Structure Activity Relationships of Novel Merocyanine Dye Derivatives

Structure Activity Relationships of Novel Merocyanine Dye Derivatives: Effects on Acid-Base Chemistry and Fluorescence Energy Transfer Capabilities with G-Quadruplex Forming DNA with an Application for Detection

Delaney E. Armstrong-Price

A Thesis

Presented to

The University of Guelph

In partial fulfilment of requirements

For the degree of

Master of Science

Chemistry and Toxicology

Guelph, ON, Canada

©Delaney Armstrong-Price, December 2017

ABSTRACT

Structure Activity Relationships of Novel Merocyanine Dye Derivatives: Effects on Acid Base Chemistry and Fluorescence Energy Transfer Capabilities with G-Quadruplex Forming DNA with an Application for Detection

Delaney E. Armstrong-Price Advisor: University of Guelph, 2017 Professor R. Manderville

The merocyanine family of dyes was first introduced in the 1950s as photosensitizing agents for the development of colour photographs. Since this time, this family of chromophores has gained much more attention in the realms of materials science, dye chemistry as well as in biotechnology. Demonstrating impressive optical changes as a function of pH, solvent polarity, and solvent viscosity, these dyes present as prime candidates for use in a wide variety of detection platforms. Herein, novel phenolic merocyanine dye derivatives were synthesized and characterized based on changes observed in the dyes’ absorbance and fluorescence capabilities as a function of acid-base exchange in aqueous environments and their ability to undergo tautomeric exchange processes in a number of polar protic solvents.

Additionally, molecular rotor examples of these dyes were synthesized and assessed for their ability to undergo fluorescence emission enhancement in high viscosity solvents as well as in the presence of guanine-quadruplex (GQ) DNA. It was observed that these dye derivatives were capable of undergoing an energy transfer process in the presence of a DNA GQ, providing a means to determine the binding affinity of the dyes with various GQ topologies. This was achieved through the use the human telomeric repeat sequence, a sequence demonstrating high polymorphism in its ability adopt a variety of GQ topologies.

Finally, this energy transfer phenomenon was applied to the detection of Ochratoxin A

(OTA) with a GQ forming 36-mer DNA aptamer. In this label free assay, energy transfer from

the aptamer to the toxin allowed for a 3-fold enhancement in the visible fluorescence of OTA, providing a sensitive, simple and elegant strategy for the detection of this small molecule target.

ACKNOWLEDGEMENTS

I have to thank my family first and foremost. Without their constant love and support, I would have never even tried to pursue graduate studies. Because they knew I could do it, I mustered up the courage to try. To my mother, Sharon, thank you for your unwavering support and love through every single level of my post secondary education. To my Aunt Dawn and cousin Cari, second Mom and the closest thing to a sister I was the luckiest girl to have, again, your support, love, and care packages fuelled the completion of this document. You can keep the care packages coming if you’d like. To the Aunts I met just this year (a long story), Karen and Ann, it was meeting you where I finally got to see the whole picture of where I come from. Thank you for reaching out to me and I hope the end of this chapter in my life opens a new one where I get to know both of you a whole lot better. On the thread of family, Cayla, Sabine, and Jess, the best friends I have know idea how I got so blessed to have, thank you for your patience when I ranted, your support through and through and for providing me with an outlet when the going got tough. I love each and everyone of you. Thank you, thank you, thank you!

I would like to thank my advisor Dr. Richard Manderville. Thank you for taking me on as a Master’s student in your lab. I have learned so much and I can’t thank you enough for giving me the opportunity to continue my studies at the graduate level. There were many peaks and many valleys along the way when trying to piece together this story, but we made it! Thank you for all of your support over these three years I have spent in your lab. This thesis would not have been possible without your guidance. Thank you. I would like to thank my advisory committee, Dr. Adrian Schwan and Dr. William Tam. Throughout undergrad and now to the end of my graduate degree, I cannot thank you enough for imparting your wisdom and for all of your guidance. Thank you.

I would like to thank the Grain Farmers of Ontario for providing their financial support.

I would also like to thank the members of the Manderville research group. Everyone whom I have crossed paths with in my time with the group has helped to shape the person that I am today. I have learned many valuable skills and life lessons from some and I have grown not only as a scientist and as a student but as a person overall. I owe so much of that to my lab mates. Truly, from the bottom of my heart, thank you. To those who I consider friends, I cannot thank you enough for all of your help, support, and for your unique understanding of this crazy life we call grad school. Special mention goes out to Dr. Kaila Fadock and Thomas Cservenyi. Kaila, without your guidance and wisdom, this thesis would not exist. It was a privilege to be mentored by you and I am honoured and so glad to now call you a best friend. Thomas, I feel as though we had similar journeys traversing these ropes. You understood best how the wins and loses could really make or break your motivation and spirit. Thank you for listening and for your understanding. Thank you both so much.

I have to thank Dr. Kate Stuttaford and Dr. Rob Reed for being the biggest, baddest (by “baddest” I mean best!) bosses on the block. Kate, thank you for life chats and good times. Thank you for always being there and lending an ear when all I needed was for someone to listen. I also cannot thank you enough for giving me the opportunity to teach in your labs. I am a better analytical chemist for it. I come away from this with not only a degree but also a life long colleague

iv and friend. Thank you. Rob, thank you for letting me run with the wind in the Organic III labs. I have to say, teaching this lab was one of the brightest highlights I experienced in my time as a graduate student in Guelph. Thank you for believing in me to do it justice after Kaila set the bar so high and thank you for your unfailing guidance with regards to teaching and even in my own academic endeavours. It is thanks to you both that I was able to teach the best way I know how: with a little tough love, a little seriousness and maybe even a little silliness. I learned from the best in this regard.

A few others that I have to thank: Dr. David Josephy for the use of his UV-Vis spec, Dr. A. Rod Merrill for the use of his CD spec, Dr. France-Isabelle Auzanneau for her support and entrusting me with so many of the “hats” I was able to wear throughout my time as a grad student, to Dr. Chris Whitfield for giving me my first shot at research as an undergraduate research assistant, Dr. Iain Mainprize of the Whitfield research group for showing me initially, way back in 2013, that I just might have a knack for this research thing, Dr. Paula Russell for life chats, support, and making me the writer that I am today, Debbie and Linda for fulfilling my caffeine and sass requirements each day, to the D-Tox crew through undergrad and even now, to The Brass Taps for always having a pint ready and waiting after work on a tough day, to all of the members of the GSA, past and present that I had the privilege to work with. My graduate experience was enriched beyond mention thanks to my involvement with this organization. Special mention here goes out to Jane Ferguson. I am so glad to have gained a great friend in my journeys with the GSA. It was an honour and a total blast to represent the chemistry department with you. To everyone who contributed in some small way to this journey and who is not mentioned above, THANK YOU!!

TABLE OF CONTENTS

Chapter 1: Introduction ...... 1 1.1 Deoxyribonucleic Acid ...... 2 1.1.1 Components of DNA ...... 2 1.2 The structure of DNA ...... 3 1.2.1 The DNA Double Helix ...... 3 1.2.2. Hoogsteen Base Pairs ...... 5 1.2.3. The Guanine Quadruplex ...... 6 1.3. Analysis of DNA Structure ...... 8 1.3.1. An Introduction to the Methods ...... 8 1.3.2. Analysis of DNA using UV-Vis Spectroscopy ...... 9 1.3.4. Circular Dichroism Spectroscopy ...... 10 1.4 An Introduction to Biosensors ...... 12 1.5. Aptamers ...... 13 1.5.1. Aptamers: A Definition...... 13 1.5.2. SELEX ...... 14 1.5.3. GQ Forming Aptamers ...... 15 1.6. Fluorescent Probes for Aptamer Based Detection ...... 16 1.6.1. An Introduction to Fluorescence ...... 16 1.6.2. Internal Base Modification ...... 17 1.6.2.1. An Example of a Fluorescently Modified Internal Base ...... 18 1.6.3. Molecular Rotor Probes ...... 19 1.6.4. Cyanine Dyes: Examples of Molecular Rotor Probes ...... 21 1.6.5. pH Sensing Probes ...... 22 1.6.6. Excited State Proton Transfer Fluorescence ...... 24 References ...... 26 Chapter 2: Synthesis and Characterization of Novel Phenolic Merocyanine Dye Derivatives ... 29 2.1 Introduction ...... 30 2.1.1. Brooker’s Merocyanine ...... 30 2.1.2. Introduction to the Hammett Equation ...... 31

2.1.3. Effects of Keto-Enol Tautomerism on Solvatochromic and Fluorosolvatochromic Properties of Hydroxy Merocyanine Dyes ...... 32 2.2. Materials and Methods ...... 34 2.2.1. General Synthesis of Merocyanine Dyes ...... 34 2.2.1.1. General Procedure - Synthesis of Activated Methylene Compounds ...... 34 2.2.1.2. Synthesis of Activated Compounds ...... 35 2.2.1.3. General Procedure - Synthesis of Merocyanine Dyes ...... 36 2.2.1.4. Synthesis of Merocyanine Dyes...... 37 2.2.2. Determination of Photophysical Properties ...... 39 2.2.3. pKa Determination ...... 41 2.3. Results and Discussion ...... 41 2.3.1. Solvatochromic Properties of the Merocyanine Derivatives ...... 41 2.3.2. Fluorosolvatochromic Properties of the Merocyanine Dyes ...... 49 2.3.3. Absorbance Characteristics as a function of pH ...... 52

2.3.4. pKa Determination ...... 55 2.3.5. Hammett Analysis ...... 57

2.3.6. Insight into Hammett Analysis and pKa Results: What Does it All Mean? ...... 62 2.3.7. Fluorescent Characteristics as a function of pH ...... 68 2.3.7.1. Excited State Electron Transfer of the Merocyanine Dyes ...... 71 2.3.7.2. Explaining Fluorescence Enhancement of PhOH Btz and PhOH Ind in Basic Conditions ...... 71 2.4. Conclusions ...... 73 References ...... 76 Chapter 3: Probing Structure Activity Relationships Contributing to GQ Binding Ability of Novel Merocyanine Dyes: Energy Transfer Fluorescence and CD Spectroscopy Studies ...... 79 3.1. Introduction ...... 80 3.1.1. Human Telomeric Repeat Sequence ...... 80 3.1.2. Thrombin Binding Aptamer ...... 82 3.1.3. GQ Binding Fluorophores...... 83 3.1.4. Energy Transfer: From DNA into Fluorophores ...... 85 3.2. Materials and Methods ...... 86 3.2.1. General Synthesis of the Merocyanine Dyes ...... 86

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3.2.2. Preparation of Stock Solutions and UV-Vis and Fluorescence Experiments ...... 87 3.2.3. Glycerol Studies ...... 87 3.2.4. TBA:C2 Fluorescence Titrations ...... 88 3.2.5. CD Studies ...... 88 3.2.6. UV-Vis and Fluorescence Titrations of the Merocyanine Dyes with Htelo ...... 88

3.2.7. Determination of the Binding Affinity of the Merocyanine Dyes for Htelo DNA – Kd Determination ...... 89 3.3. Results and Discussion ...... 89 3.3.1. Characterization of the Rotor Merocyanine Dyes ...... 89 3.3.2. Duplex to Quadruplex Exchange: The Thrombin Example ...... 92 3.3.3. CD Studies ...... 94 3.3.4. UV-Vis and Fluorescence Titrations of Merocyanine Dyes with Htelo ...... 107 3.3.5.1.1. Making Sense of the Differences in Optical Output from the Merocyanine Dyes as They Relate to Varied Conditions...... 122 3.3.5.2. ET Emission with GQ Binding ...... 127 3.3.6. Binding Affinity of Merocyanine Dyes with Htelo ...... 130 3.4. Conclusions ...... 137 References ...... 139 Chapter 4: Developing an Aptamer Based Detection Platform for a Small Molecule Target: An Energy Transfer Approach ...... 142 4.1. Introduction ...... 143 4.1.1. Ochratoxin A ...... 143 4.1.2. Current OTA Detection Methods...... 143 4.1.3. Aptamer Based Platforms for OTA Detection ...... 144 4.1.4. Inspiration for the Current Study: A FRET Based Label Free Assay for OTA Detection ...... 145 4.2. Materials and Methods ...... 147 4.2.1. Preparation of OTA Binding Buffer...... 148 4.2.2. Fluorescence Titration of OTA with OTAA36...... 148

4.2.3. Kd Determination...... 148 4.2.4. Circular Dichroism Studies...... 148 4.2.5. Calibration Curves...... 149

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4.2.6. Determination of LOD and LOQ...... 150 4.2.7. Control Fluorescence Titrations...... 150 4.3. Results and Discussion ...... 150 4.3.1. Fluorescence Titration of OTA with OTAA36 ...... 150 4.3.2. Dissociation Constant Determination and Circular Dichroism Binding Studies ...... 153 4.3.3. Limit of Detection and Limit of Quantitation ...... 155 4.3.4. Selectivity and Specificity of the Assay ...... 158 4.4. Conclusion ...... 160 References ...... 162 Appendix A: General Methods ...... 165 Appendix B: 1H and 13C NMR Characterization of Synthesized Products ...... 169 Appendix C: Additional Data ...... 181

LIST OF TABLES

Table 2.1: Summary of solvatochromic data for the novel merocyanine dye derivatives...... 45 Table 2.2: Summary and Comparison of keto and enol fluorescent properties ...... 52 Table 2.3: A summary of the absorbance characteristics of the phenolic merocyanine dyes...... 55

Table 2.4: Calculated Phenolic pKas for the merocyanine derivatives ...... 57 Table 2.5: Summary of the calculated Hammett substituents constants for the activated nitrogen heteroaryl substituents...... 59 Table 2.6: Comparison of pKa and σ- values of the merocyanine dyes to the reference compounds p-nitrophenol, p-cynophenol and p-propenyl phenol...... 60 Table 2.7 HOMA values for literature example compounds pertinent to the current study...... 63 Table 2.8: NICS(1) Values for a Number of Reference Compounds ...... 67 Table 2.9: Summary of the fluorescent data obserevd for the merocyanine dyes as a function of pH...... 70 Table 3.1: Summary of Molecular Rotor Characterization Data ...... 91 Table 3.2: Comparing excitation maximum wavelengths for the free merocyanine to merocyanine dyes bound to Htelo DNA...... 107 Table 3.3: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer without the presence of a metal cation ...... 111 Table 3.4: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer with 50 mM NaCl...... 116 Table 3.5: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer with 50 mM KCl...... 121 Table 3.6: Summary of Stoke’s Shift data for all of the merocyanine dyes under all experimental conditions employed: 50 mM Tris, 50 mM Tris with 50 mM NaCl or 50 mM Tris with 50 mM KCl ...... 124

Table 3.7: Summary of Kd values determined for all of the merocyanine dyes in 50 Mm Tris in the presence of: no metal cation, 50 mM Na+, or 50 mM K+ ...... 131 Table 4.1: Comparison of the fluorescence data obtained for free OTA and the OTA:OTAA36 ...... 158

LIST OF FIGURES

Figure 1.1: The structure of a single nucleotide subunit of DNA which also demonstrates the difference in structure between a nucleotide, nucleoside and nitrogenous base. The numbering scheme for the 2’deoxyribose sugar as well as the N-glycosidic bond (χ) are shown...... 2 Figure 1.2: An oligonucleotide demonstrating the phosphodiester backbone of DNA where linkages occur at the 3’hydroxyl of one nucleotide and the 5’hydroxyl of an adjacent one. In addition, all purine and pyrimidine nitrogenous bases are shown...... 3 Figure 1.3 A): A diagram of a B-form DNA double helix in which the major and minor grooves are highlighted.3 B): The W-C base pairs that occur within the double helix of DNA. A and T share 2 W-C H bonds while G and C share 3 H bonding contacts. The anti-parallel nature of DNA is also shown; one strand runs in the 5’ to 3’ direction where its complement runs from 3’ to 5’...... 4 Figure 1.4 A): The conformational equilibrium about the glycosidic bond experienced by purine nucleosides demonstrated here using dG as an example. The more favoured anti conformer exposes the W-C H-bonding face; this allows for bonding within the DNA helix. In comparison, flipping of the nucleobase by 180° about χ, exposes the Hoogsteen face, allowing for alternative H-bonding contacts to occur. B) : An example of a Hoogsteen H-bonding pair between dG and dC+. Note that it is required for dC to be protonated...... 6 Figure 1.5A): The planar structure of a G-tetrad. The presence of a metal ion (M+) is shown here. The metal is most often found to be K+, Na+ is also capable of stabilizing this structure. B) Various structures of GQ (top, from left to right): Tetramolecular parallel, bimolecular anti- parallel, intramolecular anti-parallel, (bottom, from left to right), intramolecular antiparallel and an intramolecular hybrid GQ structure...... 8 Figure 1.6: Representative CD spectra for (A) anti-parallel, (B) parallel and (C) hybrid GQ structures...... 12 Figure 1.7: The SELEX process. A random library of DNA oligos is applied to a column containing a target molecule, tethered to a solid support. Unbound DNAs that show no binding capacity to the target are washed off of the column after application buffers containing high concentrations of salt. Bound DNAs will be washed off of the column following the addition of water. DNA sequences that demonstrate binding to the target will be amplified by PCR and re- applied to the column. This process will proceed until the oligo with the best binding capacity is obtained...... 15 Figure 1.8: Jablonski diagram describing the energy level of a fluorophore and possible route for relaxation following excitation by a light photon...... 16 Figure 1.9: A) The structure of the 2fur dU B) Schematic representing the detection strategy employed using the 2fur dU base modification in TBA for the detection of thrombin protein. .. 18 Figure 1.10: A schematic representation of a molecular rotor probe with the example of known molecular rotor fluorophore, p-(dimethylamino) stilbazolium (p-DAPSMI).27 ...... 20 Figure 1.11: Jablonski diagram representing A) LE emission from a molecular rotor and B) relaxation from a TICT state of a molecular rotor probe...... 21

Figure 1.12: The structures of a number of cyanine dyes that can be classified as molecular rotor probes...... 22 Figure 1.13: Acid-base exchange of the Brooker’s merocyanine derivative as reported by Wu et al.37 ...... 23 Figure 1.14: The ESIPT process illustrated using the known ESIPT fluorophore, HBT.41 ...... 24 Figure 1.15: The tautomeric exchanges proposed for a prodigiosin analogue, inspiring the analysis of the novel dyes presented here.42 ...... 25 Figure 2.1 : The structure of Brooker’s merocyanine ...... 30 Figure 2.2: Structural comparison between azo dye (left) 5-(2-benzothiazolylazo)-8-hydroxy quinoline (H-BTHQ) and a merocyanone derivative synthesized for this work (right), 4-[2-(4- hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q)...... 33 Figure 2.3: The enol-keto or azo-hydrazone tautomerism that occurs for the azo dye, H-BTHQ.17 ...... 34 Figure 2.4: The UV absorbance traces of A) PhOMe P, B) PhOMe Q and C) PhOMe Btz in solvents of decreasing polarity: H2O (---), MeOH (---), EtOH (---) and iPrOH (---) ...... 42 Figure 2.5: Illustrating the charge separated state of the p-anisaldehyde derivatives, illustrated here with the PhOMe P example. The more delocalized form of the dye is favoured in less polar solvents...... 43 Figure 2.6: Absorbance traces of (top row, from left to right) PhOH P, Q and Btz and (bottom row, left to right) PhOMe P, Q and Btz in solvents of decreasing polarity: H2O (---), MeOH (---), EtOH (---) and iPrOH (---) ...... 44 Figure 2.7 A) The enol form of PhOH Q that predominates in an aqueous environment of water that is stabilizes by H-bonding of the dye with the solvent. B) Tautomerization of the enol form of the dye to the keto form in a less polar environment of isopropyl alcohol...... 47 Figure 2.8: Fluorescence emission traces of A) PhOH P, B) PhOH Q, C) PhOH Btz and D) PhOH Ind in solvents of decreasing polarity: H2O (---), MeOH (---), EtOH (---) and iPrOH (---). This emisson corresponds to enol emission as solvent polatiry decreases. Irel values are representative in the fluorescence emission change in water versus iPrOH...... 50 Figure 2.9: A comparison of the fluorescence characteristics of the enol and keto tautomers of A) PhOH Btz and B) PhOH Ind in ethanol (---) and methanol (---). Dashed lines are representative of excitation spectra. There is no excitation spectra shown for the enol form of the indolinine derivative as its fluorescent emission is almost non-existant and there is no evidence of an excitation band for this emission wavelength observed in the excitation spectra presented for the keto emission at λem 560 nm...... 52 Figure 2.10: Visible changes in absorbance maxima for (from the top) PhOH P, PhOH Q, PhOH Btz and PhOH Ind. The pH of the solution is denoted below each respective vial...... 53 Figure 2.11: The resonance forms for merocyanine dyes as they become deprotonated, represented here by PhOH Ind...... 54 Figure 2.12: Absorbance traces for (from the top) PhOH P, PhOH Q, PhOH Btz, and PhOH Ind as a function of pH. The arrows indicate an increase in red shifted absorbance as pH increases...... 56

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Figure 2.13: The Hammett plot obtained by plotting literature σ- values for a variety of para substituted phenols against the log of the ratio between the equilibrium constant of the ionization of phenol and the equilibrium constant of the test reactions (depicted as black diamonds). Key p- phenol compounds are highlighted in blue and green. The novel merocyanine dyes are represented on this plot as yellow circles (σ- values calculated using the Biggs ρ value) and red squares (σ-values calculated using the ρ value presented here)...... 58 Figure 2.14: The structures of p-propenylphenol, p-nitrophenol and p-cyanophenol. The colour here represents the colour they are represented in on the Hammett plot in Figure 2.12...... 58 Figure 2.15: Chemical structure of the HOMA reference compound, 6-bromo-2-methylsulfanyl- 1,3-benzothiazole...... 64 Figure 2.16: Fluorescence emission spectra for PhOH P (top left), PhOH Q (top right), PhOH Btz (bottom left) and PhOH Ind (bottom left) in solutions of increasing pH, excited at the wavelength corresponding to the isosbestic point observed in the UV data...... 70 Figure 2.17: Jablonski diagrams demonstrating ET process undergone by PhOH P in the excited sate to result in an excited TICT state of the molecule. Relaxation from the TICT state is achieved by non-radiative decay pathways. This figure uses PhOH P as an example. This figure is also meant to represent the quinoline merocyanine derivative...... 72 Figure 2.18: Jablonski diagram describing the emission of PhOH Btz and PhOH Ind in solutions of pH greater than 7. Here, PhOH Ind represents both dyes...... 73 Figure 3.1: A) Antiparallel GQ topology of Htelo in the presence of Na+. B) Parallel GQ topology of Htelo in the presence of K+, crystallographic interpretation. C) Hybrid GQ topology of Htelo in the presence of K+...... 82 Figure 3.2: A) The crystal structure of TBA in complex with its protein target thrombin. B) A simplified rendering of the TBA G4. The G residues involved in this G4 are found in alternating syn- and anti-conformations. The G residues that make up the G4 are bolded within the aptamer sequence ...... 83 Figure 3.3: Chemical structures of a number of GQ binding fluorophores...... 84 Figure 3.4: Structural comparison of ThT to one of the novel merocyanine dyes presented in this work...... 85 Figure 3.5: Glycerol studies of the merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBTz at a concentration of 5 µM dye in solution, increasing glycerol content in solution from 0 to 80%...... 91 Figure 3.6: Duplex to quadruplex exchange assay of 1 µM (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Orange traces represent the fluorescence emission of the free dye, blue traces represent the fluorescence intensity of the dye with the addition of 0.5 µM TBA:C2 duplex DNA and purple traces represent the emission of the dye in the presence of TBA:C2 duplex after the addition of 1 µM thrombin protein...... 93 Figure 3.7: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the absence of metal cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration...... 98 Figure 3.8: Comparing antiparallel and parallel GQ structures. The antiparallel-basket topology (A) demonstrates a barrier to pi-stacking as the loop region crests the top of the GQ. The parallel

xiii topology (B) boasts smaller loop regions, none of which interfere with the planar faces of the GQ making them readily amenable to pi-stacking interactions...... 99 Figure 3.9: Demonstrating the preferred resonance forms of PhBtz and PhQ respectively. The Btz derivative prefers the iminium form due to the electron withdrawing ability of the benzothiazole acceptor. The quinoline example prefers the resonance form that maintains aromaticity within the acceptor moiety, making the Q derivative a better candidate to under go pi-stacking with G-tetrads of the parallel GQ. Larger arrow head signifies the preferred resonance structure...... 99 Figure 3.10: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the presence of 50 mM Na+ cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration...... 102 Figure 3.11: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the presence of 50 mM K+ cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration...... 106 Figure 3.12: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer. The increase in colour saturation correlates with the increase in Htelo concentration...... 109 Figure 3.13: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer. The increase in colour saturation correlates with the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra...... 110 Figure 3.14: The visible change in absorbance maximum wavelength for (top) PhBtz and (bottom) ThBtz as well as the intense fluorescence turn on for both dye derivatives in the presence of Htelo DNA, no metal cation in solution...... 113 Figure 3.15: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM NaCl. The increase in colour saturation correlates with the increase in Htelo concentration...... 114 Figure 3.16: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM NaCl. The increase in colour saturation correlates with the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra...... 115 Figure 3.17: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM KCl. The increase in colour saturation correlates with the increase in Htelo concentration...... 119 Figure 3.18: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM KCl. The increase in colour saturation correlates with

xiv the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra...... 120 Figure 3.19: Structure of the non-rotor analogue of ThT...... 123 Figure 3.20: ET excitation and emission spectra of 5 µM ThBtz in (A) 50 mM Tris (B) 50 mM Tris with 50 mM NaCl an (C) 50 mM Tris with 50 mM KCl with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.5, 2.5, 4, and 5 µM) using λex 256 nm...... 129 Figure 3.21: A) The conformational equilibrium that occurs for the ThBtz dye derivative. Because the planar state of the dye occurs when the dye is free in solution, this explains the high baseline fluorescence for the dye. The planar conformation is suspected to prefentialy bind with Htelo, explaining the enhancement in ET emission and low Kd B) Explaining the increase in fluorescence emission of PhQ in the presence of Htelo. Although this dye is not expected to undergo through resonance, binding with the GQ will induce a more planar conformation of the dye, explaining the increase in ET fluorescence emission. C) Binding isotherms for ThBtz (blue trace) and PhQ (red trace)...... 134 Figure 3.22: A) Binding of ThT to the parallel GQ of Htelo. The side view of the complex shows the distortion of the top tetrad that results from binding of the fluorophore. The top view demonstrates how ThT stack on the tetrad in the presence of Na+. B) Binding of ThT to the anti- parallel GQ of Htelo. It is readily observed from the side view that there is no distortion in the tetrad interacting with the fluorophore. The top view shows the staking of ThT with the top tetrad. Reprinted with permission from Mohanty, J.; Barooah, N.; Dhamodharan, V.; Harikrishna, S.; Pradeepkumar, P. I.; Bhasikuttan, A. C. J. Am. Chem. Soc. 2013, 135, 367. ©2013 American Chemical Society. C) A comparison of the structure of the dyes discussed in these studies with that of ThT...... 135 Figure 4.1: The molecular structure of Ochratoxin A ...... 143 Figure 4.2: A) The DNA sequence of OTAA36 B) Pictorial demonstration of a G-tetrad C) The proposed GQ structure of OTAA36 upon target binding, shown here housing a potassium metal ion ...... 146 Figure 4.2: A schematic demonstrating the “turn on” platform. The OTA:OTAA36 complex is subject to λex=256 nm. ET from the DNA to the toxin causes an enhancement in emission at λem= ~430 nm...... 147 Figure 4.4: A) Fluorescence titration of 1 µM OTA with increasing concentrations of OTAA36 (0 to 1.2 µM) with λex 375.). B) Fluorescence titration of 1 µM OTA with increasing concentrations of OTAA36 (0 to 1.1 µM) with λex 256 nm ...... 152 Figure 4.5: The binding isotherm describing the affinity with which OTA binds to OTAA36. OTA (1 µM) was excited at λex 256 nm as increasing concentrations of OTAA36 (0 to 1.2 µM) were added to the solution...... 154 Figure 4.6: The CD spectra of 3 µM of OTAA36 (black trace) and equimolar amounts of OTA and OTAA36 (3 µM) to give the OTA:OTAA36 complex in 3 µM concentration (red trace) in OTA binding buffer obtained at 10 oC...... 155

λex 256 nm. The log of OTA:OTAA36 complex is plotted against the log of the fluorescence emission intensity. The linear range is found from 0.01 to 1 µM of toxin: aptamer complex .. 157 Figure 4.8: Calibration curve obtained by plotting the concentration of OTA:OTAA36 complex against fluorescence emission intensity at 430 nm...... 157 Figure 4.9: The molecular structure of OTB...... 159

Figure 4.10: A comparison of the Irel values obtained for all of the fluorescence titrations carried out for this study: the titration of OTA with OTAA36 (green), the titration of OTA with TBA (blue) and the titration od OTB with OTAA36 (red)...... 160 Figure AB-1: 300 MHz 1H NMR of 3-(3-hydroxypropyl)-2-methylbenzothiazolium bromide (1) in DMSO-d6 ...... 170 Figure AB-2: 300 MHz 13C NMR of 3-(3-hydroxypropyl)-2-methylbenzothiazolium bromide (1) in DMSO-d6 ...... 170 Figure AB-3: 300 MHz 1H NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOH Btz, 5a) in DMSO-d6 ...... 171 Figure AB-4: 300 MHz 13C NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOH Btz, 5a) in DMSO-d6 ...... 171 Figure AB-5: 300 1H NMR of 2-[2-(4-methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOMe Btz, 5b) in DMSO-d6 ...... 172 Figure AB-6: 300 MHz 13C NMR of 2-[2-(4-methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOMe Btz, 5b) in DMSO-d6 ...... 172 Figure AB-7: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-4-methylpyridinium bromide (2) in DMSO-d6 ...... 173 Figure AB-9: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-4-methylpyridinium bromide (2) in DMSO-d6 ...... 173 Figure AB-8: 300 MHz 1H NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOH P, 6a) in DMSO-d6 ...... 174 Figure AB-10: 300 MHz 13C NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOH P, 6a) in DMSO-d6 ...... 174 Figure AB-11: 300 MHz 1H NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOMe P, 6b) in DMSO-d6 ...... 175 Figure AB-12: 300 MHz 13C NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOMe P, 6b) in DMSO-d6 ...... 175 Figure AB-13: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-4-methylquinolium bromide (3) in DMSO-d6 ...... 176 Figure AB-14: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-4-methylquinolium bromide (3) in DMSO-d6 ...... 176 Figure AB-15: 300 MHz 1H NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q, 7a) in DMSO-d6 ...... 177 Figure AB-16: 300 MHz 13C NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q, 7a) in DMSO-d6 ...... 177 xvi

Figure AB-17: 300 MHz 1H NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOMe Q, 7b) in DMSO-d6 ...... 178 Figure AB-18: 300 MHz 13C NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOMe Q, 7b) in DMSO-d6 ...... 178 Figure AB-19: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-2,3,3-trimethylindolenium bromide (4) in DMSO-d6...... 179 Figure AB-20: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-2,3,3-trimethylindolenium bromide (4) in DMSO-d6...... 179 Figure AB-21: 300 MHz 1H NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl indolineuim bromide (PhOH Ind, 8) in DMSO-d6...... 180 Figure AB-22: 300 MHz 13C NMR 300 MHz of 2-[2-(4-hydroxyphenyl)ethenyl]-1- hydroxypropyl indolineuim bromide (PhOH Ind, 8) in DMSO-d6 ...... 180 Figue AC-1: Energy transfer excitation and emission spectra for PhP in 50 mM Tris buffer with the addition of increasing concentrations of Htelo...... 182 Figure AC-2: Energy transfer excitation and emission spectra for PhP in 50 mM Tris with 50 mM Na+ with the addition of increasing concentrations for Htelo...... 182 Figure AC-3: Energy transfer excitation and emission spectra for PhP in 50 mM Tris with 50 mM K+ with the addition of increasing concentrations of Htelo...... 182 Figure AC-4: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with the addition of increasing concentrations of Htelo...... 183 Figure AC-5: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with 50 mM Na+ with the addition of increasing concentrations of Htelo...... 183 Figure AC-6: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with 50 mM K+ with the addition of increasing concentrations of Htelo...... 183 Figure AC-7: Energy transfer excitation and emission spectra for PhBtz in 50 mM Tris buffer with the addition of increasing concentrations of Htelo...... 184 Figure AC-8: Energy transfer excitation and emission spectra for PhBtz in 50 mM Tris buffer with 50 mM Na+ with the addition of increasing concentrations of Htelo...... 184 Figure AC-9: Energy transfer excitation and emission spectra for PhBtz in 50 mM tris buffer with 50 mM K+ with the addition of increasing concentrations of Htelo...... 184 Figure AC-10: Binding isotherms for PhP with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+...... 185 Figure AC-11: Binding isotherms for PhQ with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+...... 185 Figure AC-12: Binding isotherms for PhBtz with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+...... 186 Figure AC-13: Binding isotherms for ThBtz with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+...... 186

AC-14: Calibration curve for free OTA using λex of 256 nm...... 187

xvii

Figure AC-15: Negative control titration of OTAA36 with OTB...... 187 Figure AC-16: Negative control titration of OTA with the thrombin binding aptamer (TBA). . 188

xviii

LIST OF SCHEMES

Scheme 2.1: Synthesis of Activated Compounds ...... 35 Scheme 2.2: Synthesis of Merocyanine Dyes ...... 37 Scheme 3.1: Structure of the Merocyanine Dyes ...... 87

xix

LIST OF ABBREVIATIONS

2fur dU 2-Furan deoxyuridine

A adenine au arbitrary unit

C cytosine

CAPS N-cyclohexyl-3-aminopropanesulfonic acid

CD circular dichroism

CT charge transfer

CV Crystal violet dA deoxyadenosine p-DASPMI p-Dimethylamino stilbazolium dC deoxycytidine dG deoxyguanosine

DNA deoxyribonucleic acid

DMSO dimethyl sulfoxide dsDNA double-stranded DNA dU deoxyuracil

EM electromagnetic

ESIPT excited state intramolecular proton transfer

ESPT excited state proton transfer

ET energy transfer eT electron transfer

EtOH ethanol

FRET fluorescence (Forster) resonance energy transfer

G guanine

GQ guanine quadruplex

H-bond hydrogen bond

HBT 2-(2’-hydroxyphenyl)benzothiazole

H-BTHQ 5-(2-benzothiazolylazo0-8-hydroxyquinoline

HeLa immortal cell line from Henrietta Lacks

HOMO highest occupied molecular orbital

HPLC high performance liquid chromatography

HRMS high resolution mass spectrometry

Htelo human telomeric repeat sequence

ICD induced circular dichroism

ICT intramolecular charge transfer iPrOH isopropyl alcohol (2-propanol)

Irel relative intensity

IUPAC International Union for Pure and Applied Chemistry

Kd dissociation constant

L-CPL left-handed circularly polarized light

LC liquid chromatography

LE locally excited

LUMO lowest unoccupied molecular orbital

MeOH methanol

MES 2-(N-morpholino)ethanesulfonic acid

MOPS 3-(N-morpholino)propanesulfonic acid

MS mass spectrometry

NICS nucleus independent chemical shift

xxi nm nanometer

NMR nuclear magnetic resonance

Oligo oligonucleotide

OTA ochratoxin A

OTAA36 ochratoxin A Aptamer

OTABB ochratoxin A binding buffer

OTB ochratoxin B

PAGE polyacrylamide gel electrophoresis

PBS phosphate buffered saline

PCR polymerase chain reaction

PhBtz 2-[2-(4-N,N methylhydroxyethylphenyl) ethenyl]-1-methyl benzothiazolium iodide PhP 4-[2-(4-N,N methylhydroxyethylphenyl) ethenyl]-1-methyl pyridinium iodide

PhQ 4-[2-(4-N,N methylhydroxyethylphenyl) ethenyl]-1-methyl quinolium iodide

PhOH Btz 2-[2-(4-hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide

PhOMe Btz 2-[2-(4-methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazoliumbromide PhOH Ind 2-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl indolineuim bromide

PhOH P 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide

PhOMe P 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide

PhOH Q 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide

PhOMe Q 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide pKa negative log of the acid ionization constant

QBS quinine bisulfate

R-CPL right-handed circularly polarized light

Rh 101 rhodamine 101

xxii

RNA ribonucleic acid s seconds

S0 ground state

S1 and S2 singlet excited states

SELEX Systematic Evolution of Ligands by Exponential enrichment

ΔS Stokes' shift ssDNA single-stranded DNA

T thymine

TAPS tris[(hydroxymethyl)methyl amino] propanesulfonic acid

TBA thrombin binding aptamer

ThBtz 2-[2-(2-N,N methylhydroxyethylthienyl)ethenyl]-1-methyl benzothiazolium iodide

ThT Thioflavin T

TICT twisted intramolecular charge transfer

TMPyP4 5,10,15,20-tetrakis (N-methyl-4-pyridl) porphyrin

TO thiazole orange

Tris tris(hydroxymethyl)aminomethane

U uracil

UV ultraviolet

UV-Vis ultraviolet visible v/v volume per volume

WC Watson-Crick

ε molar extinction coefficient

λ wavelength

ρ Hammett reaction constant

σ Hammett substituent constant

xxiii

σ- substituent constant for electron withdrawing groups

ϕ quantum yield

χ glycosidic bond angle

xxiv

Chapter 1: Introduction

1.1 Deoxyribonucleic Acid 1.1.1 Components of DNA Deoxyribonucleic acid (DNA) is a naturally occurring biopolymer that contains the genetic information required of all living organisms to necessitate growth and development. Each strand of DNA is composed of individual subunits called nucleotides which are comprised of three building blocks: a phosphate moiety, a 2’deoxyribose sugar as well as a nitrogenous base.1 The nitrogenous base is adjoined to the deoxyribose sugar by an N-glycosidic bond (χ) (Figure 1.1).1

In the absence of the phosphate moiety, the molecule is then referred to as a nucleoside.1 When removing both the phosphate and the sugar entities, what remains is known as a either a nitrogenous base or a nucleobase.1 The DNA strand itself is held together by phosphodiester bonds which are achieved when the 3’ hydroxyl group of one nucleotide is linked to the 5’ hydroxyl group of a subsequent nucleotide to give rise to the sugar phosphate backbone.2 When nucleotides are arranged in a single strand in this manner, where the sequence of bases is clearly discernable when read from the 5’ to 3’ end of the strand, this arrangement is also referred to as the primary structure of DNA.2

There exist four different nitrogenous bases that are observed in DNA: adenine (A), thymine

(T), cytosine (C), and guanine (G) (Figure 1.2).2 These bases can be categorized into two different groups as either a pyrimidine or purine nitrogenous base.1,2 The pyrimidines, C and T, exhibit a single heterocyclic ring structure while the purine bases, A and G, exhibit a fused heterocyclic ring motif.1,2

Figure 1.2: An oligonucleotide demonstrating the phosphodiester backbone of DNA where linkages occur at the 3’hydroxyl of one nucleotide and the 5’hydroxyl of an adjacent one. In addition, all purine and pyrimidine nitrogenous bases are shown.

1.2 The structure of DNA 1.2.1 The DNA Double Helix The global structure of DNA was elucidated in 1952 by Drs. Watson and Crick with major contributions from Rosalind Franklin.2 It was discovered that the most biologically relevant form

3 of DNA exhibits a double helical secondary structure.2 Two separate strands of DNA that run in antiparallel directions to form the double helix (Figure 1.3 A).2 The antiparallel nature of DNA is defined by the fact that one strand runs in the 5’ to 3’ direction while its complement runs from 3’ to 5’.2 DNA strands can demonstrate free hydroxyl groups at both of their 5’ and 3’ ends.2 Within the body, the 5’ end of DNA will often carry a phosphate group as DNA replication carries out in the 5’ to 3’ direction.3 The right handed helix of DNA, in its most biologically important B-form, exhibits both a major and a minor groove.3 The major groove presents a larger solvent exposed surface area of the interior of the helix in comparison to the minor groove.3

Major [

Minor [

PDB 1BNA

Figure 1.3 A): A diagram of a B-form DNA double helix in which the major and minor grooves are highlighted.3 B): The W-C base pairs that occur within the double helix of DNA. A and T share 2 W-C H bonds while G and C share 3 H bonding contacts. The anti-parallel nature of DNA is also shown; one strand runs in the 5’ to 3’ direction where its complement runs from 3’ to 5’.

The interior of the helix is composed of nucleobase pairs.2 The DNA bases hold the double helix together by forming Watson-Crick (WC) hydrogen bonds in which each purine base will pair with a pyrimidine.2 G will pair with C sharing three W-C H-bonding contacts.2 T will pair with

A, demonstrating only two H-bonding interactions (Figure 1.3 B).2 WC H-bonding is only

4 possible when a purine nitrogenous base is found in an anti-conformation about the N-glycosidic bond it shares with the 2’deoxyribose sugar.2 This is defined by a chi angle between 110° to 180°.2

This conformation is favoured as it limits steric clashing between the nucleobase and the sugar moiety.2

1.2.2. Hoogsteen Base Pairs WC H-bonding that gives rise to the traditional base pairs as they are known is perhaps the most biologically relevant form of H-bonding undergone by the DNA nitrogenous bases.

However, as these bases demonstrate a variety of functional groups within their structure, it is not surprising that these molecules demonstrate the ability to form alternate H-bonding contacts than those that form the canonical WC base pairs of the double helix. For instance, it is also possible for the DNA bases to produce what are known as Hoogsteen base pairs. Hoogsteen base pairs are made possible by flipping of a purine base about the N-glycosidic bond, χ, it shares with the 2’ deoxyribose sugar by 180°, resulting in the syn conformer (Figure 1.4 A).4 The first instance of these contacts was reported by Hoogsteen in 1963 after characterizing the H-bond contacts between the nucleoside mimics 1-methylthymine and 9-methyladenine.5 Hoogsteen base pairs have been observed between A and T as well as between G and C. The latter example is only observed under acidic conditions, when deoxycytidine is in its protonated form (Figure 1.4 B).4

This alternative H-bonding capacity of the DNA bases gives rise to a variety of secondary structures. An example of one such structure, the guanine quadruplex (G-quadruplex) that is key to this work will be discussed in detail in sections 1.2.4.

Figure 1.4 A): The conformational equilibrium about the glycosidic bond experienced by purine nucleosides demonstrated here using dG as an example. The more favoured anti conformer exposes the W-C H-bonding face; this allows for bonding within the DNA helix. In comparison, flipping of the nucleobase by 180° about χ, exposes the Hoogsteen face, allowing for alternative H-bonding contacts to occur. B) : An example of a Hoogsteen H-bonding pair between dG and dC+. Note that it is required for dC to be protonated.

1.2.3. The Guanine Quadruplex As discussed previously, DNA oligonucleotides can adopt a variety of different conformations based on the high degree of chemical functionality observed within their structure.

Examples of alternative DNA tertiary structures include triplex structures, stem-loops or hairpins as they are also known, as well as i-motif structures.6 A DNA tertiary structure that is of utmost interest to the Manderville laboratory research efforts is the G-quadruplex (GQ). DNA sequences that exhibit G rich sequences have the potential to form GQ structures.6,7 GQs are comprised of stacked layers of planar structures called G-tetrads.6,7 A G-tetrad is composed of four G residues that interact with each other via Hoogsteen H-bonding (Figure 1.5 A).6,7 The central cavity of the

G-tetrad houses a cationic metal ion which is most often observed to be potassium (K+). The affinity GQs have for K+ ions can be explained by the lower solvation energy K+ demonstrates in comparison to other monovalent ions such as Li+ and Na+.6 The presence of this cationic metal ion is required to neutralize repulsive forces by the O6 oxygens of guanine involved in forming this coordinate structure.7

GQs are highly polymorphic in their ability to adopt a variety of conformations. This structure can form both inter- and intramolecularly.7 Tetra- and bimolecular GQs have been observed where four and two strands come together to form this structure respectively.7 A single strand of DNA that exhibits a large proponent of G residues within its sequence can fold in upon itself to give rise to a GQ.6 Similar to the DNA double helix, GQs can display an anti-parallel directionality but are capable of demonstrating a parallel topology as well. In a unimolecular anti- parallel GQ, the G residues of each tetrad will be found in alternating syn and anti conformations.

In contrast, within a parallel GQ, the G residues will all be found in the preferred anti configuration about the N-glycosidic bond of its structure.7 Additionally, GQs can be found as hybrid structures in solution.7-9 This is observed when a GQ is not found in strictly anti-parallel of parallel conformations, but rather the GQ exhibits characteristics of both of these at once. Examples to illustrate the diversity observed among GQ structures can be seen in Figure 1.5 B. An example of a DNA sequence capable of adopting a number of GQ topologies, the human telomeric repeat sequence, will be discussed in detail in Chapter 3.

Figure 1.5A): The planar structure of a G-tetrad. The presence of a metal ion (M+) is shown here. The metal is most often found to be K+, Na+ is also capable of stabilizing this structure. B) Various structures of GQ (top, from left to right): Tetramolecular parallel, bimolecular anti-parallel, intramolecular anti-parallel, (bottom, from left to right), intramolecular antiparallel and an intramolecular hybrid GQ structure.

1.3. Analysis of DNA Structure 1.3.1. An Introduction to the Methods A variety of analytical techniques have been used to analyse the structural facets DNA possesses. The highly conjugated nature of the DNA nucleobases lend this molecule to analysis via ultraviolet-visible (UV-vis) spectroscopy. In addition, based on the chiral nature of DNA, the use of circular dichroism has been employed to study the nature of the double helix based on the

8 molecules response to circularly polarized light. These methods are keystone techniques when studying DNA in any context and are utilized exhaustively within this work.

1.3.2. Analysis of DNA using UV-Vis Spectroscopy UV-Vis spectroscopy is often referred to synonymously as absorption spectroscopy.

Molecules that possess pi and/or non-bonding electrons have the ability to absorb light energy and become excited from their ground state to occupy an excited state.10 The readout obtained when the absorbance of a molecule is measured indicates the wavelength of light the molecule can absorb as well as how absorbent the molecule is.10 This refers to how much light energy the molecule in question is capable of taking in. The lower the energy gap that exists between the ground state and the excited state of a molecule indicates that the molecule will be able to absorb photons of a lower energy that vibrate at longer wavelengths. 10

DNA exhibits absorbance within the UV range of the electromagnetic (EM) spectrum around 260 nm. This characteristic lends DNA to analysis via UV-vis spectroscopy. In fact, UV-

Vis is the standard protocol used to quantify the amount of DNA within a sample. This is done through the use of the Beer-Lambert Law which describes the ability of a molecule to absorb light energy as a function of concentration of the analyte in solution.10 This is a linear relationship where the amount of light absorbed by the molecule is directly proportional to its concentration.

The concentration of analyte can be determined from the equation:

퐴 = 휀푐푙

Where A represents the absorbance of the analyte, a unitless measurement, c the concentration often expressed in molarity (M), l the path length the incident beam will travel through the sample, measured in centimetres (cm) and ε the molar extinction coefficient which has units of M-1 cm-1.

The extinction coefficient is a constant value that is unique to the analyte. This constant represents the amount of light a molecule attenuates at a specific wavelength of light. It is important to note that different DNA oligonucleotides will exhibit different extinction coefficients based on the make up of the base sequence.11 The molar extinction coefficient for DNA is dependent on monitoring the absorbance of the molecule at 260 nm. At this wavelength only can an accurate concentration of single stranded DNA be determined.11 Interestingly, GQ DNA also demonstrates the ability to absorb light at a slightly more red-shifted wavelength than duplex or single stranded DNA.12 GQ structures have a propensity to absorb light of wavelengths from 260 nm to 290 nm.12 This feature of the GQ has been utilized to determine accurate melting temperatures for the structure.12 Monitoring the melting curve at 290 nm as opposed to 260 nm with increasing temperatures provides a better understanding of the dynamics of the GQ specifically.12 Although GQ structures demonstrate the ability to absorb at 290 nm provides a more accurate means to determine thermal melting temperatures for this structure, absorbance can still be observed from GQ DNA at 260 nm. This absorbance characteristic of the GQ allows for visualization of GQ formation in solution as a band around 260 nm becomes more apparent in absorbance and fluorescence excitation spectra. This characteristic of the GQ will be discussed and exploited in Chapter 3.

1.3.4. Circular Dichroism Spectroscopy Circular dichroism (CD) is a property that is inherent to molecules that bear chiral chromophores. This characteristic describes how the molecule will react to both right-handed (R-

CPL) and left-handed circularly polarized light (L-CPL).13 The term “dichroism” in this sense refers to how the molecule will absorb both types of polarized light. Depending on the nature of the chromophore, one type of polarized light will be absorbed preferentially over the other giving rise to both positive and negative readings.13 CD is a function of the wavelength of light the analyte 10 is exposed to.13 This explains why both positive and negative signals can be observed for the same molecule; the amount of either L-CPL or R-CPL that will be absorbed by the molecule changes as the wavelength of light varies.

CD spectroscopy is a useful tool when it comes to characterizing the tertiary structures of macromolecules. Two biological examples that this technique is best applied to are proteins and

DNA.13-15 Considering the chiral nature of these species, CD spectroscopy is easily applied to the study of the global structure of these biological molecules. What is of most interest to these studies is utilizing this technique to determine what type of GQ is predominantly found in solution based on a variety of factors the DNA under study is exposed to.14,15 This is easily done as GQs in solution are readily distinguished from one another based on their characteristic CD spectra. For instance, an anti-parallel GQ exhibits positive CD bands at 290 and 240 nm and a negative signal around 260 nm.15,16 In contrast, a parallel GQ will only demonstrate two CD bands, a positive band around 260 nm and a negative signal at 240 nm.15,16 Hybrid GQ structures will demonstrate a broad CD signal centered at 290 nm.16 Additionally, a hybrid GQ will have a small negative signal around 240 nm (Figure 1.6).16 The obvious differences observed in the spectral readout of varying GQ structures makes CD spectroscopy the primary method for distinguishing what conformation of the structure predominates in solution.

A) 1

0.5

0 220 240 260 280 300 320

-0.5 CD (mdeg) CD -1

-1.5 Wavelength (nm)

1.5 B) 1

0.5

CD (mdeg) CD 220 230 240 250 260 270 280 -0.5

-1 Wavelength (nm)

1.5 C)

0.5

CD (mdeg) CD 0 220 240 260 280 300 320 -0.5 Wavelength (nm)

Figure 1.6: Representative CD spectra for (A) anti-parallel, (B) parallel and (C) hybrid GQ structures.

1.4 An Introduction to Biosensors A large topic of research in the field of biotechnology today is developing platforms that provide simple, fast, sensitive, and reliable methods for detecting a variety of target molecules.

Presently used techniques, such as high-performance liquid performance chromatography

(HPLC), optical spectroscopy, and mass spectrometry are elaborate, expensive and time consuming approaches to indicate the presence of a target entity.17 The development of novel biosensor platforms presents an opportunity to create more cost effective, simpler and more user

12 friendly means for detection in comparison to the traditional and more complex methods currently used today.

The International Union of Pure and Applied Chemistry (IUPAC) defines a biosensor as a device that will provide a quantitative analytical signal upon recognition of a target entity by a biological recognition element.18 The biological recognition element will elicit a signal by acting through a transducer element.18 Target molecules can range from small molecules, viruses, to proteins as well as toxic metal ions.17 The recognition element within the system is most often a biological macromolecule.17 This can include protein antibodies, nucleic acids, or protein enzymes. Each of these demonstrate great selectivity for their target molecules.17 The exchange that occurs between the target molecule and its recognition element will result in some quantitative analytical signal by acting through a physical transducer.17 There are many different classes of transducer elements. A few examples being: optical, which results in changes in UV absorbance or fluorescence within the system, electrochemical, which registers differences in current or potential in the presence or absence of a target and colorimetric transducers, exhibiting the appearance, disappearance or change of colour upon target binding.17 The signal that it is produced by the systems’ transducer element in the presence of a target molecule is the basis for detection.17

A bourgeoning area of research in the vein of developing new methods for detection is creating nucleic acid based platforms through the use of DNA aptamers.

1.5. Aptamers 1.5.1. Aptamers: A Definition Although DNA is an essential macromolecule to all forms of life, it can also be used for various biotech applications such as developing aptamers. Aptamers receive their name from the latin word aptus which means “to fit.”19 They are single stranded nucleic acids which bind to a target molecule with high affinity and selectivity.19,20 These nucleic acids can be oligomers of

13 either RNA or DNA and are synthesized in vitro.19,20 Aptamers are capable of binding to a variety of target molecules. In fact, when this concept was first introduced in the early 1990s, RNA oligonucleotides were shown to bind to small molecules, such as dyes, as well as protein targets.19,20 Aptamers will adopt a specific structural geometry upon binding to a target molecule and are selected for using a process called in vitro selection which uses affinity chromatography and series of wash steps to obtain an oligonucleotide which bind to a desired target. 19 Many also refer to this process as the systematic evolution of ligands by exponential enrichment (SELEX).20

1.5.2. SELEX The SELEX process has shown that a random library of oligonucleotides (oligos) bind to a molecule of interest with increasing affinity after each subsequent cycle of the process.19,20 The affinity with which an oligo binds to the target increases exponentially as the process progresses.19,20 The selection of an oligo that binds to a target molecule proceeds via affinity chromatography to discriminate candidate oligos from oligos that show no binding capacity for the target what so ever.19,20 A target molecule is tethered to the solid support of a column to which a random library of oligonucleotides are applied.19,20 Oligos demonstrating little to no affinity for the target molecules are washed away after applying buffers that contain high salt concentrations.19,20 Nucleic acids, demonstrating a substantial net negative charge, and that show no binding capacity are easily washed away with the addition of cations to the column. Bound oligos demonstrating inherent affinity for the target are eluted from the column after the application of water.19,20 Oligos that demonstrate inherent binding are amplified using the polymerase chain reaction (PCR).19,20 This new library of amplified oligos are applied to a column containing the target molecule again, and the process continues.19,20 In order for this process to work, various conditions such as salt concentrations and the use of negative controls are varied as the cycles

14 progress.19,20 These steps ensure that the DNA aptamer with the highest affinity for the target molecule possible is obtained while discarding nucleic acids that are less apt to bind to the molecule of interest (Figure 1.7).

PCR and Reapplication to column

Figure 1.7: The SELEX process. A random library of DNA oligos is applied to a column containing a target molecule, tethered to a solid support. Unbound DNAs that show no binding capacity to the target are washed off of the column after application buffers containing high concentrations of salt. Bound DNAs will be washed off of the column following the addition of water. DNA sequences that demonstrate binding to the target will be amplified by PCR and re- applied to the column. This process will proceed until the oligo with the best binding capacity is obtained.

1.5.3. GQ Forming Aptamers As mentioned previously, once an aptamer binds to a target molecule, the single stranded oligo will undergo a conformational change that allows for favourable contacts to occur between the aptamer strand and the target entity. One such structure the aptamer strand can adopt upon target binding, granted that the DNA sequence is rich in G residues, is the GQ. Two aptamer systems pertinent to this work are the thrombin: thrombin binding aptamer (TBA) and the

Ochratoxin A: Ochratoxin A aptamer (OTAA) examples.21-23 Both of these aptamers are either

15 known to or are hypothesized to adopt a GQ upon target binding. These examples be discussed in detail in Chapters 3 and 4.

1.6. Fluorescent Probes for Aptamer Based Detection 1.6.1. An Introduction to Fluorescence Although a variety of analytical techniques are amenable to aptamer based detection of a variety of target molecules, one method of particular interest to this work is the use of fluorescence spectroscopy. Fluorescence is defined as the emission of a light photon from a compound after it

10,24 has been excited by a light source. The molecule is excited from a ground state (So) to a singlet

24 excited state (S1 or S2). If excited to an S2 state, the molecule can undergo internal conversion to an S1 state. The molecule can then relax back to its ground state from an S1 excited state via loss of thermal, vibrational, or light energy.24 The latter of these is referred to as radiative decay and one such path a molecule can follow to relax is by way of fluorescence (Figure 1.8). 24 Fluorescence spectroscopy measures the fluorescence intensity of emitted light photons from a molecule when it is excited by light of a specific wavelength.24

Figure 1.8: Jablonski diagram describing the energy level of a fluorophore and possible route for relaxation following excitation by a light photon.

The use of fluorescent probes in developing aptamer based detection platforms presents many advantages over other strategies. Fluorescence can be dependent upon the environment in which a fluorophore resides. This indicates that any changes in the probes environment brought about by target binding will change the probes’ fluorescent behaviour.12 This can be caused by the direct interaction of the probe with a target entity or changes in the probe’s microenvironment as a result of nucleic acid topology changes, indicating that fluorescence offers a more sensitive approach to detection in comparison to other techniques. The changes in fluorescent behaviour experienced by the probe can also be referred to as the probe’s ability to switch on or off.

Switching capability is essential when developing probes that will be used for detection purposes.6

In addition, because DNA is not inherently fluorescent and does not exhibit strong absorbance outside of the UV region of the EM spectrum, it is necessary to introduce a transducer element of some kind. This can be done through the use of internal base modification, as well as through the use of free dye assays.

1.6.2. Internal Base Modification Arguably, the modification of a DNA base that resides within an aptamer sequence itself or replacement of a base with a fluorescent base mimic provides an assay with more sensitivity.

This is explained by the ability to strategically place the fluorescent modification at a location within the aptamer strand that optimizes contact with the target entity. Additionally, the modification can be placed at a position where the topology of the aptamer strand readily changes upon target binding. This can either lead to an enhanced turn on or turn off signal. In either case, the optical changes as a result of target binding in comparison to the unbound state would be starkly different, signalling a binding event. Drawbacks to modifying an internal base are often costly and elaborate synthetics pathways. The use of an internally modified aptamer assay, however, will

17 often prevail as this methodology allows for an increased sensitivity to the method that can sometimes be absent in end label and free dye approaches.

1.6.2.1. An Example of a Fluorescently Modified Internal Base An example of an internal base modification, in this case, a modification that is able to sense the presence of a target molecule upon incorporation into an aptamer sequence is that of a furan modified deoxyuridine (dU) base (2fur dU) (Figure 1.9).25 This internal base modification has been used within the Manderville research laboratory for the detection of the protein target thrombin. The 2fur dU probe was strategically incorporated into a site within TBA at a position within the sequence that was known to interact with thrombin protein.22,25 Upon introduction of the target protein to a TBA: complementary strand duplex, as thrombin is able to bind with its aptamer stripping the duplex structure apart, a turn on fluorescence response is observed from the

2fur dU internal probe.25,26 In this case, the 2fur dU probe behaves as a molecular rotor probe.

Briefly, as the protein binds to the aptamer strand, interacting with 2fur dU in doing so, free rotation within the molecule is restricted. This forces the fluorophore into a planar conformation, allowing for the observed increase in fluorescence emission.27,28

B) A)

Figure 1.9: A) The structure of the 2fur dU B) Schematic representing the detection strategy employed using the 2fur dU base modification in TBA for the detection of thrombin protein.

Although the work presented here does not involve the use of internally modified DNA, it is important to provide background on past strategies utilized in our laboratory in order to set the stage for the current work. In most cases, the use of internally modified fluorescent base mimics will be of preference to use in aptamer-based detection platforms due to the inherent sensitivity they possess. However, it is also of interest to develop simple, elegant and cost-effective detection strategies as well. This provides a rationale for the studies presented here. It was the 2fur dU studies that provided great insight into the prospect of molecular rotor probes and their use in nucleic acid based detection platforms. More details on molecular rotor probes will follow in section 1.6.3. The application of molecular rotor probes to the detection of DNA topologies will be discussed in depth in Chapter 3.

1.6.3. Molecular Rotor Probes Fluorescent probes that exhibit a different fluorescent behaviour that occurs as a result of environmental changes it experiences are of utmost interest. One class of probes that behave in this manner are molecular rotor fluorescent dye molecules. There are a number of structural characteristics that define a fluorophore as a molecular rotor. These dyes exhibit at least one bond about which free rotation can occur. In addition, there will be an electron donor moiety adjoined to an electron acceptor moiety, often linked together through an electron rich alkene bridge. The presence of the bridging olefin allows for electron movement between the donor and acceptor units

(Figure 1.10).27

In the ground state, a rotor dye will often be found in the planar state. Upon excitation, the fluorophore can undergo an intramolecular charge transfer (ICT) process as the donor moiety donates charge density to the acceptor.29 This can induce rotation within the molecule to give rise to a twisted intramolecular charge transfer (TICT) state.29 From the TICT state, the preferred route

19 for the fluorophore to relax back to a ground state is via non-radiative pathways.27,29 Ultimately, in the twisted state, it is preferential for the molecule to relax via thermal and rotational means.29

Relaxing in this manner indicates that there will be no emission of a photon from the analyte, making this TICT state of the molecule non-fluorescent. In addition, there is the possibility for the fluorophore to become locally excited to give rise to a locally excited (LE) state of the molecule.27,29 LE emission of the dye is achieved when the fluorophore is locked into its planar state in the ground state.27 Upon excitation by a photon, the planar state of the molecule will be excited and relax in the planar state, allowing for LE emission from the fluorophore resulting in a fluorescent signal.27 For molecular rotor probes, forcing the dye into its planar conformation in the ground state by increasing the viscosity of the solvent or by introducing a binding agent that stabilizes the planar form of the dye by pi stacking interactions will allow for an enhancement in the fluorescence emission of the dye (Figure 1.11).27,29,30

Figure 1.10: A schematic representation of a molecular rotor probe with the example of known molecular rotor fluorophore, p-(dimethylamino) stilbazolium (p-DAPSMI).27

Figure 1.11: Jablonski diagram representing A) LE emission from a molecular rotor and B) relaxation from a TICT state of a molecular rotor probe.

1.6.4. Cyanine Dyes: Examples of Molecular Rotor Probes The cyanine dyes are a class of dye that have been used historically as photosensitizers for developing photographs.31,32 Structurally they are characterized by two heteroatom containing aryl moieties connected by one or more methine groups.31 Symmetrical cyanine dyes contain the same heteroaryl rings adjoined by a polymethine chain, while asymmetric cyanine dyes contain two different heteroaryl moieties.31

The cyanine family of dyes are classified as molecular rotor probes owing to the presence of the electron acceptor component, a cationic nitrogen containing aromatic ring adjoined with an electron donor moiety by at least one carbon-carbon double bond. Examples of cyanine dyes that have been used in various nucleic acid detection platforms are SYBR Green as well thiazole orange

(TO) (Figure 1.12).33,34 In fact, these dyes are widely accepted alternatives for use in nucleic acid staining.33,34 These dyes will exhibit a turn on fluorescence response upon interaction with DNA.

This can be explained by the ability for the dyes to be locked into a planar conformation upon stacking with DNA in solution.34 Because the cyanine family of dyes have demonstrated the ability to effectively bind to DNA, it makes these fluorophore prime candidates for application in aptamer based detection platforms and even for visualization of nucleic acids structures within cells. For

21 these reasons, expanding the knowledge on how the structure of a dye affect its ability to interact with DNA provides a precedence for the studies presented herein.

In addition, the cyanine family of dyes are not only capable of behaving as molecular rotor probes. A cyanine dye of particular interest to this work is Brooker’s merocyanine (Figure 1.12).35

The rotor characteristics of this probe are no exploited in these studies, however, the ability for this dye to participate in acid-base exchange in aqueous media make it an interesting target in the development of pH probes. This will be explained in depth in Section 1.6.5.

Figure 1.12: The structures of a number of cyanine dyes that can be classified as molecular rotor probes. 1.6.5. pH Sensing Probes pH sensing probes that are able to elicit a colorimetric or fluorescence response as a function of the pH of its microenvironment are highly sought-after tools, particularly within the realm of biology. Most cancer cells demonstrate a lower intracellular pH as a result of an increased expression of lysosome structures within the cell. Where healthy cells exhibit an intracellular pH around 7.2, cancerous cells demonstrate a pH inside the cell wall at values as low as 6.7.36 An important goal when it comes to cancer imaging is to develop probes that demonstrate the ability to distinguish healthy cells from cancerous ones with a high degree of selectivity and sensitivity in doing so.37,38 Additionally, the ability to create probes that absorb and emit light within the

22 visible region of the EM spectrum, so as not to destroy tissues upon irradiation with a UV light source, is another important facet to consider when designing dyes for this purpose.

Wu and colleagues were able to synthesize an analogue of Brooker’s merocyanine dye that

37 demonstrates a pKa value of 6.44. This dye derivative possesses an indolinine acceptor component in comparison to Brooker’s dye that bears a pyridine acceptor.37 This dye was readily up taken by a culture of cancerous cells, namely HeLa cells, to achieve a turn on fluorescence response by being deprotonated within the malignant cells.37 This dye presents as an easier synthetic target than other pH probes that have been utilized for the same purpose.37,38 In addition, it bears a pKa value consistent with the intracellular pH of cancer cells, allowing for the added level of sensitivity for the assay as well as boasting excitation and emission maximum wavelength that reside with in the visible region of the EM spectrum (Figure 1.13).

Figure 1.13: Acid-base exchange of the Brooker’s merocyanine derivative as reported by Wu et al.37

It was because of these studies that allowed for the current work presented here.

Ultimately, it is a goal of the research presented in this work to create pH sensing fluorophores that are simple to synthesize and that provide an intense fluorescence response upon participating in an acid-base exchange within an aqueous environment. It is also a goal with this research to understand what structural facets of the dye species contribute to the best switching properties of the pH sensing fluorophore.

1.6.6. Excited State Proton Transfer Fluorescence Fluorophores that bear acid-base labile functional groups can under go excited-state proton transfer (ESPT) processes in solution.39 It is possible for the fluorophore in question to undergo proton transfer with its surrounding environment.40 It is also possible for the fluorophore to undergo excited-state proton transfer intramolecularly (ESIPT).39,41 When a fluorophore becomes excited, take for example a phenolic compound, the observed pKa of the molecule when it is in an

* 39,41 excited state (pKa ) has been observed to readily decrease. Because of this phenomenon, it is possible for the ionizable functional group to transfer this proton within it structure as a result.39,41

Essentially, ESIPT allows for a tautomeric exchange to occur within the structure allowing for a ground state enol species to, once it is excited, undergo tautomerism to give rise to a keto form of the molecule. This is illustrated in Figure 1.14 using a well known ESIPT fluorophore, 2-(2’- hydroxyphenyl)benzothiazole (HBT).41

Figure 1.14: The ESIPT process illustrated using the known ESIPT fluorophore, HBT.41

The ability for a molecule to undergo ESIPT or proton transfer with its surrounding environment allows for large changes to be observed in the fluorescence and absorbance behaviour of the fluorophore. For one, as the molecule adopts its keto form, it is documented in the literature that the fluorescence emission wavelength of the chromophore red-shifts in comparison to the emission of the enol tautomer.41,42 The same is true of the absorbance maximum and excitation maximum wavelength for the fluorophore as well.39,42 This is readily explained by an increase in conjugation within the system as a result of keto formation. The possibility of an increase in

24 conjugation within the system as a result of keto formation readily explains the red-shift observed as the enol form undergoes this exchange to the keto.

The phenomenon of ESPT as it relates to the optical changes that are observed from a given fluorophore as the hypothesized tautomeric exchange occurs has been noted within the

Manderville laboratory previously.42 For example, it has been observed that analogues of the natural product prodigiosin can undergo keto-enol tautomerism as a function of solvent polarity

(Figure 1.15).42 This is attributed to an ESIPT process that is favoured to occur in protic solvents.42

This knowledge is what inspired the in-depth analysis of the novel merocyanine dyes presented within this work and their ability to partake in ESPT processes. Although this has not been noted in the literature for this class of dyes, it was of interest to analyze the impressive solvatochromic changes of the phenolic merocyanine dyes to determine if these impressive spectral changes could be explained by ESPT and by the possibility of a keto-enol tautomeric exchange.

Figure 1.15: The tautomeric exchanges proposed for a prodigiosin analogue, inspiring the analysis of the novel dyes presented here.42

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Chapter 2: Synthesis and Characterization of Novel Phenolic Merocyanine Dye Derivatives

2.1 Introduction 2.1.1. Brooker’s Merocyanine Brooker’s merocyanine was first introduced in the 1950s.1 The search for photosensitizing agents to develop colour photographs in this era lead to the synthesis of Brooker’s dye, 4-[2-(4- hydroxyphenyl)ethenyl]-1-methylpyridinium iodide.1 The synthetic pathway to achieve this dye has remained similar over the decades, requiring activation of the heterocyclic component with an alkylhalide to give the quaternary amine salt.1-3 This activated methylene compound then undergoes condensation with an aldehyde donor component via a Knovenegel mechanism in the presence of an organic base in an alcoholic solvent to give the desired dye.1,4 The name

“merocyanine” is derived from the Greek word “mero” which means “part.” The traditional cyanine dye is comprised of two heterocyclic ring components joined by a polymethine bridge.5

The merocyanine does not have the second heterocycle moiety, explaining the origins of its given name. Instead, in the case of Brooker’s example, the donor component is a phenol ring (Figure

2.1). The family of dyes presented here have a similar structural framework, explaining the direct comparison to Brooker’s dye.

Figure 2.1 : The structure of Brooker’s merocyanine

Brooker’s dye is a highly interesting molecule. It demonstrates astonishing changes in its absorbance and fluorescence characteristics as a function of the solvent the dye is exposed to.6

This merocyanine dye is known to exhibit negative solvatochromism, meaning that as the polarity of the solvent decreases, the absorbance maximum wavelength of the chromophore increases.5,7

As a result, Brooker’s dye demonstrates vast colour changes in a variety of solvents. It had previously been hypothesized that the dye undergoes resonance in solution between its zwitterionic form to give a covalent quinoid species.8 Studies have ultimately shown that this is not the case, and that the covalent quinoid species does not exist.9 However, in low polarity solvents there is literature precedence to suggest that based on the C-O bond length of the of the phenol moiety that the dye exists as an amalgam of both zwitterionic and quinoid forms.9 This alludes to the generation of a more delocalized, charged separated state of Brooker’s dye in non-polar solvents, explaining the observed negative solvatochromism.7,9 As polarity of the solvent decreases, the charge separated form of the molecule is favoured and explains why absorbance wavelengths are red-shifted; the increase in electron delocalization requires less energy, or light of longer wavelengths, to excite the neutral form. Further insight into why this is observed will be explained in section 2.2.1.

2.1.2. Introduction to the Hammett Equation The novel merocyanine dyes presented in this work bear phenol as the donor component within the dye structure. Unsubstituted phenols are known to participate in acid-base exchange in aqueous solution, demonstrating an approximate pKa of 10. Because these dyes bear phenol substituents, it was of interest to probe how the spectral properties of these chromophores would change as a function of pH and to quantify the ability of these derivatives to partake in acid-base processes based on the acceptor moiety present. This was achieved through the use of the Hammett equation. The Hammett equation is used to describe how the structure of a compound and the substituents present within a molecules core structure relate to its reactivity.10,11 This relationship was first explored using the dissociation of benzoic acid.10,12 The equation can be shown as:

퐾 푙표푔 ( ) = 𝜎𝜌 퐾0

31 where K0 is the K value of the reference reaction, K, the equilibrium constant for the test reaction, sigma (σ) the substituent constant and rho (ρ) the reaction constant.11-13 This relationship can ultimately be used to describe any chemical equilibrium.

The substituent constant describes how substituents on a benzene ring affect the observed

11,13 pKa of the ionizable functional group within the structure of the molecule. Substituents can have effects on the test reaction through induction or through resonance.11 For our purposes, there is reason to believe that inductive forces play a limited role in the electron withdrawing ability of the acceptor component of the dyes. Primarily, the electron withdrawing ability through resonance of the merocyanine dyes is explored and quantified in section 2.3.5. through the use of σ- values.

The σ- value is used to describe the resonance electron withdrawing ability of a given substituent based on the generation of a negative charge at the benzylic position of the molecule.13,14

2.1.3. Effects of Keto-Enol Tautomerism on Solvatochromic and Fluorosolvatochromic Properties of Hydroxy Merocyanine Dyes As mentioned in Chapter 1, a variety of dyes that possess ionizable functional groups that can readily participate in acid-base exchange can undergo tautomeric exchange in solution as well as ESIPT processes to result in vast changes in their absorbance and fluorescence capabilities.

Examples previously discussed are that of HBT and various prodigiosin analogues.15,16 The general trend observed is that when the enol form of the dye tautomerizes to the keto form, this results in a red-shift in absorbance maximum wavelength of the dye as well as an enhancement of the absorbance capability of the chromophore.15,16 Additionally, there is also often a red shift in fluorescence emission wavelength.15,16 To the best of our knowledge, the ability for Brooker’s dye and the varying analogues that will be discussed herein to undergo keto-enol tautomerism has not been explored in depth. Because of the phenol functional group and the ability for these dyes

32 to undergo acid-base exchange, it is highly probably that this equilibrium process will be observed for the phenolic merocyanine family of dyes presented within this work.

To further support the hypothesis that phenolic merocyanine dyes are capable of undergoing keto-enol exchange in solution, the hydroxy azo family of dyes can be used to provide a rationale.17,18 Azo dyes are a well classified family of dyes that have been used in a variety of applications: as colourants, in non-linear optics and various other applications in materials chemistry.17 The reason azo dyes are used in this scenario to illustrate the possibility for keto-enol tautomerism to occur for the merocyanines is due to the similarity in their structure (Figure 2.2).

Although the donor and acceptor components of the azo dyes are separated by a N-N double bond where the merocyanine are joined by a C-C double bond, the comparison stems from the similarity in their structural framework where the donor and acceptor moieties are joined by a pi bond, allowing for a fully conjugated dye species.

Figure 2.2: Structural comparison between azo dye (left) 5-(2-benzothiazolylazo)-8-hydroxy quinoline (H-BTHQ) and a merocyanone derivative synthesized for this work (right), 4-[2-(4- hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q).

The tautomerism of azo dyes is well documented in the literature.17-19 Taking H-BTHQ as an example, the solvatchromism that results when exposing this dye to solvents of varying polarity indicates the enol/azo form of the dye is preferentially formed in non-polar environments.17

Intuitively, the keto/hydrazone species predominates in more polar solvents (Figure 2.3).17 This

33 is known to be the case as the absorbance maximum wavelength of the dye red-shifts substantially as solvent polarity increases.17

Figure 2.3: The enol-keto or azo-hydrazone tautomerism that occurs for the azo dye, H-BTHQ.17

Although the merocyanine dyes presented in these studies do not bear the bridging N-N double bonds the azo dyes do, these dyes are only used here to aid in the illustration of the potential keto-enol tautomerism the phenolic dyes are capable of participating in. Because this is observed for this class of chromophore, it is highly likely that the solvatchromic properties observed for the novel merocyanines dyes can be rationalized by a similar process as seen in the azo family of dyes.17,18

2.2. Materials and Methods 2.2.1. General Synthesis of Merocyanine Dyes The group of merocyanine dye derivatives presented here were designed to contain a variety of cationic nitrogen containing heterocyclic acceptor moieties, while the phenolic donor component remained the same for most of these examples. These compounds were synthesized according to the literature with some modifications.1,2

2.2.1.1. General Procedure - Synthesis of Activated Methylene Compounds The heterocyclic acceptor of choice (10.8 mmol) was combine with 3-bromo-1-propanol

(7.2 mmol) in acetonitrile (5 mL) and heated to reflux for 18 hrs. Alternatively, the reaction mixture was exposed to microwave radiation through the use of a CEM microwave reactor to a

34 temperature of 90 oC under variable pressure for 30 min-1 hr. The activated methylene compound was isolated as a precipitate or as a viscous liquid after the addition of ether and cooling at -18 oC.

Scheme 2.1: Synthesis of Activated Compounds

2.2.1.2. Synthesis of Activated Compounds

3-(3-Hydroxypropyl)-2-methylbenzothiazolium bromide (1)20: Synthesis was performed as

1 described to afford 712.9 mg of a white solid (34% yield). H NMR (300 MHz, DMSO-d6) δ =

8.48 (d, J= 8.7 Hz, 1H), 8.45 (d, J= 8.1 Hz, 1H), 7.87 (m, 1H), 7.85 (m, 1H), 4.78 (t, J= 7.2 Hz,

13 2H), 3.51 (t, J= 5.7 Hz, 2H), 3.23 (s, 3H), 2.03 (m, 2H). C NMR (300 MHz, DMSO-d6) δ =

177.03, 140.61, 129.06, 128.81, 127.76, 124.43, 116.55, 57.12, 46.68, 30.21, 16.64.

1-(3-Hydroxypropyl)-4-methylpyridinium bromide (2)21: Synthesis was performed as described to afford 1.32 g of a viscous, orange liquid (80% yield). 1H NMR (300 MHz, DMSO- d6) δ = 8.94 (d, J= 6.0 Hz, 2H), 7.97 (d, J= 6.6 Hz, 2H), 4.61 (t, J= 7.2 Hz, 2H), 3.42 (t, J= 6.0 Hz,

13 2H), 2.59 (s, 3H), 2.04 (m, 2H). C NMR (300 MHz, DMSO-d6) δ = 158.65, 143.92, 128.17,

57.72, 57.06, 33.16, 21.31.

1-(3-Hydroxypropyl)-4-methylquinolium bromide (3)22: Synthesis was performed as described

1 to afford 2.6 g of a grey solid (100% yield). H NMR (300 MHz, DMSO-d6) δ = 9.44 (d, J= 6.0

Hz, 1H), 8.54 (m, 2H), 8.24 (m, 1H), 8.05 (m, 2H), 5.08 (t, J= 7.2 Hz, 2H), 3.50 (t, J= 5.7 Hz, 11.4

13 Hz, 2H), 2.99 (s, 3H), 2.06 (m, 2H). C NMR (300 MHz, DMSO-d6) δ= 158.46, 148.71, 136.72,

135.02, 129.49, 128.88, 127.13, 122.57, 119.30, 57.38, 54.72, 32.04, 19.69.

1-(3-Hydroxypropyl)-2,3,3-trimethylindolenium bromide (4)23: Synthesis was performed as described to afford 1.05 g of a viscous, purple liquid (49% yield). 1H NMR (300 MHz, DMSO- d6) δ = 7.94 (dd, J= 2.4 Hz, 4.2 Hz, 1H), 7.82 (m, 1H), 7.61 (m, 2H), 4.53 (t, J= 6.9 Hz, 2H), 3.53

(t, J= 5.7 Hz, 2H), 2.83 (s, 3H), 2.04 (s, 6H), 2.01 (t, J= 6.0 Hz, 2H). 13C NMR (300 MHz, DMSO- d6) δ = 195.54, 141.83, 141.11, 129.29, 128.89, 123.43, 115.37, 57.86, 54.09, 45.58, 29.62, 21.76,

13.91, 1.11.

2.2.1.3. General Procedure - Synthesis of Merocyanine Dyes The activated methylene compound (3.5 mmol) was solubilized in ethanol (5 mL) with the addition of catalytic amounts of piperdine. The reaction was stirred at room temperature for 20 minutes, after which 4-hydroxybenzaldehyde or p-anisaldehyde (5.3 mmol) was added to the vessel. The reaction mixture was then heated to reflux for 1 hr. The product was isolated as a solid after the addition of ether and/or dichloroacetic acid with cooling to -18 oC to facilitate precipitation of the dye from solution. When necessary the product was recrystallized from acetonitrile.

Scheme 2.2: Synthesis of Merocyanine Dyes

2.2.1.4. Synthesis of Merocyanine Dyes 2-[2-(4-Hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (5a) Synthesis was performed as described to afford 126.2 mg of purple solid (47% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 10.63 (bs, 1H), 8.42 (d, J= 7.2 Hz, 1H), 8.20 (m, 2H), 7.93 (d, J= 8.7 Hz, 2H), 7.82-

7.71 (3H, m), 6.93 (d, J= 8.7 Hz, 2H), 4.92 (t, J= 6.9 Hz, 2H), 3.54 (2H, bt), 2.03 (2H, m). 13C

NMR (300 MHz, DMSO-d6) δ = 172.06, 162.18, 149.58, 141.23, 132.56, 129.23, 127.99, 127.71,

+ 125.36, 124.25, 116.40, 116.28, 109.72, 57.30, 46.15, 31.35. HRMS Calculated for C18H18NO2S

[M+] 312.1053; found 312.1046.

2-[2-(4-Methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazoliumbromide (5b) Synthesis was performed as described to afford 142.5 mg of yellow solid (48% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 8.43 (d, J= 7.5 Hz, 1H), 8.24 (m, 2H), 8.04 (d, J= 8.7 Hz, 2H), 7.88-7.73 (3H, m),

7.12 (d, J= 8.7 Hz, 2H), 4.95 (t, J= 6.9Hz, 2H), 4.86 (bs, 1H), 3.54 (bt, 2H), 2.03 (t, J= 6.0 Hz,

13 2H). C NMR (300 MHz, DMSO-d6) δ = 172.08, 162.92, 148.96, 141.27, 132.15, 129.32, 128.14,

127.90, 126.81, 124.33, 116.54, 114.83, 110.93, 57.31, 55.69, 46.29, 31.38. HRMS Calculated

+ + for C19H20NO2S [M ]: 326.1209; found 326.1203.

4-[2-(4-Hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (6a) Synthesis was

1 carried out as described to give 1.86 g of red solid (77% yield). H NMR (300 MHz, DMSO-d6) δ

= 8.73 (d, J= 6.9 Hz, 2H), 8.01 (d, J= 6.6 Hz, 2H), 7.89 (d, J= 15.9 Hz, 1H), 7.53 (d, J= 8.7 Hz,

2H), 7.13 (d, J= 15.9 Hz, 1H), 6.73 (d, J= 8.4 Hz, 2H), 4.47 (2H, t, J= 7.2Hz), 3.43 (2H, t, J= 5.7

13 Hz), 2.01 (2H, m). C NMR (300 MHz, DMSO-d6) δ = 163.54, 153.51, 143.76, 141.95, 130.60,

+ 124.35, 122.41, 117.83, 117.16, 116.85, 57.09, 56.94, 33.14. HRMS Calculated for C16H18NO2

[M+]: 256.1332; found: 256.1329.

4-[2-(4-Methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (6b) Synthesis was carried out as described to give 953.3 mg of yellow-brown solid (83% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 8.91 (d, J= 6.9 Hz, 2H), 8.18 (d, J= 6.9 Hz, 2H), 8.02 (d, J= 16.2 Hz, 1H), 7.73 (d,

J= 8.7 Hz, 2H), 7.40 (d, J= 16.5 Hz, 1H), 7.06 (d, J= 9.0 Hz, 2H), 4.55 (t, J= 6.9 Hz, 2H), 3.81

13 (3H, s), 3.46 (t, J= 5.4 Hz, 2H), 2.05 (m, 2H). C NMR (300 MHz, DMSO-d6) δ = 161.19, 153.20,

144.24, 140.77, 129.99, 127.80, 123.21, 120.71, 114.62, 57.33, 57.13, 55.41, 33.14. HRMS

+ + Calculated for C17H20NO2 [M ]: 270.1489; found: 270.1467

4-[2-(4-Hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (7a) Synthesis was performed as described to give 546.5 mg of dark purple solid (83% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 9.27 (d, J= 6.0 Hz, 2H), 9.04 (d, J= 9.0 Hz. 1H), 8.46 (m, 2H), 8.21 (t, J= 7.2 Hz,

1H), 8.12 (bd, 2H), 8.00 (m, 1H), 7.87 (d, J= 7.5 Hz, 2H), 6.91 (d, J= 8.1 Hz, 2H), 4.99 (t, J= 6.3

13 Hz, 2H), 3.52 (bt, 2H), 2.11 (bm, 2H). C NMR (300 MHz, DMSO-d6) δ = 160.57, 153.25,

147.28, 143.82, 137.86, 134.92, 131.22, 128.81, 126.75, 126.46, 118.97, 116.03, 115.31, 57.49,

+ + 54.13, 31.96. HRMS Calculated for C20H20NO2 [M ]: 306.1489; found 306.1464.

4-[2-(4-Methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (7b) Synthesis was performed as described to afford 334.2 mg of dark brown solid (48% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 9.33 (d, J= 3.9 Hz, 1H), 9.07 (d, J= 7.5 Hz, 1H), 8.52 (m, 2H), 8.23 (m, 3H), 7.99

(bm, 3H), 7.09 (d, J= 6.9 Hz, 2H), 5.02 (bt, 2H), 4.81 (bs, 1H), 3.84 (bs, 3H), 2.11 (bt, 2H). 13C

NMR (300 MHz, DMSO-d6) δ = 161.55, 153.12, 147.48, 143.22, 137.85, 134.99, 130.91, 128.92,

128.25, 126.84, 126.58, 117.20, 115.68, 114.57, 57.50, 55.48, 54.252, 31.98. HRMS Calculated

+ + for C21H22NO2 [M ]: 320.1645; found 320.1627

2-[2-(4-Hydroxyphenyl)ethenyl]-1-hydroxypropyl indolineuim bromide (8) Synthesis was performed as described to afford 455.0 mg of dark purple solid (35% yield). 1H NMR (300 MHz,

DMSO-d6) δ = 10.85 (bs, 1H), 8.44 (d, J= 16.2 Hz, 2H), 8.12 (d, J= 8.7 Hz, 2H), 7.86 (m, 2H),

7.60 (m, 3H), 6.80 (d, J= 8.7 Hz, 2H), 4.66 (t, J= 6.9 Hz, 2H), 3.54 (t, J= 5.4 Hz, 2H), 2.01 (m,

13 2H), 1.77 (s, 6H). C NMR (300 MHz, DMSO-d6) δ = 181.52, 173.86, 164.06, 163.39, 159.56,

154.32, 143.39, 140.90, 133.62, 128.93, 128.71, 126.02, 122.93, 116.45, 114.63, 108.91, 65.09,

+ + 57.48, 51.78, 43.67, 30.85, 25.84. HRMS Calculated for C21H24NO2 [M ]: 322.1802; found

322.1799.

2.2.2. Determination of Photophysical Properties Stock solutions of each dye were made by dissolving the solid compound in DMSO to stock concentrations of 2 mM or 4 mM. Solvatochromic measurements were made in solutions of less than 1% DMSO in reagent grade solvent. UV measurements were made in 1000 µL 100-QS quartz cells and observed from 800 nm to 200 nm. Fluorescence measurements were made in

1000 µL 101-QS quartz fluorescence cells using excitation and emission slit widths of 5 nm for

39 the solvatochromic studies and slit widths of 10 nm for the pH studies. For the solvatochromic studies, each molecule was excited at the absorbance maximum centered around 430 nm and the emission monitored to 750 nm. Excitation spectra were obtained using the emission maximum wavelength and monitored from 200 nm to the emission maximum. For the pH studies, the dyes were excited at the wavelength corresponding to the isosbestic point observed in the absorbance traces used to determine pKa. Molar extinction coefficients (ε) were calculated using Beer-

Lambert Law:

퐴 = 휀푐푙

Quantum yields for the merocyanine dyes were calculated using a comparative method, as

24 described in the literature. Quinine bisulfate (QBS, ϕ in 0.5 M H2SO4 = 0.546) and Rhodamine

101 (Rh 101, ϕ in EtOH = 1) were used as fluorescence standards. Values were derived using the equation:

2 퐴푠퐹푥 푛푥 휙푓푥 = ( ) 휙푓푠 퐴푥퐹푠 푛푠

Where the subscripts x and s represent the test molecule and the fluorescence standard respectively.

Ai is the wavelength at which the test compound and the standard overlap in their absorbance spectra, each compound of concentration in solution to give a similar amount of absorbance signal.

Fi is the integration under the curve of the fluorescence emission of the molecule using λex equal to the wavelength of overlap in the absorbance spectrum. The variable ni is representative of the solvent polarity index of the solvent used in the study. ϕfs is the quantum yield of the fluorescence standard.

2.2.3. pKa Determination Citrate buffer (pH 3.0, 3.5, 4.0, 4.5, 5.0), MES buffer (pH 5.5, 6.0, 6.5), MOPS buffer (pH

7.0, 7.5), TAPS buffer (pH 8.0, 8.5, 9.0) and CAPS buffer (pH 9.5 and 10.0) were prepared in

Milli Q water (18.2 MΩ) to a final salt concentration of 50 mM. Phenolic pKz values of the dyes were determined using the equation:

퐴 − 퐴푚 푝퐾푎 = 푝퐻 − 푙표푔 ( ) 퐴푖 − 퐴

Where A is the absorbance of the dye at an analytical wavelength that varies for each dye. This wavelength is the point at which the most change in the absorbance capability of the dye is observed. For example, the absorbance of the dye at an analytical wavelength corresponding to the deprotonated dye species would be used for readings made in all buffered solutions, of pH< 7 through to the most basic pH solutions. Am is the absorbance of the dye in its protonated form

(solution of pH < 7). Ai is the absorbance of the dye in its deprotonated state (solution of pH > 7). pKa values were determined for pH values between 3.0 and 10.0, contingent on the dye under study, and an average of the values was taken.

2.3. Results and Discussion 2.3.1. Solvatochromic Properties of the Merocyanine Derivatives As discussed, Brooker’s merocyanine demonstrates impressive spectral changes when it is exposed to a variety of different solvents, often attributed to the H-bonding capacity of the phenolic oxygen.1,8 It was of interest with this work to assess the solvatochromic properties of the new derivatives to compare these dyes to the classical example of Brooker’s dye. The solvatochromic data for the p- methoxy and p- phenol derivatives are shown in Figure 2.4 and 2.6 and summarized in Table 2.1.

0.3 0.3 A) B) 0.25 0.25 0.2 0.2 0.15 0.15

0.1 0.1 Abaorbance(au) Abaorbance(au) 0.05 0.05 0 0 300 400 500 600 300 400 500 600 Wavelength (nm) Wavelength (nm)

0.45 0.4 C) 0.35 0.3 0.25 0.2 0.15

Absorbance (au) 0.1 0.05 0 300 350 400 450 500 550 Wavelength (nm)

Figure 2.4: The UV absorbance traces of A) PhOMe P, B) PhOMe Q and C) PhOMe Btz in solvents of decreasing polarity: H2O (---), MeOH (---), EtOH (---) and iPrOH (---)

There are a few key observations to be made with these data. First, in each instance of the the p-anisaldehyde donor moitey, as the polarity of the solvent decreases, the maximum absorbance wavelength of the molecule shifts toward the red. As the dye is introduced to water, followed by methanol and increasing the alkyl chain length to ethanol and isopropyl alcohol, the trend is clear.7 This is explained by an increase in charge separation within the dye’s structure as solvent polarity decreases (Figure 2.5) As previously discussed, within the charge separated state, the pi electrons within the system are more delocalized than in the polar ground state of the molecule. Increased delocalization within the chromophore is what allows for a more red shifted

42 absorbance maximum. Although the changes in absorbance wavelength are minimal, it is clear that the methoxy deivatives are still capable of responding to solvent polarity. There are far more marked differences in the absorbance spectra for the phenolic derivatives as they are exposed to solvents of varying polarity.

Figure 2.5: Illustrating the charge separated state of the p-anisaldehyde derivatives, illustrated here with the PhOMe P example. The more delocalized form of the dye is favoured in less polar solvents.

As shown in Figure 2.6, the absorbance traces for the phenolic derivatives demonstrate spectacular changes as the dyes are exposed to varying solvents in comparison to the methoxy derivatives. It is hypothesized that this is a result of the ability for the phenolic merocyanine dyes to undergo prototropic tautomerism in solution.25 It is speculated that the largely red-shifted absorbance bands observed for the phenolic dye derivatives in solvents of decreasing polarity is caused by the existence of a keto form of the merocyanine. In more polar solvents, it is suspected that the enol form of the dye predominates.25

It is possible to use the absorbance characteristics of the methoxy compounds to explain the absorbance characteristics of the perceived enol tautomer observed for the phenolic derivatives.

As previously mentioned, as the PhOMe examples of the dyes are exposed to solvents of decreasing polarity, their absorbance shifts toward the red. In the absorbance spectra of all of the phenolic derivatives, there is an absorbance band observed centered around 400 nm, similar to that of the methoxy compounds. The p- methoxy derivative can be likened to an enol tautomer that is

0.5 A) 0.3 B) 0.4 0.25 0.2 0.3 0.15 0.2

0.1 Absorbance (au) Absorbance (au) 0.1 0.05

0 0 300 400 500 600 350 450 550 650 Wavelength (nm) Wavelength (nm)

0.5 0.4 0.45 C) 0.35 D) 0.4 0.3 0.35 0.3 0.25 0.25 0.2 0.2 0.15 0.15

Absorbance (au) 0.1 Absorbance (au) 0.1 0.05 0.05 0 0 300 400 500 600 350 450 550 650 Wavelength (nm) Wavelength (nm) Figure 2.6: Absorbance traces of (top row, from left to right) PhOH P, Q and Btz and (bottom row, left to right) PhOMe P, Q and Btz in solvents of decreasing polarity: H2O (---), MeOH (-- -), EtOH (---) and iPrOH (---)

“locked” in place; it is an enol form of the molecule that will never undergo exchange to give the keto tautomer. It is clear that the absorbance bands observed for the phenolic derivatives occurring around 400 nm corresponds directly to its enol form based on comparison to the methoxy derivatives. This absorbance band observed for the phenolic compounds also red-shifts in more non-polar environments, similar to the methoxy compounds. A similar argument can be made for the PhOH series of dyes as for the PhOMe dyes. As the phenolic dyes are exposed to solvents of

44 decreasing polarity, the pi electrons of the enol tautomer would also become more delocalized, explaining this red shift in enol absorbance (Figure 2.5).7

Table 2.1: Summary of solvatochromic data for the novel merocyanine dye derivatives.

Solvent Donor Acceptor λmax Solvent Donor Acceptor λmax (nm) (nm) H2O P 376 H2O P 374 Q 420 Q 417 Btz 409, 498 Btz 406 Ind 429, 526 ------MeOH P 393 MeOH P 384 PhOH Q 442 PhOMe Q 431 Btz 424 Btz 414 Ind 445, 543 ------EtOH P 401, 519 EtOH P 385 Q 450, 606 Q 434 Btz 432, 554 Btz 418 Ind 451, 548 ------iPrOH P 405,553 iPrOH P 386 Q 454,643 Q 438 Btz 435 Btz 420 Ind 451 ------

It is important to note that in the absorbance traces taken in water for the PhOH Ind and

PhOH Btz derivatives that the emergence of the red shifted absorbance band is a result of an acid- base exchange undergone by the dye and that it is not necessarily the direct result of keto-enol tautomerism in solution. However, the absorbance changes observed from the phenolic derivatives as they are exposed to solvents of decreasing polarity can be explained by prototropic tautomerism.

As previously mentioned, the azo family of dyes are a class of dye compounds that are known to undergo keto-enol tautomerism in solution. Taking this fact and relating it to the merocyanine dyes discussed herein, it is possible to argue that the same equilbirum process is occuring with these novel merocyanine dyes as well.17,18,25

In the instance of the pyridine and quinoline examples, when they are found in a polar environment of water, they exhibit one absorbance band centered around 370 nm for the pyridine example and 420 nm for the quinoline. It is not until these dyes are introduced to ethanol and subsequently isopropyl alcohol that more substantial differences are noted. In the case of iPrOH, both PhOH P and PhOH Q exhibit largely red shifted absorbance maxima occurring at 553 nm and

643 nm respectively. It is in this non-polar environment that the enol state of the dye would preferenetially tautomerize to give the keto species. The rationale as to why this is observed is due to the H-bonding capacity the dye has with its surrounding environment. When the pyridine and quinoline examples are in a polar environment like water, it is possible for the the phenolic alcohol to H-bond with any surrounding water molecules. This limits the ability for the phenolic moiety to act as an effective donor, preventing the oxygen lone pair from participating in through resonance within the dye’s structure (Figure 2.7).

Conversely, in a more non-polar environment such as isopropanol, there is no longer as much propensity for the solvent to H-bond with the phenolic oxygen of the dye. Here, the oxygen lone pair of the phenol can readily participate in resonance to give rise to a quinoid species. In this state, the positive charge found on the ring nitrogen of the acceptor is maintained on the quinone of the phenolic donor. From this stage, it is possible for the dye to undergo solvent assisted intramolecular proton transfer to alleviate the positive charge found on the phenolic oxygen and to reform the cationic charge on the ring nitrogen of the acceptor moiety (Figure 2.7). 26-28 The keto tautomer of the dyes corresponds to a more red shifted absorbance as the conjugation of the dye increases in comparison to the enol state. With through resonance, the dye is capable of absorbing light of a lower enrgy and longer wavelength, observed in the red-shifted absorbance band that corresponds to the keto form.

Figure 2.7 A) The enol form of PhOH Q that predominates in an aqueous environment of water that is stabilizes by H-bonding of the dye with the solvent. B) Tautomerization of the enol form of the dye to the keto form in a less polar environment of isopropyl alcohol.

As the solvent polarity decreases it is the general trend, at least for the pyridine and quinoline phenolic derivatives, that the amount of keto tautomer observed relates exactly to how non-polar its surrounding environment is. Although this is the opposite of what is observed for the azo family of dyes, it can be argued that the enol form of the merocyanine dyes are favoured in more polar environments due to the H-bonding ability of the phenolic alcohol substituent coupled with the cationic charge located on the ring nitrogen in the enol state. Taking the quinoline derivative as an example, in isopropyl alcohol the keto form of the dye is readily observed based on the large absorbance band at 643 nm. What differs among the derivatives when it comes to the benzothiazole and the indolinine examples, however, is that these dyes preferentially form the keto tautomer in ethanol for PhOH Btz and in methanol for PhOH Ind. It would be expected that these derivatives would also prefer the keto form in the most non-polar solvent of iPrOH. More experiments as to why this is the case will be required going forward. It can be hypothesized however that the electron withdrawing ability of the acceptor component of the dye plays a role in why the keto tautomer is favoured in a variety of different solvents. It is hypothesized that the

47 aromaticity of the acceptor is at play here as well.29 This fact and how it pertains to the acid-base chemistry of the dyes will be discussed in detail in section 2.2.6.29 It is also important to note that in most cases, prototropic tautomerism is often observed in highly polar solvents such as water.16,17,25 In these studies, however, only polar protic solvents were used to assess the solvatochromic properties of the dyes. Although the argument has been made that non-polar solvents favour the keto side of this equilbrium, this is only the case because all of the solvents used here are capable of aiding in proton transfer, due to the presence of the alcohol functional group with in the solvents’ structure. It will be necessary to study these dyes in purely non-polar and polar aprotic environments to fully understand this tautomeric exchange.

What is interesting about this hypothesized tautomeric exchange exhibited by the phenolic dyes is that it quite easily explains the results obeserved by Morley et al in the late 1990s and the perceived presence of a resonance hybrid of the zwitterion and the covalent quinoid species of

Brooker’s dye.9 They justify the presence of this hybrid based on NMR experiments and by comparing the experimental C-O bond length to a literature value of known C=O bond lengths.

Based on their observations, the experimental phenolic C-O bond length does not correspond to either a true carbonyl bond length nor to a carbon-oxygen single bond. This rationalizes the argument that a resonance hybrid of the dye exists in low polarity solvents.9 However, due to the nature of NMR spectroscopy, the time scale within which this equilibrium can be observed is small in comparison to the time scale UV absorbance spectroscopy allows. It is highly probable that in the previous study, mixtures of the enol and keto form of Brooker’s dye were observed in the NMR spectrum. In the UV studies presented here, the time scale is longer such that it is possible for the keto form of the dye species to predominate in solution once the solution is analyzed. However, their findings still indicate that the solvatochromic properties of Brooker’s dye are contingent on

48 the H-bonding capability of the phenolic alcohol.9 With these studies, this finding is certainly the case and we offer an explanation as to why these vast solvatochromic changes are observed.

2.3.2. Fluorosolvatochromic Properties of the Merocyanine Dyes The general trend observed for the phenolic dyes shows that as the polarity of the solvent decreases, going from water to isopropyl alcohol, the fluorescence emission of the enol form of the dyes also increases. This further demonstrates that as the polarity of the solvent decreases, the charge separated enol species predominates leading to an increase in fluorescence emission. This can be explained based on the molecular rotor characteristics of the dyes. As more charge separated dye species is found in solution as solvent polarity decreases which, due to the delocalization of pi electrons, is a more planar form of the dye, there is an increased oppourtunity for the dye to under go LE emission once excited by a photon.30-32 As shown in Figure 2.8, all of the merocyanines demonstrate an increase in fluorescence emission intensity going from water to isopropyl alcohol. As illustrated by the relative intensity (Irel ) values, the pyridine, quinoline, and indoline dyes undergo emission enhancement of no less than 2 intensity units. The quinoline example responds most intensely, demonstrating an Irel value of 10.5.

It was of interest to also probe the fluorescence capability of the keto tautomer of the dyes and, if possible, to utilize the fluorescence characteristics of the dyes when exciting at the most red shifted absorbance maximum wavelength to confirm the formation of the keto tautomer in solution. This was done using the benzothiazole and indolinine phenolic dye derivatives as examples. Although for PhOH Btz and PhOH Ind there was no clear trend as to what solvent allowed for preferential formation of the keto form of the dye, there was clear formation of the speculated keto species in ethanol for the benzothiazole example and in methanol for the indolinine.

140 250 B) 120 A) 200 100 150 80 Irel =2.0 Irel =10.5 60 100 40 50 20

0 Fluorescence Intensity(au) 0 Fluoresceence Fluoresceence Intensity(au) 450 500 550 600 650 450 550 650 Wavelength (nm) Wavelength (nm)

35 C) 14 D) 30 12 25 10 20 Irel =2.2 8 Irel =3.3 15 6 10 4 5 2

0 0 Fluorescence Fluorescence Intensity(au) 450 500 550 600 650 700 Fluorescence INtensity(au) 470 520 570 620 670 Wavelength (nm) Wavelength (nm)

Figure 2.8: Fluorescence emission traces of A) PhOH P, B) PhOH Q, C) PhOH Btz and D) PhOH

Ind in solvents of decreasing polarity: H2O (---), MeOH (---), EtOH (---) and iPrOH (---).

This emisson corresponds to enol emission as solvent polatiry decreases. Irel values are representative in the fluorescence emission change in water versus iPrOH.

When exciting the benzothiazole and indolinine phenolic dyes at the wavelength corresponding to the suspected keto absorbance (Table 2.2), there is a very clear increase in the fluorescence emission intensity for both dyes in comparison to enol emission (Figure 2.9) where

PhOH Btz demonstrates an Irel of 2.3 and PhOH Ind an Irel of 12.8. Additionally, there is a distinct red shift in the emission wavelength. These observations alone provide good evidence to support the formation of the keto tautomer for these examples of the merocyanine dyes. As the keto form of the dye is observed, there is an increase in conjugation within the dye’s structure. This can explain the red shift in emission maxima and the increase in emission intensity; once the keto form

50 predominates in solution, this more conjugated species will be more apt to under go LE emission to relax back to a ground state following excitiation with light.30,32 What is perhaps the most compelling piece of evidence that can justify that the keto-enol equilibirum process is occuring is the decrease in Stoke’s shift when monitoring the fluroescence excitation and emission characteristics of the keto species in comparison the the enol tautomer. This decrease in Stoke’s shift indicates that the ground state of the molecule very closely resembles the excited state of the moelcule as it relaxes.33 In this instance, this indicates that in the ground state, the keto tautomer is present. As the keto form of the dye is excited, because in this form the dyes are far more conjugated and planar, in this state the dye will undergo LE emission with very little change to its global structure as it relaxes, emitting a photon.30,31 In comparison to the enol state, where there is far more oppourtunity for the dyes to undergo a variety of conformational changes in their excited states, there is a much larger difference between the excitation and emission wavelengths.

Based on these observations, it is clear that keto-enol taoutomerism plays a role in both the solvatochromic and the fluorosolvatochromic characteristics of these novel merocyanine dyes.

Although it will be necessary to determine the fluorosolvatochromic properties of the pyridine and quinoline dyes as well, using the benzothiazole and indolinine derivatives to describe the hypothesized existence of keto-enol equilbria is at least the beginnings of understanding the spectacular properties these dyes possess.

40 20 A) B) 35 18 16 30 14 25 12 20 10

15 8 6 10

Fluorescence Fluorescence Intensity(au) Fluorescence Intensity(au) 5 2 0 0 350 450 550 650 750 300 400 500 600 700 Wavelength (nm) Wavelength (nm)

Figure 2.9: A comparison of the fluorescence characteristics of the enol and keto tautomers of A) PhOH Btz and B) PhOH Ind in ethanol (---) and methanol (---). Dashed lines are representative of excitation spectra. There is no excitation spectra shown for the enol form of the indolinine derivative as its fluorescent emission is almost non-existant and there is no evidence of an excitation band for this emission wavelength observed in the excitation spectra presented for the keto emission at λem 560 nm.

Table 2.2: Summary and Comparison of keto and enol fluorescent properties

a Compound λex λem λex λem Emission Emission Irel ΔS ΔS (nm) (nm) (nm) (nm) Enol Keto Enol Keto (enol) (enol) (keto) (keto) (au) (au) (nm) (nm) PhOH Btz 432 507 554 578 14.2 33.0 2.3 75 24 PhOH Ind 445 511 543 560 1.5 19.0 12.8 66 17 aUsed to denote Stoke’s Shift

2.3.3. Absorbance Characteristics as a function of pH The phenolic functional group that contributes to the solvatochromism of these dyes also provides precedent to suspect that these dyes can act as pH-sensing molecules. Indeed, the work presented here demonstrates that these dyes provide a colorimetric signal when exposed to solutions of varying pH. As shown in Figure 2.10, all of the phenolic dyes experience a red-shift in their absorbance maxima as the pH of their environment increases. In the case of PhOH P, the

52 absorbance of this dye shifts from the UV at a λmax value of 378 nm into the visible region where

λmax is found at 449 nm, going from colourless in solutions of pH less than 7 to having a yellow hue under basic conditions. The same thing is observed for PhOH Q, PhOH Btz and PhOH Ind.

These dyes appear red and orange with λmax values found at 513, 498 and 526 nm respectively in solutions of pH greater than 7. Conversely, these dyes appear light green-yellow in solutions that have acidic pH values where these λmax values occur at 420 nm for the quinoline derivative, at 407 nm for the benzothiazole framework and at 425 nm for the indolinine example (Table 2.3)

All of these dyes experience a red-shift of more than 70 nm in their λmax values upon deprotonation (Table 2.3). As the dye becomes deprotonated, this generates a negative charge on the phenolic oxygen. This theoretically would give rise to a zwitterionic dye species as the ring nitrogen bears a positive charge. This zwitterion could become neutral through resonance to result in an uncharged, fully conjugated species (Figure 2.11).

5.5 6 6.5 7 7.5 8 8.5 9 9.5 10

4.5 5 5.5 6 6.5 7 7.5 8 8.5 9

3 3.5 4 4.5 5 5.5 6 6.5 7 7.5 8 Figure 2.10: Visible changes in absorbance maxima for (from the top) PhOH P, PhOH Q, PhOH Btz and PhOH Ind. The pH of the solution is denoted below each respective vial.

The increase in conjugation within the dye’s structure allows the compound to absorb light of a longer wavelength and of lower energy to result in promotion of a ground state electron within the molecule’s highest occupied molecular orbital (HOMO) to its lowest unoccupied molecular orbital

(LUMO).33 As the pi system within the molecule increases, less energy is required for the delocalized electrons within this system to be promoted to an excited state.33

Figure 2.11: The resonance forms for merocyanine dyes as they become deprotonated, represented here by PhOH Ind.

Additionally, there is a noted increase in the absorbance capability of the dyes under basic conditions. This is illustrated by the experimentally determined extinction coefficients for the phenolic derivatives (Table 2.3). The extinction coefficient can also be referred to as the molar absorptivity of a molecule. This value describes how much of the incident beam of light a molecule will prevent from being transmitted through the sample as it becomes absorbed by the analyte of interest.33 In all cases, the extinction coefficient of the dyes increases in basic solution. The ability for the molecule to absorb light is directly related to the level of conjugation observed within its structure. In addition, the λmax values and extinction coefficients observed for the p- methoxy derivatives can be likened to the protonated phenol derivatives (Table 2.3). The fact that these values are so similar for the PhOMe and PhOH derivatives, where the methoxy examples mimic the phenol compounds frozen in a permanently protonated state, further illustrates that the changes observed in the spectroscopic properties for the phenolic compounds are a direct result of deprotonation when they are exposed to basic media.

Table 2.3: A summary of the absorbance characteristics of the phenolic merocyanine dyes.

a a,i b b,i c c,i Compound λmax ε λmax ε Δ λmax Compound λmax ε (nm) (nm) (nm) (nm)

PhOH P 378 35 391 449 40 551 71 PhOMe P 374 33 614 PhOH Q 420 20 614 513 26 414 93 PhOMe Q 413 18 160 PhOH Btz 407 56 380 498 86 526 91 PhOMe 406 37 638 Btz PhOH Ind 425 34 538 526 68 912 101 ------aThese values were compiled in acidic media (pH < 7) bThese values were compiled n basic media (pH >7) cThese values were compiles in MOPS buffer (pH=7) iUnits for ε are M-1cm-1

2.3.4. pKa Determination It is evident that the absorbance properties of these dyes vary substantially with changes in pH. Figure 2.12 shows the absorbance traces of all four of these dyes as they are exposed to buffered solutions of increasing pH. These traces corroborate the claim that absorbance maxima for the molecules experience a red-shift. Additionally, there is a clear isosbestic point in all of the traces. This is representative of the fully protonated dye undergoing resonance to result in the fully conjugated, neutral species once deprotonated. From this data it is possible to determine what the phenolic pKa is for each of the dye molecules.

The benzothiazole example demonstrates a phenolic pKa of 7.76 while the indolinine derivative has a calculated pKa of 7.25. What is perhaps most surprising about the calculated values is that the pyridine and quinoline derivatives boast overtly similar pKa values at 8.50 and

8.54 respectively (Table 2.4), which correspond to values already reported in the literature for the

Brooker’s pyridine dye framework.34-36 An explanation of the observed differences and similarities between the calculated pKa values for the merocyanine dyes will be provided in section

2.2.5. However, what is important to note about the calculated pKa values for these chromophores is that they reside close to if not well within the physiological pH range. This gives a precedent to

55 suspect that these dyes will be amenable to application in the detection of peptidic target molecules, among other biological applications.

2 A) 2 B) 1.8 1.8 1.6 1.6 1.4 1.4 1.2 1.2 1 1 0.8 0.8 0.6 0.6

0.4 0.4 Normalized Absorbance(au) Normalized Absorbance(au) 0.2 0.2 0 0 300 350 400 450 500 550 350 450 550 650 Wavelength (nm) Wavelength (nm)

3 2.5 C) D) 2.5 2 2 1.5 1.5

1 1

0.5 0.5

Normalized Absorbance(au) Normalized Absorbance(au)

0 0 300 400 500 600 300 400 500 600 Wavelength (nm) Wavelength (nm)

Figure 2.12: Absorbance traces for (from the top) PhOH P, PhOH Q, PhOH Btz, and PhOH Ind as a function of pH. The arrows indicate an increase in red shifted absorbance as pH increases.

Table 2.4: Calculated Phenolic pKas for the merocyanine derivatives

Compound pKa

PhOH P 8.50 ± 0.07 PhOH Q 8.54 ± 0.09 PhOH Btz 7.76 ± 0.10 PhOH Ind 7.25 ± 0.11

2.3.5. Hammett Analysis

For the merocyanine dyes, the pKa values can be used to quantify the electron withdrawing ability of the acceptor component of each molecule. This can be done through the use of a

Hammett plot. A number of examples of para substituted phenols and their literature pKa and their substituent constants, σ-, were compiled to generate the Hammett plot shown in Figure 2.13.14

Namely, the literature examples used here are: phenol, p-nitrophenol, p-bromophenol, p-cresol, p- methoxyphenol, p-fluorophenol, p-propenylphenol and p-cyanophenol.37,38 This plot was constructed based on previous studies performed by Biggs and Robinson in the 1960s.37 They sought to apply the Hammett relationship to para and meta substituted phenols and anilines. The majority of substituted phenols used for this study remain the same with some omissions and additions. Because the dyes discussed here bear para vinyl substituents, it was key that at least one of the points along this standard curve be representative of a phenol bearing a similar substituent. The point highlighted in green is representative of p- propenylphenol (Figure 2.14).39

In addition, it was of interest to select examples of para-substituted phenols that demonstrates a similar pKa value to the derivatives presented in this work. For this purpose, p-nitrophenol and p- cyanophenol were kept within this sample as they have phenolic pKas of 7.15 and 7.97 respectively

(Figure 2.14).37-39 They are represented on this Hammett plot in blue.

In order to determine Hammett substituent constants for the novel dye molecules, it is necessary to determine the reaction constant, ρ. This value is the slope of the line that is generated by plotting σ- versus the log of the equilibrium constant for the test reaction over the equilibrium constant for the reference reaction, in this case the ionization of phenol, (ie: log(K/Ko)). Using the literature examples stated above, the plot depicted in Figure 2.13 is achieved.

2.5

) 1.5 0 y = 2.13x + 0.05 R² = 0.99 K/K 1

Log( 0.5

0 -0.4 -0.2 0 0.2 0.4 0.6 0.8 1 1.2 1.4 -0.5

-1 σ-

Figure 2.14: The structures of p-propenylphenol, p-nitrophenol and p-cyanophenol. The colour here represents the colour they are represented in on the Hammett plot in Figure 2.12.

This relationship demonstrates good correlation with an r2 value of 0.99. The ρ value obtained from this plot is found to be 2.13. Although this value is close to the literature value

58 proposed by Biggs of 2.23 using mostly the same values as shown here, to calculate the σ- values for the novel dye derivatives in this work, it was decided to utilize both ρ values.37 Two sets of calculations were performed to determine the Hammett substituents constants of the novel merocyanine dyes using the literature ρ value and the one determined here. This was done in hopes of comparing the calculated σ- values to gain insight into the accuracy of the ρ value presented.

- 37 The Hammett Equation: 푝퐾 − 푝퐾0 = 𝜎𝜌 was used to determine σ values. The pKa values for the merocyanine dyes were determined spectroscopically (Table 2.3) and phenol, used as a

o reference, has a pKa value of 9.92 in solution. The values obtained for the substituent constants using both ρ values discussed and the Hammett equation depicted above are summarized in Table

2.5.

Table 2.5: Summary of the calculated Hammett substituents constants for the activated nitrogen heteroaryl substituents.

Substituent σ- a σ- b P 0.64 0.67 Q 0.62 0.65 Btz 0.97 1.01 Ind 1.20 1.25 a Calculated using the Biggs ρ value of 2.2337 b Calculated using the ρ value presented here of 2.13 *See Scheme 2.1 for the structure of the substituent.

In comparing the calculated σ- values for the merocyanines dyes from the two sets of calculations using the ρ values discussed, it is observed that the values obtained in both cases are very similar. There is good agreement in the substituents constants calculated from the literature

ρ value and the value determined in this work. To illustrate this agreement, data points that represent the merocyanine dyes were added to Figure 2.13 by plotting the calculated σ- versus the log of the ratio of equilibrium constants for both sets of substituent constant values. The yellow points on the graph are representative of the merocyanine dye substituent constants calculated

59 using the Biggs ρ value and the red points are representative of the σ- values calculated using the

ρ value from the newly constructed plot. Both sets of data are observed to be on trend with this plot. Based on this observation, it can be concluded that the Hammett substituent constants calculated here using the reaction constant deduced from the plot above are accurate and can be used to describe the electron withdrawing ability of the acceptor moiety of the merocyanine dyes.

Perhaps the most important comparison to draw in this study, however, is how the calculated σ- values of the acceptor moieties of these dyes compare to the aforementioned standard compounds presented, namely, p- nitrophenol, p-cyanophenol and p-propenylphenol and how this relates to their calculated pKa values. A summary of this comparison is shown in Table 2.6

Table 2.6: Comparison of pKa and σ- values of the merocyanine dyes to the reference compounds p-nitrophenol, p-cynophenol and p-propenyl phenol.

- Substituent pKa σ p-propenyl 9.82 0.08 P 8.50 0.67 Q 8.54 0.65 p-CN 7.97 0.87 Btz 7.76 1.01 Ind 7.25 1.25 p-NO2 7.15 1.27 Values for reference compounds have be bolded for clarity.13 *See Scheme 2.1 for the structure of the substituent.

- First, the calculated σ values demonstrate an inverse proportionality to the pKa. As the

- pKa value for the dyes decreases, the electron withdrawing ability described by the σ value increases. As the acceptor component becomes more able to draw electron density within the system, the pKa value is effectively lowered. Additionally, the pKa values of the dyes and their respective σ- values correlate well based on the reference standards used here.

The pyridine and quinoline derivatives demonstrate nearly equal pKa values and their

Hammett substituent constant are equally similar. In comparison to p-cyanophenol which

- - demonstrates a phenolic pKa of 7.97 and a σ value of 0.87, the σ values for PhOH P and PhOH Q at 0.67 and 0.65 respectively, are fitting and make sense when factoring in their respective pKa values of 8.50 for the pyridine compound and 8.54 for the quinoline example.13,37,38 When comparing these pKa values to that of p-cyanophenol, there is an increase in the value for the merocyanine dyes by approximately 0.5 units. It then makes sense that the σ- value for the substituents presented here are less than that of the reference standard. The same trend is observed when comparing PhOH Btz and PhOH Ind to one of the other choice references, p-nitrophenol.

The nitrophenol compound demonstrates the lowest pKa out of these three at 7.15. It, too, demonstrates the largest σ- value, 1.27.13 The σ- values for these merocyanine dyes once again show good agreement with their pKa. The benzothiazole compound is found to have a pKa of 7.76 and a Hammett substituent constant of 1.01 where the indolinine example is strikingly similar to

- p-nitrophenol, demonstrating a pKa value of 7.25 and a σ value of 1.25.

Interestingly, the p-propenylphenol standard compound demonstrates notably less electron withdrawing character than the other key standards used for comparison in this case. This illustrates the importance of the cationic heterocycle and its ability to contribute to the pKa of the dyes presented. The para vinyl substituent on its own does not contribute a great deal to the observed pKa, however the addition of an electron withdrawing substituent across this double bond allows for lowering of the observed pKa values due to the electron withdrawing of the acceptor component through resonance, through the para vinyl linker.

2.3.6. Insight into Hammett Analysis and pKa Results: What Does it All Mean? These observations beg the question, what exactly is the difference among these

- compounds that accounts for disparity among their pKa and σ values? If all of the compounds presented contain a cationic nitrogen as an electron withdrawing substituent, should their pKa values and Hammett substituent constants not be more similar? The hypothesized answer to this question takes into account the aromaticity or lack thereof observed in the acceptor moiety of the dye molecule.

In order to address the role that aromaticity of the acceptor moiety plays in the observed pKa of the molecule, the Harmonic Oscillator Model of Aromaticity (HOMA) index can be used to provide a rationale as to why the aromatic character of the acceptor is so important.29 It should be stated that for an aromatic molecule, maintaining complete delocalization within the system is far more thermodynamically favoured than to disrupt the π-system and lose aromaticity.40 Based on the resonance forms of these dye molecules as they undergo acid base exchange (Figure 2.10), upon deprotonation that leads to the fully conjugated neutral form of the dye, aromaticity is subsequently lost in the acceptor component to achieve the neutral form of the molecule. The

HOMA index can be used to quantify how aromatic the acceptor components are, explaining the

- differences in calculated pKas and σ values.

The HOMA index is a geometrical explanation of aromaticity based on an optimal C-C bond length within a delocalized π system of 1.388 Å.29,40 Previous indices prior to the introduction of HOMA utilized an average bond length within a conjugated ring system, taking the mean of the entire sum of all C-C bonds within the ring structure.29 This did not take into consideration that all C-C bond lengths would be equal if the system were truly delocalized and instead took the sum of alternating single and double C-C bonds to determine an average bond length. The HOMA is

62 the first index to be applied to heteroaryl species. Again, ideal C-X bonds (where X represents a heteroatom) were approximated and utilized in these cases. The HOMA value for a specific compound provides a quantification of the aromatic character of the given molecule.29

Utilizing literature HOMA values for a number of representative compounds, the aromatic capability of the acceptor moieties presented here can be more readily understood (Table 2.6).

Based on the HOMA index, the most classical example of an aromatic molecule, benzene, demonstrates a HOMA value of 0.98. Similarly, pyridine and quinoline are observed to have values nearing that of benzene at 0.99 and 0.93 respectively. The similarity between the heteroaryl molecules amongst themselves and in comparison to benzene can be explained by the fact that the nitrogen lone pair does not participate in the aromaticity of the ring. Without any contribution from the heteroatom, it makes sense that these values align with the value observed for benzene.

Of the dye molecules presented here, the quinoline and pyridine examples demonstrate the highest

- pKa values, lowest σ values and now are observed to be the most aromatic based on the HOMA index. Because these acceptors possess the most aromatic character, they appear to be less willing to undergo an acid base exchange than the other dyes involved in this study.

Table 2.7 HOMA values for literature example compounds pertinent to the current study.29,41,42

Compound HOMA Ref Benzene 0.98 29 Pyridine 0.99 29 Quinoline 0.93 29 Thiazole 0.73 41 6-Bromo-2-methylsulfanyl-1,3- 0.82 42 benzothiazole

Figure 2.15: Chemical structure of the HOMA reference compound, 6-bromo-2-methylsulfanyl- 1,3-benzothiazole.

The aromatic character of the acceptor can be used to explain the benzothiazole example as well.

Benzothiazole should be markedly less aromatic than both pyridine and quinoline based on the

HOMA indices of 0.73 and 0.82 observed for the reference compounds provided here, namely thiazole and an alternative benzothiazole derivative, 6-bromo-2-methysulfanyl-1,3-benzothiazole

(Figure 2.15). In addition, it has been noted in the literature that as the number of heteroatoms

41 within the system increases, the aromaticity decreases as a result. Indeed, the pKa value for

PhOH Btz at 7.76 is less than PhOH P and PhOH Q by 0.8 pH units and the σ- value also increase in comparison. Because the benzothiazole acceptor demonstrates presumably less aromaticity than the previously discussed examples, there is a lower thermodynamic threshold with this acceptor component, providing an explanation as to why its pKa is demonstrably lower than the aza-benzene derivatives.

The previous statement, whereby the aromaticity of a compound decreases as the number of heteroatoms within the system increases, does not describe the observation made in Table 2.7.

When comparing the HOMA values for pyridine and benzene, it is observed that pyridine is in fact more aromatic than benzene demonstrating a HOMA value of 0.99 where benzene is found to have a HOMA value of 0.98. Although the use of the HOMA index within this work to describe the aromaticity of the acceptor components of these dyes does illustrate, for the most part, the role

64 aromaticity plays in the observed pKa of the novel compounds, it is clear that a geometrical explanation is not the only factor at play in what defines a compound as an aromatic species.

Another criterion that can be used to describe the aromaticity of a compound is the nucleus- independent chemical shift (NICS) an aromatic compound is capable of eliciting on a dummy atom placed within the center of the ring structure.43 This is a computational method that was first devised based on the observed changes in the chemical shift of 7Li once it coordinates to the pi system of an aromatic ring structure.44 Ultimately, NICS is a measurement that determines how much the ring current within the aromatic species contributes to the observed chemical shift of the central atom of the system.43 NICS values are reported as the negative of the shielding effect observed in the test atom.43 For example, if the test atom experiences a shift upfield in the computed NMR spectrum, the observed chemical shift that resulted from exposure to the aromatic species is reported as the negative value.43 The more negative the NICS value, the more aromatic the compound of interest is.43 The upfield shift that is observed from the compound results due to a diatropic ring current within the pi system of the compound.43,45 What this indicates is that the ring current of the pi system generates a magnetic field within the ring that opposes the direction of the magnetic field outside of the aromatic species.43,45

Using this model to explain the aromaticity of the pyridine acceptor, in comparison to an arguably perfect aromatic system, the benzene ring, NICS(1) values for benzene and the pyridinium cation as discussed within this work are observed to be -10.20 and -9.89 respectively

(Table 2.8).43 Additionally, pyridine in its unprotonated state demonstrates an NICS(1) value of

-10.17, which still indicates that this species is not as aromatic at benzene.43 These NICS values corroborate the previous statement with respect to the number of heteroatoms within the aromatic system. Benzene, demonstrating a more negative NICS value is clearly more aromatic than the

65 pyridinium species, however, considering that the NICS value for protonated pyridine is still close to the value observed for benzene, it is clear that the pyridine acceptor moiety demonstrates a considerable amount of aromatic character.

In the case of the quinoline acceptor component, when attempting to use NICS values to describe the aromaticity within this ring system, the NICS model indicates that quinoline is more aromatic than benzene. NICS values can be derived for each ring component of the quinoline acceptor. The A-ring is observed to have an NICS(1) value of -10.83 where the B-ring, bearing the ring nitrogen atom, demonstrates an NICS(1) value of -10.52.43 These are demonstrably more negative than the -10.20 observed for benzene (Table 2.8).43 What this discrepancy illustrates is simply that there are multiple factors that contribute the aromaticity of a compound. Aromaticity can not be defined by one simple model that is used to quantify this characteristic. Aromaticity is described by the sum of a number of different factors that contribute to the observed aromaticity of a compound. Factors that aid in quantifying aromaticity include the geometrical contribution described by HOMA, NICS values, chemical reactivity of a compound, as well as the magnetic susceptibility of a compound.43 These parameters all play a role in describing the entire concept of how aromatic a molecule is.43 Although the HOMA index does a good job of describing the observations made in these studies, it is not the only model that contributes or that can be used to describe the observed aromaticity within the acceptor components of the dyes presented here.43

Table 2.8: NICS(1) Values for a Number of Reference Compounds

Compound NICS(1) Ref Benzene -10.20 43 Pyridinium -9.89 43 Cation Pyridine -10.17 43 Quinoline A ring: -10.83 43 B-ring: -10.52

It is no mistake that there is no HOMA reference compound or NICS values to describe the indolinine acceptor within the data shown above. The PhOH Ind merocyanine dye

- demonstrates the lowest pKa value at 7.25 and the largest σ value at 1.25. This acceptor is decidedly the most electron withdrawing within this series based on the aforementioned parameters. This can be explained by the fact that the indolinine acceptor is in fact nonaromatic.

Although a variety of indole derivatives are aromatic, this indolinine acceptor, because of the presence of the sp3 hybridized carbon bearing the 2 methyl groups within the heterocyclic component of the acceptor, this acceptor moiety is not aromatic as it does not demonstrate delocalized pi system. Without this thermodynamic threshold to overcome upon deprotonation under basic conditions, it is clear why PhOH Ind demonstrates the lowest pKa value. This pKa can also be likened to that of p-nitrophenol. The indolinine is almost as equally electron withdrawing, based on its pKa. These cationic nitrogen groups with PhOH in and that of p-nitrophenol are not involved in aromaticity at all and simply act as electron withdrawing substituents through resonance.

Although all of the acceptors discussed here undergo alkylation to achieve the necessary activated methylene compound, as discussed previously, it is known that the nitrogen lone pairs of the pyridine and quinoline examples do not participate in aromaticity. Alkylating these

67 acceptors has little bearing on the aromatic character of the ring system. In fact, the HOMA value for pyridine in its protonated form, the pyridinium cation, is calculated to be 0.98.46 This is virtually the same as what is observed for pyridine in its neutral form. Additionally, although alkylating the ring nitrogen of the benzothiazole example does eliminate this lone pair, it is hypothesized that a lone pair found on the ring sulfur atom can also play a role in aromaticity in this case, filling the void created by generating the cationic nitrogen species.

2.3.7. Fluorescent Characteristics as a function of pH The change in absorbance characteristics of the dyes as a function of pH led to speculation that the fluorescence properties likely change in a similarly dramatic manner. This is the case as shown in Figure 2.16. Each of the merocyanine dyes were subject to solutions of increasing pH and the fluorescence emission spectra obtained. The excitation wavelength used in each case corresponds to the isosbestic point observed in the absorbance spectra for each individual molecule. The excitation and emission wavelengths as well as intensity data for each merocyanine are summarized in Table 2.8.

It is observed that the pyridine and quinoline merocyanines experience fluorescence quenching as the pH of the solution increases. The Irel under acidic conditions when the dyes are completely protonated compared to when the dyes are deprotonated under basic conditions is observed to be -6.3 and -17.6 for the pyridine and quinoline derivatives respectively. In contrast, the benzothiazole and indolinine examples demonstrate positive Irel values at 1.4 and 2.3, experiencing an enhancement in fluorescence emission under basic conditions. The Irel values are not the only parameters that describe the enhancement or deficits in fluorescence emission of the merocyanine dyes. The calculated fluorescence quantum yield values for each derivative, under acidic and basic conditions also demonstrate similar findings (Table 2.8).

The fluorescence quantum yield of a fluorophore describes the ratio of photons absorbed to photons emitted by the molecule.33 Often quantum yield is calculated using a comparitive approach, through the use of standard fluorophores that have a known quantum yield. The closer the quantum yield of a chromophore is to unity, the more efficient the translation of the light energy input in to the system into an emitted light photon from the fluorophore.

Although the quantum yields of the merocyanine dyes do not indicate an efficient fluorescence emission, there is still a clear trend that follows what is observed from the Irel values.

For the pyridine and quinoline derivatives, the fluorescence quantum yield when going from acidic to basic media demonstrates a decrease. This is not surprising given the quenching observed in the fluorescence emission spectra for these dyes. In the case of the benzothiazole and indolinine derivatives, the efficiency of their fluorescence emission increases as the dyes become deprotonated in basic conditions in comparison to acidic conditions. This is reflected in the quantum yields observed for PhOH Btz and PhOH Ind. After determining the acceptor ability of the various heterocycles by consulting the HOMA index, it is possible to rationalize why the pyridine and quinoline examples of these merocyanine dyes will experience fluorescence quenching in basic conditions, while the other examples demonstrate an enhancement in their fluorescence emission capability. It is hypothesized, in conjunction with the acceptor ability of the pyridine and quinoline derivatives, that under basic conditions, the fluorescence quenching observed can be explained by excited state electron transfer (eT).

300 180 B) A) 160 250 140 as pH 200 120 100 increases 150 80

100 60 Intensity(au) Intensity(au) 40

50 Fluoresence Fluoresence Emission Fluoresence Fluoresence Emission 20 0 0 400 450 500 550 600 650 450 550 650 750 Wavelength (nm) Wavelength (nm)

120 120 D) 100 C) 100 80 80 as pH 60 60 increases

40 40

Intensity (au) Intensity Intensity(au)

20 20

Fluorescence Fluorescence Emission Fluorescence Emission 0 0 450 500 550 600 650 700 475 525 575 625 675 Wavelength (nm) Wavelength (nm)

Figure 2.16: Fluorescence emission spectra for PhOH P (top left), PhOH Q (top right), PhOH Btz (bottom left) and PhOH Ind (bottom left) in solutions of increasing pH, excited at the wavelength corresponding to the isosbestic point observed in the UV data.

Table 2.9: Summary of the fluorescent data obserevd for the merocyanine dyes as a function of pH.

a b a b a b Compound λex λem λem Intensity Intensity Irel ϕ ϕ (nm) (nm) (nm) (au) (au) PhOH P 404 508 581 252.7 39.8 -6.3 0.0061 0.0013 PhOH Q 450 564 577 132.1 7.5 -17.6 0.0040 0.0003 PhOH Btz 439 512 559 69.0 96.1 1.4 0.0010 0.0025c PhOH Ind 460 531 554 44.1 101.4 2.3 0.0005 0.0044c aIndicates values obtained in acidic conditions bIndicates values obtained in basic conditions cQY calculated with Rh 101 as a fluorescence standard. All other calculations used QBS as a standard.

2.3.7.1. Excited State Electron Transfer of the Merocyanine Dyes As discussed in Section 2.2.7., the pyridine and quinoline acceptors are the least electron withdrawing of this series attributed to their high aromatic character. Because these acceptors are the most aromatic, the propensity for the phenolic alcohol to become deprotonated giving rise to a fully conjugated, neutral dye species is lowered as doing so would result in a loss of aromaticity within the heterocyclic moiety of the dye structure. It is hypothesized that, under basic conditions, instead of becoming completely deprotonated to give a fully conjugated form of the dye, the pyridine and quinoline dyes will alternatively undergo eT in the S1 excited state. The donor moiety of the dyes, the deprotonated phenol functional group, will transfer an electron to the acceptor component of the molecule. This generates a biradical species which is what allows for the twisted conformation of the fluorophore (Figure 2.17).30,31 From this TICT excited state, the merocyanine dyes will relax back to a TICT ground state through non-radiative pathways, primarily through the loss of rotational and thermal energy.31 Because the dyes preferentially undergo non-radiative relaxation under basic conditions, the quenching observed from the dyes when they are exposed to solutions of increasing pH, the ability for the dyes to adopt a TICT state provides an explanation for the observed loss of fluorescence emission capability for PhOH P and PhOH Q in basic media.

2.3.7.2. Explaining Fluorescence Enhancement of PhOH Btz and PhOH Ind in Basic Conditions The opposite trend is observed for the PhOH Btz and PhOH Ind dyes. Under acidic conditions, these dyes exhibit limited fluorescence emission in comparison to the quinoline and pyridine examples. The turn on fluorescence respsonse observed under basic conditions can, however, be explained by the acceptor ability of the heterocyclic component of the dye. As stated, the Btz and Ind acceptors are the most electron withdrawing, resulting in the lowest pKa values of the series.

Figure 2.17: Jablonski diagrams demonstrating ET process undergone by PhOH P in the excited sate to result in an excited TICT state of the molecule. Relaxation from the TICT state is achieved by non-radiative decay pathways. This figure uses PhOH P as an example. This figure is also meant to represent the quinoline merocyanine derivative.

There is no thermodynamic barrier to deprotonation in these examples as the benzothiazole and indolinine acceptor components are the least aromatic or exhibit no aromaticity at all. As a result, once PhOH Btz and PhOH Ind are deprotonated, they will undergo through resonance to give the fully conjugated, planar state of the dye in the ground state. This annihilates the cationic charge found on the ring nitrogen, giving rise to an arguably more stable molecule. Exciting this delocalized, planar system given the stability it has gained as a result of being deprotonated, the molecule will relax back to a ground state from an S1 state purely by a radiative pathway, by emission of a light photon (Figure 2.18).31

Figure 2.18: Jablonski diagram describing the emission of PhOH Btz and PhOH Ind in solutions of pH greater than 7. Here, PhOH Ind represents both dyes.

2.4. Conclusions The solvatochromic properties of the dyes were effectively characterized and compared to

Brooker’s merocyanine. It was observed that the drastic changes in the absorbance characteristics of the phenolic dyes is attributed to the H-bonding ability of the phenolic alcohol group and, ultimately, the ability for the dyes to undergo keto-enol tautomerization in solution. This was proven to be the case when comparing the phenol derivatives to the results obtained from the p- methoxy derivatives, which acts as an enol form of the dyes that are “frozen” in place. In all cases, the absorbance maximum wavelength of the dyes experiences a bathochromic shift as solvent polarity decreases. Additionally, fluorescence data corroborates the hypothesis that the keto form of the dye forms in solvents of decreasing polarity, particularly results obtained or the benzothiazole and indolinine examples. Not only do these dyes experience a large enhancement in fluorescence emission as a result of changes in solvent polarity, but the decrease in Stoke’s shift observed for these derivatives also indicates the formation of the dye’s keto tautomer in solution .

The pKa values were determined for all of the phenolic dyes demonstrating that the pyridine and quinoline derivatives are the least acidic with pKa values of 8.50 and 8.54 respectively. These

73 examples were followed next by the benzothiazole derivative with a calculated pKa of 7.76. The most acidic of these dyes was the indolinine example demonstrating a pKa value of 7.25. Through the use of the Hammett equation and these calculated pKa values, the electron withdrawing ability of each acceptor component was characterized. Through the use of the HOMA index and a variety of reference compounds, it was determined that the pyridine and quinoline acceptor components are the most aromatic of the acceptor moieties used here. As such, the willingness to break aromaticity is not thermodynamically favoured explaining why the pKa values for these derivatives are higher than the other two; the pyridine and quinoline examples are not as apt to become deprotonated to undergo through resonance to give the fully conjugated, neutral dye species.

Conversely, PhOH Ind has the lowest phenolic pKa which results from the fact that the indolinine acceptor, once activated, is not aromatic making deprotonation far more favourable.

Finally, the fluorescence characteristics of the dyes and how they change as a function of pH was also investigated. It was observed that the fluorescence emission ability of the pyridine and the quinoline derivatives drastically decreased as the pH of solution increased. The opposite was observed for the benzothiazole and indolinine merocyanine dyes, that as the pH of the solution increases, the fluorescence emission ability of these examples also increases. A suspected electron transfer in the excited sate for the PhOH P and PhOH Q derivatives under basic conditions leads to a TICT state of the dye. This explains the loss of fluorescence emission intensity as the TICT state of the molecule will relax via non-radiative pathways. This is linked to the aromaticity of the acceptor component of the dye as well. Because the pyridine and quinoline acceptors are less inclined to undergo through resonance upon deprotonation, the oxygen anion under basic conditions becomes a good donor, explaining electron transfer to result in fluorescence quenching.

PhOH Btz and PhOH Ind do not experience fluorescence quenching as a result of becoming deprotonated but instead exhibit a “turn on” fluorescence response to an increase in pH. Because the acceptor moieties of these dyes show limited aromaticity or are not aromatic at all, upon deprotonation, through resonance will result giving a more planar, fully delocalized dyes species.

This offers an explanation for the enhancement of fluorescence emission observed from the benzothiazole and indolinine dyes.

The overarching goal of this work will be to incorporate these phenolic merocyanine dyes into aptamers sequences that are selected to bind to protein targets. Because of their acid base capability and the vast colorimetric and fluorescent changes that are observed as a result of any acid base exchange the dye partakes in, it will be of interest to assess the applicability of these merocyanine dyes to signal the presence of peptidic target molecules. Amino acids, the building blocks of all proteins, bear ionizable side chains that can also readily undergo acid-base exchange processes. By strategically placing these phenolic dyes into aptamer sequences where the protein target is known to interact with the DNA strand, it is conceivable to believe that an exchange between the fluorophore and the amino acid side-chains of the protein could occur, eliciting a response from the dye to signal target binding. The studies presented here provide a good foundation of knowledge with regards to how the dyes behave in a variety of conditions. However, a long-term goal of this research will include the application of these dyes into aptamer based detection platforms.

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Chapter 3: Probing Structure Activity Relationships Contributing to GQ Binding Ability of Novel Merocyanine Dyes: Energy Transfer Fluorescence and CD Spectroscopy Studies

3.1. Introduction 3.1.1. Human Telomeric Repeat Sequence The human telomeric repeat sequence (Htelo) is a DNA sequence found on the 3’end of human chromosomal DNA.1 The repeating sequence (5’-TTAGGG-3’) represents a non-coding region of DNA that repeats itself on the order of 102 times at the end of human DNA.1 The Htelo repeat sequence plays a pivotal role in the overall functionality of the cell as it aids in protecting the genome from degradation, from end to end fusion of DNA as well as aiding in recognition by repair enzymes when DNA damage has occurred.2,3 As the cell undergoes multiple rounds of

DNA replication and mitosis, chromosomal DNA begins to shorten as this process continues for multiple cycles.4 The benefit that the Htelo sequence provides is that, as it is a non-coding region of DNA, should these repeats be shortened over multiple courses of DNA replication, no genetic information is lost as a result. However, as more of the Htelo sequence is lost over multiple rounds of DNA replication, there is a critical point where, once too much of the Htelo repeating DNA is lost, the cell will undergo senescence or programmed cell death, namely apoptosis.4 This process is hypothesized to cause the observed effects of aging in humans.4

There is an enzyme responsible for maintaining the length of telomeres in eukaryotic cells.5

Telomerase, a ribonuclease enzyme, can replicate and extend the Htelo sequence with the aid of an RNA primer.5 This enzyme is often upregulated and far more active in many different kinds of cancer cells.6 Interestingly, there is virtually no telomerase activity in healthy, somatic cells.6,7

This upregulated process of extending the Htelo repeat sequence at the end of chromosomal DNA is a factor in what can make cancer cells so potent.6 Telomerase activity often allows a cell to become “immortal.”6 By constantly replacing the lost repeats after rounds and rounds of DNA replication, there is virtually no point at which the cell will naturally enter the process of apoptosis.

In fact, the famous HeLa cell line exhibits enhanced telomerase activity, explaining the infallible

80 nature of these cells as they can thrive under varying conditions, being the first human cell line to be successfully cultured outside of the human body.8,9

Considering the G-rich nature of the Htelo repeat sequence, under physiological conditions it is not surprising that the sequence can be found in a GQ topology.10 This sequence demonstrates a high degree of polymorphism in the GQ structures it can adopt as well.11-14 This is often contingent on the cation present that will stabilize the G-tetrads of the GQ structure. The GQ structures observed in potassium are perhaps the most biologically important, considering that intracellular levels of K+ are higher than that of Na+.15 However, for the purposes of this work, the varied GQ topologies that are possible within the Htelo sequence as a result of varying metal ions is a key feature to understanding the novel dyes presented herein. In solutions containing sodium, Htelo will adopt a basket-type anti-parallel GQ topology (Figure 3.1).12 Interestingly,

Htelo can adopt a variety of GQ topologies in the presence of K+. Most notably, it is possible for

Htelo to adopt a parallel structure as elucidated through crystallographic data.13 Additionally,

Htelo can adopt a hybrid type of GQ structure in solutions of K+, demonstrating structural facets of both parallel and anti-parallel topologies (Figure 3.1).14

The fact that Htelo can adopt these GQ structures under physiological conditions plays a role in inhibiting telomerase activity.16 If the DNA is folded into this tertiary structure, the telomerase enzyme cannot effectively replicate the DNA because it is not in its linear conformation. This makes GQ structures interesting targets for anti-cancer drugs.17 If a given medication can bind to and stabilize the Htelo GQ, there would be no way for telomerase to continue replicating the telomere, allowing for an end of the cell’s immortality.17 In addition to creating drugs that would act as GQ stabilizing entities, the synthesis of GQ binding fluorophores is also a research area of high interest. Novel fluorophores designed to bind to a variety of GQ

81 structures would provide a means to visualize key biological processes relating to telomeres and telomerase, shining a light on the role these processes play in the proliferation and growth of a number of cancers.18,19 The work presented here attempts to introduce novel dye derivatives that selectively bind to GQ structures, possibly providing novel avenues to be used in the future of telomere research.

Figure 3.1: A) Antiparallel GQ topology of Htelo in the presence of Na+. B) Parallel GQ topology of Htelo in the presence of K+, crystallographic interpretation. C) Hybrid GQ topology of Htelo in the presence of K+.

3.1.2. Thrombin Binding Aptamer Thrombin is a serine protease that is involved in the blood cascade pathway within mammalian plasma.20 Thrombin enzymatically cleaves the soluble glycoprotein fibrinogen to fibrin and leads to clot formation upon injury to stop bleeding.20 An aptamer was selected against thrombin in the early 1990s.21 The thrombin binding aptamer (TBA) is a 15mer DNA oligo that demonstrates a G-rich sequence (Figure 3.2).21 Upon target binding, as proven by the crystal structure of the nucleic acid in complex with its protein target, TBA folds into a basket type, antiparallel GQ.21,22 As a very well-studied and characterized aptamer, the TBA example is often used as a proof-of-concept system when developing aptamer based detection platforms. For this reason, the Manderville laboratory carries out preliminary studies using novel fluorescent probes

82 within TBA prior to exploring less studied DNA aptamers. TBA was used initially in this work in hopes of developing a free dye detection platform employing a duplex to quadruplex exchange approach, making use of the novel fluorophores synthesized in house. Results of these studies indicated that the dyes presented here had more of a propensity to interact with GQ DNA. These findings will be discussed in detail in section 3.3.2.

A) B)

PDB: 4DIH

Thrombin Binding Aptamer (TBA) 5’- GGTTGGTGTGGTTGG-‘3

Figure 3.2: A) The crystal structure of TBA in complex with its protein target thrombin. B) A simplified rendering of the TBA G4. The G residues involved in this G4 are found in alternating syn- and anti-conformations. The G residues that make up the G4 are bolded within the aptamer sequence

3.1.3. GQ Binding Fluorophores At present there are a wide variety of GQ binding fluorophores that allow for visualization of this DNA tertiary structure based on a turn on fluorescence response. Mostly, these dyes are hypothesized to either end stack on the top or bottom tetrad of a GQ or intercalate between G- tetrads.23,24 A few examples of such fluorophores are shown in Figure 3.3.

Figure 3.3: Chemical structures of a number of GQ binding fluorophores.

A number of these fluorophores have been used in a variety of applications. For instance,

NMM has been utilized to determine the ability for GQ structures to fold within duplex DNA structures.25 CV has been used to explore the ability for extrinsic fluorophores to undergo energy transfer fluorescence based on interactions the fluorophore has with GQ DNA.24 The GQ binding fluorophore of most interest to this work is that of ThT. This small molecule, which is essentially non-fluorescent in a purely aqueous environment, will undergo a 210-fold increase in fluorescence emission in the presence of the 22-mer Htelo sequence.19 In addition, this dye demonstrates the ability to induce specific GQ topologies in the presence of Htelo DNA.19 The dyes assessed in this work are structurally very similar to ThT. The only difference in their structures being the presence of a bridging olefin group in the novel dyes discussed herein. Based on these similarities, 84 it was of utmost interest to compare these dyes to ThT in order to have a full grasp on why these dyes behave as they do in the presence of Htelo DNA (Figure 3.4)

Figure 3.4: Structural comparison of ThT to one of the novel merocyanine dyes presented in this work.

3.1.4. Energy Transfer: From DNA into Fluorophores The ability for energy transfer to occur between Htelo DNA and fluorescently modified internal base mimics has been explored and noted in the literature.26,27 In fact, past research in the

Manderville laboratory utilizing a furan modified dG base within the 22mer Htelo DNA sequence demonstrated that an energy transfer band becomes apparent around 290 nm in the excitation spectrum once the modified DNA strand is properly folded into a GQ tertiary structure.26 This band centered at 290 nm indicates that there is a transfer of energy occurring from the GQ structure of the DNA into the fluorescent base mimic.26 It makes sense that a fluorophore that is covalently linked to the GQ structure would be able to undergo energy transfer with the DNA it is bound to.

However, it is also possible for extrinsic fluorophores to under go energy transfer processes with

GQ DNA as well. This has been noted for the GQ binding chromophore, CV.24 CV demonstrates the ability to undergo energy transfer emission once bound to GQ DNA in solution upon excitation at 280 nm.24 This facet of the fluorophores characteristics has been utilized to differentiate GQ structures in solution from duplex DNA and from single stranded DNA as well.24 It was of interest with these studies to determine whether or not the merocyanine dyes presented in this work were capable of undergoing energy transfer fluorescence in the presence of GQ DNA and whether or

85 not the energy transfer emission could be of use in determining which dyes preferentially interact with what GQ topologies of Htelo DNA.

3.2. Materials and Methods Htelo, (5’-AGGGTTAGGGTTAGGGTTAGGG-3’), TBA (5’- GGTTGGTGTGGTTGG-

3’), mismatched TBA complementary strand (C2) (5’-CCAACCACAACAACC-3’) were purchased from Sigma-Aldrich Ltd and purified by reverse phase cartridge purification. Tris base buffer, NaCl, KCl, and K2HPO4 were purchased from Fisher Scientific. Ultra Pure Glycerol was purchased from Invitrogen. Thrombin protein was purchased from BioPharm Laboratories LLC.

Three Tris buffer solutions were prepared with final salt concentrations of : (a) 50 mM Tris base (b) 50 mM Tris base with 50 mM NaCl and, (c) 50 mM Tris base with 50 mM KCl. All buffer solutions were prepared in Milli Q (18 MΩ) water and adjusted to a pH of 7.2.

3.2.1. General Synthesis of the Merocyanine Dyes The merocyanine dyes utilized for these studies were synthesized and fully characterized by Andrew Chung, a fellow MSc student in the Manderville research laboratory and used as confirmed. The synthetic approach utilized is similar to the pathway explained in Chapter 2. The structure of the dyes are shown in Scheme 3.1.

Scheme 3.1: Structure of the Merocyanine Dyes

3.2.2. Preparation of Stock Solutions and UV-Vis and Fluorescence Experiments Stock solutions of each dye were made by dissolving the solid compound in DMSO to stock concentrations of 4 mM. UV measurements were made in 1000 µL 100-QS quartz cells and observed from 800 nm to 200 nm. Fluorescence measurements were made in 1000 µL 101-QS quartz fluorescence cells using excitation and emission slit widths of 5 nm. Emission spectra were obtained by exciting the sample at either the excitation maximum wavelength for the dye or at a suspected energy transfer band centered at 256 nm and monitoring the spectrum from this wavelength to 800 nm. Excitation spectra were obtained using the emission maximum observed in the emission spectrum and monitored from 200 nm to the emission maximum wavelength. DNA oligonucleotides were resuspended in Milli-Q water and quantified using their respective extinction coefficients. Thrombin protein was resuspended in Milli-Q water and quantified using

ε = 72 150 M-1 at 280 nm.

3.2.3. Glycerol Studies Baseline fluorescence emission for the merocyanine dyes were determined in Milli-Q water at a dye concentration of 5 µM. The response of the dyes to an increase in rigidity of the solution was determined by increasing the glycerol content in solution to 80% and observing the

87 fluorescence emission. Irwl values were determined by taking the ratio between the fluorescence intensity of the dye in 80% glycerol to the intensity of dye in 100% water.

3.2.4. TBA:C2 Fluorescence Titrations Baseline fluorescence emission of the merocyanine dyes was determined in potassium phosphate buffer at a dye concentration of 1 µM. TBA:C2 duplex DNA, previously heated to 90

°C and slow cooled to room temperature overnight to anneal the DNA strands, was added to the dye in solution to a concentration of 0.5 µM and the fluorescence emission of the dyes was determined once more. Thrombin protein was then added to the DNA, dye sample to a concentration of 1 µM and the fluorescence emission of the dye determined once more.

3.2.5. CD Studies CD analyses were performed on a Jasco J-815 CD spectrophotometer equipped with a thermal controlled 1 x 6 multicell block. Solutions of 3 µM Htelo with varying concentrations of dye (0, 1, 3 and 5 µM) were prepared in each Tris buffer solution. Spectra were obtained at 20 oC in quarts cells (110-QS) with a path length of 1 mm and monitored between 200 and 600 nm at a bandwidth of 1 nm and scanning speed of 100 nm/min. A minimum of four scans were collected and combined. Data was corrected against a blank measurement of the varying Tris buffers. The spectra were smoothed using the Jasco Spectra Analyzer software by Savitzky-Golay function with

15 convolutions.

3.2.6. UV-Vis and Fluorescence Titrations of the Merocyanine Dyes with Htelo The baseline absorbance and fluorescence of the merocyanine dyes at a concentration of 3

µM in solution were obtained prior to the addition of Htelo DNA. Increasing concentrations (0,

0.12, 0.25, 0.75, 1.5, 2.5, 4 and 5µM) of DNA were added to the dyes in solution. Absorbance and fluorescence emission and excitation spectra were obtained following each addition of DNA.

Irwl values were determined by taking the ratio between the fluorescence intensity of the dye in the presence of the highest concentration of DNA and the intensity of the dye in the absence of DNA.

3.2.7. Determination of the Binding Affinity of the Merocyanine Dyes for Htelo DNA – Kd Determination Dissociation constants (Kd) were determined by using the fluorescence emission observed from the fluorophores when excited at the ET excitation wavelength of 256 nm. A plot of the fraction of dye bound against the concentration of Htelo DNA, using a single site ligand binding model in Sigma Plot 12.0, generated a binding isotherm allowing for the elucidation of the Kd values from the equation:

퐵 푥 푦 = 푚푎푥 퐾푑 + 푥

Where y represents the fraction of dye bound, x, the concentration of Htelo DNA and Bmax the maximum fraction of dye able to bind the DNA in solution.

3.3. Results and Discussion 3.3.1. Characterization of the Rotor Merocyanine Dyes As discussed in Chapter 1, molecular rotor probes will demonstrate an enhancement in their fluorescence emission ability when they are effectively locked into a planar, or LE, ground state.28 It was of interest with this work to determine whether or not these novel merocyanine dyes are capable of achieving LE emission upon binding to GQ DNA. However, in order to classify these dyes as molecular rotors in the first place, a more traditional approach was taken. Molecular rotor probes will respond to an increase in rigidity of their microenvironment.29,30 Increasing the viscosity of the solvent the dye resides in can also “lock” the fluorophore into a more planar conformation, allowing for an increase in fluorescence emission.30 An increase in the rigidity of the solution can be achieved with the addition of glycerol. As shown in Figure 3.2, all of the

89 merocyanine dyes demonstrate an increase in fluorescence emission when comparing their performance in solutions of 0% versus 80% glycerol. This is illustrated by the Irel values determined for each dye as the glycerol content of their surrounding environment increases (Table

3.1).

Although the phenyl quinoline derivative exhibits the largest fluorescence enhancement in glycerol, boasting an Irel value of 16.5, this dye is the least fluorescent of all of the dyes presented, only increasing in emission intensity from 2.1 to 34.4. The remaining dye derivatives all demonstrate good improvement in fluorescent emission with the increased percentage of glycerol.

What is important to note is that, although Th Btz does not demonstrate the largest increase in fluorescence emission with an Irel value of 6.7, it is definitively the most fluorescent of this series demonstrating fluorescence emission intensity of 500.2 in the more rigid environment that 80% glycerol provides.

These glycerol studies certainly allude to the idea that the novel merocyanine dyes are capable of behaving like molecular rotor probes. By forcing the dyes into their planar conformation, by increasing the rigidity of their external environment, the observed enhancement of fluorescence emission provides evidence that the dyes will behave appropriately when they are exposed to duplex as well as GQ DNA. The idea here is that when the dyes intercalate between the base pairs of the DNA double helix, or when the dye stacks with GQ DNA or intercalates between G-tetrads, the dyes will once again adopt their planar conformation, allowing for an enhancement in fluorescence emission.31

Table 3.1: Summary of Molecular Rotor Characterization Data

Acceptor Donor λmax λem Emission Emission Irel (nm) (nm) Intensity Intensity 0% 80% Glycerol Glycerol (a.u.) (a.u.) N-Methyl Phenyl OH 576 685 2.1 34.4 16.5 Quinoline N-Methyl Phenyl OH 457 607 34.2 460.9 13.5 Pyridine N-Methyl Thiophene 551 588 74.8 500.2 6.7 Benzothiazole OH Phenyl OH 505 593 19.9 210.8 10.6

500 A) 40 B) 35 400 30 300 25 20

200 15 Intensity(au) Intensity(au) 10 100

Fluorescence Fluorescence Emission Fluorescence Emission 0 0 480 580 680 780 575 675 775 Wavelength (nm) Wavelength (nm)

250 600 C) D) 200 500 400 150 300 100

200

Intensity(au) Intensity(au)

50 100

Fluorescence Fluorescence Emission Fluorescence Emission 0 0 500 600 700 800 550 600 650 700 750 Wavelength (nm) Wavelength (nm)

Figure 3.5: Glycerol studies of the merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBTz at a concentration of 5 µM dye in solution, increasing glycerol content in solution from 0 to 80%.

3.3.2. Duplex to Quadruplex Exchange: The Thrombin Example The previous molecular rotor studies inspired the idea that these dyes could be used to create a free dye, aptamer based detection platform. The design was to create an “on-off” fluorescence signal, utilizing the thrombin example and a duplex to GQ exchange approach. A similar approach has been explored with the use of CV as the reporter molecule.32 Briefly, TBA and a complementary strand, annealed to form a duplex, would be added to a solution containing the molecular rotor dyes.33 The TBA complementary strand was designed to bear a base mismatch in its sequence to encourage disassembly of the duplex in the presence of thrombin. Upon intercalation of the dye with the duplex DNA and exciting the fluorophore at its maximum absorbance wavelength, the fluorescence emission of the dye would, ideally, increase or “turn on.”

Addition of thrombin protein to the mix would see the duplex DNA pulled apart as TBA would preferentially bind to thrombin, forming a GQ structure in the process. As the dye is liberated from the rungs of the DNA double-helical ladder, the fluorescence emission of the dye should decrease as a result because the dye is no longer being forced into it’s planar, LE ground state.

25 30

20 A) 25 B) 20 15 15

10 Emission Emission

Emission Emission 10

Intensity(au) Intensity(au) 5 5 0 0 500 600 700 800 600 650 700 750 800 Wavelength (nm) Wavelength (nm)

140 250 120 200 D) 100 C) 80 150

60 100

Emission Emission Emission Emission Intensity(au) Intensity(au) 40 50 20 0 0 500 600 700 800 560 610 660 Wavelength (nm) Wavelength (nm)

Figure 3.6: Duplex to quadruplex exchange assay of 1 µM (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Orange traces represent the fluorescence emission of the free dye, blue traces represent the fluorescence intensity of the dye with the addition of 0.5 µM TBA:C2 duplex DNA and purple traces represent the emission of the dye in the presence of TBA:C2 duplex after the addition of 1 µM thrombin protein.

Although a well conceived idea, these experiments did not provide the hypothesized result.

An interesting observation that arose from these experiments, however, was when the pyridine quinoline merocyanine derivative was utilized for the assay. Instead of fluorescence quenching that was expected with the addition of thrombin to the assay, the fluorescence emission intensity of the dye increased with GQ formation. As shown in Figure 3.6 D, when PhQ is free in solution, there is virtually no fluorescence associated with the dye. With the addition of duplex DNA, however, there is a 3-fold enhancement in fluorescence emission. Upon addition of thrombin, which allowed for GQ formation of the TBA strand, there is a nearly 4-fold increase in

93 fluorescence emission intensity in comparison to the free dye. This same trend is observed for the

PhP and PhBtz, dyes as well, although to a much lower degree. With regards to the ThBtz dye, although there is no real enhancement in fluorescence emission, there is a 5 nm red-shift in λem with the addition of duplex DNA to the dye in solution. This new emission wavelength remained consistent with the addition of thrombin to the system.

These results certainly did not encourage further investigation into the utility of the dyes in a duplex to quadruplex exchange assay. However, the enhancement in fluorescence emission observed for a number of the dyes in this series as a result of GQ formation facilitated by addition of thrombin protein inspired an alternative study. The TBA assay provided inspiration to assess instead the ability of the novel merocyanine dyes to bind to GQ DNA. It was of interest to determine how the structure of the dye enhanced or eliminated the ability for the fluorophore to interact with a GQ. In addition, it was of interest to determine whether or not the dyes would preferentially interact with a certain topology of GQ over another. The use of Htelo DNA based on its ability to adopt various GQ topologies, contingent upon the cation present in solution made this DNA sequence a prime candidate to characterize the GQ binding ability of the merocyanine dyes.12-14 The structure activity relationships of the dyes and their ability to bind to GQ DNA was investigated by studying a variety of merocyanine dyes, varying the donor and acceptor components within the structure to elucidate their role in the dyes’ potential to favourably interact with quadruplex DNA.

3.3.3. CD Studies CD spectroscopy was used in these studies to determine if the merocyanine dyes would preferentially bind to a certain topology of GQ over another. Because of the distinct differences in the CD spectra of anti-parallel, parallel and hybrid GQ structures, it was within the best interest

94 of this work to monitor any changes in the CD spectra of Htelo DNA as increasing amounts of the merocyanine dyes are added to the DNA in solution. In addition to determining the GQ topology that allowed for the best dye binding, another goal of these studies was to determine whether or not the novel dyes could induce the formation of a certain GQ topology over another. These studies were done in Tris buffer with either no cation, Na+ or K+ present to facilitate folding of the appropriate Htelo GQ.

50 mM Tris – No Cation

As can be seen in the CD spectra below in Figure 3.7, when Htelo is found alone in solution with no counter ion to stabilize the formation of a GQ, the amplitude of the peaks that are detected by the CD is very low. It is possible to argue that the DNA strand is tending toward the formation of a parallel GQ based on the small negative signals centered around 240 nm in conjunction with the small positive bands around 260 nm. However, the lack of signal intensity still suggests that there is a limited amount of CD active geometries being formed in this solution.

There is a distinct difference in the Btz dye derivatives’ ability to interact with Htelo in the absence of a cation in comparison to the P and Q dye examples. PhBtz and ThBtz when they are added to the DNA in solution do not invoke any large or pronounced changes in the overall topology of Htelo. In the case of PhQ, however, there are notable differences in the CD spectrum with its addition to the solution. As more and more PhQ is added to Htelo, the negative CD band around 240 nm begins to sharpen and its amplitude increases as well. The largest change observed is the distinct emergence of a positive peak centered at 260 nm with the addition of PhQ. Although some positive CD bands around 250 nm were observed prior to the addition of increasing amounts of the chromophore, the fact that the peak maximum red-shifts toward 260 nm coupled with the overall increase in signal strength indicates that the quinoline dye derivative of this series

95 preferentially binds to a parallel GQ topology. In addition, these observations provide evidence of the dye’s ability to induce a conformational change within the DNA strand. PhQ will force the

DNA into a parallel GQ as this is the dye’s preferred topology to interact with (Figure 3.7 B)

There is light evidence that suggests, as well, the ability for PhP to force the parallel GQ topology (Figure 3.7 A). When there is no dye added to the Htelo DNA, there is very no clear evidence of a preferred GQ topology. The general trend observed, as more and more PhP is added to the DNA in solution, the signature CD bands at 240 nm and 260 nm of a parallel GQ structure become more evident. Although the changes in signal intensity are not nearly as pronounced as what was observed in the case of the PhQ chromophore, it is clear that the PhP dye still has some bearing on the GQ topology Htelo will adopt

What is it, then, that allows the pyridine and quinoline dye derivatives of this series to preferentially interact with and even induce the parallel GQ topology? It is hypothesized that there is more of an ability for the pyridine and quinoline dye examples to effectively pi-stack with the

GQ. In the instance of the parallel GQ structure, the loop regions associated with this topology are much smaller in comparison to those observed in the anti-parallel GQ conformation. The loop regions of parallel GQ structures, to this end, are incapable of cresting the top tetrad of the structure in the same manner that the loops of an anti-parallel GQ can (Figure 3.8). Because of this, it is possible that the pyridine and quinoline dyes are able to readily interact with the more accessible pi-stacking interface of the parallel GQ. With particular reference to the PhQ derivative, considering the size of the dye and its structural geometry it is not surprising that under these conditions, a parallel GQ is preferred. Based on literature examples of parallel quadruplex preferring chromophores such as NMM in certain circumstances as well as a macromolecular

96 molecule, BPBC, considering the bulky nature of the quinoline acceptor component, it makes sense that a parallel topology would be preferred.25,34

As discussed in Chapter 2, the pyridine and quinoline acceptor components are more aromatic than the benzothiazole framework. This characteristic of the acceptor directly relates to its electron withdrawing ability. In this series of dyes, as well, the tertiary amine group found on the donor component of the dye can readily delocalize its lone pair through the conjugated system of the dyes’ structure. In the case of the Btz dyes, this iminium form would be the more favoured resonance form for the dye to adopt (Figure 3.9). Because the acceptor component of the dye is so electron withdrawing, the tertiary amine of the donor would preferentially donate its lone pair to participate in through resonance. In the case of PhQ, the quinoline acceptor is not as electron withdrawing as the benzothiazole. This is attributed to the aromaticity of the quinoline ring; quinoline would rather maintain aromaticity instead of undergoing through resonance (Figure 3.9).

As a result, the dye maintaining full aromatic character will more readily be able to interact with the parallel GQ via aromatic, pi-stacking interactions as opposed to the dye derivative that loses any aromaticity associated with it’s structure as it undergoes resonance.

2.5 A) 2 1.5 0 uM Dye 1 1 uM Dye 0.5 3 uM Dye CD (mdeg) CD 0 -0.5 200 220 240 260 280 300 320 340 5 uM Dye -1 Wavelength (nm)

2 B) 1.5 1 0 uM Dye 0.5 1 uM Dye 3 uM Dye CD (mdeg) CD 0 200 220 240 260 280 300 320 340 -0.5 5 uM Dye -1 Wavelength (nm)

1.5 C)

1 0 uM Dye 0.5 1 uM Dye 0 3 uM Dye CD (mdeg) CD 200 220 240 260 280 300 320 340 -0.5 5 uM Dye

-1 Wavelength (nm)

2 D) 1.5 1 0 uM Dye 0.5 1 uM Dye 3 uM Dye CD (mdeg) CD 0 200 220 240 260 280 300 320 340 -0.5 5 uM Dye -1 Wavelength (nm)

Figure 3.7: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the absence of metal cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration.

A) B)

Figure 3.8: Comparing antiparallel and parallel GQ structures. The antiparallel-basket topology (A) demonstrates a barrier to pi-stacking as the loop region crests the top of the GQ. The parallel topology (B) boasts smaller loop regions, none of which interfere with the planar faces of the GQ making them readily amenable to pi-stacking interactions.

Figure 3.9: Demonstrating the preferred resonance forms of PhBtz and PhQ respectively. The Btz derivative prefers the iminium form due to the electron withdrawing ability of the benzothiazole acceptor. The quinoline example prefers the resonance form that maintains aromaticity within the acceptor moiety, making the Q derivative a better candidate to under go pi-stacking with G-tetrads of the parallel GQ. Larger arrow head signifies the preferred resonance structure.

50 mM Tris with 50 mM Na+

As mentioned in section 3.1.1., in the presence of sodium cations, Htelo DNA will adopt an anti-parallel GQ topology.12 This is observed in the CD spectra of Htelo without the addition

99 of any dye in all cases. The anti-parallel topology is characterized by positive CD bands at 290 nm and 240 nm as well as a negative CD signal centered at 260 nm. These characteristic CD signals are observed in all cases, confirming the presence of anti-parallel GQ structures in solution.

As increasing concentrations of PhP are added to the Htelo DNA, there are no obvious changes in the CD spectra to speak of. This indicates an inability for the pyridine dye derivative to affect this GQ topology (Figure 3.10 A). In contrast, the PhQ derivative does demonstrate some ability to interact with the GQ in this conformation. As increasing amounts of the quinoline dye derivative is added to Htelo, the amplitude of the negative signal decreases and the wavelength corresponding to the peak maximum shifts toward the blue (Figure 3.10 B). In addition, although a subtle difference, the peak centered at 290 nm begins to widen slightly with the presence of PhQ.

The changes observed in the CD spectra of Htelo with increasing concentrations of PhQ present could indicate that the quinoline dye is shifting the anti-parallel GQ topology toward favouring a parallel conformation. Although it is clear that the presence of PhQ has only minute effects on the overall structure of the Htelo GQ in solution, it is clear that the subtle changes observed in the CD spectrum indicate that the anti-parallel topology is not ideal for the quinoline derivative to bind to the DNA. The changes in the CD signal of Htelo are then a direct result of

PhQ forcing the equilibrium more toward a parallel GQ structure, allowing for a more favourable interaction between the dye and the DNA.

In the case of both benzothiazole examples of the dyes, there is clear evidence that PhBtz and ThBtz favourably interact with the anti-parallel topology of the Htelo GQ provided by the presence of Na+ in solution (Figure 3.10 C and 3.10 D). This is particularly the case when discussing the thiophene-benzothiazole example. When looking at the CD spectra for Htelo as

100 increasing concentrations of ThBtz are added, it is clear that the intensity of the negative CD signal increases as well. This not only alludes to a direct interaction between ThBtz and the anti-parallel

GQ but this observation also provides evidence that this chromophore is effectively “locking” the parallel topology into place, based on the changes in the CD signal. A similar observation can also be made for the PhBtz although not to the same degree as has been seen as ThBtz is added to Htelo in solution. Although it is more difficult to hypothesize the binding mode that the benzothiazole derivatives have with the GQ DNA, the CD data could be used to aid in determining how the dye interacts with Htelo going forward.

Probing the CD signal past 400 nm, and looking for an induced CD (ICD) band roughly in the region of the absorbance maximum wavelength of the dye would indicate whether or not the dyes intercalate or act as groove binders with the GQ. A positive ICD band corresponds to a groove binding mode of the dye where a negative ICD indicates intercalation of the dye between

G-tetrads of the tertiary structure.35 Although there is no clear band in the CD spectrum, in all cases for all dyes presented, the CD signal trends downward toward the negative, indicating a possible intercalation or stacking mode of binding (data not shown). It would be necessary, in the future and in future experiments to fully explore the presence of ICD bands and to utilize this data to fully characterize how the dye’s bind to GQ DNA. However, in the presence of a sodium cation to stabilize the anti-parallel GQ structure of Htelo DNA, it is clear that the Btz examples of the dyes still exhibit some ability to interact with this GQ topology where the pyridine and quinoline examples do not.

101

1.5 A) 1 0.5 0 uM Dye 0 1 uM Dye 200 220 240 260 280 300 320 340 3 uM Dye CD (mdeg) CD -0.5 -1 5 uM Dye -1.5 Wavelength (nm)

2 B) 1.5 1 0 uM Dye 0.5 1 uM Dye 0 3 uM Dye CD (mdeg) CD -0.5 200 220 240 260 280 300 320 340 -1 5 uM Dye -1.5 Wavelength (nm)

2.5 C)

1.5 0 uM Dye 0.5 1 uM Dye

3 uM Dye CD (mdeg) CD -0.5 200 220 240 260 280 300 320 340 5 uM Dye

-1.5 Wavelength (nm)

2.5 D) 2 1.5 0 uM Dye 1 1 uM Dye 0.5 3 uM Dye CD (mdeg) CD 0 -0.5 200 220 240 260 280 300 320 340 5 uM Dye -1 Wavelength (nm)

Figure 3.10: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the presence of 50 mM Na+ cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration.

102

50 mM Tris with 50 mM K+

When Htelo DNA is found in solution with potassium cations, the DNA will adopt a hybrid

GQ structure.14 As shown in section 3.1.3., this GQ topology bears structural characteristics that belong to both parallel and anti-parallel conformations of this tertiary structure. Hybrid GQ structures are characterized in their CD spectrum by a broad peak centered around 290 nm. There is also a negative peak observed at 240 nm for this GQ conformation. As shown in the CD spectra in Figure 3.11, it is evident that Htelo adopts this mixed GQ structure in solutions containing K+.

In the case of PhP, the negative signal centred around 230 nm begins to shift closer toward

240 nm. This negative signal also decreases in amplitude with the addition of more PhP to the solution. Although there are no real differences observed in the large peak centered around 290 nm, the shifting of the negative CD signal toward 240 nm, indicates that the pyridine dye derivative is, once again, interacting with this GQ topology and effectively securing the hybrid GQ structure in solution (Figure 3.11 A).

As shown in Figure 3.11 B), a large amount of fine structure associated with the positive

CD band centered around 290 nm disappears with increasing concentrations of PhQ. This loss of fine structure indicates that the dye is forcing the DNA to adopt a more defined structure, in comparison to the hybrid GQ topology. The pyridine and quinoline dye derivatives seem to favourably interact with a well defined hybrid structure and can themselves cause a more rigid conformation to predominate in solution based on the aforementioned observations. This is not the case for the benzothiazole dye derivatives.

When PhBtz is added to the Htelo in the presence of potassium, there are notable changes to the negative CD band centred at 240 nm (Figure 3.11 C). As increasing concentrations of this

103 dye are added to the DNA, the negative signal at 240 nm completely disappears where there is the emergence of a slight valley beginning to form in the CD signal around 260 nm. Also, with the addition of ThBtz to the Htelo sequence, some fine structure is lost in the spectrum around 250 nm

(Figure 3.11 D). The signal at 240 nm also broadens to give a large, slightly negative signal from approximately 235 nm all the way 245 nm. What these results suggest is that the dyes boasting the benzothiazole acceptor moiety are able to induce a switch in the GQ topology from the hybrid structure that exists in potassium to an anti-parallel conformation. The CD signatures of an anti- parallel GQ are positive CD bands at 290 nm and 240 nm with a negative band at 260 nm. The broadening of the negative signal with the addition of ThBtz to the DNA coupled with the emergence of a slight negative trend in the CD spectrum of Htelo in the presence of PhBtz around

260 nm indicates that the conformational equilibrium of the DNA is being forced toward forming an anti-parallel GQ. This has been noted in the literature with a known GQ binding fluorophore,

ThT.19 This dye, similar to the dyes presented here, possesses the same benzothiazole moiety. In solutions containing K+, ThT has demonstrated the ability to force an anti-parallel GQ topology within the Htelo sequence. It is clear that these dyes behave in a similar fashion and that the benzothiazole component of the dyes has a role in inducing anti-parallel GQ structures.

As was shown by Mohanty et al.19 using MD simulations to determine potential binding modes of ThT to Htelo quadruplex structures, it was determined that binding of the chromophore to anti-parallel structures in an end stacked fashion gave a far more stable complex than when simulations were derived for ThT end stacked with a parallel GQ structure. The stability of the anti-parallel GQ:ThT complex is a direct result of aromatic interactions between the the fluorophore and the G-tetrad of the DNA. In addition, electrostatic interactions between the cationic nitrogen of the dye’s structure and the DNA contribute to the stability of the complex.19

104

These calculations also indicate that the parallel GQ topology is destabilized by end stacking of

ThT as a result of a disruption in the ability of the guanine residues within the G-tetrad to coordinate the central metal ion.19 Based on these data using ThT as a dye analogue to aid in describing the behaviour of the dyes presented in this study, it is hypothesized that the Btz derivatives behave in a similar fashion as ThT, providing the same stabilization to anti-parallel GQ structure while destabilizing anti-parallel structures in the same manner. Because the novel merocyanine dyes bear the same benzothiazole moiety as ThT with the very same cationic N- methyl component, it is no surprise that these dyes, like ThT can induce anti-parallel GQ formation. More details on this observation will be discussed in section 3.3.6.

105

2 A) 1.5 1 0 uM Dye 0.5 1 uM Dye 3 uM Dye CD (mdeg) CD 0 200 220 240 260 280 300 320 340 -0.5 5 uM Dye -1 Wavelength (nm)

2 1.5 B) 1 0 uM Dye 0.5 1 uM Dye 3 uM Dye CD (mdeg) CD 0 200 220 240 260 280 300 320 340 -0.5 5 uM Dye -1 Wavelength (nm)

3 C)

2 0 uM Dye

1 1 uM Dye

CD (mdeg) CD 3 uM Dye 0 200 220 240 260 280 300 320 340 5 uM Dye -1 Wavelength (nm)

2.5 D) 2 1.5 0 uM Dye 1 1 uM Dye 0.5 3 uM Dye CD (mdeg) CD 0 -0.5 200 220 240 260 280 300 320 340 5 uM Dye -1 Wavelength (nm)

Figure 3.11: CD spectra of 3 µM Htelo22 in 50 mM Tris buffer in the presence of 50 mM K+ cations, in the presence of 0, 1, 3 and 5 µM concentrations of (A) PhP (B) PhQ (C) PhBtz and (D) ThBtz. Increasing colour saturation corresponds to an increase in dye concentration.

106

3.3.4. UV-Vis and Fluorescence Titrations of Merocyanine Dyes with Htelo The CD studies provided an in-depth look at how the merocyanine chromophores affect the overall topology and structure of Htelo GQ DNA in solution. However, it was of utmost interest to determine what effects, if any, does the interaction with GQ DNA have on the dyes’ optical properties. Ultimately, it is any changes in the absorbance or fluorescence characteristics of the dyes that will provide a signal to indicate binding to the DNA going forward, applying these chromophores to the development of detection platforms. The changes in optical properties of the dyes were monitored by UV-Vis and fluorescence titration experiments in which increasing concentrations of Htelo DNA were added to the designated chromophore in Tris buffer in the absence of a metal cation as well as in the presence of sodium and potassium ions to determine the effects on the dye’s optical output as a function of GQ topology. Before going into the details on how the optical outputs of each dye varied as a function of GQ topology, a phenomenon was observed in the fluorescence excitation spectra for all of the dyes as increasing concentrations of

Htelo DNA were added to the solution. As more DNA was added to the dyes in solution, the excitation maximum wavelength shifted toward the red, particularly in the case when no metal ions were present. These observations are summarized in Table 3.2.

Table 3.2: Comparing excitation maximum wavelengths for the free merocyanine to merocyanine dyes bound to Htelo DNA.

a b Dye λex λex Δ λex (nm) (nm) (nm) PhP 457 500 43 PhQ 576 588 12 PhBtz 505 559 54 ThBtz 551 582 31 a λex of the free chromophore b λex of the chromophore in the presence of Htelo DNA, no metal cation

107

All of the dyes experience a red-shift in λex by at least 10 nm when they are exposed to

Htelo DNA. Additionally, there is a red-shift in absorbance maximum for all of the dyes as increasing concentrations of Htelo is added to the dye in solution. This is illustrated in the UV-

Vis titration data below. Although these observations were initially made when Htelo was added without the presence of a metal cation to stabilize GQ formation, the red-shifted excitation wavelengths were used in these studies because it is believed that this red-shift is a direct result of the dye interacting with the DNA, regardless of GQ topology. This red-shift in λex can be explained by the interaction of the dye with the more non-polar interface provided by DNA in comparison to the very polar environment of the aqueous buffered solution.36 The interaction between the dye and the DNA can conceivably encourage the dye to adopt a more planar conformation, particularly if the dye is stacking with the tetrads of any GQ structure in solution. It is in this more planar state that the dye can also become a more delocalized pi system, allowing for more red-shifted absorbance and excitation wavelengths.37 While these observations encourage the notion that the dyes are in fact stacking with GQ structures in solution, these changes in optical properties of the dyes can readily be applied to determining whether or not these merocyanines will preferentially interact with a certain GQ topology over another.

50 mM Tris – No Cation

When Htelo DNA was introduced to the merocyanine dyes without the presence of a cationic metal ion, there were clear changes in the optical properties of the dyes as a result of interactions with the DNA strand. In most cases, there were changes in the absorbance characteristics of the merocyanine dyes. In all cases, some level of fluorescence enhancement was observed with the addition of Htelo. The results are illustrated in figures 3.12 and 3.13 and the fluorescent data is summarized in Table 3.3.

108

0.1 A) 0.14 B) 0.09 0.08 0.12 0.07 0.1 0.06 0.08 0.05

0.04 0.06 Absorbance (au) Absorbance (au) 0.03 0.04 0.02 0.02 0.01 0 0 350 400 450 500 550 600 350 450 550 650 750 Wavelength (nm) Wavelength (nm)

0.2 C) 0.4 D) 0.18 0.35 0.16 0.3 0.14 0.12 0.25 0.1 0.2

0.08 0.15 Absorbance (au) Absorbance (au) 0.06 0.1 0.04 0.02 0.05 0 0 400 450 500 550 600 450 500 550 600 650 Wavelength (nm) Wavelength (nm)

Figure 3.12: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer. The increase in colour saturation correlates with the increase in Htelo concentration.

109

30 A) 120 B)

25 100

20 80

15 60

10 40

5 20

Fluorescence Fluorescence Intensity(au) Fluorescence Intensity(au)

0 0 400 500 600 700 400 500 600 700 800 Wavelength (nm) Wavelength (nm)

1000 C) 1000 D) 900 800 800 700

600 600 500 400 400 300

200 200 Fluorescence Fluorescence Intensity(au) Fluorescence Fluorescence Intensity(au) 100 0 0 400 500 600 700 400 500 600 700 Wavelength (nm) Wavelength (nm)

Figure 3.13: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer. The increase in colour saturation correlates with the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra.

110

Table 3.3: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer without the presence of a metal cation

Dye λex λem Emission Emission Irel (nm) (nm) Intensitya Intensityb (au) (au) PhP 500 607 2.9 27.0 9.3 PhQ 588 685 0.7 106.0 151.4 PhBtz 559 588 9.1 912.1 100.2 ThBtz 589 593 10.2 876.0 85.9c a Emission intensity corresponding to the free dye in 50 mM Tris buffer b Emission intensity corresponding to the dye:DNA complex at the highest added concentration of Htelo DNA (in most cases 5 µM Htelo to 3 µM dye) c Only 2.5 µM of Htelo was added in to prevent saturation of the fluorescence signal.

This data provides some insight into how well the dyes are interacting with Htelo DNA in the absence of a metal cation to stabilize any GQ structures. In the case of PhP, there are no real changes in the absorbance capabilities of the dye. PhQ undergoes a red-shift in absorbance maximum, as seen in in Figure 3.12 B, while not demonstrating any change in the chromophore’s absorbance ability. In the case of the benzothiazole derivatives, both PhBtz and ThBtz demonstrate a red-shift in absorbance maximum wavelength as well as a decrease in absorbance ability at the red-shifted λmax (Figure 3.12 C and D). The red-shift in absorbance maximum reflects what has been in observed in the excitation wavelength for the fluorophores, although not to the same extent. As previously discussed, this red-shift in absorbance and excitation wavelength corresponds to an increase in electron delocalization within the ground state of the dye. This occurs as a result of the dye stacking with the G-tetrads of GQ structures that exist in solution. In stacking, the dye is locked into a more planar state, encouraging more electron delocalization attributed to the donor acceptor character of the dyes.37

111

In all cases, when monitoring the fluorescence and emission spectra of the dyes as increasing concentrations of Htelo were added to the free dye in solution, there was an enhancement in the fluorescence output of the dye. This is illustrated in the Irel values determined for all of the dyes, comparing the emission of the free dye to the emission of the dye in the presence of excess DNA (Table 3.3). The pyridine derivative performs underwhelmingly, exhibiting a relative fluorescence intensity of only 9.3. The remaining dye derivatives, PhQ, PhBtz and ThBtz. all exhibit impressive Irel values at 151.4, 100.2 and 85.9 respectively. The quinoline example undergoes the largest fluorescence enhancement of the dyes in this series. Although it is not nearly as fluorescent as the benzothiazole examples, the observed enhancement in fluorescence emission is not unexpected. As discussed in section 3.3.3, based on the CD spectroscopy studies, it was observed that PhQ could readily induce formation of a parallel GQ topology as it was added to

Htelo DNA in the absence of a cationic metal ion. It is not surprising then that this fluorophore demonstrates the largest increase in fluorescence output under these conditions; it induces a preferred topology of GQ, allowing for a more favourable interaction between the dye and the

DNA to give this enhancement in fluorescence emission.

The benzothiazole examples, although they do not boast Irel values that are quite as impressive as the quinoline derivative, these dyes still experience a turn on fluorescence response as a result of being exposed to increasing concentrations of Htelo DNA. In the case of ThBtz, found at a concentration of 3 µM in solution, the titration data presented only includes additions of Htelo up to 2.5 µM to avoid saturating the fluorescence detector (Figure 3.13 D). This fact alludes to a rather favourable interaction between the fluorophore and the DNA under these conditions, considering that the dye species remains in excess within this titration. Even so, the enhancement observed to this point is quite astonishing. Similarly, the PhBtz example, with the

112 addition of 5 µM Htelo, nearly saturates as well demonstrating an intensity of 912.1 au (Figure

3.13 C). There is a key feature that the benzothiazole examples demonstrate that the pyridine and quinoline examples do not. The difference in excitation, or nearly synonymously, absorbance maximum wavelength for these derivatives as increasing concentrations of Htelo DNA are added to the dye in the absence of a metal cation are detectable by the naked eye (Figure 3.14).

Additionally, upon exposure to a UV light through the use of a benchtop light source, the difference in the fluorescence capability of the dye is readily observed by the naked eye as well (Figrue 3.14).

Figure 3.14: The visible change in absorbance maximum wavelength for (top) PhBtz and (bottom) ThBtz as well as the intense fluorescence turn on for both dye derivatives in the presence of Htelo DNA, no metal cation in solution.

These visible changes in the dyes’ behaviour makes them interesting probes for future development. Although this is not the goal of this research, the enhancement in fluorescence

113 output and the absorbance characteristics of these benzothiazole derivatives allow for an academic discussion as to why this may be the case. Details on this can be found in section 3.3.5.

50 mM Tris with 50 mM Na+

0.1 0.14 0.09 A) B) 0.08 0.12 0.07 0.1 0.06 0.08 0.05

0.04 0.06 Absorbance (au) Absorbance (au) 0.03 0.04 0.02 0.02 0.01 0 0 350 400 450 500 550 600 350 450 550 650 750 Wavelength (nm) Wavelength (nm)

0.2 0.40 C) D) 0.18 0.35 0.16 0.30 0.14 0.12 0.25 0.1 0.20

0.08 0.15 Absorbance (au) Absorbance (au) 0.06 0.10 0.04 0.02 0.05 0 0.00 400 450 500 550 600 450 500 550 600 650 Wavelength (nm) Wavelength (nm)

Figure 3.15: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM NaCl. The increase in colour saturation correlates with the increase in Htelo concentration.

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25 A) 140 B) 120 20 100

15 80

10 60 40

5 Fluorescence Fluorescence Intensity(au) Fluorescence Fluorescence Intensity(au) 20

0 0 375 475 575 675 400 500 600 700 800 Wavelength (nm) Wavelength (nm)

800 500 700 C) 450 D) 400 600 350 500 300 400 250 300 200 150 200

100 Fluorescence Fluorescence Intensity(au) Fluorescence Fluorescence Intensity(au) 100 50 0 0 400 500 600 700 800 400 500 600 700 Wavelength (nm) Wavelength (nm)

Figure 3.16: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM NaCl. The increase in colour saturation correlates with the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra.

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Table 3.4: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer with 50 mM NaCl.

Dye λex λem Emission Emission Irel (nm) (nm) Intensitya Intensityb (au) (au) PhP 500 607 3.7 23.8 6.4 PhQ 588 685 0.9 94.0 103.3 PhBtz 559 588 8.4 248.4 29.6 ThBtz 589 593 9.0 436.8 48.5 a Emission intensity corresponding to the free dye in 50 mM Tris buffer b Emission intensity corresponding to the dye:DNA complex at the highest added concentration of Htelo DNA (in most cases 5 µM Htelo to 3 µM dye)

When the merocyanine dyes were exposed to Htelo DNA in a Tris buffered solution containing sodium chloride, providing the metal cation required to stabilize an antiparallel GQ topology within the Htelo DNA sequence, some changes are observed in the optical characteristics of the dyes.12 It is clear that the dyes are still able to interact with the anti-parallel GQ topology, however, it is also clear that the interaction between the dye and the DNA has been affected by the presence of the sodium cation.

Based on the UV-Vis titration data shown in Figure 3.15 A), the pyridine derivative demonstrates a limited response as increasing concentrations of Htelo are added. There is no shifting in absorbance maximum wavelength and a very minimal increase in the absorbance of the dye around 450 nm. In the case of the quinoline derivative, however, the red-shifting in absorbance wavelength of the dye as was observed in the titration without a metal cation present is still observed in the presence of Na+. There is, however, a decrease in the absorbance ability of PhQ in this case which is was not noted in the previous study (Figure 3.15 B). In the case of PhBtz and

ThBtz, similar changes in their UV-Vis absorbance characteristics that were seen in the titration without a cation prevail in the Na+ study. However, when comparing Figure 3.12 A) and B), representing the interaction of the benzothiazole derivatives with Htelo in the absence of a metal

116 cation to the titration data provided above in Figure 3.15 A) and B), representing the changes in the dyes’ in absorbance in the presence of Na+, there are subtle, albeit clear, differences.

In the UV-Vis titrations of ThBtz and PhBtz without Na+, there is a clear red-shift in absorbance wavelength that occurs in conjunction with a decrease in absorbance capability.

Although the same trend is observed for the benzothiazole derivatives in the titration data where

Na+ is present, these changes are not observed nearly to the same extent as was observed from the interaction of these chromophores with Htelo in the absence of a metal ion.

The fluorescence data for the merocyanine dyes tells a similar story as the UV-Vis data.

When discussing the Irel values for all of the dye derivatives in this study as they are titrated against

Htelo in the presence of Na+, all of the dyes still undergo fluorescent enhancement. PhP and PhBtz exhibit the lowest Irel values at 9.4 and 29.6 respectively. The PhQ dye once again exhibits the highest Irel value at 103.3. The largest shock here is the Ire; value observed for ThBtz. When the sodium ion is present, allowing for an anti-parallel GQ structure in solution, the relative intensity nearly decreases by half in comparison to what is observed when there is no metal ion present. In

+ the Na titration data, it is observed that the Irel value for ThBtz is 48.5 when 1.7 equivalents of

DNA were added to the free dye in solution. This value decreased 1.8x in comparison to the Irel observed for the fluorophore in the absence of a metal cation when only 0.8 equivalents of Htelo were added to this dye derivative in solution. It is clear that the presence of the sodium ion plays a role in the ability or inability for the fluorophore to interact with Htelo in solution (Table 3.4).

Considering there was a drop in Irel in all cases of the dyes, the ThBtz example undergoing the most dramatic changes with the presence of Na+, there is very clearly a correlation between these observed decreases in fluorescence out put from the dyes and the presence of the sodium ion.

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It is hypothesized that in the absence of a metal ion to stabilize any GQ structures in solution, the cationic charge found on the ring nitrogen of these merocyanine derivatives will interact with the

G-tetrads of a putative GQ structure in solution, stabilizing it like a cationic metal ion would.

50 mM Tris with 50 mM K+

Although it is clear that the presence of Na+ impacts the ability for the merocyanine dyes to interact with the anti-parallel GQ topology of Htelo in solution, it was of interest to determine if the optical outputs of the dyes would suffer in the presence of potassium ions as they have with the presence of sodium. It may be that the dyes do not have as much of an affinity for an anti- parallel topology and that in the presence of a hybrid GQ structure that K+ would allow, binding may still occur.

As shown in Figure 3.17, the UV-Vis titration data is more indicative of dye binding in the presence of K+ in comparison to the sodium titration data. In all cases, there is a noted red-shift in absorbance maximum as well as a decrease in absorbance ability of chromophore. Particularly for the PhP dye derivative, where under the previous experimental conditions there were no real changes in the absorbance ability of the dye, although it is subtle, there is a slight red-shift and decrease in the absorbance of the dye as more Htelo is added to the solution in the presence of

K+(Figure 3.17 A). With respect to the other derivatives, PhQ, PhBtz and ThBtz,, there is the noted red-shift and decrease in absorbance as has been noted in previous experiments. However, in comparison to the Na+ titrations, the response to Htelo in the presence of K+ is nearly restored to what was observed when there was no cation present at all (Figure 3.17).

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0.10 0.14 0.09 A) B) 0.08 0.12 0.07 0.1 0.06 0.08 0.05

0.04 0.06 Absorbance (au) Absorbance (au) 0.03 0.04 0.02 0.02 0.01 0.00 0 350 400 450 500 550 600 350 450 550 650 750 Wavelength (nm) Wavelength (nm)

0.2 0.4 D) 0.18 C) 0.35 0.16 0.3 0.14 0.12 0.25 0.1 0.2

0.08 0.15 Absorbance (au) Absorbance (au) 0.06 0.1 0.04 0.02 0.05 0 0 400 450 500 550 600 450 500 550 600 650 Wavelength (nm) Wavelength (nm)

Figure 3.17: UV-Vis titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM KCl. The increase in colour saturation correlates with the increase in Htelo concentration.

119

40 100 35 90 80 30 70 25 60 20 50 15 40 30 10

20 Fluorescence Fluorescence Intensity(au) Fluorescence Fluorescence Intensity(au) 5 10 0 0 350 450 550 650 750 400 500 600 700 800 Wavelength (nm) Wavelength (nm)

300 1000 250 800 200 600 150 400 100

50 200

Fluorescence Fluorescence Intensity(au) FLuorescence FLuorescence Intensity(au)

0 0 400 500 600 700 400 500 600 700 800 Wavelength (nm) Wavelength (nm)

Figure 3.18: Fluorescence titrations of 3 µM merocyanine dyes (A) PhP, (B) PhQ, (C) PhBtz and (D)ThBtz with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.25, 2.5, 4 and 5 µM) in 50 mM Tris buffer with 50 mM KCl. The increase in colour saturation correlates with the increase in Htelo concentration. Dashed lines correspond to excitation spectra where solid lines correspond to emission spectra.

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Table 3.5: Summary of the fluorescence data for the titration of each merocyanine dye against Htelo DNA in 50 mM Tris buffer with 50 mM KCl.

Dye λex λem Emission Emission Irel (nm) (nm) Intensitya Intensityb (au) (au) PhP 500 607 4.1 36.5 8.9 PhQ 588 685 0.8 127.6 159.5 PhBtz 559 588 9.0 715.4 79.5 ThBtz 589 593 10.1 944.4 93.5c a Emission intensity corresponding to the free dye in 50 mM Tris buffer b Emission intensity corresponding to the dye:DNA complex at the highest added concentration of Htelo DNA (in most cases 5 µM Htelo to 3 µM dye) c Only 4 µM of Htelo was added in to prevent saturation of the fluorescence signal.

Given that the absorbance data suggested that the merocyanine derivatives were favourably interacting with the hybrid GQ structure of Htelo provided by K+, it was not surprising that the fluorescence data corroborates this claim. As shown in Figure 3.18 and summarized in Table 3.5, once again it is the PhQ merocyanine derivative that demonstrates the largest Irel value with the introduction of Htelo DNA, at 159.5. Again, the phenyl pyridine derivative performs underwhelmingly, demonstrating an Irel that is not even within the same order of magnitude of the other dyes presented here. ThBtz demonstrates a restored turn on in the presence of K+ in

+ comparison to the Na titrations. The observed Irel is 93.5 after the addition of 4 µM Htelo to 3

µM of ThBtz in solution. Surprisingly, in the case of the phenyl benzothiazole dye derivative, the

Irel in the presence of potassium is observed to be only 79.5. Although this is an increase in fluorescence emission in comparison to the sodium trials, this is a noted decrease to what was observed for the dye when there is no cation present when the Irel was found to be 100.9.

It is clear that the dyes show a distinct preference in their ability to interact with Htelo in the absence of a metal cation. In the presence of metal cations, it is clear that there is less of an aptitude for the dyes to bind with the DNA, most notably when sodium is present in the solution.

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It is hypothesized that it is not only pi stacking interactions with the various GQ topologies observed in solution that indicates a favourable interaction between the dyes and Htelo but that it is also an ability for these cationic dyes to stabilize the tetrads of the Htelo GQ that allows for binding to occur. This will be discussed in detail in section 3.3.5.1.1.

3.3.5.1.1. Making Sense of the Differences in Optical Output from the Merocyanine Dyes as They Relate to Varied Conditions The optical changes observed in the dyes can be explained, once again, by the fact that these dyes are molecular rotor probes. The increase in fluorescence output of all of the dyes, in all cases, is a direct result of the dyes being locked into a planar state by interacting with Htelo

DNA.38 In adopting this more planar, LE ground state there is an increased ability for the dyes to undergo LE emission when excited, explaining the fluorescence enhancement.38,39 Based on these observations, as well, there is a strong argument to be made for the fact that these merocyanine dye derivatives are stacking with the GQ of Htelo in solution as opposed to groove binding.24 In stacking with the G-tetrads, the aromatic interactions that occur between the DNA and the dye would favourably induce the planar conformation of the dye, providing the observed turn on fluorescence response. In addition, previous studies performed by Liu et al, assessing the ability for ThT to selectively bind to hybrid GQ structures make it clear that the rotor character of a fluorophore is key in the observed turn on signal when the dye binds to the GQ structure.31 Using a non-rotor analogue of ThT, shown in Scheme 3.2, the same light up is not observed.31 Further evidence that supports the idea that the binding mode of the dyes with Htelo is by stacking will be discussed in 3.3.5.2.24

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Figure 3.19: Structure of the non-rotor analogue of ThT.

To further support the hypothesis that the dyes are stacking with the GQ DNA, locking them into their more planar conformation, is the fact that in all cases, the Stoke’s shift observed as more and more DNA is added to the dyes in solution decreases as a result (Table 3.6). This indicates that there is very little difference between the ground state and excited state structures of the dye.37 If, in its ground state, the dye is planar as a result of interactions with Htelo DNA, upon excitation by a light photon, the dye will relax by radiative means as the planar conformation is stabilized by stacking interactions with the DNA quadruplex.37 The decrease in Stoke’s shift as it relates to the addition of DNA to the system supports the argument that the dyes are effectively stacking with the GQ of Htelo in solution.

Although a brief rationale as to why some of the dye derivatives are more fluorescent than others was made in Section 3.3.3, as this fluorescence relates to the ability for the dyes to interact with GQ DNA, a more indepth discussion as to why different turn on fluorescent responses were observed for different dyes is warranted. Primarily, it is important to mention why the fluorescence of the benzothiazole derivatives is so much more impressive, particularly when DNA is introduced to the dye in solution in comparison to the pyridine and quinoline examples.

Relating to the molecular rotor character of the dyes, once again, the benzothiazole acceptor component is the strongest by far of this series. Based on the Hammett analysis performed in Chapter 2, it is known that the benzothiazole component demonstrates the most electron withdrawing ability. In addition, the N,N-dimethyl mimic phenyl and thienyl donors demonstrate the ability to effectively donate electrons within the system as well. Because of the

123 coupling of a good electron acceptor with a good donor, it is not surprising that the baseline fluorescence of these molecules is quite high. Even in the ground state, there is a certain amount of electron delocalization, contributing to the fluorescence output of the fluorophore. Upon binding of ThBtz and PhBtz to the Htelo GQ, under all conditions, it is also not surprising that the fluorescence increases to the extent it does. Binding to the DNA to lock the dye into it’s planar conformation, allows for an increase in LE emission, explaining the large enhancement in emission.38,39

Table 3.6: Summary of Stoke’s Shift data for all of the merocyanine dyes under all experimental conditions employed: 50 mM Tris, 50 mM Tris with 50 mM NaCl or 50 mM Tris with 50 mM KCl

a a a b b b Compound Metal Ion λex λem ΔS λex λem ΔS (nm) (nm) (nm) (nm) (nm) (nm) PhP n/a 461 608 147 506 608 102 Na+ 471 605 134 495 602 107 K+ 472 606 134 503 605 102 PhQ n/a 548 679 131 587 681 94 Na+ 548 686 138 585 676 91 K+ 552 678 126 585 677 92 PhBtz n/a 531 603 72 564 608 44 Na+ 538 602 64 560 606 46 K+ 535 603 68 560 607 47 ThBtz n/a 560 600 40 584 595 11 Na+ 551 600 49 581 600 19 K+ 554 600 46 574 600 26 a Denotes dye in its free state (no DNA added) b Denoted dyes in its fully bound state (2.5 – 5 µM of DNA added) ΔS denotes Stoke’s Shift

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Comparing the benzothiazole derivatives to the quinoline and pyridine derivatives, although the baseline fluorescence is not as pronounced for PhP and PhQ, there is still a noted increase in fluorescence for these dyes as Htelo is added to the solution, particularly in the case of the quinoline example. Based on the observations made in Chapter 2, the ability for the pyridine and quinoline examples to effectively act as electron acceptors is not as good as their benzothiazole counterpart. This is attributed to the aromatic character of these acceptors. Because pyridine and quinoline are more aromatic than benzothiazole, these derivatives are not as willing to undergo through resonance as doing so would break aromaticity within the acceptor moiety. This readily explains the lack of fluorescence before the addition of DNA to these dyes in solution. Without any interaction to increase planarity within the dye’s structure, the dye can freely rotate in solution, allowing for relaxation via non-radiative pathways following excitation by a light photon.39

However, the turn on response in the presence of Htelo DNA still indicates a stacking interaction between PhP and PhQ, locking the dye into a planar state, explaining the turn on fluorescence in the presence of the GQ structure.

In comparing the UV-Vis titration data and the fluorescence data for all of the dyes under all of the varying conditions employed in these studies, it is clear that a majority of the merocyanine dyes were able to preferentially interact with the DNA in the absence of a metal cation. This is rationalized on the basis that when sodium was introduced to the solution, the Irel values for all of the merocyanine dyes decreased in comparison. In addition, although the addition of K+ to the solution allowed for most of the dyes to regain some of the fluorescence intensity lost when sodium metal was present, particularly in the case of the benzothiazole derivatives, the fluorescence output still does not reach the levels that were observed when no metal cation is present. It is hypothesized that the cationic nitrogen group of the dyes has the ability to interact with the G-tetrads of the Htelo

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GQ in solution through stacking to stabilize the GQ structure of Htelo.19 In the absence of a metal cation, the only entity in the solution bearing a positive charge to effectively stabilize a GQ tertiary structure would be the dye itself. This explains why the most fluorescence enhancement from the merocyanine derivatives is observed in the absence of metal cations; there is nothing to out compete the dye in it’s ability to place the cationic charge it bears on the ring nitrogen of its structure into the central cavity of the G-tetrad of a putative GQ in solution, stabilizing its structure.

Without the sodium or potassium ion to compete with, it makes it easier for the dye to stack with the GQ in this case as there is no interfering cation to prevent the interaction of the dye with the

Htelo DNA.

It has been noted previously in the literature that ThT, a known GQ binding fluorophore, can flip the hybrid structure of the Htelo GQ that occurs in solutions containing potassium to an antiparallel topology.19 In the case of the ThBtz and PhBtz dye derivatives, the recovery of fluorescence output that is observed in the presence of K+ versus what is observed in Na+ can be explained by the ability for these dyes to behave in a similar manner. The ability for the dyes to flip the conformation of a GQ in solution to a topology that allows for an increased ability for both entities to interact with one another explains the enhancement in UV-Vis and fluorescent response of the chromophore. Flipping the DNA to a more favourable conformation allows for a more optimal stacking interaction, explaining the enhancement in optical outputs from the dyes in the presence of potassium. This is not surprising, as well, considering that ThT and the ThBtz and

PhBtz dye derivatives bear the same benzothiazole donor component.19

Based on the above results, it is clear that the merocyanine dyes are capable of interacting with all of the Htelo GQ topologies based on the changes in the LE emission of the dyes. What is an interesting facet to these studies, however, is to determine the ability for the dyes to undergo

126

ET with the DNA in solution and how does this transfer of energy relate to the fluorescence ability of the dyes. Given the literature precedent of ET between quadruplex DNA and free fluorophores as discussed in section 3.1.4, it was of interest to determine if these dyes were at all capable of undergoing similar process.

3.3.5.2. ET Emission with GQ Binding An interesting observation was made in the previous fluorescence titration experiments. It was noted that upon monitoring the excitation spectra for the merocyanine dyes as more Htelo

DNA was added to the solution, there was the emergence of an excitation band centered around

256 nm. Probing this finding further, it was determined that by exciting the chromophore in the presence of GQ DNA at 256 nm, there was a corresponding emission band within the visible region of the EM spectrum. This is illustrated in Figure 3.19 using the ThBtz example. When the dye is free in solution and excited at it’s absorbance maximum of 551 nm there is fluorescence emission observed at 588 nm. However, if the free dye were to be excited at 256 nm, there would likely be minimal fluorescence emission observed within the visible region of the EM. The visible emission of the chromophore observed upon excitation at 256 nm in the presence of DNA that corresponds to its natural fluorescence is due to energy transfer from the DNA to the merocyanine dye.

The ability for the dyes to respond to the presence of the DNA in this manner provides a means to determine the affinity with which the dyes will bind to the various conformations of GQ

Htelo DNA can adopt. As shown in Figure 3.19 using the ThBtz example, it is clear to see that the fluorescence intensity in both the excitation and emission bands of the fluorophore begins to plateau as the amount of DNA added to the dye in solution nears and surpasses one equivalent under all sets of buffer conditions. This phenomenon was observed for all dye derivatives. What this indicates is that the dye and the GQ bind in a 1:1 stoichiometry and that, at least under these

127 experimental conditions, no more than one dye molecule will interact with a single GQ in solution.

This observation differs from what is observed for the LE emission of the dye. Exciting at the excitation maximum wavelength for the dye leads to a linear increase in fluorescence emission within the visible region of the EM spectrum. Using the ET excitation band demonstrates that the dye does in fact bind to the DNA and does so at a 1:1 ratio. This finding allows for the determination of dissociation constants for each of the dyes, for each topology of GQ Htelo can adopt based on the experimental conditions of the given ET fluorescence titration.

In addition, it is possible to utilize this finding to fully characterize the binding mode between the dye and the Htelo GQ. Based on the examples explained in section 3.1.4, namely the ability for the fluorophore Crystal Violet to interact with G-tetrads in a manner that allows for energy transfer to be observed.23,24 Based on the literature precedent provided, it is clear that a similar process is occurring with these novel merocyanine dyes. In order for ET to occur between a GQ of the Htelo DNA and the merocyanine dye, it is crucial that the dye and any DNA base pairing interaction within the GQ be in close proximity with one another. This is only possible when the fluorophore can either stack on the top of the bottom tetrad of the GQ of to intercalate between G-tetrads.23 If the fluorophore had been groove binding or adopting another mode of binding to the GQ outside of the core structure, the ET emission observed in these studies would not have occurred.

128

250 A) 200

150

100

0 Fluorescence Fluorescence Intensiy(au) 200 300 400 500 600 700 Wavelength (nm)

120 B) 100 80 60 40 20 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

300 C) 250 200 150 100 50 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelenght (nm)

Figure 3.20: ET excitation and emission spectra of 5 µM ThBtz in (A) 50 mM Tris (B) 50 mM Tris with 50 mM NaCl an (C) 50 mM Tris with 50 mM KCl with increasing concentrations of Htelo DNA (0, 0.12, 0.25, 0.75, 1.5, 2.5, 4, and 5 µM) using λex 256 nm.

Clearly, the ET phenomenon as it relates to the merocyanine dyes and their ability to interact with the Htelo GQ, provides a clearer picture of how the dyes are able to interact with the DNA. It is clear that the dyes are effectively stacking with the GQ, explaining the ability for

129 the dyes to undergo ET fluorescence. Although the ET fluorescence aids in determining how the dyes bind to the DNA, this fluorescent characteristic can also be used to determine how much affinity these novel dyes have with the Htelo DNA. How the ability for the dyes to interact favourably with the varying topologies of the Htelo GQ, contingent on what metal cation is present can be explained by examining the Kd values of the novel chromophores calculated using the ET emission for each dye with the addition of Htelo DNA. How the binding affinity of the dyes changes as a result of GQ topology will be discussed further in section 3.3.6.

3.3.6. Binding Affinity of Merocyanine Dyes with Htelo Based on the various optical responses observed from the dyes as a result of the Htelo GQ found in solution, it was of interest to determine if the presence of either sodium of potassium metal ions in solution affected the ability for the dyes to bind to Htelo. By calculating the Kd values for each of the dyes, under all of the experimental conditions described, using the ET fluorescence observed through out these studies, the ability for the dyes to effectively bind to the

GQ DNA can be quantified. The Kd values for the dyes and Htelo DNA in the absence of a metal cation and in the presence of both sodium and potassium are highlighted and summarized in Table

3.7.

The observation that becomes immediately clear is the fact that the ThBtz example of the dyes binds with the highest affinity to the Htelo DNA in all conditions. Particularly, when there is no metal ion present in solution, the ThBtz dye demonstrates a Kd value of 0.9 µM, the lowest dissociation constant among all of the dyes, for all of the conditions explored. What is also

+ + intriguing for this dye is that when either K of Na is present, the Kd value does not change as a result, staying consistent at 1.1 µM. These results aid in explaining what has already been observed throughout the UV-Vis and fluorescence experiments. The ThBtz dye example not only

130 demonstrates the lowest Kd value as demonstrated using the ET emission of the fluorophore to calculate the binding affinity, but throughout the optical studies, it demonstrates the largest changes in LE emission as well.

Table 3.7: Summary of Kd values determined for all of the merocyanine dyes in 50 mM Tris in the presence of: no metal cation, 50 mM Na+, or 50 mM K+

Compound Metal Ion Kd Standard (µM) Error a PhP n/a 1.6 0.2 Na+ 1.8 0.1 K+ 2.3 0.1 PhQ n/a 2.7 0.3 Na+ 2.1 0.2 K+ 2.7 0.3 PhBtz n/a 1.0 0.2 Na+ 1.8 0.2 K+ 1.5 0.2 ThBtz n/a 0.9 0.1 Na+ 1.1 0.1 K+ 1.1 0.1 a Standard error values were determined from the curve fitting macro used for a one-site ligand binding mode, generated in SigmaPlot 12.0.

As previously mentioned, because the benzothiazole acceptor moiety of this dye demonstrates the best electron withdrawing ability, it is hypothesized that there is some level of planarity observed in the ground state of this structure, explaining the impressive baseline fluorescence it demonstrates even in the absence of Htelo DNA. This coupled with the high affinity the dye has with the DNA in solution, it is hypothesized that the high donor acceptor character of the ThBtz merocyanine is what contributes to its ability to interact with GQ DNA. If

131 the dye is found in a planar conformation to some extent prior to the addition of DNA to the solution, it is not surprising that the dye would have a much higher aptitude for interacting with the DNA. If there is no energetic barrier to over come to achieve a more planar conformation, for example, if there is no twisting required to achieve a planar state, it is conceivable to believe that a stacking interaction would be far more favourable to occur between an already planar species and the G-tetrads of the GQ DNA (Figure 3.20 A).

In contrast, the above data also readily distinguishes PhQ as the worst binder overall, under most conditions utilized in these studies, using ET emission to calculate the Kd of the dye with

+ + Htelo in solution. With Kd values of 2.1 in solutions containing Na and 2.7 in the presence of K and in the absence of a cation as well, it is clear that this molecule does not bind with Htelo as well as the other dye derivatives presented here. It is certainly possible to relate the ability for the PhQ dye to bind to DNA based on the acceptor ability of the quinoline moiety. As previously discussed, the quinoline and pyridine acceptors demonstrate the highest aromatic character, describing their inability to effectively draw electron density with the dye structure. This indicates that there is a higher percentage of twisted dye species in solution as the propensity for the dye to adopt a planar conformation is not as strong as what has been observed from the benzothiazole examples of the merocyanine dyes.

Although in all cases PhQ demonstrates the largest increase in LE emission as it is introduced to more DNA in solution, it is clear that this is a direct result of interactions occurring with Htelo DNA. The baseline fluorescence of PhQ is basically non-existent, lending the increases in observed fluorescence output to be caused by the introduction of DNA. Because the dye exists in its twisted state prior to the addition of DNA, it is required then, that the dye undergo a twisting motion to give rise to the planar conformation in order to effectively bind to the DNA. Although

132 adding DNA to the solution makes this a more favourable interaction, it is clear that this added barrier to binding contributes to the observed affinity of the dye for the Htelo GQ (Figure 3.20 B).

To illustrate the disparity among Kd values for the ThBtz and PhQ dye derivatives, the binding isotherms are shown in Figure 3.20 C).

When it comes to describing how the various dyes preferentially bind to the varying topologies of Htelo GQ, it is clear that there are differences in the ability for the dyes to bind, contingent on their structure. For instance, when looking at the benzothiazole derivatives, both

PhBtz and ThBtz clearly prefer interacting with Htelo in the absence of a metal cation with Kd values of 1.0 and 0.9 respectively. What this indicates is what has already been hypothesized; in the absence of a metal ion to stabilize the DNA tertiary structure, the cationic nitrogen of the benzothiazole moiety is able to act as the positive charge required to stabilize the G-tetrads of a putative GQ in solution.19 Additionally, the benzothiazole dye derivatives demonstrate a propensity to favourable interact with the hybrid GQ structure found in solution, provided by the presence of K+. Particularly in the case of PhBtz, when comparing the ability for the dye to interact with the GQ in the presence of sodium versus that of potassium, there is a decrease in Kd from 1.8 to 1.5 µM. As mentioned previously, a known GQ binding fluorophore, ThT, is capable of binding to the hybrid GQ structure of Htelo found in solutions containing K+ and flipping the topology from a hybrid species to an anti-parallel GQ.19 It is believed that the dyes presented here, bearing the same benzothiazole component as ThT, are capable of carrying out a similar process (Figure

3.21). As the dye is able to interact with the hybrid GQ topology and flip to a conformation that is more favourable, this readily explains why the Kd values for the benzothiazole derivatives is lower in K+ than in Na+.

133

C) 1.2

0.8

0.6

0.4

Fractionof Dye Bound 0.2

0 0 1 2 3 4 5 [Htelo] (µM)

Figure 3.21: A) The conformational equilibrium that occurs for the ThBtz dye derivative. Because the planar state of the dye occurs when the dye is free in solution, this explains the high baseline fluorescence for the dye. The planar conformation is suspected to prefentialy bind with Htelo, explaining the enhancement in ET emission and low Kd B) Explaining the increase in fluorescence emission of PhQ in the presence of Htelo. Although this dye is not expected to undergo through resonance, binding with the GQ will induce a more planar conformation of the dye, explaining the increase in ET fluorescence emission. C) Binding isotherms for ThBtz (blue trace) and PhQ (red trace).

Why the benzothiazole derivatives are capable of interacting favourably with the anti- parallel topology observed in K+ that is caused by interactions of the dye with the DNA can be

134 explained using the ThT example. Primarily, in the case of ThT, upon stacking with a parallel GQ structure of Htelo, the fluorophore severely affects the co-ordination of the G residues within the tetrad with the central sodium metal ion.19 In doing this, the stacking distance between the tetrads of the GQ is also affected, leading to the overall destabilization of the overall structure.19

When ThT binds to the anti-parallel GQ, none of the destabilizing features are observed as in the former case (Figure 3.21). Although the merocyanine dyes presented here are not exact replicas of ThT, their structures remain similar with only a single vinyl group separating the donor and acceptor moieties of the dyes. ThT is used to model how the merocyanine dyes of this work are perceived to behave in the presence of Htelo DNA under the various conditions employed throughout these studies.

Figure 3.22: A) Binding of ThT to the parallel GQ of Htelo. The side view of the complex shows the distortion of the top tetrad that results from binding of the fluorophore. The top view demonstrates how ThT stack on the tetrad in the presence of Na+. B) Binding of ThT to the anti- parallel GQ of Htelo. It is readily observed from the side view that there is no distortion in the tetrad interacting with the fluorophore. The top view shows the staking of ThT with the top tetrad. Reprinted with permission from Mohanty, J.; Barooah, N.; Dhamodharan, V.; Harikrishna, S.; Pradeepkumar, P. I.; Bhasikuttan, A. C. J. Am. Chem. Soc. 2013, 135, 367. ©2013 American Chemical Society. C) A comparison of the structure of the dyes discussed in these studies with that of ThT.

135

Although it is clear that the benzothiazole dye derivatives prefer to interact with Htelo without the presence of a metal ion or in the presence of K+, this same trend is not what is observed for the pyridine and quinoline merocyanine derivatives. Based on the calculated Kd values for PhP and PhQ with Htelo in the presence of Na+, PhQ demonstrates more of an affinity with this topology in comparison to topologies present in solutions containing no cation or K+. The quinoline dye derivative demonstrates a higher affinity in sodium with a Kd of 2.1 µM in comparison to 2.7 µM under the other conditions described. Similarly, although PhP does show a propensity to bind to Htelo in the absence of a metal cation with a Kd of 1.6 µM, it clearly prefers to bind to the defined anti-parallel topology provided by Na+ in comparison to the hybrid topology

+ in K demonstrating Kd values of 1.8 and 2.3 µM respectively. What is interesting about these results is the fact that, from the CD studies, it was observed that PhQ induced a parallel GQ topology in the absence of a metal cation. PhP, it could be argued, shows a slight propensity to do the same. It is possible that in the presence of sodium, with the defined anti-parallel GQ structure in solution, that PhP and PhQ will preferentially bind to this topology over the hybrid topology found in the presence of K+. As previously discussed, it is hypothesized that PhQ can more readily interact with a parallel GQ topology as the there are no large loop regions interfering with the tetrads of this GQ conformation, allowing for an optimal interaction between this example of the dyes and the GQ DNA.25 However, even though these dyes seems to prefer the anti-parallel GQ topology, if the opportunity for the dyes to bind to a parallel GQ presents itself, as it does when no metal ions are present in solutions, these merocyanine dyes will induce a topology that maximizes preferential interactions between the fluorophore and the GQ of Htelo DNA.

Even though there are distinct differences in the ability for these dyes to interact with a variety GQ topologies, these differences are readily explained by the structure activity

136 relationships discovered through out these studies. It is clear that the structure of the dye plays a significant role in the chromophore’s ability to interact with the Htelo DNA in solution. These studies provide a solid foundation of knowledge on what structural facets lead to an increased ability for GQ: fluorophore binding. It is the hope of this work that it will contribute to future research efforts in determining what fluorophores will be best suited for GQ binding and how will this property will aid in developing novel detection platforms, novel therapeutic uses among many other applications.

3.4. Conclusions The studies presented above demonstrate that this series of merocyanine dye derivatives are able to interact with Htelo GQ DNA under a variety of experimental conditions. This has been observed through the use of CD spectroscopy as well as through changes observed in both the absorbance and fluorescence capabilities of the dyes as they are introduced to Htelo DNA in solution. It is clear that these dyes can readily be classified as molecular rotor probes seeing as stacking with the GQ structure in solution effectively forces the dyes into their planar conformations, leading to the observed enhancement in LE fluorescence emission as a result of dye:GQ binding.

Although these dyes are capable of undergoing LE emission, it has been demonstrated that upon GQ binding, ET from the DNA to the fluorophore leads to ET fluorescence emission. This facet of the dye’s characteristics not only provides proof that the binding mode of the dyes with the Htelo GQ is in fact a stacking interaction, this ET emission also provides a means to quantify the ability for the dyes to interact with the various topologies of Htelo GQ as it relates to the cation present in solution. This is done by calculating the Kd values of the dyes with Htelo DNA under all of the experimental conditions explored in this work. The Kd values calculated here aid in

137 understanding why certain merocyanine dyes preferentially bind to certain GQ topologies over another. It is clear that the structure of the dyes plays a pivotal role in GQ binding ability. As such, it is our hope that this research will aid in understanding what structural features and what dye properties can increase the ability for a given fluorophore to effectively bind to GQ DNA, paving the way for novel, even more potent GQ binding fluorophores in the future.

138

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Chapter 4: Developing an Aptamer Based Detection Platform for a Small Molecule Target: An Energy Transfer Approach

142

4.1. Introduction 4.1.1. Ochratoxin A OTA is a mycotoxin that is produced by several fungal species from the genera Aspergillus and Penicillium (Figure 1).1 These fungi are most commonly found to grow in various grains and cereals, making OTA a common food contaminant in a variety of consumer products.1 Warranting a class 2B ranking from IARC, OTA is possibly carcinogenic to humans.1,2 In addition, OTA has demonstrated teratogenic, nephrotoxic and immunosuppressive capabilities in a number of animal models, demonstrating true toxicological risks to human health upon consumption of this natural product.3,4,5 Due to the grave toxicological outcomes of consuming OTA, the European

Commission has gone so far as to instill guidelines of acceptable OTA levels in varying categories of food stuffs.6 For example, the maximum allowable levels of OTA acceptable ine wine is on the order of 2 µg/L.6 With such guidelines in place, the ability to detect OTA in a variety of samples has become a fervently studied area of research.

Figure 4.1: The molecular structure of Ochratoxin A

4.1.2. Current OTA Detection Methods A variety of methods have been used to detect and quantify OTA at concentrations on the order of ng/mL. For example, HPLC, LCMS and immunoaffinity based platforms are often regarded as gold-standards for the detection of this toxin.7,8,9 Recently, the use of aptamer based platforms for the detection of a variety of targets has become a popular research topic in the realm of biotechnology. Aptamers, synthetic DNA or RNA oligonucleotides that are selected to bind to

143 a specific target, are rapidly becoming a sought after means for the detection of OTA.10,11

Aptasensor platforms are an attractive alternative to other detection methods as aptamers are readily synthesized in vitro and can replace antibodies as a recognition element within a bioassay while still maintaining similar selectivity and specificity for the target molecule without eliciting an immune response in a host organism. Additionally, aptamers are a more cost-effective method for the detection of OTA in comparison to the aforementioned methods.

4.1.3. Aptamer Based Platforms for OTA Detection A number of aptamers have been selected to bind to OTA as of late.12,13,14 The first instance of the selection against this target produced a 36-mer DNA oligonucleotide that boasts a

12 dissociation constant (Kd) with OTA of 0.2 µM. This is not dissimilar to the Kd determined for the antibody-OTA example which is found to be 0.6 µM.15

As this methodology for OTA detection becomes more and more prominent, the OTA:

OTA aptamer (OTAA) platform is consistently being regarded as a proof-of-concept system when developing aptasensor assays for the detection of small molecules.16 In fact, the OTAA platform has been applied to impedimetric and other electrochemical methods to detect this toxin with success.17,18, 19,20 In addition, there exist a number of free-dye assays as well as platforms that make use of carbon nanostructures that use the OTA:OTAA example.14,21,22 These examples often rely on fluorescence or colorimetric changes of an external chromophore to report the presence of

OTA in a given sample. An ideal platform would remove this added level of complexity in the assay by relying on changes that occur in the optical characteristics of either the target itself or an internal base modification made to the aptamer strand.

144

4.1.4. Inspiration for the Current Study: A FRET Based Label Free Assay for OTA Detection OTA is an inherently fluorescent molecule. It possesses an excitation wavelength of 344 nm and an emission maximum wavelength of 460 nm in water when the phenolic alcohol within its structure is fully protonated.23,24 This makes OTA readily amenable to analysis via fluorescence spectroscopy. In fact, HPLC protocols that utilize fluorescence detection provide limits of detection for this toxin that are lower than one of the decidedly most sensitive methods available, namely, the use of LC-MS/MS protocols.25,26 The ability for OTA to behave as its own reporter within a fluorescence based assay is an avenue that is often overlooked when developing novel bioassays for the detection of this target. However, studies performed by Li et al. 27 capitalize on the inherent fluorescence of OTA and provide a precedent for the current study presented here.

Their studies have shown the successful use of fluorescence resonance energy transfer (FRET) to signal OTA binding to its protein antibody.27 Exciting the tryptophan residues of the antibody at a wavelength of 280 nm resulted in an enhancement of visible emission from the toxin itself at 430 nm. Translating this ability to an aptamer based platform is the overarching goal of this work.27

Previous studies in our laboratory, using a 31-mer analogue of the OTAA propose that the aptamer will fold into an intramolecular anti-parallel G-quadruplex (GQ) upon target binding.28

The OTAA aptamer sequence is rich in guanine (G) residues, indicating high potential for GQ formation.28 GQs are composed of stacked layers of G-tetrads that house a K+ ion in the central cavity of their planar structure.29 It is hypothesized that OTA stacks with the GQ by interacting with one of the tetrads of this DNA secondary structure (Figure 2).28

145

Figure 4.2: A) The DNA sequence of OTAA36 B) Pictorial demonstration of a G-tetrad C) The proposed GQ structure of OTAA36 upon target binding, shown here housing a potassium metal ion

Herein, we demonstrate a turn-on response of OTA fluorescence upon binding to OTAA36.

By exciting the DNA, a non-fluorescent molecule, at a wavelength of 256 nm, energy transfer (ET) from the DNA to the target results in a 3-fold enhancement of visible fluorescent emission of OTA at 425 nm (Figure 4.2). The photophysical capabilities of cation-housing GQs and their ability to participate in ET has been explored through the use of fluorescent internally modified G bases.30,31

The principle of ET between DNA and a variety of non-covalent GQ targeting fluorophores such as Crystal Violet and TMPyP has been reported providing a rationale for the current study.32,33

The affinity with which OTAA36 binds to OTA will be explored and the specificity and selectivity of this assay will be discussed. Finally, a limit of detection for the platform will be presented.

146

Figure 4.2: A schematic demonstrating the “turn on” platform. The OTA:OTAA36 complex is subject to λex=256 nm. ET from the DNA to the toxin causes an enhancement in emission at λem= ~430 nm.

4.2. Materials and Methods The 36mer OTA aptamer (5’GATCGGGTGTGGGTGGCGTAAAGGGAGCATCGG

ACA3’) was purchased from Sigma-Aldrich Ltd with polyacrylamide gel electrophoresis specified for purification. The 15mer thrombin binding aptamer (5’GGTTGGTGTGGTTGG3’) was purchased from Sigma-Aldrich Ltd with reverse-phase cartridge chromatography specified for purification. OTA (99.5%) was supplied by a collaborator at the University of Münster, Germany.

A stock solution of OTA was prepared in MeOH and quantified by UV-Vis at 333 nm with an extinction coefficient of 6400 mol-1 cm-1. A 100 µM solution of OTA was prepared from this stock and used for these studies. OTB was purchased from Toronto Research Chemicals Canada. A stock solution was prepared in MeOH by UV-Vis at 318 nm with an extinction coefficient of 6900

-1 -1 mol cm . CaCl2, NaCl, tris phosphate, KCl, KOH, HCl, Na2HPO4 ad KH2PO4 used for the preparation of the OTA binding buffer and phosphate buffered saline (PBS) were purchased from

Fisher Scientific and used as received. All buffer and stock solutions were prepared using Milli-

Q water that was produced using a Milli-Q system (18 MΩ). All fluorescence spectra were obtained using a Cary Eclipse Fluorescence Spectrophotometer equipped with a 1 x 4 Multicell block and Peltier thermal control. Excitation and emission slit widths were set to 5 nm and 5 nm

147 and all measurements were taken at room temperature. All prepared buffers were adjusted to a pH of 7.2.

4.2.1. Preparation of OTA Binding Buffer. The OTA binding buffer (OTABB) was prepared as a 10x concentrated stock solution containing 0.2 M CaCl2, 1.2 M NaCl, 0.1 M Tris phosphate and 0.05 M KCl. This solution was standardized to pH 8 using a Thermo Orion 320 Basic PerpHect LogR pH meter.and used as required.

4.2.2. Fluorescence Titration of OTA with OTAA36. OTA was prepared as a 1000 µL solution with a concentration of 1 µM in a Hellma

Analytics 119.004F-QS quartz cuvette. A 1x concentration of OTABB was added to the solution.

OTAA36 was titrated into the OTA solution until a concentration of 1.2 equivalents of aptamer to

OTA had been added. Fluorescent readings were taken after manual mixing and 5 minutes of incubation at room temperature following the addition of aptamer to the solution. Fluorescence emission spectra were obtained at excitation wavelengths of 375 nm and 256 nm. Fluorescence excitation spectra were obtained at an emission wavelength of 430 nm.

4.2.3. Kd Determination. From the fluorescence titration data, the Kd was determined for the OTA:OTAA36 complex using an excitation wavelength of 256 nm by monitoring the increase in fluorescence emission intensity around 430 nm. A plot of the fraction of bound OTA versus the concentration of

OTAA36 generated a binding isotherm that was analyzed with SigmaPlot 12.0 using the one site saturation binding macro to obtain the Kd values.

4.2.4. Circular Dichroism Studies. CD spectra were performed on a Jasco J-815 CD spectrophotometer equipped with a thermal controlled 1 x 6 multicell block. Two 200 µL solutions, one of only OTAA36 and one of

148

OTA and OTAA36 were prepared to a final concentrations of 3 µM aptamer and 3 µM of toxin where stated. A 1x concentration of OTABB was added to each solution. Spectra were obtained at 10 oC in quarts cells (110-QS) with a path length of 1 mm and monitored between 200 and 450 nm at a bandwidth of 1 nm and scanning speed of 100 nm/min. A minimum of four scans were collected and combined. Data was corrected against a blank measurement of the OTABB. The spectra were smoothed using the Jasco Spectra Analyzer software by Savitzky-Golay function with

25 convolutions.

4.2.5. Calibration Curves. Solutions of free OTA and OTA:OTAA36 complex were prepared to final volumes of 100

µL. A 1x concentration of OTABB was added to all solutions. Seven solutions of OTA were prepared ranging in concentration from 1.0x10-4 µM to 100 µM toxin, each subsequent solution increasing by an order of magnitude. Three fluorescence readings were taken for each solution at

λex= 256 nm in a Hellma Analytics ultra micro cuvette 105.253-QS with a path length of 10 x 2 mm. The emission intensity was averaged over the three determinations. A plot of log OTA concentration versus log fluorescence emission intensity was constructed to determine the linear range for OTA at λex= 256 nm. Six additional solutions of OTA were prepared to concentrations of 0.25, 0.5, 0.75, 2.5, 5 and 7.5 µM in order to construct the calibration curve. Three readings were taken for these solutions and the emission intensity averaged. A plot of OTA concentration against fluorescence emission intensity was used to obtain the calibration line. The same methodology was employed for the OTA:OTAA36 complex, maintaining the aptamer concentration at 0.7 µM throughout to allow for a known concentration of OTA:OTAA36 complex in solution. This was determined using the Kd equation:

[푂푇퐴][푂푇퐴퐴36] 퐾푑 = 0.7 µ푀 = [푂푇퐴: 푂푇퐴퐴36]푐표푚푝푙푒푥 149

The six additional solutions required to complete the calibration curve for the complex and were prepared to concentrations of: 0.025, 0.05, 0.075, 0.25, 0.5 and 0.75 µM. A plot of OTA:OTAA36 concentration against fluorescence intensity was used to obtain the calibration line.

4.2.6. Determination of LOD and LOQ. Ten fluorescence readings were obtained for a blank solution of OTABB. The standard deviation (σ) of the emission intensities was determined and used to determine the LOD and LOQ for both free OTA and OTA:OTAA36 complex at λex = 256 nm. LOD was determined using:

3σblank/slope of the calibration line (m). LOQ was determined using: 10σblank/m.

4.2.7. Control Fluorescence Titrations. 1 µM solutions of OTB and OTA were prepared to final volumes of 1000 µL containing

1x OTABB. OTAA36 was added to the OTB solution until an equimolar concentration of aptamer had been added. Fluorescence emission was monitored at λex= 256 nm and excitation was monitored at λem= 420 nm. TBA was added to the OTA solution until an equimolar concentration of aptamer had been added. Fluorescence emission was monitored at λex= 256 nm and excitation was monitored at λem= 432 nm.

4.3. Results and Discussion 4.3.1. Fluorescence Titration of OTA with OTAA36 Figure 4.4 shows the changes in fluorescence excitation and emission of OTA with the addition of increasing amounts of OTAA36. Excitation spectra for both studies were obtained using an emission wavelength of 430 nm as increasing concentrations of OTAA36 were added to the free toxin in solution. The excitation band observed around 375 nm decreases in intensity and red shifts to 389 nm as OTA binds to the aptamer strand. As more OTAA36 is added, the emergence of a broad, prominent peak with a maximum wavelength of approximately 254 nm is observed.

150

Using an excitation wavelength of 375 nm, the emission intensity of the toxin at 432 nm not only decreases but the wavelength also shifts toward the blue to 425 nm demonstrating Δλem

=8 nm (Figure 3A). In contrast, Figure 3B demonstrates that when using an excitation wavelength of 256 nm, the fluorescence emission intensity around 430 nm undergoes a 3-fold increase in intensity while still experiencing a blue shift to 425 nm. The blue shift that is observed in the emission spectrum for both of these studies indicates that OTA does in fact pi stack with the GQ of the aptamer strand.34 This is further corroborated by the red shifting observed in the excitation spectrum as the toxin enters a more non polar environment in interacting with the GQ of the DNA as opposed to the aqueous environment it had previously resided.35

151

1000 900 800 700 600 500 400 300

200 Fluorescence Intensity (au) Intensity Fluorescence 100 0 200 250 300 350 400 450 500 Wavelength (nm)

1000 900 800 700 600 500 400 300

200 Fluorescence Intensity (au) Intensity Fluorescence 100 0 200 250 300 350 400 450 500 Wavelength (nm) Figure 4.4: A) Fluorescence titration of 1 µM OTA with increasing concentrations of OTAA36 (0 to 1.2 µM) with λex 375.). B) Fluorescence titration of 1 µM OTA with increasing concentrations of OTAA36 (0 to 1.1 µM) with λex 256 nm

The explanation for the observed decrease in OTA emission at 430 nm when exciting at

375 nm is two-fold. First, the excitation wavelength of 375 nm corresponds to the excitation of free, deprotonated toxin in solution.23 As more OTAA36 is added, the concentration of free OTA in solution decreases as the toxin binds to the aptamer strand. This results in an increase in concentration of OTA:OTAA36 complex. Secondly, and as stated previously, there is the

152 emergence of a large band around 256 nm in the excitation spectrum as increasing amounts of aptamer are added to the toxin in solution. This peak corresponds to an increase in GQ formation within the system and can be described as an ET band that represents energy transfer occurring between the aptamer and its target.36,37 As more of this toxin-aptamer complex begins to form in solution, there is an increased ability for energy transfer to occur between the two entities.36 Using the excitation wavelength of 256 nm provides emission of not only the toxin but of the

OTA:OTAA36 complex as a whole. The addition of more aptamer results in the formation of more target bound complex which ultimately leads to an increased amount of ET within the complex. This provides the observed enhancement of visible emission centered at 430 nm.

4.3.2. Dissociation Constant Determination and Circular Dichroism Binding Studies Using the fluorescence titration data and the ET excitation wavelength of 256 nm, a Kd for the OTA:OTAA36 complex was determined. Monitoring the fluorescence emission at an analytical wavelength of 433 nm, the Kd was found to be 0.7±0.1 µM. Initial studies performed

12 by Cruz-Aguado and Penner find the Kd for the complex using equilibrium dialysis to be 0.2 µM.

Differences in the experimental method can account for the discrepancies between these values.

However, the data presented in this work is considered to be consistent with the literature value, indicating that the ET fluorescence-aptamer based approach to OTA detection is both sensitive and reliable (see Figure 4.5).

It was of interest to study the circular dichroism (CD) of the free OTAA36 compared to aptamer that is bound to OTA in hopes of observing an induced CD (ICD) band to provide further insight into the binding mode of OTA to OTAA36. Although no apparent ICD was found, the

153

1.2

1.0

0.8

0.6

0.4 Fraction of OTA bound ofOTA Fraction 0.2

0.0 0 0.2 0.4 0.6 0.8 1 1.2 [OTAA36] (µM)

Figure 4.5: The binding isotherm describing the affinity with which OTA binds to OTAA36. OTA (1 µM) was excited at λex 256 nm as increasing concentrations of OTAA36 (0 to 1.2 µM) were added to the solution.

spectra comparing free OTAA36 to that of the OTA:OTAA36 complex does demonstrate marked differences. In the trace that corresponds to only aptamer in solution (Figure 4.6, black trace), it is evident that an anti-parallel GQ is the dominant DNA structure in solution. This is characterized by positive peaks at 250 and 290 nm along with the presence of a negative signal around 260 nm.38

The positive peak occurring around 290 nm is noticeably broad. The broadness of this peak indicates the possibility of hybrid GQ structures occurring in this equilibrium.39 With the addition of equimolar amounts of OTA to OTAA36, although the presence of an anti-parallel GQ is still obvious, there are noticeable increases in peak intensity as well as decreases in peak widths in comparison to the free aptamer spectrum (Figure 4.6, red trace). Based on these results, it can be concluded that OTA does in fact bind to OTAA36 and that it will preferentially bind to the anti- parallel GQ conformation of the aptamer strand in solution.39 Additionally, there is evidence to

154 suggest that OTA facilitates the formation of the anti-parallel GQ. The increase in overall peak sharpness upon addition of OTA to the aptamer strand establishes that the toxin may play a role in inducing the preferred GQ geometry for binding to occur.

1.5

0.5 3 uM OTAA36

3uM OTA:OTAA36 CD (mdeg) CD 0 230 250 270 290 310 330 350 complex

-0.5

-1 Wavelength (nm)

Figure 4.6: The CD spectra of 3 µM of OTAA36 (black trace) and equimolar amounts of OTA and OTAA36 (3 µM) to give the OTA:OTAA36 complex in 3 µM concentration (red trace) in OTA binding buffer obtained at 10 oC.

4.3.3. Limit of Detection and Limit of Quantitation Although the previous studies provide insight into the binding capability of OTAA36 for its target, it is critical with this work to demonstrate why using ET fluorescence emission of the

OTA:OTAA36 complex presents an attractive new detection method for OTA. To illustrate the effectiveness of the ET method, a limit of detection (LOD) and limit of quantitation (LOQ) were determined for free OTA and OTA bound to OTAA36 using a fluorescence spectrometer at an excitation wavelength of 256 nm and by monitoring emission at 425 nm.

155

Prior to constructing the calibration curve for the OTA:OTAA36 complex, to ensure that all of the OTA in solution would become bound to OTAA36 upon addition, the Kd equation:

[푂푇퐴][푂푇퐴퐴36] 퐾푑 = 0.7 µ푀 = [푂푇퐴: 푂푇퐴퐴36]푐표푚푝푙푒푥 was used. The assay was designed in such a way to ensure that there would be a known concentration of the toxin: aptamer complex in solution throughout.

When assessing the fluorescence emission of free OTA in solution as a function of its concentration, the linear range for the toxin was found to occur from 0.1 to 10 µM (Figure 4.7 A)).

In the case of the OTA:OTAA36 complex, the linear range decreases by an order of magnitude on either end of the spectrum and is found to occur between 0.01 to 1.0 µM (Figure 4.7 B)), This finding illustrates that the dynamic range for the complex occurs over a lower range of concentrations than that of free OTA in solution, indicating that the detection limit for OTA and the complex as a whole should decrease as well.

The calibration curve obtained by monitoring the increase in fluorescence emission at 425 nm when using an excitation wavelength of 256 nm for the OTA:OTAA36 complex demonstrates great correlation with an r2 value of 0.99 (Figure 4.8). When comparing the LOD of free OTA in solution to that of the toxin aptamer complex, it is observed that relying on the fluorescent emission of OTA alone provides a detection limit of 9.6 nM where utilizing ET fluorescence of the

OTA:OTAA36 complex results in an LOD of 1.6 nM. Similarly, LOQ values for free OTA and for the OTA:OTAA complex are calculated to be 31.9 and 5.3 nM respectively. In the case of both LOD and LOQ, addition of the aptamer strand results in a 6-fold decrease in the levels of

OTA that can detected by this method (Table 4.1).

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3 3

2.5 2.5 2 Linear Range Linear Range 2 1.5 1.5

1 (au) (au) 1 0.5 0 0.5 -4 -2 0 2 -0.5 0

-4 -2 0 2 Log Fluorescence Intensity Intensity Fluorescence Log Log Fluorescence Intensity Intensity Fluorescence Log -1 Log [OTA] (µM) Log [OTA:OTAA36] (µM)

Figure 4.7: A) Linear range determination for free OTA using λex 256 nm. The log of OTA concentration is plotted against the log of the fluorescence emission intensity. The linear range is found from 0.1 to 10 µM of free toxin. B) Linear range determination for OTA:OTAA36 using λex 256 nm. The log of OTA:OTAA36 complex is plotted against the log of the fluorescence emission intensity. The linear range is found from 0.01 to 1 µM of toxin: aptamer complex

180

160

140

120 y = 162.01x + 4.91

100 R² = 0.99

20 Fluorescence Emission Intensity (au) EmissionIntensity Fluorescence 0 0 0.2 0.4 0.6 0.8 1 1.2 [OTA:OTAA36] complex (µM)

Figure 4.8: Calibration curve obtained by plotting the concentration of OTA:OTAA36 complex against fluorescence emission intensity at 430 nm.

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Table 4.1: Comparison of the fluorescence data obtained for free OTA and the OTA:OTAA36

Parameter

Linear Range LOD LOQ (µM) (nM) (nM)

Free OTA 0.1 - 10 9.6 31.9

OTA:OTAA36 0.01 - 1 1.6 5.3

What is important to note is that this aptamer based assay provides a detection limit for the

OTA:OTAA36 complex that corresponds to a concentration of 0.6 µg/L of toxin. The maximum allowable concentration of OTA in wine samples as set by the European Commission is 2 µg/L.6,40

The LOD demonstrated by this assay is well below this allowable concentration of OTA. This platform presents an attractive alternative method for the detection of OTA as compared to more traditional approaches as it boasts a sensitive detection limit which is, in most instances, lower than the allowable levels of OTA prescribed for a variety of consumer goods. In addition, this assay relies on the inherent fluorescence of OTA which makes the platform elegantly simple while maintaining cost efficiency.

4.3.4. Selectivity and Specificity of the Assay With the improvements observed for the levels of OTA that can be detected using the aptamer-ET fluorescence method, it is important to demonstrate that not only does this assay provide a sensitive approach to OTA detection but that it is also a selective and specific method.

To illustrate the specificity of the assay, two negative control experiments were employed. The first of which was the fluorescence titration of OTAA36 with Ochratoxin B (OTB) using the ET excitation wavelength of 256 nm. The structures of OTA and OTB are not dissimilar, the only difference between the two being the absence of the para chlorine atom of the phenol function in

158

OTB (Figure 4.9). This small structural difference has demonstrably large effects on the aptamer’s ability to bind to OTB.12 The OTA aptamer has a 100-fold lower affinity for OTB than for the toxin it was selected to bind to, OTA.12 The second control experiment was the fluorescence titration of OTA with an alternative anti-parallel GQ forming aptamer.41,42 The aptamer used in this case was the 15mer oligonucleotide that has been selected to bind to thrombin protein, namely the thrombin binding aptamer (TBA).41 These control experiments demonstrate that OTAA36 is selective to OTA only and that OTA will not simply bind to any anti-parallel GQ.

Figure 4.9: The molecular structure of OTB.

In the titration of OTB with OTAA36, there are no observable changes in the excitation spectrum as increasing amounts of aptamer are added to OTB in solution. Most notably, there is no emergence of a peak centered around 260 nm, demonstrating that no ET is occurring between the aptamer and the toxin (Appendix C). This clearly indicates that OTB does not effectively bind to OTAA36 to give a fluorescence response using the ET excitation parameters. Similarly, in the titration of OTA with TBA, there are no differences in the fluorescence spectra when TBA is added to OTA in solution, again indicating that the toxin does not bind to this aptamer (Appendix C).

When comparing the relative fluorescence intensity (Irel) values for these control titrations with the titration of OTAA36 against OTA values of -1.3 for the OTB:OTAA36 titration and -1.2 for the OTA:TBA titration are obtained. In contrast, the Irel value determined for the emission of

159

OTA when titrating against OTAA36 using the ET excitation wavelength of 256 nm is found to be 3.0 (Figure 4.10). The differences in Irel values for the control experiments in comparison to the value obtained for the OTA:OTAA36 titration further justify the conclusion that OTAA36 is selective to OTA and that OTA will only bind specifically to the GQ of the 36mer aptamer.

3.5 3.0 3 2.5 2 1.5 1 0.5 0 -0.5 -1

-1.5 -1.3 -1.2 Relative Fluorescence Intensity (au) Intensity Fluorescence Relative

OTB-OTAA36 OTA-TBA15 OTA-OTAA36

4.4. Conclusion To the best of our knowledge, we present for the first time a label free turn on fluorescence aptasensor platform for the detection of OTA. Upon binding of OTA to OTAA36, ET can occur between the GQ of the aptamer and the target toxin. When exciting at a wavelength of 256 nm, a

3-fold enhancement of visible emission of the OTA:OTAA36 complex at 425 nm is observed attributed to ET from the DNA to OTA. Fluorescence titration data and CD spectroscopy indicate that OTA does in fact bind to the anti-parallel GQ secondary structure of OTAA36.

160

When constructing a calibration curve for the OTA:OTAA36 complex in solution the LOD is determined to be 1.6 nM or 0.6 µg/L. Additionally, the LOD and LOQ values for the toxin:aptamer complex experience a 6-fold improvement in comparison to free OTA in solution.

Not only does this ET fluorescence assay provide a sensitive and simple means for OTA detection, the assay is also proven selective and specific for OTA in comparison to two negative control experiments.

This ET fluorescence-aptamer based assay for the detection of OTA provides a robust, simple, cost effective and efficient methodology to detect this molecule of interest. The fact that this assay is label free and capitalizes on the inherent fluorescence of OTA while providing a turn on fluorescence signal makes it a straight forward and tremendously appealing protocol for the detection of OTA. While maintaining sensitive detection limits, it is our hope that this new protocol will be of use and an important new method in the future of OTA detection and that this method will have implications within industrial as well as government and regulatory sectors going forward.

161

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Appendix A: General Methods

165

The majority of the methods used for these studies have been outline in detail within each chapter.

The more general procedures and equipment used follow within this appendix.

Materials: Commercial compounds were used as received and were purchased from either Sigma-

Aldrich Ltd or from Fisher Scientific. Oligonucleotides were obtained from Sigma-Aldrich. They were synthesized and purified by reverse phase cartridge purification (RP1) or by polyacrylamide gel electrophoresis (PAGE). All buffers were prepared using high-purity water, obtained from an in-house reverse osmosis filtration system (18 MΩ).

Equipment: All NMR spectra (1H and 13C) were obtained through the use of a Bruker Avance 300 spectrometer. All samples were run in DMSO-d6 and all spectra were referenced to this solvent.

Experiments were performed at room temperature and processed using TopSpin 2.1 software. All pH measurements were made using a Thermo Orion 320 Basic PerpHect LogR pH meter. UV-vis spectra were obtained using a Cary 300-Bio Spectrometer, fit with a 6×6 multicell block with

Peltier thermal control and stirring. Fluorescence experiments were performed using a Cary

Eclipse Fluorescence Spectrophotometer, fit with a 1×4 multicell block and Pelletier thermal control and stirring.

Optical Measurements: Stock solutions of the dyes were made to concentrations of 2 mM or 4 mM in DMSO and used as prepared in all optical studies. All measurements were made a room temperature. UV-Vis spectra were obtained from 800 nm to 200 nm. Fluorescence spectra were obtained starting from a wavelength 10 nm past the excitation wavelength used to a final wavelength of 800 nm, unless otherwise states. Excitation spectra were obtained from 200 nm to the 5 nm before the emission wavelength, unless otherwise noted.

166

Oligonucleotide Quantification: Oligonucleotides were quantified using UV-Vis absorbance spectroscopy, using the extinction coefficient provided by: http://www.idtdna.com/analyzer/applications/ oligoanalyzer, analyzed at 260 nm. A minimum of three absorbance readings were taken. Beer-Lambert Law was used to solve for the concentration of the oligonucleotide solution for each reading. The average of the calculated concentrations was taken and used as the stock oligo concentration.

Quantification of titrants: Titrants (OTA, OTB, bovine thrombin) were quantified using UV-Vis absorbance spectroscopy, using extinction coefficients provided by the supplier:

OTA ε= 6400 M-1 cm-1 at 333 nm in methanol, OTB ε= 6900 M-1 cm-1 at 318 nm in methanol and bovine thrombin ε= 72 150 M-1 cm-1 at 280 nm in water.

A minimum of three absorbance readings were taken. Beer-Lambert Law was used to solve for the concentration of the titrant for each reading. The average of the calculated concentrations was taken and used as the stock titrant concentration.

Mass Spectrometry Analysis: High-resolution mass spectrometry experiments for novel compounds were performed using an Agilent Q-TOF instrument, using an electrospray ionization source, monitoring for ions in the positive mode. Experiments were performed by Dr. Armen

Charcoglycan of the Advanced Analysis Center at the University of Guelph.

Circular Dichroism Spectroscopy: CD spectra were performed on a Jasco J-815 CD spectrophotometer equipped with a thermal controlled 1×6 multicell block. Measurements were made at 20 oC in quartz cells (110-QS) unless otherwise stated. Spectra were obtained using a light path of 1 nm, monitored from 500 nm to 200 nm at a bandwidth of 1 nm and scanning speed of 100 nm/min. A minimum of four scans were averaged to give the spectra as shown. Data was

167 corrected against a blank measurement of the appropriate buffer and smoothed using Jasco software using at least 15 convolutions unless otherwise noted.

Kd Determination: Kd values were determined by plotting the fraction of analyte bound as a function of DNA or aptamer concentration in solution, determining the fraction of analyte bound using the equation:

퐹 − 퐹 퐹푟푎푐푡푖표푛 퐵표푢푛푑 = 표푏푠 푖푛 퐹푚푎푥 − 퐹푖푛

Where Fobs is the observed fluorescence emission intensity of the analyte at a specific concentration of DNA/aptamer in solution, Fin is the fluorescence emission intensity of the analyte in the absence of DNA and Fmax is the fluorescence emission intensity of the analyte at the highest concentration of DNA added. A plot of the fraction of bound analyte versus the concentration of DNA/aptamer generated a binding isotherm that was analyzed with SigmaPlot 12.0 using the one site saturation binding macro to obtain the Kd values, using the equation:

퐵 푥 푦 = 푚푎푥 퐾푑 + 푥

Where y represents the fraction of dye bound, x, the concentration of DNA/aptamer and Bmax the maximum fraction of analyte able to bind the DNA in solution.

168

Appendix B: 1H and 13C NMR Characterization of Synthesized Products

169

Figure AB-1: 300 MHz 1H NMR of 3-(3-hydroxypropyl)-2-methylbenzothiazolium bromide (1) in DMSO-d6

Figure AB-2: 300 MHz 13C NMR of 3-(3-hydroxypropyl)-2-methylbenzothiazolium bromide (1) in DMSO-d6

170

Figure AB-3: 300 MHz 1H NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOH Btz, 5a) in DMSO-d6

Figure AB-4: 300 MHz 13C NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOH Btz, 5a) in DMSO-d6 171

Figure AB-5: 300 1H NMR of 2-[2-(4-methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOMe Btz, 5b) in DMSO-d6

Figure AB-6: 300 MHz 13C NMR of 2-[2-(4-methoxyphenyl)ethenyl]-3-hydroxypropyl benzothiazolium bromide (PhOMe Btz, 5b) in DMSO-d6 172

Figure AB-7: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-4-methylpyridinium bromide (2) in DMSO-d6

Figure AB-9: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-4-methylpyridinium bromide (2) in DMSO-d6

173

Figure AB-8: 300 MHz 1H NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOH P, 6a) in DMSO-d6

Figure AB-10: 300 MHz 13C NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOH P, 6a) in DMSO-d6

174

Figure AB-11: 300 MHz 1H NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOMe P, 6b) in DMSO-d6

Figure AB-12: 300 MHz 13C NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl pyridinium bromide (PhOMe P, 6b) in DMSO-d6

175

Figure AB-13: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-4-methylquinolium bromide (3) in DMSO-d6

Figure AB-14: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-4-methylquinolium bromide (3) in DMSO-d6

176

Figure AB-15: 300 MHz 1H NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q, 7a) in DMSO-d6

Figure AB-16: 300 MHz 13C NMR of 4-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOH Q, 7a) in DMSO-d6

177

Figure AB-17: 300 MHz 1H NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOMe Q, 7b) in DMSO-d6

Figure AB-18: 300 MHz 13C NMR of 4-[2-(4-methoxyphenyl)ethenyl]-1-hydroxypropyl quinolium bromide (PhOMe Q, 7b) in DMSO-d6 178

Figure AB-19: 300 MHz 1H NMR of 1-(3-hydroxypropyl)-2,3,3-trimethylindolenium bromide (4) in DMSO-d6

Figure AB-20: 300 MHz 13C NMR of 1-(3-hydroxypropyl)-2,3,3-trimethylindolenium bromide (4) in DMSO-d6

179

Figure AB-21: 300 MHz 1H NMR of 2-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl indolineuim bromide (PhOH Ind, 8) in DMSO-d6

Figure AB-22: 300 MHz 13C NMR 300 MHz of 2-[2-(4-hydroxyphenyl)ethenyl]-1-hydroxypropyl indolineuim bromide (PhOH Ind, 8) in DMSO-d6

180

Appendix C: Additional Data

181

ET Emission for Htelo: Merocyanine Dye Titrations

40 35 30 25 20 15 10 5 0 Fluorescene Fluorescene Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figue AC-1: Energy transfer excitation and emission spectra for PhP in 50 mM Tris buffer with the addition of increasing concentrations of Htelo.

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figure AC-2: Energy transfer excitation and emission spectra for PhP in 50 mM Tris with 50 mM Na+ with the addition of increasing concentrations for Htelo.

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figure AC-3: Energy transfer excitation and emission spectra for PhP in 50 mM Tris with 50 mM K+ with the addition of increasing concentrations of Htelo.

182

70 60 50 40 30 20 10 0 Fluoresence Fluoresence Intensity(au) 200 300 400 500 600 700 800 Wavelength (nm)

Figure AC-4: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with the addition of increasing concentrations of Htelo.

80 70 60 50 40 30 20 10 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 800 Wavelength (nm)

Figure AC-5: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with 50 mM Na+ with the addition of increasing concentrations of Htelo.

120 100 80 60 40 20 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 800 Wavelength (nm)

Figure AC-6: Energy transfer excitation and emission spectra for PhQ in 50 mM Tris buffer with 50 mM K+ with the addition of increasing concentrations of Htelo.

183

350 300 250 200 150 100 50 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figure AC-7: Energy transfer excitation and emission spectra for PhBtz in 50 mM Tris buffer with the addition of increasing concentrations of Htelo.

120 100 80 60 40 20 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figure AC-8: Energy transfer excitation and emission spectra for PhBtz in 50 mM Tris buffer with 50 mM Na+ with the addition of increasing concentrations of Htelo.

400 350 300 250 200 150 100 50 0

Fluorescence Fluorescence Intensity(au) 200 300 400 500 600 700 Wavelength (nm)

Figure AC-9: Energy transfer excitation and emission spectra for PhBtz in 50 mM tris buffer with 50 mM K+ with the addition of increasing concentrations of Htelo.

184

Kd Binding Curves for Htelo: Merocyanine Dye Titrations

1.2

0.8

0.6 50 mM Tris with K+ 50 mM Tris 0.4

50 mM Tris with Na+ Fraction Dye Bound FractionDye

0.2

0 0 1 2 3 4 5 [Htelo] (µM)

Figure AC-10: Binding isotherms for PhP with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+.

1.2

0.8

0.6 50 mM Tris 50 mM Tris with K+ 0.4

50 mM Tris with Na+ FractionDye Bound

0.2

0 0 1 2 3 4 5 [Htelo] (µM)

Figure AC-11: Binding isotherms for PhQ with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+.

185

1.2

0.8

0.6 50 mM Tris 50 mM Tris with K+ 0.4

50 mM Tris with Na+ FractionDye Bound

0.2

0 0 1 2 3 4 5 [Htelo] (µM)

Figure AC-12: Binding isotherms for PhBtz with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+.

1.2

0.8

0.6 50 mM Tris 50 mM Tris with K+ 0.4

50 mM Tris with Na+ FractionDye Bound

0.2

0 0 1 2 3 4 5 [Htelo] (µM)

Figure AC-13: Binding isotherms for ThBtz with Htelo in (a) 50 mM Tris buffer, (b) 50 mM Tris buffer with 50 mM Na+ and (c) 50 mM Tris buffer with 50 mM K+.

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300

250

200

150 y = 36.994x + 5.8141 R² = 0.9982 100

50 Fluoresence Fluoresence Emission Intensity(au)

0 0 1 2 3 4 5 6 7 8 [OTA] (µM)

AC-14: Calibration curve for free OTA using λex of 256 nm.

500

450

400

350

300

250

200

150

100

Fluorescence Intensity (au) Intensity Fluorescence 50

0 200 250 300 350 400 450 500 Wavelength (nm)

Figure AC-15: Negative control titration of OTAA36 with OTB.

187

400

350

300

250

200

150

100 Fluorescence Intensity (au) Intensity Fluorescence 50

0 200 250 300 350 400 450 500 Wavelength (nm) Figure AC-16: Negative control titration of OTA with the thrombin binding aptamer (TBA).

188