M IPTG

4 M NaCl Bradford reagent

2 M Imidazole SDS-PAGE reagents

183

E.2.2 Equipment and Consumables

500 mL centrifuge bottles 37 °C bench-top shaker

50 mL centrifuge tubes Fermenter

Empty gravity chromatography column Dialysis tubing

Inoculation loop Cell Centrifuge

15 mL and 50 mL Falcon tubes Swing bucket centrifuge

37 °C incubator Spectrophotometer

E.2.3 Solutions

Table E.1 Lysis Buffer (500 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 10.00 mL 20.00 mM 4 M NaCl 37.50 mL 300.00 mM 2 M Imidazole 1.25 mL 5.00 mM 50 % Glycerol 50.00 mL 5.0 % 14.3 M β-Mercaptoethanol 70.00 μL 2.00 mM ddH2O Up to 500 mL

Table E.2 Washing Buffer (200 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 4.00 mL 20.00 mM 4 M NaCl 25.00 mL 500.00 mM 2 M Imidazole 500.00 μL 5.00 mM 50 % Glycerol 40.00 mL 10.0 % 14.3 M β-Mercaptoethanol 28.00 μL 2.00 mM ddH2O Up to 200 mL

Table E.3 Elution Buffer 1 (100 mM Imidazole, 50 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 1.00 mL 20.00 mM 4 M NaCl 6.25 mL 500.00 mM 2 M Imidazole 2.50 mL 100.00 mM 50 % Glycerol 10.00 mL 10.0 % 14.3 M β-Mercaptoethanol 7.00 μL 2.00 mM ddH2O Up to 50 mL

184

Table E.4 Elution Buffer 2 (200 mM Imidazole, 50 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 1.00 mL 20.00 mM 4 M NaCl 6.25 mL 500.00 mM 2 M Imidazole 5.00 mL 200.00 mM 50 % Glycerol 10.00 mL 10.0 % 14.3 M β-Mercaptoethanol 7.00 μL 2.00 mM ddH2O Up to 50 mL

Table E.5 Elution Buffer 3 (300 mM Imidazole, 50 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 1.00 mL 20.00 mM 4 M NaCl 6.25 mL 500.00 mM 2 M Imidazole 7.50 mL 300.00 mM 50 % Glycerol 10.00 mL 10.0 % 14.3 M β-Mercaptoethanol 7.00 μL 2.00 mM ddH2O Up to 50 mL

Table E.6 Elution Buffer 4 (400 mM Imidazole, 50 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 1.00 mL 20.00 mM 4 M NaCl 6.25 mL 500.00 mM 2 M Imidazole 10.00 mL 400.00 mM 50 % Glycerol 10.00 mL 10.0 % 14.3 M β-Mercaptoethanol 7.00 μL 2.00 mM ddH2O Up to 50 mL

Table E.7 Elution Buffer 5 (500 mM Imidazole, 50 mL) Reagent Volume [Final concentration] 1 M Tris-HCl, pH 7.9 1.00 mL 20.00 mM 4 M NaCl 6.25 mL 500.00 mM 2 M Imidazole 12.50 mL 500.00 mM 50 % Glycerol 10.00 mL 10.0 % 14.3 M β-Mercaptoethanol 7.00 μL 2.00 mM ddH2O Up to 50 mL

Table E.8 Storage Buffer (2000 mL) Reagent Volume [Final concentration] 1 M K HPO 18.20 mL 2 4 20 mM, pH 7.9 1 M KH2PO4 1.84 mL 4 M NaCl 25.00 mL 100 mM 0.5 M EDTA 0.40 mL 0.10 mM Glycerol 1000 mL 50 % 14.3 M β-Mercaptoethanol 0.84 mL 2.00 mM ddH2O Up to 2000 mL

185

E.2.4 Procedures

Day 1

 Gently streak cells onto LB-agar plate from glycerol stock of BL21-pLysS

transformed with plasmid containing gene of native T7 RNA polymerase. Plasmid

possesses ampicillin resistance gene and the gene is under T3 promoter.

 Incubate the plate upside down in 37 °C incubator overnight.

Day 2

 Pick a single colony from the overnight LB-agar plate with the inoculation loop,

seed the colony into 5 mL LB medium with ampicillin in a 15 mL Falcon tube.

 Loosen the cap and shake the Falcon tube at 37 °C 225 rpm overnight in the

bench-top shaker.

Day 3

 Aliquot 5 mL of LB medium, put aside as control.

 Transfer the 5 mL overnight culture into 1000 mL LB medium, shake the cell culture

at 37 °C 225 rpm in the fermenter.

 Closely monitor the optical density at 600 nm using spectrophotometer. When the

reading reaches 0.6, add 1 mL of 1 M IPTG to induce T7 RNA polymerase

expression.

 After 4 hour expression induction, transfer the cell culture into 500 mL centrifuge

bottles, pellet the cells at 4 °C 5,000 rpm for 15 minutes.

 Re-suspend the cell pellet in 50 mL Lysis buffer, then centrifuge again with the

same settings as the previous step. Store the cell pellet in -20°C freezer.

Day 4

 Re-suspend cell pellet in ~30mL Lysis buffer, normally most cells lysate

themselves, however running through homogenizer is still necessary.

186

 Rinse the homogenizer with ice-cold Lysis buffer, then pump cell suspension

through homogenizer until cell lysate is less viscous and almost clear.

 Centrifuge at 15,000 rpm for 15minutes at 4 °C to remove the cell debris, transfer

the supernatant into another centrifuge tube, and centrifuge again under the same

condition for 15 minutes. This step further separates the protein from the debris

and ensures the smooth gravity flow during the purification. Keep the cleared

lysate on ice.

 Aliquot 6 mL Ni-NTA (~3 mL resin), incubate with 20 mL Lysis buffer via end-over-

end rotation for 15min at 4 °C, then pellet Ni-NTA beads for 5 minutes in swing-

bucket centrifuge at 4,500 rpm 4 °C, discard the supernatant slowly without

disturbing the beads. Repeat the wash step for two more times.

 Combine the cell lysate with the equilibrated Ni-NTA resin, rotate end-over-end for

2 hours at 4 °C.

 Spin down the resin, transfer the supernatant into a clean 50 mL Falcon tube, label

as “Unbound”, if the total purification yield is low, this fraction can re-apply to affinity

column.

 Re-suspend the resin in 40 mL cold washing buffer, rotate for 10 minutes at 4 °C ,

centrifuge down the resin and carefully transfer the liquid into another tube marked

as “Wash 1”. Perform additional 2 washes, and label the supernatant as “Wash 2”

and “Wash 3” individually.

 Transfer the resin into an empty gravity column, drain the residual buffer by gravity,

then start imidazole step elution. Apply 2 mL of Elution buffer 1 to the column and

collect the eluent in 1 ml fraction; once the level of Elution buffer 1 hits the resin

bed, start adding 2 ml of Elution buffer 2, collect the eluent and repeat this step

until all five Elution buffer have been applied.

187

 Identify fractions containing protein by mixing 5 μL of the eluent with 100 μL of

Bradford reagent5, the intensity of blue color indicates the protein concentration

within the fraction.

 Take aliquots of protein positive fractions, denature with 2x Laemmli buffer

supplemented with 10% β-Mercaptoethanol and resolve on a 4% - 12% SDS

acrylamide gel. Stain the gel with coomassie blue to identify pure fractions (E.3

Protocol 2)

 Combine all the pure fractions and dialyze against 1 L Storage buffer overnight.

Day 5

 Change the reservoir into another 1 L of Storage buffer and dialyze for another 6

hours.

 Recover the concentrated protein solution from dialysis tubing, make 50 μL protein

aliquots into microcentrifuge tubes, and then flash freeze with liquid nitrogen and

store in the -80 °C deep freezer.

 Leave one tube of the protein out for concentration determination via Bradford

Assay and activity using in vitro transcription (E.4 Protocol 3).

E.3 Protocol 2: SDS-PAGE for Purity Analysis

E.3.1 Material

29:1 Acrylamide: Bis-acrylamide 0.5 M Tris-HCl, pH 6.8 solution 10x Tris-glycine-SDS buffer

10% Ammonium persulfate (APS) 2x Laemmli buffer

Tetramethylethylenediamine (TEMED) β-Mercaptoethanol

20% SDS Bio-Safe stain (Bio-Rad 161-0786)

1.5 M Tris-HCl, pH 8.8

188

E.3.2 Apparatus

Digital heating block Protein electrophoresis system

E.3.3 Procedure

 Prepare resolving gel according to the table below. T7 RNA polymerase is ~ 99

kDa in molecular weight, hence 12% resolving gel is most appropriate.

 Pour resolving gel solution into the plate until the gel surface is about 8 mm away

from the bottom of the wells.

 Seal the gel with a layer of ddH2O to generate a smooth and flat gel surface.

 Once the resolving gel is solidified, prepare 4% stacking gel solution.

 Remove the water layer with filter paper.

 Layer the stacking gel solution on top of solidified resolving gel, then insert the

comb.

 Prepare protein samples while the gel is setting. First, add 14.3 M β-

Mercaptoethanol to 2x Laemmli buffer to a final concentration of 10% v/v. Then

mix the buffer with protein samples at the ratio of 1:1, heat samples at 100 °C for

10 minutes to ensure total denaturation, then chill on ice.

189

Table E.9. Resolving Gel Recipes Gel Volume Gel % Reagent [Final] MW 5mL 10mL 15mL 20mL ddH2O 2.921 5.842 8.763 11.684 29:1 Acryl : Bis 0.75 1.50 2.25 3.00 6% 1.5M Tris pH 1.25 2.50 3.75 5.00 0.375M 60 kDa 6% 8.8 ~ 600 kDa 20% SDS 0.025 0.05 0.075 0.10 0.1% 10% APS 0.05 0.1 0.15 0.2 0.1% TEMED 0.004 0.008 0.012 0.016 -- ddH2O 2.671 5.342 8.013 10.684 29:1 Acryl : Bis 1.00 2.00 3.00 4.00 8% 1.5M Tris pH 1.25 2.50 3.75 5.00 0.38M 40 kDa 8% 8.8 ~ 400 kDa 20% SDS 0.025 0.05 0.075 0.10 0.1% 10% APS 0.05 0.1 0.15 0.2 0.1% TEMED 0.004 0.008 0.012 0.016 -- ddH2O 2.421 4.842 7.263 9.684 29:1 Acryl : Bis 1.25 2.50 3.75 5.00 10% 1.5M Tris pH 1.25 2.50 3.75 5.00 0.38M 20 kDa 10% 8.8 ~300 kDa 20% SDS 0.025 0.05 0.075 0.10 0.1% 10% APS 0.05 0.1 0.15 0.2 0.1% TEMED 0.004 0.008 0.012 0.016 -- ddH2O 2.171 4.342 6.513 8.684 29:1 Acryl : Bis 1.50 3.00 4.50 6.00 12% 1.5M Tris pH 1.25 2.50 3.75 5.00 0.38M 10 kDa 12% 8.8 ~ 200 kDa 20% SDS 0.025 0.05 0.075 0.10 0.1% 10% APS 0.05 0.1 0.15 0.2 0.1% TEMED 0.004 0.008 0.012 0.016 -- ddH2O 1.796 3.592 5.388 7.184 29:1 Acryl : Bis 1.875 3.75 5.625 7.50 15% 1.5M Tris pH 1.25 2.50 3.75 5.00 0.38M 3 kDa 15% 8.8 ~ 100 kDa 20% SDS 0.025 0.05 0.075 0.10 0.1% 10% APS 0.05 0.1 0.15 0.2 0.1% TEMED 0.004 0.008 0.012 0.016 --

Table E.10 Stacking Gel Recipes Gel Volume Gel % Reagents [Final] 1mL 2mL 3mL 4mL ddH2O 0.609 1.218 1.827 2.436 29:1 Acryl : Bis 0.125 0.25 0.375 0.50 5% 4% 0.5M Tris pH 6.8 0.25 0.50 0.75 1.00 0.125M Stacking 20% SDS 0.005 0.010 0.015 0.020 0.1%

10% APS 0.01 0.02 0.03 0.04 0.1% TEMED 0.001 0.002 0.003 0.004

190

 Make 10x Tris-glycine-SDS buffer following the recipe below:

Table E.11. 10x Tris-Glycine-SDS Buffer Reagent Mass [Final concentration] Tris Base 30.30 g 0.25 M Glycine 144.00 g 1.92 M SDS 10.00 g 1%

ddH2O Fill up to 1 L

 Load samples as well as the protein ladder onto the gel, electrophorese with

constant current of 25 mA until blue dye migrates into resolving gel, then increase

the current to 60 mA until the blue dye reaches the bottom of the gel.

 Take of the gel, rinse twice with distilled water to remove residual SDS, then stain

the gel with Bio-Safe coomassie stain for 30 min, de-stain with distilled water until

the background is low and clear bands appear.

 Measure the T7 RNA polymerase concentration by UV absorption at 280 nm and

6 5 - calculate the protein molarity using extinction coefficient ε280nm= (1.4±0.1) × 10 M

1cm-1.

 Purity is estimated at 95%, lower molecular weight impurities at 75 kDa and 23

kDa are known proteolytic fragments.7, 8 The proteolytic cleavage reduces the

processivity of the fragments even though the polymerase activity is partially

retained,7, 8 when calculate yield and units of specific activity, mass of the

fragments needs to be subtracted from the total mass.

191

Figure E.1. Purity evaluation through SDS-PAGE. Top: positive fractions confirmed using Bradford assay during elution and dialyzed stock; Bottom: (from left to right) cell lysate crude, supernatant after clearance of cell debris, unbound, wash 1, wash 2, wash 3 and three different loadings from dialyzed stock.

192

E.4 Protocol 3: T7 RNA Polymerase Activity Test

E.4.1 Material

2 M HEPES-KOH, pH 7.5 5 μM DNA template

2 M MgCl2 Inorganic pypophosphatase

0.5 M Spermidine-HCl T7 RNA polymerase

1 M DTT 10x TBE buffer rNTP set – 25 mM for each nucleotides Agarose

2 mg/mL BSA 10 mg/mL Ethidium Bromide

Blue juice nucleic acid sample buffer 0.5 M EDTA

E.4.2 Equipment

Digital heating block

Horizontal gel electrophoresis system

UV transmission gel scanner

193

E.4.3 DNA Template Design for T7 Transcription

T7 RNA polymerase is highly promoter specific, a single mutation in the promoter sequence significantly reduces its catalytic activity possibly due to enfeebled transcription

9 initiation resulted from loose promoter binding and a larger KM. Besides promoter sequences, +1 and +2 position on the template as well play crucial role in proper initiation, it is suggested that positioning dC at both sites warrants the robust transcription start.10

In laboratory, the 17-base promoter sequence is most commonly used in in vitro transcription with T79. Further, though T7 RNA polymerase contains helicase activity, which unwinds double stranded genome to form the transcription bubble and ensures enzymatic read-through in vivo, studies have shown that T7 shows normal initiation with single-stranded DNA template that only has upstream double-strand promoter region4, 11.

Promoter: 5’-TAATACGACTCACTATA-3’ (non-coding strand)

3’-ATTATGCTGAGTGATATCC------template------5’ (coding strand) +1

E.4.4 Procedure

 Estimate the total mass of purified T7 RNA polymerase using extinction coefficient,

molar mass and volume of protein stock.

OD g Total mass = 280 × Stock volume in Liter × 99,000 ε mole

 T7 RNA polymerase generally has typical specific activity of 300,000 to 500,000

units/mg6. According to the total mass of the purified T7, dilute purified T7 into

about 50 units/μL, which is the commercial concentration of T7 RNA polymerase

from New England Biolabs (Catalog # M0251S).

194

 Melt 2 g of agarose in 100 mL 1x TBE buffer, add 2 μL of Ethidium bromide solution

to reach a final concentration of 0.2 μg/mL. Cast the gel when the gel solution is

sufficiently cooled.

 Make 10x High Yield Transcription Buffer

Table E.12. 10x High Yield Transcription Buffer Reagent Volume [Final concentration] 2 M HEPES-KOH, pH 0.50 mL 1 M 7.5

2 M MgCl2 50.00 μL 100 mM 0.5 M Spermidine-HCl 40.00 μL 20 mM 1 M DTT 0.40 mL 400 mM

RNase-free H2O 10.00 μL 1.00 mL

Store the buffer in -20 °C freezer and the buffer is generally good for 2 weeks,

multiple freeze-thaw cycles will expedite oxidation of DTT.

 Set up two parallel 50 μL in vitro transcriptions with commercial enzyme and

purified enzyme.

Table E.13. in vitro T7 Transcription Protocol Reagent Volume [Final concentration] 10x High Yield Transcription Buffer 5.00 μL 1x 1 M DTT 0.50 μL 10 mM

2 M MgCl2 0.65 μL 26 mM 25 mM rNTP set 15.00 μL 7.5 mM each 1 unit/μL Inorganic pyrophosphatase 0.50 μL 0.5 unit 5 μM DNA template 5.00 μL 500 nM 50 unit/μL T7 RNA polymerase 5.00 μL 5 unit/μL 2 mg/mL BSA 2.50 μL 0.1 mg/mL

RNase-free H2O 15.85 μL 50.00 μL

195

Incubate the reaction at 37 °C for 90 minutes. During the course of transcription,

take 5 μL aliquots of reaction mixture and quench into 0.5 M EDTA, this will

efficiently chelate the catalytic Mg2+. Monitoring transcription product through the

time course will help in assess the overall transcription processivity.

 Mix quenched reaction aliquots with blue juice nucleic acid sample buffer at 4:1

ratio, then load onto the agarose gel. Apply constant voltage at 10 V/cm for 1~2

hours until the front bromophenol blue dye migrates to ¾ of the whole gel.

 Scan the gel with UV transmission scanner.

Figure E.2. T7 RNA polymerase activity estimation. Two in vitro transcriptions were performed parallel with same condition, time points were taken to evaluate enzymatic performance thoroughly at initiation and elongation phase. Left: commercial T7 RNA polymerase from New England Biolabs; Right: purified T7 RNA polymerase, which showed a slightly higher activity.

 Estimate the specific activity by comparing the RNA yields. According to the gel

shown above, purified T7 RNA polymerase has slightly higher catalytic activity

compare to the commercial T7.

196

E.5 References for Appendix E

1. Studier, F. W.; Moffatt, B. A., Use of bacteriophage T7 RNA polymerase to direct

selective high-level expression of cloned genes. Journal of molecular biology 1986,

189, 113-30.

2. Sydow, J. F.; Cramer, P., RNA polymerase fidelity and transcriptional proofreading.

Current opinion in structural biology 2009, 19, 732-9.

3. Rosenberg, A. H.; Lade, B. N.; Chui, D. S.; Lin, S. W.; Dunn, J. J.; Studier, F. W.,

Vectors for selective expression of cloned DNAs by T7 RNA polymerase. Gene

1987, 56, 125-35.

4. Milligan, J. F.; Groebe, D. R.; Witherell, G. W.; Uhlenbeck, O. C.,

Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA

templates. Nucleic acids research 1987, 15, 8783-98.

5. Kruger, N. J., The Bradford method for protein quantitation. Methods in molecular

biology 1994, 32, 9-15.

6. King, G. C.; Martin, C. T.; Pham, T. T.; Coleman, J. E., Transcription by T7 RNA

polymerase is not zinc-dependent and is abolished on amidomethylation of

cysteine-347. Biochemistry 1986, 25, 36-40.

7. Ikeda, R. A.; Richardson, C. C., Enzymatic properties of a proteolytically nicked

RNA polymerase of bacteriophage T7. The Journal of biological chemistry 1987,

262, 3790-9.

8. Muller, D. K.; Martin, C. T.; Coleman, J. E., Processivity of proteolytically modified

forms of T7 RNA polymerase. Biochemistry 1988, 27, 5763-71.

9. Martin, C. T.; Coleman, J. E., Kinetic analysis of T7 RNA polymerase-promoter

interactions with small synthetic promoters. Biochemistry 1987, 26, 2690-6.

197

10. Weston, B. F.; Kuzmine, I.; Martin, C. T., Positioning of the start site in the initiation

of transcription by bacteriophage T7 RNA polymerase. Journal of molecular

biology 1997, 272, 21-30.

11. Maslak, M.; Martin, C. T., Kinetic analysis of T7 RNA polymerase transcription

initiation from promoters containing single-stranded regions. Biochemistry 1993,

32, 4281-5.

198

APPENDIX F

Drosophila Schneider 2 Cells

Culture and Protein Expression

199

F.1 Introduction

Drosophila melanogaster Schneider 2 (S2) cell line1 is considered a valuable alternative host for mammalian protein expression. Advantages of recombinant protein expression via Drosophila S2 system include but are not limited to: i) High density cell culture in suspension can be easily attained, easy cultural conditions at room temperature;1 ii)

Expression is tightly controlled under metallothionein promoter, which is only inducible via heavy metal addition,2 suitable for expressing concentrated heterologous protein in short period of time, minimizing lethal effects and proteolysis;3, 4 iii) Introduction of recombinant gene through plasmid transfection, cells can stably transfected with multiple copies of foreign gene and express heterologous protein in a continuous bioprocess, compare to lytic Baculovirus expression system only feasible for transient expression;3, 5 iv) Most importantly, S2 cells are able to deliver nearly authentic mammalian and viral proteins with proper type and level of post-translational modifications due to the apposite signal recognition from eukaryotes.6-8

Drosophila melanogaster S2 cells are easy to culture, neither CO2 nor antibiotics are needed. Cells can be effortlessly adapted to serum free condition, ease the cultural expense and purification process by avoiding using huge amount of FBS. Below are the detailed protocols for Drosophila melanogaster S2 cell culture, and recombinant protein purification utilizing Drosophila expression system.

200

F.2 Materials

F.2.1 Cell Culture Reagents

Schneider’s Drosophila medium (Life Technology Cat # 21720-024)

Fetal Bovine Serum, US origin, heat inactivated.

Sf-900 II Serum Free Medium (Life Technology Cat # 10902-096)

F.2.2 Buffers and Stock Solutions

100 mg/mL Hygromycin B 500 mM CuSO4

5 mg/mL MTT solution in DPBS 1 M Tris-Cl, pH 7.8

MTT solvent Nonidet P-40

2M HEPES-NaOH, pH 7.05-7.12 4x Laemmli sample buffer

0.1M Na2HPO4 4%-16% SDS acrylamide gel

5M NaCl β-mercaptoethanol

2M CaCl2 Bio-safe coomassie stain

F.2.3 Consumables and Apparatus

Centrifuge

T-25 cm2 flask, or T-75 cm2 and T-162 cm2 flask

15 mL Falcon tubes

Serological pipettes

Cryovials

12-well culture plate, 96-well culture plate

M5 Spectrophotometer

201

F.3 Protocol 1: General Culture of Drosophila melanogaster S2

Cells

F.3.1 Thawing Cells From Frozen Stock:

 Warm up the complete Schneider’s Drosophila medium supplemented with 10%

heat inactivated FBS to room temperature. Aliquot 5 mL medium into a 15 mL

sterile Falcon tube.

 Remove one cryovial from liquid nitrogen storage tank, and quickly thaw the cells

in a 37 °C water bath in about 1-2 minutes.

 Decontaminate the surface of the vial with 70% ethanol when cells are about 85%

thawed, and then transfer the vial into laminar flow hood.

 Transfer the thawed cell stock into the 5 mL complete medium that prepared in the

first step. Pipette up and down twice to suspend the cells, then centrifuge at 130 g

for 3 minutes. This step is to remove DMSO in the freezing medium9.

 Carefully remove the supernatant, re-suspend the cell pellet with 5 mL complete

culture medium and transfer the cell suspension into a T-25 cm2 cell culture flask.

 Incubate the cells in a 28 °C incubator without CO2 and non-humidified, loosen the

cap to ensure sufficient air exchange.

 Drosophila melanogaster S2 cells generally forms a loosely attached monolayer

first when using a new culture flask, cells will gradually grow into suspension once

the density is about 2×106 cells/mL. Cells are sensitive to density9, so keep track

of cell density and viability and only subculture when the density is above 2×106

cells / mL and 95% viable, subculture cells into density > 5×105 cells/mL.

202

F.3.2 Subculturing Drosophila melanogaster S2 Cells

 To maintain cells, subculture when the density is above 6×106 cells/mL, cell

viability will decrease with density is below 5×105 cells/mL or above 20×106

cells/mL.

 Gently tap the bottom of the culture flask to dislodge the cells, use serological

pipettes to break up the cell clumps. Determine the cell density and viability using

Trypan blue and a hemacytometer.

 Calculate and subculture cells into density ≥ 5×105 cells/mL. Simply calculate the

dilution factor then dilute cell culture with fresh medium, no centrifugation is

needed. Medium carried over to new cell passage is called conditioned medium, it

contains metabolites that can facilitate the cell growth.

 For T-25 cm2 flask, the total culture volume should not exceed 6 mL. However, if

large cell culture volume is needed, cells can be subcultured into T-75 cm2 flask

for maximum 15 mL or T-162 cm2 flask for 50 mL.

F.3.3 Generating Frozen Stocks of Drosophila melanogaster S2 Cells

 After 3-4 passages and cell density is ≥ 6×106 cells/mL and viability is >90%,

subculture 1/3 of the cells to maintain continuous culture. Transfer the other 2/3 of

cell culture into a Falcon tube, pellet cells at 130 g for 3 minutes.

 Carefully pipette out the supernatant, DO NOT discard the supernatant, which is

the conditioned medium needs to be supplemented into freezing medium.

 Wash the cell pellets once with complete medium, this is to further remove the

dead cells.

 During 2nd centrifugation process, prepare freezing medium following the recipe

below:

203

Table F.1. Freezing Medium Recipe

Reagent Volume (mL) Concentration (v/v %) Fresh complete medium w/ 10% FBS 2.25 mL 45% Conditioned medium 2.25 mL 45% DMSO 0.50 mL 10% 5.00 mL

 Cells need to be frozen at a density of ≥ 1×107 viable cells /mL, calculate and re-

suspend cell pellet with appropriate amount of freezing medium. If cells were

grown in large culture vessels, scale up the freezing medium.

 Aliquot cell suspension into cryovials, generally fill the cryovials up to 75% full (1

mL into 1.5 mL vials, and 1.5 mL into 2 mL vials), label the vials on the cap and

place them in a programmable freezing device, which allows gradual cooling that

minimizes cell damage.

 To set up a programmable freezing device, place a cardboard tube divider in a

Lock-Lock box, fill the box with 2-propanol until the liquid level is slightly higher

than the cell suspension level inside the cryovials. High heat capacity and low

freezing point of 2-propanol will ensure cells undergo a slowly cooling process and

2-propanol will remain as liquid that makes downstream vial transfer expedient.

 Place the whole gradual freezing device containing cryovials in the -80 °C deep

freezer overnight. Transfer the vials into liquid nitrogen storage tank for long term

storage10.

204

F.4 Protocol 2: Hygromycin Kill Curve for Hygromycin B Selection

F.4.1 Overview

In order to select transfected S2 cells from surrounding non-transfected cells, transfection is conducted with the presence of a selection vector. pCoHygro selection vector that carries the hygromycin B resistant gene is commonly used. Hygromycin B belongs to aminoglycoside family of antibiotics, it functions by ceasing ribosomal translocation in both prokaryotes and eukaryotes.11-15 Cells co-transfected with hygromycin resistant gene produces hygromycin B phosphotransferase that inactivates hygromycin B via phosphorylation.

F.4.2 Procedures

Before transfection, it is important to establish the optimal hygromycin B concentration selection by generating a kill curve for blank S2 cells. Kill curve experiment is a dosage- dependent assay, where cells are exposed to different amount of antibiotic and cell viability is monitored through the course of a week, optimal concentration is usually the minimum concentration that is needed to kill all the cells in one week.

Note: With any new cell line, new antibiotics, even a new frozen stock, optimal concentration might be different, always perform a kill curve before setting a selection concentration.

 Seed cells into 2 12-well cell culture plates, cell density at 1×106 cells/mL, and

each well seed 0.6 mL of cell suspension.

 Make hygromycin B stock solution in ddH2O at the concentration of 100 mg/mL.

Keep hygromycin B stock in 4 °C refrigerator protected from exposure to light.

205

 Add hygromycin into each well following the pattern below, incubate two plates at

regular S2 culture condition, and monitor cell viability daily.

 On Day 3 and Day 7 assay the percentage viability of cells in each well using MTT

assay.

Control 25 μg/mL 50 μg/mL 100 μg/mL

150 μg/mL 200 μg/mL 250 μg/mL 300 μg/mL

Control

350 μg/mL 400 μg/mL 450 μg/mL 500 μg/mL

 Add 1.0 mL MTT solvent into each well to dissolve the purple-colored formazan,

incubate at room temperature for 10 minutes. Then transfer 100 μL aliquots into a

96-well plate for absorption readings.

 Calculate percentage viability via comparison to the control well; plot the dosage

dependent viability curve to determine the optimal concentration. An examples plot

is shown below in Figure F.1

 Optimal dosage is determined as the minimum concentration that kills all cells in 7

days; High dosage is determined as the concentration showed evident cytotoxicity

in 2-3 days.

206

100 7-day Viability % 3-day Viability %

)

( 80

i 60

a 40

e 20

0 0 100 200 300 400 500 600 Concentration (mg/mL)

Figure F.1. Hygromycin B kill curve result. As indicated in the chart, 100 μg/mL hygromycin B led to 100% cell death in 7 days, thus can be set as optimal selection concentration.

F.5 Protocol 3: Calcium Phosphate Transfection of pMT/V5-HER2-

His A into Drosophila melanogaster S2 Cells

F.5.1 Overview

Calcium phosphate transfection utilizes the electrostatic interaction between calcium cation and DNA anionic phosphate backbone to form fine precipitations that can be uptaken into the cells.

Note: 1. Plasmids pMT/V5-HER2ECD-His A and selection vector pCoHygro are needed

for co-trasnfection

207

2. Plasmids have to be free from phenol, chloroform and sodium acetate, avoid high

salt and dissolve all plasmid in ddH2O.

F.5.2 Procedure

Day 1

 Prepare cells for transfection by seeding 3×106 S2 cells in Schneider’s Drosophila

medium supplemented with 10% FBS into a 35 mm culture dish, cell density should

be around 1×106 cells/mL.

 Culture cells under regular condition, check cell density 12-24 hours post-seeding,

viable cell density should be between 2×106 to 4×106 cells/mL; if cell density

doesn’t reach desired number, culture for additional 6-12 hours, or re-seed the

plate with cells in mid-log growth phase.

Day 2

 Make Ca2+-DNA mixture in one 1.5 mL microcentrifuge tube:

Table F.2. Ca2+-DNA Mixture Reagent Volume Concentration/ Mass

2 M CaCl2 36.00 μL 240 mM 1 μg/μL pMT/V5-HER2ECD-His A 19.00 μL 19.00 μg 1 μg/μL pCoHygro selection vector 1.00 μL 1.00 μg

Sterilized H2O 244.00 μL 300.00 μL

208

 Make 2× HEPES-Buffered Saline in another microcentrifuge tube:

Table F.3. 2x HEPES-Buffered Saline Reagent Volume Concentration/ Mass 2 M HEPES-NaOH, pH 7.05-7.1216 7.50 μL 50.00 mM

0.1 M Na2HPO4 4.50 μL 1.50 mM 5 M NaCl 16.80 μL 280.00 mM

Sterilized H2O 271.20 μL 300.00 μL

 Slowly add Ca2+-DNA mixture into 2× HEPES-Buffered Saline drop wise over the

course of 2 minutes with continuous vortexing. This ensures the formation of fine

precipitations that are necessary for efficient transfection.

 Incubate the mixture at room temperature for 30 minutes, fine precipitation should

be observable under microscope.

 Add totally 600 μL precipitation containing solution drop wise into cell culture dish,

gently swirl the dish to mix after each drop.

 Incubate cells in 28 °C incubator for 16 hours.

Day 3

 Dislodge cells from the culture dish by pipetting, centrifuge cells at 130 g for 3

minutes to remove calcium phosphate solution.

 Wash cells twice by re-suspending cell pellet in complete Schneider’s Drosophila

medium containing 10% FBS, then centrifuge at 130 g for 3 minutes.

 Replate cells into the transfection plate, add apposite amount of fresh medium

according to viable cell count, continue to incubate cell at 28 °C for 48 hours.

209

Day 5:

 Dislodge and then pellet cells at 130 g for 3 minutes, re-suspend cells in complete

growth medium at the density of 1×106 cells/mL, divide cell suspension into 2 equal

parts.

 Add CuSO4 to a final concentration of 500 μM into one part of the cell suspension

to induce HER2 expression (see protocol 6); add hygromycin B stock solution to

another part of the cell suspension to reach the optimal selection concentration

that is determined via kill curve assay, and start the process for selecting stably

transfected S2 cells.

F.6 Protocol 4: Stable Transfection of S2 Cells

 Following the last step in “protocol 3”, replace selection medium containing optimal

concentration of hygromycin B every 4-5 days depending on the density and

viability of the cells. To replace selection medium, simply dislodge cells, pellet cells

via centrifugation and re-suspend cells in fresh selection medium.

 In about 3-4 weeks, strong resistant cells will appear; replate the resistant cells

into new plate and only subculture into selection medium when cell density reaches

6×106-20×106 cells/mL. At this point, try to scale up the cell culture volume for the

subsequent protein expression and purification.

210

F.7 Protocol 5: Adapting Stably Transfected S2 Cells

After successfully selecting for stably transfected S2 cells, adapting resistant cells into serum free culture condition can further simplify the purification process. Expression of transfected recombinant gene will produce extracellular domain 1-4 of HER2 (human epidermal growth factor 2) with a C-terminal hexa-histidine affinity tag in secreted form, expression and purification under serum free condition can amplify affinity tag binding specificity due to the absence of large amount of albumin and other serum proteins.

In order to use serum free condition, stable S2 cells need to be gradually adapted into serum free medium.

 Subculture stable S2 cells into Sf-900 II serum free medium supplemented with 8%

FBS, cell density needs to be ≥5×105 cells/mL. Incubate cells in 28 °C incubator

until cell density reaches 6×106-20×106 cells/mL.

 Subculture cells again, meanwhile reduce the percentage of FBS to 6%; repeat

this process of gradually decreasing FBS percentage until cells remain viable and

in mid-log growth phase when using 100% Sf-900 II medium.

F.8 Protocol 6: HER2ECD(1-4) Expression and Purification via S2

Cells

 Following “Protocol 3”, after 48 hours post-induction, pellet cells at 130 g for 3

minutes, collect the supernatant since it contains the secreted HER2 extracellular

domain.

 Gradually add (NH4)2SO4 to the supernatant to 80% saturation, incubate the

mixture on ice for 20-30 minutes, this is to ensure efficient precipitation of secreted

protein.

211

 Prepare Cell Lysis buffer using the recipe below:

Table F.4. Lysis Buffer Recipe Reagents Volume Concentration 1 M Tris-Cl, pH 7.8 50.00 μL 50 mM 5 M NaCl 30.00 μL 150 mM Nonidet P-40 10.00 μL 1.0 %

ddH2O 910.00 μL 1.00 mL

 Add 50 μL of Cell Lysis buffer to the cell pellet, vortex to mix and incubate at room

temperature for 10-15 minutes. Then vortex again and pellet nuclei and cell debris

at 16,000 g for 10 minutes.

 Take aliquot of cleared cell lysate, mix with 4x Laemmli buffer supplemented with

20% β-mercaptoethanol at 3:1 ratio, boil at 100 °C for 10 minutes then quickly chill

on ice, label the sample as “lysate”.

 Pellet (NH4)2SO4 precipitation at 15,000g for 10 minutes at 4 °C, dissolve the

precipitation in DPBS then mix with 4x Laemmli buffer supplemented with 20% β-

mercaptoethanol at 3:1 ratio, boil at 100 °C for 10 minutes then quickly chill on ice,

label the sample as “secreted”.

 Load samples and protein ladder onto gel, and later on stain with Bio-safe

Coomassie blue to visualize.

212

F.9 References for Appendix F

1. Schneider, I., Cell lines derived from late embryonic stages of Drosophila

melanogaster. Journal of embryology and experimental morphology 1972, 27,

353-65.

2. Bunch, T. A.; Grinblat, Y.; Goldstein, L. S., Characterization and use of the

Drosophila metallothionein promoter in cultured Drosophila melanogaster cells.

Nucleic acids research 1988, 16, 1043-61.

3. Johansen, H.; van der Straten, A.; Sweet, R.; Otto, E.; Maroni, G.; Rosenberg, M.,

Regulated expression at high copy number allows production of a growth-inhibitory

oncogene product in Drosophila Schneider cells. Genes & development 1989, 3,

882-9.

4. Angelichio, M. L.; Beck, J. A.; Johansen, H.; Ivey-Hoyle, M., Comparison of several

promoters and polyadenylation signals for use in heterologous gene expression in

cultured Drosophila cells. Nucleic acids research 1991, 19, 5037-43.

5. Bishop, D. H., Gene expression using insect cells and viruses. Current opinion in

biotechnology 1990, 1, 62-7.

6. Ivey-Hoyle, M., Recombinant gene expression in cultured Drosophila

melanogaster cells. Current opinion in biotechnology 1991, 2, 704-7.

7. Ivey-Hoyle, M.; Culp, J. S.; Chaikin, M. A.; Hellmig, B. D.; Matthews, T. J.; Sweet,

R. W.; Rosenberg, M., Envelope glycoproteins from biologically diverse isolates of

immunodeficiency viruses have widely different affinities for CD4. Proceedings of

the National Academy of Sciences of the United States of America 1991, 88, 512-

213

8. Ivey-Hoyle, M.; Rosenberg, M., Rev-dependent expression of human

immunodeficiency virus type 1 gp160 in Drosophila melanogaster cells. Molecular

and cellular biology 1990, 10, 6152-9.

9. Rogers, S. L.; Rogers, G. C., Culture of Drosophila S2 cells and their use for RNAi-

mediated loss-of-function studies and immunofluorescence microscopy. Nature

protocols 2008, 3, 606-11.

10. Simcox, A., Progress towards Drosophila epithelial cell culture. Methods in

molecular biology 2013, 945, 1-11.

11. Gonzalez, A.; Jimenez, A.; Vazquez, D.; Davies, J. E.; Schindler, D., Studies on

the mode of action of hygromycin B, an inhibitor of translocation in eukaryotes.

Biochimica et biophysica acta 1978, 521, 459-69.

12. Gritz, L.; Davies, J., Plasmid-encoded hygromycin B resistance: the sequence of

hygromycin B phosphotransferase gene and its expression in Escherichia coli and

Saccharomyces cerevisiae. Gene 1983, 25, 179-88.

13. Borovinskaya, M. A.; Shoji, S.; Fredrick, K.; Cate, J. H., Structural basis for

hygromycin B inhibition of protein biosynthesis. Rna 2008, 14, 1590-9.

14. Cabanas, M. J.; Vazquez, D.; Modolell, J., Dual interference of hygromycin B with

ribosomal translocation and with aminoacyl-tRNA recognition. European journal of

biochemistry / FEBS 1978, 87, 21-7.

15. Cabanas, M. J.; Vazquez, D.; Modolell, J., Inhibition of ribosomal translocation by

aminoglycoside antibiotics. Biochemical and biophysical research communications

1978, 83, 991-7.

16. Kingston, R. E.; Chen, C. A.; Rose, J. K., Calcium phosphate transfection. Current

protocols in molecular biology / edited by Frederick M. Ausubel ... [et al.] 2003,

Chapter 9, Unit 9 1.

214

APPENDIX G

Protein Expression via Baculovirus System

215

G.1 Introduction

G.1.1 Overview

Baculovirus protein expression system is helper independent viral system that is employed to express recombinant proteins from many different sources, including human proteins.1

This system utilizes baculovirus infection cycle (Figure G.1), replaces non-essential viral genes with recombinant genes under strong promoters, yields comparatively large amount of proteins.

Autographa californica nuclear polyhedrosis virus (AcNPV) is the most studied baculovirus strain,2-4 its polyhedrin-encoding gene under the very strong late polh promoter is overexpressed at the late stage of viral infection for packaging occluded virus before insect

Lysis.1 It has been reported that polyhedrin is the major protein in occlude viruses which could take up to 30-50% of the total insect protein mass at the end of the infection.5 In recombinant baculovirus, polyhedrin gene in AcNPV is replaced by gene encoding heterologous protein; infection of recombinant baculovirus to sf9 cells will generate occlusion body negative viruses that overexpress recombinant protein instead.

Protein expression via baculovirus system has many advantages: i) baculovirus genome is double-stranded DNA package into rod-shaped nucleocapsid6 that can vary in size without interfering baculovirus life cycle, it is generally 130 kbp in length and contains hundreds of open reading frames, this character makes baculovirus capable of accommodating large protein-encoding genes that cannot be processed by bacteria

216

system; ii) BEVS is suitable for parallel expression of multiple proteins or multi-subunit proteins, which can substantially promotes eukaryotic protein complex structural and functional investigation;7 iii) Proteins from baculovirus system are nearly authentically modified, such like glycosylation, phosphorylation, acylation as well as proteolytic cleavage;8-10 iv) Baculovirus system can achieve high levels of recombinant expression compare to other eukaryotic expression systems, viral infection generates budding viruses that expand infection exponentially, and strong polh promoter will facilitate the recombinant protein accumulation during the late phase of infection;11-14 v) Compare to

Drosophila expression system, protein production timeline is significantly shortened, mainly because cell culture scale-up and virus stock generation can be proceeded simultaneously, and no selection for stable transfection is required.

In summary, baculovirus system is versatile and basically holds no limitation on the categories of protein being expressed; it produces protein in native/close to native state, provides extensive support for antibody generation and anti-cancer therapeutics.14

G.1.2 Bac-to-Bac HBM TOPO Expression System

Multiple methods are available for generating recombinant baculovirus. One well recognized method employ co-transfection of a transfer vector that imbeds foreign gene and baculovirus genome deficient in infectivity; homologous recombination inside insect

217

Viral Genome replication

Viral Genome Budding replication Original Digestion by mid-gut cell Baculovirus

Endocytosis Infection expansion Budding

Original Original Digestion by mid-gut cell baculovirus Baculovirus Occluded Virus

Endocytosis Cell lysis Infection expansion

Late stage infection

Occluded Virus

Occlusion body formation Cell lysis

Late stage infection

Transfection Transfection Cell lysis Protein release

Recombinant Cell lysis Baculovirus Occlusion body Protein release formation Recombinan Recombinant t baculovirus Baculovirus Secondary infection

Recombinant protein expression

Endocytosis Secondary infection

Recombinant p rotein Figure G.1. Baculovirus life cycle in vivo and in vitro. Top: native baculoviruses are protectedexpress ion by occlusion body, which is majorly composedEn ofd ocpolyheytosis drin proteins. Occluded viruses once ingested by larva, occlusion body becomes soluble and releases viral DNA into the cell. Viral DNA replicates inside host nuclei, and is subsequently packaged and budded out of host cell; budding viruses spread infection area through secondary infection; only at very late phase of infection, polh promoter (orange) is turned on and initiate overexpression of polyhedrin proteins (green), which is used in forming occlusion body before insect cells lysis. Bottom: Laboratory recombinant baculovirus, polyhedrin-encoded gene is replaced by target heterologous gene (red); gene replacement has minimum effect on virus life cycle until the very late phase of infection, instead of expression polyhedrin proteins, viruses are occlusion-negative and the heterologous protein will be overexpressed and can be harvested after cell lysis. Note: If secretion signal is fused to recombinant gene, protein can be harvested before cell lysis, reduce the risk of protease degradation.

218

cell nuclei will generate positive recombinant baculovirus that is able to infect adjacent cells, followed by plaque assays positive strain can be separated and cultured into a high infectivity titer.14-16 Although well established, this method generally produces low infectivity viruses that require multiple rounds of plaque assays to amplify.

Bac-to-Bac HBM TOPO expression system is more advanced in: i) More efficient cloning via TOPO isomerase; ii) Tn7 transposition enables recombinant viral genome purification through bacteria cells;17 iii) High infectivity titer is generated after transfection; iv) Honey bee mellitin secretion signal facilitates protein secretion out of host cells before cell lysis.

G.1.3 TOPO Cloning

Type I topoisomerase breaks one DNA strand of a supercoiled double-stranded DNA during replication and transcription, it helps unwind DNA and release tension on the

DNA.18 The tyrosyl oxygen in the active site attacking the phosphorus on the DNA backbone initiates strand-breakage, produces enzyme-DNA conjugate through the covalent phosphotyrosine link. Once the strand unwinds, they are re-joined through another transesterification .19, 20

In this protocol, blunt-end TOPO cloning is used to insert gene encoding HER2 extracellular domain 1-3 into the pFastBac vector. TOPO isomerase I is covalently linked onto the linearized vector backbone, activated and ready to ligate blunt-end PCR product.

G.1.4 Tn7 Transposition

Transposition, compare to homologue recombination, is a more sequence specific gene recombination. Tn7 transposition utilizes E. coli Tn7 transposon, which specifically recognizes attTn7 sequence on E. coli genome,21 resulted in gene insertion is exactly 25 bp downstream of attTn7 sequence.21 In this protocol, pFastBac transfer vector with

219

inserted HER2 extracellular domain 1-3 (HER2ECD(1-3)) will be transformed into

DH10Bac E. coli strain, where Tn7 transposition will occur that exchange the β- galacosidase gene on the internal baculovirus genome with HER2ECD(1-3), the successfully recombined gene will show negative result in blue-white screening due to the deficiency of β-galacosidase to digest Bluo-Gal.

Gene recombination via E.coli system through transposition, rather than homologue recombination in insect cell host significantly shorten the time to generate recombinant baculoviruses, basically all transfected bacmid are the recombinant version thus positive infectious virus can be acquired soon after transfection.

G.1.5 Honey Bee Melitin Secretion Signal Peptide

Have recombinant protein secreted into culture medium and harvested before cell lysis can simplify the purification process, and most importantly avoid proteolytic degradation upon intracellular protease release during lysis. Although the extracellular domain for

HER2 possibly contains secretion signal, addition of secretion signal to its N-terminus can facilitate protein transportation to endoplasmic reticulum,22 which is the traffic center for protein secretion. N-terminus fusion of honey bee melitin (HBM) signal peptide can facilitate the recombinant protein secretion23 yields up to 5-fold increase in protein purification yield.

220

G.2 Protocol 1: General Methods in Sf9 Cell Culture

G.2.1 Material

SF-900 II Serum Free Medium 28 °C incubator, no CO2, non-humidified

Sf9 cells, adapted to Sf-900 II SFM 125 mL shaker flask

Trypan blue Plate shaker

Hemacytometer Serological pipette

Cryovials DMSO

G.2.2 Recover Frozen Sf9 Cells in SF-900 II SFM

 Pre-warm SF-900 II SFM to 28 °C or room temperature. Transfer 25 mL medium

into a 125 mL sterile shaker flask in the laminar hood.

 Take one tube of frozen sf9 cells, quickly thaw the frozen stock in a 37 °C water

bath in about 1-2 minutes.

 Once the cells are about 80% thawed, decontaminate the vial surface with 70%

ethanol and transfer the vial into laminar hood.

 Triturate and aseptically transfer all the cell suspension into the shaker flask.

 Loosen the shaker flask cap to ensure sufficient air exchange. Incubate the cells

at 28 °C in the incubator for 4 days on the plate shaker. Orbital shaking speed

should be set between 125 rpm to 150 rpm.

 On day 5, check the cell density and viability using trypan blue stain and

hemacytometer. If viable cell density exceeds 2×106 cells/mL, subculture by

seeding cells into a new 125 mL shaker flask at with SF-900 II SFM at 4×105 viable

cells/mL in a volume of 30 mL; if cell density is lower than 2×106 cells/mL, pellet

cells at 130 g for 3 minutes and re-suspend cells with 25 mL SF-900 II SFM and

221

culture for another 2-3 days until the density reaches 2×106 cells/mL, then

subculture following the procedure mentioned above.

G.2.3 Maintain Sf9 Cell Culture

 Subculture cells when viable cell density reaches 2×106 cells/mL with the seeding

density as 4×105 viable cells/mL.

 Cells can be scaled up by sub-culturing into a larger volume of medium.

G.2.4 Cryopreservation of Sf9 Cells

 Cells can be cryopreserved once cells are maintained at 2×106 viable cells/mL and

have gone through at least 3 passages. It is necessary to scale up the culture

volume since cryopreservation requires density to be 10× higher compare to

regular cell culture.

 Count cell density, which should exceed 2×106 viable cells/mL. Transfer cell

suspension into 50 mL Falcon tubes, remove culture medium by centrifugation at

130 g for 3 minutes. Carefully transfer conditioned medium (supernatant) in to

another tube.

 Prepare freezing medium following the recipe below:

Table G.1 Freezing Medium Recipe

Reagent Concentration SF-900 II SFM conditioned 46.25% SF-900 II SFM 46.25 DMSO 7.5%

 Re-suspend cell pellet using freezing medium to reach a density of 2×107 viable

cells/mL.

 Transfer 1 mL of cell suspension into 1.5 mL cryovials, place vials into 2-propanol

bath and freeze cell stocks in a -80 °C freezer slowly overnight.

 Transfer frozen stocks into liquid nitrogen storage tank the second day.

222

G.3 Protocol 2: Acquire HER2ECD(1-3)-encoding Sequence via

PCR of pMT/V5-HER2ECD Plasmid with Designed Primers

DNA sequence encoding HER2 extracellular domain 1-3 has been determined following known crystal structure and article24, totally 1527 bp and 509 amino acids long, starting with Serine 1 to Asparagine 509.

G.3.1 Material

Primer set for PCR reaction PCR thermocycler

Q5 High fidelity DNA polymerase PCR tube

Q5 reaction buffer 1.0% agarose gel with ethidium bromide

Q5 GC enhancer Gel scanner

10 mM dNTPs set QIAquick PCR clean-up kit

G.3.2 Procedure

 Design PCR primer set in order to amplify sequence of domain 1-3 from plasmid

used in Drosophila system.

 Generally, forward primer should have the same sequence as the coding strand

from 5’, while the reverse primer should be complementary to the coding strand

from 3’. The length of primers is generally from 18 bases to 25 bases, primers too

short will cause insufficient annealing, as too long will have undesired template

binding and secondary structure. Further, to ensure specific annealing, try to have

primer sequence starts and ends with G and C.

223

Reverse Primer

5' 3' Coding strand

3' 5' Non-coding strand

Forward Primer

Figure G.2. PCR primer set design scheme

 In order to achieve decent PCR result, check the primer Tm using Tm Calculator

from NEB website: http://tmcalculator.neb.com/#!/. Tm difference of primers

should stay within 5 °C.

 For PCR reactions on gene fragments that require high accuracy during

amplification, Taq DNA polymerase from Thermus aquaticus doesn’t fulfill the

fidelity requirement, it has an average single-base substitution frequency of 1 for

each 9000 bases incorporated, as well as a frame-shift error at the frequency of

1/4100025. To attain protein with correct sequence, Q5 high fidelity DNA

polymerase is used in this protocol, which is a fusion protein with high DNA

template binding affinity and has accurate proof reading activity. Set up PCR

reaction following the recipe below:

Table G.2 HER2ECD(1-3) PCR Cloning Protocol

Reagents Volumn (μL) Final 5× Q5 Reaction buffer 10.00 1× 5× Q5 GC enhancer 10.00 1× 10 mM dNTPs 1.00 200 μM 10 μM HER2 forward primer 2.50 0.5 μM 10 μM HER2 reverse primer 2.50 0.5 μM 0.5 ng/μL pMT/V5-HER2ECD plasmid 2.00 1 ng 2 unit/μL Q5 High fidelity DNA polymerase 0.50 1 unit

ddH2O Up to 50.00

224

 Amplify using the following thermocycling parameters:

Table G.3 PCR Thermocycle Step Temperature (°C) Time (second) Initial denaturation 98.0 60 Denaturation 98.0 60 35 cycles Annealing 67.0 30 Elongation 72.0 30 Polish 72.0 120 Hold 4.0 ----

 Analyze PCR reaction on 1.0% agarose gel with ethidium bromide. Extracellular

domain 1-3 of HER2 contains totally 509 amino acids, which is 1527 bp after PCR.

Gel image in Figure. 3 indicates the correct PCR product was produced majorly.

 Purify PCR reaction with QIAquick PCR clean-up kit, and measure the PCR

product concentration in ng/μL. Measurement was taken by UV spectrometer at

260 nm, concentration is calculated using Equation G.1

Equation G.1 dsDNA in ng⁄μL = OD260 × Dilution factor × 50 ng⁄μL

225

Figure. G.3. PCR product yield examination with 1.0% agarose gel in 1× TBE, stained with ethidium bromide. Lane 1: DNA ladder; Lane 2: PCR reaction mixture after thermocycling, PCR product co-migrate with 1500 bp standard, indicating production of correct PCR product; Lane 3: pMT/V5-HER2ECD plasmid.

226

G.4 Protocol 3: TOPO Cloning HER2ECD(1-3) into pFastBac/HBM

Vector and Selection for Correctly Oriented Insertion.

G.4.1 Overview

For TOPO cloning, it requires the mole ratio of PCR insert: TOPO vector at 1:1 to 2:1.

Calculate the amount of PCR insert needed in the reaction following the rational described below.

 1 μL pFastBac TOPO vector is needed in the reaction according to the manual.

Mass of pFastBac TOPO = 10 ng

Length of pFastBac = 4824 bp

 Length of PCR insert = 1527 bp

 Length ratio of insert : vector should be close to their mass ratio

Length of PCR insert 1527 1 Mass ratio = = = Length of pFastBac TOPO vector 4824 3.16

 Moles of vector Mass of vector = ⁄Molecular weight of vector

10 푛𝑔 = ⁄3.16 × Molecular weight of PCR insert

10 푛𝑔  Mass of PCR insert at 1: 1 = Moles of vector × MW of insert = ⁄3.16 = 3.16 ng

20 푛𝑔  Mass of PCR insert at 2: 1 = 2 × Moles of vector × MW of insert = ⁄3.16 = 6.3 ng

G.4.2 Material

1.5 ng/μL PCR insert EcoRI-HF and XhoI restriction enzymes

10 ng/μL pFastBac/HBM-TOPO vector S.O.C. medium

Salt solution 10× CutSmart buffer

Mach1-T1 chemically competent E. Coli LB-agar plate with 100 μg/mL Ampicillin

227

LB medium with 100 μg/mL Ampicillin 15 mL Falcon tubes

1% agarose gel in 1×TBE Inoculation loop

Ethidium bromide Plasmid MiniPrep Kit

42 °C water bath UV gel scanner

G.4.3 Procedure

Day 1

 Set up TOPO cloning reaction with 1:1 insert: vector ratio. Leave reaction at room

temperature for 5 minutes, this is sufficient for insertion. Afterwards, place cloning

mixture on ice until transformation.

Table G.4 TOPO Ligation Protocol Reagent Volume (μL) Final 1.5 ng/μL PCR insert 2.10 3.15 ng 10 ng/μL pFastBac/HBM-TOPO vector 1.00 10 ng Salt solution 1.00 ----

ddH2O 1.90 ---- 6.00

 Transform TOPO cloning product into Mach1-T1 chemically competent E.coli cells

1. Thaw Mach1-T1 chemically competent cells on ice.

2. Add 2 μL of the TOPO cloning mixture into cell suspension, mix gently and

incubate on ice for 30 minutes.

3. Heat shock the cell-DNA mixture for 30 seconds in a 42 °C water bath

without shaking.

4. Quick move the tube onto ice bath, and chill for 2 minutes.

5. Add 250 μL S.O.C. medium to the cell-DNA mixture, shake the tube in the

bench-top shaker at 37 °C for 1 hour with orbital shaking rate at 225 rpm.

228

6. 20 minutes before seeding, pre-warm LB-agar plate containing ampicillin

in the 37 °C incubator.

7. Spread 25 μL of cell suspension onto the LB-agar plate, incubate upside-

down in the 37 °C incubator overnight.

Day 2

 Take out the LB-agar plate, there should be colonies on the plate. If not, try the

transformation again with different volume of TOPO reaction mixture; If re-

transformation still produce negative result, try the TOPO cloning reaction again

with 2:1 ratio of insert : vector.

 Pick 12 single colonies from the LB-agar plate, seed the cells into 12 individual

15ml Falcon tubes containing 5 mL LB medium with ampicillin. Shake overnight in

a 37 °C bench-top shaker at 225 rpm overnight.

Day 3

 Harvest the overnight cultures by centrifugation.

 Purify plasmid from each overnight culture using Plasmid MiniPrep kit from Qiagen.

 Measure the purified plasmid concentration in ng/μL via optical density at 260 nm.

 Dilute all the plasmid solutions to 100 ng/μL for the convenience of double

digestion.

 HER2 extracellular domain 1-3 has an intrinsic unique EcoRI cutting site,

meanwhile an XhoI site is also unique and positioned outside inserted fragment on

the vector backbone. Double digestion of purified plasmid from blunt-end TOPO

cloning with EcoRI and XhoI helps determine the orientation of the insertion, only

the correct orientation produces correct protein.

229

Table G.5. Double Digestion Recipe for Orientation Check

Reagent Volume (μL) Final 10× CutSmart buffer 5.00 1× 100 ng/μL purified pFastBac/HBM plasmid 10.00 1.0 μg 2 units/μL XhoI 1.00 2 units 2 units/μL EcoRI-HF 1.00 2 units

ddH2O 33.00 50.00

Incubate the reaction mixtures at 37 °C for 1 hour, then mix the reaction with 10

μL 6× gel loading buffer and load 5 μL mixture into each well on the agarose gel.

Electrophorese at 120 V for 1 hour, then visualized the image.

 Rational behind utilizing double digestion to determine inserted fragment

orientation is presented in the following Figure G.4 and a virtual digestion gel is

shown in Figure G.5.

230

EcoRI

EcoRI Correct Insert Reversed Insert XhoI XhoI

Correct orientation GAATTC CTTAAG EcoRI EcoRI Reversed GAATTC CTTAAG EcoRI Correct Insert Reversed Insert XhoI EcoRI XhoI Correct Insert XhoI Reversed Insert Correct orientation XhoI GAATTC CTTAAG --

Reversed GAATTC Restriction analysis of HER2 Reverse InsertionCorrect ori [Circular]entation CTTAAG Incubated with EcoRI + XhoI EcoRI GAATTC 2 fragments generated. CTTAAG

1: 5,060 bp - From XhoI[1791]EcoRI To EcoRI[500] Correct Insert 2: 1,291 bp - From EcoRI[500] To ReXhoI[1791]versed Insert XhoI Reversed XhoI GAATTC CTTAAG

Correct orientation Figure G.4. Double digestion for inserted fragment orientation determination. GAATopTTC: correct orientation, double digestion will generate a 338 bp short DNA fragment and a 6013 bp large DNA fragment. Bottom: reversed insertion, after doubel digestion, CTTAteoA G fragment are 1291 bp and 5060 bp. Significant length difference in digestion products plays the essential role in orientation determination.

Reversed GAATTC CTTAAG

231

Figure G.5. Gel image of double digested plasmid minipreps. Sample 1, 3, 5, 6, 9, 10, 11, and 12 showed digestion pattern of correct insertion orientation.

G.5 Protocol 4 - Generating Recombinant Bacmid via Tn7

Transposition.

G.5.1 Material pFastBac/HBM-HER2ECD(1-3) plasmid 10 mg/mL Gentamicin pFastBac Gus positive control plasmid 200 mg/mL IPTG pUC19 control plasmid 10 mg/mL Tetracycline in 100% ethanol

~20 LB-agar selection plates 20 mg/mL Bluo-Gal in DMSO

1 LB-agar plate with ampicillin 42 °C water bath

DH10Bac chemically competent cells 37 °C incubator and shaker

10 mg/mL Kanamycin Inoculation loop

PureLink™ Hipure Plasmid Midiprep UV spectrometer

G.5.1 Procedure

Day 1:

 Autoclave 600 mL LB-agar to prepare selection plate. Once the solution is cooled

down to room temperature, add the selection reagents following the recipe below.

232

Table G.6. Selection Agar Plate Recipe

Reagent Volume (mL) Final (μg/mL) 10 mg/mL Kanamycin 3.00 50.00 10 mg/mL Gentamicin 0.42 7.00 200 mg/mL IPTG 0.12 40.00 10 mg/mL Tetracycline in 100% ethanol 0.60 10.00 20 mg/mL Bluo-Gal in DMSO 3.00 100.00

Pour the agar plates, once the agar solidifies, invert the plate upside-down and

store them in 4 °C shielded from light exposure.

 Transform pFastBac/HBM-HER2ECD(1-3), pFastBac Gus positive control and

pUC19 control plasmid into DH10Bac chemically competent cells.

1. Thaw 3 tubes of DH10Bac chemically competent cells on ice.

2. Add DNA to DH10Bac cells, gently tap to mix, incubate on ice for 30

minutes.

pFastBac/HBM-HER2ECD(1-3): 1 ng

pFastBac Gus positive control: 1 ng

pUC19 control: 50 pg

3. Heat shock the cell-DNA mixture at 42 °C for 45 seconds without shaking.

4. Quickly place cells on ice to chill for 2 minutes.

5. Add 900 μL S.O.C. medium into each transformation tube and place tubes

in the 37 °C bench-top shaker at a rate of 225 rpm. pUC19 transformation

needs 1 hour recovery in the bench-top incubator, meanwhile all pFastBac

transformations require up to 4 hour incubation for efficient Tn7

transposition.

6. Seed pUC19 transformed cells onto ampicillin containing LB-agar plate

after 1 hour of shaking incubation.

233

Dilute the cells 10 folds, then spread 100 μL of diluted cell suspension onto

the plate, invert the plate and incubate overnight in a 37 °C incubator.

7. Make 10-fold, 100-fold and 1000-fold dilutions using S.O.C. medium of both

pFastBac/HBM-HER2ECD(1-3) transformed cells and pFastBac Gus

transformed cells after 4 hours incubation in the 37 °C bench-top shaker.

8. Spread 100 μL of each dilution as well as the original cell culture onto 4

selection LB-agar plates for each transformation. Invert the plate and

incubate for 48 hours at 37 °C.

Day 2

 Take out plate has pUC19 transformation control, the number of colonies grown

on the plate indicates the efficiency of transformations conducted on Day 1. If no

colonies on the plate, it is possible that selection plates will be negative, then it is

necessary to repeat the transformation. Calculate the transformation efficiency (TE)

using the equations below:

Equation G.2

Total plasmid mass 1 Seeding mass (μg) = × × Seeding volume Total culture volume Dilution factor

50 pg 1 = × × 100 μL = 0.5 pg = 0.5 × 10−6 μg 1000 μL 10

Equation G.3

Colony number 1 Seeding volume TE = × × μg of plasmid Dilution factor Diluted volume

96 colonies 1 100 μL = × × = 2 × 108 cfu/μg 0.5 × 10−6 μg 10 1000 μL

Day 3

 After 48-hour incubation, take out the selection plate. Colonies with dark blue and

white color should be distinguishable.

234

 Pick 10 white colonies from pFastBac/HBM-HER2ECD(1-3) transformation, and 3

white colonies from pFastBac Gus transformation, re-streak them onto new

selection plate, incubate again in 37 °C overnight. Colonies remain white color

after re-plating are true positive colonies, which contain desired transposed

Bacmid.

Blue-White Screen

Blue-white screen is a powerful molecular cloning method that allows identification of

insert without plasmid purification and sequencing.

The assay originates from the observation of β-galacosidase function restoration via

α-complementation. Gene encoding α-peptide, which contains first 59 amino acids of

β-galacosidase, was transformed into a strain expessing mutant β-galacosidase with

N-terminus deletion,26-28 function of β-galacosidase is recovered and digestion of Bluo-

Gal will generate blue colored colonies. However, with multiple cloning sites embedded within lacZ α-peptide sequences, insertion of the gene will interrupt the expression of functional α-peptide, thus transformation of the recombined plasmid will not restore β-galacosidase activity, and positive colonies will be white.

Since blue-white screen correlates genotype with phenotype, evaluation of ligation or transposition results can be accomplished within 2 days. However, since blue-white screen depends on the color development inside E.coli cells, one should provide sufficient time to ensure color to be distinguishable; further, occasionally, white colonies turn out to be negative of insertion due to mutations or other cellular repair mechanisms when linear plasmid is transformed. Therefore, always re-plate multiple white colonies to select for the correct genotype.

235

lacI Promoter Operator lacZ

Repressor OH OH OH OH OH O O HO S HO O O OH OH OH OH IPTG Lactose

lacI Promoter lacZ

b-Galactosidase

H H H OH OH NH N N N O HO O OH Br Br Br HO O O OH OH Br O Bluo-Gal HO OH Blue color OH

Figure G.6. lac operon tightly regulates the expression of β-galacosidase. Once induced, by lactose or lactose analog IPTG, repressor release the transcription site of β-galacosidase; production of β-galacosidase will initiate the hydrolysis of Bluo-Gal, whose product will dimerized via oxidation and shows blue color.

236

lacZaL Insert lacZaR lacZDM15

Recombinant

DH10Bac Genome

lacZa

Active

Inactive

Bacmid

Figure G.7. Blue-white screen basics. Positive recombination interrupts lacZα sequences, expressed lacZα fragments fail to fold into active conformation, thus α-complementation is impeded. No active β-galacosidase exists intracullularly, hence recombinant colonies appears to be white.

Day 4:

 Pick 3 true white colonies from from each transformation, seed each colony into 5

mL LB supplemented with 50 μg/mL kanamycin, 10 μg/mL tetracycline and 7

μg/mL of gentamicin.

Table G.7. DH10Bac growing medium For 50 mL of LB 10 mg/mL kanamycin 250 μL 50 μg/mL 10 mg/mL tetracycline in 100% ethanol 50 μL 10 μg/mL 10 mg/mL gentamicin 35 μL 7 μg/mL

 Incubate the liquid cell culture at 37 °C at the rate of 225 rpm overnight.

237

Day 5

 Purify recombinant bacmid from overnight cell culture, using “PureLink™ Hipure

Plasmid DNA Midiprep Kit”.

 Follow kit instruction, pay attention DO NOT pipette to dissolve bacmid after

precipitation, shear force will cause strand breaks on bacmid due to its large size.

 Measure bacmid midiprep concentration with UV spectrometer.

G.6 Protocol 5: Analyzing Recombinant Bacmid DNA by PCR

Tn7 transposition of HER2ECD(1-3), as mentioned above, is proceeded via recognition of attTn7 on DNA sequences. In DH10Bac, the intrinsic basmid has attTn7 located in between pUC/M13 forward primer and pUC/M13 reverse primer, this design enables the final analysis of bacmid recombination though PCR amplification of transposed region.

Tn7R Insertion Tn7L

pUC/M13 Forward attTn7 pUC/M13 Reverse

139 bp 157 bp

pUC/M13 Forward Tn7R Insertion Tn7L pUC/M13 Reverse

139 bp 157 bp

Figure. G.8. Transposed area is primed under pUC/M13 forward primer and reverse primers. PCR amplification using this primer set can further identify the construct of bacmid, since insertion will generate a longer piece PCR product.

238

The estimated PCR product length can be calculated as follows:

Table G.8. Estimation on Product Length from Bacmid PCR Bacmid transposed with Size of PCR product Actually size Bacmid alone ~ 300 bp --- pFastBac/HBM-HER2ECD(1-3) ~ 2,500 bp + insertion ~ 2,500 + 1,527 = 4.027 bp pFastBac Gus ~ 4,200 bp ----

G.6.1 Material

Purified Bacmid DNA pUC/M13 primer set

5× Q5 reaction buffer

5× Q5 GC enhancer

Q5 high fidelity DNA polymerase

10 mM dNTPs

1% agarose gel in 1× TBE

Ethidium bromide

Thermocycler

UV gel scanner

239

G.6.2 Procedure

 Set up PCR reaction for each purified bacmid:

Table G.9. Bacmid PCR Protocol Reagent Volume (μL) Final 5× Q5 reaction buffer 10.00 1× 5× Q5 GC enhancer 10.00 1× 10 mM dNTPs 1.00 0.2 μM 10 μM pUC/M13 forward primer 1.25 0.25 μM 10 μM pUC/M13 reverse primer 1.25 0.25 μM 100 ng/μL bacmid 1.00 100 ng 2 units/μL Q5 high fidelity DNA polymerase 0.50 1 unit

ddH2O Up to 50.00

 Amplify using the following thermocycling parameters:

Table G.10. Bacmid PCR Thermocycle Step Temperature (°C) Time (second) Initial denaturation 98.0 180 Denaturation 98.0 45 35 cycles Annealing 55.0 45 Elongation 72.0 300 Polish 72.0 420 Hold 4.0 ----

 Aliquot 5-10 μL from the reaction, mix with gel loading solutions and analyze on

agarose gel contain ethidium bromide. Apply constant voltage at 120 V for 1 hour,

and scan gel with UV gel scanner.

 Store bacmid in 4 °C until sf9 cells are ready to be transfected, freeze-thaw cycle

may shear bacmid and lower the transfection efficiency. One can also dilute and

aliquot bacmid into ready-to-use format, and freeze the bacmid in -20 °C.

240

 Once the construct of recombinant bacmid has been determined, pick another true

white colony from the re-plate, and grow it overnight into a 5 mL liquid culture.

Glycerol stock can be prepared the second day, and can be used for bacmid

preparation once the current batch is depleted.

Figure G.9. PCR analysis of recombinant bacmid. Top: PCR product of Bac-to-Bac HER2ECD(1-3) is around 4027 bp for all 3 bacmid preparations, construct is confirmed to be correct. Bottom right: PCR product of Bac-to-Bac Gus is above 4000 bp marker and can be confirmed as correct construct. Bottom left: DNA ladder information provided by the vendor (Gold Biotechnology).

241

G.7 References for Appendix G

1. David R. O'Reilly, L. K. M., Verne A. Luckow, Baculovirus expression vectors: a

laboratory manual. W.H. Freeman and Company: New York, 1992.

2. Ayres, M. D.; Howard, S. C.; Kuzio, J.; Lopez-Ferber, M.; Possee, R. D., The

complete DNA sequence of Autographa californica nuclear polyhedrosis virus.

Virology 1994, 202, 586-605.

3. Harrap, K. A., The structure of nuclear polyhedrosis viruses. 3. Virus assembly.

Virology 1972, 50, 133-9.

4. Kool, M.; Vlak, J. M., The structural and functional organization of the Autographa

californica nuclear polyhedrosis virus genome. Archives of virology 1993, 130, 1-

16.

5. Drugmand, J. C.; Schneider, Y. J.; Agathos, S. N., Insect cells as factories for

biomanufacturing. Biotechnology advances 2012, 30, 1140-57.

6. Kool, M.; Ahrens, C. H.; Vlak, J. M.; Rohrmann, G. F., Replication of baculovirus

DNA. The Journal of general virology 1995, 76 ( Pt 9), 2103-18.

7. Berger, I.; Fitzgerald, D. J.; Richmond, T. J., Baculovirus expression system for

heterologous multiprotein complexes. Nature biotechnology 2004, 22, 1583-7.

8. Hoss, A.; Moarefi, I.; Scheidtmann, K. H.; Cisek, L. J.; Corden, J. L.; Dornreiter, I.;

Arthur, A. K.; Fanning, E., Altered phosphorylation pattern of simian virus 40 T

antigen expressed in insect cells by using a baculovirus vector. Journal of virology

1990, 64, 4799-807.

9. Kloc, M.; Reddy, B.; Crawford, S.; Etkin, L. D., A novel 110-kDa maternal CAAX

box-containing protein from Xenopus is palmitoylated and isoprenylated when

expressed in baculovirus. The Journal of biological chemistry 1991, 266, 8206-12.

242

10. Kuroda, K.; Veit, M.; Klenk, H. D., Retarded processing of influenza virus

hemagglutinin in insect cells. Virology 1991, 180, 159-65.

11. Baixeras, E.; Roman-Roman, S.; Jitsukawa, S.; Genevee, C.; Mechiche, S.;

Viegas-Pequignot, E.; Hercend, T.; Triebel, F., Cloning and expression of a

lymphocyte activation gene (LAG-1). Molecular immunology 1990, 27, 1091-102.

12. Brandt-Carlson, C.; Butel, J. S., Detection and characterization of a glycoprotein

encoded by the mouse mammary tumor virus long terminal repeat gene. Journal

of virology 1991, 65, 6051-60.

13. Caroni, P.; Rothenfluh, A.; McGlynn, E.; Schneider, C., S-cyclophilin. New member

of the cyclophilin family associated with the secretory pathway. The Journal of

biological chemistry 1991, 266, 10739-42.

14. Schaefer, G.; Haber, L.; Crocker, L. M.; Shia, S.; Shao, L.; Dowbenko, D.; Totpal,

K.; Wong, A.; Lee, C. V.; Stawicki, S.; Clark, R.; Fields, C.; Lewis Phillips, G. D.;

Prell, R. A.; Danilenko, D. M.; Franke, Y.; Stephan, J. P.; Hwang, J.; Wu, Y.;

Bostrom, J.; Sliwkowski, M. X.; Fuh, G.; Eigenbrot, C., A two-in-one antibody

against HER3 and EGFR has superior inhibitory activity compared with

monospecific antibodies. Cancer cell 2011, 20, 472-86.

15. Vuori, K.; Pihlajaniemi, T.; Marttila, M.; Kivirikko, K. I., Characterization of the

human prolyl 4-hydroxylase tetramer and its multifunctional protein disulfide-

isomerase subunit synthesized in a baculovirus expression system. Proceedings

of the National Academy of Sciences of the United States of America 1992, 89,

7467-70.

16. Hasemann, C. A.; Capra, J. D., High-level production of a functional

immunoglobulin heterodimer in a baculovirus expression system. Proceedings of

243

the National Academy of Sciences of the United States of America 1990, 87, 3942-

17. Luckow, V. A.; Lee, S. C.; Barry, G. F.; Olins, P. O., Efficient generation of

infectious recombinant baculoviruses by site-specific transposon-mediated

insertion of foreign genes into a baculovirus genome propagated in Escherichia

coli. Journal of virology 1993, 67, 4566-79.

18. Wang, J. C., Cellular roles of DNA topoisomerases: a molecular perspective.

Nature reviews. Molecular cell biology 2002, 3, 430-40.

19. Champoux, J. J., DNA topoisomerases: structure, function, and mechanism.

Annual review of biochemistry 2001, 70, 369-413.

20. Wang, J. C., DNA topoisomerases. Annual review of biochemistry 1996, 65, 635-

92.

21. Kuduvalli, P. N.; Mitra, R.; Craig, N. L., Site-specific Tn7 transposition into the

human genome. Nucleic acids research 2005, 33, 857-63.

22. Ailor, E.; Betenbaugh, M. J., Modifying secretion and post-translational processing

in insect cells. Current opinion in biotechnology 1999, 10, 142-5.

23. Tessier, D. C.; Thomas, D. Y.; Khouri, H. E.; Laliberte, F.; Vernet, T., Enhanced

secretion from insect cells of a foreign protein fused to the honeybee melittin signal

peptide. Gene 1991, 98, 177-83.

24. Garrett, T. P.; McKern, N. M.; Lou, M.; Elleman, T. C.; Adams, T. E.; Lovrecz, G.

O.; Kofler, M.; Jorissen, R. N.; Nice, E. C.; Burgess, A. W.; Ward, C. W., The crystal

structure of a truncated ErbB2 ectodomain reveals an active conformation, poised

to interact with other ErbB receptors. Molecular cell 2003, 11, 495-505.

25. Tindall, K. R.; Kunkel, T. A., Fidelity of DNA synthesis by the Thermus aquaticus

DNA polymerase. Biochemistry 1988, 27, 6008-13.

244

26. Langley, K. E.; Villarejo, M. R.; Fowler, A. V.; Zamenhof, P. J.; Zabin, I., Molecular

basis of beta-galactosidase alpha-complementation. Proceedings of the National

Academy of Sciences of the United States of America 1975, 72, 1254-7.

27. Messing, J.; Gronenborn, B.; Muller-Hill, B.; Hans Hopschneider, P., Filamentous

coliphage M13 as a cloning vehicle: insertion of a HindII fragment of the lac

regulatory region in M13 replicative form in vitro. Proceedings of the National

Academy of Sciences of the United States of America 1977, 74, 3642-6.

28. Ullmann, A.; Jacob, F.; Monod, J., Characterization by in vitro complementation of

a peptide corresponding to an operator-proximal segment of the beta-

galactosidase structural gene of Escherichia coli. Journal of molecular biology

1967, 24, 339-43.

245

References

Chapter 1

1. Arnott, S., Historical article: DNA polymorphism and the early history of the double

helix. Trends Biochem. Sci. 2006, 31, 349-354.

2. Bochman, M. L.; Paeschke, K.; Zakian, V. A., DNA secondary structures: stability

and function of G-quadruplex structures. Nature Reviews Genetics 2012.

3. Tsai, Y. C.; Qi, H.; Liu, L. F., Protection of DNA ends by telomeric 3′ G-tail

sequences. J. Biol. Chem. 2007.

4. Lingner, J.; Cooper, J. P.; Cech, T. R., Telomerase and DNA end replication: no

longer a lagging strand problem? Science 1995, 269, 1533-4.

5. Paeschke, K.; Simonsson, T.; Postberg, J.; Rhodes, D.; Lipps, H. J., Telomere

end-binding proteins control the formation of G-quadruplex DNA structures in vivo.

Nat. Struct. Mol. Biol. 2005, 12, 847-54.

6. Gilson, E.; Géli, V., How telomeres are replicated. Nature Reviews Molecular Cell

Biology 2007.

7. Neidle, S.; Parkinson, G., Telomere maintenance as a target for anticancer drug

discovery. Nature Reviews Drug Discovery 2002.

8. Huppert, J. L.; Balasubramanian, S., G-quadruplexes in promoters throughout the

human genome. Nucleic Acids Res. 2007.

246

9. Burge, S.; Parkinson, G. N.; Hazel, P.; Todd, A. K.; Neidle, S., Quadruplex DNA:

sequence, topology and structure. Nucleic Acids Res. 2006, 34, 5402-15.

10. Agris, P. F.; Vendeix, F. A. P.; Graham, W. D., tRNA's wobble decoding of the

genome: 40 years of modification. J. Mol. Biol. 2007.

11. Varani, G.; McClain, W. H., The G· U wobble base pair. EMBO reports 2000.

12. Nissen, P.; Ippolito, J. A.; Ban, N.; Moore, P. B.; Steitz, T. A., RNA tertiary

interactions in the large ribosomal subunit: the A-minor motif. Proc. Natl. Acad. Sci.

U. S. A. 2001, 98, 4899-903.

13. Theimer, C. A.; Blois, C. A.; Feigon, J., Structure of the human telomerase RNA

pseudoknot reveals conserved tertiary interactions essential for function. Mol. Cell

2005, 17, 671-82.

14. Fedor, M. J.; Williamson, J. R., The catalytic diversity of RNAs. Nat. Rev. Mol. Cell

Biol. 2005, 6, 399-412.

15. Smith, A. M.; Fuchs, R. T.; Grundy, F. J.; Henkin, T. M., Riboswitch RNAs:

regulation of gene expression by direct monitoring of a physiological signal. RNA

Biol. 2010, 7, 104-10.

16. Stern, S.; Powers, T.; Changchien, L. M.; Noller, H. F., RNA-protein interactions in

30S ribosomal subunits: folding and function of 16S rRNA. Science 1989, 244,

783-90.

17. Adams, P. L.; Stahley, M. R.; Gill, M. L.; Kosek, A. B., Crystal structure of a group

I intron splicing intermediate. RNA 2004.

18. Sripathi, K. N.; Banas, P.; Reblova, K.; Sponer, J.; Otyepka, M.; Walter, N. G.,

Wobble pairs of the HDV ribozyme play specific roles in stabilization of active site

dynamics. Phys. Chem. Chem. Phys. 2015, 17, 5887-900.

247

19. Felsenfeld, G.; Davies, D. R.; Rich, A., Formation of a Three-Stranded

Polynucleotide Molecule. J. Am. Chem. Soc. 1957, 79, 2023-2024.

20. Morgan, A. R.; Wells, R. D., Specificity of the three-stranded complex formation

between double-stranded DNA and single-stranded RNA containing repeating

nucleotide sequences. J. Mol. Biol. 1968.

21. Beal, P. A.; Dervan, P. B., Second structural motif for recognition of DNA by

oligonucleotide-directed triple-helix formation. Science 1991, 251, 1360-3.

22. Radhakrishnan, I.; Patel, D. J., Solution structure of a purine· purine· pyrimidine

DNA triplex containing G· GC and T· AT triples. Structure 1993.

23. Ghosh, I.; Stains, C. I.; Ooi, A. T.; Segal, D. J., Direct detection of double-stranded

DNA: Molecular methods and applications for DNA diagnostics. Mol Biosyst 2006,

2, 551-60.

24. Oussedik, K.; Halby, L.; Arimondo, P. B., The triple helix: 50 years later, the

outcome. Nucleic acids … 2008.

25. Plum, G. E.; Park, Y. W.; Singleton, S. F.; Dervan, P. B.; Breslauer, K. J.,

Thermodynamic characterization of the stability and the melting behavior of a DNA

triplex: a spectroscopic and calorimetric study. Proc. Natl. Acad. Sci. U. S. A. 1990,

87, 9436-40.

26. Lyamichev, V. I.; Mirkin, S. M.; Frank-Kamenetskii, M. D.; Cantor, C. R., A stable

complex between homopyrimidine oligomers and the homologous regions of

duplex DNAs. Nucleic Acids Res. 1988, 16, 2165-78.

27. Lyamichev, V. I.; Voloshin, O. N.; Frank-Kamenetskii, M. D.; Soyfer, V. N.,

Photofootprinting of DNA triplexes. Nucleic Acids Res. 1991, 19, 1633-8.

28. Thomas, T.; Thomas, T. J., Selectivity of polyamines in triplex DNA stabilization.

Biochemistry 1993, 32, 14068-74.

248

29. Thomas, T. J.; Kulkarni, G. D.; Greenfield, N. J.; Shirahata, A.; Thomas, T.,

Structural specificity effects of trivalent polyamine analogues on the stabilization

and conformational plasticity of triplex DNA. Biochem. J. 1996, 319 ( Pt 2), 591-9.

30. Arimondo, P. B.; Barcelo, F.; Sun, J. S.; Maurizot, J. C.; Garestier, T.; Helene, C.,

Triple helix formation by (G,A)-containing oligonucleotides: asymmetric sequence

effect. Biochemistry 1998, 37, 16627-35.

31. Blume, S. W.; Lebowitz, J.; Zacharias, W.; Guarcello, V.; Mayfield, C. A.;

Ebbinghaus, S. W.; Bates, P.; Jones, D. E., Jr.; Trent, J.; Vigneswaran, N.; Miller,

D. M., The integral divalent cation within the intermolecular purine*purine.

pyrimidine structure: a variable determinant of the potential for and characteristics

of the triple helical association. Nucleic Acids Res. 1999, 27, 695-702.

32. Floris, R.; Scaggiante, B.; Manzini, G.; Quadrifoglio, F.; Xodo, L. E., Effect of

cations on purine.purine.pyrimidine triple helix formation in mixed-valence salt

solutions. Eur. J. Biochem. 1999, 260, 801-9.

33. Goni, J. R.; de la Cruz, X.; Orozco, M., Triplex-forming oligonucleotide target

sequences in the human genome. Nucleic Acids Res. 2004, 32, 354-60.

34. Goni, J. R.; Vaquerizas, J. M.; Dopazo, J.; Orozco, M., Exploring the reasons for

the large density of triplex-forming oligonucleotide target sequences in the human

regulatory regions. BMC Genomics 2006, 7, 63.

35. Durland, R. H.; Kessler, D. J.; Gunnell, S.; Duvic, M.; Pettitt, B. M.; Hogan, M. E.,

Binding of triple helix forming oligonucleotides to sites in gene promoters.

Biochemistry 1991, 30, 9246-55.

36. Duval-Valentin, G.; Thuong, N. T.; Helene, C., Specific inhibition of transcription

by triple helix-forming oligonucleotides. Proc. Natl. Acad. Sci. U. S. A. 1992, 89,

504-8.

249