Design of Sensors for Highly Phosphorylated Natural Compounds

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Design of sensors for highly phosphorylated natural compounds

Dissertation

Submitted for the Degree

Doctor rerum naturalium (Doctor rer. nat.)

at the Faculty of Chemistry and Pharmacy

University of Freiburg by

Rahel Hinkelmann

Rheinfelden (Baden)

2020 Vorsitzender des Promotionsausschusses: Prof. Dr. Stefan Weber Dekan: Prof. Dr. O. Einsle Referent/in: Prof. Dr. H. J. Jessen Korreferent/in: Prof. Dr. M. Müller Datum der mündlichen Prüfung: 30.10.2020 Abstract

Highly phosphorylated metabolites, e.g. inositol polyphosphates (InsPs), diphosphoinositol phosphates (PP-InsPs), inorganic polyphosphate (polyP) and Magic spot nucleotides (MSN) regulate various biological processes. Their distinctive role in biology remain still elusive particularly concerning their dynamic turnover. The investigation of their contributions in metabolic processes asks for the development of, in vivo and in vitro, real time analytical detection. In this work the design, synthesis and evaluation of a set of new ﬂuorescent chemosensors is presented, since inositol polyphosphates and polyP are lacking an intrinsic chromophore. The synthesized chemosensors can be divided into the following three categories - Pyrene-excimer based, disassembly approach and DAPI derivatives. A 4’,6-diamidin-2-phenylindol (DAPI) synthesis was developed in this work. The photophysical properties of the respective set of synthesized chemosensors in combination with the above mentioned phosphorylated metabolites was evaluated. Fe3+-Salen and PyDPA was successfully used as a polyacrylamide gel electrophoresis (PAGE) stain and the latter dye was applied as a post-column staining reagent for Ins(1,2,3,4,5,6)-hexakisphosphate

(InsP6) on an ion chromatography system. In addition, the investigation and application of a new 31P-NMR based method using a chiral solvating agent revealed the stereoselectivity of the ﬁrst described naturally occurring 1-phytase. Zusammenfassung

Hoch phosphorylierte Metabolite, z.B InsPs, PP-InsPs, polyP und MSN regulieren ver- schiedene biologische Prozesse. Ihre besondere Rolle in der Biologie ist jedoch nach wie vor unklar, insbesondere was ihren dynamischen Umsatz betrifft. Die Untersuchung ihrer Beiträge zu Stoffwechselprozessen erfordert die Entwicklung einer in vivo und in vitro Echtzeitdetektion. In dieser Arbeit wird das Design, die Synthese und die Auswertung einer Reihe neuer fluoreszierender Chemosensoren vorgestellt. Die synthetisierten Chemosensoren können in die folgenden drei Kategorien unterteilt werden - Pyren-Excimer-basiert, «disassembly approach»und DAPI-Derivate. In dieser Arbeit wurde eine DAPI Synthese entwickelt. Die photophysikalischen Eigen- schaften des jeweiligen Sets der synthetisierten Chemosensoren in Kombination mit den oben genannten phosphorylierten Metabolite wurden diskutiert. Salen-Fe3+ und PyDPA wurden erfolgreich als PAGE-Färbung verwendet, und der letztgenannte Farbstoff konn- te als Nachsäulen-Färbereagenz für InsP6 auf einem Ionenchromatographiesystem ange- wendet werden. Darüber hinaus ergab die Anwendung eines neuen 31P-NMR basierten Verfahrens unter Verwendung eines chiralen Solvatisierungsmittels die Stereoselektivität der ersten beschriebenen natürlich vorkommenden 1-Phytase. Acknowledgements

First of all, I want to thank Prof. Dr. Henning Jessen for the opportunity to be part of this exciting research from the very beginning in Freiburg. We started in empty laboratories and they were inspiring and productive years in which the group accomplished a great deal. I learned a lot from you about positive thinking, staying motivated and solving problems, in science and beyond. Thank you for your constant support, helpfull discussions and being such a great supervisor in the last years.

I am grateful to Prof. Dr. M. Müller and Prof. Dr. P. Kurz for being part of my PhD committee.

Furthermore, I want to thank the staﬀ of the department of chemistry and pharmacy of the Albert-Ludwigs University. Especially Dr. M. Keller for his help regarding NMR and Dr. C. Warth for the HRMS measurements.

Also, I want to thank Regine Schandera and Dr. Richard Krieger for the constant support during the last years. Especially to Regine for being the good soul of the group.

Many thanks goes to all those who worked with me on this thesis: Larissa Pfennig and Paul Ebensperger for their support with the DAPI project. Ann-Kathrin Mündler for being the best Master student and the fun we had all the time. Alexander Ripp and Sophia Rauscher for their support in the Excimer project. Last but not least I have to express my deepest gratitude to Stephan Mundinger. Thank you for your invaluable help.

v Special thanks go to Tamara Bittner, Verena Eisenbeis Nikolaus York, Dr. Jyoti Singh and Kathrin Hönerlage for proof-reading parts of my thesis.

For the help in endless LaTeX questions I would like to say a big thank you to Kevin Ritter and Nicole Steck.

A heartfelt thank you goes to all AK Jessen members, former and present. Dr. Jyoti Singh, Dr. Christopher Wittwer which whom I started this work years ago. Thomas Haas, Paul Ebensperger, Tobias Dürr, Isabel Pruker, Alexander Ripp, Dr. Danye Qui, Markus Häner, Sandra Moser for the good times in the lab. I enjoyed the time with you so much.

I want to thank Paraskevi Fouka, Verena Eisenbeis, Jiahui Ma and Xuang Wang for the enjoyable company and conversations during lunch time.

Also, I want to thank the friday evening group Tamara Bittner, Stephan Mundinger, Kevin Ritter, Nikolaus Jork and Dominik Bezold for the good times we had and the countless discussions about chemistry and more.

Most importantly, I want to thank my friends and family for supporting me during the last years. Especially, my mother Andrea Hinkelmann and my father Horst Hinkelmann. Thank you for being with me for the whole journey and for always believing in me. A very special thanks goes to my love Claudio Werner. I cant ﬁnd enough words to thank you for your support inside and outside the lab. And for all the great food!

vi Contents

Abbreviations vii

1. General Introduction 1 1.1. Biological importance of phosphates ...... 2 1.1.1. Myo-inositol phosphates ...... 5 1.1.1.1. Myo-inositol...... 5 1.1.1.2. Inositol polyphosphates ...... 5 1.1.1.3. Biosynthesis of inositol polyphosphates ...... 7 1.1.1.4. Diphosphoinositol phosphate signalling ...... 9 1.1.1.5. IP6K1 as a drug target ...... 10 1.1.2. Inorganic polyphosphate (polyP) ...... 12 1.1.2.1. Biosynthesis of polyP ...... 12 1.1.2.2. PolyP signalling ...... 13 1.1.2.3. PPK as a drug target ...... 13 1.1.2.4. Phosphates in wastewater ...... 14 1.1.3. Magic Spot Nucleotides (MSN) ...... 15 1.1.3.1. Biosynthesis of (p)ppGpp ...... 16 1.1.3.2. (p)ppGpp signalling ...... 17 1.1.3.3. Rel as a drug target ...... 19 1.2. A brief history of ﬂuorescence ...... 20 1.3. Supramolecular chemistry ...... 21 1.3.1. Fluorescent chemosensors ...... 22

i Contents

1.3.2. Anion sensing ...... 23 1.3.3. Ratiometric fluorescent probes ...... 23 1.3.3.1. Monomer/Excimer-based fluorescent probes ...... 24 1.3.3.2. ESIPT-based fluorescent probes ...... 25 1.3.4. Overview of chemosensors for myo-inositol polyphosphates, polyP and Magic Spot Nucleotides ...... 27 1.3.4.1. Chemosensor for myo-inositol polyphosphates ...... 27 1.3.4.2. Chemosensor for inorganic polyP ...... 32 1.3.4.3. Chemosensor for Magic Spot Nucleotides ...... 33 1.3.5. References...... 35

2. State of the Art - Analytics 43 2.1. Radioactive labelling ...... 44 2.1.1. Inositol polyphosphates ...... 44 2.1.2. Magic Spot Nucleotides ...... 45 2.2. Nuclear Magnetic Resonance Spectroscopy (NMR) ...... 45 2.2.1. 31P-Nuclear Magnetic Resonance Spectroscopy ...... 46 2.2.1.1. Inorganic Polyphosphate (polyP) ...... 46 2.2.1.2. Inositol polyphosphates ...... 47 2.2.2. 13C-Nuclear Magnetic Resonance Spectroscopy ...... 48 2.2.2.1. Inositol polyphosphates ...... 48 2.3. EnzymeAssays ...... 49 2.3.1. Inorganic Polyphosphate (polyP) ...... 49 2.4. Electrophoresis ...... 50 2.4.1. Polyacrylamide Gel Electrophoresis (PAGE) ...... 51 2.4.2. Inorganic Polyphosphate (polyP) ...... 51 2.4.3. Inositol polyphosphates ...... 52 2.4.4. Capillary Electrophoresis (CE) ...... 54 2.4.5. Inositol polyphosphates ...... 55 2.4.6. Inorganic polyphosphate (polyP) ...... 55 ii Contents

2.5. Chromatography ...... 55 2.5.1. Ionchromatography(IC)...... 55 2.5.2. Post column detection methods ...... 59 2.5.3. References...... 62

3. Goals of the Thesis 67

4. Excimer based sensing 71 4.1. Background ...... 72 4.1.1. Excimer ﬂuorescence based chemosensors ...... 72 4.2. Results and Discussion ...... 74 4.3. Synthesis of excimer based chemosensors ...... 74 4.3.1. Tripodal chemosensors ...... 74 4.3.2. PyDPA chemosensor ...... 77 4.3.3. Linked chemosensor - Intramolecular excimer formation ...... 78 4.4. Sensor evaluation ...... 82 4.4.1. Photophysical properties ...... 82 4.4.1.1. Tripodal chemosensors ...... 83 4.4.1.2. Linked chemosensors ...... 86 4.4.2. PAGE-Gel staining ...... 89 4.4.3. Post column derivatisation - Ion Chromatography ...... 91 4.5. Summary & Outlook ...... 97 4.6. Experimental ...... 101 4.6.1. Generalremarks ...... 101 4.6.2. Synthesis ...... 104 4.6.3. References...... 123

5. Sensing via a disassembly approach 125 5.1. Background ...... 126 5.1.1. Disassembly Approach ...... 126

iii Contents

5.2. Results and Discussion ...... 128 5.2.1. Synthesis of disassembly based fluorescent probes ...... 128 5.2.1.1. Salicylaldehyde-based fluorescent probes ...... 128 5.2.1.2. Naphthol-based fluorescent probes ...... 130 5.2.2. Excited state intramolecular proton transfer - ESIPT ...... 131 5.2.2.1. HBO-based fluorescent probes ...... 131 5.3. Sensor evaluation ...... 137 5.3.1. Photophysical properties ...... 138 5.3.1.1. Fe3+-Salen...... 138 5.3.1.2. HBO-based chemosensors ...... 139 5.3.1.3. Naphthol-based chemosensors ...... 144 5.3.1.4. PAGE-Gel staining ...... 145 5.4. Summary & Outlook ...... 148 5.5. Experimental ...... 151 5.5.1. Generalremarks ...... 151 5.5.2. Synthesis ...... 154 5.5.3. References...... 167

6. Sensing via hydrogen bonding 169 6.1. Background ...... 170 6.1.1. DAPI ...... 170 6.1.2. Hydrogen bonds in sensing ...... 171 6.2. Results and Discussion ...... 173 6.2.1. DAPI Synthesis ...... 173 6.2.2. DAPI analogue Synthesis ...... 178 6.2.2.1. DAPI isomer ...... 178 6.2.2.2. guanidine DAPI ...... 178 6.2.2.3. Elongated structures ...... 179 6.2.2.4. Biguanide moiety ...... 183

iv Contents

6.2.3. C-3 modiﬁed indole ...... 184 6.2.3.1. Koshland modiﬁcation ...... 185 6.2.4. Sensor evaluation ...... 186 6.2.4.1. Photophysical properties ...... 187 6.3. Summary & Outlook ...... 190 6.4. Experimental ...... 193 6.4.1. Generalremarks ...... 193 6.4.2. Synthesis ...... 195 6.4.3. References...... 207

7. Assignment of the stereoselectivity of a new phytase 211 7.1. Background ...... 212 7.1.1. Phytases...... 212 7.1.2. Xanthomonas campestris pv. vesicatoria (Xcv) ...... 213 7.2. Results and Discussion ...... 214 7.2.1. References...... 219

A. Analytical Data of Chapter 4 iii

B. Analytical Data of Chapter 5 xxv

C. Analytical Data of Chapter 6 xlvii

Abbreviations

ADP adenosine diphosphate. ATP adenosine-5’-triphosphate. Boc tert-butoxycarbonyl. CE capillary electrophoresis. CEC capillary electrochromatography. CGE capillary gel electrophoresis. cyclen 1,4,7,10-Tetraazacyclododecane. CZE capillary zone electrophoresis. DABCO 1,4-diazabicyclo[2.2.2]octane. DAPI 4’,6-diamidin-2-phenylindol. DIPEA N,N -diisopropylethylamine. DIPP diphosphoinositol polyphosphate phosphohydrolase. DMAP 4-(dimethylamino)pyridine. DNA deoxyribonucleic acid. DOSY diﬀusion-ordered spectroscopy. DPA dipicolylamine. dppf 1,1’-Bis(diphenylphosphino)ferrocen. E. coli Escherichia coli. EDTA ethylenediaminetetraacetic acid. EE energy expenditure.

vii Abbreviations

EOF electroosmotic ﬂow. eq. equivalent. ER endoplasmic reticulum. ESI electrospray ionization. ESIPT excited-state intramolecular proton transfer. FLD Fluorescence detector. GDP guanosine-5’-diphosphate. GTP guanosine-5’-triphosphate. h hour. HBO 2-(2’-hydroxyphenyl)benzoxazole. HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid. HPLC high performance liquid chromatogpraphy. HRMS high-resolution mass spectrometry. IC ion chromatography. INPP5 inositol polyphosphate 5-phosphatase.

Ins(1,4,5)P3 inositol-1,4,5-trisphosphate.

InsP2 inositol bisphosphate.

InsP3 inositol trisphosphate.

InsP5 Ins(1,3,4,5,6)-pentakisphosphate.

InsP6 Ins(1,2,3,4,5,6)-hexakisphosphate.

InsP7 diphosphoinositol pentakisphosphate. InsPs inositol polyphosphates. IP3K IP3 3-kinase. IP5K inositol pentakisphosphate 2-kinase. IP6K inositol hexakisphosphate kinase. IPMK inositol polyphosphate multikinase. IR infrared absorption spectroscopy. ITPK1 inositol tetrakisphosphate 1-kinase. LArgN L-arginine amide hydrochloride salt. viii Abbreviations

LDA lithium diisopropylamide. LHMDS lithium hexamethyldisilazane. LoD limit of Detection. MDD metal-dye detection. min minute. MSN magic spot nucleotides. NMR nuclear magnetic resonance. NOESY nuclear overhauser enhancement spectroscopy. PAGE polyacrylamide gel electrophoresis. PAR 4-(2-pyridylazo)resorcinol. PCSR postcolumn staining reagent.

Pd(dba)2 Palladium(0) bis(dibenzylideneacetone).

Pd(OAc)2 Palladium((II) acetate).

Pd(PPh3)4 Palladium-tetrakis(triphenylphosphine). PH pleckstrin homology.

Pi orthophosphate. PIPPS 1,4-Piperazinedipropanesulphonic acid. PKB protein kinase B. PLC phospholipase C. polyP inorganic polyphosphate.

PPi pyrophosphate. PP-InsPs diphosphoinositol phosphates. PPIP5K diphosphoinositol pentakisphosphate kinase. PPK1 polyphosphate kinase 1. PPK2 polyphosphate kinase 2. ppm parts per million. PPX1 exopolyphosphatase 1. PtdInsPs phosphatidylinositol polyphosphates. RNA ribonucleic acid.

ix Abbreviations

RNAP ribonucleic acid polymerase. RPT reverse proton transfer. rRNA ribosomal ribonucleic acid. SAX strong anion exchange. T2D type-2 diabetes. T3E type III effector. TB Tris/Borate. TBE Tris/Borate/EDTA. TBTU N,N,N ’,N ’-tetramethyl-O-(benzotriazol-1-yl)uronium tetrafluoroborate. TLC thin layer chromatography. TNP N2-(m-Trifluorobenzyl),N6-(p-nitrobenzyl)purine. Tris tris(hydroxymethyl)aminomethane. tRNA transfer ribonucleic acid. UV ultraviolet.

x CHAPTER 1

General Introduction

1 1. General Introduction

1.1. Biological importance of phosphates

Phosphates play an essential role in biochemistry and are ubiquitous in nature. In his paper from 1987, Westheimer posed the question ”Why nature chose phosphates” and gave some pioneering answers to this question.[1] Research was done on this topic showing that phosphate esters are the ideal structures to facilitate life as we know it.[1–3] Deoxyribonucleic acid (DNA), consisting of sequentially phosphodiester linked nucleo- sides (Scheme 1.1, right), stores the genetic information of all organisms on earth. Enzymes, regulating and catalysing chemical processes which are crucial for life, are commonly controlled by phosphorylation or dephosphorylation (Scheme 1.1, middle).[4] This type of post-translational modiﬁcation diversiﬁes the chemical nature of proteins. It is estimated that at any given time, one third of the proteome is phosphorylated.[4]

How can phosphate be that versatile? Phosphorus is a group 15 element, thus bearing ﬁve electrons in its outer shell. Consequently, phosphorus can form ﬁve covalent bonds.

3- With four oxygen atoms orthophosphate (PO4 ) is formed which is highly water soluble. Of importance are the pKa values of phosphate and its esters (see Scheme 1.1) which lie for all forms approximately at 2. This ensures negative charge at pH 7, the physiological intracellular pH, resulting in good water solubility and localization inside cell compartments due to hindered diﬀusion through lipid membranes. To retain, for example, nucleic acids within phospholipid membranes is pivotal for life. As outlined in Scheme 1.1 phosphate can be esteriﬁed and form mono-, di- or even tri-esters (not shown). Therefore, the phosphate moiety can serve as a linker of two compounds, for example, as it does in DNA. The formed esters are stable in aqueous solution and need enzyme activity to be cleaved.[2] Formation or cleavage of P-O bonds is essential for biochemical energy release (hydrolysis of adenosine-5’-triphosphate (ATP)) or protein regulation through phosphorylation and dephosphorylation.[3]

2 1.1. Biological importance of phosphates

Scheme 1.1: Inorganic phosphate and examples of biochemically relevant phosphates. Phosphoric acid, mono- and diester with its pKa values in water.

Phosphoryl transfer plays key roles in signalling, protein synthesis, energy transduction and maintaining the integrity of ribonucleic acid (RNA) and DNA.[3] The negative charge on the phosphate is repelling nucleophiles and with that prevents hydrolysis assuring a comparable stability to the organisms lifetime.

A substantial incidence for the importance of tight control of phosphorylation in cells is that abnormal changes in enzymes, which are responsible for phosphorylation and dephosphorylation, can have devastating eﬀects. Numerous human diseases can be a consequence, such as diabetes, type II obesity and diﬀerent forms of cancer.[3] The fact that phosphates play a fundamental role in the human body as well as in other organisms highlights the importance of an in-depth understanding of phosphorylated natural compounds and their metabolic pathways. A deep understanding of the processes might lead to targeted drug therapies and improvements in environmental aspects. An important player in this are the second messengers.

3 1. General Introduction

Second messengers are a fundamental signalling principle in biology. They are intracellular molecules which are released in response to extracellular stimuli - the ﬁrst messengers, for example hormones. They trigger signal cascades inside the cell ensuring signal transduction.

In the next three sections the biological background and importance of three phosphate containing metabolite classes are presented: Inositol phosphates, PolyPs and MSN. Scheme 1.2 shows an overview of the three compound classes. These molecules will serve as analytes of interest within this thesis.

N = O

O O O N NH O P O P O P O N N N NH 2- 2 OPO3 O O O O 2- 2- O3PO 2 OPO3 O O O 3 1 NH O O OH 2 4 6 O P O PO P O 2- 5 2- N O3PO OPO3 O O O O P O P O N 2- OPO3 O O n N N

InsP6 n = 3 - 1000 pppNpp Myo-inositol phosphates Inorganic polyP Magic spot nucleotides

Scheme 1.2: Examples of the three categories: InsP6 (for myo-inositol phosphates), PolyP and nucleoside-3’,5’-polyphosphate (pppNpp) (for Magic Spot Nucleotides).

4 1.1. Biological importance of phosphates

1.1.1. Myo-inositol phosphates

1.1.1.1. Myo-inositol

Their story has its origin in 1850 with Joseph Scherer who isolated an optically inactive cyclohexanol from muscle tissue.[5] He named it ”inosit”, later extended to ”inositol” the umbrella term for all nine possible stereoisomeric members of this family. These stereoisomers can be distinguished depending on the position of the respective hydroxyl groups. Almost all of them occur in nature and the myo-inositol carbon scaﬀold is a well studied isomer (Scheme 1.3).[6]

OH OH HO OH HO 3 1 4 3 4 6 HO OH 6 HO OH HO OH1 OH

Scheme 1.3: Chemical structure of myo-inositol (1) depicted in: left: natta and right: chair structure. Plane of symmetry is shown as red dotted line.

Nomenclature of these compounds has caused much confusion and therefore 1-D- configuration, with counter-clockwise numbering is used as a standard in literature since 1976, using the symbol ”Ins”.[7]. The number of phosphates bound to the inositol scaffold is indicated by a suffix after ”P”. In Scheme 1.3 a red dotted line, dissecting the 2- and 5-positions of myo-inositol (1) is indicating the plane of symmetry. Modification of the hydroxyl groups, except the 2 and 5 position, is causing the formation of enantiomers.

1.1.1.2. Inositol polyphosphates

Myo-inositol is the structural building block for InsPs: A multifaceted family of signalling molecules.

5 1. General Introduction

O O OH O O P HO O O O

2- O3PO OH 2- OPO3 PtdIns(4,5)P2 2

Scheme 1.4: Chemical structure of phosphatidylinositol polyphosphates (PtdInsPs) exempliﬁed with PtdIns(4,5)P2 (2)

In cells, inositol phosphates exist either in lipid bound form, called PtdInsPs (Scheme 1.4), or in their free water soluble, cytosolic form, called InsPs (Scheme 1.5).

Lipid bound PtdInsPs consist of a phospholipid (Figure 1.4, in red) anchored in a lipid bilayer and a myo-inositol head group pointing inside the cell. The myo-inositol head group is attached to the fatty acid through a phosphate diester at the 1-position (Phos- phatidylinositol). Phosphorylation of the inositol head group leads to phosphoinositides where 3-,4- and 5-hydroxyls can be involved in phosphorylation, resulting in a maximum of seven isomers (Example of phosphoinositide in Scheme 1.4).

Water soluble InsPs can form 63 possible isomers upon phosphorylation of the myo- inositol scaﬀold.[8] These can even be extended by attaching pyrophosphate moieties instead of monophosphates, leading to the formation of so-called PP-InsPs. Two major classes of cytosolic inositol phosphates can be distinguished1: InsPs and PP-InsPs (Scheme 1.5).

It was only in the 1980s, with the discovery of the signalling pathway of inositol-1,4,5- trisphosphate (Ins(1,4,5)P3) (3) and its role in cells, that research interest on inositol polyphosphates raised to a new level. Hormone stimulated hydrolysis of PtdIns(4,5)P2 [9] (2) yields (within 5 seconds) diacylglycerol and Ins(1,4,5)P3 (3). It is shown that the released 3 acts as a second messenger that mobilizes intracellular calcium stores from the endoplasmic reticulum (ER).[9,10]

1The term ”inositol polyphosphates” will be used in the present context as a generalization term including both classes of cytosolic inositol phosphates (InsPs & PP-InsPs).

6 1.1. Biological importance of phosphates

Inositol polyphosphates Diphosphoinositol phosphates

OPO 2- O O 3 P 2- 2- O O3PO OPO3 2- 2- O O OPO3 OPO3 P 2-O PO OPO 2- 2-O PO O O 2- 2- 3 3 3 O3PO OPO3 2- OPO3 2- 2- 2- 2- O3PO OPO3 O3PO OPO3 Inositol-(1,2,3,4,5,6)-hexakisphosphate O O 4 O P O O P O O O OH O P O O P O 2- HO OPO3 O O

2- 5-diphosphoinositol-pentakis-phosphate 1,5-bis-diphosphoinositol-tetrakis-phosphate O3PO OH 2- OPO3 5 6

Inositol-(1,4,5)-triphosphate 3

Scheme 1.5: Chemical structure of prominent examples of inositol phosphates: InsP6 (4) and inositol trisphosphate (InsP3)(3) and diphosphoinositol phosphates: 5-PP-InsP5 (5) and 1,5-InsP8 (6).

Since the discovery of 3 as a second messenger many soluble inositol phosphates and their role in metabolism have been discovered and studied.[8] Inositol polyphosphates are found in organisms ranging from slime moulds, fungi and plants to mammals.[11] One interesting question is:

How do we beneﬁt from the metabolic diversity of inositol polyphosphates and their regulating enzymes?

1.1.1.3. Biosynthesis of inositol polyphosphates

For answering this question, one needs ﬁrst to understand the biosynthesis of inositol polyphosphates inside mammalian cells and the enzymes involved.[8,12–15] In general, biosynthesis in mammals can be divided into two major pathways. The ﬁrst pathway comprises synthesis of PP-InsPs precursor Ins(1,3,4,5,6)-pentakisphosphate

(InsP5)(7)(Scheme 1.6). Starting with hormone or growth factor mediated hydrolysis of PtdIns(4,5)P2 (2) by phospholipase C (PLC) Ins(1,4,5)P3 (3) is released into the cy- [8] [16] tosol. Released 3 is phosphorylated by IP3 3-kinase (IP3K) to Ins(1,3,4,5)P4 (8) and subsequently converted to InsP5 (7) via inositol polyphosphate 5-phosphatase (INPP5) and inositol tetrakisphosphate 1-kinase (ITPK1) or sequentially converted to 7 by inos-

7 1. General Introduction

[17] itol polyphosphate multikinase (IPMK). 7 is the precursor of InsP6 (4) (also known as phytic acid), a key compound in PP-InsPs biosynthesis (shown in Scheme 1.7 as the second pathway. InsP6 is highlighted in red).

P OH OH HO OH HO P P P 1 PLC IPMK / IP3K 1

1 HO P P OH P OH

P P P

PtdIns(4,5)P2 Ins(1,4,5)P3 Ins(1,3,4,5)P4 2 3 8

O INPP5 P = O P O O IMPK

OH OH OH P P P P P P IPMK ITPK1 1 1 1

P P OH P P P OH OH P

Ins(1,3,4)P3 Ins(1,3,4,6)P4 Ins(1,3,4,5,6)P5 9 10 7

Scheme 1.6: First pathway: Synthesis of Ins(1,3,4,5,6)P5 from PtdIns(4,5)P2 (2) in mammals.

[18] Inositol pentakisphosphate 2-kinase (IP5K) converts InsP5 (7) to fully phosphorylated

InsP6 (4) (Scheme 1.7). Based on this conversion, inositol hexakisphosphate kinase [15] (IP6K) can add a β-phosphate at the 5-position to yield 5-PP-InsP5 (Scheme 1.7).

Additionally, IP6K converts InsP5 to 5-PP-InsP4 (11). In the same manner, diphosphoinositol pentakisphosphate kinase (PPIP5K) adds a phosphate to the 1-position. Both ways lead to the formation of eightfold phosphorylated 1,5-[PP]2-Ins(2,3,4,6)P4 (6) also referred to as 1,5-InsP8. The reverse reaction, hydrolysis of the pyrophosphate moiety, is regulated by diphosphoinositol polyphosphate phosphohydrolase (DIPP) accounting for the rapid turnover of PP-InsPs.[13,19] Intracellular levels of PP-InsPs in mammalian tissues are turning over at least 10 times

[19] every 40 minutes. Approximately 50 % of InsP6 and 20 % of InsP5 are converted to PP-InsPs per hour.[20]

8 1.1. Biological importance of phosphates

As inositol polyphosphates do not adsorb ultraviolet (UV) or emit visible wavelength light they are virtually undetectable by spectrophotometric or ﬂuorescence methods. This is the reason why studies, measuring the inositol phosphate turnover in intact cells, are carried out using e.g. [3H]inositol radioactive labelling followed by ion chromatography (IC) based separation.

OH P P P P P P P P 1 IP5K IP6K 1

P P DIPP P P P P 1

P P P P

Ins(1,3,4,5,6)P5 Ins(1,2,3,4,5,6)P6 5PP-Ins(1,2,3,4,6)P5 O 7 4 5 P = O P O O DIPP IP6K DIPP PPIP5K DIPP PPIP5K

O O P P = P P OH P P O O O P P O O P P P P P P 1 IP6K 1 1

P P P P DIPP P P P P P P P

5PP-Ins(1,3,4,6)P4 1PP-Ins(2,3,4,5,6)P5 1,5[PP]2-Ins(2,3,4,6)P4 11 12 6

Scheme 1.7: Second pathway: Synthesis of 5-PP-Ins(1,3,4,6)P4 (11), 1/5PP-Ins(1,2,3,4,6)P5 (5 and 12) sometimes referred to as 1/5-InsP7 and 1,5[PP]2-Ins(2,3,4,6)P4 (6) termed 1,5-InsP8 in mammals.

1.1.1.4. Diphosphoinositol phosphate signalling

Two principle mechanisms for inositol polyphosphate signalling are known today (Figure 1.1): protein-binding and pyrophosphorylation. 1.) Pyrophosphate bonds in PP-InsPs are estimated to possess similar free energy of hydrolysis to that of ATP. This high-energy β-phosphate group can be transferred to a pre-phosphorylated serine residue in a non-enzymatic manner, generating pyrophosphorylated proteins.[21–23]

9 1. General Introduction

2.) Inositol polyphosphates can bind to several proteins, for example proteins containing the pleckstrin homology (PH)-domain, regulating the catalytic activity or membrane translocation, or both.[24–26]

Figure 1.1.: Proposed mechanism of pyrophosphorylation and protein binding. Picture taken from reference. [27]

Returning finally to the above-posed question - how we can benefit from the metabolic diversity of inositol phosphates and their regulating enzymes - we can give the answer now with: By either inhibiting specific kinases or preventing pyrophosphorylation of proteins to suppress the downstream metabolic pathways.

1.1.1.5. IP6K1 as a drug target

[13] 5-InsP7 (5) is synthesized by IP6Ks in mammals (Scheme 1.7). This enzyme class consists of three members: IP6K1, IP6K2 and IP6K3, of which IP6K1 is the major isoform with tissue-speciﬁc distribution.[28,29]

IP6K1 is involved in various cellular processes, such as vesicular traﬃcking, chromatin remodelling, DNA damage and repair, haemostasis, spermatogenesis, cell-migration- mediated neuronal development and cancer metastasis, behavioural responses, neu- trophil functions, insulin signalling and energy metabolism.[11]

Protein kinase B (PKB), also known as Akt, is a kinase that plays a role in glucose metabolism. 5-InsP7 (5) is a physiological inhibitor of Akt and promotes insulin resistance.[30] N2-(m-Triﬂuorobenzyl),N6-(p-nitrobenzyl)purine (TNP) (Scheme 1.8) is a

IP6K inhibitor which reduces the intracellular 5-InsP7 levels.

10 1.1. Biological importance of phosphates

NH N O2N N

N N N H H

CF3 13

Scheme 1.8: Chemical structure of TNP a IP6K inhibitor.

Ghoshal et al. demonstrated in 2016 that TNP, injected prior to high fat diet-feeding, decelerates the beginning of diet induced obesity and insulin resistance in mice (Figure 1.2).[31] TNP induced inhibition of synthesis of 5 leads to an increased Akt activity which promotes global insulin sensitivity and enhances energy expenditure (EE) in adipose tissue, thus preventing the mice from diet induced obesity and type-2 diabetes (T2D).

With these findings the therapeutic significance of the PP-InsPs biosynthetic pathway in T2D and obesity moves into the spotlight. This example highlights the importance of effective, sensitive and reliable biochemical detection methods to investigate further targets for potential treatments.

Figure 1.2.: Graphic account of TNP mediated insulin sensitivity. left: IP6K1 generates 5-InsP7 from InsP6. Kinase Akt is inhibited by 5-InsP7 as well as EE in adipose tissue. Resulting in weight gain and decreased insulin sensitivity in mice on high fat diet. right: TNP inhibits IP6K1 and thus 5-InsP7 formation. Low levels of 5-InsP7 increase Akt activity promoting insulin sensitivity and EE. Picture taken from reference. [31]

11 1. General Introduction

1.1.2. Inorganic polyphosphate (polyP)

Another class of natural phosphoanhydrides is a polyanionic linear molecule called inorganic polyphosphate (polyP). PolyP is a polymer of up to thousands of orthophosphate

(Pi) residues which are connected by high-energy phosphoanhydride bonds (Scheme 1.9).

O O O OP O P O P O O O O n n = 3 - 1000s Inorganic polyP (PolyP)

Scheme 1.9: Chemical structure of inorganic polyphosphate (polyP)

PolyP was ﬁrst isolated by Liebermann from yeast in 1890[32]. The mounting evidence of essential metabolic functions of polyP decries the missing tools for detecting them. There is evidence about the biological importance of polyP depending on its localization. Amongst them are: substitution for ATP, capsule of bacteria and physiological adjustments to growth, development and stress.[33]

1.1.2.1. Biosynthesis of polyP

One enzyme participating in polyP synthesis is the bacterial polyphosphate kinase 1 (PPK1).[34] This enzyme catalyses the transfer of the γ-phosphate of ATP to extend the polyphosphate chain in bacteria (Figure 1.3). Polyphosphate kinase 2 (PPK2) is another kinase involved in polyP generation in bacteria.[35]

PPK1/2 PolyPn ATP/GTP PolyPn+1 ADP/GDP

Figure 1.3.: Biosynthesis and degradation of polyP by PKKs in bacteria.

Whereas PPK1 is speciﬁc to ATP, PPK2 uses guanosine-5’-triphosphate (GTP) and ATP equally well. Both enzymes are also able to catalyse the reverse reaction to degrade polyP.[36]

12 1.1. Biological importance of phosphates

Enzymes for polyP hydrolysis (phosphatases) have also been discovered in prokaryotes as well as in eukaryotes. In bacteria, one example is the exopolyphosphatase 1 (PPX1) which hydrolyses long-chain polyPs sequentially.[37] PPX1 is not limited to polyP but can also hydrolyse the alarmone guanosine pentaphosphate (pppGpp, see Figure 1.5 structure: 14 )

1.1.2.2. PolyP signalling

There is a connection between the accumulation of polyP in Escherichia coli (E. coli) and (p)ppGpp.2 The breakdown of polyP by exopolyphosphatase 1 (PPX1) is blocked by (p)ppGpp in nutritionally stressed cells resulting in extensive accumulation of polyP.[38] Although there are some theories about the polyphosphate metabolism in E. coli the exact mechanism is still unknown. In-depth understanding of the metabolic processes might lead to therapies with clinical signiﬁcance in future. One example leading in this direction is given in the next section.

1.1.2.3. PPK as a drug target

The high degree of sequence conservation of PPK1 in various bacterial species, including pathogens, such as Mycobacterium tuberculosis, Pseudomonas aeruginosa and Vibrio cholerae, has led to knockout studies which showed that PPK1 is essential for bacterial virulence.[39] Furthermore, the absence of PPK1 in eukaryotes suggests that it might be an attractive target for antimicrobial drugs.[39]

Pseudomonas aeruginosa is an representative of the Pseudomonas genus in human medicine. The pathogen causes infections in immunocompromised hosts, patients with cystic fibrosis, severe burns and wounds and is a colonizer of medical devices. Rashid et al. demonstrated that PPK1 is essential for various forms of motility[40], quorum sensing, development of biofilms and production of virulence factors.[41] Exopolysaccharides also called alginate, are synthesized by the bacteria forming the biofilm, in response to

2(p)ppGpp is the collective term for: guanosine-3’,5’-bis(diphosphate) (ppGpp) and guanosine-3’- diphosphate-5’-trisphosphate (pppGpp). For detailed information see subsection 1.1.3.

13 1. General Introduction environmental conditions. Bioﬁlms, and their resistance to antimicrobial agents, are the reason for the persistence of chronic diseases like cystic ﬁbrosis.[42]

Figure 1.4.: Scanning electron micrographs showing that mutants fail to produce comparable bioﬁlms to the wild type. Left: P. aeruginosa wild type. Right: PPK1 mutant. Picture taken from reference. [43]

Thus, to control bioﬁlm formation, PPK1 is a potential therapeutic target (Figure 1.4). The future inhibitor of PPK1 might enjoy a broad activity spectrum and little toxicity at the same time, because until now the enzyme has not been observed in mammalian cells.[39]

1.1.2.4. Phosphates in wastewater

Another aspect worth mentioning, with respect to polyP, is the environmental impact of orthophosphate. Phosphates have a particular importance as fertilizer in agriculture.[44] However, fertilizer run-oﬀ containing water is an issue. It results in increased algae blooms in bays, lakes and waterways.[39] The process is called eutrophication and represents a threat to the ecosystem.[45] Biological removal through aerobically activated

[46,47] bacteria, that take up the Pi and convert it to polyP, is one approach.

14 1.1. Biological importance of phosphates

1.1.3. Magic Spot Nucleotides (MSN)

Magic Spot Nucleotides (MSN) or alarmones play a role as regulatory signalling molecules in bacteria. Research on this compounds started in 1969 with the ﬁnding of Cashel and Gallant that an unknown spot appears on an 2D-thinlayer autoradiogram of phosphorylated

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

O O OH O P O P O O O 14 O

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

O O OH O P O P O O O 15 (a) (b) 32 3- Figure 1.5.: a) Autoradiograms of the amino-acid dependence of PO4 incorporation into diverse acid soluble metabolites in E. coli. Right: Under amino acid supplementation and left: amino acid starvation. [48] b) Chemical structure of guanosine-3’,5’-bis(diphosphate) (15) and guanosine-3’- diphosphate-5’-triphosphate (14) metabolites when E. coli cells were grown under amino acid starvation (Figure 1.5 a)). This spot is composed of two highly phosphorylated nucleotides, which were as- signed to guanosine-3’,5’-bis(diphosphate) (ppGpp (15)) and guanosine-3’-diphosphate- 5’-triphosphate (ppGppp (14)) (Figure 1.5 b)); Collectively known as (p)ppGpp. Since then, research was done on revealing (p)ppGpp’s role in bacteria. It was found that they are second messenger molecules which are regulating nutrient stress response also called ”stringent response”. Deprivation of nutrients leads to accumulation of (p)ppGpp and subsequently down-regulation the synthesis of DNA, stable RNAs, ribosomal proteins and membrane components followed by rapidly producing stress resistance factors, gly- colysis and amino acid synthesis.[49] This instantaneous response to changes in external stress factors, by global cellular metabolism modiﬁcation, optimizes growth and promotes survival of bacteria.

15 1. General Introduction

1.1.3.1. Biosynthesis of (p)ppGpp

In E. coli two enzymes, RelA and SpoT, are responsible for the synthesis of (p)ppGpp when triggered by diﬀerent starvation signals (Figure 1.6). The nucleotides are synthesized by enzymatic pyrophosphoryl group transfer of ATP to the ribose 3’OH of either GTP or guanosine-5’-diphosphate (GDP). The enzymes are summarized as RSH- proteins (RelA/SpoT-homologues) containing catalytic activity domains for hydrolase and synthase.[49] In RelA the hydrolase activity is inactive thus it only shows synthase activity. However, SpoT is specialized for hydrolase activity but also shows weak synthase function. Reciprocal intramolecular regulation of the dual activities assures a balance of (p)ppGpp levels and prevents a futile cycle of catalysis.[50]

Figure 1.6.: Regulation of (p)ppGpp synthesis by RelA/SpoT-homologues. Nutritional stress induces RelA or SpoT to synthesize (p)ppGpp by phosphorylation of GDP (or GTP) by transferring pyrophosphate from ATP. Bifunctional SpoT hydrolyses (p)ppGpp to GDP (or GTP) and pyrophosphate (PPi).

Amino acid starvation causes curtailment of ribosomal ribonucleic acid (rRNA) synthesis due to uncharged transfer ribonucleic acid (tRNA) binding the ribosomes. This is sensed by RelA and enables the synthesis of (p)ppGpp from GDP/GTP using ATP as a

16 1.1. Biological importance of phosphates phosphate donor (Figure 1.6).[51] In contrast, SpoT synthase activity is initiated under fatty acid, phosphate, carbon or iron starvation.[52] The capability of SpoT to degrade (p)ppGpp under restored nutrient availability balances the stress response in adaption to changes in environmental conditions (Figure 1.6).

1.1.3.2. (p)ppGpp signalling

The exact way how (p)ppGpp regulates cell growth is incompletely understood as the full set of effector proteins remains unknown. Wang et al. identified potential targets controlling a wide range of metabolic processes in E. coli.[53] These findings indicate that (p)ppGpp is responsible for the shutdown of many cellular processes in parallel.[53] Ribonucleic acid polymerase (RNAP) can be directly bound by (p)ppGpp and thus RNAP is often considered as the primary target of (p)ppGpp (see Figure 1.7). When nutrition is fully available the σ70-factor directs the RNAP to promoter regions encoding essential enzymes for bacterial replication and growth. Direct inhibition (Figure 1.7 (i)) of rRNA synthesis results from the destabilization of formed promoter-RNAP open complex. Figure 1.7 (ii) shows the indirect way to influence translation by direct binding of (p)ppGpp to RNAP facilitating the use of alternative σ-factors encoding proteins important for bacterial survival.

17 1. General Introduction

Figure 1.7.: (p)ppGpp regulates transcription in a (i) direct or (ii) indirect manner: (i) direct inhibition of the transcription of rRNA genes through destabilization of the formed promoter-RNAP open complexes by (p)ppGpp/DksA. (ii) indirect σ-factor competition. (p)ppGpp induces RNAP to be released from σ70-dependent promoters thus activating the expression of stress-induced genes by facilitating the use of alternative σ-factors. Picture taken from reference. [54]

However, Wang et al. showed that cells which produce a form of RNAP which does not bind (p)ppGpp are still growth inhibited. This indicates that (p)ppGpp is able to regulate cell growth through other eﬀector proteins than RNAP.[53]

The impact of the stringent response is not limited to bacteria. RelA/SpoT homologues have also been identiﬁed in plants and algae.[55,56] For example, plays stringent response in Xanthomonas citri, the causal agent of citrus canker, a role in virulence, nutrition uptake and host adaption.[57]

The insights in how alarmones modulate a plethora of processes involved in bacterial survival point out that inhibition of (p)ppGpp synthesis suppresses bacterial survival strategies. Hence, inhibition of the stringent response underlying mechanism would pave the way to new antibacterial agents.[58]

18 1.1. Biological importance of phosphates

1.1.3.3. Rel as a drug target

Approximately one quarter of the global population is asymptomatically infected with Mycobacterium tuberculosis.[59] Although the standard 6-month course treatment is effective, tuberculosis (TB) remains a global health emergency because e.g. gaps in treatment lead to the emergence of drug-resistant TB.[60] Even during active disease, non replicating cells are present displaying drug resistance (termed ”persiters”). The molecular mechanisms underlying M. tuberculosis persistence remain largely elusive.[60] M. tuberculosis possesses one homolog of RSH-protein (RelMtb) translating the stress of nutrient starvation into accumulation of (p)ppGpp. It was found that RelMtb is crucial for long-time survival of the pathogenic mycobacteria within the eukaryotic host.[60]

Dutta et al. screened pharmaceutical libraries for an inhibitor of RelMtb. They found that the lead compound X9 (16) was able to kill nutrition starved M. tuberculosis and enhanced the killing ability of the antibiotic isoniazid (17) currently used in TB treatment (Scheme 1.10). This indicates that inhibition of RelMtb is a promising strategy for targeting persisters thus shortening the treatment time.[60]

O HO NH N 2 N H N O S 17 16

Scheme 1.10: Chemical structure of antibiotic isoniazid (17) and lead compound X9 (16).

The above discussed biological importance of highly phosphorylated natural compounds substantiate the need for a reliable analytical method for such labile phosphates and its anhydrides. Methods which are currently in use are described in Chapter: State of the Art - Analytics. However, there is still interest in developing sensing approaches for example ﬂuorescence chemosensors. The next part of this chapter will describe strategies to achieve this endeavour.

19 1. General Introduction

1.2. A brief history of ﬂuorescence

In 1565, Nicolas Monardes reported fluorescence emission from water containing a mexican medicinal wood called Lignum nephriticum (Figure 1.8). A crude fluorescence emission spectrum was reported in 1845 by John Herschel from a solution of sulphate quinine.[62] Nowadays the vivid, blue fluorescence of quinine can be admired as an essential part of tonic water under UV light. Herschel used a prism to show that the blue colour could be only observed by illumi- nating with the blue end of the spectra but not with the red end. Emission spectrum analysis with a prism revealed emitted lights nature as blue, green and little yellow. However, Herschel did not conclude the emitted light to possess higher wavelengths than the incident light. Figure 1.8.: Fluorescence of infu- sion of Lignum nephriticum. Picture Seven years later in 1852, Sir George Gabriel taken from reference [61] Stokes launched the technique of observing fluorescence in his paper ”On the Refrangibility of Light”.[63] He used two coloured filters, one for the excitation beam and one through which to observe the emitted light. One of Stokes’ experiment was to move a test tube, filled with quinine sulphate, through the visible part of the light spectrum. The solution remained transparent until it reached the violet range of spectrum and beyond where it started to glow. Stokes wrote[63]: It was certainly a curious sight to see the tube instantaneously light up when plunged into the invisible rays: it was literally darkness visible. Altogether the phenomenon had something of an unearthly appearance.

He concluded that the emitted light was always of longer wavelength than the excitation. With that, Stokes contributed to the progress in history of photoluminescence and later his observations became known as the Stokes law. He called the new phenomenon

20 1.3. Supramolecular chemistry

fluorescence.[64] Additionally, association of concentration with the fluorescence intensity and quenching at high concentrations and in presence of different compounds were explained by Stokes.[64,65] These findings provided the conceptual basis of the sensitivity of fluorometric analyses.

1.3. Supramolecular chemistry

A prerequisite of life is the mutual recognition of molecules. It is crucial for the speciﬁcity of enzymes with their substrates, for antibodies, for the regulation of ion transport through ion channels and transport across cell membranes. Emil Fischer explained these processes in 1894 with a lock and key analogy.[66]

In 1987 the Nobel Prize for chemistry was awarded jointly to Donald J. Cram, Charles J. Pedersen and Jean-Marie Lehn for their development and use of molecules with structure-specific interactions of high selectivity (Figure 1.9). Peder- sen[67] published the synthesis of crown ethers which showed selective binding of spher- ical metal ions depending on the different sizes of the ”hole” they offer (Figure 1.9 structure 18). Cram[68] and Lehn[69] subsequently developed organic molecules with different cavities which could serve as hosts for molecules of different geometries (Figure 1.9 structure 19 and 20). With this work, a new chemical field called Supramolecular chemistry (or ’host-guest’ chemistry) was born.

O R R O O O O O O O N O O N R R O O O O O O O Me R O

18 20 19

Figure 1.9.: Examples of the work of Nobel Prize laureates C. Pedersen (18) [67], J. Lehn (20) [69] and D. Cram (19) [68].

Contrary to traditional chemistry is supramolecular chemistry dealing with weaker and reversible non-covalent interactions between molecular species. Lehn deﬁned it as the

21 1. General Introduction

”chemistry beyond the molecule” (Figure 1.10).[69] The term supermolecule describes complexes of two molecules which are not covalently bonded.

Figure 1.10.: Deﬁnition of supramolecular ”host-guest” chemistry according to Lehn. Picture adapted from reference. [70]

Today, supramolecular chemistry is divided into different interdisciplinary fields that interface with biological chemistry, materials science, synthetic chemistry and analytical methods.[71] One of these fields addresses the design of molecules as chemosensors.

1.3.1. Fluorescent chemosensors

”Compounds incorporating a binding site, a fluorophore, and a mechanism for commu- nication between the two are called fluorescent chemosensors.”[72] This definition is taken from Czarnik who is, together with de Silva, considered as the father of modern chemosensors.[73] Fluorescent chemosensors bear advantages, such as high sensitivity ”on-off” switchability and cost effectiveness. The modular approach allows for individual sensor designs.

22 1.3. Supramolecular chemistry

1.3.2. Anion sensing

Anions play an important part in life as we know it. In the section Biological importance of phosphates, the importance of myo-inositol , Inorganic polyphosphate (polyP) and Magic Spot Nucleotides (MSN) in biological processes have been discussed. Conse- quently, the development of facile, reliable anion sensors can be considered of importance as well. Even so, more sensors were developed for the detection of metal cations rather than for anions. This is a consequence of the selective binding of metal ions in aqueous media being significantly easier than binding to anions.[74] Thus, the design of a selective anion sensor is more challenging. Not only are anions larger than their isoelectronic cations (lower charge to radius ratio) but they are also pH sensitive (protonated at low pH and losing negative charge) and have a wide range of geometries (higher degree of design required).[74] The high solvation energy of anions is an additional difficulty because of the high influence of the environment on the molecular recognition process. The more polar the environment the higher is its ability to compete with the binding site of the chemosensor for binding the anion.

1.3.3. Ratiometric ﬂuorescent probes

One subset of optical sensing methods are the ratiometric ﬂuorescent probes. This method relies on simultaneous detection of two (or more) emission wavelengths and calculation of their ratio to detect changes to the local environment, such as ion concentration, pH or polarity. Since monitoring of a single emission wavelength can be fraught with diﬃculty because of interference from analyte independent factors, this technique bears the potential to provide precise and quantitative analysis. Such factors can be micro-environment around the probe molecule, its concentration or photobleaching. Ra- tiometric probes provide direct information about the analytes concentration without the need for calibration.[75]

23 1. General Introduction

1.3.3.1. Monomer/Excimer-based ﬂuorescent probes

Some fluorescent organic compounds exhibit a change in the ratio of emission wavelength maxima with increasing concentration instead of a concentration quenching of the excited monomer. This phenomenon was described by Förster in 1954 as a transition from a structured violet fluorescence band to a blue structureless band and is called excimer which is derived from ”excited dimer”.[76] In Figure 1.11b the monomer and excimer emission of pyrene is shown as a black line and a red dotted line respectively. Additionally, no change in the absorbance spectra is observed. Figure 1.11a summarizes the excimer formation of pyrene in two steps. I) The excitation of a ground stated pyrene (M) results in an exited state monomer (M*). The emission of its relaxation can be found at ˜ 370 nm. II) The fluorescence of this excited monomer is subject to self-quenching due to the collisional interaction of M* with M and yields an excited dimer (D*). Dissociation to its ground state gives the structureless

[77] fluoresence emission at ˜ 470 nm. Diskoidal π-systems possess extended excited singlet state lifetimes making pyrene (21) suitable as a fluorophore for excimer formation.[78] (Exiplexes are fluorescent pairs of non identical fluorophores.[78])

Monomer Excimer fluorescence fluorescence

120

l = 470 nm Monomer lEx = 345 nm lEm = 370 nm lEx = 345 nm Em

Excimer

100

60 hv M

40 Fluorescence Intensity Fluorescence

Ground stated pyrene Excited pyrene Excited dimer 0 M IM * II D * 350 400 450 500 550 600 650 21 Wavelength [nm ] (a) (b) Figure 1.11.: Excimer ﬂuorescence of pyrene: a concept of excimer formation. I) excitation of ground stated pyrene (M) forms M* with an emission at 370 nm. II) M* collides with M to D*. Emission of excited dimer at 470 nm.

Pyrene is used as a spatial ﬂuorescence probe because two ﬂuorophores needs to be in close proximity to yield the excimer emission.[79] This concept is not only powerful in studying protein spatial organization but also in anion recognition. Excimer emission,

24 1.3. Supramolecular chemistry triggered by analyte recognition, offers the possibilities to sense these analytes, without background fluorescence. Several design possibilities exist for monomer/excimer ratiometric chemosensors. Two concepts are depicted in Figure 1.12. The left part summarizes chemosensors, which rely on a guest induced folding of their flexible backbone, resulting in π-stacking of the fluorophores and consequently in an enhanced excimer signal. The second example shows a sensor where an excimer band can be observed in the free state and the binding event of the guest will increase the monomer fluorescence and decrease the excimer fluorescence.[80]

Figure 1.12.: Schematic depiction of monomer/excimer sensor design with the example of pyrene. a) Guest induced folding and stacking resulting in excimer turn-on fluorescence. b) guest induced unstacking of a tethered bifluorophore resulting in an excimer turn-off. Excitation wavelength for both sensors is 345 nm and emission 370 nm [Monomer] and 470 nm [Excimer].

1.3.3.2. ESIPT-based ﬂuorescent probes

Excited-state intramolecular proton transfer (ESIPT) is a fast proton transfer from a hydrogen bond donor, such as hydroxyl (or amino) to a hydrogen bond acceptor, e.g. a carbonyl (or imine nitrogen) atom in an excited state ﬂuorophore. One example is 2-(2’-hydroxyphenyl)benzoxazole (HBO) (Figure 1.13).

25 1. General Introduction

Figure 1.13.: Commonly used ESIPT ﬂuorophore: 2-(2’-hydroxyphenyl)benzoxazole (HBO).

The ESIPT process is a four-step photochemical sequence (Figure 1.14). The electronic ground state ﬂuorophore is typically existing in its enol (E) (Figure 1.14 left) form. Photoexcitation leads to a redistribution of the electronic charge, resulting in greater acidity for the hydrogen bond donor unit (here hydroxyl) and an increased basicity for the hydrogen bond acceptor moiety (here imine).

Figure 1.14.: The ESIPT process for the example 2-(2’-hydroxyphenyl)benzoxazole (HBO). ESIPT: Exited-state intramolecular proton transfer; RPT: Reverse proton transfer.

12 -1 This results in a fast enol to keto phototautomerization (kESIPT > 10 s ) and the excited state enol (E∗) is converted to its excited keto form (K∗) (Figure 1.14 right). After radiative decay back to the electronic ground state (K), a reverse proton transfer (RPT) takes place forming again the enol (E) form.[81] This four-step process explains why these ﬂuorescent probes are used for ratiometric sensing. ESIPT sensors possess

26 1.3. Supramolecular chemistry a large Stokes shift of approximately 200 nm. This avoids self-absorption and lowers interfering background ﬂuorescence. However, these probes are also sensitive to polar or hydrogen-bond donating solvents or to acidic analytes.[81] This can result in inhibition of the ESIPT process without formation of the keto (K∗) form as well as the emission at higher wavelengths. Therefore many ESIPT probes require a high degree of organic solvent limiting their application in aqueous media.

1.3.4. Overview of chemosensors for myo-inositol polyphosphates, polyP and Magic Spot Nucleotides

Examples of developed chemosensors from the literature are presented divided into three main categories (myo-inositol polyphosphates, inorganic polyphosphates (polyP) and Magic Spot Nucleotides (MSN)).

1.3.4.1. Chemosensor for myo-inositol polyphosphates

Inositol (1,4,5)-trisphosphate

Ins(1,4,5)P3 (3) is a member of the inositol polyphosphate family (Scheme 1.11) and controls the intracellular Ca2+ stores.

OH 2- HO OPO3 1

2- 2- O3PO OPO3 OH 3

Scheme 1.11: Chemical structure of Ins(1,4,5)P3 (3)

Niikura et al. reported a chemosensor, working in methanol, (Scheme 1.12) based on an anion sensing strategy: the indicator displacement assay.[82] The basic idea of this method is that the sensor is pre incubated with an indicator dye (here 5-carboxyﬂuorescein (22)) for which the analyte can compete in binding. The released dye is providing the optical output, which can be measured.[83] The chemical structure of the receptor

27 1. General Introduction is shown in Scheme 1.12. Hexasubstituted benzene, 1,3,5-tris(aminomethyl)-2,4,6- triethylbenzene is functioning as a rigid platform and helps building a cleft for analyte hosting. The platform is functionalized by three hexasubstituted benzene rings carrying six guanidinium (23 Scheme a)) or amine (23 Scheme b)) groups as binding moieties.

R R R R O O O R R HN N H - CO2 HN NH NH3 R = NH H - N a) b) CO2

23 22

Scheme 1.12: Chemical structure of chemosensor 23 and 5-carboxyﬂuorescein (22). An ensemble of receptor 23 and 5-carboxyﬂuorescein (22) is used in the indicator displacement assay to sense the [82] presence of Ins(1,4,5)P3 in methanol. Picture adapted from reference.

The 1,3,5-triethyl-2,4,6-trimethylbenzene (24) moiety possesses a conformational mobility which is impeded by spatial crowding (Scheme 1.13). Steric interactions of adjacent substituents at the benzene ring favours an alternating arrangement and is called the steric-gearing.[84,85]

R R

Scheme 1.13: Chemical structure of trialkylbenzene motif 24.

A convergent binding cavity is formed by the three ”arms” in a three-up, three-down manner.[86] This moiety provides a balance between preorganization and ﬂexibility thus serving as a spacer for chemosensors. A disadvantage of the indicator displacement method is the necessity of an excess of indicator dye, due to low receptor aﬃnity, thus lowering the signal-to-noise ratio and

28 1.3. Supramolecular chemistry sensitivity to low analyte concentrations.[87] However, this example shows the useful- ness of guanidinium moieties and a preorganized structure in an inositol polyphosphate chemosensor.

Do-Thanh et al. reported the tweezer shaped chemosensor 25 for Ins(1,4,5)P3 in methanol/water (1:1 v/v) showing a decrease in emission intensity upon titration with

[88] Ins(1,4,5)P3 (Scheme 1.15). Acridine, substituted at the 4- and 5-positions serves as backbone introducing rigidity and producing direct signal transduction at the same time. Click chemistry was used to attach the binding moieties to the backbone forming triazole. Triazole adds rigidity to the backbone and can further form additional hydrogen bonds to the guest.[88] 1,4,7,10-Tetraazacyclododecane (cyclen) was used as the binding moiety. Azamacrocycle complexes of cyclen[89–92] and Zn2+ are widely used binding sites in sensor design not least because of the improved water solubility. Placing a transition metal ion, such as Zn2+, into the azamacrocycle cyclen leads to coordination of the amines to the metal leaving one coordination site vacant for anions. This might give the selectivity for an analyte of interest.

O N O N 2+ 2+ N Zn N N Zn N N N N N N N H H N N N N 2- O3PO OH Ins(1,4,5)P3 2- N N HO OPO3

2-O PO OPO 2- N 3 N 3 N H N H 2+ 2+ N N N Zn N N N Zn N N N N O N O N 25

(a) (b) Figure 1.15.: a) Chemical structure of chemosensor 25 and proposed binding complex with Ins(1,4,5)P3 in methanol/water (1:1, v/v). b) Fluorescent sepctra of 25 (10 µM) in methanol/TRIS buﬀer (40 mM, pH 7.4) (1:1, v/v) upon addition of anions. Picture taken from reference.[93]

In 2012 Jung et al. reported chemosensors 26 and 27 for Ins(1,4,5)P3 using a ﬂuores- cent imidazolium receptor (Figure 1.16a).[93] Imidazolium-containing sensors making charge-charge electrostatic interactions dominating as the positive charge is delocalized in the imidazole ring. This can induce ionic hydrogen bonding between the imidazolium (C-H)+ and phosphate groups.[94–96] Sensor 26 is an example for an selective ”on-oﬀ”

29 1. General Introduction

chemosensor as the ﬂuorescence is quenched upon binding of Ins(1,4,5)P3 with an stoichiometry of 1:1, but not for other inositol phosphates, PPi and ATP in DMSO–HEPES buﬀer (20 mM, pH 7.4) (1:9, v/v) (Figure 1.16b).

N N

N N N

N N N N N N N N

26 27

(a) (b) Figure 1.16.: a) Chemical structure of chemosensors 26 and 27 for Ins(1,4,5)P3. b) Fluorescent sepctra of 26 (10 mM) in DMSO/HEPES buﬀer (20 mM, pH 7.4) (1:9, v/v) upon addition of anions. Picture taken from reference. [93]

Yoon and co worker published in 2014 a tetranaphthoimidazolium receptor 28 selective

[97] for InsP6 in HEPES buﬀer (20 mM, pH 7.4) with 1:1 stoichiometry (Scheme 1.17). Chemosensor 28 bears naphthoimidazolium moieties which can serve as anion binding site with their interactions between (C-H)+ and anions.

N N

(a) (b) Figure 1.17.: a) Chemical structure of chemosensors 28 for InsP6.b) Fluorescent sepctra of 28 (10 µM) in HEPES buﬀer (20 mM, pH 7.4) upon addition of anions. Picture taken from reference.[75]

30 1.3. Supramolecular chemistry

In addition naphthoimidazoliums are inherent fluorescent which circumvents the synthetic introduction of an chromophore into the sensor. Only addition of InsP6 lead to an selective fluorescence enhancement at 465 nm (Figure 1.17b) whereas PPi, Ins(1,4,5)P3, ATP and adenosine diphosphate (ADP) did not induce a significant change in emission intensity. Lee et al. applied sensor 28 for live cell imaging in HeLa cells. The probe successfully passed the cell walls and incubation with InsP6 induced a strong fluorescence.

[98] Ahn and co-workers developed 2008 a sensor 29 for Ins(1,4,5)P3 sensing through a competition assay.[82] The benzene-based tripodal system was again applied in this sensor as it readily provides three binding sites in a matched geometry with the three phosphates of Ins(1,4,5)P3.

N 2+ Zn N N N 2+ COO Zn Br Br 2+ N N Zn N N O O O N Br Br 29 30

(a) (b) Figure 1.18.: a) Chemical structure of chemosensors 29 for Ins(1,4,5)P3 and EosinY (30).b) Fluo- rescent sepctra of 29 (2.0 µM) and 30 (3.0 µM) in HEPES buﬀer (10 mM, pH 7.0) with addition of [99] Ins(1,4,5)P3 (0-6.0 µM). Picture taken from reference.

An ensemble system of receptor 29 (2.0 µM) and EosinY 30 (3.0 µM) in HEPES buffer (10 mM, pH 7.0) was chosen. The fluorescence of EosinY 30 was quenched when binding into the receptor 29. Titration with Ins(1,4,5)P3 (0-6.0 µM) resulted in a increase of emission intensity of 30 as it is replaced by Ins(1,4,5)P3 in the receptor. Addition of - PPi and HPO4 gave no significant change in fluorescence emission.

31 1. General Introduction

1.3.4.2. Chemosensor for inorganic polyP

Since 1980 DAPI (31) has found application in polyP staining.[100] The chemical structure is shown in Scheme 1.14. This chemosensor was initially used as a dye for DNA but an emission shift was observed with added polyP as analyte.[100] For In-depth information about this sensor see chapter Sensing via hydrogen bonding.

O O O NH OP O P O P O HN O O O N NH2 H n NH2 DAPI polyP

Scheme 1.14: left: Chemical structure of chemosensor DAPI. right: Chemical structure of polyP.

Free DAPI emits at approximately 460 nm and bound into the minor groove of A-T rich sequences of DNA a 20 fold enhancement of the fluorescence intensity can be observed. It was realised that the polyanionic polymer polyP could also interact with DAPI resulting in an enhanced fluorescence intensity and a bathochromic emission shift to 520 nm, whereas small anions, such as orthophosphate showed no effect. This enables a signal transduction with low background fluorescence , thus enhancing the sensitivity. In 2014 Angelova et al. published two benzimidazolium sensors for sensing of polyP (Figure 1.19 b)) in assays of purified polyP as well as polyP staining in living cells.[101]

(a) (b) Figure 1.19.: a) Emission spectra of JC-D7 (10 µM) and JC-D8 (10 µM) with 0-80 µM polyP (average chain length 60) in 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (20 mM, pH 7.4) [101] with λEx = 390 nm.b) Chemical structure of JC-D7/D8. Picture taken from reference.

32 1.3. Supramolecular chemistry

They screened a known library of 96 fluorescence molecules which was earlier tested for polyanions Heparin[102] and GTP[103]. The screening resulted in JC-D7 and JC-D8 (Figure 1.19 b)) as suitable polyP probes. Both probes showed an significant enhance- M ment in the emission intensity at ˜ 520 nm (Figure 1.19 a)) in HEPES (20 m , pH 7.4) with addition of polyP, whereas other anions, such as orthophosphate, nucleotides (ADP, ATP, AMP, GTP), DNA and RNA could not induce an increase in fluorescence intensity. It is particularly interesting that JC-D7 and JC-D8 did not show an fluorescence response in presence of RNA as this polymer is very abundant in organisms.[104]

1.3.4.3. Chemosensor for Magic Spot Nucleotides

In 2008, Rhee et al. published a selective (p)ppGpp chemosensor called PyDPA.[105] This chemosensors can be used for MSN detection besides 32P in thin layer chromatography (TLC) or UV detection by high performance liquid chromatogpraphy (HPLC).[106,107] Figure 1.20 shows the free sensor 33 and the formed complex upon (p)ppGpp addition. PyDPA 33 consists of two parts, as the name indicates (Py = pyrene, DPA = dipicolylamine).

O O

O ppGpp N

N N 2 N 2+N O N N N O Zn 2+ P O N 2+N Zn O N N N Zn - O O 2+ N 2+N O O HP Zn N P N Zn O P O O 2+ O- O N Zn O HO O 33 N N N

H2N N O H

Figure 1.20.: Chemical structure of PyDPA (33). Addition of (p)ppGpp forms an excimer complex with ratio of 2:1 (Sensor:Analyte). Picture adapted from reference.[105]

Two Zn2+-dipicolylamine (DPA) moieties acting as binding units as they are known for

[108] their strong aﬃnity to PPi in water. Pyrene serves as the sensing unit as it has the

33 1. General Introduction

ability of forming excimer complexes which are ﬂuorescent at ˜ 470 nm. The binding of chemosensor and (p)ppGpp can be seen in Figure 1.20 and requires a 2:1 ratio of sensor to analyte.

34 1.3. Supramolecular chemistry

1.3.5. References

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42 CHAPTER 2

State of the Art - Analytics

43 2. State of the Art - Analytics

In this chapter a brief summary of today’s most commonly applied analytic methods of the introduced polyanionic molecules is delineated.

2.1. Radioactive labelling

2.1.1. Inositol polyphosphates

One of the initially used method for detecting complex mixtures of cellular InsPs and PP-InsPs is metabolic labelling with [3H]-labelled inositol.[1] This method is based on the ability of eukaryotic cells to take up the tritiated inositol from the extracellular medium.[2] The protocol works in four steps: radioactive labelling, extraction of soluble inositol polyphosphates, chromatographic separation and scintillation counting (Figure 2.1).

Figure 2.1.: Schematic representation of workﬂow for inositol extraction and strong anion exchange (SAX) HPLC separation. Picture taken from reference [3]

Cells are grown in a medium containing [3H]-inositol (20 µCi mL-1 for mammalian cells) until isotopic equilibrium is reached (5 days). Then, the cells are lysed and the metabolites extracted with 1 m perchloric acid. The tritiated inositol-containing samples are puriﬁed on a SAX column and subsequently the total [3H] present in inositol is counted by scintillation counting of the collected fractions.[4] Recently, Desfougères et al. published a study proposing the existence of an alternative ”soluble” (nonlipid) pathway for

[5] the synthesis of InsP6 from glucose-6-phosphate in eukaryotic cells. As mentioned above, metabolic labelling, using [3H]-inositol, is a method for inositol phosphate detec-

44 2.2. Nuclear Magnetic Resonance Spectroscopy (NMR) tion. However, this method is based on the exogenously addition of [3H]-inositol, thus not able to detect inositol phosphates originating endogenously from glucose.[5]

2.1.2. Magic Spot Nucleotides

To asses whether (p)ppGpp is formed in bacteria it is possible to label total cellular

32 nucleotides by Pi during growth in vitro in low phosphate media. The nucleotide response can be analysed by 2D-thinlayer autoradiogram as shown in Figure 2.2 using the example of M. tuberculosis.[6]

32 Figure 2.2.: 2D-TLC analysis of Pi-labeled intracellular (p)ppGpp in M. tuberculosis. C) Nucleotides extracted from wild type cells under starvation. D) Nucleotides extracted from cell with deleted RelMtb under starvation conditions. E) Schematic representation of 2D-TLC. Image taken from reference. [6]

However, 2D-TLC is not able to discriminate between ppGpp and pppGpp and furthermore high concentrations are needed for the analysis.[6]

2.2. Nuclear Magnetic Resonance Spectroscopy (NMR)

Nuclear magnetic resonance (NMR) is another method for studying phosphorylated compounds and their metabolism. This technique provides detailed information about the nucleus’ chemical and structural environment based on the chemical shift.

45 2. State of the Art - Analytics

2.2.1. 31P-Nuclear Magnetic Resonance Spectroscopy

Phosphate compounds possess chemical shifts, in the 31P-NMR, which can span over a wide parts per million (ppm) range. Additionally, no solvent suppression is needed, since no water signal appears in 31P resonance region.

2.2.1.1. Inorganic Polyphosphate (polyP)

It is possible to determine by NMR the connectivity (linear, cyclic[7] or branched[8]) of phosphate chains. PolyP with a chain length ≥7-units display four major signals that correspond to the respective position of the phosphate in the chain. Terminal phosphate groups (PP1, -6 ppm), the penultimate (PP2, -20 ppm,; PP3, -21 ppm) and the core phosphates (PP4, -22 ppm) (Figure 2.3).[9]

31 Figure 2.3.: P-NMR spectrum of a mixture of Pi, polyP (PP1-4) and cyclic polyP. Picture taken from reference [10]

Recently a study was published where 31P-diffusion-ordered spectroscopy (DOSY) was used to determine specific chain lengths of individual polyphosphates within a mixture (Figure 2.4) whereas conventional 31P-NMR only measures the average chain length.[11] DOSY reports the diffusion coefficient and links it with the individual chemical shift.[12] Molecules in solution possess a translational motion. By the use of a gradient the diffusion of molecules with different size can be marked depending on their position in the tube because small molecules have faster translational motion in solution than larger ones. The measured signal is the integral over the whole sample volume.

46 2.2. Nuclear Magnetic Resonance Spectroscopy (NMR)

Figure 2.4.: 31P-DOSY-NMR spectrum of Algae (Chlorella vulgaris) alkaline extract. Picture taken from reference. [11]

2.2.1.2. Inositol polyphosphates

Myo-inositol and phytate are meso-compounds thus possessing a mirror plane. In 31P- NMR, phytate displays four distinct resonances (see Figure 2.5b), for the 2- and 5 positions each phosphate possess one resonance. The 1/3 and 4/6 positions result in one distinct resonance for each pair. Therefore, the integrals display a ratio of 1 (C2):2 (C1/3):2 (C4/6):1 (C5) (Figure 2.5b).

2- OPO3 2- 2- O3PO OPO3 3 1 4 6 2- 2- O3PO OPO3 2- OPO3

(a) (b) Figure 2.5.: a) InsP6 with mirror plane (red dotted) dissecting the 2- and 5- positions. 1/3 and 4/6 31 positions are chemically identical. b) Proton decoupled P-NMR of InsP6 which displays four distinct resonances, one for the 2/5 position each and one for the 1/3 and 4/6 positions each.

Breaking InsP6 symmetry will result in ﬁve diﬀerent signals for every single phosphate.

There are six possible InsP5 isomers. Among these six isomers are two enantiomeric pairs: InsP5 [1/3-OH] and [4/6-OH] (Scheme 2.1). With achiral counter ions, such as Na+, the enantiomers can not be distinguished by 31P-NMR.

47 2. State of the Art - Analytics

2- 2- OPO3 OPO3 2- 2- O3PO OH HO OPO3 3 1 3 1 4 6 4 6 2- 2- 2- 2- O3PO OPO3 O3PO OPO3 2- 2- OPO3 OPO3

InsP5 [1-OH] InsP5 [3-OH]

Scheme 2.1: Chemical structure of InsP5 [1-OH] and InsP5 [3-OH].

To address this problem the Jessen-group developed a new NMR-method using L- arginine amide hydrochloride salt (LArgN) as a chiral solvating agent. In this thesis, this method was applied to show that the enantiomeric inositol polyphosphates, 1/3-OH

31 InsP5 are distinguishable in the presence of an excess of LArgN by P-NMR. This work is described in detail in Chapter Assignment of the stereoselectivity of a new phytase.[13]

2.2.2. 13C-Nuclear Magnetic Resonance Spectroscopy

The fact that the 13C isotope has a natural abundance of approximately 1 %, makes labelling of selected compounds a tool for targeted information on these molecules, even in complex cellular environments.

2.2.2.1. Inositol polyphosphates

Recently, the Fiedler group developed a NMR method, which is based on 13C-labelled compounds that can be applied for the detection and quantiﬁcation of InsPs and PP- InsPs within complex mixtures and at physiological concentrations (Figure 2.6).[14] In their work they demonstrated that metabolic labelling of mammalian cell lines with

13 13 [ C6]myo-inositol and subsequent detection of in vivo generated [ C6]InsP enables monitoring of InsPs/PP-InsPs levels in complex mixtures.[14]

48 2.3. Enzyme Assays

Figure 2.6.: Representation of 13C-labelled InsPs and PP-InsPs which can be detected without the need of puriﬁcation in complex media. Picture taken from reference[14]

NMR is a non invasive method for analysing phosphorus species quantitatively in solution. However, this technique requires expensive equipment and high sample concentrations (millimolar range). Lower concentrations are possible but increase the time which is needed for the measurement.

2.3. Enzyme Assays

2.3.1. Inorganic Polyphosphate (polyP)

Enzymatic methods, using PPX1 and polyphosphate kinase 2 (PPK2) are highly specific in determining polyP concentration quantitatively in bacterial samples only requiring a plate reader. (i) Assay one is a two-enzyme assay: the PPK2 reaction, in which polyphosphate is converted into ATP in presence of an excess of ADP, is used and subsequent hydrolysis of the generated ATP by standard firefly luciferase assay is generating light. This method is able to detect picomolar amounts of phosphate residues but prior polyP extraction is required.[15,16] (ii) PPX1 assay works with Saccharomyces cerevisiae exopolyphosphatase 1 which hydrolyses polyP and the liberated Pi is colorimetric determined by either ascorbate or malachite green standard assay Figure 2.7b.

49 2. State of the Art - Analytics

PPX1 PolyPn n Pi

(a) (b) Figure 2.7.: a) PolyP is hydrolysed to Pi by PPX1. b) Example of a malachite green assay. The shown wells contain: 1 and 2: enzyme reactions; 3: 0.04 mM PPi; 4 negative control. Picture taken from reference. [17]

For enzyme assays commonly available laboratory equipment is required, such as a plate reader and a multichannel pipet. The throughput capacity is high because the assays are done in 96-well plates and impurities like DNA, RNA are not interfering the analysis. However these assays reach their limits with shorter polyP chains.[16]

2.4. Electrophoresis

Electrophoresis is a technique, which uses the ability of charged molecules to migrate through a porous matrix, driven by a homogeneous electric ﬁeld. Smaller molecules will migrate faster than larger ones. Analytes can be separated according to their intrinsic electrophoretic mobility (µep), in an applied electrical ﬁeld (Equation 2.1). Where q is the charge on the analyte, η is the viscosity of the media and rs is the hydrodynamic radius.

µep = q/6πηrs (2.1)

The electrophoretic mobility (µep) depends on charge and radius of the analyte, viscosity of the media and gives information about the rate of migration of a given compound. Velocity, at which an ion moves, is proportional to the applied electric ﬁeld. The greater the ﬁeld strength, the faster migration.

50 2.4. Electrophoresis

2.4.1. Polyacrylamide Gel Electrophoresis (PAGE)

Another method for the quantitative determination of highly phosphorylated compounds is PAGE analysis. This technique doesn’t require expensive equipment.

Maxam and Gilber invented 1977 a technique for DNA sequencing, using gel electrophoresis, which was adapted seven years later for polyPs by Robinson et al..[18,19] A typical polyP PAGE analysis uses tall chambers (14 x 40 cm) in which the vertical slab gel is poured between two glass plates and equipped with a comb, building the loading pockets. Polymerized for a sufficient time the gel is pre-electrophoresed. Subsequently, a mixture of analyte solution and orange G (OG) is loaded into the prepared cavities. Orange G functions here as a horizontal, visual reference. A ladder containing different length of polyP functions as a measure of vertical separation. The gel is electrophoresed typically in Tris/Borate/EDTA (TBE) buffer under external cooling, allowing the anions to migrate along with the electric current. After removal of the glass plates, the gel is stained with a dye visualizing the anionic compounds.

NH N Cl HN N NH2 H H2N S N NH 2 34 35

Scheme 2.2: Commonly used gel staining dyes for visualization of anionic compounds a) 4’,6-Diamidin- 2-phenylindol (DAPI) (34) b) Toluidine blue (35).

2.4.2. Inorganic Polyphosphate (polyP)

Staining protocols for polyP have been developed using toluidine blue[20] and DAPI[21] (see Scheme 2.2). The former is a positively charged metachromatic dye, which shifts its absorption to 520 nm when binding to an anionic polymer like polyP. The gels are stained with 0.05% toluidine blue, 25% methanol and 1% glycerol, followed by destaining in 25% methanol and 5% glycerol. Stained gels can be scanned in white light on a ﬂatbed scanner and analysed with picture analysis software.

51 2. State of the Art - Analytics

(a) (b) Figure 2.8.: Comparison of DAPI and Toluidine blue staining of PAGE analysis. Analytes per lane from left to right: A polyP, B heparin, C heparan sulfate, D chondroitin sulfate A, E chondroitin sulfate B, F chondroitin sulfate C (10 µg of each). a) DAPI positive stain with p-phenylenediamine and ethylenediaminetetraacetic acid (EDTA) and b) Toluidine blue staining. Pictures are taken from reference. [21]

DAPI staining is based on the fact, that this dye exhibits a blue emission when bound to DNA but a greenish-yellow when bound to polyP. 2 µg/mL DAPI, 1 mg/mL p- phenylenediamine and 10 mM EDTA pH 8.0 and the usage of foil-covered containers are needed for this procedure. Destaining is achieved by using the same solution but omitting DAPI. Imaging these gels requires an UV transilluminator plate or a hand lamp with 365 nm light and a digital camera. Bound to polyP, DAPI undergoes photobleaching providing even more sensitivity and selectivity.

Both staining methods have their advantages and disadvantages: Toluidine blue is suitable for universal staining of anionic species but the LOD, for polyP, is, with 250 pmol per band, higher than 4 pmol for photobleached DAPI.[21] DAPI, on the other hand, stains with higher sensitivity and speciﬁcity for inorganic polyphosphate but is not applicable for a broader analyte spectrum (Figure 2.8).

2.4.3. Inositol polyphosphates

Recently, studies from Losito et al. proofed that PAGE analysing and subsequent staining with the dyes mentioned above (Scheme 2.2) is also applicable for highly phosphorylated inositol polyphosphates in nanomolar quantities.[22] For enzymatic studies

52 2.4. Electrophoresis they took advantage of the fact that DAPI stains diphosphoinositol pentakisphosphate

(InsP7) (< 100 pmol) positively and InsP6 (> 200 nmol) negatively.

A ratiometric fluorescent chemosensor (36) was reported by Williams et al. in 2015 (Scheme 2.9).[23] As mentioned above (chapter General Introduction), at any given time on third of the living systems proteome is phosphorylated. Complementary to this post translational modification, protein pyrophosphorylation is a signalling mechanism where PP-InsPs transfer their β-phosphoryl group to a phosphoserine group on the protein. Until today, this mechanism has only been characterized in vitro. Williams et al. developed a chemosensor 36 based on a dizinc(II)bis(dipicolylamine) binding moiety and pyrene as a fluorescent reporter for pyrophosphorylated peptides. They were separated on SDS-PAGE and stained using 36 in DMSO/HEPES 1:1 (100 mM, pH 7.16).[23]

NHHN NH HN N OH N N H2O N Zn Zn N O N

(a) (b) Figure 2.9.: a) Chemical structure of ratiometric ﬂuorescent chemosensor (36) and b) Pyrophospho- rylated protein and a gel extract stained with 36. Picture taken from reference. [23]

PAGE provides qualitative results and with help of polyP or DNA[24] standards it can be interpreted quantitatively and polyP as well as inositol phosphates can be analysed without any degradation. However, samples have to be puriﬁed in advance, to get rid of interfering compounds, and rather high sample volumes and time consuming runs are needed.

53 2. State of the Art - Analytics

2.4.4. Capillary Electrophoresis (CE)

Another separation technology is capillary electrophoresis (CE). This method is performed within capillaries (standard inner diameter 50µm). In a typical set-up the electroosmotic ﬂow (EOF) is directed to the negatively charged cathode and the result of that is a decreased migration velocity of the anions. It can be explained with the example of an silica capillary which is in contact with the water solution. If the pH is higher than 1.5 the silanole groups are deprotonated. A consequence of this dissociation is the formation of a double layer with negative charge on the silica surface and positive charge on the liquid side.[25] The charge on the surface is stationary while the mobile charge follows the electrical ﬁeld towards the cathode.

veo = µeo × E (2.2)

Equation 2.2 shows that the electroosmotic velocity (veoeo) is dependent on the electrophoretic mobility of the solute (µeo) and the ﬁeld strength (E). The apparent mobility

(µapp) of a solute is the sum of the electrophoretic mobility (µep) and the electroosmotic mobility (µeo) (Equation 2.3) which is important for the separation power of a CE system.

µapp = µeo + µep (2.3)

The inner surface of the capillary can be coated with small molecules to reverse the EOF and with that restoring the normal direction of migration - anions towards the anode. Separated analytes can be detected by using indirect UV-, MS- or conductivity- detection.

54 2.5. Chromatography

2.4.5. Inositol polyphosphates

In 1992 Henshall et al. used CE for the detection of inositol phosphates with indirect photometric detection achieving a LOD of approximately 200 ng/mL .[26] Since then, numerous studies were published on inositol phosphate analysis using CE from InsP1 to [27–34] InsP8.

2.4.6. Inorganic polyphosphate (polyP)

Almost simultaneously, research on polyP using CE technique was reported by three independent groups.[35–37] PolyPs with longer chain length than four phosphates display nearly the same electrophoretic mobility and thus are challenging to separate. Using

[35] cationic EOF modiﬁer, Frederick et al. reported in 1997 separation up to P30. Capillary electrophoresis bears several advantages: short analyse times, low sample consumption, no need of pre- or post-column derivatization using indirect UV detection and various separation modes like capillary zone electrophoresis (CZE), capillary gel electrophoresis (CGE) and capillary electrochromatography (CEC). Nevertheless biological samples have to be extracted prior to measurement. This extraction methods use strong acidic conditions which might damage the analytes and falsify the results.

2.5. Chromatography

Chromatography allows the separation of a substance mixture, based on their diﬀerent distribution between a mobile and a stationary phase.

2.5.1. Ion chromatography (IC)

HPLC is a technique which is ubiquitous in laboratories all over the world. Development of exchange resins, in the 1930s, made separation of ions possible but the breakthrough came 40 years later with the establishment of IC.[38] IC utilizes the diﬀerence of the charged molecules aﬃnity to a stationary phase. The stationary phase is an anion

55 2. State of the Art - Analytics exchange resin that carries charged functional groups. In case of anion exchange chromatography these are quaternary ammonium groups, which interact with the oppositely charged anions.

The general set-up of an ion chromatographic system is shown in Figure 2.10. The diﬀerence to an conventional HPLC-system is the usage of an eluent generating cartridge followed by a suppressor and conductivity detection.

Figure 2.10.: General set-up of an IC-system. Picture taken from reference [39]

Deionized water is pumped through the system by a dual pump, passing an eluent generator cartridge (EGC) (invented in 1998 by Dionex). The EGC contains an electrolyte solution and generates the hydroxide eluent from deionized water in high ﬁdelity. The operation principle can be seen in Figure 2.11, water is pumped through the eluent generation chamber and a current is applied between the anode located in the eletrolyte reservoir and the cathode located in the eluent generation chamber. At the cathode, water is oxidized to form H+ ions which displace K+ ions in the reservoir. The displaced K+ ions migrate towards the eluent generation chamber. At the anode water is reduced to form OH- ions which combine with K+ in the eluent generation chamber to form the KOH eluent. Precisely applied current determines the concentration of generated KOH.

56 2.5. Chromatography

Figure 2.11.: Illustration of the operation principle of an Dionex EGC Cartridge for the generation of an KOH eluent. A current is applied between the cathode and anode resulting in the electrolysis of water. H+ ions, generated at the anode, replace K- which combine with OH- in the eluent generation chamber to produce the KOH eluent. Picture taken from reference. [40]

Downstream connected are the continuously reagent trap column (CR-TC) for remov- ing any contamination of the deionized water and a gas removal device that removes electrolysis gases created during eluent generation. Figure 2.12 depicts the principle of a CR-TC which consists of an anion exchange bed and a cathode at the eluents in- let. Anode and cathode are separated through an anion exchange membrane. Anionic impurities cross the membrane and hydroxide generated at the cathode continuously regenerates the anion exchange resin. H+ generated at the anode combines with the anions to form acids.

Figure 2.12.: Operation principal of an continuously reagent trap column (CR-TC). Anionic impurities cross the anion exchange membrane and the continously generated hydroxide from the cathode regenerates the anion exchange resin. Picture taken from reference. [41]

57 2. State of the Art - Analytics

The sample is injected via an autosampler to the system and passes through the guard- and analytical column where the anions are separated based on their retention to the quaternary ammonium groups. Increasing the ionic strength, or by variation of the eluents pH, the equilibrium of the phases is aﬀected, causing the retained anions to be sequentially eluted. Some anions, such as inositol polyphosphates, lack an intrinsic chromophore. However, conductivity is an universal property of ionic species in solution. Detection of the analyte of interest is hindered by the presence of much more abundant eluting electrolyte (for example OH-). Development of a combination of resins, which separates the ions of interest and neutralizes the eluent, laid the foundation for a suppressor based IC system.[42] Electrolysis of water forms hydronium ions at the anode and hydroxide at the cathode. Cation exchange membranes allow the hydronium ions to pass into the analyte chamber to neutralize the hydroxide eluent generating neutral water with low conductivity (see Figure 2.13). Sample or eluent counter ions are driven towards the electric ﬁeld to the cathode where they combine with the formed hydroxide. This process lowers the background resulting in increased sensitivity.

Figure 2.13.: Operation principal of a suppressor. Electrolysis of water is used to remove cations (for example Na+ or K+) ions and replace it with H+, generating neutral water with low conductivity. Picture taken from reference. [43]

58 2.5. Chromatography

Although conductivity detection is a good method to detect anions, the system is limited to the usage of diﬀerent concentrations of the hydroxide eluent. Compounds like polyP or inositol phosphates are not stable under these conditions. Therefore, a possibility to circumvent this limitation is a system which allows free selection of the eluent. As conductivity detection is not possible under these conditions, UV/Vis detection is an option, but, as mentioned before, some anions possess no chromophores.

2.5.2. Post column detection methods

One solution could be the usage of a postcolumn staining reagent (PCSR). This works with external addition of a molecule which interacts with the analytes generating a readable UV/Vis or ﬂuorescence response. Fluorescent dyes are of interest because of the increased sensitivity of the signal output. For the reasons given above, it is challenging to detect anionic species like inositol phosphates or polyP. Whereas for polyP analysis some successful IC methods were reported[44–47], using a hydroxide gradient and conductivity detection, diﬀerent ways have to be found for inositol phosphates. 1986 Meer et al. reported a non-radiometric PCSR method which uses on-line enzymatic hydrolysis of the phosphate esters and subsequent detection of the formed

[48] Pi. This method allowed the determination of concentrations of inositol bisphosphate

(InsP2) and InsP3 in living animals with a LOD less than 1 nmol. This procedure suf- fered from the high amounts of tissue needed and from interference of incompletely dephosphorylated higher inositol phosphates.

During the past few years a number of ion-exchange chromatography methods,with gradient elution for separation of inositol phosphates in biological tissues, have been developed.[49–51] In 1988 Mayr et al. published a method called the ”metal-dye detection (MDD)” method which did not require for dephosphorylation.[49] Polyanions, like inositol phosphates, bind with high aﬃnity to trivalent transition-metals. Mayr used this fact and developed a dye-based ternary-complexometric technique which relies on two essential

59 2. State of the Art - Analytics components: 4-(2-pyridylazo)resorcinol (PAR) and trivalent transition metal cations (Figure 2.14). PAR (37) (Scheme 2.3) is a dye which shows little absorbance at 520 nm in absence of cations. However, transition-metal complexes of this dye absorb strongly at 520 nm.

N NN

HO OH 37

Scheme 2.3: Chemical structure of 4-(2-pyridylazo)resorcinol PAR 37

The trivalent transition metal cations are provided with the buﬀer and the dye PAR is continuously added post-column to the eluate (Figure 2.14b). The signal at 520 nm decreases when competing ligands, e.g. InsPs, are present and the mixture has a pH above 7.5 (Figure 2.14a). The resulting chromatograms show negative peaks which are routinely inverted to facilitate integration.[49] A strong acidic elution protocol was used in order to prevent precipitation of the metal with phosphorylated compounds on the column. The post-column added dye solution was buﬀered to bring the mixture to alkalinity to assure detection.

(a) (b) Figure 2.14.: a) InsPs bind to trivalent transition-metal ions (M = Y,La,Nd,Gd,Ho,Lu). b) Build-up of the HPLC system using the post-column derivatisation with trivalent metal. Pictures taken from reference. [49]

Mayr used an HCl-eluent (0.2-0.4 mM) containing YCl3 (9-14 µM). The post-column stain solution contained PAR (200 µM) and by UV-detection at 520 nm they could detect inositol phosphates in a pico molar range. Figure 2.15 shows and example chromatogram of a standard mixture of nucleotides and InsPs eluted with the acidic eluent

60 2.5. Chromatography

containing YCl3 and monitored at 545 nm after mixing with post-column staining solution (upper trace) and the lower trace by UV at 245 nm.

Figure 2.15.: Chromatogram of a standard mixture of nucleotides and InsPs. The upper trace was monitored by MDD at 545 nm and the lower trace by UV at 245 nm. Picture taken from reference. [49]

Today, a common eluent system for separation of inositol phosphates is a HCl-gradient with post-column mixing of 1%Fe(NO3)3‚9*H2O in a 2% solution of HClO4. The UV signal is measured at 290 nm according to the method of Phillipy and Bland.[50]

Table 2.1.: Summary of Analytical methods and the individual applicability for polyinositolphosphates, PolyP and Magic Spot Nucleotides. 3 32 32 [ H]/[ Pi] P-NMR Enzyme Assay PAGE CE HPLC IC InsPs + + - ++-+ PolyP - + + ++-+ MSN + + - + + + +

Table 2.1 summarizes the analytical methods discussed in this chapter and shows the individual applicability of for the discussed phosphorylated metabolites. Some of the methods, described in this chapter, were used within this thesis in combination with new developed ﬂuorescent chemosensors.

61 2. State of the Art - Analytics

2.5.3. References

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CHAPTER 3

Goals of the Thesis

67 3. Goals of the Thesis

Phosphorylated metabolites are important molecules regulating various biological processes. In order to get a deeper understanding of the distinct roles of these molecules sophisticated sensors have to be developed. These sensors have to be able to give a reliable output in aqueous media, at low analyte concentrations and in complex media, such as they exist in a biological context.

To address this issue the present thesis focused on the development, synthesis and evaluation of fluorescent chemosensors for InsPs, PP-InsPs, polyP and Magic Spot Nucleotides. As our group has synthetic access to pure inositol phosphates and inositol pyrophos- phates as well as to magic spot nucleotides and PolyP, we can carefully screen the effect of these molecules, which usually is not possible due to the limited material availability. Three different sensing approaches were used for the design of the chemosensors: Disassembly approach, Pyrene-Excimer and DAPI-based (Figure 3.1).

Figure 3.1.: Diﬀerent approaches applied for sensing of phosphorylated metabolites.

Furthermore, the application of the developed chemosensors in polyacrylamide gel staining should be assessed. PAGE is a method which is commonly used in many laboratories

68 for the analysis of cell extracts for inositol polyphosphates, polyP and MSN. Developing a convenient procedure with a new chemosensor could lower the limit of detection. The developed sensors could as well be applied as post column stain for an ion chromatography system. This sophisticated machine is powerful in separation of anions and a convenient post column stain would shed some more light on this important classes of phosphorylated molecules. Scheme 3.1 shows examples of the chemosensor designs in the three categories: DAPI- based, Pyrene-Excimer and Disassembly approach.

DAPI-based Pyrene-Excimer

NH 2 NH NH N H

H2N N H N HN Disassembly approach NH N N N NH O HN NH O HN N N NH N O 2+ O O Cu O HN N O O NH N N N N N N N O N N

Scheme 3.1: Examples of novel chemosensors synthesized in this thesis

In this thesis the design, synthesis and evaluation of novel chemosensors for highly phosphorylated metabolites will be presented.

CHAPTER 4

Excimer based sensing

71 4. Excimer based sensing

4.1. Background

Some fluorescent organic compounds, such as naphthalene, anthracene and pyrene possess the ability to form excimers - exited dimers of two planar fluorophores. An excited pyrene molecule emits at approximately 370 nm and the excimers emission is red shifted to approximately 470 nm. Thus, suitable molecules, able to form excimers resulting from binding of the target phosphate would become unique phosphate sensors. (For more information see Chapter General Introduction, section Monomer/Excimer-based fluorescent probes)

4.1.1. Excimer ﬂuorescence based chemosensors

In 2011, Ni et al. reported a ratiometric fluorescent chemosensor L for inorganic phosphate which uses an excimer emission ”on-off” to ”off-on” switching mechanism in neutral solution.[1] The pyrene moieties are linked to a homooxacalix[3]arene scaffold via tria- zoles. In Figure 4.1 the molecular structure of chemosensor L is shown. Pre-organised conformation is obtained by the calixarene moiety facilitating the anion recognition event. Coordination of the Zn2+ (50 equivalent (eq.)) ion to the nitrogen atoms of the triazole moieties, prevents the pyrenyls to maintain π-π-stacking. Thus, decreasing excimer fluorescence (470 nm, ”on-off”) and increasing monomer emissiom (370 nm) can be observed in MeCN/DCM (1000:1, v/v).

Figure 4.1.: Design inspiration L for preorganized chemosensors. Molecular structure of chemosensor 2+ 2+ - L and emission spectra of L with added Zn and subsequently Zn and H2PO4 in MeCN/DCM (1000:1, v/v). Picture taken from reference. [1]

72 4.1. Background

- The emission can be switched back on by the addition of H2PO4 but not by other anions - - - [1] - 2+ such as F , NO3 , CH3COO . Selectivity for H2PO4 is only given when Zn is present. The design of selective receptors for phosphorylated molecules is initially inspired by reversible binding to Zn2+ ions in the active binding sites of metalloenzymes. The vacant coordination sites of Zn2+ bound to the ligand, enables phosphate coordination.

In 2007 Hong and co-workers developed a chemosensor for the alarmone (p)ppGpp and called it PyDPA (33). The chemical structure and binding composition of this chemosensor is depicted in Figure 4.2. Each of the two sensor molecules 33 binds with the DPA·Zn2+ to a pyrophosphate moiety at the 3’-and 5’-hydroxyl positions of (p)ppGpp.

O O

O ppGpp N

N N 2 N 2+N O N N N O Zn 2+ P O N 2+N Zn O N N N Zn - O O 2+ N 2+N O O HP Zn N P N Zn O P O O 2+ O- O N Zn O HO O 33 N N N

H2N N O H

Figure 4.2.: Binding conformation between PyDPA 33 and the alaramone ppGpp. Two sensor molecules can bind to each pyrophosphate moiety enabling π-π staking of the pyrenes and increased excimer emission in water. Picture reproduced from reference. [2]

Due to this binding the two pyrene moieties of the chemosensors come close to each other enabling π-π stacking hence generating excimer ﬂuorescence. Excimer emission is only observed with (p)ppGpp but not with other nucleotides, such as ATP, GTP, cGMP or PPi.

73 4. Excimer based sensing

4.2. Results and Discussion

This project is divided into three approaches utilising excimer ﬂuorescence - tripodal- , linked- and PyDPA- approach. Each part will be discussed in a separate section in the following chapter.

4.3. Synthesis of excimer based chemosensors

4.3.1. Tripodal chemosensors

The amino acid lysine was used as a linker between the binding- and sensing moiety. This approach allowed ﬂexibility in sensor design and peptide chemistry (see Scheme 4.1). Platform 38 is incorporated to the molecule due to the steric interactions of the adjacent substituents at the benzene ring. This favours an alternating arrangement of the sensor arms, thus building a binding cavity.

Starting from Fmoc-Lys(Boc)-OH (39) compound 40 was formed by coupling bis(2- pyridylmethyl)amine (DPA) with 41 using TBTU and DIPEA in DCM yielding the product with 58%. Cleaving the Fmoc group with piperidine afforded 42 which was further reacted with 1-pyreneacetic acid (41) using the same conditions as for the first peptide coupling step. The tert-butoxycarbonyl (Boc) deprotection was performed by stirring 43 in DCM with TFA. The linker was coupled with 44 by stirring propargylacetic acid (45) in DCM with TBTU and DIPEA. Click chemistry of alkine 46 with azide 38 was performed with CuSO4 and sodium ascorbate in a mixture of THF and TEEA buffer (100 mM, pH 7). Purification by column chromatography gave 85 mg of uncomplexed chemosensor 47 (Figure 4.1) in six steps with an overall yield of 23%.

74 4.3. Synthesis of excimer based chemosensors

N N N N N H N O N N R OH O 2 N O O 48 N R2 N H O N H TBTU, DIPEA Piperidine H = R2 HN HN DCM, RT, overnight DCM, RT, 3 h H2N R1 O O 40 42 (58%) (93%) = R 1 39 OH

O TBTU, DIPEA

DCM, RT, overnight 41

O N N N N HO N N O N R N 45 N O 2 O N O N NH2 H H TBTU, DIPEA TFA HN HN HN DCM, RT, overnight DCM O 46 O 44 O (73%) 43 (88%) (90%)

CuSO4 * 5 H2O, Na-Ascorbate, TBTA O N3 N3 O NH THF/buffer, 60 °C, overnight N O N 38 O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N

47 (75%)

Scheme 4.1: Synthesis overview of 47 in six steps with an overall yield of 23%.

Jung et al. reported 2012 an imidazolium receptor for Ins(1,4,5)P3 which could serve as a turn-off sensor. Scheme 4.2 shows the synthesis of the chemosensor. Refluxing 1- (pyren-1-ylmethyl)-1H -imidazole and 1,2,4,5-tetrakis(bromomethyl)benzene in acetonitrile gave the final sensor.

75 4. Excimer based sensing

Scheme 4.2: Synthesis of chemosensor 27 from Yoon and co-workers. Picture taken from reference. [3]

The idea was to incorporate imidazolium 49 into chemosensor 50 (see Scheme 4.3), which can serve as a binding moiety and might provide electrostatic repulsion at the same time. Scheme 4.3 shows the attempted synthesis of chemosensor 50. Following a literature known synthesis[4], 51 was synthesized by reacting 1-bromomethylpyrene (52) with 3.5 eq. of di(1H -imidazol-1-yl)methane (49) in THF. The product precipitated from the reaction mixture in 69% yield. However, reproducing this protocol was not always successful due to inseparable impurities caused by 49. No product formation could be observed by the reaction of the monoimidazolium ion 51 with the platform 53 in DMF or THF. This might be due to the bad solubility of 51.

Br Br Br N N Br Br N N N N N N Br N N 49 N 53 Br N Br N N THF DMF Br N 52 51 N N (69%) N N Br N

Scheme 4.3: Attempted synthesis of positively charged imidazol-bridged tripodal molecule 50.

76 4.3. Synthesis of excimer based chemosensors

4.3.2. PyDPA chemosensor

The above mentioned chemosensor PyDPA (33) developed by Hong and co-workers inspired us to further investigate the ability of PyDPA in PP-InsPs and polyP sensing. Therefore, the reported synthesis of chemosensor 33 was reproduced (Scheme 4.4).[5]1

O O HO OH 55 O cat. piperidine OH LiAlH4 OH Pyridine O THF

54 56 57 (52%) (39%)

DIAD, PPh3

MeO2C CO2Me THF, -78 °C → 70 °C 58 1) LiAlH4 THF

2) PBr3 O THF/DCM (2:1) O O Br OMe

60 Br 59 O OMe (49%) (63%)

N N H N DMF 48

K2CO3, KI

N N N 2+ Zn N N Zn(NO3)2⋅ 6 H2O O N O MeCN

N N N N 2+ 61 33 Zn (96%) (quant.) N N

Scheme 4.4: Synthesis overview of the reported chemosensor PyDPA 33. The product was obtained in six steps with an overall yield of 6%.

1Synthesis of chemosensor PyDPA was supported by Sophia Rauscher during her Master’s thesis.

77 4. Excimer based sensing

Starting from 1-pyrenecarboxaldehyde (54) compound 56 was obtained by a Knoeve- nagel condensation with malonic acid (55). This was followed by reduction of the double bond and the carboxylic acid yielding the alcohol 57. This step is suffering from low yield (14%) due to difficulties in purification with flash column chromatography. Subsequent Mitsunobu reaction of 57 with dimethyl-5-hydroxyisophthalate (58),

PPh3 and DIAD in THF gave compound 59 in 63% yield. The alcohol was produced by reduction of 59 with LiAlH4 in THF. The crude alcohol was directly subjected to bromination using PBr3 yielding alkylbromide 60 with 49% yield in two steps. Reaction of 60 with 2.0 eq. DPA in DMF aﬀorded 74 mg of the uncomplexed chemosensor 61 in

6% yield. The ﬁnal step was the complexation with Zn(NO3)2· 6H2O in MeCN.

4.3.3. Linked chemosensor - Intramolecular excimer formation

Chemosensor PyDPA (33) could be successfully applied as a sensor for PP-InsPs and (p)ppGpp. However, this sensing concept possesses a ratio dependency. For every analyte molecule two sensor molecules have to be present (sensor/analyte 2:1) to obtain the optimal excimer emission. If this ratio is exceeded or falls short the emission intensity decreases. To produce optimal output the exact concentration of analyte has to be known. A possible solution could be to install two pyrene moieties in one molecule. The so designed chemosensor could act in a tweezer manner thus providing a 1:1 sensor/analyte ratio. This could circumvent the ratio dependency making it possible to get a reliable output with unknown analyte concentrations. A schematic drawing of this concept is shown in Figure 4.3.

DPA-Zn2+ linker sensing unit spacer binding unit

DPA-Zn2+

Figure 4.3.: Schematic design of a chemosensor with linked pyrene moieties and DPA· Zn2+ moieties.

78 4.3. Synthesis of excimer based chemosensors

Scheme 4.5 shows an overview of the attempted synthesis of linked chemosensor 62. Starting from Boc-Gly-OH (63), peptide coupling with DPA aﬀorded compound 64 in 63% yield. Subsequent Boc deprotection with TFA gave 65 in 72% yield. Having compound 65 in hand the pyrene scaﬀold was attached in the next step.

N N H N

O 48 O O H DIPEA, TBTU H TFA O N N H2N OH R N N DCM DCM O N N N N 63 = R 64 65 (63%) (72%)

H H N N Br 6 Pd(OAc)2 Na t-butoxide dppf N N N N Toluene, 100 °C, N N Br H H overnight O O N 66 N 62

H2N NH2 6 N 68 N N H O N 67 (63%)

Scheme 4.5: Attempted synthesis of linked chemosensor 62.

To access a suitable building block for an linked chemosensor a monofunctionalization of 1,6-dibromopyrene was achieved with a Buchwald-Hartwig cross coupling of 65 and

66. The reaction was carried out in toluene with Palladium((II) acetate) (Pd(OAc)2) as catalyst, 1,1’-Bis(diphenylphosphino)ferrocen (dppf) as a ligand and sodium t-butoxide as base. The mono-coupled product 67 was obtained with 63% yield. Coupling between the pyrene moiety 67 and 1,8-octanediamine (68) was attempted to aﬀord the chemosensor 62. However, coupling of the amine with the pyrenylbromide 67 was not successful.

The reaction was screened with three diﬀerent palladium catalysts Pd(OAc)2, Palla- dium(0) bis(dibenzylideneacetone) (Pd(dba)2) and Pd(dppf)Cl2 in toluene and dioxane

79 4. Excimer based sensing with sodium tert-butoxide as base and dppf as ligand. No product formation could be observed in all cases.

As a chemosensor can ideally be synthesized in few and high yielding steps we changed our strategy of chemosensor design towards utilizing peptide coupling reactions.

O O O O HO OH HO 6 OH N 72 69 TBTU, DIPEA N TBTU, DIPEA N O O NH2 DCM DCM N H O O O O 44 HN HN 6 NH NH

N N N N N N HN N HN N NH NH O O O O O O N N O N N O

73 70 (85%) (quant.)

Zn(NO3)2 * 6 H2O Zn(NO3)2 * 6 H2O

DCM/MeOH (2:1) DCM/MeOH (2:1)

O O O O

HN HN 6 NH NH

N N N N 2+ N 2+ 2+ N N 2+ N HN Zn Zn NH HN Zn Zn NH O O O O O O N N O N N O

- - 74 4 NO3 71 4 NO3 (quant.) (quant.)

Scheme 4.6: Synthesis of linked chemosensors 71 and 74 utilizing peptide synthesis strategy.

As starting point for the synthesis of an linked sensor building block 44 can be obtained in four step with an overall yield of 43% (see Scheme 4.1). The free base of amine 44 could be coupled with glutaric acid (69) and sebacic acid respectively (72) to give chemosensor 70 and 73. Coupling conditions were TBTU as coupling reagent and DIPEA as base in DCM. Complexation with Zn2+ in DCM/MeOH 2:1 yielded 71 and 74 in quantitative yield.

80 4.3. Synthesis of excimer based chemosensors

Following this approach a sensor molecule with a hydrophilic linker should be synthesized to prevent the sensor molecule from coiling up in polar solvent (Scheme 4.7). Starting from commercially available Fmoc-Glu(OtBu)-OH (75) coupling of DPA (48) aﬀorded protected compound 76. Fmoc deprotection with piperidine gave the acid 77 which could be coupled with 1-pyrenecarboxylic acid (78) to obtain 79 with 76% yield.

= R2 N N OH N N O O H N N N O 48 O O N N HN R2 O O TBTU, DIPEA R2 O Piperidine O HN O DMF DCM H2N O R1

= R1 76 77 (98%) (80%)

75 O OH TBTU, DIPEA

DCM O O O O O N N O O N N 3 78 O NH O NH N N

N N

O HO 3 OH N N N N O OH O O 81 R2 EDC, DMAP HN O TFA HN O DCM O DCM O

80 79 (31%) (97%)

Scheme 4.7: Synthesis attempt of an PEG linked sensor

Deprotection of the tert-butyl protecting group with TFA in DCM aﬀorded 80 which was directly used without further puriﬁcation in the next step. Coupling of tetraethylene glycol (81) should give the free chemosensor 82. However, no product formation could be observed and the chemosensor was not obtained.2

2Syntheses of chemosensors in this chapter were supported by Stephan Mundinger.

81 4. Excimer based sensing

4.4. Sensor evaluation

Linked chemosensor PyDPA

O O N N 2+ Zn HN n NH N O N N N 2+ 2+ N HN Zn Zn NH O N O O N N N O 33 Zn 2+ N n = 1 71 n = 6 74

Tripodal chemosensor

O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N

Scheme 4.8: Overview of synthesized chemosensors in this chapter.

Having these chemosensors (Scheme 4.8) in hand their photophysical properties and abilities to act as chemosensors for the analytes: inositol phosphates, polyP and MSN were evaluated.

4.4.1. Photophysical properties

As these chemosensors were designed based on the ability of pyrene to form excimers thus acting as ratiometric sensors our focus was set on measuring the excimer/monomer

82 4.4. Sensor evaluation

ratio (I470 nm/I370 nm). A value above one implies a higher excimer ﬂuorescence compared to the monomer ﬂuorescence and vice versa.

4.4.1.1. Tripodal chemosensors

To utilise the above mentioned approach, from Ni et al., for phosphate sensing we drafted a design which is shown in Figure 4.4. In place of calixarene the trialkylbenzene motif as a platform was used. For the binding event of the anionic guest molecules metal- phosphate coordination interactions as the main binding force was chosen. In order to connect the sensing arms to the platform a suitable linker was chosen to employ the advantages of click chemistry giving the tripodal sensor 83

Figure 4.4.: Design for the pre-organized ratiometric phosphate chemosensor 83.

Having this concept in hand, we were able to synthesize the first candidate 83 from the preorganized chemosensors. As a linker we used 2-bromo-N -(prop-2-yn-1-yl)acetamide with cyclen as binding unit. The sensing concept is shown in Figure 4.5. Neither the exact binding of Zn2+ or analyte, nor the correct positions of the pyrene units is known. Figure 4.5 is therefore being used for illustrative purposes only. The uncomplexed chemosensor possesses a pyrene excimer fluorescence due to the flexibility of the pyrenyl units. In complex with Zn2+ this excimer fluorescence is turned off. Upon binding the analyte, excimer emission is switched on again, indicated by the red pyrene units.

83 4. Excimer based sensing

N N N

NH HN NH HN NH HN N N N N N N N N N Zn2+ - - NH NH A NH A HN NH HN NH HN NH O HN O HN O HN N N N NH N NH N NH N O O O O Zn2+ O 2+ O HN N HN N HN Zn N NH NH NH N N N N N N N N N N N N N N N N N N N N N N N N

Figure 4.5.: Design for the pre-organized ratiometric phosphate chemosensor 83.

In principle the sensing mechanism of this molecule was confirmed by experimental evidence (See Figure 4.6) but some results remain elusive: Firstly, the need of 1000 eq. Zn2+ to switch off the excimer fluoresence is disproportionately high as one could expect four eq. as sufficient, binding into the cyclene units and in between the triazole moieties. Secondly, the missing isoemissive point (indicated in Figure 4.5 with an green cycle) indicates more than one complexing mechanism for Zn2+ due to multiple binding sites (Cyclen, peptide bond and triazole) in the chemosensor.[6] An iosemissive point reflects a specific wavelength, where the emission of light does not change.[7]3

Figure 4.6.: Proof of concept of the ”on-off” to ”off-on” switching mechanism for chemosensor 83. Free sensor (0.2 µM, 10 mM 1,4-Piperazinedipropanesulphonic acid (PIPPS) pH 4.7, grey curve), complexed 2+ with Zn (1000 eq., black curve), under influence of sodium salt of InsP6 (1.0 µM, 5 eq., red curve). λEx = 330 nm.

In order to circumvent the need of 1000 eq. Zn2+ of chemosensor 83 we designed and synthesized sensor 47 using DPA as a binding unit instead of cyclen. Having sensor

3For further information see the Master’s thesis of Ann-Kathrin Mündler

84 4.4. Sensor evaluation

47 in hand the absorption spectrum of 47 (1.2 µM) was measured in HEPES (10 mM, pH 7.4) to give absorption maxima at 268, 280, 330 and 350 nm from which 330 nm was chosen as excitation wavelength for further ﬂuorescence measurements. Second, the uncomplexed sensors (12 nM) emission was measured in HEPES (10 mM, pH 7.4,

λex = 330 nm) (Figure 4.7, black). For the ﬂuorescence measurements a 1 mM stock solution in DMSO was stored in the freezer and working solutions (12 nM in HEPES) were freshly prepared before the measurements. It is recommended to use a freshly prepared stock solution from the solid because the molecule suﬀers from emission intensity loss when stored in the freezer in DMSO for several days.

Subsequently various equivalents of Zn(ClO4)2 were added to clarify whether the sensors excimer emission is aﬀected, upon metal binding.

O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N 47

Figure 4.7.: Emission spectra of sensor 47 (12 nm in 10 mm HEPES, black) and the eﬀect of Zn2+ (10, 100 and 1000 eq.). λEx = 330 nm.

In the emission scan of the uncomplexed sensor 47 (Figure 4.7, black) the excimer emission intensity is low and not further aﬀected by addition of 10, 100 or 1000 eq.

2+ Zn (Figure 4.7, red, blue, green). Similarly, the addition of analyte (PPi, InsP6 and

5-PP-InsP5 with diﬀerent equivalents) had no eﬀect on the excimer nor the monomer emission.

85 4. Excimer based sensing

This could be interpreted as the disability of the pyrene moieties to come in close proximity. Although rotation of the chemosensors arms is restricted through the platform they might be too ﬂexible and π-π-stacking does not occur. Another possibility is that the anion binding is not occurring. Basically, the binding units within this chemosensor are amides and these are not well suited for complexation of Zn2+. Without a complexed metal no binding event will take place.

4.4.1.2. Linked chemosensors

Along with tripodal sensor 47 another approach was tested - the linked chemosensors 71 and 74. Both molecules bears the same components as 47 which are linked via peptide bonds with an aliphatic backbone diﬀering in chain length (see Figure 4.8b). From the the absorption sepctra of both sensors 71 and 74 (50 µM in HEPES 10 mM, pH 7.4) the excitation wavelength was set to λEx = 330 nm. Emission measurements were carried out with a sensor concentration of 5 nM in HEPES and with addition of various equivalents of analytes (Figure 4.9). An emission intensity loss over time was observed (Figure 4.8) so it is appropriate to use freshly prepared working solutions.

12000

0 min

10 min

10000

20 min

30 min O O

8000 40 min

HN n NH

6000

time 4000 N N

Fluorescence Intensity [RFU] Intensity Fluorescence N 2+ 2+ HN Zn N 2000 Zn NH O O O N 0 N O

400 500 600 700

Wavelength [nm] n = 1 71 n = 6 74

(a) (b) Figure 4.8.: a) Emission spectra of 71 (5nM in HEPES 10 mM, pH 7.4) over time. The inset shows I470 nm/I370 nm. λEx = 330 nm λEm = 350-750 nm. b) Chemical structure of chemosensors 71 and 74

For every set of measurement a new working solution was prepared from a 100 µM stock solution in DMSO which was stored in the freezer. However, a slight decrease in overall emission intensity could be observed during the measurements. Considering

86 4.4. Sensor evaluation

I470 nm/I370 nm a trend towards increasing of the monomer emission can be seen in the fluorescence spectra of uncomplexed sensor (Figure 4.8a inset). The next step was to investigate whether addition of anions might result in an increase of the excimer fluorescence. In Figure 4.9 the effect of added anions is depicted.

Four diﬀerent anions were selected: PPi (4.9a), InsP6 (4.9b), 5-PP-InsP5 (4.9d) and pppGpp (4.9c).

(a) PPi (b) InsP6

(c) pppGpp (d) 5-PP-InsP5 Figure 4.9.: Emission scans of 71 (5nM in HEPES 10 mM, pH 7.4, black) with added anions: a) PPi (10, 100 and 1000 eq.). b) InsP6 (10, 100 and 1000 eq.). c) pppGpp (10, 100 eq.). d) 5-PP-InsP5 (10, 100 and 1000 eq.). λEx = 330 nm, λEm = 350-750 nm.

Addition of pyrophosphate PPi resulted in continuous decreasing of excimer emission

(Figure 4.9a). With InsP6, 5-PP-InsP5 and pppGpp a small enhancement of excimer emission (Figure 4.9b, c and d red curve) was observed upon the ﬁrst addition of 10 equivalents. However, the increase is too low to be signiﬁcant and addition of further equivalents of anions resulted in decrease of the excimer emission and continuous enhancing of monomer emission (Figure 4.9). This can be attributed to the beforehand observed behaviour of the free sensors emission intensity loss and indicates that anion

87 4. Excimer based sensing addition does not have any impact on the sensors emission. Fluorescence response to added anions was investigated for sensor 74 following the same procedure as described above for 71. Again, the emission intensity of the sensor decreased over time. The reason behind this observation is not known, but it might be an equilibrium reached by the sensor over time with distant pyrene units. Addition of PPi had no eﬀect on the sensors emission spectra (Figure 4.10a) as well as addition of the inositol phosphates. Only pppGpp resulted in a slight increase in excimer emission. In Figure 4.10c this eﬀect can be observed, however the small increase in excimer emission can not be further increased by adding more equivalents of analyte. However, both chemosensors are not suitable to act as inositol phosphate, pppGpp or polyP (not shown) chemosensor.

(a) PPi (b) InsP6

(c) pppGpp (d) 5-PP-InsP5 Figure 4.10.: Emission scans of 74 (5nM in HEPES 10 mM, pH 7.4) with added a) PPi (10, 100 and 1000 eq.). b) InsP6 (10 and 100 eq.). c) ppppGpp (10 and 100 eq.). d) 5-PP-InsP5 (10 and 100 eq.). λEx = 330 nm, λEm = 350-750 nm.

88 4.4. Sensor evaluation

4.4.2. PAGE-Gel staining

The first step was to determine the minimium staining time. As analyte InsP6 was chosen as it is commercially available and previous fluorescence measurements revealed that inositol phosphates showed a higher increase in excimer emission intensity than (p)ppGpp. A gel was prepared according the general procedure (see General remarks) loaded with InsP6 (150 µM, 22.5 nmol) and run in TBE buffer for 4 hours (hs) at 500 V. The gel was washed for 30 minutes (mins) in deionized water to remove EDTA and subsequently placed into the PyDPA (50 µM in water) staining solution. The staining process was followed with a transilluminator plate at λEx = 312 nm. After 20 mins dark bands could be observed (Figure 4.11).

Figure 4.11.: PAGE of InsP6 (150 µM) in TBE buﬀer for 4 hs and stained with PyDPA (50 µM in water) for 1 h. Pictures were taken every 10 mins for 60 mins and the gel was excited at λEx = 312 nm

89 4. Excimer based sensing

As EDTA can decomplex Zn2+ from the DPA binding units, thus suppressing the binding of PyDPA to the analyte, from this point on further PAGE were run in Tris/Borate (TB) buffer. This has not been done before and gels run according the general procedure (20 hs, 500 V) in TB buffer indicated that EDTA is not essential for the separation. In the next experiments the ratio dependency shall be addressed. Previous fluorescence measurements revealed that a PyDPA/analyte ratio of 1:2 results in the highest emission intensity for inositol phosphates. However, when the ratio is exceeded the signal intensity is reduced. In Figure 4.12 PyDPA (50,40 and 30 µM) stained PAGE gels are shown as well as a table which summarizes the lanes with loaded concentration of 5-PP-InsP5 (ranging from 100 µM (3 nmol) to 1 µM (0.03 nmol)), the concentration of the stain and the Py-

DPA/analyte ratios. 5-PP-InsP5 was chosen as it showed the highest emission intensity in the initial ﬂuorescence measurements. The standard procedure, used for the separation of inositol phosphates, requires a run time of 20 hs. For the next experiments shorter run time of 4 hs was chosen because the staining intensity in the gel matrix was of interest but not yet the separation of analytes. However, the poor resolution of the bands and the deteriorated band shape might be a result of the reduced run time. The added reference compound Orange G Dye (OG) can be seen as dark bands and the stained analyte is visible as a bright blue band (Figure 4.12). Staining with 50 µM PyDPA (Figure 4.12A) results in a blue band for the complete concentration series. It is even possible to distinguish a thick band and some slightly smaller bands underneath.

These might be degradation products from 5-PP-InsP5, such as InsP6 or InsP5 from the sample. In Figure 4.12 B the gels are stained for 15 mins with 40 µM PyDPA. With a lower concentration of the staining solution, 50 and 40 µM 5-PP-InsP5 (lane 8 and 9) can not be stained. Consistent with previous ﬂuorescence measurements, PyDPA/analyte ratio of approx. 1:2 gave visible bands with 30, 20 and 15 µM (lane 10,11 and 12 in Figure 4.12B) 5-PP-InsP5, but lower concentrations of 5-PP-InsP5 could not be stained.

90 4.4. Sensor evaluation

Figure 4.12.: PAGE gel run in TB buﬀer for 4 hs, stained with PyDPA and recorded on a transilluminator at λEx = 312 nm. A) 5-PP-InsP5 in sequence from left to right: 100 µM (3 nmol), 50 µM (1.5 nmol), 40 µM (1.2 nmol), 30 µM (0.9 nmol), 25 µM (0.75 nmol),20 µM (0.6 nmol) and 15 µM (0.45 nmol). stained with PyDPA (50 µM in water). B) 5-PP-InsP5 in sequence from left to right: 50 µM (1.5 nmol), 40 µM (1.2 nmol), 30 µM (0.9 nmol), 20 µM (0.6 nmol), 15 µM (0.45 nmol), 10 µM (0.3 nmol), 5 µM (0.15 nmol) and 1 µM (0.03 nmol). Stained with PyDPA (40 µM in water). C) 5-PP- InsP5 in sequence from left to right: 15 µM (0.45 nmol), 10 µM (0.3 nmol), 5 µM (0.15 nmol) and 1 µM (0.03 nmol). Stained with PyDPA (30 µM in water). All informations are summarized in the table.

Reduced stain concentration to 30 µM successfully stained 10, 5 and 1 µM (lane 17,18 and 19 in Figure 4.12C). Interestingly, lane 16 was not stained although the Py- DPA/analyte ratio was 1:2 whereas the lower concentrated bands are slightly visible.

The above discussed results underline the feasiblity and convenience fo staining PAGE with PyDPA. This approach could reach low limits of detections and the developed procedure can assure facile handling.

4.4.3. Post column derivatisation - Ion Chromatography

Ion exchange chromatography is a tool for separating charged molecules based on their aﬃnity to the column material. The stationary phase of strong anion exchange chro-

91 4. Excimer based sensing matography is an ion-exchange resin, typically described as modiﬁed quaternary ammonium groups, and partitioning occurs with ion exchanges between the analyte and the stationary phase.

Until today the common used method for detecting inositol phosphates after HPLC separation is scintillation counting of previously tritium labelled compounds.[8] In 1988 Mayr et al. developed the MDD method using trivalent transition metals which allowed them to analyse non-radioactively labelled cells via UV-detection.[9] However, separation of higher phosphorylated inositol phosphates is not possible with this method. Recently,

Shears and co-workers published a buffer system which is able to separate InsP6, InsP7 - [10] and 1,5-InsP8 using NO3 as the eluting anion at pH 4.7. Inspired by this research we envisaged to develop a post column staining method which relies on fluorescence detection. Ion chromatography combined with fluorescence detection has never been done before and this approach would enable to combine and use the benefits of both methods. High separation power combined with high sensitivity. Sensi- tivity of fluorescence measurements is greater than absorption photometric methods.[11] Thus, potentially leading to lower LoD and the usage of less sample. Additionally only fluorescent species are detected resulting in greater specificity compared to absorption measurement. For developing a suitable IC method some variables can be changed:

eluent composition/concentration/pH

column

analyte detection

ﬂow rate

Our concept is shown in Figure 4.13. The two buffer eluents are prepared separately and used for elution of the target analyte off the anion exchange column. The sample is injected and pumped through the analytical column where it comes to affinity based

92 4.4. Sensor evaluation separation of the anionic species. Simultaneously a dye solution (PCSR) is pumped into the system by a separate pump.

Figure 4.13.: Concept of post column derivatisation of polyanionic analytes (here with the example of ppGpp) via PyDPA

At this point the eluted anions from the column and the sensor solution are combined in a mixing-coil (Figure 4.14). This allows analyte and chemosensor to react with each other for approximately 1.5 mins to give a measurable ﬂuorescent output. The emission is measured by a downstream Fluorescence detector (FLD).

Figure 4.14.: Picture of a mixing coil with indicated incoming analyte from the column and PCSR from an external pump (AXP). Analyte and stain combine in the mixing coil for approximately 1.5mins to form the excimer ﬂuorescence which is measured in the FLD.

93 4. Excimer based sensing

As we know that the buffer system, used by Shears and co-workes[10], can separate higher inositol phosphates this buffer was used for initial measurements. Chemosensor PyDPA should be used as the postcolumn staining reagent (PCSR) to stain analytes which were beforehand separated on an anion-exchange column. Dionex CarboPac PA- 100 (4 x 250 mm) analytical column from Thermofisher was used as this column is specialized for the separation of neutral and anionic oligosaccharide mixtures. The idea was to utilize the ability of the pyrene-moieties, to form excimer emission which possess a shifted fluorescence emission compared to the monomer fluorescence to lower the background fluorescence.

Next, it should be investigated whether the sensor is working in the before mentioned buﬀer system from Shears and co-workers.[10] In the buﬀer contained MeOH slows

- down the elution, thus improving separation of the analytes. NO3 is the eluting anion, by increasing the nitrate concentration resin bound analytes are eluted from the column. Since the requirement of EDTA in the buﬀer is unclear and because it would decomplex

2+ Zn from the DPA moieties, thus preventing binding of the PyDPA to InsP6 the buffer was prepared without EDTA. Both buffers were prepared separately and fluorescence measurements were carried out with PyDPA (5µM) in both buffers with added InsP6 (2.5 µM). In Figure 4.15 it can be seen that in Buffer C the excimer formation of the pyrene moieties can occur (Figure 4.15, red).

150

Buffer C

Buffer C with IP

Buffer D

Buffer D with IP

100

50 Fluorescence Intensity [RFU] Fluorescence

350 400 450 500 550 600 650

Wavelength [nm]

Figure 4.15.: Emission Scans of PyDPA (5 µM) in Buﬀer C and Buﬀer D with added InsP6 (2.5 µM). λEx = 344 nm, λEm = 350-650 nm.

94 4.4. Sensor evaluation

- 2+ The NO3 , might compete with InsP6 for binding to the Zn -DPA as only slight excimer formation can be observed (Figure 4.15, green). However, as the anions are eluted using

- a gradient slowly increasing the NO3 concentration this should not be an issue.

Next it should be investigated whether this chemosensor can be applied as a post column staining reagent for inositol phosphates on the IC system. Direct injection of a separately prepared solution of InsP6 (25 µM) and PyDPA (50 µM) to the system without an analytical column, using water as the eluent, gave a peak with high intensity at

λEm = 460 nm. This proofed the utilization of recording the excimer emission with the FLD. As the emission intensity in the PIPPS buﬀer decreased a concentration test was carried out in 100% Buﬀer C.

Figure 4.16.: Stacked chromatograms of varying InsP6 concentrations (10 µM, blue; 7.5 µM, orange; 5 µM, cyan; 2.5 µM, black; 1 µM, brown; 0.8 µM, green.) mixed with 5 µM PyDPA. No analytical column, ﬂow rate 0.5 mL/min, isocratic in 100% Buﬀer C, λEx = 344 nm and λEm = 460 nm

Figure 4.16 shows stacked injections of a mixture of PyDPA (5 µM) and InsP6 in different concentrations without an analytical column measured in 100% Buﬀer C. The stacked chromatograms conﬁrm the ratio dependency as the highest emission intensity is achieved with PyDPA (5 µM) and InsP6 (2.5 µM) which corresponds to an 2:1 Py-

DPA/InsP6 ratio (Figure 4.16, black). As the proportion of InsP6 increases (Figure 4.16, blue, orange and cyan) the intensity decreases with a factor of 2 and higher pro- portions of PyDPA (Figure 4.16, blue and red) result in an emission intensity 5 times lower.

95 4. Excimer based sensing

The analyte concentration of 150 µM InsP6 was used with an injection volume of 10µL and a flow rate of 5.0 mL/min. Staining solution containing 10 µM PyDPA was prepared separately and pumped to the system with 0.2 mL/min. Both solutions are allowed to combine in the mixing coil for approximately 1.5 mins. The sample was injected and eluted from the column with Buffer C/Buffer D (20%/80%.) Recording the fluorescence signal with λEx = 344 nm and λEm = 400 and 460 nm resulted in the chromatogram shown in Figure 4.17. The black curve shows the excimer emission raising at 4.50 mins with simultaneously decreasing the monomer emission signal at 400 nm (green curve).

Figure 4.17.: Proof of concept: Detection of InsP6 (150 µM) with PyDPA (10 µM) after elution from the analytical column (Dionex PA-100) with a flow rate of 0.5 mL/min, BufferC/BufferD (20%/80%). λEx = 344, λEm = 400 and 460 nm.

Attempts to separate InsP6 and 5-PP-Ins5 with this approach were made but until now this goal could not be achieved. We could successfully proof our concept (Figure 4.13) of detecting inositol phosphates injected and eluted from a CarboPac PA-100 column with a buffer system (BufferC/BufferD (20%/80%)). Subsequently the analyte was stained with a PyDPA (10 µM) staining solution in a mixing coil.

96 4.5. Summary & Outlook

These experiments laid the foundation for a new analytical method which uses the separation power of an ion chromatography system with the high sensitivity of ﬂuorescence detection.

4.5. Summary & Outlook

In this chapter the synthesis, photophysical evaluation and first steps towards applications of a chemosensor for inositol phosphates were presented. In summary, four potential chemosensors in three categories (tripodal chemosensor, linked chemosensor and PyDPA) were synthesized. The approach of a tripodal chemosensor should provide preorganization of the sensor molecule with the trialkyl benzene platform. Fluorescence output should be achieved by pyrene forming excimers upon the analytes binding event as the shifted excimer emission pyrene, allows for fluorescence detection against low background. To ensure high affinity to oxyanions in aqueous media metal-phosphate coordination was chosen. With this design concept two sensor molecules were successfully synthesized during the course of this thesis (Figure 4.18).

N O 2+ O NH N Zn N O N N N N O NH 2+ O 2+ Zn HN N Zn N 2+ N O N Zn O N N N N 2+ N NH N Zn N HN 2+ N Zn O O N O N HN N HN NH HN N N N O N 2+ N N O Zn N N N N N N 2+ N Zn N N N N 83 47 (a) (b) Figure 4.18.: a) Chemical structure of chemosensor 83 b) Chemical structure of chemosensor 47 with proposed binding mode to Zn2+.

The excimer emission of sensor 83 could be suppressed by adding Zn2+, and switched back on upon analyte binding (InsP6) in HEPES (10 mM, pH 7.4). With that we

97 4. Excimer based sensing developed an on-off-on chemosensor with an affinity to inositol phosphates applicable in aqueous solution. However, the need of extensive amounts of Zn2+ remains puzzling . Sensor 47 was designed in a similar way as the preceding probe 83. In this case no excimer fluorescence was formed, thus the monomer emission remained higher. Addition of metal and analyte did not lead to significant changes in the emission profile.

The linked chemosensors should provide a sensor/analyte 1:1 binding and sensing ability due to the incorporation of two pyrene units into one molecule. Direct coupling of two 1-bromo-pyrene moieties, which bears the binding unit at the 6-position, was not successful. Also, the attempt to synthesize a sensor linked with PEG, to assure a linear form in aqueous solution, could not be realized. However, chemosensors 71 and 74 could be synthesized based on peptide coupling reactions (Figure 4.19). The idea was that binding of the analyte, with both DPA moieties, could bring the pyrenes close enough to support excimer formation. Fluorescence measurements of both sensors in HEPES (10 mM, pH 7.4) revealed that monomer emission of the sensor exceeds the excimer emission and even addition of 10, 100 and 1000 eq. of analyte could not induce excimer formation. However, in this design the amide might prevent coordination to Zn2+ taking the ability to act as a suitable binding unit. Additionally, during the measurements the monomer emission increased compared to the excimer emission. This could indicate that the sensor adopts a conformational state over time where no π-π-stacking occurs.

O O N N 2+ Zn HN n NH N O N N N 2+ 2+ N HN Zn Zn NH O O O N N N O 2+ N Zn n = 1 71 33 N n = 6 74

(a) (b) Figure 4.19.: a) Chemical structure of chemosensor 71 and 74 b) Chemical structure of PyDPA 33.

PyDPA (33) was published as a selective pppGpp sensor.[2] However, the attempt to use this sensor for inositol phosphates and polyP was successful. In this thesis PyDPA

98 4.5. Summary & Outlook could be applied as a stain for PAGE of inositol phosphates. Recording the excimer fluorescence on a transilluminator at 312 nm results in bright blue bands. A procedure was developed including the use of TB buffer, instead of commonly used TBE running buffer, to prevent decomplexation of Zn2+ from the DPA units. A staining time of 20 mins was evaluated which is short compared to 2 h for toluidine blue. Dilution series of 5-PP-InsP5 resulted that excimer detection at 312 nm on a transilluminator is possible down to 1 µM (30 pmol). However, the ratio dependency of pyrene is an issue that limits the applicability. Nevertheless it should be possible to estimate a concentration range of the analyte and to correspondingly adjust the molarity of the staining solution.

A promising step towards the application of PyDPA in inositol phosphate detection was made by using the sensor as a post column staining reagent. Ion chromatography (IC) has never been used in combination with fluorescence detection. The studies in this thesis constitute the first step towards a sensitive fluorescence detection of anions separated on an ion exchange column.

In future research on the linked chemosensor might provide a suitable sensor for the detection of analytes of interest. Furthermore, sensing one analyte molecule with one sensor molecule could circumvent the excimer decrease of PyDPA when a higher amount of analyte over PyDPA is present. The coordination of peptide bound DPA units to Zn2+ should be investigated in more detail and another design to attach the binding units should be found if necessary. As this sensor may not show selectivity for either pyrophosphate, inositol phosphates or magic spot nucleotides it would be a good candidate for post column staining of beforehand separated anions.

The tripodal design might introduce selectivity when the sensors cavity is less ﬂexible and suited for the anions shape, such as inositol phosphates. Pursuing this path is promising as the building block design allows the synthesis of a sensor library in future.

It is in any case worth to reﬁne the protocol for PAGE staining with PyDPA as well as with other chemosensors. The potentially time saving while sensitivity gaining protocol will open doors for the investigation of biological processes.

99 4. Excimer based sensing

The same is true for the combination of ion chromatography and ﬂuorescence detection. Up today the ratio dependency of PyDPA is an issue but it probably will be possible to solve this problem in future paving the way to new and enriching possibilities to use this powerful tool for the analysis of polyanionic compounds.

100 4.6. Experimental

4.6. Experimental

4.6.1. General remarks

Reactions were carried out in ﬂame-dried glassware under argon atmosphere, unless noted otherwise. Reaction control was performed via TLC.

Reagents were purchased from commercial suppliers and used without further puriﬁ- cation.

Solvents were dried using the “Braun Solvent Puriﬁcation System 800“ or used from commercial suppliers. Analytical grade solvents were used as received for extraction and chromatographic puriﬁcatios.

Thin layer chromatography wa performed on Merck silica gel 60 F254 plates (0.25 mm layer thickness, with ﬂuorescence indicator). TLC was analyzed by UV (λ =,254 nm), ﬂuorescence (λ =,368 nm).

Flash column chromatography was carried out using silica gel 60 (0.04 - 0.063 mm, 230 - 400 mesh) from Macherery-Nagel. The mobile phase was forced through the stationary phase using excess pressure (hand pump).

1H-NMR spectra samples were measured in the indicated deuterated solvents on a Bruker 300 MHz, a Bruker Avance 400 MHz and a Bruker 500 MHz spectrometer. Except for measurements on the Bruker 300 MHz, all spectra were recorded by Dr. M. Keller and the team from the analytical department of the Institute of Oragnic Chemistry at the University of Freibug. All signals are referenced to an internal solvent signal standard (CDCl3, δ =7.26 ppm; D2O, δ =4.79 ppm; MeOD4, δ =3.31 ppm; DMSO-d6, δ =2.50 ppm).

13C-NMR spectra samples were measured in the indicated deuterated solvents on a Bruker 101 MHz, a Bruker 126 MHz spectrometer. The spectra were recorded by Dr. M. Keller and the team from the analytical department of the Institute of Oragnic

101 4. Excimer based sensing

Chemistry at the University of Freibug. All signals are referenced to an internal solvent signal standard (CDCl3, δ =77.1 ppm; MeOD4, δ =49.0 ppm; DMSO-d6, δ =39.52 ppm).

High resolution mass spectrometry (HRMS) was performed by C. Warth from the analytical department of the Institute of Organic Chemistry at the University of Freiburg, using a Thermo LCQ Advantage (spray voltage: 2.5 - 4.0 kV; spray current: 5 µA; ion transfer tube: 250 (150) ° C, evaporation temperature: 50-400° C).

High Performance Ion Chromatography (HPIC) was performed on a Thermo Scientiﬁc Dionex ICS-5000+ HPIC system using a dual pump equipped with an 10 µL injector loop, a Dionex CarboPac PA-100 guard column (2 x 50 mm) and a Dionex CarboPac PA-100 analytical column (4 x 250 mm) and a FLD detector. Post-column reagent was pumped by an Dionex AXP Auxiliary Pump and mixed in a knitted Reaction Coil, 750 µL.

UV/Vis-Absorption Sepctra were recorded on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Transparent) or on a Shimadzu UV-1800 UV/VIS Spektrophotometer using a 2 mL cuvette.

Fluorescence Spectra were recorded on a Perkin Elmer LS55 Luminescence Spectrom- eter or on a JASCO FP-8200 Spectroﬂuorometer using a 3 mL cuvettes or on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Black).

Polyacrylamide gel electrophoresis (PAGE) PAGE was carried out on a Hoefer SE660 Tall Standard Dual Cooled Vertical Unit. The PAGE procedure was conducted according to the general procedure as described by Losito et al..[12] Following buffers and solutions for gel electrophoresis were prepared:10 × Tris/ Borate/ EDTA (TBE) buffer (0.89 M tris-HCl, 0.89 M boric acid, 20 mM EDTA, pH 8.3), 10 × Tris/ Borate (TB) buffer (0.89 M tris-HCl, 0.89 M boric acid, pH 8.3), 1 × Orange G dye(10 mM tris-HCl, 1 mM EDTA, 30 % (w/v) glycerol, 0.1 % (w/v) OrangeG, pH 7.0).

102 4.6. Experimental

During pre-run and run, the lower buffer chamber was filled with 6 L of pre-chilled 1 x TBE or TB buffer (4°C).The buffer was continuously stirred and recirculating cooler was used for chilling the buffer. Sample loading was performed with gel-loading pipet tips.

The gel sandwich was assembled by employing glass plates (24 x 18 cm) and spacers (1 cm wide, 1.0 mm thick). For the gel preparation, 35.8% (w/v) acrylamide:bis-acrylamide 19:1 (33.9 mL, 40% solution, Roth 3030), 10.0% (v/v) 10 x TBE buﬀer (3.8 mL) and

0.05%(w/v) ammonium persulfate (APS) (200 µL of 10% APS in milli-Q H2O) were stirred for 1 min at 0°C, followed by the addition of 0.05% (v/v) tetramethylethylene- diamine (TEMED) (20 µL). After stirring for 1 min, the mixture was poured between the pre-casted glass-plates and a 15 lane comb was inserted. The solution was allowed to polymerize for 25 min at room temperature. After polymerization, the comb was removed and gels were pre-run at 4°C in 1 x TBE or 1 x TE buffer for 30 min at 300 V. Samples were prepared and 1 × Orange G dye (7 µL) was added to each sample prior to loading. Wells were washed with 1 x TBE or 1 x TB buffer by using a syringe and needle to remove any precipitates and non-polymerized gel debris. The gel was then loaded and run at 4°C in 1 x TBE or 1 x TB buffer for 4 h or 20 h at 500 V. After run, the gel apparatus was disassembled and the gel was stained for 15 min with PyDPA staining solution or 30 min for toluidine blue staining with subsequent destaining for 2 h with 2-3 changes of destaining solution. Finally, the gel was recorded on a transilluminator plate with λEx = 312 nm.

Buffer C & D Table 4.1.: Composition of buffer C and D reported by Shears et al.. [10] Buffer C Buffer D

1 mM Na2EDTA 1 mM Na2EDTA 10 mM PIPPS 10 mM PIPPS pH 4.7 0.5 M NMe4NO3 MeOH (5%) pH 4.7 MeOH (5%)

103 4. Excimer based sensing

4.6.2. Synthesis

General procedure for peptide synthesis The acid (1.0 eq.) is dissolved in DMF/DCM (50 mM) and N,N,N ’,N ’-tetramethyl-O- (benzotriazol-1-yl)uronium tetrafluoroborate (TBTU) (1.0 eq./acid) is added followed by N,N -diisopropylethylamine (DIPEA) (6.0 eq.). The mixture is stirred for 30 mins until the solution turns clear. The amine (0.9 eq./acid) is added subsequently and the mixture stirred overnight at RT. The reaction is washed with NaHCO3, dried over Na2SO4 and the solvent is removed under reduced pressure and the product is purified by flash column chromatography.

General procedure for Boc deptrotection The Boc-protected amine (1.00 eq.) is dissolved in DCM (50 mM), TFA (5.0 eq./1.0 eq.Boc) is added and the mixtures stirred at RT until complete consumption of the starting material is evidenced by TLC. The solvent is removed under reduced pressure and the solid dissolved in NaOH (6N). The product is extracted with DCM and the combined organic layers dried over Na2SO4. The solvent is removed under reduced pressure.

(9H -ﬂuoren-9-yl)methyl tert-butyl(6-(bis(pyridin-2-ylmethyl)amino)-6- oxohexane-1,5-diyl)(R)-dicarbamate (40)

O N H N N O N O O HN O

Fmoc-Lys(Boc)-OH 39 (1.13 g, 2.40 mmol, 1.00 eq.) was treated with DPA (400 mg, 2.22 mmol, 0.92 eq.) according to the general procedure for peptide synthesis. Puriﬁca- tion was achieved by ﬂash column chromatography (EE/EtOH = 100:0 to 96:4) yielding the product as a yelllow solid (751 mg, 1.15 mmol, 58%).

104 4.6. Experimental

Rf (SiO2, EE/EtOH 6:1 (vol/vol)) 0.6. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.53 (dd, J = 11.3, 4.2 Hz, 2H), 7.76 (d, J = 7.5 Hz, 2H), 7.70 – 7.51 (m, 4H), 7.40 (t, J = 7.5 Hz, 2H), 7.35 – 7.27 (m, 2H), 7.25 – 7.14 (m, 3H), 5.71 (d, J = 8.4 Hz, 1H), 4.93 – 4.65 (m, 4H), 4.59 (br s, 1H), 4.34 (d, J = 7.3 Hz, 2H), 4.20 (t, J = 7.2 Hz, 1H), 3.13 – 2.99 (m, 2H), 1.88 – 1.75 (m, 1H), 1.74 – 1.62 (m, 1H), 1.57 – 1.31 (m, 13H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 173.3, 165.9, 156.7, 156.1, 155.9, 149.8, 148.7, 144.0, 143.9, 141.4 (2C), 137.6, 137.2, 127.8 (2C), 127.2 (2C), 125.3, 125.3, 122.9, 122.9, 122.8, 121.8, 120.1, 120.1, 79.2, 67.1, 53.0, 51.6, 51.2, 47.3, 40.3, 33.0, 29.7, 28.6, 22.4.

(3 Carom missing, however in molecule 42 all carbons could be detected) HRMS (ESI) m/z: calcd.: 650.3337, found: 650.3328 [M+H+]+. tert-butyl(R)-(5-amino-6-(bis(pyridin-2-ylmethyl)amino)-6-oxohexyl) carbamate (42)

N N O N O O N H

H2N

40 (1.17 g, 1.80 mmol, 1.00 eq.) was dissolved in DCM (15 mL), piperidine (3.45 g, 40.5 mmol, 22.4 eq.) was added and stirred for 2.5 hs at RT. The mixture was washed with NaHCO3 (10 mL) and the product extracted with DCM (3x10 mL). The combined organic layers were dried over MgSO4 and the solvent was removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 96:4 to 80:20) gave 42 (798 mg, 1.86 mmol, 93 %) as a pale white oil.

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.3. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.52 (dd, J = 21.0, 4.6 Hz, 2H), 7.63 (qd, J = 7.6, 1.8 Hz, 2H), 7.29 – 7.24 (m, 1H), 7.23 – 7.12 (m, 3H), 4.96 (d, J = 15.1 Hz, 1H), 4.81 (d, J = 17.1 Hz, 1H), 4.63 (d, J = 17.0 Hz, 2H), 4.51 (d, J = 15.1 Hz, 1H),

105 4. Excimer based sensing

3.93 – 3.87 (m, 1H), 3.07 (d, J = 6.2 Hz, 2H), 2.49 (br s, 2H), 1.84 – 1.70 (m, 1H), 1.66 – 1.54 (m, 1H), 1.49 - 1.29 (m, 13H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 175.6, 157.1, 156.2, 156.1, 150.1, 149.3, 137.1, 137.0, 122.9, 122.7, 122.5, 121.9, 79.2, 52.8, 51.7, 51.4, 40.3, 34.1, 29.9, 28.6, 22.9. HRMS (ESI) m/z: calcd.: 428.2656, found: 428.2657 [M+H+]+. tert-butyl(R)-(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl) acetamido)hexyl)carbamate (43)

N N O N O O N H HN O

The acid 41 (534 mg, 2.25 mmol, 1.00 eq.) was treated with 42 (798 mg, 1.86 mmol, 0.82 eq.) according to the general procedure for peptide synthesis to give the product puriﬁed by ﬂash column chromatography (pentane/EE 98:2 to 95:5) as a yellow solid (1.12 g, 1.67 mmol, quant.).

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.6. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.53 – 8.50 (m, 1H), 8.44 – 8.39 (m, 1H), 8.26 – 7.91 (m, 9H), 7.60 (td, J = 7.7, 1.8 Hz, 1H), 7.38 (td, J = 7.7, 1.8 Hz, 1H), 7.20 – 7.11 (m, 2H), 7.09 – 6.97 (m, 2H), 6.24 (d, J = 8.2 Hz, 1H), 5.10 - 5.00 (m, 1H), 4.90 – 4.50 (m, 4H), 4.41 – 4.20 (m, 3H), 2.84 -2.80 (m, 2H), 1.80 – 1.60 (m, 2H), 1.43 (s, 9H), 1.27 – 1.09 (m, 2H), 1.06 - 0.90 (m, 2H).

13 C-NMR (101 MHz, CDCl3, δ/ppm): 172.9, 170.9, 156.0, 155.8, 149.8 (2C), 137.1 (2C), 131.5, 131.2, 130.9, 129.7, 128.6, 128.6, 128.3, 127.6 (2C), 126.3, 125.6, 125.4, 125.4, 125.3, 124.8, 123.3, 122.9, 122.7, 121.9, 79.1, 53.0, 51.0, 49.4, 42.2, 40.2, 32.3, 29.4, 28.6, 22.2. HRMS (ESI) m/z: calcd.: 670.3388, found: 670.3381 [M+H+]+.

106 4.6. Experimental

(R)-6-amino-2-(2-(pyren-1-yl)acetamido)-N,N -bis(pyridin-2-ylmethyl) hexanamide (44)

N N

N O NH2

HN O

Boc deprotection of 43 (1.12 g, 1.67 mmol, 1.00 eq.) was carried out according to the general procedure for Boc deprotection to yield the product 44 (906 mg, 1.35 mmol, 81%) as a white foam.

1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.52 – 8.49 (m, 1H), 8.40 – 8.37 (m, 1H), 8.25 – 7.91 (m, 9H), 7.58 (td, J = 7.7, 1.8 Hz, 1H), 7.35 (td, J = 7.7, 1.8 Hz, 1H), 7.16 – 7.12 (m, 2H), 7.05 – 6.98 (m, 2H), 6.51 (d, J = 8.2 Hz, 1H), 5.03 (td, J = 8.1, 4.5 Hz, 1H), 4.86 (d, J = 17.0 Hz, 1H), 4.69 – 4.58 (m, 3H), 4.35 (d, J = 16.0 Hz, 1H), 4.23 (d, J = 16.0 Hz, 1H), 2.38 – 2.25 (m, 2H), 2.03 (br s, 2H), 1.75 – 1.60 (m, 1H), 1.52 - 1.37 (m, 1H), 1.22 – 1.08 (m, 2H), 1.07 – 0.92 (m, 2H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 172.9, 170.9, 156.8, 156.0, 149.9, 149.3, 136.9, 136.7, 131.4, 131.1, 130.9, 129.6, 128.8, 128.6, 128.2, 127.6, 127.7, 126.3, 125.5, 125.3, 125.2, 124.8, 123.3, 122.8, 122.3, 122.1, 121.7, 121.7, 52.9, 51.5, 49.6, 42.1, 41.5, 38.7, 32.5, 22.2. HRMS (ESI) m/z: calcd.: 570.2864, found: 570.2859 [M+H+]+.

107 4. Excimer based sensing

(R)-6-(pent-4-ynamido)-2-(2-(pyren-1-yl)acetamido)-N,N -bis(pyridin-2 -ylmethyl)hexanamide(46)

N N NH N O N H O HN O

Compound 46 was prepared using the general procedure for peptide synthesis with compound 44 (450 mg, 0.789 mmol, 1.00 eq.) and pent-4-ynoic acid (100 mg, 1.03 mmol, 1.30 eq.) to give the product as a white solid (505 mg, 0.777 mmol, 98%).

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.46 (dd, J = 33.4, 5.0 Hz, 2H), 8.27 – 7.92 (m, 9H), 7.61 (td, J = 7.6, 1.7 Hz, 1H), 7.40 (t, J = 7.6 Hz, 1H), 7.18 – 7.12 (m, 2H), 7.10 – 6.97 (m, 2H), 6.35 (d, J = 8.2 Hz, 1H), 5.06 (td, J = 8.2, 4.1 Hz, 1H), 4.85 – 4.51 (m, 4H), 4.37 (d, J = 16.0 Hz, 1H), 4.25 (d, J = 16.0 Hz, 1H), 2.97 - 2.76 (m, 2H), 2.39 (td, J = 7.2, 2.6 Hz, 2H), 2.18 – 2.09 (m, 2H), 1.96 (t, J = 2.6 Hz, 1H), 1.74 – 1.62 (m, 1H), 1.51 – 1.36 (m, 1H), 1.29 – 1.12 (m, 2H), 1.12 – 0.92 (m, 2H). Probe (47)

O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N

108 4.6. Experimental

Alkine 46 (100 mg, 0.153 mmol, 3.20 eq.) was suspended in THF (3 mL), TEEA buﬀer (3 mL, 100 mM, pH 7) was added and argon bubbled through the solution for 1 h. The azide 38 (15.7 mg, 0.048 mmol, 1.00 eq.), TBTA (13 mg, 0.024 mmol, 0.50 eq.) and

CuSO4· 5H2O (24.0 mg, 0,096 mmol, 2.00 eq.) was added and the mixtures stirred for 10 mins. Sodium ascorbate (76.2 mg, 0.384 mmol, 8.00 eq.) was added and the reaction heated to 60 ° C overnight. The mixture was cooled to RT and stirred with EDTA, extracted with DCM (3x5 mL) and the solvent removed under reduced pressure. Puriﬁ- cation was achieved by ﬂash column chromatography (DCM/MeOH 9:1 to 8:2) yielding the chemosensor 47 (85 mg, 0,036 mmol, 75%) as a bright yellow solid.

Rf (SiO2, DCM/MeOH 8:2 (vol/vol)) 0.6. 1 H-NMR (400 MHz, DMSO-d6, δ/ppm): 8.81 (d, J = 8.2 Hz, 3H), 8.46 (ddd, J = 4.9, 1.8, 0.9 Hz, 3H), 8.44 - 8.42 (m, 3H), 8.37 (d, J = 9.3 Hz, 3H), 8.27 – 8.22 (m, 6H), 8.20 (d, J = 7.8 Hz, 3H), 8.13 (d, J = 8.6 Hz, 9H), 8.04 (d, J = 7.6 Hz, 3H), 8.02 – 7.97 (m, 3H), 7.82 (t, J = 5.5 Hz, 3H), 7.68 – 7.60 (m, 6H), 7.44 (td, J = 7.7, 1.8 Hz, 3H), 7.28 – 7.22 (m, 3H), 7.19 (ddd, J = 7.6, 4.8, 1.1 Hz, 3H), 7.13 (ddd, J = 7.6, 4.8, 1.1 Hz, 3H), 7.09 (dt, J = 7.9, 1.1 Hz, 3H), 5.52 (s, 6H), 4.98 (d, J = 17.0 Hz, 3H), 4.90 – 4.73 (m, 6H), 4.52 (d, J = 17.0 Hz, 3H), 4.43 (d, J = 15.8 Hz, 3H), 4.33 – 4.16 (m, 6H), 2.99 - 2.86 (m, 6H), 2.85 - 2.78 (m, 6H), 2.78 - 2.68 (m, 6H), 2.42 - 2.33 (m, 6H), 1.76 - 1.55 (m, 6H), 1.33 – 1.09 (m, 6H), 0.66 (t, J = 6.5 Hz, 9H).

13 C-NMR (101 MHz, DMSO-d6) δ/ppm): 172.6, 170.09, 170.2, 157.2, 156.6, 149.3, 148.9, 146.2, 145.5, 136.9, 136.6, 131.0, 130.8, 130.4, 130.0, 129.7, 129.0, 128.7, 127.4, 127.2, 126.8, 126.2, 125.1, 124.9, 124.7, 124.1, 124.0, 123.9, 122.6, 122.1, 121.7 (2C), 121.0, 52.4, 51.3, 48.9, 47.3, 40.4, 38.3, 35.0, 31.4, 28.9, 22.9, 22.7, 21.4, 14.8. HRMS (ESI) m/z: calcd.: 2277.1185, found: 2277.1184 [M+H+]+.

109 4. Excimer based sensing

1-((1H -imidazol-1-yl)methyl)-3-(pyren-1-ylmethyl)-1H -imidazol-3-ium bromide (51)

N N N N

1-bromomethylpyrene (29.5 mg, 0.100 mmol, 1.00 eq.) and bisimidazole (51.8 mg, 0.350 mmol, 3.50 eq.) were suspended in anh. THF (9 mL) and reﬂuxed for 24 h under argon. After cooling to RT, the precipitate was ﬁltered and washed with ether. The desired product 51 was obtained as a white solid (31 mg, 0.069 mmol, 69%)

Analytical data were in agreement with literature.[4]

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 9.39 (s, 1H), 8.52 – 7.71 (m, 12H), 7.42 (s, 1H), 6.98 (s, 1H), 6.37 (s, 2H), 6.24 (s, 2H). HRMS (ESI) m/z: calcd.: 363.1604, found: 363.1602 [M+H+]+. (E)-3-(pyren-1-yl)acrylic acid (56)

A solution of 1-pyrenecarboxyaldehyde (54) (1.5 g, 6.4 mmol, 1.0 eq.), malonic acid (55) (1.4 g, 14 mmol, 2.2 eq.), pyridine (10 mL) and piperidine (11 drops) was reﬂuxed for 4 hs under nitrogen atmosphere. The reaction mixture was poured into a cold 3 M HCl solution. The formed precipitate was ﬁltered and washed with cold water. Recrystallization (DCM/MeOH 3:1) yielded the product as a yellow solid (906 mg, 3.30 mmol, 52%).

Analytical data were in agreement with literature.[5]

1 H-NMR (400 MHz, MeOH-d4, δ/ppm): 6.83 (d, J = 15.7 Hz, 1H), 8.11 (t, J = 7.7 Hz, 1H), 8.23 (q, J = 8.9 Hz, 2H), 8.38 – 8.29 (m, 4H), 8.52 (t, J = 8.0 Hz, 2H), 8.70 (d, J = 15.7 Hz, 1H), 12.78 – 12.45 (m, 1H).

110 4.6. Experimental

HRMS (ESI) m/z: calcd.: 271.075, found: 271.076 [M-H+]−.

3-(pyren-1-yl)propan-1-ol (57)

LiAlH4 (189 mg, 4.99 mmol, 3.40 eq.) was suspended in dry THF (60 mL) and 56 (400 mg, 1.46 mmol, 1.00 eq.) in dry THF (20 mL) was added dropwise at RT under argon atmosphere and the reaction mixture was, then, stirred for 24 hs. Water (5 mL), NaOH solution (15 %, 10 mL) and a sat. Rochelle salt solution (60 mL) were added and the mixture was stirred until all aluminium salts were dissolved. The organic layer was separated, and the aqueous layer was extracted with DCM (3x20 mL). The combined organic layers were dried over MgSO4 and the solvent was removed under reduced pressure. Puriﬁcation by silica column (DCM/MeOH 10:1) gave 57 (151 mg, 0.58 mmol, 39 %) as yellow solid.

Analytical data were in agreement with literature.[5]

Rf (SiO2, DCM/MeOH 10:1 (vol/vol)) 0.6. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.31 (d, J = 9.3 Hz, 1H), 8.19 – 8.07 (m, 5H), 8.04 – 7.96 (m, 2H), 7.89 (d, J = 7.8 Hz, 1H), 3.79 (t, J = 6.3 Hz, 2H), 3.48 – 3.39 (m, 2H), 2.19 – 2.09 (m, 2H). HRMS (APCI) m/z: calcd.: 261.1274, found: 261.128 [M+H+]+. dimethyl 5-(3-(pyren-1-yl)propoxy)isophthalate (59)

O O OMe

O OMe

57 (84.0 mg, 0.322 mmol, 1.00 eq.) was dissolved in THF (10 mL), dimethyl-5- hydroxyisophthalate (58) (108 mg, 0.516 mmol, 1.60 eq.) and PPh3 (110 mg, 0.419 mmol,

111 4. Excimer based sensing

1.30 eq.) were added and the solution cooled to -78° C and DIAD (70.0 µL, 0.355 mmol, 1.10 eq.) was added and the reaction was refluxed overnight. Water (1 mL) was added and the solvent removed under reduced pressure. The product 59 was purified by flash column chromatography (pentane/EE 9:1) (93 mg, 0.205 mmol, 63%). and obtained as a bright yellow solid.

Analytical data were in agreement with literature.[5]

Rf (SiO2, pentane/EE 9:1 (vol/vol)) 0.6. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.35 – 8.28 (m, 2H), 8.17 (d, J = 7.6 Hz, 2H), 8.14 – 8.07 (m, 2H), 8.04 – 7.96 (m, 3H), 7.89 (d, J = 7.7 Hz, 1H), 7.78 (d, J = 1.4 Hz, 2H), 4.14 (t, J = 6.0 Hz, 2H), 3.93 (s, 6H), 3.63 – 3.54 (m, 2H), 2.48 – 2.33 (m, 2H). 1-(3-(3,5-bis(bromomethyl)phenoxy)propyl)pyrene (60)

O Br

LiAlH4 (38.9 mg, 0.205 mmol, 1.00 eq.) was suspended in THF (8 mL) and 59 (93.0 mg, 0.205 mmol, 1.00 eq.) dissolved in THF (2 mL) was added dropwise and the mixture stirred overnight at RT. Water (1 mL), 6 N NaOH (1 mL) and Rochelles salt (10 mL) was added and stirred for 2 hs. The organic layer was separated and the aqueous layer extracted with DCM (3x5 mL). The combined organic layers were dried over Na2SO4 and the solvent removed under reduced pressure. The crude was dissolved in THF (5 mL), cooled to 0° C and PBr3 (58 µL, 0.616 mmol, 3.00 eq.) was added and the solution stirred overnight at RT. MeOH (1 mL) and sat. NaHCO3 were added. The aqueous phase was extracted with DCM (2x5 mL) and the solvent removed under reduced pressure. Puriﬁcation was obtained by ﬂash column chromatography (pentane/DCM 2:1) and the product 60 was obtained as a brown solid (53 mg, 0.101 mmol, 49%).

Analytical data were in agreement with literature.[5]

Rf (SiO2, pentane/DCM 2:1 (vol/vol)) 0.6.

112 4.6. Experimental

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.31 (d, J = 9.3 Hz, 1H), 8.19 – 8.07 (m, 4H), 8.06 – 7.95 (m, 3H), 7.90 (d, J = 7.8 Hz, 1H), 7.00 (d, J = 1.6 Hz, 1H), 6.88 (d, J = 1.5 Hz, 2H), 4.42 (s, 4H), 4.07 (t, J = 6.0 Hz, 2H), 3.64 – 3.52 (m, 2H), 2.35 (dt, J = 13.4, 6.2 Hz, 2H). 1,1’-(5-(3-(pyren-1-yl)propoxy)-1,3-phenylene)bis(N,N -bis(pyridin-2-ylmethyl) methanamine)(61)

N O N

N N

60 (53 mg, 0.101 mmol, 1.00 eq.) was dissolved in DMF (4 mL) and K2CO3 (51.8 mg, 0.375 mmol, 3.70 eq.), KI (45.6 mg, 0.274 mmol, 2.70 eq.) and DPA (62.6 mg, 0.314 mmol, 3.10 eq.) was added and the mixtures stirred at RT for 1 day. The solvent was removed and the solid partitioned between EE (5 mL) and H2O (5 mL). The aqueous layer was extracted with DCM (3x2 mL) the combined organic layers dried over Na2SO4 and the solvent removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 95:5 to 9:1) and the chemosensor 61 could be obtained as a yellow solid (74 mg, 0.097 mmol, 96%).

Analytical data were in agreement with literature.[5]

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.5. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.49 (d, J = 4.9 Hz, 4H), 8.30 (d, J = 9.3 Hz, 1H), 8.17 – 7.93 (m, 7H), 7.88 (d, J = 7.8 Hz, 1H), 7.59 – 7.50 (m, 8H), 7.08 (td, J = 5.1, 3.3 Hz, 5H), 6.91 (d, J = 1.4 Hz, 2H), 4.05 (t, J = 5.6 Hz, 2H), 3.81 (s, 8H), 3.66 (s, 4H), 3.56 (t, J = 7.5 Hz, 2H), 2.35 (p, J = 6.4 Hz, 2H).

113 4. Excimer based sensing

PyDPA (33)

N N 2+ Zn N O

N 2+ N Zn N

Zn(NO3)2·6H2O (15.6 mg, 0.052 mmol, 2.00 eq.) was dissolved in MeCN (2 mL) and 61 (20.0 mg, 0.026 mmol, 1.00 eq.) was added and the mixture stirred for 2.5 hs at RT. The solvent was removed and PyDPA was obtained as a yellow solid (28 mg, 0.024 ˙mmol, 95%).

Analytical data were in agreement with literature.[5] UV λ = 344/277/228 nm Fluorescence λ = 380/470 nm

N -(4-iodobenzyl)-1-(pyridin-2-yl)-N -(pyridin-2-ylmethyl)methanamine (84)

N N N

DPA (550 mg, 2.76 mmol, 1.00 eq.) and 4-Iodobenzylaldehyde (760 mg, 3.31 mmol, 1.20 eq.) were dissolved in 1,2-DCE (50 mL) and stirred for 15 mins.

NaBH(OAc)3 (820 mg, 3.86 mmol, 1.40 eq.) was added and the mixture stirred over night at RT. Saturated NaHCO3 was added and extracted with DCM (3x40 mL). The combined organic layers were dried over Na2SO4 and the solvent removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 98:2) and the product 84 was obtained as a brown oil (1.13 g, 2.72 mmol, 98%).

114 4.6. Experimental

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.52 (ddd, J = 4.9, 1.8, 0.9 Hz, 2H), 7.72 – 7.60 (m, 4H), 7.55 – 7.51 (m, 2H), 7.20 – 7.10 (m, 4H), 3.80 (s, 4H), 3.64 (s, 2H). tert-butyl (2-(bis(pyridin-2-ylmethyl)amino)-2-oxoethyl)carbamate (64)

O H O N N O N N

The acid Boc-Gly-OH (315 mg, 1.79 mmol, 1.00 eq.) was treated with DPA (271 muL, 1.50 mmol, 0.84 eq.) according to the general procedure for peptide synthesis to give the product puriﬁed by ﬂash column chromatography (pentane/EE 98:2 to 98:2) as a yellow solid (530 mg, 1.49 mmol, 99%).

Rf (SiO2, DCM/MeOH 98:2 (vol/vol)) 0.5. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.55 (d, J = 4.6 Hz, 1H), 8.50 (d, J = 4.5 Hz, 1H), 7.63 (tdd, J = 7.5, 5.2, 1.8 Hz, 2H), 7.29 (s, 1H), 7.22 - 7.10 (m, 3H), 4.76 (s, 2H), 4.64 (s, 2H), 4.18 (d, J = 4.4 Hz, 2H), 1.43 (s, 9H). HRMS (ESI) m/z: calcd.: 357.1921, found: 357.1921 [M+H+]+.

2-amino-N,N -bis(pyridin-2-ylmethyl)acetamide (65)

O H N 2 N N N

Boc deprotection of 64 (530 mg, 1.49 mmol, 1.00 eq.) was carried out according to the general procedure for Boc deprotection. The crude was dissolved in water and 6 N NaOH was added until the pH of the mixture became >9. The product was extracted with DCM (3x10 mL), the solvent was removed and the product 65 was obtained as a white solid (293 mg, 1.14 mmol, 72%).

115 4. Excimer based sensing

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.56 (d, J = 4.6 Hz, 1H), 8.50 (d, J = 4.7 Hz, 1H), 7.63 (tdd, J = 7.5, 5.2, 1.8 Hz, 2H), 7.30 (d, J = 7.8 Hz, 1H), 7.23 – 7.09 (m, 3H), 4.78 (s, 2H), 4.63 (s, 2H), 3.64 (s, 2H). 2-((6-bromopyren-1-yl)amino)-N,N -bis(pyridin-2-ylmethyl)acetamide (67)

N N N H O N

1,6-dibromopyrene (100 mg, 0.27 mmol, 1.00 eq.), Pd(OAc)2 (4.36 mg, 0.02 mmol, 0.07 eq.), Na t-butoxide (58.7 mg, 0.61 mmol, 2.20 eq.) and dppf (36.9 mg, 0.07 mmol, 0.24 eq.) were dissolved in degassed toluene (8 mL) and stirred for 15 mins at RT. The mixture was heated to 100 ° C and the amine 65 (83 mg, 0.33 mmol, 1.2 eq.) was added and refluxed overnight. Water (1 mL) was added and the mixture was filtered over celite, washed with brine (2x5 mL) and the solvent was removed under reduced pressure. The product was purified by flash column chromatography (pentane/EE 95:5) and the product 67 was obtained as a yellow solid (92 mg, 0.17 mmol, 63%).

Rf (SiO2, DCM/MeOH 95:5 (vol/vol)) 0.5. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.63 (d, J = 4.6 Hz, 1H), 8.55 (d, J = 4.9 Hz, 1H), 8.26 – 7.81 (m, 8H), 7.67 (qd, J = 7.5, 1.8 Hz, 2H), 7.37 (d, J = 7.8 Hz, 1H), 7.28 (s, 1H), 7.24 – 7.14 (m, 2H), 6.32 (s, 1H), 4.89 (s, 2H), 4.84 (s, 2H), 4.45 (d, J = 3.8 Hz, 2H). HRMS (ESI) m/z: calcd.: 535.1128, found: 535.1130 [M+H+]+.

116 4.6. Experimental

N1,N5 -bis(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl) acetamido)hexyl)glutaramide (70)

O O HN NH

N N N HN N NH O O O N N O

Glutaric acid (14 mg, 0.106 mmol, 1.00 eq.) was treated with 44 (124 mg, 0.21 mmol, 2.05 eq.) according to the general procedure for peptide synthesis to give the product puriﬁed by ﬂash column chromatography (DCM/MeOH 95:5 to 80:20) as a yellow solid (130 mg, 1.05 mmol, 99%).

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.3. 1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.47 (ddd, J = 4.8, 1.8, 0.9 Hz, 2H), 8.37 (ddd, J = 5.0, 1.8, 1.0 Hz, 2H), 8.27 – 7.88 (m, 18H), 7.56 (td, J = 7.7, 1.8 Hz, 2H), 7.33 (td, J = 7.7, 1.8 Hz, 2H), 7.16 (d, J = 7.8 Hz, 2H), 7.11 (ddd, J = 7.6, 4.8, 1.1 Hz, 2H), 7.06 – 6.96 (m, 4H), 6.94 (s, 1H), 6.92 (s, 1H), 6.16 (s, 2H), 4.99 (td, J = 8.2, 4.2 Hz, 2H), 4.85 (d, J = 17.1 Hz, 2H), 4.73 – 4.56 (m, 6H), 4.37 – 4.18 (m, 4H), 3.03 – 2.94 (m, 2H), 2.84 – 2.73 (m, 2H), 2.07 - 1.82 (m, 4H), 1.77 – 1.66 (m, 4H), 1.60 – 1.37 (m, 4H), 1.19 – 0.99 (m, 6H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 173.1, 172.8, 171.1, 156.5, 155.8, 149.7, 148.7, 137.4, 137.1, 131.4, 131.0, 130.9, 129.6, 129.0, 128.6, 128.1, 127.6, 127.5, 126.2, 125.5, 125.3, 125.3, 125.2, 124.8, 123.4, 122.9, 122.5, 122.2, 121.9, 52.8, 51.4, 49.7, 41.7, 38.5, 34.8, 32.0, 29.9, 28.8, 22.1. (each signal contains 2 carbons atoms) HRMS (ESI) m/z: calcd.: 1235.5866, found: 1235.5852 [M+H+]+.

117 4. Excimer based sensing

N1,N10 -bis(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl) acetamido)hexyl)decanediamide (73)

O O HN 6 NH

N N N HN N NH O O O N N O

Sebatic acid (21 mg, 0.11 mmol, 1.00 eq.) was treated with 44 (120 mg, 0.21 mmol, 2.05 eq.) according to the general procedure for peptide synthesis to give the product puriﬁed by ﬂash column chromatography (DCM/MeOH 96:4 to 84:16) as a yellow solid (114 mg, 0.09 mmol, 82%).

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.4. 1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.49 (ddd, J = 4.8, 1.8, 1.0 Hz, 1H), 8.37 (ddd, J = 4.8, 1.8, 1.0 Hz, 1H), 8.25 – 7.87 (m, 18H), 7.57 (td, J = 7.7, 1.8 Hz, 2H), 7.34 (td, J = 7.7, 1.8 Hz, 2H), 7.15 – 7.11 (m, 4H), 7.03 – 6.98 (m, 4H), 6.61 (s, 1H), 6.59 (s, 1H), 5.51 (s, 2H), 5.05 (td, J = 8.3, 4.2 Hz, 2H), 4.83 (d, J = 17.0 Hz, 2H), 4.70 – 4.56 (m, 6H), 4.32 (d, J = 16.0 Hz, 2H), 4.22 (d, J = 15.9 Hz, 2H), 2.98 – 2.76 (m, 4H), 1.96 – 1.85 (m, 4H), 1.73 – 1.63 (m, 2H), 1.54 – 1.37 (m, 8H), 1.29 – 1.15 (m, 8H), 1.11 – 0.95 (m, 6H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 173.4, 172.9, 171.1, 156.6, 155.8, 149.9, 149.2, 137.0, 136.9, 131.4, 131.1, 130.9, 129.6, 128.8, 128.6, 128.2, 127.6, 127.5, 126.3, 125.6, 125.4, 125.3, 125.2, 124.8, 123.3, 122.9, 122.4, 122.1, 121.7, 52.8, 51.4, 49.3, 42.0, 39.0, 36.6, 32.5, 29.2, 29.1, 28.6, 25.7, 22.2. (each signal contains 2 carbon atoms) HRMS (ESI) m/z: calcd.: 1305.6648, found: 1305.6631 [M+H+]+.

118 4.6. Experimental

70· 2Zn (71)

O O

HN NH

N N 2+ N 2+ N HN Zn Zn NH O O O N N O

4 NO3-

70 (20 mg, 0.02 mmol, 1.00 eq.) was dissolved in DCM/MeOH 2:1 (15 mL) and

Zn(NO3)2·6H2O (9.63 mg, 0.032 mmol, 2.00 eq.) and the mixture stirred for 2hs at RT. The solvent was removed under reduced pressure and the product 71 was obtained as a white solid (22 mg, 0.016 mmol, quant.).

UV λ = 344 nm Fluorescence λ = 470 nm

73· 2Zn (74)

O O

HN 6 NH

N N 2+ N 2+ N HN Zn Zn NH O O O N N O

- 4 NO3

73 (20 mg, 0.015 mmol, 1.00 eq.) was dissolved in DCM/MeOH 2:1 (15 mL) and

Zn(NO3)2·6H2O (9.12 mg, 0.031 mmol, 2.00 eq.) and the mixture stirred for 2hs at RT. The solvent was removed under reduced pressure and the product 71 was obtained as a white solid (22 mg, 0.015 mmol, quant.).

UV λ = 344 nm Fluorescence λ = 470 nm

119 4. Excimer based sensing tert-butyl(R)-4-((((9H -ﬂuoren-9-yl)methoxy)carbonyl)amino)-5-(bis (pyridin-2-ylmethyl)amino)-5-oxopentanoate (76)

N O H O N N N O

O O

Compound 76 was prepared from Fmoc-Glu(OtBu)-OH (1.03 g, 2.40 mmol, 1.00 eq.) using the procedure as for 40. Puriﬁcation was achieved by ﬂash column chromatography (EE/EtOH 100:0 to 97:3) gave the product 76 (1.19 g, 1.96 mmol, 98%) as a white solid.

Rf (SiO2, EE/EtOH 9:1 (vol/vol)) 0.4. 1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.51 (m, 1H), 8.49 (m, 1H), 7.76 (d, J = 7.5 Hz, 2H), 7.65 – 7.56 (m, 4H), 7.40 (t, J = 7.6 Hz, 2H), 7.34 – 7.28 (m, 2H), 7.23 (d, J = 7.8 Hz, 2H), 7.18 – 7.12 (m, 2H), 5.77 (d, J = 8.5 Hz, 1H), 4.95 – 4.75 (m, 4H), 4.61 (d, J = 15.3 Hz, 1H), 4.34 (dd, J = 7.4, 2.1 Hz, 2H), 4.20 (t, J = 7.2 Hz, 1H), 2.43 – 2.29 (m, 2H), 2.25 – 2.09 (m, 1H), 1.96 – 1.80 (m, 1H), 1.41 (s, 9H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 172.9, 172.4, 156.9, 156.2, 156.0, 150.0, 149.4, 144.1, 144.0, 141.4, 141.4, 136.9, 136.8, 127.8 (2C), 127.2 (2C), 125.3, 125.3, 122.8, 122.5, 122.3, 121.8, 120.1 (2C), 80.7, 67.1, 52.8, 51.4, 50.8, 47.3, 31.2, 28.7, 28.2. (Signals corresponding to 2C were identiﬁed using a known reference compound.) HRMS (ESI) m/z: calcd.: 607.2915, found: 607.2913 [M+H+]+.

120 4.6. Experimental tert-butyl (R)-4-amino-5-(bis(pyridin-2-ylmethyl)amino)-5-oxopentanoate (77)

N O N H2N N

O O

Compound 77 was prepared from 76 (1.15 g, 1.94 mmol, 1.00 eq.) using the procedure as for 42. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 96:4 to 65:35) gave 77 (598 mg, 1.55 mmol, 80%) as a white solid.

Rf (SiO2, DCM/EtOH 9:1 (vol/vol)) 0.2. 1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.58 – 8.51 (m, 1H), 8.48 (ddd, J = 4.9, 1.8, 0.9 Hz, 1H), 7.61 (tdd, J = 7.7, 6.6, 1.8 Hz, 2H), 7.30 – 7.23 (m, 1H), 7.21 – 7.10 (m, 3H), 5.05 (d, J = 15.2 Hz, 1H), 4.97 (d, J = 17.1 Hz, 1H), 4.58 (d, J = 17.1 Hz, 1H), 4.42 (d, J = 15.2 Hz, 1H), 3.93 (dd, J = 8.8, 4.6 Hz, 1H), 2.52 - 2.43 (m, 1H), 2.42 – 2.33 (m, 3H), 2.08 - 1.98 (m, 1H), 1.81 - 1.69 (m, 1H), 1.38 (s, 9H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 176.6, 173.0, 157.4, 156.4, 150.0, 149.3, 136.9 (2C), 122.7, 122.5, 122.4, 121.8, 80.5, 52.4, 51.6, 50.7, 31.6, 30.2, 28.2. (Signals corresponding to 2C were identiﬁed using a known reference compound.) HRMS (ESI) m/z: calcd.: 385.2234, found: 385.2231 [M+H+]+. tert-butyl (R)-5-(bis(pyridin-2-ylmethyl)amino)-5-oxo-4-(pyrene-1-carboxamido) pentanoate(79)

N O H N N N O

O O

121 4. Excimer based sensing

Compound 79 was prepared from 77 (580 mg, 1.51 mmol, 1.00 eq.) using the general procedure for peptide synthesis. The product was puriﬁed by ﬂash column chromatography (DCM/MeOH 9:1) and 79 (925 mg, 1.51 mmol, quant.) was obtained as a white solid.

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.3. 1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.58 (ddd, J = 4.8, 1.8, 0.9 Hz, 1H), 8.53 (ddd, J = 4.9, 1.8, 0.9 Hz, 1H), 8.30 – 7.99 (m, 9H), 7.68 (td, J = 7.7, 1.8 Hz, 1H), 7.63 (td, J = 7.6, 1.8 Hz, 1H), 7.38 (dt, J = 7.8, 1.0 Hz, 1H), 7.31 (dt, J = 7.9, 1.1 Hz, 1H), 7.23 – 7.14 (m, 3H), 5.58 - 5.51 (m, 1H), 5.03 (d, J = 3.1 Hz, 2H), 4.95 (d, J = 15.4 Hz, 1H), 4.73 (d, J = 15.4 Hz, 1H), 2.63 – 2.50 (m, 2H), 2.47 – 2.35 (m, 1H), 2.15 – 2.02 (m, 1H), 1.44 (s, 9H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 173.0, 172.5, 169.7, 162.6, 157.0, 156.1, 150.0, 149.5, 137.0, 136.9, 132.9, 131.3, 130.8, 130.5, 129.0, 128.9, 128.8, 127.3, 126.4, 125.9, 125.9, 124.9, 124.7, 124.5, 124.5, 122.9, 122.5, 122.3, 122.1, 80.8, 53.0, 51.6, 49.8, 38.7, 28.6, 28.2.

122 4.6. Experimental

4.6.3. References

Biblography Chapter Excimer based sensing

[1] X.-l. Ni, X. Zeng, C. Redshaw, T. Yamato, J. Org. Chem. 2011, 76 (14), 5696–5702.

[2] H.-W. Rhee, C.-R. Lee, S.-H. Cho, M.-R. Song, M. Cashel, H. E. Choy, Y.-J. Seok, J.-I. Hong, J. Am. Chem. Soc. 2008, 130 (3), 784–785.

[3] J. Y. Jung, E. J. Jun, Y.-U. Kwon, J. Yoon, Chem. Commun. 2012, 48 (64), 7928– 7930.

[4] Z. Xu, N. J. Singh, J. Lim, J. Pan, H. N. Kim, S. Park, K. S. Kim, J. Yoon, J. Am. Chem. Soc. 2009, 131 (42), 15528–15533.

[5] J. Oh, J.-I. Hong, Org. Lett. 2013, 15 (6), 1210–1213.

[6] B. Schazmann, N. Alhashimy, D. Diamond, J. Am. Chem. Soc. 2006, 128 (26), 8607–8614.

[7] „isoemissive point“ in IUPAC Compendium of Chemical Terminology, IUPAC, Re- search Triagle Park, NC, 2009.

[8] M. S. C. Wilson, A. Saiardi, Top. Curr. Chem. 2017, 375 (1), 1–21.

[9] G. W. Mayr, Biochem. J. 1988, 254 (2), 585–591.

[10] H. Wang, V. S. Nair, A. A. Holland, S. Capolicchio, H. J. Jessen, M. K. Johnson, S. B. Shears, Biochemistry 2015, 54 (42), 6462–6474.

[11] J. R. Lakowicz, Principles of ﬂuorescence spectroscopy, third edition, corrected at 4. printing Auﬂ., Springer, New York, NY, 2010.

[12] O. Losito, Z. Szijgyarto, A. C. Resnick, A. Saiardi, PloS one 2009, 4 (5), e5580.

123

CHAPTER 5

Sensing via a disassembly approach

125 5. Sensing via a disassembly approach

5.1. Background

Salen (N,N ’-disalicylideneethylenediamine, 85) (Scheme 5.1) and its analogues are ligands which possess a convenient synthesis and versatile coordination abilities.[1] These tetradentate ligands are employed in the range from materials to biomedical science to catalysis.[2] Complexed with a metal (e.g. M = Zn2+, Fe3+, Mn2+) the complex is stable in water. However, the imine bonds are prone to hydrolysis when the metal is missing.[1] This can be used as a sensing strategy.[3]

N N M O O 85

Scheme 5.1: Chemical structure of a salen ligand with a fourfold coordinated complexed metal.

5.1.1. Disassembly Approach

In 2011 Jung et al. reported an ensemble-based fluoregenic chemodosimeter for thiol- detection.[4,5] Chemodosimeters are a class of sensors based on analyte binding induced irreversible chemical reaction.[4] In their studies, they found that demetalation and hy- drolytic cleavage of a Schiff base generates a fluorescent species (Figure 5.1). Inspired by that work, the Zelder group developed a molecular probe detecting PPi (86) following this strategy and called it the ”disassembly approach”.[3] The concept of this approach is shown in Figure 5.2. PPi (87) sequesters the central metal ion from the salen-Fe3+ complex followed by imine hydrolysis in water. This releases two eq. of fluorogenic salicylaldehyde 88 resulting in a turn-on fluorescence.

Figure 5.1.: Molecular design of Zn2+-salen chemosensor. Demetalization of Zn2+ leads to disassembly. Picture adapted from reference. [6]

126 5.1. Background

In Figure 5.2 the time course of the reaction between complex 89 and added PPi (87) in tris(hydroxymethyl)aminomethane (Tris) buﬀer is displayed (Figure 5.2 a)) as well as the synthesis and disassembly of the complex 89 (Figure 5.2 b)). To investigate whether their chemosensor is selective for PPi (87) or disassembly is also induced by other anions Kumari et al. tested the reactions between complex 89 and various anions

- 2- - - 2- (e.g., OAc , SO4 , CN ,H2PO4 /HPO4 , ATP, ADP, AMP) However, no signiﬁcant changes could be observed upon the addition of other phosphate oxoanions, thus they

[6] concluded 89 being a selective chemosensor for PPi (87).

H H O O

OH N N a), b) Fe 2x O O 88 89 Fluorescent Non-Fluorescent O O P P O O O O O 87 (a) (b) Figure 5.2.: a) Changes in emission spectra of 89 (50 µM, in Tris buﬀer pH 7.4) with time (0-35 min) after added PPi (87) (10 eq.) b) Synthesis and disassembly of 89 with PPi. λEx = 350 nm, λEm = 300- 600 nm. Pictures adapted from reference. [6]

InsP6 and lower inositol phosphates are known to form complexes with metal cations such as Fe3+, Cu2+ and Zn2+.[7] Due to this fact, the effect of inositol polyphosphates on Fe3+-salen (89) should be investigated. We assumed that the complexed Fe3+ would sequestered by InsP6 (90) and 5-PP-InsP5 (5) resulting in hydrolysis of 89 and fluorescence emission from released salicylaldehyde (88). Recently, Winkler et al. published an study which compared the stability of Fe3+, Zn2+, Mn2+ and Mn3+ towards demetallation by different oxoanions.[8]

127 5. Sensing via a disassembly approach

5.2. Results and Discussion

5.2.1. Synthesis of disassembly based ﬂuorescent probes

The following chapter describes the synthesis of several metal complexes including the synthesis of their ligands.

5.2.1.1. Salicylaldehyde-based ﬂuorescent probes

Ligand 89 was synthesized following a literature known procedure[9] by condensation of salicylaldehyde (88) with ethylenediamine (91) in ethanol (Scheme 5.2) in 97% yield. Subsequent complexation is carried out by reﬂuxing the ligand with 1.0 eq. of

3+ the corresponding metal salt in ethanol. Complexation with FeCl3 aﬀorded the Fe - complex 89 in 52% yield.

NH H N 2 N N O 2 3+ N Fe N 91 OH FeCl3 O H EtOH HO EtOH O OH 92 93 89 (97%) (52%)

Scheme 5.2: Synthesis of complex 89 via formation of the bis-Schiﬀ base 93 and subsequent iron complexation.

Having complex 89 in hand, the next step was to investigate the effect of inositol phosphates on the fluorescence response of it. Figure 5.3 shows the fluorescence emission spectra of 89 in Tris buffer (pH 7.4) over time and added anions. Indeed, inositol py- rophosphates are also able to induce the disassembly process resulting in an enhancement of fluorescence over time.

128 5.2. Results and Discussion

2- O O OPO3 2- 2- P P O3PO OPO3 O O O O O 2- 2- O3PO OPO3 O O P O O O P O O

(a) (b) Figure 5.3.: Emission spectra of complex 89 (50 µM in Tris, pH 7.4) over time (0-90 min) after added a) PPi (50 µM, 1.0 eq.) b) 5-PP-InsP5 (50 µM, 1.0 eq.). λEx = 350 nm, λEm = 400-850 nm

Figure 5.4 depicts the increase in emission intensity at 500 nm over time (0-55 mins) induced by PPi, InsP6, 5-PP-InsP5 and 1,5-InsP8.

Figure 5.4.: Corresponding increase in emission intensity at 500 nm over time (0-55 min). Added anions: PPi (50 µM, 1.0 eq., grey), InsP6 (50 µM, 1.0eq., red), 5-PP-InsP5 (50 µM, 1.0 eq., blue), 1,5- InsP8 (50 µM, 1.0 eq., green).

These ﬁndings, and the fact that salicylaldehyde (92) possess low ﬂuorescence intensity, motivated us to synthesise a library of compounds for potential broadening of the scope of this approach.

129 5. Sensing via a disassembly approach

5.2.1.2. Naphthol-based ﬂuorescent probes

Ding et al. recently reported a naphthol based ﬂuorogenic probe for hypochlorite D1 (Figure 5.5).[10] D1 should show ﬂuorescence when the C=N bond is oxidized by ClO- and deoximation to the corresponding aldehyde is induced.

Figure 5.5.: Fluorescence response of probe D1 through deoximation induced by ClO-. Picture taken from reference. [10]

To examine if the formed aldehyde could be used for inositol phosphate detection compound 94 was synthesized (Scheme 5.3) according the published procedure.[10] Start- ing from 2,7-dihydroxynaphthalene (95) compound 96 was obtained, in 42% yield, by a Bucherer reaction using dimethylamine (97) and Na2S2O5 in water. Subsequent

Vilsmeier-Haack reaction produced the aryl aldehyde 94 using POCl3 in DMF with 93% yield. On the basis of aldehyde 94 ligand 98 was formed through condensation with ethylenediamine (91). Complexation with FeCl3 gave complex 99 in quantitative yield.

Me2NH 97 HO OH N OH Na2S2O5 POCl3 N OH

H2O DMF O 95 96 94 (42%) (93%)

H2N NH 2 EtOH 91

N 3+ N OH N N O Fe N O FeCl3

EtOH N N OH

99 N 98 (quant.) (60%)

Scheme 5.3: Synthesis overview of complex 99. [10]

130 5.2. Results and Discussion

5.2.2. Excited state intramolecular proton transfer - ESIPT

2-(2’-hydroxyphenyl)benzoxazole (HBO) (100) derivatives are a class of fluorophores possessing intrinsic ESIPT properties. The detailed mechanism of ESIPT is explained in section ESIPT-based fluorescent probes. Photoexcitation of the molecule leads to a structural reorganization and a large Stokes shift. Molecule 100 is a benzoxazole scaffold (indicated in red in Scheme 5.4) carrying a substituent at the 2-position.

HO N

O 100

Scheme 5.4: Chemical structure of 2-(2’-hydroxyphenyl)benzoxazole (HBO) 100.

Fluorescent chemosensors with a large Stokes shift allow emission measurement against low background ﬂuorescence, thus enhancing sensitivity. To investigate whether we can implement this feature in our chemosensors, two building blocks 101a and 101b (Scheme 5.5) were synthesized. Several synthetic routes to obtain benzoxazole are reported in literature.[11] Traditional approaches are reacting 2-aminophenols with carboxylic acids derivatives under acidic conditions or reacting 2-aminophenols with alde- hydes, followed by oxidative cyclization of the imine intermediates.[11] Prakash and co-workers reported the oxidative intramolecular cyclization of phenolic Schiﬀ’s bases via iodobenzene diacetate (IBD).[12]

5.2.2.1. HBO-based ﬂuorescent probes

Scheme 5.5 shows the synthesis of 101a and 101b in two steps. In the ﬁrst step, initial formation of the imine from 2-hydroxy-5-methylbenzaldehyde (102a) and (102b) and 2-aminophenol (103) in methanol is followed by oxidative cyclization with hypervalent iodine 104, forming product 105a and 105b.

131 5. Sensing via a disassembly approach

H2N N HO R R R N N O 103 O N O PhI(OAc)2 104 106 HO N N MeOH HO TFA HO O

102a R= -CH3 105a (55%) 101a (99%) 102b R = -COOCH3 105b (82%) 101b (74%)

Scheme 5.5: Preparation of building blocks (101a and 101b) using PhI(OAc)2 (104) and urotropin (106)/TFA.

Subsequent ortho-formylation of the phenol moiety is achieved via a Duff reaction with urotropin (106) in TFA yielding building blocks 101a and 101b.[13] Having these two molecules in hand, we could proceed to synthesize chemosensors 107a and 107b applying the same procedure as for the salen synthesis (Scheme 5.6). Schiﬀ base formation with ethylenediamine (91) and corresponding building block gives ligands 108a and 108b with 68% and 57% yield respectively. Complexation with FeCl3 in ethanol gave the products 107a and 107b.

R O H2N NH R 2 N O 91 HO N FeCl3 EtOH EtOH N HO N HO O N

101a R= -CH3 O R 101b R= -COOCH3 R O 108a (68%) N (57%) 108b O 3+ N Fe O N N

O R 107a (quant) 107b (93%)

Scheme 5.6: Synthesis overview of complex 107a and 107b.

Fu et al. reported, that complexing HL sodium with Cu2+ forms an ensemble with quenched ﬂuorescence through metal induced ﬂuorescence quenching (Figure 5.6).[14]

132 5.2. Results and Discussion

Figure 5.6.: Proposed sensing mechanism for S2-. Picture taken from reference. [14]

The sensing mechanism, of the complex shown in Figure 5.6, relies on the displacement of Cu2+ from the coordination sphere releasing the ligand into solution inducing ﬂuorescence enhancement. Cu2+ was the only metal showing a quenching eﬀect to the ensemble. Again, we assumed that this concept would be adaptable for our sensing in- tends. Inspired by that work we complexed building block 101a with CuCl2 in THF/H2O to obtain 33 mg of 109 (Scheme 5.7).

O CuCl N 2 O 2+ O N THF/H2O Cu HO O O O 101a N O 109 (quant.)

Scheme 5.7: Synthesis of non-ﬂuorescent complex 109.

To address solubility issues in aqueous buﬀer, it was attempted to introduce a PEG linker into 110a (Scheme 5.8). Starting with the cleavage of ester 111a with NaOH in

H2O/MeOH the carboxylic acid derivative 112 is obtained with 99% yield. Formylation of 112 is executed with urotropin (106) in TFA with 37% yield.

133 5. Sensing via a disassembly approach

N N O O N O O OH OH NaOH O O 106 O

H2O/MeOH TFA N N N HO HO HO O 105b 112 113 (99%) (37%)

O HO O 5 O 81 O 5 O O O

N HO O 114

Scheme 5.8: Attempted synthesis of compound 114

The yield may be due to the reduced electron density, through the carboxylic acid at the aromatic ring, making introduction of the formyl group more diﬃcult. Coupling of 113 with hexamethylene glycol monomethyl ether (81) was attempted with the coupling reagent and base EDC/DMAP and TBTU/DIPEA in DCM, respectively, but no conversion was observed.

[15] Cheng and co-workers reported a ESIPT based chemosensor for PPi. The concept is shown in Figure 5.7a and relies on the fact that the phenoxide in 3·2Zn is participating

2+ in Zn binding (ESIPT-OFF). Upon binding of PPi to the DPA moietes this bond is broken and an intramolecular hydrogen bond to the HBO core is formed (ESIPT- ON). This leads to an emission shift of approximately 100 nm. Figure 5.7b show superimposed emission spectra of 3·2Zn with various anions, such as PPi and ATP.

We assumed that this sensing mechanism could be adopted for analytes such as 5-PP-

InsP5 due to the pyrophosphate moiety in the molecule. Hence we decided to synthesise 3·2Zn (Scheme 5.9).

134 5.2. Results and Discussion

(a) (b) Figure 5.7.: a) Proposed sensing mechanism of PPi with ESIPT based chemosensor 3·2Zn. b) PPi induced emission shift. Picture taken from reference. [15]

Starting from commercial available 2-(2’-hydroxyphenyl)benzoxazole (115) we ﬁrst tried to execute a di-formylation of the phenol following the Duff protocol. With acetic acid in toluene the starting material was reisolated. Reﬂuxing 115 with 106 in TFA gave the product 116 in 47% yield.

N HN N N N NH2 N O N 106 O 117 O

TFA N N NaBH(OAc)3, HO O DCE HO O 116 HN N (47%) N 118 HO (88%) N N 115 N H N N N N 122 Br DIPEA, 106 N MeCN NaBH(OAc)3/NaBH4 119 Acetic Acid, Toluene

N 2+ N Zn N N N N O Zn(NO3)2*H2O O 3NO - N 3 MeOH/H2O N O N HO 2+Zn N N N N N 121 120 (98%) (62%)

Scheme 5.9: Synthesis of ESIPT based sensor 121.

135 5. Sensing via a disassembly approach

Installing the bis(2-pyridylmethyl)amino groups using 122 via reductive amination was not successful. NaBH4 in MeOH and DCE gave no product and only educt was reisolated.

Although little product was formed with NaBH(OAc)3 in DCE, impurities hampered the isolation. A diﬀerent route to obtain 120 in two steps was alternatively applied.

With 117, reductive amination was successful using NaBH(OAc)3 with 88% yield. The

ﬁnal product was obtained by a SN2 reaction with 2-(bromomethyl)pyridine (119) with

DIPEA in acetonitrile and subsequent complexation with Zn(NO3)2·H2O gave 143 mg of the ﬁnal chemosensor 121. A variation of this sensor was synthesized according to Scheme 5.10. Condensation of 2-pyridinecarboxaldehyde (123) with N,N -dimethyl- N 2-(pyridin-2-ylmethyl)ethane-1,2-diamine 124 gives 125. Attaching 125 two times to the scaﬀold 116 using NaBH4 in MeOH gave the ligand 126 with 67% yield. The formed ligand could be complexed with Zn2+ and chemosensor 127 was obtained.1

O O

N N H2N HO O 124 116 NaBH4 H NaBH N 4 O N N N MeOH MeOH 123 125 (98%)

N N Zn2+ N N N O O Zn(NO3)2 N - N N 3NO3 MeOH N HO O N N 2+Zn N 127 N 126 N (quant.) (67%)

Scheme 5.10: Synthesis of ESIPT based sensor 127.

1Syntheses of chemosensors were supported by Stephan Mundinger.

136 5.3. Sensor evaluation

5.3. Sensor evaluation

In the previous section, the syntheses of chemosensors following the disassembly approach were discussed. Diﬀerent analogues have been made accessible in the categories “Salen“, “HBO“ and “Naphthol“ (Scheme 5.11).

Salen

N 3+ Fe N O O

HBO-based

O O O O O N N N O O O N 2+ 3+ O 3+ N Cu Fe Fe O O O N O N N N N O O O 107a 107b O 109 O

N N 2+ Zn Zn2+ N N N N O O

N N O N O N 2+ Zn N 2+Zn N N N 121 127

Naphthol-based

N 3+N N O Fe O

99 N

. Scheme 5.11: Overview of synthesized chemosensors in this chapter. Salen-based: 89; HBO-based: 107a, 107b, 109, 121,& 127; Naphthol-based: 99

137 5. Sensing via a disassembly approach

5.3.1. Photophysical properties

With these chemosensors in hand, their photophysical properties and their ability to act as an anion sensor based on their ﬂuorescence response was evaluated. For all compounds, the UV absorbance spectra were meausred from which the corresponding

λEx,max were extracted for subsequent ﬂuorescence measurements.

5.3.1.1. Fe3+-Salen

3+- As Fe salen is reported as an chemosensor for PPi, it suggest itself to test this sensor on inositol pyrophosphate and (p)ppGpp as these molecules bear a pyrophosphate moiety and are known to complex Fe3+. Fluorescence measurements were carried out with probe 89 (50 µM) in Tris buﬀer (10 mM, pH 7.4) which was incubated for 30 mins with 1.0eq. analyte and subsequently the emission intensity was measured. Figure 5.8 depicts the emission response to various analytes (nucleotides, PolyP, (p)ppGpp and inositol polyphosphates) after 30 mins incubation with the anions.

Figure 5.8.: Normalized emission intensity of 89 (50 µM) at 500 nm in Tris buﬀer (10 mM, pH 7.4) in presence of various anions (1.0 eq.) after 30 mins. λEx = 350 nm, λEm = 500 nm

From the diagram, it can be seen that nucleotides are not able to induce ﬂuorescence while PPi results in a response as well as P22 and P45, however the signal is less intense for polyP. This might be because the concentration of the PolyP samples are calculated

138 5.3. Sensor evaluation according their monomoer concentration what might lead to a lower PolyP concentrations. Inositol phosphates are also able to induce the disassembly of Fe3+-salen where

5-PP-InsP5 shows higher eﬃciency as InsP6 and incubation with pppGpp results in the highest ﬂuorescence response of the tested analytes.

5.3.1.2. HBO-based chemosensors

As discussed in chapter General Introduction, HBO-based chemosensors are able to form an excited-state intramolecular proton transfer (ESIPT), with the keto- and enol-form (Figure 5.9b), thus giving the possibility to raise the sensitivity of anion detection due to the low background ﬂuorescence. First, the Cu2+ complexed sensor 109 (Scheme 5.9a) will be evaluated. In this sensor, formation of the keto form is blocked by Cu2+ and the emission intensity at 500 nm should be low. Addition of an anion would lead to demetalization of Cu2+, subsequent formation of the keto form resulting in an enhanced emission intensity at approx. 500 nm.

N O 2+ O Cu O ESIPT O O O N N N H O O O H enol tautomer keto tautomer 109 λEm ~ 420 nm λEm ~ 500 nm

(a) (b) Figure 5.9.: a) Chemical structures of chemosensors 109. b) Excited State Intramolecular Proton Transfer (ESIPT).

This assumption was conﬁrmed by ﬂuorescence experiments. Measuring complex 109 in HEPES (10mM, pH 7.4) gave a low emission at 400 nm (Figure 5.10, black curve). Next, various anions were added to track the emission change over time. The excitation wavelength was set to λEx = 350 nm and the emission was recorded from 370 - 700 nm for 15 mins. The measurements were carried out in HEPES (10 mM, pH 7.4) and with added: PPi, InsP6, 5-PP-InsP5 and pppGpp (10 µM, 10 eq., each).

139 5. Sensing via a disassembly approach

Figure 5.10 shows the emission intensity scans with respective anion over time. Mea- surements were carried out every 5 mins for 15 mins. The black curves in Figure 5.10 are emission scans of sensor 109 in buﬀer, then the anion is added and measured immediately (red curve) and every 5 mins (blue, green, purple).

While InsP6 induces only a low (3-fold) enhancement in emission intensity (Figure

5.10b) pyrophosphaste (PPi) and 5-PP-InsP5 (Figure 5.10a and 5.10d) have almost the same impact on the sensor. The emission enhancement at 520 nm is for both analytes about 10-fold. The reason for the similar results might be due to the pyrophosphate moiety contained by 5-PP-InsP5. However, the largest enhancement in emission intensity is induced by pppGpp (Figure 5.10c). Addition of 10 eq. pppGpp results in a 24-fold increase of the ﬂuorescence signal at 520 nm.

2- O O OPO3 2- 2- P P O3PO OPO3 O O O O O 2- 2- O3PO OPO3 2- OPO3

(a) PPi (b) InsP6

O O O O N 2- NH OPO3 O P O P O P O N 2- 2- O O O O N NH2 O3PO OPO3

O O OH 2-O PO OPO 2- O P O P O 3 3 O O O O P O O O P O O

(c) pppGpp (d) 5-PP-InsP5 Figure 5.10.: Emission scans over time (0-15 mins) of 109 (1 µM in HEPES 10 mM, pH 7.4) with added a) PPi (10 eq.). b) InsP6 (10 eq.). c) ppppGpp (10eq.). d) 5-PP-InsP5 (10 eq.). λEx = 350 nm, λEm = 370-750 nm.

140 5.3. Sensor evaluation

However, as for InsP6, PPi and 5-PP-InsP5 the emission plateau seemed to be achieved, the signal for pppGpp still increased. This indicates that 109 could be a convenient chemosensor for magic spot nucleotides which will be further developed in future.

Comparison of InsP6 and 5-PP-InsP5 after 15 mins gave that the signal intensity induced by 5-PP-InsP5 is approx. 3 times higher than that for InsP6. This could be an advantage in assays to determine whether higher phosphorylated inositol phosphates are generated in the presence of InsP6. Figure 5.11 shows the fluorescence enhancement over time at 520 nm. In this Figure it can be seen that pppGpp exhibits a significant higher fluorescence signal then the other analytes.

Figure 5.11.: Emission 109 (1 µM) at 520 nm with time and added anions (10 µM, 10eq.) in Tris buﬀer (pH 7.4). λEx = 350 nm, λEm = 520 nm.

Next, chemosensor 107a was investigated. In this sensor, the fluorescent core is the same as in 109, but two molecules are connected trough a ethylenediamine bridge and complexed with Fe3+. However, the same ESIPT fluorescence turn-on sensing mechanism should take place after removal of the metal from the ligand by an analyte and subsequent ESIPT formation. Figure 5.12 summarizes the fluorescence measurements of 107a with added analytes.

For all four anions (PPi, InsP6, 5-PP-InsP5, pppGpp), a 2-fold increase could be observed within 15 mins. Although the ﬂuorescence of only complex 107a in HEPES is already

141 5. Sensing via a disassembly approach high, the eﬀect by the anion to the sensor can be distinguished from the background ﬂuorescence.

O 3+ N Fe O N N

O 107a

Scheme 5.12: Chemical structure of chemosensor 107a.

Since all of the added anions induce a similar increase of the signal sensor 107a could be used as stain for methods that need non-selective sensing, such as for PAGE staining.

2- O O OPO3 2- 2- P P O3PO OPO3 O O O O O 2- 2- O3PO OPO3 2- OPO3

(a) PPi (b) InsP6

O O O O N 2- NH OPO3 O P O P O P O N 2- 2- O O O O N NH2 O3PO OPO3

O O OH 2-O PO OPO 2- O P O P O 3 3 O O O O P O O O P O O

(c) pppGpp (d) 5-PP-InsP5 Figure 5.12.: Emission scans over time (0-15 mins) of 107 (1 µM in HEPES 10 mM, pH 7.4) with added a) PPi (10 eq.). b) InsP6 (10 eq.). c) pppGpp (10eq.). d) 5-PP-InsP5 (10 eq.). Every 5 mins, λEx = 350 nm, λEm = 370-750 nm.

142 5.3. Sensor evaluation

Sensor 107b (Figure 5.13a) is a congener of the before discussed molecules. Again, a HBO-core is found in the design with an ester substituent installed at the molecule. Due to metal complexation, ﬂuorescence at 500 nm is turned-oﬀ. Emission measurements of the ESIPT sensor with added anions showed dual emission bands. Due to limited material available and the high amounts which would be needed for testing e.g. 10.0 eq., this measurements were carried out with 1.0 eq. of anions. The enol tautomer gave emission at 430 nm and the keto tautomer gave emission at 500 nm upon anion addition (Figure 5.13a). Both wavelength can be used for the detection of the analyte binding event.

O O O

O 3+ N Fe O N N

O O 107bO

(a) (b) Figure 5.13.: a) Emission spectra over time (0-10 min) of 107b (50 µM) in HEPES (10 mM, pH 7.4) with added InsP6 (50 µM, 1eq.). λEx = 350 nm, λEm = 370-750 nm. b) Normalized emission intensity of 107b (50 µM) at 500 nm in Tris buﬀer (10 mM, pH 7.4) in presence of various anions (1.0 eq.). λEx = 350 nm, λEm = 500 nm.

Figure 5.13b shows that after 30mins of incubation with 5-PP-InsP5, pppGpp and

PPi the emission intensity at 500 nm is also increasing. Chemosensor 107b shows no selectivity with the tested anions. However, the turn-on ﬂuorescence can be useful for staining techniques which do not require selectivity.

143 5. Sensing via a disassembly approach

N Zn2+ N N N Zn2+ O N N O N N O O N N 2+ N 2+Zn N Zn N N 121 127 (a) (b) Figure 5.14.: Chemical structures of chemosensors a) 121. b) 127

The ﬂuorescence emission shift from 400 nm to 500 nm of 121 (12 µM)(Scheme 5.14a) in combination with PPi (10.0 eq.) irradiated at λ = 365 nm could not be reproduced using the condition from the literature.[15] Emission measurements in HEPES (10 mM, pH 7.4) gave a peak at 420 nm which could not be further inﬂuenced by addition of

PPi (10 eq.). Carrying out the emission measurements in DMSO resulted in a second emission maximum at 520 nm. However, also after 45 mins of incubation both emission maxima were still present. Measuring chemosensor 127 (30 µM) in HEPES (10 mM, pH 7.4) with an excitation wavelength of 365 nm gives an emission maximum at 420 nm, which is not further aﬀected by the addition of PPi (10.0 eq.).

5.3.1.3. Naphthol-based chemosensors

The synthesised naphthol-derivative 99 (10 µM) in HEPES (10 mM, pH 7.4) did not show a significant change in fluorescence intensity or emission shift upon analyte addition. Measuring the fluorescent core 94 (50 µM) of the chemosensor in Tris gives an emission maximum at 600 nm. This indicates that the demetalation or hydrolysis process is slow in this sensor molecule.

144 5.3. Sensor evaluation

N 3+N N O Fe O

99 N

(a) (b) Figure 5.15.: Emission scans of 99 (10 µM in Tris 10 mM, pH 7.4) in presence of various anions (10.0 eq.). λEx = 415 nm, λEm = 475-700 nm. b) Chemical structure of sensor 99

5.3.1.4. PAGE-Gel staining

Polyacrylamide gel electrophoresis is a technique for the identification of highly phosphorylated compounds. Today, staining protocols using toluidine blue and DAPI as staining dyes are reported and staining with toluidine blue is used on a daily basis (see chapter State of the Art - Analytics). The use of the chemosensors, developed in this chapter, for staining polyacrylamide gels would give access to fast procedures as the fluorescence response to the analytes can be reported within a few minutes. Furthermore, these molecules are accessible in few synthetic steps with good yields, making preparation of staining solutions convenient. Polyacrylamide gels are commonly run in TBE buffer. As EDTA would decomplex the metal from the sensor molecules this would hinder staining of the gels. An attempt to run a gel, loaded with pppGpp, in EDTA as well as in TB buffer was made.

145 5. Sensing via a disassembly approach

Scheme 5.13: Polyacrylamide gel run in A) TBE-buﬀer for 20 h (500V), stained with toluidine blue and 89 (500 µM in Tris/HCl pH 7.4) for 1hs. pppGpp in sequence from left to right: 150 µM (4.5 nmol), 300 µM (9 nmol), 450 µM (13.5 nmol), 600 µM (18 nmol), 600 µM (18 nmol), 450 µM (13.5 nmol), 300 µM (9 nmol), 150 µM (4.5 nmol). B) TB-buﬀer for 20 hs (500V), stained with toluidine blue and 89 (250 µM in Tris/HCl pH 7.4) for 1h. pppGpp in sequence from left to right: 150 µM (4.5 nmol), 300 µM (9 nmol), 450 µM (13.5 nmol), 600 µM (18 nmol), 600 µM (18 nmol), 450 µM (13.5 nmol), 300 µM (9 nmol), 150 µM (4.5 nmol). Gel stained with 89 was recorded on a gel documentation system at λEx = 312 nm.

Figure 5.13 shows two polyacrylamide gels loaded with pppGpp and Orange G Dye (OG) as a horizontal reference. Gel A was run in TBE buffer and gel B in TB buffer. The gels were cut in the middle and the left part was stained with toluidine blue and the right part with 89 (500 or 250 µM) for 1h. It is noticeable that the staining ability for the given concentrations of pppGpp with 89 is comparable to toluidine blue. Concentrations of 600 and 450 µM give dark bands (lane 5 & 6). Also lower concentrations 300 and 150 µM results in visible bands stained with toluidine blue and 89. The influence of EDTA seems to be low in regards to staining solution (89) and migration of the analyte in the gel. Although this was not further explored it can be assumed that EDTA can be removed from the running buffer. EDTA was initially used as a component for running buffers to complex Mg2+, thus impeding DNA degrading enzymes. Enzymes which are commonly used today do not possess a DNA degrading behaviour, thus EDTA can be removed from the running buffer.[16]

146 5.3. Sensor evaluation

The next step was to screen a concentration series from pppGpp with the Fe3+-Salen complex 89 to assess the staining ability of this dye. In Figure 5.16 the salen stained gels can be seen as black and white picture. These gels are recorded on a gel-documentation system at λEx = 312 nm. One gel was stained with toluidine blue in order to varify the analytes bands (Gel C).

Figure 5.16.: Polyacrylamide gel run in TB-buﬀer for 4h (500V), stained with Fe3+-Salen 89 (250- 25 µM) in HEPES buﬀer (10 mM, pH 7.4) (A,B,D,E,F, black and white picture) and with toluidine blue (C, blue picture). A) pppGpp in sequence from left to right: 600 µM (18 nmol, 1), 150 µM (4.5 nmol, 2), 100 µM (3.0 nmol, 3). Stained for 30 mins with Fe3+-Salen (250 µM). B) pppGpp in sequence from left to right: 600 µM (18 nmol, 1), 150 µM (4.5 nmol, 2), 100 µM (3.0 nmol, 3). Stained for 30 mins with Fe3+-Salen (150 µM). C) pppGpp in sequence from left to right: 600 µM (18 nmol, 1), 150 µM (4.5 nmol, 2), 100 µM (3.0 nmol, 3). Stained for 1 h with Toluidine blue. D) pppGpp in sequence from left to right: 300 µM (9.0 nmol, 4), 150 µM (4.5 nmol, 5), 100 µM (3.0 nmol, 6). Stained for 30 mins with Fe3+-Salen (100 µM). E) pppGpp in sequence from left to right: 300 µM (9.0 nmol, 4), 150 µM (4.5 nmol, 5), 50 µM (1.5 nmol, 6). Stained for 30 mins with Fe3+-Salen (50 µM). F) pppGpp in sequence from left to right: 150 µM (4.5 nmol, 7), 100 µM (3.0 nmol, 8), 50 µM (1.5 nmol, 9). Stained for 30 mins with Fe3+-Salen (25 µM). Gels stained with 89 were recorded on a gel documentation system at λEx = 312 nm.

Gel C shows the horizontal reference Orange G dye which is the ﬁrst lane appearing and the analyte band close to it. The salen stained gels exhibit three bands which could indicate that the bands at the bottom are either a degrading product from the analyte or components from Orange G dye. Until now this was not further investigated. However, it can be seen that 89 in high concentrations (250 µM) stains less then lower concentrations. It was observed that at concentration of 250 and 150 µM the sensor

147 5. Sensing via a disassembly approach precipitates immediately after adding to the HEPES-buﬀer. With this concentration series, Fe3+-Salen 89 sensor was able to stain pppGpp down to 50 µM (1.5 nmol), The LoD of Toluidine blue for pppGpp is 0.15 nmol.[17]

5.4. Summary & Outlook

In this chapter, a library of chemosensors has been synthesized relying on the disassembly approach and a combination of latter with subsequent formation of an ESIPT sensor.

Measuring the ﬂuorescence response of the literature reported Fe3+-Salen complex[6], 89 gave a ﬂuorescence response to inositol polyphosphates, MSN and polyP. PolyP gave low signals compared to InsPs and MSN but it has to be kept in mind that the concentration of polyP might be lower due to the polymer polydispersity. These results highlight the applicability of this easy to prepare chemosensor for phosphorylated compounds. Furthermore, it is an advantage that the sensor is not responding to nucleotides, such as GTP and also has a small signal with ATP. As these molecules are present in a high concentration in cells they could impede the sensing.

The new HBO-based chemosensors 107a, 107b and 109 are very promising in terms of sensing phosphorylated compounds (Figure 5.17). Chemosensor 107a and 107b exhibits a ﬂuorescence turn-on response to all tested phosphorylated anions. However, 107b displayed dual emission maxima (420 nm enol-form and 500 nm keto-form) whereas 107a shows much more intense emission maxima at 500 nm and only a small signal at 420 nm in the ﬂuorescence emission scans. O O O O N O N O 3+ N Fe N O 3+ N O Fe O N 2+ O Cu O N N O O N N O O O O O 107a 107b 109 (a) (b) (c) Figure 5.17.: Chemical structures of chemosensors a) 107a. b) 107b. c) 109 synthesized in this chapter.

148 5.4. Summary & Outlook

It is interesting that only in 107b a dual emission is observed. As the phenol side of 107a and 107b is substituted either with a electron withdrawing or donating group it can be assumed that the observed emission spectrum is a result of diﬀerent substituents.

O ESIPT O

N N H O O H enol tautomer keto tautomer λEm ~ 420 nm λEm ~ 500 nm

Figure 5.18.: keto- and enol-tautomer of 2-(2’-hydroxyphenyl)benzoxazole (HBO) (100).

A computational study from Azarias et al. was carried out in order to rationalize why dual emission is only observed in some compounds.[18] As the ESIPT mechanism is sensitive to the substituents in the fluorophore as well as to the environment it is difficult to estimate the outcome of the emission spectra. Their studies only include sensor molecules measured in organic solvents. The ESIPT mechanism is pH sensitive which makes it further difficult to predict the sensing behaviour for sensor molecules used in aqueous buffers. However, the trend they found for HBO-based molecules is that an electron withdrawing group at the phenol side can favour both the enol and keto tautomers (Figure 5.18). This might explain our observations of a dual emission for 107b. Additionally they found that adding a donor substituent at the phenol side tend to have dual or enol emission. This is not consistent with our observations but still it might be interesting to modify the sensor scaffold with different electron withdrawing and donating groups to tune the emission spectra.

The use of the Fe3+-Salen complex 89 as a gel stain was successfully proven. Staining the polyacrylamide gel with a salen staining solution for 1h gave black bands when recorded on a gel documentation system at λEx = 312 nm. Screening of diﬀerent concentrations of staining solution with diﬀerent concentration of loaded pppGpp leads to the assumption that at higher concentrations (250 µM) the complex might decomplex or precipitate from the aqueous solution as for lower concentrations (150 and 50µM) better staining results were obtained. Although the LoD of toluidine blue for pppGpp is better with 0.15 nmol

149 5. Sensing via a disassembly approach it is advisable to proceed research on reﬁning the staining protocol with 89 as it might be a promising candidate for lowering the staining time from 2 hrs to 30 mins. Also the HBO-based ESIPT sensors should be examined in their ability to act as a gel stain. Due to their rather poor solubility in aqueous buﬀer varying concentrations of organic solvent should be accessed to yield an optimal result or an sensor molecule bearing a PEG-group should be synthesized.

150 5.5. Experimental

5.5. Experimental

5.5.1. General remarks

Reactions were carried out in ﬂame-dried glassware under argon atmosphere, unless noted otherwise. Reaction control was performed via TLC.

Reagents were purchased from commercial suppliers and used without further puriﬁ- cation.

Thin layer chromatography wa performed on Merck silica gel 60 F254 plates (0.25 mm layer thickness, with ﬂuorescence indicator). TLC was analyzed by UV (λ =,254 nm), ﬂuorescence (λ =,368 nm).

151 5. Sensing via a disassembly approach

Chemistry at the University of Freibug. All signals are referenced to an internal solvent signal standard (CDCl3, δ =77.1 ppm; MeOD4, δ =49.0 ppm; DMSO-d6, δ =39.52 ppm).

UV/Vis-Absorption Sepctra were recorded on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Transparent) or on a Shi- madzu UV-1800 UV/VIS Spektrophotometer using a 2 mL cuvette.

Fluorescence Spectra were recorded on a Perkin Elmer LS55 Luminescence Spectrom- eter using 3 mL cuvettes or on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Black).

Polyacrylamide gel electrophoresis (PAGE) PAGE was carried out on a Hoefer SE660 Tall Standard Dual Cooled Vertical Unit. The PAGE procedure was conducted according to the general procedure as described by Losito et al..[19] Following buffers and solutions for gel electrophoresis were prepared:10 × Tris/ Borate/ EDTA (TBE) buffer (0.89 M tris-HCl, 0.89 M boric acid, 20 mM EDTA, pH 8.3), 10 × Tris/ Borate (TB) buffer (0.89 M tris-HCl, 0.89 M boric acid, pH 8.3), 1 × Orange G dye(10 mM tris-HCl, 1 mM EDTA, 30 % (w/v) glycerol, 0.1 % (w/v) OrangeG, pH 7.0).

0.05%(w/v) ammonium persulfate (APS) (200 µL of 10% APS in milli-Q H2O) were

152 5.5. Experimental stirred for 1 min at 0°C, followed by the addition of 0.05% (v/v) tetramethylethylene- diamine (TEMED) (20 µL). After stirring for 1 min, the mixture was poured between the pre-casted glass-plates and a 15 lane comb was inserted. The solution was allowed to polymerize for 25 min at room temperature. After polymerization, the comb was removed and gels were pre-run at 4°C in 1 x TBE or 1 x TE buffer for 30 min at 300 V. Samples were prepared and 1 × Orange G dye (7 µL) was added to each sample prior to loading. Wells were washed with 1 x TBE or 1 x TB buffer by using a syringe and needle to remove any precipitates and non-polymerized gel debris. The gel was then loaded and run at 4°C in 1 x TBE or 1 x TB buffer for 4 h and at 500 V. After run, the gel apparatus was disassembled and the gel was stained for 15 min with staining solution. Finally, the gel was recorded on a transilluminator plate with λEx = 312 nm.

153 5. Sensing via a disassembly approach

5.5.2. Synthesis

2,2’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene))bis(methaneylylidene))diphenol (93)

N N OH HO

Salicylaldehyde (93) (1.04 mL, 10 mmol, 2.0 eq.) was dissolved in EtOH (5 mL) and ethylenediamine (91) (334 µL, 5.0 mmol, 1.0 eq.) was added. The mixture was stirred for 1 h at RT. The precipitate was ﬁltered, washed with cold EtOH and dried in vacuo. The product was obtained as a yellow solid (1.3 g, 4.8 mmol, 97%).

Analytical data were in agreement with literature.[20]

Rf (SiO2, Pentane/EE 5:1 (vol/vol)) 0.3. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 13.37 (s, 2H), 8.59 (s, 2H), 7.42 (dd, J = 7.6, 1.8 Hz, 2H), 7.37 – 7.27 (m, 2H), 7.01 – 6.79 (m, 4H), 3.92 (s, 4H). HRMS (ESI) m/z: calcd.: 269.1285, found: 269.1285 [M+H+]+.

Probe (89)

N 3+ Fe N O O

93 (1.3 g, 4.8 mmol, 1.0 eq.) was suspended in EtOH (80 mL), FeCl3 (778 mg, 4.8 mmol, 1.0 eq.) was added and the mixture was reﬂuxed for 1 h. The solvent was removed under reduced pressure, the solid was partitioned between MeOH (50 mL) and the product precipitated with Et(2)O to give the product as a purple solid (850 mg, 2.5 mmol, 52%) HRMS (ESI) m/z: calcd.: 322.0399, found: 322.0401 [M+H+]+.

UV λ = 350 nm Fluorescence λ = 500 nm

154 5.5. Experimental

2-(benzo[d]oxazol-2-yl)-4-methylphenol (105a)

N HO

A solution of 2-hydroxy-5-methylbenzylaldehyde (102) (1.00 g, 7.34 mmol, 1.0 eq.) and 2-aminopehnol (103) (0.80 g, 7.34 mmol, 1.0 eq.) in MeOH was stirred for 2 hs. (Di- acetoxyiodo)benzene (2.60 g, 8.07 mmol, 1.1 eq.) was added and the mixture stirred overnight. The solvent was removed under reduced pressure and the residue was puriﬁed by ﬂash column chromatography (pentane/EtOAc 98:2) and the product was obtained as a yellow solid (452 mg, 2.00 mmol, 27%). Analytical data were in agreement with literature.[15]

Rf (SiO2, Pentane/EE 98:2 (vol/vol)) 0.8. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 7.84 – 7.82 (m, 1H), 7.77 – 7.69 (m, 1H), 7.65 – 7.57 (m, 1H), 7.44 – 7.34 (m, 2H), 7.29 – 7.22 (m, 1H), 7.06 – 7.01 (m, 1H), 2.37 (s, 3H). HRMS (ESI) m/z: calcd.: 226.0863, found: 226.0864 [M+H+]+.

3-(benzo[d]oxazol-2-yl)-2-hydroxy-5-methylbenzaldehyde (101a)

N HO O

Compound 105a (119 mg, 0.53 mmol, 1.0 eq.) and urotropin 106 (162 mg, 1.16 mmol, 2.2 eq.) were dissolved in TFA (10 mL) and reﬂuxed over night. The reaction was cooled to RT, 4 M HCl (25 mL) was added and the solution stirred for 15 mins. The mixture was extracted with DCM (3x 25 mL) and washed with 4 M HCl (50 mL) and brine (50 mL).

The organic phase was dried over Na2SO4 and the solvent removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (pentane/EtOAc 9:1) and the product 101a was obtained as a yellow solid (97 mg, 0.38 mmol, 72%).

155 5. Sensing via a disassembly approach

Analytical data were in agreement with literature.[15]

Rf (SiO2, Pentane/EE 9:1 (vol/vol)) 0.7. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 10.60 (s, 1H), 8.15 – 8.07 (m, 1H), 7.80 (dt, J = 2.3, 0.7 Hz, 1H), 7.79 – 7.74 (m, 1H), 7.67 – 7.60 (m, 1H), 7.47 – 7.39 (m, 2H), 2.42 (s, 3H). HRMS (ESI) m/z: calcd.: 254.0812, found: 254.0812 [M+H+]+. UV λ = 356/375 nm Fluorescence λ = 520 nm

6,6’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene)) bis(methaneylylidene))bis(2-(benzo[d]oxazol-2-yl)-4-methylphenol) (108a)

N HO N

HO N N

Compound 101a (100 mg,0.395 mmol, 2.0 eq.) was suspended in EtOH and ethylenediamine (91) (0.013 mL, 0.197 mmol, 1.0 eq.) was added. The mixture was reﬂuxed until white precipitate was formed and then stirred at RT over night. The reaction was cooled to 0° C and ﬁltered. The residue was washed with cold EtOH and dried to yield the product 108a as a yellow solid (71 mg, 13.3 µmol, 68%).

1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.46 (s, 2H), 8.02 (dd, J = 2.4, 0.8 Hz, 2H), 7.88 – 7.78 (m, 2H), 7.68 – 7.55 (m, 2H), 7.40 – 7.30 (m, 6H), 4.03 (s, 4H), 2.34 (s, 6H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 163.6, 161.6, 158.6, 150.1, 142.0, 135.0, 133.4, 127.9, 125.1, 124.6, 120.3, 114.4, 110.6, 59.8, 20.4. HRMS (ESI) m/z: calcd.: 531.2027, found: 531.2025 [M+H+]+.

156 5.5. Experimental

Probe (107a)

O 3+ N Fe O N N

Compound 107a was prepared using the procedure as for 89 in DCM. The product 107a (30 mg, 55µmol, 93%) was obtained as a green solid.

UV λ = 356/375 nm Fluorescence λ = 520 nm

Methyl-3-(benzo[d]oxazol-2-yl)-4-hydroxybenzoate (105b)

O O O

N HO

Compound 105b was prepared using the procedure as for 105a. The product was ﬁltered from the reaction mixture and washed with cold MeOH to obtain 105b (610 mg, 2.2 mmol, 82%) as a white solid.

1 H-NMR (400 MHz, DMSO-d6, δ/ppm): 11.76 (s, 1H), 8.62 (d, J = 2.3 Hz, 1H), 8.07 (dd, J = 8.6, 2.3 Hz, 1H), 7.89 (t, J = 7.4 Hz, 2H), 7.56 – 7.45 (m, 2H), 7.25 (d, J = 8.7 Hz, 1H), 3.88 (s, 3H).

13 C-NMR (101 MHz, DMSO-d6) δ/ppm): 165.2, 161.3, 161.2, 149.1, 139.5, 134.3, 129.7, 126.1, 125.4, 121.3, 119.4, 117.7, 111.2, 111.1, 52.2. HRMS (ESI) m/z: calcd.: 270.0761, found: 270.0764 [M+H+]+.

157 5. Sensing via a disassembly approach

Methyl 3-(benzo[d]oxazol-2-yl)-5-formyl-4-hydroxybenzoate (101b)

O O O

N HO O

Compound 101b was prepared using the procedure as for 101a. Puriﬁed by ﬂash column chromatography (pentane/EE/DCM 9:1:1 to 1:1:1) gave the product 101b (329 mg, 1.10 mmol, 74%) as a white solid.

Rf (SiO2, Pentane/EE 9:1 (vol/vol)) 0.2. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 10.44 (s, 1H), 8.76 (d, J = 2.2 Hz, 1H), 8.43 (d, J = 2.2 Hz, 1H), 7.98 – 7.88 (m, 2H), 7.61 – 7.47 (m, 2H), 3.92 (s, 3H). HRMS (ESI) m/z: calcd.: 298.0710, found: 298.0712 [M+H+]+.

UV λ = 356/375 nm Fluorescence λ = 520 nm

Methyl 3-(benzo[d]oxazol-2-yl)-5-((E)-((2-(((E)-3-(benzo[d]oxazol-2-yl) -2-hydroxy-5-(methoxycarbonyl)benzylidene)amino)ethyl)imino)methyl) -4-methylbenzoate (108b)

O O O

N HO N

HO N N

O O O

Compound 108b was prepared using the procedure as for 108a. The product 108b (94 mg, 0.152 mmol, 61%) was obtained as a yellow solid.

158 5.5. Experimental

1 H-NMR (300 MHz, CDCl3, δ/ppm): 1H 12.87 (s, 2H), 10.62 (s, 2H), 8.97 (d, J = 2.2 Hz, 2H), 8.66 (d, J = 2.2 Hz, 2H), 7.90 – 7.64 (m, 4H), 7.55 – 7.42 (m, 4H), 3.98 (s, 10H). HRMS (ESI) m/z: calcd.: 619,1823, found: 619,1826 [M+H+]+.

Probe (107b)

O O O

O 3+ N Fe O N N

O O O

Compound 107b was prepared using the procedure as for 107a. The product 107b (10 mg, 14.8 µmol, 93%) was obtained as a red solid.

UV λ = 370 nm Fluorescence λ = 420,500 nm

7-(dimethylamino)naphthalen-2-ol (96)

N OH

2,7-Dihydroxynaphthalene (95) (3.0 g, 18.7 mmol, 1.0 eq.) and sodium disulﬁte (11.9 mL,

93.5 mmol, 5.0 eq.) were suspended in water (10 mL), diemthylamine (40 wt.% in H2O) (128) (11.9 mL, 93.5ß,mmol, 2.0eq.) was added and the reaction was reﬂuxed over night. After cooling to RT concentrated HCl was added and extracted with DCM

(3x 50 mL). The organic layer was dried over Na2SO4 and the solvent was removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (pentane/EtOAc 9:1) and the product 96 was obtained as a white solid (1.48 g, 7.90 mmol, 42%).

159 5. Sensing via a disassembly approach

Analytical data were in agreement with literature.[10]

Rf (SiO2, pentane/EE 9:1 (vol/vol)) 0.2. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 7.68 – 7.54 (m, 2H), 7.03 (dd, J = 9.0, 2.5, Hz, 1H), 6.98 (d, J = 2.5 Hz, 1H), 6.86 (dd, J = 8.7, 2.5 Hz, 2H), 3.05 (s, 6H). HRMS (ESI) m/z: calcd.: 186.0924, found: 186.0924 [M-H+]−.

6-(dimethylamino)-3-hydroxy-2-naphthaldehyde (94)

N OH

96 (700 mg, 3.74 mmol, 1.0 eq.) was added at 0° C to a solution of DMF (1.75 mL,

22.4 mmol, 6.0 eq.) and POCl3 (0.85 mL, 9.35 mmol, 2.5 eq.). The reaction was stirred at 50° C for 2.5 hs. The mixture was poured into ice water (25 mL) followed by adjusting the pH to 8 with sodium acetate. The obtained yellow precipitate was filtered and purification was achieved by flash column chromatography (pentane/EtOAc 9:1) to obtain the product 94 as a yellow solid (750 mg, 3.48 mmol, 93%).

Analytical data were in agreement with literature.[10]

Rf (SiO2, Pentane/EE 1:1 (vol/vol)) 0.3. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 10.33 (s, 1H), 8.83 (s, 1H), 7.84 (d, J = 9.1 Hz, 1H), 7.65 (d, J = 8.9 Hz, 1H), 7.13 (d, J = 9.1 Hz, 1H), 7.04 (d, J = 8.7 Hz, 1H), 3.15 (s, 6H). HRMS (ESI) m/z: calcd.: 216.1019, found: 216.1020 [M+H+]+. 3,3’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene))bis(methaneylylidene))bis(7- (dimethylamino)naphthalen-2-ol) (98)

N OH

160 5.5. Experimental

94 (100 mg, 0.464 mmol, 2.0 eq.) was suspended in EtOH and ethylenediamine (0.016 mL, 0.232 mmol, 1.0 eq.) was added. The mixture was reﬂuxed until white precipitate was formed and then stirred at RT over night. The reaction was cooled to 0° C and ﬁltered. The residue was washed with cold EtOH and dried to yield the product 98 as a white solid (63 mg, 0.138 mmol, 60%).

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 8.72 (s, 2H), 8.60 (d, J = 2.5 Hz, 2H), 7.74 (d, J = 8.9 Hz, 2H), 7.64 (d, J = 8.7 Hz, 2H), 7.11 (d, J = 8.8 Hz, 2H), 6.92 (dd, J = 8.8, 2.4 Hz, 2H), 4.11 (s, 4H), 2.65 (s, 12H).

13 C-NMR (101 MHz, DMSO-d6) δ/ppm): 161.8, 156.8, 153.6, 133.2, 131.1, 129.4, 124.1, 118.8, 116.2, 114.8, 107.2, 63.4, 45.1. HRMS (ESI) m/z: calcd.: 455.2442, found: 455.2440 [M+H+]+.

Probe (99)

N 3+N N O Fe O

Compound 99 was prepared using the procedure as for 107a. The product 99 (30 mg, 59.0 µmol, 89%) was obtained as a red solid.

UV λ = 356/412 nm Fluorescence λ = 520 nm

5-(benzo[d]oxazol-2-yl)-4-hydroxyisophthalaldehyde (116)

O O

N HO O

161 5. Sensing via a disassembly approach

2-(2-hydroxyphenyl)benzoxazole (1.75 g, 8.28 mmol, 1.0 eq.) and urotropin (106) (2.55 g, 18.22 mmol, 2.2 eq.) were dissolved in TFA (20 mL) and reﬂuxed overnight. After cooling to RT the mixture was poured into 4 M HCl (40 mL), stirred for 30 mins and extracted with DCM (3x40 mL). The combined organic layers were washed with 4 M HCl (60 mL),

H2O (60 mL) and saturated NaCl (60 mL). It was dried over Na2SO4 and the solvent removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 100:0 to 95:5) and the product 94 was obtained as a yellow solid (1.05 g, 3.92 mmol, 47%).

Analytical data were in agreement with literature.[15]

Rf (SiO2, DCM/MeOH 95:5 (vol/vol)) 0.4. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 13.06 (s, 1H), 10.65 (s, 1H), 10.03 (s, 1H), 8.84 (d, J = 2.2 Hz, 1H), 8.49 (d, J = 2.1 Hz, 1H), 7.99 – 7.78 (m, 1H), 7.74 – 7.63 (m, 1H), 7.56 – 7.41 (m, 2H). 2-(benzo[d]oxazol-2-yl)-4,6-bis(((pyridin-2-ylmethyl)amino)methyl) (120)

HN N

N HO HN N

116 (200 mg, 0.75 mmol, 1.0 eq.) and 2-(aminomethyl)pyridine (193 µL, 1.87 mmol,

2.5 eq.) were dissolved in 1,2-DCE (20 mL) and stirred for 15 mins. NaBH(OAc)3 (400 mg, 1.87 mmol, 2.5 eq.) was added and the mixture stirred over night at RT. Satu- rated NaHCO3 was added and extracted with DCM (3x40 mL). The combined organic layers were dried over Na2SO4 and the solvent removed under reduced pressure. Pu- riﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 98:2) and the product 120 was obtained as a brown oil (299 mg, 0.662 mmol, 88%).

Rf (SiO2, DCM/MeOH 9:1 (vol/vol)) 0.3.

162 5.5. Experimental

1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.58 - 8.54 (m, 2H), 7.97 (d, J = 2.2 Hz, 1H), 7.76 – 7.68 (m, 1H), 7.65 (tt, J = 7.6, 2.0 Hz, 2H), 7.61 - 7.57 (m, 1H), 7.48 (d, J = 2.3 Hz, 1H), 7.40 – 7.31 (m, 4H), 7.20 – 7.13 (m, 2H), 4.02 (s, 2H), 4.01 (s, 2H), 3.97 (s, 2H), 3.86 (s, 2H).

13 C-NMR (101 MHz, CDCl3) δ/ppm): 163.1, 159.5, 159.3, 156.4, 149.5, 149.4, 149.3, 140.2, 136.7, 134.2, 130.9, 127.5, 126.1, 125.5, 125.1, 122.6, 122.5, 122.2, 122.2, 119.4, 110.8, 110.4, 54.5, 54.5, 50.9, 48.9, 45.9. HRMS (ESI) m/z: calcd.: 452,2081, found: 452,2083 [M+H+]+.

2-(benzo[d]oxazol-2-yl)-4,6-bis((bis(pyridin-2-ylmethyl)amino)methyl)phenol (118)

N N N

N HO N N

120 (100 mg, 0.22 mmol, 1.00eq.) and 2-(bromomethyl)pyridine (140 mg, 0.55 mmol, 2.5 eq.) were dissolved in MeCN and DIPEA (300 µL, 1.76 mmol, 1.8 eq.) was added. The mixture was reﬂuxed overnight. The solvent was removed under reduced pressure and the solid partitioned between DCM and washed with water (2x20 mL). The aqueous layer was extracted with DCM (3x20 mL) and the combined organic layers dried over

Na2SO4 and the solvent was removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH 97:3 to 80:20) and the product 118 was obtained as a red oil (40 mg, 63.1 µmol, 29%).

Analytical data were in agreement with literature.[15]

Rf (SiO2, Pentane/EE 9:1 (vol/vol)) 0.35.

163 5. Sensing via a disassembly approach

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.54 – 8.48 (m, 4H), 7.93 (d, J = 2.2 Hz, 1H), 7.79 (d, J = 2.2 Hz, 1H), 7.76 – 7.70 (m, 1H), 7.67 – 7.55 (m, 10H), 7.41 – 7.34 (m, 2H), 7.18 – 7.08 (m, 4H), 3.98 – 3.92 (m, 6H), 3.90 (s, 1H), 3.86 (s, 3H), 3.72 (s, 2H). HRMS (ESI) m/z: calcd.: 634.2925, found: 634.2927 [M+H+]+; calcd.: 656.2750, found: 656.2751 [M+Na+]+. Probe (121)

N Zn2+ N N O

N O N 2+Zn N N

118 (97 mg, 0.153 mmol, 1.00 eq.) was dissolved in MeOH (15 mL) and Zn(NO3)2·6H2O (91 mg, 0.306 mmol, 2.00 eq.) was added in one portion. The mixture was stirred for 2 hs. The solvent was removed under reduced pressure and the obtained solid recrystallized from MeOH/H2O (1:1) to yield the sensor 121 as a brown solid (140 mg, 0.147 mmol, 96%)

UV λ = 360/375 nm Fluorescence λ = 430 nm

N,N -dimethyl-N 2-(pyridin-2-ylmethyl)ethane-1,2-diamine (125)

H N N N

2-Pyridinecarboxaldehyde (123) and 2-(Dimethylamino)ethylamine (124) was dissolved in MeOH (40 mL) and and stirred for 5 min. Then NaBH4 is added and the mixture stirred for 2 hs. The solvent was removed under reduced pressure, the solid was

164 5.5. Experimental

partitioned between DCM (30 mL) and washed with NaHCO3 (30 mL). The aqueous phase was extracted with DCM (3x30mL) the organic layers combined and dried over

Na2SO4.The product was obtained as a brown solid (3.30 g, 18.4 mmol, 98%)

Analytical data were in agreement with literature.[21]

1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.53 (ddd, J = 4.9, 1.8, 0.9 Hz, 1H), 7.62 (td, J = 7.6, 1.8 Hz, 1H), 7.36 – 7.30 (m, 1H), 7.18 – 7.09 (m, 1H), 3.92 (s, 2H), 2.73 (t, J = 6.2 Hz, 2H), 2.44 (t, J = 6.2 Hz, 2H), 2.20 (s, 6H).

2-(benzo[d]oxazol-2-yl)-4,6-bis(((2-(dimethylamino)ethyl)(pyridin-2-ylmethyl) amino)methyl)phenol (126)

N O N N N HO N

Compound 116 (150 mg, 0.56 mmol, 1.0 eq.) was suspended in MeOH (15 mL) and 125 (220 mmg, 1.23 mmol, 2.2 eq.) was added and the mixture stirred until the solution became clear. NaBH4 was added carefully at 0° C and the reaction stirred at RT over night. Saturated NaHCO3 (25 mL) was added and the product was extracted with DCM (6x30 mL). The combined organic layers were dried over Na2SO4 and the solvent removed under reduced pressure. Puriﬁcation was achieved by ﬂash column chromatography (DCM/MeOH/Et3N 10:10:0.1 to 0:100:0.1) and the product 118 was obtained as a brown solid (225 mg, 0.379 mmol, 67%).

1 H-NMR (400 MHz, CDCl3, δ/ppm): 8.56 – 8.51 (m, 2H), 7.94 (d, J = 2.2 Hz, 1H), 7.77 – 7.51 (m, 6H), 7.41 – 7.30 (m, 3H), 7.18 – 7.12 (m, 2H), 4.68 (s, 2H), 3.94 (s, 2H), 3.89 (s, 2H), 3.87 (s, 2H), 2.90 – 2.72 (m, 4H), 2.68 – 2.62 (m, 2H), 2.48 (t, J = 6.2 Hz, 2H), 2.29 (s, 6H), 2.24 (s, 6H).

165 5. Sensing via a disassembly approach

13 C-NMR (101 MHz, DMSO-d6) δ/ppm): 162.9, 159.6, 159.4, 156.5, 149.3, 149.3, 148.9, 140.3, 136.6, 136.5, 133.7, 132.3, 126.7, 125.3, 124.9, 123.3, 122.3, 122.1, 121.9, 119.3, 110.6, 110.6, 64.6, 60.7, 58.9, 57.1, 55.2, 53.1, 51.3, 46.8, 45.5, 45.3. Probe (127)

N Zn2+ N N O

N O N 2+Zn N

Compound 127 was prepared using the procedure as for 121. The product 127 (20 mg, 27.6 µmol, 69%) was obtained as a orange oil.

UV λ = 290/350 nm Fluorescence λ = 420 nm

Probe (109)

N O 2+ O Cu O O N

101a (30 mg, 0.118 mmol, 2.0 eq.) was dissolved in THF (2.0 mL) and CuCl2 (8.00 mg,

59.2 µmol, 1.00 eq.) dissovled in H2O (1.0 mL) was added. The mixture was heated for 1 h and the product ﬁltered (28 mg, 49.2 µmol, 83%).

UV λ = 292/402 nm Fluorescence λ = 525 nm

166 5.5. Experimental

5.5.3. References

Bibliography Chapter Sensing via a disassembly approach

[1] S. Akine, T. Taniguchi, W. Dong, S. Masubuchi, T. Nabeshima, J. Org. Chem. 2005, 70 (5), 1704–1711.

[2] Abhay Nanda Srivastva, Stability and Applications of Coordination Compounds, IntechOpen, 2020.

[3] N. Kumari, F. Zelder, Chem. Commun. 2015, 51 (96), 17170–17173.

[4] H. S. Jung, J. H. Han, Z. H. Kim, C. Kang, J. S. Kim, Org. Lett. 2011, 13 (19), 5056–5059.

[5] H. S. Jung, J. H. Han, Y. Habata, C. Kang, J. S. Kim, Chem. Commun. 2011, 47 (18), 5142–5144.

[6] N. Kumari, H. Huang, H. Chao, G. Gasser, F. Zelder, Chembiochem. 2016, 17 (13), 1211–1215.

[7] L. Bohn, A. S. Meyer, S. K. Rasmussen, J. Zhejiang Univ. Sci. B 2008, 9 (3), 165–191.

[8] D. Winkler, S. Banke, P. Kurz, Z. Anorg. Allg. Chem. 2020, 646 (13), 933–939.

[9] P. N. Borase, P. B. Thale, G. S. Shankarling, ChemistrySelect 2018, 3 (20), 5660– 5666.

[10] Y. Ding, C. Xu, Z. Li, W. Qin, X. Han, X. Han, C. Zhang, C. Yu, X. Wang, L. Li, W. Huang, Chembiochem. 2019, 20 (6), 831–837.

[11] S. Rajasekhar, B. Maiti, K. Chanda, Synlett 2017, 28 (05), 521–541.

[12] Rajender S. Varma, Rajesh K. Saini, Om Prakash, Tetrahedron Lett. 1997, 38 (15), 2621–2622.

167 5. Sensing via a disassembly approach

[13] L. F. Lindoy, Synthesis 1998, 1998 (07), 1029–1032.

[14] Y. Fu, Q.-C. Feng, X.-J. Jiang, H. Xu, M. Li, S.-Q. Zang, Dalton Trans. 2014, 43 (15), 5815–5822.

[15] W.-H. Chen, Y. Xing, Y. Pang, Org. Lett. 2011, 13 (6), 1362–1365.

[16] J. R. Brody, S. E. Kern, BioTechniques 2004, 36 (2), 214–216.

[17] T. M. Haas, P. Ebensperger, V. B. Eisenbeis, C. Nopper, T. Dürr, N. Jork, N. Steck, C. Jessen-Trefzer, H. J. Jessen, Chem. Commun. 2019, 55 (37), 5339–5342.

[18] C. Azarias, Š. Budzák, A. D. Laurent, G. Ulrich, D. Jacquemin, Chem. Sci. 2016, 7 (6), 3763–3774.

[19] O. Losito, Z. Szijgyarto, A. C. Resnick, A. Saiardi, PloS one 2009, 4 (5), e5580.

[20] F. Mohandes, M. Salavati-Niasari, New J. Chem. 2014, 38 (9), 4501–4509.

[21] S. Mundinger, U. Jakob, W. Bannwarth, Chem. Eur. J. 2014, 20 (5), 1258–1262.

168 CHAPTER 6

Sensing via hydrogen bonding

169 6. Sensing via hydrogen bonding

6.1. Background

6.1.1. DAPI

DAPI (Figure 6.1) was synthesized 1971 as a trypanocide by Dann et al..[1] However, DAPI never went to clinical trials as a drug but showed usefull DNA binding properties. In 1975 the ability of DAPI to interact with DNA and forming a ﬂuorescent complex, via displacing water molecules, was described.[2,3] Free DAPI is excited at 360 nm and emits at approximately 460 nm. Binding of the dye into the minor groove of A-T rich sequences of DNA leads to a 20 fold enhancement of ﬂuorescence intensity. Therefore the use of this dye was suited for mitochondrial DNA which is rich in A-T sequences.

(a)

(b) Figure 6.1.: a) Chemical structure of 4’,6-diamidino-2-phenylindole dichloride (DAPI). b) Crystal structure of DAPI (yellow) binding into the minor groove of a DNA helix.

Five years later Miller et al. discovered the potential of DAPI for staining the polyanionic molecule polyphosphate (polyP).[4] Conspicuous ﬂuorescent bodies in Sac- charomyces cerevisiae vacuoles, which were treated with DAPI, drew their attention since DNA was not found in these vacuoles. On the other hand it is known that the polyanion polyphosphate is abundant in yeast vacuoles. As Kapuscinski and Skoczy- las showed that the interactions of DAPI with anionic sodium dodecyl sulphate leads to an increase in ﬂuorescence intensitiy[5] they concluded that polyP might trigger the re-

170 6.1. Background sponse in this case. Indeed, the polyP-DAPI complex leads to a bathochromic emission shift of almost 60 nm (λFree = 460 nm & λpolyP = 520nm, comparison of both emission proﬁles in Figure 6.2 ).[4,6,7] In contrast, small molecules, like orthophosphate, sugar phosphates and acetate, did not aﬀect the emission of DAPI.[6]

Figure 6.2.: Emission proﬁle of DNA-DAPI (green) and polyP-DAPI (red) complexes.Picture taken from reference [8]

In Figure 6.2 the polyP induced emission shift is shown. This significant difference of the maxima can be a useful tool for the detection of highly phosphorylated compounds. Recently, the group of Saiardi investigated the influence of InsP6 on the [9] excitation-emission spetra of DAPI. They found that the InsP6-DAPI complexes emit at approximately 550 nm when excited at 410 nm. With InsP5 the same effect was observed while lower phosphorylated members of the inositol phosphate family, like InsP3 showed no effect. With this dye, problems, like high background fluorescence can be avoided which makes it a candidate of interest for the development of a chemosensor for phosphorylated compounds.

6.1.2. Hydrogen bonds in sensing

It has to be energetically favourable for the analyte to bind the host, thus one challenge of sensing phosphate in aqueous environment arises from the inherently strong hydra- tion.[10] A further issue is that modiﬁed phosphate anions exist in diﬀerent protonated states at neutral pH. These reasons lead to the need of optimized anion receptors in respect to their electrostatic and hydrogen-bond interactions. Functional groups, such

171 6. Sensing via hydrogen bonding as polyammonioum, imidazolium and guanidine moieties are used for phosphate sensors, since they are easily protonated.[11] Furthermore, functional groups like amidines, guanidines and biguanidines possess NH2 donor moieties which are appropriately spaced to form cooperative hydrogen bonds with phosphate analytes. A drawback of these common hydrogen bond donors is their limited application in aqueous solution. However, combination of hydrogen bonding interactions and positively charged binding sites, such as guanidine, might help to reduce this problem to some extent. Nature - as so often - shows us solutions, which we can adopt for our needs. In proteins, the ability of the guanidine group to build hydrogen-bonds has important functions, including bonding to phosphate anions.[12] This is due to the salt-bridges which are formed between the positively charged binding sites and the oxyanions on the biomolecules. Salt-bridges are a speciﬁc form of hydrogen bonds which uses oppositely-charged residues which are in close proximity to each other for electrostatic attraction (Figure 6.3).[13,14] The binding free energy of guanidinium-phosphate is -2.38 kJ/mol.[15]

Figure 6.3.: The formation of salt-bridges between arginine residues and phosphate groups on the DNA backbone is well known. [15] Picture shows the salt-bridge interaction between a phosphate and a guanidine moiety from arginine.

Guanidine is a stronger base than amidine.[16] This is attributed to resonance stabilisa- tion, stabilization by intramolecular hydrogen bonds, Y-aromaticity and the stabilizing eﬀect of solvent molecules on the protonated form.[16] This might give a better aﬃnity to phosphates compared to amidine in water.

172 6.2. Results and Discussion

6.2. Results and Discussion

Since DAPI seemed to be a promising candidate for sensing phosphates, we decided to synthesise it, although it is commercial available. High costs for small amounts in combination with the need to make DAPI analogues were drivers for this decision. There are not many DAPI-syntheses reported in the literature.[1,17,18]

6.2.1. DAPI Synthesis

Figure 6.1 illustrates the retrosynthetic pathway to obtain the target molecule DAPI (34).

NH HN N N NH H 2 N N H NH2 34 129

O H N 2 NH

Br Br N Br H Br 130 132 131

Scheme 6.1: Retrosynthethic pathway describing the synthesis to obtain DAPI 34

The proposed retrosynthetic route of 34 involves a three step reaction sequence. In the ﬁrst step the indole is formed via a Fischer-indole-synthesis. The second step involves a cyanation of the aryl bromides and subsequently transformation of the nitrile into an amidine group is completing the synthesis. Starting from a Fischer-indole-synthesis converting 1-(4-bromophenyl)ethan-1-one (132) and (3-bromophenyl)hydrazine (131) into the corresponding di-bromo derivative (130) by reﬂuxing in polyphosphoric acid (Scheme 6.2). During the ring closure reaction two isomers were formed which had to be separated by column chromatography. This inevitably led to low yields.

173 6. Sensing via hydrogen bonding

H N 2 NH Br Br N H O Br 131 130 (29%) Polyphosphoric acid

MeOH Br Br 132 Br N H 133 (24%)

Scheme 6.2: Fischer indole synthesis to obtain the bromo indole derivative 130

Identiﬁcation of the target isomer was carried out by a nuclear overhauser enhancement spectroscopy (NOESY)-experiment. For the preparation of aromatic nitriles[19] from aryl halides the Rosenmund-von-Braun reaction is one method. This method faces some limitations; for example, the harsh conditions which have to be applied for aryl bromides (150 - 280 ° C) or the stoichiometric amounts of CuI cyanide used, which may complicate the workup of the reaction. To circumvent these problem Buchwald et al. developed a copper-catalyzed domino halogen exchange-cyanation reaction.[20]. They reported that upon the use of 1,2-diamine ligands the copper-catalyzed reactions of aryl halides with nucleophiles are accelerated.[21] Still, aryl bromides were sensitive to the copper precatalyst and addition of 20 mol % KI to the reaction mixture improved the eﬃcacy. Even heterocyclic substrates containing a N-H group - like indoles - are well tolerated as no N-arylation takes place. This made this approach convenient for our needs.

NaCN CuI, KI, Br N,N'-Dimethylethylendediamine N Br N N H Toluene, reflux N H 130 129 (85%)

Scheme 6.3: Cyanation step of 130 by using the copper-catalyzed domino halogen exchange-cyanation protocol from Buchwald et al.. [20]

174 6.2. Results and Discussion

The di-bromo indole 130 was cyanated without the need of protecting the N-H group at the indole and 129 was obtained in 85% yield. The presence of the nitrile groups was confirmed by electrospray ionization (ESI)-high-resolution mass spectrometry (HRMS) and infrared absorption spectroscopy (IR) measurements (Figure 6.4). In the IR spectrum a strong band at 2200 cm-1, due to the stretching mode of nitrile, can be observed, confirming a successful reaction. The bottleneck of this synthesis was the final conversion of the nitrile into the amidine. Commonly, nitriles are converted into amidines via three methods: The Pinner method[22], amidoxime method[23] or the nucleophilic addition of lithium hexamethyldisilazane (LHMDS)[24,25].

100

-1

2200 cm Transmission / % / Transmission

20 N N N H

4000 3500 3000 2500 2000 1500 1000 500

-1

Wavenumbers / cm

Figure 6.4.: IR-spectrum of 129. A sharp band at 2200 cm-1 can be observed, the vibrational band of nitrile.

In the paper from 1971[1] conversion of the nitrile into the amidine was done by a Pinner-reaction. Since this method suﬀers from the need of dry HCl-gas and low yields, we tried to substitute this by modern reaction conditions for amidine formation. In Scheme 6.4 an overview of the attempts for the synthesis of arylamidines are shown.Method A is a Pinner reaction and HCl is generated in situ by controlled hydrolo- sis of thionylchloride in methanol. Workup with ammonia aﬀords the desired amidine.[26] Method B takes advantage of in situ generated lithium bis(trimethylsilyl)amide which can react with nitriles to generate persilylated amidines, such as 134. Using ethanolic

175 6. Sensing via hydrogen bonding

HCl, the intermediate can be hydrolysed directly.[25] This method enables synthesis of unsubstituted amidines derived from nitriles lacking an α-hydrogen or chlorine atom.

Method C Method A N 135

Method B

NH NH NH HCl O N(TMS)2 O 136 134 137

NH4Cl HCl, H2O NH3, MeOH

NH HCl NH2 138

[26] Scheme 6.4: Attempts for synthesis of arylamidine 138. Method A: SOCl2, MeOH, H2O, Et2O, r.t. ; [25] [27] Method B: (Me3Si)NLi*Et2O, 0°C → -78° C ; Method C: MeONa, MeOH, r.t. .

Method C is an alternative to the Pinner reaction which uses sodium methoxide as nucleophile for the formation of a carboximidate. Acetic acid and ammonium chloride were added directly to the reaction solution, quenching remaining methoxide and initiating the formation of the amidine groups.[27] After checking all of these possibilities, by using benzonitrile as an test compound, Method B turned out as the method of choice for further reactions (Scheme 6.5).

N 1) LiHMDS HN NH 2 2) HCl/H2O HCl Et2O

135 138 (50%)

Scheme 6.5: Test reaction with benzonitrile 135 to convert into amidine 138 using Method B.

Treating benzonitrile with lithium bis(trimethylsilyl)amide in Et2O at 0° C and subsequent hydrolysis of the intermediate into the amidine hydrochloride salt provided the crude product. The product could then be puriﬁed by acid-base extraction. 13C-NMR

176 6.2. Results and Discussion provided evidence of the formation of the product 138 (Figure 6.5) due to the peak at 165 ppm which is characteristic for amidine carbons.

Figure 6.5.: 13C-NMR trace of benzimidamide 138 showing chemical shift of amidine carbon.

Having the method established the amidine synthesis was applied to the dinitrile-indole derivative 129. As the LHMDS would deprotonate the indole proton this would prevent the desired reaction because of the delocalisation of the negative charge on the nitrile. The indole NH was protected transiently using the Boc group.

N N (Boc) O, DMAP N 2 N H Boc N DCM N 129 139 (85%) (75%)

1) LiHMDS 2) HCl/H2O

Et2O

NH NH 2 HCl HN HCl HN 2 HCl N NH N NH H 2 Boc 2 Dioxane NH2 NH2 34 140 (49%) (72%)

Scheme 6.6: Amidine formation of dinitrile derivative 129 with prior protection of the indole NH with the Boc group. The ﬁnal compound DAPI 34 was obtained in a overall yield of 6.5%.

Due to NH4Cl formation HCl was added stoichiometric to convert LHMDS to hexamethyldisilazane which could be removed under vacuum. Subsequently added EtOH-HCl aﬀorded the amidine salt. Deprotection of the Boc protecting group yieled the target

177 6. Sensing via hydrogen bonding molecule DAPI 34 in an overall yield of 6.5%. With that we established a DAPI synthesis enabling us to access 22 mg of DAPI.1

6.2.2. DAPI analogue Synthesis

6.2.2.1. DAPI isomer

As we obtained two isomers in the Fischer indole synthesis step (Scheme 6.2) the same procedure was applied to anaolgue 133. Starting from the dibromo isomer, cyanation was carried out with following transformation of the nitrile into the HCl-salt to obtain 141 (Scheme 6.7).

H N 2 NH

Br O 131 Br See Scheme 6.3 HN NH2 Scheme 6.6 Polyphosphorsäure Br NH 2 HCl MeOH N H N NH H 2 Br 132 133 141 (24%)

Scheme 6.7: Synthesis overview of the Fisher-Indol approach for DAPI-isomer 141.

6.2.2.2. guanidine DAPI

Starting from 6-Aminoindole (142) the synthesis of a guanidine-substituted isomer 143 was attempted. This amendment in respect of the underlying concept of DAPI might lead to a higher binding affinity or different fluorescent shifts. The indole 142 was protected by adding Boc in anhydrous dichloromethane using DMAP as a catalyst. The obtained protected indole 144 was lithiated by using lithium diisopropylamide (LDA) in tetrahydrofuran at -20 °C under anhydrous conditions. Subsequent Stille coupling using N -Boc-4-iodoaniline was not successful. The reaction was carried out with Palladium-tetrakis(triphenylphosphine) (Pd(PPh3)4) and Pd(OAc)2 as palladium

1This work was supported by Paul Ebensperger during an internship

178 6.2. Results and Discussion catalysts and 1,4-diazabicyclo[2.2.2]octane (DABCO) as a base but the desired product could not be obtained (Scheme 6.8).

Boc2O, DMAP Sn Me3SnCl, LDA H N N BocHN N BocHN N 2 H DCM Boc THF Boc 142 144 145 (99%) (27%)

I Pd(PPh3)4/ Pd(OAc)2, DABCO, KF

NHBoc 1,4-Dioxane 146 HN NH NH 2 NH NHBoc N N H2N N BocHN H H Boc 143 147

Scheme 6.8: Attempted synthesis of guanidine-DAPI isomer 143.

6.2.2.3. Elongated structures

The idea was to extend the dye and to modify it with different binding sites to achieve a broader response to the phosphate derivatives. The elongation of the aromatic system might lead to a higher fluorescence intensity and more basic binding sites, such as guanidine, could lead to a higher binding affinity and fluorescent shifts upon binding. The sensor molecule 148 was designed as an elongated derivative of DAPI. Preparation of this compound is described in Scheme 6.9. 6-Bromoindole (149) was Boc protected and the obtained indole 150 was lithiated by using LDA in anhydrous tetrahydrofuran at -20 ° C. Susequently, the lithioindole intermediate was reacted with trimethyltin chloride yielding the indole stannane 151. This stannane intermediate was converted into the 2-iodoindole using molecular iodine at room temperature. A Suzuki coupling reaction between two equivalents of 4-Cyanophenylboronic acid (152) and 153 in the presence of

5 mol% Pd(PPh3)4 aﬀorded the dinitrile 154. By using lithium bis(trimehtylsilyl)amide the diamidine of 148 as a hydrochloride salt was obtained with an overall yield of 40%.

179 6. Sensing via hydrogen bonding

LDA, Sn Br N (Boc)2O, DMAP Br N Me SnCl, Br N H 3 DCM O THF O 149 O O 151 150 (87%) (quant.)

OH I2 N B THF OH 152 Pd(PPh3)4, Na CO N 2 3 I N Toluene Br N O O O N O 154 153 (55%) (84%)

1.) LiN(TMS)2 2.) 4 M HCl in Dioxane

Et2O

N NH H 2 HN *2 HCl

NH 2 148 (quant.)

Scheme 6.9: Synthesis overview of the DAPI-derivative 148 with an overall yield of 40%.

As discussed above, guanidine moieties bear advantages when it comes to phosphate binding. For this reason we attempted to synthesize a molecule containing this binding unit (Scheme 6.10). Starting with the 2-iodoindole (153) a Suzuki coupling with two equivalents of 4-aminophenylboronic acid using 10 mol% Pd(PPh3)4 and Na2CO3 in toluene gave the Boc-protected compound 155. Deptrotection of this molecule aﬀorded the HCl-salt of the diamine 156.

180 6.2. Results and Discussion

OH BocHN B OH 157 Pd(PPh3)4, I Na2CO3 NHBoc Br N N O Toluene O O BocHN O 153 155 (63%)

4 M HCl Dioxane

NH2 N H 2 HCl H2N 156 (quant.)

Scheme 6.10: Synthesis overview of diamine 156.

Having this molecule in hand, the transformation of the amino group into a guanidine congener was attempted. In literature various ways to achieve this are reported.[28–34] These methods can be divided roughly into three categories: urea- or amidine-based reagents and Lewis-acid activated carbodiimides. Scheme 6.11 provides a brief overview of the guanidinylation reagents which were used in this work.

2 i) NR R2 = Boc 2 R HN X X = NTf, N N 158 159

1 R -NH2 Base

iii) NH 1 NH 1 ii) R -NH2 R -NH2 R1 N NH 2 H2N Cl H2N N Sc(OTf) H 3 161 162 160

Scheme 6.11: Concise overview of guanidinylation reagents used in this work forming the guanidine core structure: i) amidine-based reagents [29,35]; ii) lanthanide activated carbodiimide[36]; iii) Chloro- formamidine hydrochloride.[37]

Many methods, commonly used for the synthesis of substituted guanidines, reported in the literature, often perform well with electron-rich amines but it is a diﬀerent situation

181 6. Sensing via hydrogen bonding when it comes to electron-deficient aromatic amines. We choose the three guanidinylation reagents (Scheme 6.11) based on their ability to react with weakly nucleophilic amines. i) Diprotected triflylguanidine 158 is a bench stable crystalline substance, which is commercial available and which allows the preparation of protected guanidines with exceptional ease and efficiency.[35] In a typical reaction, a slight excess of amine is added to a solution of 158 and 1.0 eq. triethylamine in dichloromethane. A simple aqueous workup removes the byproducts and the crude product is usually obtained in high purity. Isothioureas are common used guanidinylation reagents but a drawback is the need of highly toxic mercury salts. To avoid the use of toxic metals pyrazole carboximidamide transfer reagents has been developed. Reagent 159 was used for the synthesis of arginine containing peptides.[38] ii) Treatment of free cyanamide with catalytical amounts of scandium triflate gives the activated carbodiimides which react readily with amines in water.[36] The advantage of this method is the ability to guanidinylate substrates which are not soluble in organic solvents. Additionally, this method selectively guanidinylates the anilin nitrogen in presence of an indole N1 nitrogen. iii) As mentioned above the use of isothiourea derivatives suffer from the need of toxic metals. Direct guanidinylation of electron deficient amines by using preformed 161 in acetonitrile bypasses this problem and provides a one-pot synthesis of guanidines from aromatic amines.[37]

182 6.2. Results and Discussion

CF3 O S O N BocN BocHN NHBoc NHBoc 158 NH NBoc N NEt3 H 156 BocHN N DCM, reflux H 163 (31%)

HCl (4 M in Dioxane)

HN NH2 NH 2 HCl NH N H

H2N N H 164 (quant.)

Scheme 6.12: Guanidinylation of diamine 156 by using guanidinylation reagent 158 and 1.0 eq. triethylamine in dichloromethane.

In our case method i) using diprotected triflylguanidine 158 was successfully applied to diamine 156 affording the Boc-protected 163. Deprotection of the Boc groups with HCl (4 M in dioxane) gave the guanidinylated DAPI-isomer 164 (Scheme 6.12). Besides this method ii) could be used for bisguanidinylation of the C6-position of an indole scaffold (see Figure 6.13). However, Method iii) gave no results.

6.2.2.4. Biguanide moiety

As a biguanide consists of two condensed guanides (Figure 6.6) this compound is also classiﬁed as a strong base (pKa > 10) and so exists in a protonated state at physiological pH. Biguanides are also capable of polyvalent anion binding through multiple-site hydrogen bondings and delocalized positive charge in aqueous solution.[39]

Figure 6.6.: Binding moieties used in this work

183 6. Sensing via hydrogen bonding

Method ii) was used to exchanged cyanamide with dicyanamide to achieve formation of a biguanidine moiety. Indeed, this approach worked as was proven in a test reaction with 6-Aminoindole 142 (Scheme 6.13).

NH NH NH N Sc(OTf)3 (10 mol%) N H N N H2N N H2N N N 2 H H H2O H H H 142 165 166 (91%)

Scheme 6.13: Test reaction for forming of biguanidine moiety.[36]

Treating 142 with dicyandiamide (165) under Sc(TOf)3 catalysis in refluxing water gave the pure product without any further need of purification. However, the attempt to apply this method to diamine 156 was not successful as only educt was isolated from the reaction mixture. It might be possible that the amines were not sufficiently converted to the free base, thus not being nucleophilic enough to proceed the reaction. Nevertheless, for the reasons given above it is an interesting binding moiety and worth to be kept in mind.

6.2.3. C-3 modiﬁed indole

The electron-rich indole core has its highest reactivity towards electrophilic substitution at the C3 position given its enamine substructure..[40]

Figure 6.7.: Numbering of the indole core.

In this section a possibility for further modiﬁcations on the DAPI scaﬀold is discussed. The so gained modularity provides opportunities for further sensor design.

184 6.2. Results and Discussion

6.2.3.1. Koshland modiﬁcation

2-hydroxy-5-nitrobenzyl bromide (167) was used 1964 by Koshland et al. for selective modiﬁcation of tryptophane residues in proteins.[41]

OH OH O2N

O2N NH NH 167 HN N NH HN H2O/acetone 2 N NH 2 H H NH 2 NH2 34 2 HCl 168 2 HCl (71%)

HCO2H, 10% Pd/C 50% TFA in H2O

NH OH OH H2N N N NH2 H2N H 160 NH Sc(OTf)3 NH HN HN N NH H O N NH H 2 2 H 2 NH NH 2 170 2 169 3 TFA (31%)

Scheme 6.14: Modiﬁcation of DAPI with Koshland reagent 167.

Today, this reagent is known as “Koshland’s reagent I“ and was already used for further derivatisation of the DAPI scaﬀold.[42,43] Reaction of the phenylindole core with 167 gave the C3 alkylated product 168. Further treatment with formic acid in presence of 10% Pd/C allowed the reduction of the nitro group to an amine 169 (Scheme 6.14). This compound 169 is a versatile building block in sensor design. Conversion of the amine into a guanidine moiety was tried with cyanamide 160. Workup for the poly cationic molecule is tedious and material is lost during washing steps. However, after workup educt was reisolated, thus up to now the transformation of the amine into a utilizable moiety could not be achieved.

185 6. Sensing via hydrogen bonding

6.2.4. Sensor evaluation

In the previous section 6.2.1 the syntheses of DAPI-derived chromophores were discussed. Diﬀerent congeners have been made accessible in the subcategories “Amine“, “Amidine“ and “Guanidine“ (Scheme 6.15).

Amine NH2 N H H2N 156

Amidine NH HN NH 2 HN N NH NH H 2 NH 2 N NH 34 H 2

NH 141

N NH H 2 HN 148 NH2

HN Guanidine NH2 NH NH N H

H2N N 164 H

Scheme 6.15: DAPI-derived sensors synthesised in this work. All compounds are tested in their HCl-salt form.

With this chemosensors in hand we evaluated their photophysical properties and their ability to act as an anion sensor based on their ﬂuorescence response in water.

Firstly we determined the excitation (λEx) and emission wavelength (λEm) at an appropriate concentration of the sensor. In the next step we searched for suitable buffer systems and as a last step investigated their fluorescence intensity/wavelength change upon addition of analytes of interest. It was mentioned above (section 6.1.1) that the studies of Saiardi et al. showed that besides polyP also InsP6 induces a fluorescence shift of DAPI. For their fluorescence intensity measurements they used 10 mM Tris/HCl (pH 7.4) to mimic cytosolic conditions and 150 mM NaClO4 due to its minimised interaction with cations like DAPI compared

186 6.2. Results and Discussion to chloride (Cl-).[9] Due to the fact that in the literature similar experiments were carried out, it was decided to use this buﬀer composition for our experiments.

6.2.4.1. Photophysical properties

Determination of the absorbance maxima of the ﬂuorophores gives information about the wavelength with which the speciﬁc dyes has to be exited to obtain the maximal emission intensity. In Figure 6.8a the stacked UV/Vis-spectra of 34 (43 µM), 141 (49 µM) and 148 (50 µM) obtained in water are shown. In Scheme 6.16 the tested amidine-sensors are shown.

HN NH2 NH NH NH N HN NH2 N NH2 H H N NH HN NH2 H 2

NH2 148 34 141

Scheme 6.16: Amidine sensors

(a) (b) Figure 6.8.: a) Absorbance spectra of 34 (43 µM), 141 (49 µM) and 148 (50 µM) obtained in water from λ=300-700 nm. b) Changes in the 148 (50 µM) absorption sepctrum in the presence of 1.0 eq. of polyP, InsP6 and DNA in water from λ=300-700 nm.

From these spectra λmax can be extracted for each compound and with that the excitation wavelength for recording the ﬂuorescence intensity spectra is known. In Figure 6.8b changes in the absorbance spectra upon anion binding can be seen by the example of 148. The anions cause a small red shift accompanied by an increase of the signal near 350 nm. The small redshift indicates that the sensor can be excited in this region only

187 6. Sensing via hydrogen bonding with an anion present. These effects could be observed in all absorbance spectra of the investigated chemosensors. To get an idea about the sensors behaviour in different media we measured the fluorescence intensity from the sensor molecule itself in buffer (blank) and with added anion.

For consistent comparison λEx was set to 410 nm for all samples, following the protocol from Saiardi et al.[9]. The concentration of the working solution of the dye was 40 µM and the anion were added in a 1:1 ratio. Following buﬀer systems were used:

• Tris/HCl (10 mM, pH 7.6) NaClO4

• Tris/HCl (10 mM, pH 7.6)

• Triethylammonium acetate (pH 7)

• HEPES (10 mM, pH 7.3)

Initial measurements were done in water and the results are shown below in Figure 6.9. The blank measurements of the respective sensor already have a high emission intensity (see Figure 6.9, solid lines) which was also mentioned in Saiardis studies. DAPI (34) emits at approximately 550 nm even without adding of polyP. Whereas polyP induces only a small increase in intensity. InsP6 results in a heavy quenching of the signal (Figure 6.9 b)).

HN NH2 NH NH NH N HN NH2 N NH2 H H N NH HN NH2 H 2

NH2 148 34 141 HN NH2 NH NH N H

H2N N H 164

Scheme 6.17: Structures of the evaluated chemosensors.

188 6.2. Results and Discussion

(a) (b) Figure 6.9.: a) Superimposed emission spectra of evaluated sensors (40 µM, water) with added P12 (1.0 eq.) from 450 - 800 nm with λEx = 410 nm. b) Overlayed emission spectra of the evaluated sensors (40 µM, water) with added InsP6 (1.0 eq.) from 450 - 800 nm with λEx = 410 nm.

It is noteworthy that sensor 141 exhibits a small emission red shift with polyP and a larger blue shift with added InsP6. Sensor molecules 141 and 164 show no significant change in the emission intensity of fluorescence shift in water. As the binding moieties of the used sensor family are pH sensitive we investigated the emission intensity hereafter in buffered systems. In Tris/HCl (10 mM, pH 7.6) buffer the expected emission shift for DAPI accompanied by an intensity increase could be observed (Figure 6.10a, dotted black line). However, none of the other chemosensors showed any response to polyP or InsP6 either.

(a) (b) Figure 6.10.: a) Superimposed emission spectra of the evaluated sensors (40 µM, Tris/HCl 10 mM, pH 7.6) with added P12 (1.0 eq.) from λEm=450 - 800 nm with λEx = 410 nm. b) Superimposed emission spectra of the evaluated sensors (40 µM, Tris/HCl 10 mM, pH 7.6) with added InsP6 (1.0 eq.) from λEm=450 - 800 nm with λEx = 410 nm.

189 6. Sensing via hydrogen bonding

In Tris/HCl (10 mM, pH 7.6) NaClO4, DAPI showed the expected emission shift from

480 nm to 520 nm (showed in black in Figure 6.11a) upon addition of P12 while 148 and 164 remained broadly unchanged except for a small decrease in intensity (orange, green). Interestingly, elongated sensor 148 showed a red shifted emission maxima at approximately 530 nm. This maxima showed a small change with polyP to 580 nm (orange). Although in Tris/HCl DAPI was not aﬀected by inositol phosphate chemosensor

34 shows an increased emission with NaClO4 being a component of the buﬀer (Figure 6.11b, black). Indicating that Cl- is competing with the analytes for binding.

(a) (b) Figure 6.11.: a) Superimposed emission spectra of the evaluated sensors (40 µM, Tris/HCl 10 mM, pH 7.6, NaClO4) with added P12 (1.0 eq.) from λEm=450 - 800 nm with λEx = 410 nm. b) Superimposed emission spectra of the evaluated sensors (40 µM, Tris/HCl 10 mM, pH 7.6, NaClO4) with added InsP6 (1.0 eq.) from λEm=450 - 800 nm with λEx = 410 nm.

For all of the chemosensors no significant changes in the fluorescence profiles could be observed for triethylammonium acetate (pH 7.0), HEPES (10 mM, pH 7.3) and HCl 0.5 M.

6.3. Summary & Outlook

In this chapter a family of DAPI-based chemosensors was designed, synthesized and evaluated. Few syntheses of DAPI are reported in literature. Within this work we could establish a route for synthesising this dye in a ﬁve step sequence with an overall yield of 6.5% (see section 6.2.1). The core was obtained via an Fischer-indole synthesis and

190 6.3. Summary & Outlook subsequent cyanation gave the dinitrile compound. The nitrile as transformed into the amidine by using the LHMDS-method and so bypassing the need of dry HCl-gas.

The preparation of elongated sensor molecules was attempted to achieve a more po- tent chemosensor in respect to enhanced fluorescence intensity due to the potentially elongated π-system. However, the biaryl might possess a twisted state, thus being out of conjugation. These molecules were synthesised with amidine and guanidine binding sites. Guanidylation was accomplished by using diprotected triflylguanidine (Scheme 6.12). However, a significant increase of emission intensity upon an analyte binding event was not observed with the guanylated chemosensors.

Besides the preparation of sensor molecules, some possible modifications for further sensor development were also presented. The biguanide moiety can be implemented into a molecule by using an lanthanide catalysed method and the usage of the so-called “Koshland reagent I“ gave C3 modified indole scaffolds.

Inspired from the work of Saiardi et al. we evaluated the photophysical properties of the sensors summarized in Figure 6.15 in respect to polyP and InsP6. Absorbance measurements showed that the sensor molecules were interacting with the analytes (see Figure 6.8b). Emission intensity measurements in diﬀerent buﬀer systems with added analytes (P12 and InsP6) showed that in water the emission intensities were already high.

However, sensor 141 showed interesting eﬀects with P12 as addition of the anion results in a emission shift from 513 nm to 525 nm and InsP6 results in a blue shift from 513 nm to 470nm (see Figure 6.12a).

In Tris/HCl (pH 7.6) no effect on the emission profile with InsP6 or P12 on any sensor could be observed. Solely the expected emission maxima shift with 34 and P12 was detected. Although the only difference between Figure 6.10 and 6.11 is the added perchlorate in the buffer the results for 34 with InsP6 are significantly different in the latter. The emission maxima of the blank can be found at 520 nm and upon addition of

InsP6 the signal increases and is red shifted to 560 nm whereas P12 leads to an decreasing signal and blue shift to 460 nm (Figure 6.12b).

191 6. Sensing via hydrogen bonding

(a) (b) Figure 6.12.: a) Eﬀects of 1.0 eq. of added analyte (P12 and InsP6) on 141 (40 µM) in water. b) Eﬀects of 1.0 eq. analytes (P12 and InsP6) on 34 (40 µM) in Tris/HCl (10 mM, pH 7.6, NaClO4).

In conclusion, the herein synthesised and evaluated chemosensors show a utilizable emission response to polyP and inositol phosphates. DAPI 34 as well as sensor 141 shows a ﬂuorescence response to InsP6 and PolyP. The high water solubility is advantageous because it makes these dyes feasible for applications, such as PCSR for IC, staining Gels or for plate reader assays in 96-well plates. Convenient syntheses strategies for amidine and guanidinium-sensors were established during this work which could be applied for further sensor design in future.

192 6.4. Experimental

6.4. Experimental

6.4.1. General remarks

Reactions were carried out in ﬂame-dried glassware under argon atmosphere, unless noted otherwise. Reaction control was performed via TLC.

Reagents were purchased from commercial suppliers and used without further puriﬁ- cation.

Thin layer chromatography wa performed on Merck silica gel 60 F254 plates (0.25 mm layer thickness, with ﬂuorescence indicator). TLC was analyzed by UV (λ =,254 nm), ﬂuorescence (λ =,368 nm).

193 6. Sensing via hydrogen bonding

Chemistry at the University of Freibug. All signals are referenced to an internal solvent signal standard (CDCl3, δ =77.1 ppm; MeOD4, δ =49.0 ppm; DMSO-d6, δ =39.52 ppm).

Analytical HPLC was performed on an Agilent 1100 (binary pump, autosampler, DAD).

Preparative HPLC was performed on a Knauer Azura Semipreparative HPLC (binary pump, ASM 2.1L, DAD).

MPLC was performed on an Interchim PF430.

UV/Vis-Absorption Sepctra were recorded on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Transparent).

Fluorescence Spectra were recorded on a Tecan Spark 10M Microplate Topreader using 96-well plates (Thermo Fisher Scientiﬁc-Nunclon 96 Flat Black).

194 6.4. Experimental

6.4.2. Synthesis

General procedure for the amidine synthesis

To a solution of 1.1.1.3.3.3-hexamethyldisilazane (2.5 eq./Nitrile) in diethyl ether (50 mM) at 0 ° C was added n-BuLi (3.0 eq.). The mixture was stirred at RT until white precipitate formed. The suspension was cooled to 0 ° C and the nitrile dissolved in diethyl ether was added and the mixture was stirred at RT until complete consumption of the starting material was evidenced by TLC. The mixture was cooled to 0 ° C and quenched with a solution of 4.0 M HCl in Dioxane (1 eq./LHMDS). The solvent was evaporated and the residue was suspended in EtOH, followed by addition of EtOH-HCl until the mixture became acidic and stirred at RT for 2 h to give precipitate. The mixture was concentrated to half, precipitated with ether and centrifuged. The solid was suspended in H2O N (˜ 3mL/100 mg) and 6 NaOH was added until the pH of the mixture became >9. The precipitate was centrifuged and dried. The solid was suspended in a small amount of EtOH and treated with EtOH-HCl. Precipitation with ether and centrifugation gave the amidine-HCl salt.

6-Bromo-2-(4-bromophenyl)-1H -indole (130)

Br Br N H

A mixture of 4-Bromacetophenone (527 mg, 2.64 mmol, 1.00 eq.) and 171 (557 mg, 2.97 mmol, 1.12 eq.) was heated to 100 ° C for 10 min. Then MeOH (3 mL) was added and the solution refluxed for 1 h. After cooling down to RT the precipitated product could be isolated by filtration as a yellow greenish powder. No analytical data considering this molecule were collected because it was used in the next step without further purification. Polyphosphoric acid (8.00 g, 8 x the weight) was added at 0 ° C. After heat- ing the solution to 120 - 130 ° C for 20 min it was stirred at this temperature for 1 h. Then, water (50 mL) was added to the hot mixture and everything was poured into water (100 mL). After a few minutes at RT the precipitated product could be separated

195 6. Sensing via hydrogen bonding by ﬁltration. Separation of both isomers 130 and 133 was achieved by ﬂash column chromatography (cyclohexane:EE 10:1) and the product 130 (316 mg, 0.90 mmol, 34 %) was received as a brownish solid.

Rf (SiO2, Hexane/EE 10:1 (vol/vol)) 0.6. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.26 (br.s, 1H), 7.63 – 7.44 (m, 6H), 7.12 (dd, J = 8.4, 1.7 Hz, 1H), 6.78 (d, J = 1.7 Hz, 1H).

13 C-NMR (126 MHz, CDCl3, δ/ppm): 137.8, 137.5, 132.4 (2C), 130.9, 128.2, 126.7 (2C), 124.0, 122.1, 122.1, 116.2, 114.0, 100.7. HRMS (ESI) m/z: calcd.: 349.9008, found: 349.9009 [M-H+]−.

2-(4-Cyanophenyl)-1H -indole-6-carbonitrile (129)

N N N H

A flask was charged with NaCN (152 mg, 3.10 mmol, 2.4 eq.), CuI (50 mg, 0.26 mmol, 20 mol%), aryl bromide (450 mg, 1.29 mmol. 1.0 eq.), and KI (86 mg, 0.52 mmol, 40 mol%). Anhydrous toluene (2 mL) and N,N’-dimethylethylenediamine (277 µL, 2.58 mmol, 2.0 eq.) were added under argon. The reaction mixture was stirred at 120 ° C for 24 h. The resulting suspension was allowed to reach RT, diluted with 30% aq. ammonia (6 mL), and extracted with ethyl acetate (15 mL). The combined organic phases were dried over Na2SO4, concentrated and the residue was purified by flash chromatography (pentane/EE 5:1 to 3:1) to provide the product as a yellow solid (266 mg, 1.09 mmol, 85%)

Rf (SiO2, Pentane/EE 5:1 (vol/vol)) 0.3. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 12.34 (s, 1H), 8.12 (d,J = 8.4 Hz, 2H), 7.98 (d,J = 8.4 Hz, 2H), 7.89 (s, 1H) 7.77 (d,J = 8.25 Hz, 1H), 7.37 (dd, J = 8.26, 1.38 Hz, 1H), 7.30 (d,J = 2.03 Hz, 1H).

13 C-NMR (101 MHz, DMSO-d6, δ/ppm): 139.6, 136.2, 135.3, 132.9 (2C), 131.4, 126.1 (2C), 122.3, 121.6, 120.3, 118.7, 116.3, 110.4, 103.7, 101.9. HRMS (ESI) m/z: calcd.: 242.0723, found: 242.0723 [M-H+]−.

196 6.4. Experimental

IR(neat, cm-1) 2200, 1580, 1350, 1133, 822, 612.

Benzimidamide · HCl (138)

HN NH 2 HCl

Benzonitrile (247 µL, 2.4 mmol, 1.0 eq.) was treated according to the general procedure for amidine synthesis to give the HCl-salt as a white solid (146 mg, 1.2 mmol, 50%).

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 8.79 (s, 4H), 7.98 (s, 2H), 7.80 – 7.67 (m, 1H), 7.62 - 7.54 (m, 2H).

13 C-NMR (126 MHz, DMSO-d6, δ/ppm): 165.8, 133.71, 128.8, 128.1 HRMS (ESI) m/z: calcd.: 121.0761, found: 121.0761 [M+H+]+.

1-(tert-Butoxycarbonyl)-6-cyano-2-(4-cyanophenyl)-1H -indole (139)

N N N Boc

4-(dimethylamino)pyridine (DMAP) (4.0 mg, 0.033 mmol, 0.08 eq.) was added at 0 ° C in one portion to a stirred mixture of 2-(4-cyanophenyl)-1H -indole-6-carbonitrile (129)

(100 mg, 0.41 mmol, 1.0 eq.) suspended in dichloromethane (1.5 mL) and Boc2O (119 mg, 0.54 mmol, 1.3 eq.). After 2 h the reaction mixture was quenched with ice water (2 mL). The organic layer was extracted with dichloromethane (5 mL). The combined organic phase was washed with 1 M HCl (5 mL), brine (5 mL) and H2O (5 mL), respectively, and dried over Na2SO4. The solution was ﬁltered, passed through a bed of silica gel and concentrated under reduced pressure to give the product as a yellow solid (106 mg, 0.308 mmol, 75%).

Rf (SiO2, Pentane/EE 5:1 (vol/vol)) 0.3.

197 6. Sensing via hydrogen bonding

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 8.47 (d, J = 1.4 Hz, 1H), 7.96 (d, J = 8.3 Hz, 2H), 7.86 (d, J = 8.1 Hz, 1H), 7.75 (d, J = 8.3 Hz, 2H), 7.70 (dd, J = 8.1, 1.5 Hz, 1H), 7.02 (s, 1H), 1.30 (s, 9H).

DAPI · 2HCl (34)

NH HN N NH H 2 NH 2 2 HCl

The nitrile 139 (192 mg, 0.56 mmol) was treated according to the general procedure for amidine synthesis to give the HCl-salt as a green solid (61 mg, 0.17 mmol, 31%).

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 12.63 (s, 1H), 9.45 (s, 2H), 9.32 (s, 2H), 9.18 (s, 2H), 8.98 (s, 2H), 8.24 (d,J = 8.2 Hz, 2H), 7.99 (d, J = 8.4 Hz, 3H), 7.79 (d,J = 8.3 Hz, 1H), 7.47 (dd, J = 8.4, 1.6 Hz, 1H), 7.32 (d, J = 1.9 Hz, 1H). HRMS (ESI) m/z: calcd.: 278.1400, found: 278.1404 [M+H+]+.

4-Bromo-2-(4-bromophenyl)-1H -indole (133)

Br N H

Compound 133 was prepared using the procedure as for 130 on page 6.4.2. The product 133 (301 mg, 0.862 mmol, 32%) was obtained as a brownish solid.

Rf (SiO2, Hexane/EE 10:1 (vol/vol)) 0.4. 1 H-NMR (500 MHz, DMSO-d6, δ/ppm): 11.97 (s, 1H), 7.87 (d, J = 8.7 Hz, 2H), 7.67 (d, J = 8.6 Hz, 2H), 7.42 (dt, J = 8.1, 0.8 Hz, 1H), 7.23 (dd, J = 7.5, 0.8 Hz, 1H), 7.05 (dd, J = 8.1, 7.6 Hz, 1H), 6.92 (dd, J = 2.3, 0.9 Hz, 1H).

13 C-NMR (126 MHz, DMSO-d6, δ/ppm): 137.5, 137.4, 131.9 (2C), 130.7, 129.0, 127.2 (2C), 123.0, 122.2, 121.0, 113.2, 111.0, 99.0.

198 6.4. Experimental

2-(4-Cyanophenyl)-1H -indole-4-carbonitrile (172)

N N H

Compound 172 was prepared using the procedure as for 129 and the product was obtained as a yellow solid (266 mg, 1.09 mmol, 85%).

Rf (SiO2, Hexane/EE 10:1 (vol/vol)) 0.4. 1 H-NMR (500 MHz, DMSO-d6, δ/ppm): 12.38 (s, 1H), 8.17 (d, J = 8.4 Hz, 2H), 7.96 (d, J = 8.5 Hz, 2H), 7.78 (d, J = 8.2 Hz, 1H), 7.58 (d, J = 7.2 Hz, 1H), 7.36 – 7.32 (m, 1H), 7.31 (d, J = 7.9 Hz, 1H).

13 C-NMR (126 MHz, DMSO-d6, δ/ppm): 138.7, 137.2, 135.3, 133.0 (2C), 129.1, 126.2 (2C), 125.5, 122.5, 118.7, 118.5, 116.9, 110.4, 101.5, 99.5. 2-(4-Carbamimidoylphenyl)-1H -indole-4-carboximidamide (141)

HN NH2

N NH H 2

Compound 141 was prepared according the procedure as for 34 and the product was obtained as a yellow solid (58 mg, 0.16 mmol, 87%).

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 12.45 (s, 1H), 9.40 (s, 2H), 9.31 (s, 2H), 9.14 (s, 2H), 9.10 (s, 2H), 8.21 (d, J = 8.4 Hz, 2H), 7.98 (d, J = 8.5 Hz, 2H), 7.80 (d, J = 7.9 Hz, 1H), 7.56 – 7.28 (m, 3H). 6-Bromo-1-(tert-butoxycarbonyl)-1H -indole (150)

Br N Boc

199 6. Sensing via hydrogen bonding

Compound 150 was prepared according the procedure as for 139 on page 6.4.2 and the product was obtained as a white solid (1.23 g, 4.16 mmol, 82%).

Analytical data were in agreement with literature.[44]

Rf (SiO2, Pentane/EE 40:1 (vol/vol)) 0.7. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 8.22 (s, 1H), 7.69 (d, J = 3.7 Hz, 1H), 7.60 (d, J = 8.3 Hz, 1H), 7.41 (d, J = 8.4, 1.8 Hz, 1H), 6.73 (d, J = 3.7 Hz, 1H) 1.63 (s, 9H).

6-Bromo-1-(tert-butoxycarbonyl)-2-(trimethylstannyl)-1H -indole (151)

Sn Br N Boc

To a solution, cooled to -40 ° C, of 6-Bromo-1-(tert-butoxycarbonyl)-1H -indole (978 mg,

3.30 mmol, 1.0 eq.) and Me3SnCl (789 mg, 3.96 mmol, 1.2 eq.) in anhydrous THF (15 mL) was slowly added freshly prepared 2.0 M LDA in anhydrous THF (2 mL) while keeping the temperature below -20 ° C. The reaction mixture was warmed up to RT and stirred for 23 h. After quenching the reaction with water (10 mL), the solvent was removed under reduced pressure and the residue was extracted with Et2O (15 mL). The organic phase was washed with 10% NaF solution (3 mL) and water (20 mL), dried over

Na2SO4 and solvent removed under reduced pressure to yield the product as a brown solid (1.35 g, 2.94 mmol, 89%).

Analytical data were in agreement with literature.[44]

Rf (SiO2, Pentane/EE 40:1 (vol/vol)) 0.9. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 8.11 - 8.05 (m, 1H), 7.53 (d, J = 8.4 Hz, 1H), 7.37 (dd, J = 8.3, 1.7 Hz, 1H), 6.81 (d, J = 0.9 Hz, 1H) 1.67 (s, 9H), 0.29 (s, 9H).

6-Bromo-1-(tert-butoxycarbonyl)-2-(iodo)-1H -indole (153)

I Br N Boc

200 6.4. Experimental

A molecular iodine (2.09 g, 8.22 mmol) solution in anhydrous THF (10 mL) was added slowly to a solution of the indole stannane 151 (1.88 g, 4.11 mmol, 1.0 eq.) in anhydrous THF (10 mL). The mixture was stirred at RT for 19 h. The solvent was removed under reduced pressure, the residue dissolved in ethyl acetate (15 mL); washed with sodium thiosulfate solution (10%) to remove any excess of iodine, then with water, dried over

Na2SO4 and the solvent was removed under reduced pressure to yield the product as a yellow oil (1.17 g, 2.78 mmol, 67%).

Analytical data were in agreement with literature.[17]

Rf (SiO2, Pentane/EE 40:1 (vol/vol)) 0.7. 1 H-NMR (300 MHz, CDCl3, δ/ppm): 8.32 - 8.31 (m, 1H), 7.32 - 7.30 (m, 2H), 6.94 (d, J = 0.7 Hz, 1H), 1.72 (s, 9H).

4, 4’-(1H -indole-2, 6-diyl) dibenzonitrile (173)

N N H

An aqueous solution of Na2CO3 (3 mL, 2M) and 4-Cyanophenylboronic acid (351 mg, 2.39 mmol, 2.02 eq.) were added to a stirred solution of 153 (500 mg, 1.18 mmol, 1.0 eq.), in DMF (9 mL) and Pd(PPh3)4 (136 mg, 0.118 mmol, 10 mol%) was added. The vig- orously stirred mixture was warmed up to 130 ° C over night. After evaporation of the solvent under reduced pressure, the solid was partitioned between ethyl acetate (15 mL), then washing with water, passed through celite to remove the catalyst, dried over Na2SO4 and evaporated. Puriﬁcation was achieved by ﬂash column chromatography (pentane/EtOAc 10:1) and the product was obtained as a yellow solid (86 mg, 0.26 mmol, 23%).

Rf (SiO2, Pentane/EtOAc 10:1 (vol/vol)) 0.3.

201 6. Sensing via hydrogen bonding

1 H-NMR (500 MHz, DMSO-d6, δ/ppm): 11.97 (s, 1H), 8.12 – 8.07 (m, 2H), 7.99 – 7.93 (m, 2H), 7.92 (s, 4H), 7.75 – 7.69 (m, 2H), 7.44 (dd, J = 8.3, 1.7 Hz, 1H), 7.21 (d, J = 1.9 Hz, 1H).

13 C-NMR (126 MHz, DMSO-d6, δ/ppm): 145.8, 138.2, 137.3, 136.1, 132.9 (2C), 132.8 (2C), 128.8, 127.5 (2C), 125.6 (2C), 121.4, 119.4, 119.3, 119.0, 118.8, 110.0, 109.5, 109.1, 101.6. HRMS (ESI) m/z: calcd.: 318.1036, found: 318.1036 [M-H+]−. tert-butyl 2,6-bis(4-cyanophenyl)-1H -indole-1-carboxylate (174)

N N Boc

Compound 174 was prepared using the procedure as for 139 on page 6.4.2 and the product was obtained as a yellow solid (84 mg, 0.2 mmol, 74%).

The product was used in the next step without further puriﬁcation

4,4’-(1H -indole-2,6-diyl)dibenzimidamide · 2 HCl (148)

N NH H 2 HN 2 HCl

NH 2

The nitrile compound 174 (84 mg, 0.2 mmol, 1.0 eq.) was treated according to the general procedure for amidine synthesis to give the HCl-salt as a yellow solid (80 mg, 0.18 mmol, 94%).

Analytical data were in agreement with literature.[17]

1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 12.17 (s, 1H), 9.40 (s, 4H) 9.12 (s, 4H) 8.17 (d, J = 8.4 Hz 2H), 8.02 - 7.91 (m, 6H), 7.79 (s, 1H), 7.73 (d, J = 8.4 Hz 1H), 7.47 (dd, J = 8.3, 1.6 Hz, 1H), 7.24 (s, 1H).

202 6.4. Experimental

4,4’-(1H -indole-2,6-diyl)dianiline · 2 HCl (156)

NH 2 N H 2 HCl

H2N

Compound (155) (960 mg, 1.60 mmol, 1.0 eq.) was stirred in HCl (4 mL, 16 mmol, 4 M in dioxane) at RT for 6 h. After complete consumption of the starting material shown by HPLC, the solvent was removed to yield the product as red solid (428 mg, 1.15 mmol, 72%)

1 H-NMR (500 MHz, DMSO-d6, δ/ppm): 11.94 (s, 1H), 10.38 (br, 6H), 8.01 (d, J = 8.6 Hz, 2H), 7.79 (d, J = 8.5 Hz, 2H), 7.66 (s, 1H), 7.63 (d, J = 8.3 Hz, 1H), 7.48 (dd, J = 12.5, 8.6 Hz, 4H), 7.33 (dd, J = 8.3, 1.7 Hz, 1H), 6.97 (d, J = 2.1 Hz, 1H).

13 C-NMR (126 MHz, DMSO-d6, δ/ppm): 141.2, 137.8, 137.7, 132.9, 131.6, 131.3, 130.4, 129.7, 128.3, 127.9, 127.7, 126.2, 123.8, 123.5, 123.5, 123.3, 120.7, 119.0, 109.4, 99.2. HRMS (ESI) m/z: calcd.: 300.1496, found: 300.1497 [M+H+]+.

HPLC (C18, MeCN/H2O, 1:99 to 99:1 in 20 min) rt: 10.1 min.

1-(tert-Butoxycarbonyl)-2,6-bis(4-N,N’-Bis-(tert-Butyloxycarbonyl)phenyl) -1H -indole (163)

BocN NHBoc NH NBoc N H BocHN N H

The dihydrochloride salt of 156 (100 mg, 0.268 mmol, 1.0 eq.) was added to a solution of N,N’-di-Boc-N”-triﬂylguanidine (158) (189 mg, 0.48 mmol, 1.8 eq.) and triethylamine (74 µL, 0.537 mmol, 2.0 eq.) in dichloromethane (2 mL). The mixture was reﬂuxed for 24 h until the starting material was consumed as evidenced by TLC. The mixture was diluted with dichloromethane (3 mL) and washed with 2M sodium bisulfate (2x),

203 6. Sensing via hydrogen bonding

saturated sodium bicarbonate (2x), and brine (1x). After drying with Na2SO4 and filtering the solvent was evaporated under reduced pressure. Purification was achieved by flash column chromatography (pentane/EtOAc 4:1) and the product was obtained as a yellow oil (66 mg, 0.084 mmol, 31%).

Rf (SiO2, Pentane/EtOAc 4:1 (vol/vol)) 0.3. 1 H-NMR (300 MHz, DMSO-d6, δ/ppm): 11.59 (s, 1H), 11.42 (s, 1H), 11.41 (s, 1H), 10.10 (s, 1H), 10.07 (s, 1H), 7.87 (d, J = 8.6 Hz 2H), 7.74 - 7.53 (m, 8H), 7.33 (dd, J = 8.3, 1.6 Hz, 1H), 6.91 (d, J = 2.0 Hz, 1H), 1.53 (s, 18H), 1.43 (s, 18H). HRMS (ESI) m/z: calcd.: 884.4553, found: 884.4566 [M+H+]+. 1-(tert-Butoxycarbonyl)-2,6-bis(4-N,N’-Bis-(tert-Butyloxycarbonyl)phenyl)-1H - indole · 2 HCl (164)

NH 2 NH NH N H 2 HCl

H2N N H

163 (55 mg, 0.76 mmol, 1.0 eq.) was stirred in HCl (0.382 mL, 1.53 mmol, 4 M in dioxane) at RT for 6 h. The solvent was removed, the crude dissolved in water (4 mL) and NaOH (6 N) added until pH > 9. The precipitate was ﬁltered and redissolved in EtOH. EtOH- HCl was added until pH < 3 and the precipitate was ﬁltered to yield the product as green solid (25 mg, 0.54 mmol, 72%)

1 H-NMR (400 MHz, DMSO-d6, δ/ppm): 7.97 (d, J = 8.7 Hz, 1H), 7.81 (d, J = 8.6 Hz, 1H), 7.75 (d, J = 8.6 Hz, 1H), 7.69 – 7.57 (m, 4H), 7.40 – 7.30 (m, 4H), 7.26 (d, J = 8.6 Hz, 1H), 7.07 (s, 1H). HRMS (ESI) m/z: calcd.: 384.1932, found: 384.1934 [M+H+]+. UV λ = 292/404 nm Fluorescence λ = 530 nm

204 6.4. Experimental

2-(4-Amidinophenyl)-3-(2-hydroxy-5-nitro) benzyl-6-amidinoindole · 2 HCl (168)

OH O2N

NH HN N NH H 2 NH 2 2 HCl

DAPI dihydrochloride (350 mg, 1.00 mmol, 1.0 eq.) was dissolved in water (15 mL) and acetone (10 mL). A solution of Koshland’s reagent 167 (278 mg, 1.2 mmol, 1.2 eq.) in acetone (9 mL) was added and the mixture was stirred at RT. After 22 h, a second portion of 167 (47 mg, 0.202 mmol, 0.2 eq.) was added and the reaction mixture was stirred for another 22 h. Then, the solvents were evaporated and the residue dissolved in methanol. The mixture was evaporated again, and the resulting yellow precipitate suspended in chloroform and stirred overnight. The solid was ﬁltered and washed twice with chloroform and once with water to give the product as a yellow solid (305 mg, 0.711 mmol, 71%)

Analytical data were in agreement with literature.[43]

Rf (SiO2, Pentane/EtOAc 1:1 (vol/vol)) 0.4. 1 H-NMR (300 MHz, Methanol-d4, δ/ppm): 8.07 (s, 1H), 8.02 (dd, J = 8.9,2.9 Hz, 1H), 7.96 (d, J = 8.5 Hz, 2H), 7.88 (d, J = 8.5 Hz, 2H), 7.67 (d, J = 8.4 Hz, 1H), 7.59 (d, J = 2.8 Hz, 1H), 7.49 (dd, J = 8.4,1.7 Hz, 1H), 6.99 (d, J = 8.9 Hz, 1H), 4.36 (s, 2H). 3-(5-amino-2-hydroxybenzyl)-2-(4-carbamimidoylphenyl)-1H - indole-6-carboximidamide · 3 TFA (169)

OH H2N

NH HN N NH H 2 NH 2 3 TFA

205 6. Sensing via hydrogen bonding

Product 168 (200 mg, 0.406 mmol, 1.0 eq.) was dissolved in 22 mL of formic acid. Then 10% Pd/C (857 mg, ) and 7.4 mL of a 1 : 1 mixture of trifluoroacetic acid and water were added successively. The reaction mixture was heated to 100 ° C for 18 h. It was filtered through celite, the residue was washed with formic acid, and the filtrate was evaporated to yield a yellow solid (121 mg, 0.63 mmol, 31%).

HRMS (ESI) m/z: calcd.: 399.1928, found: 399.1926 [M+H+]+.

206 6.4. Experimental

6.4.3. References

Bibliography Chapter Sensing via hydrogen bonding

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[3] D. H. Williamson, D. J. Fennell, Methods Cell Biol. 1975, 12, 335–351.

[4] R. A. Allan, J. J. Miller, Can. J. Microbiol. 1980, 26 (8), 912–920.

[5] J. Kapuściński, B. Skoczylas, Anal. Biochem. 1977, 83 (1), 252–257.

[6] J. Tijssen, H. W. Beekes, J. van Steveninck, Biochim. Biophys. Acta, Mol. Cell Res. 1982, 721 (4), 394–398.

[7] B. Kjeldstad, M. Heldal, H. Nissen, A. S. Bergan, K. Evjen, Can. J. Microbiol. 1991, 37 (7), 562–567.

[8] S. Omelon, J. Georgiou, Z. J. Henneman, L. M. Wise, B. Sukhu, T. Hunt, C. Wyn- nyckyj, D. Holmyard, R. Bielecki, M. D. Grynpas, PLoS ONE 2009, 4 (5), e5634.

[9] B. Kolozsvari, F. Parisi, A. Saiardi, Biochem. J. 2014, 460 (3), 377–385.

[10] H. J. Jessen, Phosphate Labeling and Sensing in Chemical Biology, Springer Inter- national Publishing, Cham, 2017.

[11] N. Busschaert, C. Caltagirone, W. van Rossom, P. A. Gale, Chem. Rev. 2015, 115 (15), 8038–8155.

[12] C. Trujillo, V. Previtali, I. Rozas, Theor. Chem. Acc. 2016, 135 (12), 1–12.

[13] R. Mogaki, P. K. Hashim, K. Okuro, T. Aida, Chem. Soc. Rev. 2017, 46 (21), 6480–6491.

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[14] B. Springs, P. Haake, Bioorg. Chem. 1977, 6 (2), 181–190.

[15] K. A. Schug, W. Lindner, Chem. Rev. 2005, 105 (1), 67–114.

[16] T. Ishikawa, Superbases for organic synthesis: Guanidines, amidines and phosp- hazenes and related organocatalysts, John Wiley & Sons, Chichester, UK, 2009.

[17] A. A. Farahat, A. Kumar, M. Say, T. Wenzler, R. Brun, A. Paul, W. D. Wilson, D. W. Boykin, Eur. J. Med. Chem. 2017, 128, 70–78.

[18] S. D. Kuduk, R. K. Chang, J. M.-C. Wai, C. N. Di Marco, V. Cofre, R. M. DiPardo, S. P. Cook, M. J. Cato, A. Jovanovska, M. O. Urban, M. Leitl, R. H. Spencer, S. A. Kane, G. D. Hartman, M. T. Bilodeau, Bioorg. Med. Chem. Lett. 2009, 19 (15), 4059–4063.

[19] G. P. Ellis, T. M. Romney-Alexander, Chem. Rev. 1987, 87 (4), 779–794.

[20] J. Zanon, A. Klapars, S. L. Buchwald, J. Am. Chem. Soc. 2003, 125 (10), 2890– 2891.

[21] A. Klapars, J. C. Antilla, X. Huang, S. L. Buchwald, J. Am. Chem. Soc. 2001, 123 (31), 7727–7729.

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[23] F. Eloy, R. Lenaers, Chem. Rev. 1962, 62 (2), 155–183.

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[27] J. Jänichen, M. Presiss, C. Alonso-alija, Straub, Alexander, Method for the production of substituted pyrimidine derivatives, , WIPO 2000, (WO2001017998A2).

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[31] K. S. Kim, L. Qian, Tetrahedron Lett. 1993, 34 (48), 7677–7680.

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210 CHAPTER 7

Assignment of the stereoselectivity of a new phytase

211 7. Assignment of the stereoselectivity of a new phytase

7.1. Background

Phosphate is a key nutrient of cells. It is essential for normal plant growth and survival and thus of importance in fertilizers in agriculture.[1] Phytate is the main phosphate storage molecule in plants and owes its name by the fact that it is of plant origin (Figure 7.1).[2] Furthermore, it is known to be involved in plant defence against fungal, viral and bacterial pathogens.[3] It contains myo-inositol, a member of the inositol family (Figure 7.1). The number of substituted phosphate groups on the inositol scaﬀold varies from

[4] one to six resulting in up to 63 possible isomers. The phosphate ester linkage in InsP6 (also called phytate) is inert and abiotic degradation is almost impossible. This makes the stored phosphate unutilized either by humans or farm animals. Additionally, metals such as calcium (Ca), zinc (Zn), copper (Cu), magnesium (Mg) and iron (Fe) form insoluble complexes with phytate thus limiting bioavailability of these minerals.

Figure 7.1.: Chemical structure of myo-inositol and the fully phosphorylated phytate.

7.1.1. Phytases

Certain enzymes (phytases - myo-inositol (1,2,3,4,5,6) hexakisphosphohydrolase) are able to release this stored phosphate by catalysing the hydrolysis of phytate in a step- wise manner and release inorganic phosphate (Figure 7.2). Presence of these enzymes was firstly reported 1907 in rice bran,[5] followed by diverse species including bacteria, fungi and plants. Phytases are classified by the position on the myo-inositol scaffold, at which the first dephosphorylation takes place. According to this, a 3-phytase will initiate dephosphorylation at the C3-position and has InsP5 [3-OH] as product, which

212 7.1. Background then can be further degraded depending on the enzyme. All types of phytases have been detected in nature, except for the 1-phytase[6,7], which is described in this work.[8]

Figure 7.2.: PDB structure of a phytase (blue) with complexed phytate (red). InsP6 is the main phosphate storage in nature. Phytases degrade progressively InsP6 to lower inositol phosphate derivatives and release orthophosphate. Picture taken from reference. [9]

7.1.2. Xanthomonas campestris pv. vesicatoria (Xcv)

Members of the genus Xanthomonas represent a group of phytopathogenic bacteria, which infect economically important crop plants causing substantial crop loss.[10] Xan- thomonas campestris pv. vesicatoria (Xcv) is the causative agent of bacterial spot disease on tomato and pepper plants.[10] The pathogenicity of Xcv depends, in many cases, on a conserved type III secretion system that translocates effector proteins directly into the plant cells.[11,12] These effector proteins are crucial for the virulence. The pepper and tomato pathogen encodes more than 30 type III effector (T3E) proteins, called ”Xops” (Xanthomonas outer proteins).[13] Once inside the plant cell, effector proteins interfere with the plant signalling and metabolic pathways to the benefit of the pathogen.

A hitherto unknown type III effector protein was identified and named ”XopH” by the group of professor Bonas. Preliminary studies indicated that XopH shows high specificity for phytate. With that XopH would be the first T3E protein with phytate degrading behaviour.

The central questions, which had to be answered, was: What is the speciﬁcity of XopH in this process and which is the absolute conﬁguration of the product (Figure 7.3)?

213 7. Assignment of the stereoselectivity of a new phytase

Figure 7.3.: The unknown type III effector protein XopH is able to degrade InsP6. Key questions: What is the specifity of XopH and which is the absolute configuration of the dephosphorylation product?

7.2. Results and Discussion

We answered this question by combining the results of PAGE, IC and 31P-NMR analysis. As discussed in chapter ”State of the Art - Analytics” - IC combined with PCSR enables UV detection of inositol phosphates. These compounds do not possess intrinsic UV activity, however, binding to a trivalent transition metal ion (Fe3+) results in a UV detectable complex.

PAGE analysis veriﬁed that the digest of XopH with InsP6 showed the same electrophoretic mobility as InsP5. This was unambiguously conﬁrmed by IC (Figure 7.4).

For the analysis InsP6, InsP5 (all six isomers) and a phosphate standard were measured using IC to determine the retention times under acidic conditions measured by UV after

3+ PCSR with Fe . Phytate elutes at 39 mins whereas all the lower InsP5 isomers elute in a range of 30 to 37 mins. Enantiomeric 1/3 and 4/6-OH InsP5 display the same retention times as enantiomers can not be separated under these conditions. Comparison of

XopH digest to the InsP5 isomers revealed that the identity had to be either one of the enantiomeric InsP5 [1-OH] or [3-OH] isomers or a mixture of both (Figure 7.4).

Additional experiments with the enzyme and all six available InsP5 isomers gave the result that every isomer, ecxept of InsP5 [1-OH], readily undergoes further degradation.

This was the initial hint that InsP5 [1-OH] represents the product of XopH-mediated

InsP6 hydrolysis.

214 7.2. Results and Discussion

Figure 7.4.: Stacked ion chromatograms of InsP6 (2 nmol), XopH (0.13 µg/µL) + InsP6 (2 nmol), phosphate standard (35 nmol) and all InsP5 isomers (2 nmol) eluted on CarboPac PA100: eluent 0.5 M HCl and water (18 MΩ∗cm); detection: post-column reaction with Fe3+, UV: 290 nm, injection volume 10 µL.

To unambiguously identify the isomeric identity, we developed a 31P-NMR using a chiral solvating agent based method which can be applied to samples without interference of high salt concentrations or buﬀers. As discussed in chapter State of the Art - Analytics, myo-inositol and phytate are meso-compounds which possess a mirror plane dissecting the 2- and 5-position. This results in one distinct signal for each phosphate in the 2 and 5 position, one signal for the 1/3 and one for the 4/6 positions due to internal symmetry. Breaking this symmetry will consequently result in 5 diﬀerent signals for every single phosphate. Our method is based on the addition of enantiopure LArgN. Guanidinium moieties are known to strongly interact with phosphates, thus LArgN acts here as a chiral solvating agent. Enantiodiscrimination is achieved by ionic interactions of the enantiopure amino acids guanidinium group and the inositol phosphate. This interaction generates diastereomers, which can be distinguished by 31P-NMR (Figure 7.5).

215 7. Assignment of the stereoselectivity of a new phytase

Figure 7.5.: Strategy for enantiodiscrimination using 31P-NMR: Achiral counterions, such as Na+ cre- ate identical chemical environments, whereas enantiopure counterions form diastereomeric pairs giving rise to diﬀerent signals in the 31P-NMR. Picture adapted from reference. [8]

31P-NMR shifts are matrix and pH sensitive. To ensure that this technique can be used to identify the isomers, we performed spiking experiments by mixing InsP5 [1-OH] with

InsP5 [3-OH] with diﬀerent ratios (1:1.5, 1:1, 1.5:1) in ammonium acetate buﬀer (pH 7.1).

Figure 7.6.: Upper trace: mixture of InsP5 [1-OH] (A) and InsP5 [3-OH] (B) without LArgN in ammonium acetate buﬀer (pH 7.1) gave no peak separation but integration 1:1:2:1 (from left to right). In presence of an excess of LArgN (approx. 100 fold) peak separation is observed in all three diﬀerent ratios (A/B: 1:1.5, 1:1, 1.5:1). Asterisk marks phosphate as an impurity.

216 7.2. Results and Discussion

Without added LArgN the chiral mixture showed four peaks with a ratio of 1:1:2:1 (Figure 7.6, upper trace), whereas addition of an excess (approx. 100 fold) of LArgN caused shifts in all signals. This led to distinct observable diﬀerences for four of the ﬁve resonances Figure 7.6 and proved 31P-NMR, in combination with an excess of LArgN as chiral solvating agent, a suitable technique for the enantiodiscrimination of InsP5 [1-OH] and [3-OH].

Analysis of the reaction product of the enzyme with InsP6 could be performed without prior puriﬁcation in ammonmium acetate buﬀer (pH 7.1). The enzyme digest was

31 spiked with commercial enantiopure InsP5 [1-OH]. The P-NMR showed the expected

five resonances for InsP5 (Figure 7.7a upper trace) and one additional peak that was identified as phosphate impurity by recording of a proton-coupled spectrum (Figure 7.7a lower trace). For phosphates bound to the inositol scaffold the proton-coupled

3- spectrum showed peak splitting (doublets) while the PO4 appeared as a singlet as no protons to couple to were available.

(a) (b) Figure 7.7.: a) Digest of InsP6 (600 nmol) by XopH in ammonium acetate buffer (pH 7.1), spiking with InsP5 [1-OH] (400 nmol) (ratio A/InsP5 [1-OH] 1.5:1) and addition of LArgN in excess (approx. 100 fold). No additional peaks can be seen in a proton-decoupled spectrum (upper trace). A proton- coupled spectrum identifies all resonances belonging to inositol-bound phosphates (A). b) Digest of InsP6 (600 nmol) by XopH (A) in ammonium acetate buffer (pH 7.1) in presence of LArgN (approx. 100 fold). Addition of InsP5 [3-OH] (B) in different ratios (A/B: 1.5:1, 1:1, 1:1.5) leads to the appearance of new resonances.

217 7. Assignment of the stereoselectivity of a new phytase

Equivalent spiking experiments were carried out with InsP5 [3-OH] in diﬀerent ratios in presence of an excess of LArgN. As with the InsP5 [1-OH/3-OH] mixture, a separation of four of the ﬁve signals could be observed (Figure 7.7b). This resolves any doubts about the identity of the build inositol phosphate isomer.

Combination of PAGE, IC and 31P-NMR with chiral solvating agent gave us the possibility to unambiguously reveal InsP5 [1-OH] as the reaction product of the newly identiﬁed T3E protein XopH from the phytopathogenic Xanthomonas campestris pv. vesicatoria. Hence, XopH is the ﬁrst described naturally occuring 1-phytase (Figure 7.8).[8]

Figure 7.8.: XopH is the ﬁrst described naturally occurring 1-phytase, initiating dephosphorylation of phytate at C-1 of the inositol scaﬀold. InsP5 [1-OH] is the reaction product of this enzyme.

How does degrading of phytate to InsP5 [1-OH] aﬀect the plant cells and the bacterial pathogen? The nutrition of the pathogen could be improved by the released phosphate or the plants hormone pathway could be disturbed by aﬀecting the availability of potential co-factors. Additionally, InsP6 has been suggested to be crucial for plants resistance against pathogens, thus XopH could interfere into this defense mechanism.

This discovered mechanism of the phytopathogen could be a commonly used strategy of host immune system avoidance for bacteria and could therefore provide a basis for development of new strategies to overcome or attenuate crop disease caused by bacterial pathogens.

Additionally the phytate activity can be advantageous for further biotechnological ma- nipulation of phytate levels. Due to the above mentioned high aﬃnity of phytate to minerals including zinc and iron, InsP6 is known as an antinutrient for humans and some animals. Thus, phytases are important to signiﬁcantly improve the bioavailability of phosphate and minerals.1

1This work was published [8]

218 7.2. Results and Discussion

7.2.1. References

Bibliography Chapter Assignment of the stereoselectivity of a new phytase

[1] P. J. White, E. J. Veneklaas, Plant Soil 2012, 357 (1-2), 1–8.

[2] U. Schlemmer, W. Frølich, R. M. Prieto, F. Grases, Mol. Nutr. Food Res. 2009, 53 Suppl 2, 330–375.

[3] A. M. Murphy, B. Otto, C. A. Brearley, J. P. Carr, D. E. Hanke, Plant J. 2008, 56 (4), 638–652.

[4] R. F. Irvine, M. J. Schell, Nat. Rev. Mol. Cell Biol. 2001, 2 (5), 327–338.

[5] U. Suzuki, K. Yoshimura, M. Takaishi, „About the enzyme “phytase”, which splits “anhydro-oxy-methylene diphosphoric acid”“.

[6] V. Kumar, A. K. Sinha, „General aspects of phytases“ in Enzymes in human and animal nutrition: Principles and perspectives / edited by Carlos Simões Nunes, Vikas Kumar (Hrsg.: C. S. Nunes, V. Kumar), Academic Press, Amsterdam, 2018, S. 53–72.

[7] L. Bohn, A. S. Meyer, S. K. Rasmussen, J. Zhejiang Univ. Sci. B 2008, 9 (3), 165–191.

[8] D. Blüher, D. Laha, S. Thieme, A. Hofer, L. Eschen-Lippold, A. Masch, G. Bal- cke, I. Pavlovic, O. Nagel, A. Schonsky, R. Hinkelmann, J. Wörner, N. Parvin, R. Greiner, S. Weber, A. Tissier, M. Schutkowski, J. Lee, H. Jessen, G. Schaaf, U. Bonas, Nat. Commun. 2017, 8 (1), 1–14.

[9] D. S. Goodsell, „PDB, zuletzt geprüft am 19.09.2020. http://pdb101.rcsb.org/motm/225“.

219 7. Assignment of the stereoselectivity of a new phytase

[10] Frederik Leyns, Marcel de Cleene, Jean-Guy Swings, Jozef de Ley, Bot. Rev. 1984, 305–355.

[11] D. Büttner, U. Bonas, EMBO J. 2002, 21 (20), 5313–5322.

[12] G. R. Cornelis, F. van Gijsegem, Annu. Rev. Microbiol 2000, 54, 735–774.

[13] F. F. White, N. Potnis, J. B. Jones, R. Koebnik, Mol. Plant Pathol. 2009, 10 (6), 749–766.

220 APPENDIX A

Analytical Data of Chapter 4

iii A. Analytical Data of Chapter 4

(9H -ﬂuoren-9-yl)methyl tert-butyl(6-(bis(pyridin-2-ylmethyl)amino)-6- oxohexane-1,5-diyl)(R)-dicarbamate (40)

N N O N O O N H HN O O

1 Figure A.1.: H-NMR (300 MHz, CDCl3) of compound 40

N N O N O O N H HN O O

13 Figure A.2.: C-NMR (101 MHz, CDCl3) of compound 40

iv N N O N O O N H HN O O

Figure A.3.: HRMS (ESI) m/z: calcd.: 650.3337, found: 650.3328 [M+H+]+ of compound 40. tert-butyl(R)-(5-amino-6-(bis(pyridin-2-ylmethyl)amino)-6-oxohexyl) carbamate (42)

N N O N O O N H H2N 42

1 Figure A.4.: H-NMR (300 MHz, CDCl3) of compound 42

v A. Analytical Data of Chapter 4

N N O N O O N H H2N 42

13 Figure A.5.: C-NMR (101 MHz, CDCl3) of compound 42

N N O N O O N H H2N 42

Figure A.6.: HRMS (ESI) m/z: calcd.: 428.2656, found: 428.2657 [M+H+]+ of compound 42.

vi tert-butyl(R)-(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl) acetamido)hexyl)carbamate (43)

N N O N O O N H HN O 43

1 Figure A.7.: H-NMR (300 MHz, CDCl3) of compound 43

N N O N O O N H HN O 43

13 Figure A.8.: C-NMR (101 MHz, CDCl3) of compound 43

vii A. Analytical Data of Chapter 4

N N O N O O N H HN O 43

Figure A.9.: HRMS (ESI) m/z: calcd.: 670.3388, found: 670.3381 [M+H+]+ of compound 43.

(R)-6-amino-2-(2-(pyren-1-yl)acetamido)-N,N -bis(pyridin-2-ylmethyl) hexanamide (44)

N N

N O NH2

HN O 44

1 Figure A.10.: H-NMR (300 MHz, CDCl3) of compound 44

viii N N

N O NH2

HN O 44

13 Figure A.11.: C-NMR (101 MHz, CDCl3) of compound 44

N N

N O NH2

HN O 44

Figure A.12.: HRMS (ESI) m/z: calcd.: 570.2864, found: 570.2859 [M+H+]+ of compound 44.

ix A. Analytical Data of Chapter 4

(R)-6-(pent-4-ynamido)-2-(2-(pyren-1-yl)acetamido)-N,N -bis(pyridin-2 -ylmethyl)hexanamide(46)

N N O N O N H HN O 46

1 Figure A.13.: H-NMR (300 MHz, CDCl3) of compound 46

Probe (47)

O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N 47

1 Figure A.14.: H-NMR (400 MHz, DMSO-d6) of compound 47

x O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N 47

13 Figure A.15.: C-NMR (101 MHz, DMSO-d6) of compound 47

O O NH N O N O NH O N HN O N N N N N HN O N HN O HN N O N N N N N N N N 47

Figure A.16.: HRMS (ESI) m/z: calcd.: 2277.1184, found: 2277.1185 [M+H+]+ of compound 47.

xi A. Analytical Data of Chapter 4

4,0 12000

3,5

10000

3,0

O O NH 8000

2,5 N O N O NH O N HN O N N N 2,0 N 6000 N HN O N HN HN 1,5 O N O 4000 N N N Absorbance [O.D.] Absorbance N N N N 1,0 N Fluorescence Intensity [RFU] Intensity Fluorescence 47 2000

0,5

0,0 0

300 400 500 600 700

Wavelength [nm]

Figure A.17.: Absorbance & Fluorescence Scan of compound 47 in water (HEPES 10 mM pH 7.4).

1-((1H -imidazol-1-yl)methyl)-3-(pyren-1-ylmethyl)-1H -imidazol-3-ium bromide (51)

N N N N

1 Figure A.18.: H-NMR (300 MHz, DMSO-d6) of compound 51

xii Br

N N N N

Figure A.19.: HRMS (ESI) m/z: calcd.: 363.1604, found: 363.1602 [M+H+]+ of compound 51.

N -(4-iodobenzyl)-1-(pyridin-2-yl)-N -(pyridin-2-ylmethyl)methanamine (84)

N N N 84

1 Figure A.20.: H-NMR (300 MHz, CDCl3) of compound 84

xiii A. Analytical Data of Chapter 4 tert-butyl (2-(bis(pyridin-2-ylmethyl)amino)-2-oxoethyl)carbamate (64)

O H O N N O N N

1 Figure A.21.: H-NMR (300 MHz, CDCl3) of compound 64

O H O N N O N N

Figure A.22.: HRMS (ESI) m/z: calcd.: 357.1921, found: 357.1921 [M+H+]+ of compound 64.

xiv 2-amino-N,N -bis(pyridin-2-ylmethyl)acetamide (65)

O H N 2 N N N 65

1 Figure A.23.: H-NMR (300 MHz, CDCl3) of compound 65

2-((6-bromopyren-1-yl)amino)-N,N -bis(pyridin-2-ylmethyl)acetamide (67)

N N N H O N 67

1 Figure A.24.: H-NMR (300 MHz, CDCl3) of compound 67

xv A. Analytical Data of Chapter 4

N N N H O N 67

Figure A.25.: HRMS (ESI) m/z: calcd.: 535.1128, found: 535.1130 [M+H+]+ of compound 67.

N1,N5 -bis(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl) acetamido)hexyl)glutaramide (70)

O O HN NH

N N N HN N NH O O O N N O

1 Figure A.26.: H-NMR (400 MHz, CDCl3) of compound 70

xvi O O HN NH

N N N HN N NH O O O N N O

13 Figure A.27.: C-NMR (101 MHz, CDCl3) of compound 70

O O HN NH

N N N HN N NH O O O N N O

Figure A.28.: HRMS (ESI) m/z: calcd.: 1235.5866, found: 1235.5852 [M+H+]+ of compound 70.

xvii A. Analytical Data of Chapter 4

N1,N10 -bis(6-(bis(pyridin-2-ylmethyl)amino)-6-oxo-5-(2-(pyren-1-yl)acetamido)hexyl) decanediamide (73)

O O HN 6 NH

N N N HN N NH O O O N N O

1 Figure A.29.: H-NMR (400 MHz, CDCl3) of compound 73

O O HN 6 NH

N N N HN N NH O O O N N O

13 Figure A.30.: C-NMR (101 MHz, CDCl3) of compound 73

xviii O O HN 6 NH

N N N HN N NH O O O N N O

Figure A.31.: HRMS (ESI) m/z: calcd.: 1305.6648, found: 1305.6631 [M+H+]+ of compound 73.

70· Zn (71)

4,0 12000

O O

3,5 HN NH

10000

N 3,0 N 2+ N 2+ N HN Zn Zn NH O 8000 O O 2,5 N N O 71 4 NO3-

2,0 6000

1,5

4000 Absorbance [O.D.] Absorbance

1,0 Fluorescence Intensity Intensity Fluorescence [RFU ]

2000

0,5

0,0 0

300 400 500 600 700

Wavelength [nm]

Figure A.32.: Absorbance & Fluorescence Scan of compound 71 in water (HEPES 10 mM pH 7.4).

xix A. Analytical Data of Chapter 4

73· Zn (74)

12000 O O 4,0 HN 6 NH

10000 3,5

N N N 2+ 2+ HN Zn Zn N 3,0 NH O O O 8000 N N O

2,5 - 4 NO3 74

6000

2,0

1,5

4000 Absorbance [O.D.] Absorbance

1,0 Fluorescence Intensity [RFU] Intensity Fluorescence

2000

0,5

0,0 0

300 400 500 600 700

Wavelength [nm]

Figure A.33.: Absorbance & Fluorescence Scan of compound 74 in water (HEPES 10 mM pH 7.4). tert-butyl(R)-4-((((9H -ﬂuoren-9-yl)methoxy)carbonyl)amino)-5-(bis (pyridin-2-ylmethyl)amino)-5-oxopentanoate (76)

N O H O N N N O

76 O O

1 Figure A.34.: H-NMR (400 MHz, CDCl3) of compound 76 xx N O H O N N N O

76 O O

13 Figure A.35.: C-NMR (101 MHz, CDCl3) of compound 76

N O H O N N N O

76 O O

Figure A.36.: HRMS (ESI) m/z: calcd.: 607.2915, found: 607.2913 [M+H+]+ of compound 76.

xxi A. Analytical Data of Chapter 4 tert-butyl (R)-4-amino-5-(bis(pyridin-2-ylmethyl)amino)-5-oxopentanoate (77)

N O N H2N N

O O 77

1 Figure A.37.: H-NMR (400 MHz, CDCl3) of compound 77

N O N H2N N

O O 77

13 Figure A.38.: C-NMR (101 MHz, CDCl3) of compound 77

xxii N O N H2N N

O O 77

Figure A.39.: HRMS (ESI) m/z: calcd.: 385.2234, found: 385.2231 [M+H+]+ of compound 77. tert-butyl (R)-5-(bis(pyridin-2-ylmethyl)amino)-5-oxo-4-(pyrene-1-carboxamido) pentanoate(79)

N O H N N N O

O O 79

1 Figure A.40.: H-NMR (400 MHz, CDCl3) of compound 79

xxiii A. Analytical Data of Chapter 4

N O H N N N O

O O 79

13 Figure A.41.: C-NMR (101 MHz, CDCl3) of compound 79

xxiv APPENDIX B

Analytical Data of Chapter 5

xxv B. Analytical Data of Chapter 5

2,2’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene))bis(methaneylylidene))diphenol (93)

N N OH HO 93

1 Figure B.1.: H-NMR (300 MHz, DMSO-d6) of compound 93

N N OH HO 93

Figure B.2.: HRMS (ESI) m/z: calcd.: 269.1258, found: 269.1258 [M+H+]+ of compound 93.

xxvi Probe (89)

N 3+ Fe N O O 89

Figure B.3.: HRMS (ESI) m/z: calcd.: 322.0399, found: 322.0401 [M+H+]+ of compound 89.

2-(benzo[d]oxazol-2-yl)-4-methylphenol (105a)

N HO 105a

1 Figure B.4.: H-NMR (300 MHz, CDCl3) of compound 105a

xxvii B. Analytical Data of Chapter 5

N HO 105a

Figure B.5.: HRMS (ESI) m/z: calcd.: 226.0863, found: 226.0864 [M+H+]+ of compound 105a.

3-(benzo[d]oxazol-2-yl)-2-hydroxy-5-methylbenzaldehyde (101a)

N HO O 101a

1 Figure B.6.: H-NMR (300 MHz, CDCl3) of compound 101a

xxviii O

N HO O 101a

Figure B.7.: HRMS (ESI) m/z: calcd.: 254.0812, found: 254.0812 [M+H+]+ of compound 101a.

N HO O 101a

Figure B.8.: Absorbance & Fluorescence Scan of compound 101a in water (Tris pH 7.4, 10 mM) .

xxix B. Analytical Data of Chapter 5

6,6’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene))bis(methaneylylidene)) bis(2-(benzo[d]oxazol-2-yl)-4-methylphenol) (108a)

N HO N

HO N N

O 108a

1 Figure B.9.: H-NMR (400 MHz, CDCl3) of compound 108a

N HO N

HO N N

O 108a

13 Figure B.10.: C-NMR (101 MHz, CDCl3) of compound 108a xxx O

N HO N

HO N N

O 108a

Figure B.11.: HRMS (ESI) m/z: calcd.: 531.2027, found: 531.2025 [M+H+]+ of compound 108a.

Probe (107a)

O 3+ N Fe O N N

O 107a

Figure B.12.: Absorbance & Fluorescence Scan of compound 107a in water (Tris pH 7.4, 10 mM) .

xxxi B. Analytical Data of Chapter 5

Methyl-3-(benzo[d]oxazol-2-yl)-4-hydroxybenzoate (105b)

O O O

N HO 105b

1 Figure B.13.: H-NMR (400 MHz, DMSO-d6) of compound 105b

O O O

N HO 105b

13 Figure B.14.: C-NMR (101 MHz, DMSO-d6) of compound 105b

xxxii O O O

N HO 105b

Figure B.15.: HRMS (ESI) m/z: calcd.: 270.0761, found: 270.0764 [M+H+]+ of compound 105b.

Methyl 3-(benzo[d]oxazol-2-yl)-5-formyl-4-hydroxybenzoate (101b)

O O O

N HO O 101b

1 Figure B.16.: H-NMR (300 MHz, DMSO-d6) of compound 101b

xxxiii B. Analytical Data of Chapter 5

O O O

N HO O 101b

Figure B.17.: HRMS (ESI) m/z: calcd.: 298.0710, found: 298.0712 [M+H+]+ of compound 101b.

Methyl 3-(benzo[d]oxazol-2-yl)-5-((E)-((2-(((E)-3-(benzo[d]oxazol-2-yl)-2- hydroxy-5-(methoxycarbonyl)benzylidene)amino)ethyl)imino)methyl)-4- methylbenzoate (108b)

O O O

N HO N

HO N N

O O 108bO

1 Figure B.18.: H-NMR (300 MHz, CDCl3) of compound 108b xxxiv O O O

N HO N

HO N N

O O 108bO

Figure B.19.: HRMS (ESI) m/z: calcd.: 619,1823, found: 619,1826 [M+H+]+ of compound 108b.

Probe (107b)

O O O

O 3+ N Fe O N N

O O O 107b

Figure B.20.: Absorbance & Fluorescence Scan of compound 107b in water (HEPES pH 7.4, 10 mM).

xxxv B. Analytical Data of Chapter 5

7-(dimethylamino)naphthalen-2-ol (96)

N OH

1 Figure B.21.: H-NMR (300 MHz, CDCl3) of compound 96

N OH

Figure B.22.: HRMS (ESI) m/z: calcd.: 186.0924, found: 186.0924 [M+H+]+ of compound 96.

xxxvi 6-(dimethylamino)-3-hydroxy-2-naphthaldehyde (94)

N OH

O 94

1 Figure B.23.: H-NMR (300 MHz, CDCl3) of compound 94

N OH

O 94

Figure B.24.: HRMS (ESI) m/z: calcd.: 216.1019, found: 216.1020 [M+H+]+ of compound 94.

xxxvii B. Analytical Data of Chapter 5

3,3’-((1E,1’E)-(ethane-1,2-diylbis(azaneylylidene))bis(methaneylylidene))bis (7-(dimethylamino)naphthalen-2-ol) (98)

N OH

N OH 98

1 Figure B.25.: H-NMR (300 MHz, DMSO-d6) of compound 98

N OH

N OH 98

13 Figure B.26.: C-NMR (101 MHz, DMSO-d6) of compound 98

xxxviii N OH

N OH 98

Figure B.27.: HRMS (ESI) m/z: calcd.: 455.2442, found: 455.2440 [M+H+]+ of compound 98.

Probe (99)

N 3+N N O Fe O

99 N

Figure B.28.: Absorbance & Fluorescence Scan of compound 99 in water (Tris pH 7.4, 10 mM).

xxxix B. Analytical Data of Chapter 5

5-(benzo[d]oxazol-2-yl)-4-hydroxyisophthalaldehyde (116)

O O

N HO O 116

1 Figure B.29.: H-NMR (300 MHz, CDCl3) of compound 116

2-(benzo[d]oxazol-2-yl)-4,6-bis(((pyridin-2-ylmethyl)amino)methyl)phenol (120)

HN N

N HO HN N

120

1 Figure B.30.: H-NMR (300 MHz, CDCl3) of compound 120 xl HN N

N HO HN N

120

13 Figure B.31.: C-NMR (101 MHz, CDCl3) of compound 120

HN N

N HO HN N

120

Figure B.32.: HRMS (ESI) m/z: calcd.: 452,2081, found: 452,2083 [M+H+]+ of compound 120.

xli B. Analytical Data of Chapter 5

Probe 118

N N N

N HO N N

N 118

1 Figure B.33.: H-NMR (300 MHz, CDCl3) of compound 118

N N N

N HO N N

N 118

Figure B.34.: HRMS (ESI) m/z: calcd.: 634.2925, found: ,634.2927 [M+H+]+; 656.2750, found: 656.2751 [M+Na+]+ of compound 118.

xlii Probe (121)

N Zn2+ N N O

N O N 2+Zn N N

121

Figure B.35.: Absorbance & Fluorescence Scan of compound 121 in water (HEPES pH 7.4, 10 mM).

N,N -dimethyl-N 2-(pyridin-2-ylmethyl)ethane-1,2-diamine (125)

H N N N 125

1 Figure B.36.: H-NMR (300 MHz, CDCl3) of compound 125

xliii B. Analytical Data of Chapter 5

2-(benzo[d]oxazol-2-yl)-4,6-bis(((2-(dimethylamino)ethyl) (pyridin-2-ylmethyl) amino)methyl)phenol (126)

N O N N N HO N

N 126

1 Figure B.37.: H-NMR (400 MHz, CDCl3) of compound 126

N O N N N HO N

N 126

13 Figure B.38.: C-NMR (101 MHz, CDCl3) of compound 126

xliv Probe (127)

N Zn2+ N N O

N O N 2+Zn N

N 127

Figure B.39.: Absorbance & Fluorescence Scan of compound 127 in water (HEPES pH 7.4, 10 mM).

Probe (109)

N O 2+ O Cu O O N

O 109

Figure B.40.: Absorbance & Fluorescence Scan of compound 109 in water (Tris pH 7.4, 10 mM).

xlv

APPENDIX C

Analytical Data of Chapter 6

xlvii C. Analytical Data of Chapter 6

6-Bromo-2-(4-bromophenyl)-1H-indole (130)

Br Br N H 130

1 Figure C.1.: H-NMR (300 MHz, CDCl3) of compound 130

Figure C.2.: NOESY (300 MHz, CDCl3) of compound 130. Proton 9 can couple to the adjacent protons 6, 15 and 11.

xlviii Br Br N H 130

13 Figure C.3.: C-NMR (126 MHz, CDCl3) of compound 130

Br Br N H 130

Figure C.4.: HRMS (ESI) m/z: calcd.: 349.9008, found: 349.9009 [M-H+]− of compound 130

xlix C. Analytical Data of Chapter 6

2-(4-Cyanophenyl)-1H -indole-6-carbonitrile (129)

N N N H 129

1 Figure C.5.: H-NMR (300 MHz, DMSO-d6) of compound 129

N N N H 129

13 Figure C.6.: C-NMR (126 MHz, DMSO-d6) of compound 129

l N N N H 129

Figure C.7.: HRMS (ESI) m/z: calcd.: 242.0723, found: 242.0723 [M-H+]− of compound 129

Benzamidine · HCl (138)

HN NH2

138

1 Figure C.8.: H-NMR (300 MHz, DMSO-d6) of compound 138

li C. Analytical Data of Chapter 6

HN NH2

138

13 Figure C.9.: C-NMR (126 MHz, DMSO-d6) of compound 138

HN NH2

138

Figure C.10.: HRMS (ESI) m/z: calcd.: 121.0761, found: 121.0761 [M+H+]+ of compound 138

lii 1-(tert-Butoxycarbonyl)-6-cyano-2-(4-cyanophenyl)-1H -indole (139)

N N N Boc 139

1 Figure C.11.: H-NMR (300 MHz, DMSO-d6) of compound 139

DAPI · 2HCl (34)

NH HN N NH H 2 NH2 34

1 Figure C.12.: H-NMR (300 MHz, DMSO-d6) of compound 34

liii C. Analytical Data of Chapter 6

NH HN N NH H 2 NH2 34

Figure C.13.: HRMS (ESI) m/z: calcd.:278.1400 , found:278.1404 [M+H+]+ of compound 34

4-Bromo-2-(4-bromophenyl)-1H-indole (133)

Br N H 133

1 Figure C.14.: H-NMR (300 MHz, DMSO-d6) of compound 133

liv Br

Br N H 133

13 Figure C.15.: C-NMR (126 MHz, DMSO-d6) of compound 133

2-(4-cyanophenyl)-1H-indole-4-carbonitrile (172)

N N H 172

1 Figure C.16.: H-NMR (300 MHz, DMSO-d6) of compound 172

lv C. Analytical Data of Chapter 6

N N H 172

13 Figure C.17.: C-NMR (126 MHz, DMSO-d6) of compound 172

2-(4-Carbamimidoylphenyl)-1H -indole-4-carboximidamide (141)

HN NH2

N NH H 2 141

1 Figure C.18.: H-NMR (300 MHz, DMSO-d6) of compound 141

lvi 6-Bromo-1-(tert-butoxycarbonyl)-1H -indole (150)

Br N Boc 150

1 Figure C.19.: H-NMR (300 MHz, DMSO-d6) of compound 150

6-Bromo-1-(tert-butoxycarbonyl)-2-(triemthylstannyl)-1H -indole (151)

Sn Br N Boc 151

1 Figure C.20.: H-NMR (300 MHz, DMSO-d6) of compound 151

lvii C. Analytical Data of Chapter 6

6-Bromo-1-(tert-butoxycarbonyl)-2(iodo)-1H -indole (153)

I Br N Boc 153

1 Figure C.21.: H-NMR (300 MHz, CDCl3) of compound 153

4,4‘-(1H -indole-2,6-diyl) dibenzonitrile (173)

N N H 173 N

1 Figure C.22.: H-NMR (300 MHz, DMSO-d6) of compound 173

lviii N N H 173 N

13 Figure C.23.: C-NMR (126 MHz, DMSO-d6) of compound 173

N N H 173 N

Figure C.24.: HRMS (ESI) m/z: calcd.: 318.1036, found: 318.1036 [M-H+]− of compound 173

lix C. Analytical Data of Chapter 6

4,4‘-(1H -indole-2,6-diyl) dianiline · 2HCl (148)

N NH H 2 HN 148 2 HCl

NH 2

1 Figure C.25.: H-NMR (300 MHz, DMSO-d6) of compound 148

4,4‘-(1H -indole-2,6-diyl) dibenzimidamide · 2HCl (156)

NH 2 N H 2 HCl

H2N 156

1 Figure C.26.: H-NMR (500 MHz, DMSO-d6) of compound 156

lx NH 2 N H 2 HCl

H2N 156

13 Figure C.27.: C-NMR (126 MHz, DMSO-d6) of compound 156

NH 2 N H 2 HCl

H2N 156

Figure C.28.: HRMS (ESI) m/z: calcd.: 300.1496, found: 300.1497 [M+H+]+ of compound 156

lxi C. Analytical Data of Chapter 6

NH 2 N H 2 HCl

H2N 156

Figure C.29.: HPLC: C18, MeCN/H2O, 1:99 to 99:1 in 20 min) rt: 10.1 min

1-(tert-Butoxycarbonyl)-2,6-bis(4-N,N’-Bis-(tert-Butyloxycarbonyl) phenyl)-1H -indole (163)

BocN NHBoc NH NBoc N H BocHN N 163 H

1 Figure C.30.: H-NMR (300 MHz, DMSO-d6) of compound 163

lxii BocN NHBoc NH NBoc N H BocHN N 163 H

Figure C.31.: HRMS (ESI) m/z: calcd.: 884.4553, found: 884.4566 [M+H+]+. of compound 163

1-(tert-Butoxycarbonyl)-2,6-bis(4-N,N’-Bis-(tert-Butyloxycarbonyl) phenyl)-1H -indole · 2 HCl (164)

NH 2 NH 2 HCl NH N H

H2N N 164 H

1 Figure C.32.: H-NMR (400 MHz, DMSO-d6) of compound 164

lxiii C. Analytical Data of Chapter 6

NH 2 NH 2 HCl NH N H

H2N N 164 H

Figure C.33.: HRMS (ESI) m/z: calcd.: 384.1932, found: 384.1934 [M+H+]+ of compound 164

2-(4-Amidinophenyl)-3-(2-hydroxy-5-nitro) benzyl-6-amidinoindole · 2 HCl (168)

OH O2N

NH HN N NH H 2 NH2 168 2 HCl

1 Figure C.34.: H-NMR (300 MHz, Methanol-d4) of compound 168

lxiv 3-(5-amino-2-hydroxybenzyl)-2-(4-carbamimidoylphenyl)-1H -indole- 6-carboximidamide · 3 TFA (169)

OH H2N

NH HN N NH H 2 NH2 169 2 HCl

Figure C.35.: HRMS (ESI) m/z: calcd.: 399.1928, found: 399.1926 [M+H+]+. of compound 169

lxv