Sequence-Addressed Assemblies of Trisoligonucleotides into Nanoscale Motifs and Structural Studies

Christos Panagiotidis

Dissertation

zur Erlangung des Grades

“Doktor der Naturwissenschaften”

an der Fakultät für Chemie und Biochemie der Ruhr-Universität Bochum

Bochum/Wuppertal 2016

This dissertation is based on the research work carried out during the period April 2012 to December 2015 at the chair of Organic Chemistry I, Bioorganic Chemistry of the Ruhr- University Bochum, Germany.

First referee: Prof. Dr. Günter von Kiedrowski

Second referee: Prof. Dr. Frank Schulz

Day of submission: 2. Dec. 2015

Day of disputation: 29. Feb. 2016

Herewith I declare that the following work has been carried out independently by myself and all the sources of help and services used during this work have been reported herein. I further declare that I have not submitted this thesis in this or in a similar form to any other university or college. No competing interest is declared.

Christos Panagiotidis, December 2015

Danksagungen (Acknowledgements)

Ich danke Herrn Prof. Dr. Günter von Kiedrowski für das interessante Forschungsgebiet, die Möglichkeit zur freien Gestaltung und Durchführung der Arbeit, der Diskussionsbereitschaft und interessanten Gesprächen über der Arbeit hinaus. Auch danke ich Herrn Prof. Dr. Frank Schulz für die Übernahme des Koreferates.

Herrn Dr. Wolf Matthias Pankau danke ich ebenfalls für die ständige Diskussionsbereitschaft und Herrn Dr. Volker Patzke für das Korrekturlesen des Manuskripts.

Herrn Michael „Stoßrohr“ Wüstefeld danke ich für die Durchführung der automatisierten Synthesen, für das offene Ohr und die regen Unterhaltungen.

Frau Katja Schulz danke ich für ihre Tätigkeit als chemisch-technische Assistentin, ihre stete Unterstützung beim bioorganischen Praktikum und der freundlichen Atmosphäre.

Florian Kaschuba und Carsten Lodwig danke ich für die Behebung technischer Ungereimtheiten. Frau Stephanie Nolte aus der Hollmann-Gruppe danke ich für Einweisung und Nutzung des Bioimagers.

Der Bürofee Stefanie Wittmann und ihren zwei Hunden wünsche ich alles Gute im weiteren Leben. Lache immer laut und herzhaft.

Meinen Kollegen und Mitstreitern sei herzlichst für die recht entspannte Arbeitsatmosphäre unter Berücksichtigung des allwährenden Wahnsinns gedankt. Wir sind bekloppt und das ist auch gut so. Das wären Daniel Kramer, Nora Leistner, Chris Nielinger, Elena Palmieri, Miriam Patzke, Jana Pyka, Berit Sorge, aber vor allem Markus Rethmeier, der am meisten von meinem Gejammer abbekommen hat und Ilhan Sevim für die gelegentlichen philosophischen Unterhaltungen.

Allen Studenten (bzw. das billige Kanonenfutter) aus den Praktika, den Abschlussarbeiten und dem Labor sei gedankt, aber vor allem Sebastian Michalski, Elric Engelage, Holger Rabuske und Ivan Grgic. Bei Herrn Dimitri Kolmanovsky erinnere ich mich gerne an den hilfreichen E-Mail-Austausch speziell am Anfang der Arbeit.

Ich danke meiner Familie.

“Just a heads up: We're gonna have a superconductor turned up full blast and pointed at you for the duration of this next test. I'll be honest, we're throwing science at the wall here to see what sticks. No idea what it'll do. Probably nothing. Best-case scenario, you might get some superpowers. Worst case, some tumors, which we'll cut out.”

Cave Johnson,

CEO of Aperture Science, Inc.

(Portal 2)

Abstract

Trisoligonucleotides have been previously established as building blocks in the construction of DNA-based nanostructures, like tetrahedral and dodecahedral scaffolds, possible by a sequence-addressed self-assembly. The resulting care-like shapes are potentially usable as nanoscale containers.

Trisoligonucleotides may contain C3h-symmetrical linkers that connect three oligonucleotide arms. One previously established linker was based on 1,3,5-trishydroxypropylbenzene, but required a lengthy and laborious synthesis over up to twelve steps. This is a bottle-neck in subsequent studies with DNA. Six steps of the previously applied linker synthesis pathway have been cut by following a Heck-coupling route, which improved the overall yield. Additionally, a new generation of isocyanurate-based linkers was introduced, which even further shortened preparation to just two or three steps, respectively.

Linkers require alkylene chains to allow folding of the single-strands into polyhedral scaffolds. So far studies on the flexibility of methylene- and propylene-chains exist and not for ethylene-chains. Sets of trisoligonucleotides for tetrahedral assemblies were prepared via automated synthesis, purified via preparative polyacrylamide gel electrophoresis and confirmed by MALDI-TOF-MS. Hybridisation products were studied with agarose gel electrophoresis and showed relative stability against enzymatic digestion with mung bean nuclease, which in our hands is a proof for the existence of discrete closed nanoobjects. Likewise, isocyanurate-based trisoligonucleotides shared similar results.

An alternative sequence-pattern for trisoligonucleotides was studied, namely, two of three arms sharing an identical oligonucleotide sequence leading to situations below so called maximal instruction. A set of these semi-addressable building blocks were assembled and studied with native agarose gel electrophoresis. New conceivable motifs were accessible expanding the repertoire of trisoligonucleotide-based nanostructures.

Tetrahedral scaffolds were intercalated with the anti-cancer drugs dauno- and doxorubicin. UV/VIS spectroscopy and fluorescence quenching experiments were done in comparison to linear oligonucleotides and hint at additional binding pockets close to the vertices of the tetrahedron enhancing the maximally possible binding capacity of duplex DNA.

Keywords: structural DNA nanotechnology; C3h-symmetrical linker; isocyanurate; trisoligonucleotide; Sequence- addressability; self-assembly; DNA tetrahedron; nanosynthesis; anthracycline; intercalation; fluorescence quenching Content Content

Prologue ...... 1

I. Theoretical Background...... 2

I.1. Nanotechnology ...... 2

I.2. Deoxyribonucleic acid ...... 3

I.3. DNA Nanotechnology ...... 8

I.4. Chemically unmodified DNA Nanostructures ...... 10

I.5. Chemically modified DNA Nanostructures ...... 13

I.6. Static DNA Nanostructures for Drug Delivery ...... 21

II. Aim of Work ...... 23

III. Results and Discussion ...... 25

III.1. Synthesis of C3h-Symmetrical Trislinkers ...... 25

III.2. Oligonucleotide Synthesis ...... 34

III.2.1. Trisoligonucleotide Synthesis Protocol ...... 34

III.2.2. Phosphoramidite Oligonucleotide Synthesis ...... 37

III.2.3. Sequence Design ...... 39

III.2.4. Purification and Quality Control ...... 43

III.2.5. Long-Term Stability Control ...... 47

III.2.6. Solid Supports for Trisoligonucleotide Synthesis ...... 48

III.2.7. Fluorous Affinity Purification Studies ...... 55

III.3. DNA-Tetrahedron Assembly Experiments ...... 68

III.3.1. Hybridization Conditions ...... 68

III.3.2. Effect of short Alkylene Chains in T1, T2 and T3 Assembly Studies...... 70

III.3.3. Assembly and Digestion Experiments of TN ...... 75

III.3.4. Intermixing Assembly Experiments between T1, TN and T3 ...... 76

III.4. The Concept of UNO-, DOS- and TRE- Sequence Patterns...... 78 Content

III.4.1. Introduction ...... 78

III.4.2. Hybridization and Digestion of the TRE-Pattern TY ...... 79

III.4.3. Hybridization and Digestion of the DOS-Pattern TD ...... 82

III.4.4. Sequence-Addressed Motif Designs via Intermixing Hybridizations ...... 86

III.5. Anthracycline Intercalation ...... 92

III.5.1. Qualitative Analysis ...... 92

III.5.2. Quantitative Analysis I ...... 95

III.5.3. Quantitative Analysis II: Fluorescence Spectroscopy ...... 98

IV. Summary and Outlook ...... 106

V. Experimental Section...... 112

V.1. Methods and Equipment ...... 112

V.1.1. Chromatographic Methods ...... 112

V.1.2. Spectroscopic Methods ...... 114

V.1.3. Mass Spectrometric Methods ...... 115

V.1.4. Electrophoretic Methods ...... 116

V.1.5. Oligonucleotide Synthesis ...... 119

V.1.5. Enzymatic Digestion of DNA Single-Strands with Mung Bean Nuclease ...... 121

V1.7. Further Equipment and Software ...... 122

V.2. Chemicals ...... 124

V.2.1. Solvents ...... 124

V.2.2. Educts, Reagents and Other Chemicals ...... 125

V.2.3. Automated Oligonucleotide Synthesis ...... 127

V.2.4. Biomolecular Reagents...... 127

V.3. Organic Synthesis ...... 129

V.3.01. AOC-DMT-PNO-T1 (05) ...... 129

V.3.02. 1,3,5-triacetybenzene (12) ...... 130 Content

V.3.03. 2,2',2''-(benzene-1,3,5-triyl)triacetic acid (08) ...... 131

V.3.04. 2,2',2''-(benzene-1,3,5-triyl)triacetic acid (09) ...... 132

V.3.05. 1,3,5-trishydroxyethylbenzene (10) ...... 132

V.3.06. AOC-AOC-T2 (18) ...... 133

V.3.07. AOC-AOC-PNO-T2 (19) ...... 134

V.3.08. DMT-DMT-T2 (16) ...... 135

V.3.09. DMT-DMT-PNO-T2 (17) ...... 136

V.3.10. DMT-T2 (13) ...... 137

V.3.11. AOC-DMT-T2 (14) ...... 138

V.3.12. AOC-DMT-PNO-T2 (15) ...... 139

V.3.13. 1,3,5-tris(E- methyl acryoyl) benzene (28) ...... 140

V.3.14. 1,3,5-tris(methyl propyloyl) benzene (24) ...... 141

V.3.15. Allyl benzyl ether (31) ...... 142

V.3.16. 1E,3E,5Z-1,3,5-tris(benzyl oxy isoallyl) benzene (32) ...... 143

V.3.17. 1,3,5-tris(3-hydroxy propyl) benzene (25) ...... 144

V.3.18. AOC-T3 (33) ...... 145

V.3.19. AOC-DMT-T3 (34) ...... 146

V.3.20. AOC-DMT-PNO-T3 (35) ...... 147

V.3.21. AOC-TN ...... 148

V.3.22. DMT-TN (37) ...... 149

V.3.23. AOC-DMT-TN (38) ...... 150

V.3.24. AOC-DMT-PNO-TN (39) ...... 151

V.3.25. DMT-DMT-PNO-TN (40) ...... 153

V.3.26. DMT-DMT-PNO-TN (41) ...... 154

V.3.27. Perfluorooctyl ethylthioethyl hydroxide (47) ...... 155

V.3.28. F-TAG amidite (48) ...... 155 Content

VI. References ...... 157

VII. Appendix ...... 168

VII.1 Glossary ...... 168

VII.2 Parr Hydrogenation Apparatus German Instruction Manual ...... 174

VII.3 Full Sequence Pool ...... 175

VII.4 Dimer Energies ...... 179

VII.5 Full MALDI-TOF-MS Data Set ...... 182

VII.6 Alternative Visualisation of Anthracycline Binding Efficiencies ...... 187

VII.7 NMR-Spectra ...... 188

V.3.01. AOC-DMT-PNO-T1 (05) ...... 188

V.3.02. 1,3,5-triacetybenzene (12) ...... 191

V.3.03. 2,2',2''-(benzene-1,3,5-triyl)triacetic acid (08) ...... 193

V.3.04. 2,2',2''-(benzene-1,3,5-triyl)triacetic acid (09) ...... 195

V.3.05. 1,3,5-trishydroxyethylbenzene (10) ...... 197

V.3.06. AOC-AOC-T2 (18) ...... 199

V.3.07. AOC-AOC-PNO-T2 (19) ...... 201

V.3.08. DMT-DMT-T2 (16) ...... 204

V.3.09. DMT-DMT-PNO-T2 (17) ...... 206

V.3.10. DMT-T2 (13) ...... 209

V.3.11. AOC-DMT-T2 (14) ...... 210

V.3.12. AOC-DMT-PNO-T2 (15) ...... 212

V.3.13. 1,3,5-tris(E- methyl acryl) benzene (28) ...... 215

V.3.14. 1,3,5-tris(methyl ester propyl) benzene (24)...... 217

V.3.15. Allyl benzyl ether (31) ...... 219

V.3.16. 1E,3E,5Z-1,3,5-tris(benzyl oxy isoallyl) benzene (32) ...... 221

V.3.17. 1,3,5-tris(3-hydroxy propyl) benzene (25) ...... 223 Content

V.3.18. AOC-T3 (33) ...... 225

V.3.19. AOC-DMT-T3 (34) ...... 227

V.3.20. AOC-DMT-PNO-T3 (35) ...... 229

V.3.21. AOC-TN ...... 232

V.3.22. DMT-TN (37) ...... 234

V.3.23. AOC-DMT-TN (38) ...... 236

V.3.24. AOC-DMT-PNO-TN (39) ...... 238

V.3.25. DMT-DMT-PNO-TN (40) ...... 241

V.3.26. DMT-DMT-PNO-TN (41) ...... 243

V.3.27. Perfluorooctyl ethylthioethyl hydroxide (47) ...... 246

V.3.28. F-TAG amidite (48) ...... 248

VII. Curriculum Vitae ...... 251

Prologue

Prologue

Above screenshot is taken from the videogame Transcripted (2012; By Alkemi). Set in the near-future, the player assumes quite fittingly the role of a PhD student named Adam who remotely controls a ‘Nano Probe’ inside mutated human tissue in front of his computer to study a pathogen of unknown origin. The major goal in the game is to destabilize the ‘pseudo-DNA’ in the mutated cells by gathering ‘pseudo-nucleotides’ in the plasma and adding them to the ‘DNA-chain’, whilst fending off antibody-like entities.

This fictional scenario is not far-fetched from reality though. Scientific and technical advancements in the field of nanotechnology already deal with the concepts of nanoconstruction, nanomachinery and the precise manipulation of matter on the nanoscale level. Science-fiction yet again just serves as an inspiration for future technological developments.

The following thesis also deals with the concept of nanoconstruction by humbly contributing to the field of DNA nanotechnology with studies on the construction of DNA nanocages, sequence-controlled building-block assemblies on the nanoscale into different DNA motifs and potential drug transport capabilities of aforementioned DNA cages in intercalation studies. Admittedly, this “Nano Probe” would have been useful as a tool in this work.

1 I. Theoretical Background

I. Theoretical Background

I.1. Nanotechnology

Nanotechnology is an interdisciplinary field of research, which spans and connects the areas of semiconductor physics, supramolecular and surface chemistry, molecular biology and medicine. Major goals lie in the manufacturing, manipulation und analysis of structure-, function- and scaffolding units on the nanometre scale (~1-100 nm). In this order of magnitude the influence of the surface properties of an object start to overtake the volume properties, which progressively require quantum mechanical effects to be taken into account. The shape and/or the size of an object therefore determines its properties.

Major goals in research involve the fabrication of new materials, the development of nanomachines and nanorobots, the downscaling of circuitry, quantum computing, new data- and energy storage devices and efficient power generation.[1]

The production of nanoobjects distinguishes two basic methodologies that encompass the top-down and bottom-up approach with methods like lithographic patterning of nanostructures meeting somewhere in the middle.[2] The top-down approach starts with microscale objects and aims to shape them into smaller ones. This approach is a continuation of the microtechnological field utilizing lithographic methods. These approaches employ large machinery such as electron beam microscopes. In contrast, bottom-up approaches start with small objects like atoms or molecules and arrange those single components to larger assemblies. This utilizes the physical and chemical principles of self-assembly and self-organization. Exemplary materials of the bottom-up approach include gold nanoparticles,[3,4] quantum dots,[5] fullerenes[6] and carbon nanotubes.[7] Scanning Probe Microscopy encompasses typical methods in the structural analysis of nanomaterials.[8]

Deoxyribonucleic acid (DNA) is another viable material exploiting the specificity of nucleobase pairing, which leads to the area of DNA nanotechnology. The next paragraphs introduce DNA and the associated subfield of DNA nanotechnology.

2 I. Theoretical Background

I.2. Deoxyribonucleic acid

Nucleic acids are biopolymers built up of nucleotides.[9] The most prominent representative is the deoxyribonucleic acid (DNA), whose most common use in nature lies in the storage of genetic information in terrestrial life forms and DNA-viruses. The transport of information is accomplished via transcription of this information to a ribonucleic acid (RNA), which releases the information on a destined target via translation leading primarily to the formation of proteins.

The polymeric backbone of DNA consists of the sugar 2’-deoxyribose (Fig. I-1). The sugar units are bound as a hydrophilic phosphate diester at their C-3’- and C-5’-hydroxy- functions. Via an N- glycosidic bond a distinct set of two purine- derivates and two pyrimidine-derivates are attached laterally to the backbone. Those four Fig. I-1: The building blocks of DNA. Purine and pyrimidine are added for reference. nucleobases entail the two purine-derivates adenine (A) and guanine (G) and the two pyrimidine-derivates thymine (T) and cytosine (C). The former two attach at their N-9 position and the latter at their N-1 position onto the C-1 position. The structure of RNA contains a ribose and uracil instead of 2’-deoxyribose and thymine (Not pictured).

A unit of a nucleobase and the sugar is called a nucleoside. The addition of a phosphate- group at the C-3’ and/or C-5’ position turns it into a nucleotide. A DNA single-strand is consensually read in the direction of C-5’ of the first nucleotide unit to C-3’ of the last unit.

3 I. Theoretical Background

Fig. I-2: Layout of an exemplary DNA double-strand: The single-strands contain the sugar-phosphate backbone and the nucleobases A, C, G and T. Hydrogen bridge bonds on the nucleobases lead to an anti-parallel double-strand. (Modified image taken from reference [10]) Under certain conditions defined by temperature, pH and salt concentration DNA forms a double-strand (Fig. I-2). This is due to non-covalent, hydrophobic interactions between nucleobases of two anti-parallel strands. Those interactions contain primarily nucleobase to nucleobase hydrogen bridge bonds and π–stacking of the aromatic cycles. Under physiological conditions the nucleobases hybridize in two distinct pairs named after Watson and Crick:[11] Adenine and thymine form two hydrogen bridge bonds, whereas cytosine and guanine form three. Different numbers of hydrogen bridge bonds lead to energetically favoured A/T and C/G constellations. A common double-strand consists of two fully complementary single-strands. The additional hydroxide-functionality in RNA in turn favours an intramolecular hybridization into folded structures. The double-stranded nature of DNA allows for molecular recognition and self-organization of the genetic information.

Double-strand formation results in a helical twist, in which both strands intertwine around a common axis. The nucleobase pairs lie in the inside, whereas the sugar-phosphate chain

4 I. Theoretical Background forms the outside backbone. Depending on certain conditions the most common DNA forms are called A-DNA, B-DNA and Z-DNA.

Fig. I-3: Tilted side view of A-, B- and Z-DNA. (Modified image taken from reference [12]) B-DNA is the typical form under physiological conditions (Fig. I-3).[12] It has 0.34 nm of rise per base pair, a helix diameter of 2.37 nm and requires 10.4 base pairs for a complete turn of a helix with a 3.54 nm pitch. Dehydration leads to the formation of the broader shaped A- DNA with a shorter rise per base pair of 0.23 nm, a helix diameter of 2.55 nm and requires 11 base pairs for a complete turn of a helix with 2.53 nm pitch. Both forms have a right- handed screw sense unlike the Z-DNA form with a left-handed one. The narrower shaped Z-DNA type requires 0.38 nm of rise per base pair, a narrower helix diameter of 1.84 nm and requires 12 base pairs per turn of its helix with a 4.56 nm pitch. Z-DNA tends to form in high salt concentrations. The DNA sequence is G and C rich and lies in an alternating pattern such

[13] as CGCGCG. This meta-stable conformation has its phosphoryl-groups zigzag along the backbone. Dehydrated B-DNA in the presence of metal ions like Li+ or Mg2+ can lead to the less common C-DNA conformation similar to B-DNA.[14,15] Another less common

[16] conformation is D-DNA, which forms in an A and T rich sequence pattern in the likes of ATATAT and requires only 8 base pairs for a complete turn.

Double-stranded B-DNA possesses with a persistence length of 53 nm (~150 bp; ~1 nm for ssDNA)[17] a minimum stiffness required in nanoconstruction qualifying it as a semi-rigid polymer.

A variety of different other hydrogen bridge bonds are conceivable like reverse Watson- Crick-, Hoogsteen- or reverse Hoogsteen base pairs (Fig. I-4). Some of them can be achieved via protonation of a nucleobase-nitrogen. A guanine-rich sequence for example tends to

5 I. Theoretical Background

[18] form a G-quadruplex motif under Hoogsteen base pairing. A Watson-Crick double helix exposes its nucleobases at its major groove allowing for further non-standard interactions.[19,20]

Fig. I-4: A: Examples of different base pairings: a) G-C reverse Watson-Crick b) G-C Hoogsteen via protonation c) A-T reverse Watson-Crick d) A-T reverse Hoogsteen e) A-C reverse Hoogsteen B: a) G-quadruplex motif b) Various strand orientations of stacked G-quadruplexes. (Image taken from reference [18]) The combination of a Watson-Crick base pairing and a normal or reverse Hoogsteen base pairing, for instance, leads to the formation of a triple helix, whereby G and A in a homopurine strand hybridize with two C and two T in an homopyrimidine strand (Fig. I-5). The third strand attaches along the major groove to the Watson-Crick double-strand under acidic conditions (pH: 5-6) in the pattern T-A ∙ ∙ ∙ T and C-G ∙ ∙ ∙ CH+.

Fig. I-5: The homopurine and homopyrimidine strands forming double-strand A. The third strand B attaches onto A via Hoogsteen base pairing to triplex DNA C. This process is pH-dependent. The non-covalent nature of base pair interactions allows for full reversibility of the double- strand conformation. Temperatures above the melting point of the single-strands lead to denaturation. An effect also inducible via altering the pH or adding denaturing agents like

[21] urea. A DNA double-strand can be viewed in an equilibrium once a third complementary

6 I. Theoretical Background competing single-strand is introduced. A high excess of a competing strand can push one strand out of the double-strand and replace it.

DNA biosynthesis is an in vivo DNA amplification process described as the DNA replication. Template DNA can be extracted out of microorganisms and then multiplied in quantity via the in vitro DNA amplification method polymerase chain reaction. Enzymes such as Taq- polymerase replicate nucleic acids in the presence of nucleotide mixtures (dNTPs) and a template as initiator. Other methods include molecular cloning[22] and artificial gene synthesis, in which a gene is synthesized in vitro without the need for initial template DNA samples. It is an automated de novo solid-phase DNA synthesis procedure[23] that allows chain lengths of up to 100 nucleobases. For lengthier chains smaller synthesized fragments

[24,25] [26,27] can be connected post-synthetically either by chemical or enzymatic ligation.

The availability of biocatalysts allows DNA to be processed further in manifold ways (Fig. I- 6):[28] Endonucleases cleave duplex DNA at sequence-specific patterns. Telomerase elongates a single-strand by adding TTAGGG sequence repeats to the 3’-end. Sequence- specific nicking enzymes cleave one strand of duplex DNA structures. Exonucleases, like Exo III, cleave specific ends of duplex DNA domains through digestion of one strand. Ligase acts contrary to nicking enzymes by ligating several shorter nucleic acid strands. Helicase (not pictured) catalyses separation of duplex oligonucleotides into single-strands by an ATP- dependent reaction. Nucleic Fig. I-6: Examples of enzymatically biocatalysed transformations on DNA. (Modified Image taken from reference [28]) acid strands coupled chemically with new moieties like chemical functional groups, (bio)polymers and inorganic nanoobjects alter the molecules structural, morphological and/or self-assembly properties.[29] All these above-mentioned operations and modifications serve as important tools in the field of DNA nanotechnology.

7 I. Theoretical Background

I.3. DNA Nanotechnology

DNA nanotechnology makes use of the sequence-related self-recognition and self- organization of DNA in order to build functional structures on a molecular scale.[30,31] This is due to the predictable hybridization behaviour of DNA. The choice of the base sequence allows for a programmable self-assembly. The development started with the assembly of

[32–34] [35] static structures with applications in structural biology and computer science and other three-dimensional architectures.[36–39] Many applications in the field of nanomedicine and nanorobotics, however, require additional capabilities for controlled movement in three-

[40–45] dimensional spaces.

[46,47] [48] [49] [50] RNA and nucleic acid analogues like PNA, pRNA or LNA are valid alternatives in the construction of nanostructures,[51–54] but DNA is most commonly used because of availability.

The following schematic (Fig. I-7) summarizes exemplary areas of interest in the DNA nanotechnological field:[28]

Fig. I-7: Schematic overview of areas associated with DNA nanotechnology. (Modified image taken from reference [18]) Sensors: Sequence-specific DNA, equipped with a biosensor (probe), is immobilized on a transducing surface. DNA serves as a recognition ligand for sensing events. A stimulus like the hybridization of complementary DNA (target) leads to conformational changes on the

8 I. Theoretical Background probe, which alters the interaction between the biosensor and the transducing surface resulting in a change of signal output. Sensing can be based on electrochemical,[55–59] electronic,[60–63] optical,[64–67] microgravimetric[68–70] or mechanical[71–74] effects.

Functional enzyme/DNA structures are multi-component systems that are cross-linked on nanoparticles via nucleic acids and aim to mimic the cascaded biotransformations of intracellular catalytic processes. A typical dual enzyme cascade is glucose oxidase and horseradish peroxidase converting oxygen first into hydrogen peroxide and into water subsequently. This conversion runs in presence of glucose and a hydrogen peroxide probe like ABTS or Amplex Red.[75]

DNA Hydrogels: Polymerase elongated DNA chains may entwine non-covalently into hydrogel meshes. This metamaterial behaves in presence of high contents of water like a gelatinous solid and with reduced contents like an amorphous liquid. A pH change may also lead to a stimulus-induced gelation. The hydrogel has a hierarchical internal structure meaning that a moulded gel can return to its original shape. Additionally a hydrogel may encapsulate molecules or particles during the gelation process. Loaded drugs turn hydrogels possibly into biodegradable drug-carrier systems.[76]

DNA Machines: A set of nucleic acids with a partially overlapping sequence design perform mechanical operations that entail pulling, stretching or rotating motions.[77] These nanodevices require an external “input” as an environmental stimulus to trigger a

[41,45,78] [79–82] [83] [84–88] mechanical process like other nucleic acid strands, pH, ions, light or

[42,44,89–92] catalysing molecules. The first DNA machine driven by a hybridization force

[45] resembled molecular tweezers. In another example DNA walkers execute stepwise movements along linear tracks driven by a sequential addition of controls strands or by enzymatic hydrolysis.[93] The transport of gold nanoparticles along a fixed path was possible this way.[94]

DNA machines are dynamic structures with mechanical capabilities as opposed to static nano- and microobjects like scaffolds or cages. The latter example may lead to discrete structures of a fixed size suitable for defined cargo transport and nanoscopic scale synthesis. The assembly of these constructs either purely relies on the specificity of base pairing or it expands the constructional capabilities by adding chemical modifications like branches in the strand. A further insight into the concept of chemically unmodified and modified DNA nanostructures is given in the next chapter.

9 I. Theoretical Background

I.4. Chemically unmodified DNA Nanostructures

Monomeric DNA building units can assemble into complex DNA nanoobjects. The controlled self-assembly of DNA subunits results in the formation of one-dimensional templates and precise shapes of two-dimensional lattices and three-dimensional nanostructures. Seeman, a pioneer in the field of DNA nanotechnology, proposed DNA hybridization as an artificial strategy for complex nanoscopic scale assembly by generating periodic lattices from DNA.[34] Branched DNA structures occur in living systems as unstable intermediates in the replication and recombination process called the Holliday-junction (Fig. I-8).[95] The local symmetry at the junction causes a branch-migration avoidable by an appropriate sequence design. The first constructs were 4-way-junction analogues of the Holliday-junction structures. The fixed junction was an important starting point in the development of DNA nanoarchitectonics.

Fig. I-8: Holliday junctions. A: Symmetry at the vicinity of the branch point leads to branch migration in the junction. B: Structurally fixed 4-way-junction with minimized sequence symmetry. (Modified image taken from reference [96]) Motifs generated include 3-arm,[97,98] 4-arm,[99–101] 5- and 6- arm[102] and 8- and 12 arm[103] junctions (Fig. I-9). This type of structures may be seen as the simplest designs in DNA nanoconstruction. The important goal of sequence design lies in the minimization of symmetry leading to better assembly control. Issues arise in a case like the 12-arm junction, where it is not possible to flank the branch point with different base pairs. AT and GC pairs alternate symmetrically around the junction instead.[104] Fig. I-9: Close-up of a 12 arm junction branch point (Image taken from reference [103])

10 I. Theoretical Background

Two double-strands can combine along their helical axis by using so called sticky ends (Fig. I-10) resulting in an elongated yet still one-dimensional double-strand.[105,106] The structure still coheres to the B-DNA form regardless of the existing gaps between the stick end regions.[107]

Fig. I-10: Sticky end hybridization requires double-strands with single-strand overhangs. Complementary overhangs hybridize to a new combined double-strand.

A set of 4-way-junctions with complementary sticky-ends may assemble into a two- dimensional crystalline lattice (Fig. I-11).[30] Enzymatic ligation closes the overlap gaps that turns the complex into a covalently bonded structure. Unpaired sticky ends at the edges allow for more units to be added later and further extend the lattice.

Fig. I-11: A: Assembly of a two-dimensional lattice consisting of four Holliday junctions with sticky end overhangs. B: Six interlocked cyclic single-strands green, yellow, pink, red blue, and dark blue form a cube-like object. (Modified image taken from reference [30])

Ligation increases stability in the formation of the first three-dimensional structures like cubes. Six cyclic intermeshed single-stands hybridize twice to their four neighbours, thereby forming double-stranded edges. A truncated octahedron made of 14 oligonucleotides was also assembled correspondingly.[108] Other examples of constructional DNA self-assembly can be found in the literature including contributions by Mao and Turberfield.[38,109–116]

Inspired by the Japanese paper folding technique Rothemund reported a versatile computational folding method of large DNA single-strands into precise nanostructures.[117] The concept of scaffolded DNA origami heavily expanded the capabilities of structural DNA

11 I. Theoretical Background nanotechnology going beyond Seemans crystalline lattices made of repeating sub units (Fig. I-12). One long single-stranded viral DNA (M13mp18; 7249 nucleotides) serves as a scaffold strand and is folded into a flat array of antiparallel helices by over 200 shorter complementary strands described as staple strands. These are usually 16- or 32-mers, form stable crossover motifs and may bridge more than two separate scaffold strand segments. The flexibility in the design of folding patterns leads to a vast variety of shapes like squares, stars, triangles et cetera.

Fig. I-12: A: Theoretical folding pattern of a long DNA stand (black) with several smaller staple strands (coloured). Some staple strands divide in up to five different helix sections. In box: 4-T hairpin loop used for blunt-end capping; B: Exemplary origami shapes. The colouring indicates the base-pair index with orange being the first base and purple being the last one in the scaffold. The middle row of AFM images are 165x165 nm in scale. (Image taken from reference [117]) 4-T hairpin loops may cap blunt-ended helices, which reduces the amount of stacking between several shape units. The idea of folding a large DNA single-strand with smaller single-strands resembles an earlier concept of Shih and co-workers in which a ~1.7 kb DNA strand and five oligomers assembled into an octahedral shape via crossover motifs.[37,118] The DNA origami technique results in constructs with much larger diameters around 100 nm and a spatial resolution of 6 nm making it highly addressable for atomic force microscopy. Additionally, the assembly method works with unpurified starting materials. Douglas et al. extended the origami technique to build three- dimensional objects (Fig. I- 13).[119] A set of three double Fig. I-13: Hierarchical assembly of three origami monomer tiles into an icosahedron. Cryo-EM images on the right (Modified image taken from triangular shapes with ten reference [119])

12 I. Theoretical Background half-struts each serve as tiles for an icosahedral wireframe. Each strut is a nanotube made out of six helices. In a hierarchical fashion the double triangular origami tiles form first and subsequently assemble at specific positions at the half-struts into an icosahedron with diameters of over 100 nm. A more recent example is a DNA origami three-point- star motif by Iinuma et al.[120] stabilized with three double- strand struts resembling a tripod (Fig. I-14). The angle Fig. I-14: Struts stabilize a tripod motif. Sticky ends “click” a number of units into between the legs is tuneable polyhedral structures. AFM- and cryo-EM images show a cube and a tetrahedron motif. (Modified image taken from reference [120]) via the strut length. Six short DNA double helices at the vertex mask blunt duplex ends to avoid aggregation. Sticky ends at the end of every leg connect tiles together in a hierarchical assembly into polyhedral structures comprising tetrahedrons and cubes with edge widths of 100 nm larger than previous constructs.

Curved single-layer 3D origami further extended the structural repertoire with shapes that resemble gears,[121] flasks[122] and spheres with a gridiron like pattern.[123]

This chapter only focused on static structural DNA nanotechnology. Literature provides a more comprehensive look into this topic.[104,124–131]

I.5. Chemically modified DNA Nanostructures

The design philosophy of DNA nanoconstruction is based on sequence-controlled assemblies into higher order hybridization motifs. Chemically unmodified DNA nanostructures rely only on the formation of hydrogen bridge bonds and the specificity of Watson-Crick base pairing as major forces of structural assembly. The DNA backbone is completely unaltered in all cases, which might lead to structural limitations like the fixed persistence length. Chemical modifications at the backbone or at the nucleobases can deliberately change the chemical or structural behaviour of a construct. They can connect several native oligonucleotides together and modify the structural information of a design by acting as flexible hinges. They allow for decreased sizes of DNA nanoconstructs compared to DNA origami based designs if necessary.

13 I. Theoretical Background

Chemical modifiers are introduced in the automated oligonucleotide synthesis as protected phosphoramidite building blocks or, alternatively, the oligonucleotide synthesis starts with modified solid supports and subsequent oligonucleotide syntheses on top.

Fig. I-15: Schematic linkage of two or three DNA strands with variable angles and strand orientations. Linking modifiers usually connect two, three of four single strands together and can introduce a fixed symmetry (Fig. I-15). They also allow to change the orientation of the strands. For instance, two connected strands can both end with 3’-positions.

The upcoming described linking methods are sorted by the number of linked oligonucleotides to illustrate the dimension of different branching strategies.

Different design philosophies for connecting two strands exist[96,132,133] including the addition of flexible spacers at the backbone (Fig. I-16; e1, e2),[134] rigid linkers which induce fixed spatial angles (e3, e4)[39,135,136] and metallo-organic coordination complex-based linking (e5, e6).[137–141] Conceivable motifs based on two connected strands are for example circular shapes[136] that, moreover, can stack into prismatic shapes by adding linear oligonucleotides as connectors.[39,135]

A selection of chemical modifiers that can link up to three oligonucleotides, in the context of this work termed trislinkers, are shown in the next figure. Linkage of three oligonucleotides is the basis of this work and therefore of most importance in this context. They allow for more accessible designs of polyhedral objects than bislinker-based designs.

Examples e7[142–146] and e8[147–150] (Fig. I-17) are part of a distinct group of linkers based on nucleotide derivates (branching monomers) and either introduce the branch at a modified nucleobase or at the 2-position of the sugar backbone.

14 I. Theoretical Background

Fig. I-16: Flexible (e1, e2), rigid (e3, e4) and complex-based (e5, e6) DNA bislinking designs; Names refer to precursor compounds.

Fig. I-17: Examples of nucleotide-based linkers for linking three oligonucleotides. Names refer to precursor compounds.

15 I. Theoretical Background

The following four examples introduce non-nucleoside DNA branching units. The Ψ-linker (Fig. I-18) by Scheffler et al.[151,152] was part of a self-assembly experiment based on trisoligonucleotidyls.[153,154] In the automated synthesis the triple DMT-protected linker amidite attaches on a CPG solid support with a nucleotide as a spacer (Fig. I-18; A), which is followed by the oligonucleotide synthesis of three identical self-complementary sequences. Cleavage with ammonia does not affect the amido functionality and cleaves the starter nucleotide at the solid support instead, leaving this nucleotide as a residue in the final product.

Fig. I-18: A: Branch synthesis strategy. B; Agarose gel show discrete hybrids with equal numbers of units being more predominant; Two strand lengths D and G show a slight change in mobility. C: Assembly of two units into nano-acetylene and nano-cyclobutadiene (Modified image taken from reference [151]) Analysis of hybridization products in native agarose gels revealed defined bands instead of polymeric networks (Fig. I-18; B). This was achieved by a special hybridization protocol, which included a fast cooling step after denaturation and prior to reaching the annealing- temperature. Hybrids with an even number of building blocks are favoured because in those cases all arms are paired. In accordance to the isolobal principle the two most distinct products were named after resembling hydrocarbons (Fig. I-18; C). The trisoligonucleotidyl dimer was coined nano-acetylene and the tetramer nano-cyclobutadiene.

[152,155] The Vas-linker by Dorenbeck, is based on a (2S)-pentan-1,2,5-triol and differs in direct comparison to other non-nucleoside trislinkers with its asymmetrical design and the presence of a stereocenter (Fig. I-19). A symmetrical analogue, the non-orthogonally

16 I. Theoretical Background protected V-linker, predates this linker in trisoligonucleotidyl-chemistry.[156,157] The protected phosphoramidite attaches to the solid support, which is functionalized with a nucleotide. The selective deprotection of DMT allows the oligonucleotide synthesis on one position of the linker. After the selective deprotection of AOC a second linker amidite attaches on the other position of the first linker. Stepwise deprotection of both protection groups allows the synthesis of two more strands with different sequence designs. This is an early example of orthogonal protection group deployment (DMT + AOC) in linker systems allowing for full sequence control on a branched DNA design.

Fig. I-19: A: Abbreviated trisoligonucleotidyl. Synthesis. It contains a deprotection step and strand synthesis; B: Sequence- addressed self-assembly of four trisoligonucleotidyls into a tetrahedral shape. Each colour represents a unique complementary sequence. (Modified image taken from reference [155]) The V-shaped linker is bendable and a flexible component in nanoconstruction. A set of four trisoligonucleotidyls with this flexible linking modifier can shape into a tetrahedron in a sequence-addressed self-assembly (Fig. I-19; B). Every edge of the structure contains a unique complementary DNA sequence, thus the set of buildings blocks can only self- organize in one specific way. Every one of the four vertices is the trislinker.

Zimmermann et al. introduced an aromatic C3h-symmetrical linker component with flexible propyl-arms (Fig. I-20).[158,159] The synthesis starts with the first oligonucleotide strand in the reverse order 5’-3’ and introduces the trislinker afterwards. Consecutive strand synthesis of the second and third arm continues in 3’-5’-direction using orthogonal deprotection steps (Fig. I-20; B). This strategy omits the use of nucleotide anchors that remain as a residue in the final product as in the aforementioned strategy for trisoligonucleotidyls by Scheffler and Dorenbeck resulting in so-called trisoligonucleotides.

17 I. Theoretical Background

Fig. I-20: A: 1,3,5-Trishydroxypropylbenzene core, orthogonal protection group and amidite synthesis; B: Trisoligonucleotide synthesis: 1) First strand synthesis in 5’-3’-direction. 2) Trislinker amidite coupling and subsequent detritylation. 3) Synthesis of second strand. 4) Pd-catalysed AOC cleavage and final stand synthesis. Release from solid support with ammonia. C: Native agarose gel (2%) of the full dodecahedron assembly and partial subsets. Digestion experiment of single-strands with mung bean endonuclease. (Modified image taken from reference [158]) A set of twenty unique trisoligonucleotide modules were part of a self-assembly into a dodecahedral design with 12 pentagonal faces, 30 vertices and 20 edges (Fig. I-20; C). The full dodecahedron set and partial subsets were assembled and studied on native agarose gel electrophoresis. The full dodecahedron did not degrade in presence of mung bean endonuclease that preferably digests single-strands. All strands were therefore hybridized, which proofed the structural integrity of a fully closed nanocage. Atom force microscopic studies confirmed an expected diameter of ~20 nanometres.

Tumpane et al. (Fig. I-21)[160] offered a structurally closely related to the previous trislinker and accessible through a Sonogashira coupling step. A linker by Kuroda et al. used the same reaction.[161]

Fig. I-21: A: Protected alkinylbenzene linker. B: Assembly into a circular motif CT and a linear motif LT by altering one strand. C: Gel electrophoresis shows structure-dependent mobility. (Modified image taken from reference [160])

18 I. Theoretical Background

The stiffness of the alkyne moiety is compensated by the flexibility of the attached alkylene chain. The orthogonal protection-group strategy in this case involves the Lev- and DMT- groups. This linker served in the addressable construction of a flat hexagonal shape with protruding arms at the vertices. These arms were additionally complemented by single- strands. The sequence-addressed specific assembly is demonstrated by altering one sequence to be non-complementary in the set. This prevents the cyclization resulting in a linear structure with six branches. Gel-electrophoresis show how structural changes can affect properties. Both shapes have the same mass yet run at different speeds through the gel.

Further strategies in linking three oligonucleotides involve template-controlled triple crosslinking as described by Nielsen et al.,[162] Gothelf et al.,[163] and Eckardt et al..[164,165]

Fig. I-22: Linking modifiers connecting four strands (e9-e11) and six strands (e12).

19 I. Theoretical Background

Linkage of four or more oligonucleotides also exists, but orthogonal protection strategies are not available as of yet. A brief overview of different strategies is given for reasons of completion (Fig. I-22). Maleimide-modified porphyrins can connect up to four oligonucleotides modified with cystamine-functionalities.[166] Both connect in a click-type reaction after activation with dithiothreitol (e9; Endo et al.[167–171]). In a similar fashion Clavé et al.[172] connected an azide-functionalized porphyrin to four single-strands with alkyne- termini in a copper-catalysed Huisgen cycloaddition. Ueno et al.[173] describe with example e10 a strategy based on pentaerythritol as the linking core and Stewart et al.[174] demonstrated with example e11 a four-way-branch with a substituted nickel(II)-cyclam complex. The nickel-ion has a rigidifying effect on the complex, which stabilizes a tetrahedral orientation of all four strands. With example e12[174,175] the same authors synthesized six-arm DNA-

2+ [Ru(bpy)3] -conjugates and successfully dimerized two complementary units. So far, it is one the few examples of six-arm-branches in oligonucleotide nanoconstruction. Full sequence-addressed assembly of unique complementary sequences becomes increasingly more difficult with the introduction of more oligonucleotides and the limitations of orthogonal protection group chemistry. Additionally, the presence of unique sequences introduces chirality at the complex due to different possible orientations of the ligands hampering distinct assemblies into precise nanoconstructs.

Structural DNA nanotechnology shows potential applications in areas like directed material assembly, structural biology, biocatalysis, DNA computing, nanorobotics and cargo transport possible by spatial addressability and multivalent properties.[124] Several applications for DNA wireframe and tensegrity nanostructures are reported in the literature:[125] For instance, two-dimensional lattices are used as periodic grids that allow to position molecules and particles in a periodic fashion. Examples include aptamer arrays fixed on a platform for sensitive and multiplexed biosensing,[176,177] the positioning of proteins to facilitate cryo-electron microscopy[178] or the use of origami frames housing singular DNA duplexes to study binding effects and efficiencies of enzymes like EcoRI methyltransferase on a per molecule basis.[179] Three-dimensional arrangements of gold-nanoparticles of different sizes at the vertices of DNA tetrahedrons enable the building of chiral assemblies.[180] The encapsulation of cargo is also reported, like gold nanoparticles into DNA icosahedra,[181] as well as the controlled release of encapsulated cargo, like gold nanoparticles from triangular DNA nanotubes[182] or the temperature-controlled release of the enzyme horseradish peroxidase in a truncated octahedron.[183] The edges of this cage are composed of twelve double-strands connected by truncated corners consisting of short

20 I. Theoretical Background single-stranded thymidine linkers and one different longer single-strand capable of folding into hairpin structures. This design imposes a temperature-controlled conformational change, in which the hairpin is unstable at 37 °C, but stable at 4 °C. The absence of the hairpin leads to a more flexible structure and allows for entrance of the enzyme into the cage. A low temperature of 4 °C ensures the cargo to stay retained inside the cage. The enzyme is still catalytically active inside the cage.

I.6. Static DNA Nanostructures for Drug Delivery

Unlike single- and double-stranded oligonucleotides, DNA nanostructures exhibit good water solubility, biocompatibility and high permeability into mammalian cells, which makes them very attractive for medicinal applications like in cancer treatment.[184,185] The defined sizes and shapes of DNA nanostructures allow for predictable loading patterns and binding efficiencies.

Turberfield and co-workers assembled four oligonucleotides into a tetrahedral shape in presence of cytochrome c, which was conjugated at a 5’-position of one of the strands (Fig. I-23; A).[186] By sequence-design the protein was pointing to the inside of the cavity. The same tetrahedral shape was applied by Anderson and co-workers[187] to successfully transport small interfering RNA in vivo for gene suppression in mouse model studies (Fig. I-23; B) and by Liu et al.[188] as a vaccine platform that contained an antigen and CpG oligonucleotides as an immunostimulating adjuvant. (Fig. I-23; C) The bound complex of antigen, adjuvant and DNA induced stronger and longer-lasting antibody responses in tests with immunized mice than unbound mixtures of antigen and CpG oligonucleotides alone.

Though static DNA nanostructures are not as sophisticated as dynamic theranostic[189,190] devices such as switchable origami nanobots[191,192] and boxes,[193,194] they can serve as masking agents, bioactivity enhancers and, theoretically, as drug stabilizers. Knudsen et al. demonstrated a temperature-controlled encapsulation and release of the enzyme horseradish peroxidase. Ding and co-workers loaded triangular-shaped origami with the anthracycline doxorubicin via intercalation (Fig. I-23; D).[195] In cancer treatment it inhibits the topoisomerase II and effectively stops the process of replication.[196,197] An uptake of loaded origami into human breast adenocarcinoma cancer cells (MCF7) was observed. Cargo was released through partial digestion of the origami inside the cell. The loaded origami was even able to bypass doxorubicin-resistant MCF7 cancer cells, unlike unbound doxorubicin,

21 I. Theoretical Background and showed cytotoxic effects in vitro.[198] This was possible by masking doxorubicin inside the origami carrier and thereby circumventing drug resistance.

Fig. I-23: A: The tetrahedral shape comprises four oligonucleotides. Black arrow indicates the protein attachment site at one of the 5’-ends. Molecular model of the tetrahedron packed with cytochrome c antigen. B: Tetrahedron containing six siRNA duplexes. The tetrahedrons binds with folic acid tags (grey) to folate receptors of the tumor cell surface. C: Tetrahedron carrying a streptavidin antigen (red) and CpG oligonucleotide adjuvants (yellow ribbons). They bind specifically to B cells, which then bind to T cells to induce antibody production. D: Triangular origami loaded after incubation with doxorubicin via intercalation. Uptake into cancer cells and release through partial origami digestion. (Modified image taken from references [186], [187], [188], [195])

22 II. Motivation

II. Aim of Work

Over the last two decades the von Kiedrowski group explored capable linker designs to be used in the field of trisoligonucleotide-based nanoconstruction. Their structural and functional potential enabled subsequent constructs like Schefflers’ motifs of nano-acetylene and nano-cyclobutadiene[151] or Zimmermanns’ dodecahedral design.[158] The 1,3,5- tris(hydroxypropyl)benzene linker by Zimmermann and Cebulla[199,200] has proven to be a very potent linker-design so far, which iteratively streamlined the orthogonal synthesis approach

[155] based on Dorenbecks Vas-linker. However, the chemical linkers themselves are not commercially available and, so far, require a lengthy and laborious manual synthesis, which is a bottleneck in succeeding trisoligonucleotide-based assembly studies. An optimized linker design is needed for that matter.

Previous work also included the preparation of the shorter alkylene chained linkers 1,3,5- tris(hydroxymethyl)benzene (Anscheidt,[201] Schorr[202]) and 1,3,5-tris(hydroxyethyl)benzene (Castonguay[203]). The alkylene chains are needed for flexibility at the joint. All three arms in a trisoligonucleotide set need to bend in order to fold into a polyhedral shape. The influence of the alkyl chain length in nanoconstruction in terms of stability is best tested on a tetrahedral scaffold because of being the smallest possible object of three-dimensional topology. This implies in the case of tetrahedral trisoligonucleotide scaffolds that all three arms bend with a higher constrain then in other polyhedra like cubes or icosahedra. If a linker forms stable tetrahedra under those constrained conditions it is also suitable for less constrained higher polyhedral motifs. Though initial studies conducted by Schorr exist for the case of the methylene chain, there is no full systematic examination of all three linkers available. The question is how the alkylene chain length (C1-C3) at the linker influences the stability of trisoligonucleotide- based tetrahedral scaffolds.

23 II. Motivation

Schefflers assembly motifs[151] were based on a trislinker with three identical arms (“XXX”), whereas the orthogonal protection group approach allowed for full sequence-control with three unique sequences (“XYZ”). Unexplored is a gap between those two sequence-designs that is a trislinker with two identical and one unique arm (“XXY”). Theoretically, a set of four trisoligonucleotides of this setup could still form a tetrahedral scaffold depending on an appropriate sequence design. It is not clear if either a discrete object or a dendrimeric

network will arise since complementary twin arms might chain up and polymerize.

Another area of concern in trisoligonucleotide chemistry is purification. Preparative polyacrylamide gel electrophoresis was previously established as a purification method for mutant removal.[152] Preparative liquid chromatography in contrast would allow for an automated and faster purification process. Unfortunately, trisoligonucleotides cannot satisfyingly be separated from mutants via HPLC and other DMT-on purification methods.[155] A potential alternative is the strategy of fluorous affinity purification. The separation principle is based on the specific fluorophilic interaction between fluorous-tagged compounds and a fluorous stationary phase. The idea is to apply fluorous tags to trisoligonucleotides as part of their synthesis and then study the effect on separation via HPLC on fluorous and reverse stationary phases. Conditions need to be found on how to cleave the tag after successful separation.

DNA nanocages show potential use as cargo carriers. One possible means of transport is using the DNA itself via an intercalation mechanism. DNA origami was recently established as a carrier for doxorubicin able to permeate drug-resistant cancer cells and improve cytotoxicity.[195] Trisoligonucleotide-based tetrahedral scaffolds are of much smaller size then origami constructs (~100 bp against ~10 kbp), which could potentially improve cellular permeability even further. It is not clear how the intercalation has an effect on trisoligonucleotide-based tetrahedrons in terms of stability and how the nanoconstructs might influence binding in comparison to linear DNA.

24 III. Results and Discussion

III. Results and Discussion

III.1. Synthesis of C3h-Symmetrical Trislinkers

The synthesis of trislinkers can be divided in two parts: The first part is the synthesis of the achiral linker core itself that contains three alkylhydroxide arms in a C3h-symmetrical orientation. A goal is to keep this part as short as possible to reduce preparation time and to maximize yields. The second part contains the protection group strategy and the phosphoramidite reaction. With three chemically identical hydroxides at the linker core double- and triple-substitutions are unavoidable, which typically reduces the overall yield. The orthogonally protected trislinkers of 1,3,5-tris(hydroxymethyl)benzene and 1,3,5-tris(2- hydroxyethyl)benzene and the Bannwarth phosphitylation reagent were previously synthesized and were therefore readily available. The trislinkers based on 1,3,5-tris(3- hydroxypropyl)benzene and isocyanurate, though, required synthesis.

Fig. III-1: Established pathway to 1,3,5-tris(hydroxymethyl)benzene by Anscheidt and Schorr. A hypothetical linker based on phloroglucinol 00 (1,3,5-trihydroxybenzene), which bears no alkyl chains, was considered too rigid. Derived trisoligonucleotides would most likely be planar and are severely hindered to fold into polyhedral shapes. The trislinker with the shortest alkylene chains is 1,3,5-tris(hydroxymethyl)benzene. The established synthesis by Anscheidt[201] and Schorr[202] is given in Fig. III-1.

25 III. Results and Discussion

The established preparative pathway of 1,3,5-tris(2-hydroxyethyl)benzene (Fig. III-2) was given by Castonguay.[203] Bromine was substituted with a cyanide in dimethylsulfoxide (DMSO) to 07, which was the crucial chain elongation step to turn from methyl- to ethyl chains. Basic treatment with aqueous sodium hydroxide converted the nitrile-groups to acids in 08. The following steps were the acidic esterification to 09 and reduction with lithium alanate to 10.

Fig. III-2: Established pathway to 1,3,5-tris(2-hydroxyethyl)benzene by Castonguay.

Different strategies were investigated in an attempt to improve on the synthesis. One strategy involved a direct chain elongation of mesitylene with formaldehyde. Mesitylene was treated at -50 °C with the Schlosser superbase LICKOR. In presence of an equivalent amount of potassium tert-butoxide a benzylic α-methylation is highly preferred over an aromatic o- methylation.[204] No o-hydroxymethylated product was found in the mixture. However, only traces of the desired trialcohol formed. Ostensibly, a highly charged species like the trilithium mesitylene was not stable enough under the given conditions. Nonetheless, successful reactions with oxiranes were reported, albeit with low yields.[205] Other pathways were considered, but ultimately rejected, like a Wittig reaction[206] with triformylbenzene,[207– 209] the hydroboration of trivinylbenzene[210] or corresponding trimethylsilylenolethers,[211] Suzuki-couplings[212,213] and cobalt-catalysed cyclotrimerizations of 3-butyn-1-ol.[214,215] All of those considerations required preparation of either reactants, reagents or catalysts first, which defeated the purpose of cutting down the overall number of reaction steps.

26 III. Results and Discussion

Additionally, reactants like trivinylbenzene are prone to polymerization and therefore difficult to handle in bulk amounts.

An alternative route was investigated, which starts with the trimerization of E-4-methoxy- 3-buten-2-one 11 to triacetylbenzene 12[216,217] continued by a Wilgerodt-Kindler reaction[218,219] into 08 (Fig. III-3). Older procedures of the trimerization incorporate an in-situ generation of the sodium salt of a 3-buten-2-one with acetone, ethyl formate and sodium ethanolate.[220] This reaction requires dry conditions because of the generation of ethanolate. Dry conditions are not mandatory for the actual trimerization step. A reaction in water is possible by starting directly with the methoxybutenone 11. After synthesis crude product can then be purified by recrystallization in ethanol.

Fig. III-3: Trimerization of E-4-methoxy-3-buten-2-one, followed by a Wilgerodt-Kindler reaction. Esterification and reduction with LiAlH4 gives 1,3,5-Tris(2-hydroxyethyl)benzene.

In the subsequent Wilgerodt-Kindler reaction the acetyl-residues of 12 were converted to 2- thioamides in the presence of morpholine and sulphur. Under acidic conditions and subsequent basic workup the intermediate species was hydrolysed to the tricarbocylic acid 08. Michalski further elaborated this pathway by acidic esterification in methanol and the reduction with lithium alanate to the triple alcohol as part of his bachelor thesis.[221] It is possible to skip the esterification step and directly reduce from the tricarboxylic acid 08 to 10 according to literature,[218] but overall yields are lower (42 % in literature) and chromatography in acetonitrile required very long elution times. An alternative eluent for

27 III. Results and Discussion good separation was not found. This new synthetic route saves one step in comparison to the established pathway and increased the overall yield.

With this linker 12 three separate phosphoramidites bearing two allyloxy carbonyl-groups (19), two dimethoxytrityl-groups (17) and both protection groups (15) were prepared (Fig. III- 4). The first two were specifically prepared for compatibility studies on CPG solid support described in chapter III.2.3.

Fig. III-4: Orthogonal protection group strategy with allyloxycarbonyl chloride (AOC-Cl) and dimethoxytrityl chloride (DMT- Cl) and phosphoramidite reaction of 1,3,5-tris(hydroxyethyl)benzene.

All three hydroxide groups were chemically equivalent, which made selective single- or double hydroxide substitutions difficult resulting in mixtures of single-, double-, and triple substituted products. In order to minimize yields leading to by-product, a shortfall of the protection group chlorides was added. The phosphoramidite reaction used chloro-2-

28 III. Results and Discussion cyanoethyl-N,N-bisisopropylphosphoramidite (“Bannwarth-reagent”) and N,N- diisopropylethylamine (DIPEA) as a base.

The established synthesis of 1,3,5-tris(3-hydroxypropyl)benzene 25 by Zimmermann and Cebulla[199,200] offers two different openings (Fig. III-5). Treatment of 04 with hydrogen bromide in acetic acid gives a clean conversion to 20. Alternatively, an immediate bromination of mesitylene with NBS is possible. The first route gives a cleaner product with a higher overall yield, whereas the second offers a faster single step reaction. The crucial chain elongation step occurs in the subsequent malonic ester synthesis with diethylmalonate and sodium hydride in tetrahydrofuran to yield 21. This was then followed by a saponification to 22 and a decarboxylation to 23 at high temperatures. The last steps are an esterification and a reduction into 25.

Fig. III-5: Established synthesis of 1,3,5-tris(3-hydroxypropyl)benzene by Zimmermann and Cebulla.

A closer look into this strategy reveals redundancy: The chain elongation with malonate was followed by two “deconstruction steps” and the full pathway even repeated the esterification and reduction steps. In an attempt to simplify the preparation a retrosynthetic analysis revealed the possibility of a Heck-type coupling reaction by introducing a double-bond in the

29 III. Results and Discussion chain (Fig. III-6). A suitable aromatic starting material was 1,3,5-tribromobenzene. The appropriate allyl alcohol synthon needed a masked alcohol to not interfere with the catalyst in the Heck reaction. An alternative retrosynthetic analysis suggested the treatment of trilithium mesitylene with oxirane.[205] This was disregarded due to the previously mentioned difficulties in quantitative lithiation of mesitylene.

Fig. III-6: Retrosynthesis of 1,3,5-tris(3-hydroxypropyl)benzene

The Heck coupling reaction[222] shortened the synthesis to three steps in an overall good yield (Fig. III-7). Tribromobenzene 26 was coupled with methacrylate 27 to 28 in the presence of palladium(II)acetate, tri(o-tolyl)phosphine (TOTP) and triethylamine in acetonitrile. Acetonitrile was used in p.a. grade and not further dried prior to synthesis since the presence of water traces in the reaction can improve the catalytic activity.[223] The product 28 recrystallized in dioxane. Hydrogenation over palladium/charcoal led to 24 and was purified via distillation at high temperatures.

Fig. III-7: Synthesis of 1,3,5-tris(3-hydroxypropyl)benzene involving a Heck coupling reaction, an alkyl chain hydrogenation and an ester reduction.

30 III. Results and Discussion

The final step was the reduction with lithium-alanate to 1,3,5-tris(3-hydroxypropyl)benzene 25. A recrystallization procedure in THF/pentane at -78 °C was developed and simplified the purification of bulk amounts.

The synthesis was further shortened by replacing methacrylate with allylbenzylether 31[224] in the Heck reaction (Fig. III-8).

Fig. III-8: Heck coupling of 1,3,5-tribromobenzene and allylbenzylether and single step hydrogenation of alkyl chains and hydrogenolysis of the O-benzyl group into 1,3,5-tris(3-hydroxypropyl)benzene.

The reagent 31[224] was readily prepared in a base mediated Williamson ether synthesis between allyl alcohol 29 (b.p. ~98 °C) and benzyl chloride 30 (b.p. ~180 °C). The reagent was purified by distillation (b.p. ~204 °C). The same reaction with allyl chloride (polymerizes at 45 °C) and benzyl alcohol (b.p. ~206 °C) complicates the separation by distillation primarily due to the similar boiling points of the benzyl alcohol and the reagent 31. The Heck reaction resulted in a noticeable mixture of E- and Z-conformers of 32 indicated by thin layer chromatography in cyclohexane and ethyl acetate. It was not possible to extract all conformers in column chromatography in pure quality. The ligands TOTP and the cheaper triphenylphosphine were compared in similar setups and showed similar results in terms of yield or selectivity. O-benzyl groups were selectively deprotected in a palladium-catalysed hydrogenation.[225] Since this step was already necessary to saturate the alkene chain this

31 III. Results and Discussion leads to a one-step hydrogenation and hydrogenolysis[226] of 32 to the 1,3,5-tris(3- hydroxypropyl)benzene 25. The hydrogenolysis was slower than the hydrogenation and requires overnight reaction times.

The orthogonal protection groups were then introduced, namely the allyloxycarbonyl- and the dimethoxytrityl-group (Fig. III-9). The final step was the phosphitylation with Bannwarth- reagent in DIPEA.

Fig. III-9: Reversed orthogonal protection group strategy with allyloxycarbonylchloride (AOC-Cl) and dimethoxytrityl- chloride (DMT-Cl) and phosphoramidite reaction of 1,3,5-tris(hydroxypropyl)benzene.

The search for alternative linker designs with smaller preparative effort revealed the promising candidate 1,3,5-tris(2-hydroxyethyl)-N,N’,N’’-isocyanurate 36 (Fig. III-10; THEIC). It is based on cyanuric acid and bears three alkyl hydroxide chains. In the industrial context THEIC is, for example, part of crosslinking agents in urethane foams and used as a flame- retardant additive in polymers.[227] Solubility of THEIC in pyridine was greatly improved by the addition of dimethylformamide (DMF) serving as a cosolvent. Reactivity with DMT-Cl was improved by an increase of the reaction temperature to 60 °C. A single- (37) and a double- (40) DMT-protected linker was prepared. The former was orthogonally protected with AOC- Cl (38). Easy detection on thin layer chromatography made the introduction of the dimethoxytrityl as the first protection-group preferable. Both underwent the same

32 III. Results and Discussion phosphitylation reaction to the final trislinkers 39 and 41. 41 was studied in trisoligonucleotides with semi-addressable sequence patterns (see chapter III.4).

Fig. III-10: Synthesis of two isocyanurate-based linker amidites.

33 III. Results and Discussion

III.2. Oligonucleotide Synthesis

III.2.1. Trisoligonucleotide Synthesis Protocol

On the DNA/RNA synthesizer Gene Assembler Plus (Pharmacia Biotech) the reverse 5’-3’ oligonucleotide arm is built on the custom reverse polystyrene solid support based on Primer Support 200 Amino. The scale is 1.3 µmol and standard protocols for detritylation, capping and trivalent phosphorous oxidation are applied. 5-benzylmercaptotetrazole activates all phosphoramidites. The protocol is summarized in the next table (Tab. III-1).

Step Coupling time Arm 1 (Reverse 5‘-3‘-15mer DNA-synthesis) First and second 5‘-amidite (0.1 M ACN) 2x 5 min

Further 5‘-amidites (0.1 M ACN) 5 min Trityl-on Arm 2 (3‘-5‘-15mer DNA-synthesis)

C3h-Linker (0.2 M in ACN) 2x 5 min

3‘-amidites (0.1 M ACN) 1.5 min* Trityl-on

Arm 3 (3‘-5‘-15mer DNA-synthesis) AOC-deprotection (Flow: 0.5 mL/min) 15 min

First 3‘-amidite after AOC-deprotection (0.1 M ACN) 3x 15 min 3‘-amidites (0.1 M ACN) 3x 2 min Trityl-off

Tab. III-1: Automated synthesis protocol for trisoligonucleotides. * = 2 x 2 min for simultaneous synthesis of the second and third arm using double DMT-protected linkers. The first few nucleoside 5’-amidites are coupled twice for 5 minutes each and every further amidite once. The step after the synthesis of the first arm is divided in two parts: The detritylation with 3 % dichloroacetic acid in 1,2-dichloroethane and the subsequent introduction of the trislinker amidite as a 0.2 M solution in acetonitrile (a). It couples twice for 5 minutes each and is then detritylated (b). Afterwards, the sequence direction changes to 3’-5’ and requires 3’-amidites following standard procedures (c). A freshly prepared AOC- deprotection mixture is then added manually to the system with a flow rate of 0.5 mL per minute (d). The mixture typically contains 17.1 mg 1,2-bis(diphenylphosphino)ethane, 24.7 mg

34 III. Results and Discussion bis(dibenzylideneacetone)palladium(0) and 10.7 µL pyrrolidine in 10 mL of ACN. The first 3’- amidite to synthesize the third arm (e) necessitates longer coupling times for optimal yields. Trityl-on purification methods are insufficient to purify trisoligonucleotides therefore the automation ends in a trityl-off step. In the final step manual treatment with concentrated ammonia (33 %) liberates the trisoligonucleotide from solid support (f).

The following figure (Fig. III-11) shows the general strategy in orthogonal trisoligonucleotide synthesis with an introduced C3h-symmetrical trislinker.

Fig. III-11: General concept of orthogonal trisoligonucleotide synthesis: After 5’-3’ oligonucleotide synthesis of strand 1 trislinker amidite is introduced (a). Detritylation at the linker with dichloroacetic acid (b) and synthesis of second strand at newly exposed hydroxide with acetate-capping (c). Deprotection of AOC with a Pd(0) catalyst (d), synthesis of third strand (e) and post-synthetic deblocking with concentrated ammonia into final trisoligonucleotide (f).

Fig. III-12: General concept of semi-addressable trisoligonucleotide synthesis: After 5’-3’ oligonucleotide synthesis of strand 1 the double O-DMT protected trislinker amidite is introduced (a). Detritylation at both DMT positions at the linker with dichloroacetic acid (b). Simultaneous synthesis of two identical arms (c). Post-synthetic deblocking with concentrated ammonia releases the trisoligonucleotide (f).

35 III. Results and Discussion

A semi-orthogonal trisoligonucleotide synthesis introduced double-DMT protected C3h- symmetrical trislinkers (Fig. III-12). This allowed for a simultaneous synthesis of the second and third arm, sharing an identical sequence. 3’-nucleoside phosphoramidites were coupled twice for two minutes in order to improve coupling efficiencies in the parallel synthesis of two arms.

Fig. III-13: General chemical structure of a trisoligonucleotide with an isocyanurate linker core.

36 III. Results and Discussion

III.2.2. Phosphoramidite Oligonucleotide Synthesis

The chemical synthesis of oligonucleotides historically includes the H-phosphonate-,[228,229] the phosphodiester-,[230] the phosphotriester-[231,232] and the phosphite triester[233,234] methods and evolved into the today most commonly used phosphoramidite approach.[235,236] Previous methods were originally conducted in solution and were later adapted for solid support[237– 240] inspired by Merrifields’ solid phase synthesis of peptides.[241] The preparation of up to 200- mers are possible this way depending on the porosity of the chosen solid support. The growing oligonucleotide sequence stays immobilized throughout the entire synthesis cycle, which simplifies purification procedures and allows for full process automation. Additionally, the ability to use high excesses of reagents lead to increased yields after each step. The solubility of an oligonucleotide in organic solutions decreases with its length due to the added polarity of the phosphate ester groups. This effect is circumvented with the immobilization of the product. Typical solid supports are macroporous polystyrene (MPPS) or controlled pore glass (CPG) of defined pore sizes between 500 and 3000 Å. MPPS is crosslinked with divinylbenzene and aminomethylated for further functionalization. CPG is treated with (3-aminopropyl)triethoxysilane to give aminopropyl CPG and is further functionalized to give long chain alkyl amines (LCAA-CPG). It serves as a spacer and anchor for typically a succinyl linker on which the first 5’-OH- and nucleobase-protected nucleoside is already attached to the so called starter nucleoside.

The standard synthesis goes from 3’ to 5’ direction. The synthesis cycle of the phosphoramidite method is shown in Fig. III-14 and starts in step 1 with the removal of the 5'-O-DMT protecting group using 3 % dichloroacetic acid or 2 % trichloroacetic acid. The orange-red coloured DMT cation is flushed and allows for on-line photometric monitoring of coupling efficiencies. (Conducting detritylation for an extended period of time leads to depurination, which reduces the yield.) In step 2 the nucleoside phosphoramidite is introduced typically as a 0.2 M solution in acetonitrile and activated using 4,5- dicyanoimidazole[242] or 5-benzylmercaptotetrazole.[243,244] Step 3 is the coupling of the now free 5'-hydroxyl-position (B1‘) to the activated phosphoramidite (B2‘). All nucleotide phosphoramidites bearing an exocyclic amino functionality at the heterobases as well as the backbone are protected to avoid unwanted side-reactions. Coupling times are usually fast with only 2 minutes for each nucleotide amidite. In step 4 all unreacted nucleotides (B1‘) are capped, using acetic anhydride with DMAP and 2,6-lutidine. The process of capping blocks unreacted 5’-OH groups and minimizes the risk of generating deletion mutants.

37 III. Results and Discussion

Fig. III-14: Synthetic cycle of the phosphoramidite method.

The reaction continues in step 5 with the oxidation of the trivalent phosphorous to the pentavalent one using an aqueous iodine solution and the base collidine. This step is necessary given the instability of the trivalent phosphorous in subsequent steps. It is possible to interchange step 4 and 5 depending on the used protocol. The cycle repeats until the desired oligonucleotide length is reached. Cleavage of the oligonucleotide from the succinyl linker to the CPG resin, the cyanoethyl-groups at the phosphorous and the deprotection of protected nucleoside functional groups is done using concentrated ammonium hydroxide. The final 5’-DMT-group is either left intact (step 6a) or removed (step 6b) before cleavage. The increased mass and lipophilicity of DMT improves separation of linear <150mer oligonucleotides in RP-HPLC or cartridge based methods.[245] 2 % trifluoroacetic acid removes the group post-synthetically.

38 III. Results and Discussion

Fig. III-15: Nucleoside phosphoramidites on solid support for normal 3’-5’- synthesis orientation and reverse 5’-3’. Protection of exocyclic amino function with benzoyl- (dA’, dC’) and isobutyryl- (dG’) groups. 2-cyanoethyl groups protect the free oxygen at the phosphoramidite.

Synthesis in 5’-3’ direction requires the reversely configured nucleoside phosphoramidites and polystyrene solid support (Fig. III-15). They are both commercially available at a much higher price than the corresponding 3’-5’ reagents. This is explained in the increased effort in preparation. The selective DMT-protection of the secondary 3’-OH is aggravated by the presence of the more reactive primary 5’-OH. In reversed solid supports the starter nucleoside connects at their 5’-OH group as succinic esters. The 3‘-OH end is DMT-protected instead. The DMT-deprotection of the less reactive secondary alcohol takes longer than on a primary alcohol. The synthesis in reverse direction is therefore slower in comparison to the standard direction at slightly lower coupling efficiencies.

III.2.3. Sequence Design

With the software tool DNASequenceGenerator (1.01b) by Feldkamp et al.[246] a pool of one hundred 15mer sequences was created for a 1.25 µM solution in 110 mM NaCl with a set melting point range of TM = 58.3-58.4 °C at a neutral pH. The program uses SantaLucia parameters[247] and the Nearest Neighbour-model for duplex energy calculations. The first and the final nucleotide in the sequences were G and C to reduce fraying at the blunt ends.

39 III. Results and Discussion

Six unique sequences were picked out of the pool as listed below with their complementary counterparts (marked with an asterisk).

No. Sequence 5’-3’ GC % Tm [°C]

11. CGC TTA TGA GTC CTC 0.53 58.4 11* GAG GAC TCA TAA GCG 12. GAA TGC GAG AGG TAG 0.53 58.4 12* CTA CCT CTC GCA TTC 16. GGT GTG TTA GGT CGG 0.60 58.4 16* CCG ACC TAA CAC ACC 18. CGA AGC ATA GAC TCC 0.53 58.4 18* GGA GTC TAT GCT TCG 73. CCA GTG TCC TTC CTC 0.60 58.4 73* GAG GAA GGA CAC TGG 84. CAG TCC GCT CTA ATC 0.53 58.4 84* GAT TAG AGC GGA CTG Tab. III-2: Six chosen 15mer sequences out of 100. Full sequence pool in appendix. Numbers with asterisk indicate derived complementary sequences to the same number.

Sequences with more than two G nucleotides in a row were avoided to minimize the risk of G-quadruplex formations. The selection criteria of sequences include homogeneity of theoretical melting points and minimization of hairpin- and palindrome-motifs. The energies of self-pair dimers, mismatch pair dimers and of hairpins were calculated with PrimerSelect (3.11; DNASTAR). The program used the Nearest-Neighbour model with the older Breslauer parameter set.[248] [c = 1.25 µM; c(Salt): 110 mM] The calculated free energies of the duplexes showed values between 27-28.2 kcal/mol and were more than three times higher compared to the most stable mismatch energy found with -8.3 kcal/mol. The calculations for hairpins indicated no thermodynamic stability. Data are available in the appendix.

The sequences were part of a trisoligonucleotide-based tetrahedral design. A tetrahedron has four triangular faces, four vertices and six edges. A set of four complementary trisoligonucleotides (T.1-T.4) form a tetrahedral shape as described by Dorenbeck [155] in which the four trislinkers assume the role of the vertices and the double helices correspond to the six edges. Schlegel-diagrams help in the assignment of sequences to the tetrahedral grid (Fig. III-16).

40 III. Results and Discussion

Fig. III-16: A: Schematic trisoligonucleotide-based tetrahedral shape; B: Schlegel-diagram of a flattened tetrahedron with all arranged complementary sequences at display.

Four different trislinkers shared the same orthogonal sequence design with three unique sequences (XYZ), namely 1,3,5-tris(hydroxymethyl)benzene T1, 1,3,5- tris(hydroxyethyl)benzene T2, 1,3,5-tris(hydroxyethyl)benzene T3 and THEIC TN. They were part of assembly studies that involve the influence of the chain length in trisoligonucleotide base tetrahedral scaffolds. The semi-orthogonal design (XXY) has two out of three strands that shared the same sequence and uses THEIC as a trislinker (TD). It offered an alternative sequence motif design for later hybridization experiments.

Fig. III-17: Variation of trislinkers (T1, T2, T3) and different sequence patterns (TN, TD, TY).

41 III. Results and Discussion

T1, T2, T3, TN Sequences 5’-3’ Vertex (Arm 1, 2, 3)

CGC TTA TGA GTC CTC T.1 GAA TGC GAG AGG TAG GGT GTG TTA GGT CGG CTA CCT CTC GCA TTC T.2 CGA AGC ATA GAC TCC GAG GAA GGA CAC TGG GAG GAC TCA TAA GCG T.3 CCA GTG TCC TTC CTC CAG TCC GCT CTA ATC CCG ACC TAA CAC ACC T.4 GGA GTC TAT GCT TCG GAT TAG AGC GGA CTG

Tab. III-3: Assignment of unique sequences to four XYZ-trisoligonucleotides and calculated extinction coefficients for later reference. Theoretical melting point of a 15mer ≈ 58.4 °C

TD Sequences 5’-3’ Vertex (Arm 1, 2, 3)

CGC TTA TGA GTC CTC TD.1 CGA AGC ATA GAC TCC CGA AGC ATA GAC TCC GAG GAC TCA TAA GCG TD.2 CGA AGC ATA GAC TCC CGA AGC ATA GAC TCC GAA TGC GAG AGG TAG TD.3 GGA GTC TAT GCT TCG GGA GTC TAT GCT TCG CTA CCT CTC GCA TTC TD.4 GGA GTC TAT GCT TCG GGA GTC TAT GCT TCG

Tab. III-4: Assignment of sequences to four XXY-trisoligonucleotides and calculated extinction coefficients for later reference. Theoretical melting point of a 15mer ≈ 58.4 °C

TY was also based on THEIC and consisted of three identical 14mer arms (XXX). The sequence was manually designed and is self-complementary to allow hybridizations with itself.

42 III. Results and Discussion

TY Sequence 5’-3’ Vertex (Arm 1, 2, 3)

TTA ACC GC GGT TAA TY TTA ACC GC GGT TAA TTA ACC GC GGT TAA

Tab. III-5: Sequence design of a self-complementary XXX-trisoligonucleotide. Theoretical melting point of one 14mer is ≈ 52.1 °C.

The sequences of the original trisoligonucleotide pool were additionally used in the synthesis of twelve individual 15mer oligonucleotides as a control group in drug loading experiments (Tab. III-6).

Vertex Sequences 5’-3’ A. CGC TTA TGA GTC CTC A’ GAG GAC TCA TAA GCG B. GAA TGC GAG AGG TAG B’ CTA CCT CTC GCA TTC C. GGT GTG TTA GGT CGG C’ CCG ACC TAA CAC ACC D. CGA AGC ATA GAC TCC D’ GGA GTC TAT GCT TCG E. CCA GTG TCC TTC CTC E’ GAG GAA GGA CAC TGG F. CAG TCC GCT CTA ATC F’ GAT TAG AGC GGA CTG

Tab. III-6: Set of twelve 15mer oligonucleotides sharing the same sequences as the orthogonal XYZ trislinker sets. Listed as complementary pairs. Theoretical melting point of each 15mer ≈ 58.4 °C.

III.2.4. Purification and Quality Control

After synthesis all polynucleic products were treated with concentrated ammonia for deblocking. In the case of linear DMT-on 3’-5’-oligonucleotides the solution was applied to Glen-Pak DNA purification cartridges (Glen Research) using standard procedures[245] for isolation and desalting. Denaturing polyacrylamide gel electrophoresis showed no traces of mutants (Fig. III-18). Fainter bands of higher mobility are contributed to the added bromophenol loading dye in each sample.

43 III. Results and Discussion

Fig. III-18: Quality control of linear 3’-5’-oligonucleotides: 16 % denat. PAGE; 1xTBE; 100 V; 100 min; 1:5 in sat. urea; Gel Loading Buffer: 0.05 % bromophenol blue, 40 % sucrose 0.1 M EDTA pH: 8.0, 0.5 % SDS; Marker dye: 0.25% bromophenol blue (fast), 0.25 % xylene cyanol (slow) and 40 % sucrose.

15mer Calculated Experimental ε (254 nm) Yield -1 -1 Oligonucleotide Mass [g/mol] Mass [m/z] [Lcm mol ] [nmol]

A. 4519.0 4526.7 136392 452 A’ 4626.1 4636.8 168518 306 B. 4706.1 4718.4 182794 420 B’ 4438.9 4442.1 122116 284 C. 4695.1 4699.0 165423 258 C’ 4451.0 4460.0 139257 272 D. 4546.0 4550.3 154242 337 D’ 4599.0 4602.1 150668 301 E. 4454.9 4456.0 122595 347 E’ 4691.1 4699.8 182085 357 F. 4488.0 4490.7 135204 367 F’ 4657.1 4660.6 169706 410

Tab. III-7: Experimental masses of 3’-5’-oligonucleotides via MALDI-TOF-MS (THAP-matrix) and photometrically determined yields.

Masses were measured with MALDI-TOF-MS in a 2,4,6-trihydroxyacetophenone matrix. Besides protonation effects the experimental masses matched the calculated expectations. Discrepancies are explained by varying ion-exchange efficiencies and not by sequences errors. Complementary sequences fully hybridized as shown in the chapter about anthracycline loading (Fig. III-77). Yields of all nucleotide products were determined with UV/VIS-spectroscopy. The necessary extinction coefficients were calculated from of

44 III. Results and Discussion nucleobase increments.[159,249] 3’-5’-oligonucleotides show an expected yield range around 250-450 nmol for a 1.3 µmol scale synthesis.

Increment Nucleotide [molcm-1L-1] C 6541 T 7250 A 13200 G 13679

Tab. III-8: Nucleobase increment chart to calculate extinction coefficients at 254 nm; pH: 7; 25 °C

Branched oligonucleotides are insufficiently purified by RP-HPLC.[155,159,165] For trisoligonucleotides preparative PAGE was deployed instead.

Fig. III-19: Visualisation of trisoligonucleotide purification. Product is the slowest running thick band “3 arms”. Typical conditions for preparative gel electrophoresis: 12 % PAGE; 1x TBE; 450 V; 4 h; 15 °C; Dimensions: 20x20x0.3 cm.

All preparative gels showed a common pattern with two thick bands and several thinner bands in between (Fig. III-19). For purity reasons, only the upper three quarters of the product band were extracted. MALDI-TOF usually resulted in broad signals, which was explained in a poor crystallization behaviour of branched oligonucleotides (Tab. III-9). The relatively small size of the trislinker had negligible absorption effects in trisoligonucleotides and was therefore not taken into account in quantitative analysis. Introduction of the branching trislinkers reduced the average yield range to 60-165 nmol compared to the linear oligonucleotide synthesis. Yields of T1.1, TN.2 and TN.3 are low because of faulty detritylation solutions used in the oligonucleotide synthesis. Coupling of the T1 trislinker appeared to be

45 III. Results and Discussion less efficient in average than the others, which can be explained by its higher rigidity and the more spacious requirements of the corresponding trisoligonucleotide.

Trisoligo- Calculated Experimental ε (254 nm) Yield -1 -1 nucleotide Mass [g/mol] Mass [m/z] [Lcm mol ] [nmol]

T1.1 14320.3 14340.9 484609 26.4

T1.2 14076.1 14121.1 458443 66.2

T1.3 13969.0 14004.4 426317 67.4

T1.4 14107.1 14120.2 459631 81.6

T2.1 14362.4 14364.5 484609 98.6

T2.2 14118.2 14114.7 458443 129.5

T2.3 14011.1 14068.8 426317 165.9

T2.4 14149.2 14182.8 459631 142.8

T3.1 14404.5 14441.6 484609 121.0

T3.2 14160.3 14242.0 458443 160.4

T3.3 14053.2 14100.0 426317 111.5

T3.4 14191.3 14262.9 459631 142.8

TN.1 14413.4 14429.4 484609 208.7

TN.2 14169.2 14192.7 458443 39.9

TN.3 14062.1 14093.0 426317 35.3

TN.4 14200.2 14215.1 459631 102.7

TD.1 14061.1 14121.8 444876 120.4

TD.2 14168.1 14257.8 477002 73.6

TD.3 14354.2 14387.3 484130 163.4

TD.4 14087.0 14163.5 432452 157.4

TY.1 13281.6 13528.8 427380 111.7

Tab. III-9: All trisoligonucleotides: MALDI-TOF-MS in a 3-hydroxypicolinic acid matrix and yields. See appendix for full MALDI-TOF-MS data.

46 III. Results and Discussion

III.2.5. Long-Term Stability Control

All aqueous trisoligonucleotide stock solutions were originally stored at ~8 °C for practical reasons. This allowed to test the relative stability of the building blocks in a long-term study with denaturing polyacrylamide gel electrophoresis (Fig. III-20). The figure displays the age of each building block at the moment of analysis and after additional 12 months.

Fig. III-20: Control of long-term building block stability. 12 % denat. PAGE; 1xTBE; 120 V; 80 min; 1:2 in sat. urea.

All sets showed varying degrees of decomposition as seen in the appearance of oligonucleotide fragments on the gels. Trisoligonucleotides based on the T1 (T1.1-4) were less stable than the ones of T2 and T3. The T2 set showed only very minor decomposition at the trisoligonucleotide T2.1 making it the most stable and most favourable set for long-term experiments. The T3 set showed more noticeable partial decomposition at T3.2 and T3.4, but was otherwise still useable in assembly experiments. Isocyanurate-based trisoligonucleotide sets TN, TD and TY seemed to decompose stronger with increased age. TY was the latest isocyanurate building block showed less decomposition than the older isocyanurates TN and TD. A slow hydrolysis of the alkyl chains is assumed.[250] All building blocks were then stored at ~20 °C and analysed again after one year. Altered storage conditions effectively stopped further noticeable decomposition of all building blocks as seen on the gels. Deeper cooling is especially important for the more easily hydrolysable isocyanurate-based trisoligonucleotides.

47 III. Results and Discussion

III.2.6. Solid Supports for Trisoligonucleotide Synthesis

Synthesis of trisoligonucleotides was performed on reversed polystyrene solid supports. They were previously prepared in the group and used without further modification.[159] The starter nucleosides for this work include dC’, dG’ and dT as part of 5’-O-succinic ester-3’-O- DMT-nucleosides 42a, 43a and 44a. The general synthesis strategy of those is as follows: An excess of DMT-Cl is added to protect both 3’- and 5’-hydroxy groups of the corresponding deoxynucleosides to give the double DMT-protected products. Selective detritylation at the more reactive 5’-terminus occurs with zinc(II)bromide in nitromethane at low temperatures. Treatment with equivalent amounts of succinic anhydride in pyridine under DMAP catalysis yields the 5’-O-succinic ester-3’-O-DMT-nucleosides. 0.25 equivalents of the succinates 42b, 43b and 44b are activated with HBTU and then coupled to the macroporous polystyrene (Primer Support 200 Amino, GE Healthcare; Fig. III-21). Free amino groups of the solid support are then capped with acetic acid anhydride. Loading capacities are typically around 50 µmol/g.

Fig. III-21: Coupling of HBTU-activated 5’-O-succinic ester-3’-O-DMT-deoxynucleosides with macroporous polystyrene solid support MPPS. The loading capacity is ~50 µmol/g solid support. Established synthesis.

Commercially available CPG solid supports are typically not used in the automated synthesis of trisoligonucleotides due to an unfavourable allyl ether side reaction in the AOC- deprotection step leading to low coupling efficiencies of nucleotides at the third arm as originally investigated by Scheffler[152] and Dorenbeck.[155] This was quantified by studying the

48 III. Results and Discussion intensity of detritylation signals of the coupling steps after the AOC-deprotection step, which showed a noticeable drop-off in signal intensity.

Fig. III-22: Mechanism of AOC-deprotection with Pd(0).

At the start of the AOC-deprotection mechanism[251] palladium(0) forms a π-complex with the allyl-moiety. A allyl-Pd(0)-complex by decarboxylation releasing carbon dioxide and liberating the alcoholate (Fig. III-22). In the presence of a nucleophilic scavenger an allyl transfer is possible (1). Absence of the scavenger, on the contrary, highly favours a side reaction between the alcoholate and the π-allylcation into an allyl ether effectively capping the alcoholate for further reactions. Deprotection of the AOC-group for trisoligonucleotides has proven to show best results with bis(dibenzylideneacetone)palladium(0), 1,2- bis(diphenylphosphino)ethane (dppe) as a coligand and pyrrolidine as a nucleophile.[152][251] HPLC analysis and subsequent mass spectrometry via MALDI-TOF by Dorenbeck showed in several test runs a mixture of the desired product and about equal amounts of the corresponding allyl ether. In addition, traces of the free 3’-OH species were also found that are explained by an insufficient conversion. However, no traces of the AOC-protected species were found, whose absence is indicative of a quantitative AOC-deprotection. The switch to polystyrene support significantly improved the formation of the desired three-arm product. It was speculated that somehow the cavities of the CPG solid support retain the π- allylcation close to the reaction side in the sense of a contact ion pair leading to capping of 3’-OH into an allyl ether regardless of the presence of a nucleophile.

This was later confirmed by Zimmermann[159] in a synthesis of trisoligonucleotides with 9mer arms that showed high coupling efficiencies of up to 98.8 % on reversed 500 Å and 1000 Å CPG support following the same protocol described in the last chapter. Unlike the setup of Dorenbeck coupling of the linker and deprotection starts after the synthesis of the first arm. It was assumed that the added distance between the solid support and the AOC-group might have a positive effect on the deprotection. After AOC-deprotection, however, a strong drop in coupling efficiencies was detected regardless of pore size and the altered synthesis protocol, which again revealed the presence of the allyl ether species (Fig. III-23). Reverse

49 III. Results and Discussion polystyrene solid support gave good yields across all coupling steps including the AOC- deprotection thus turning it into the solid support of choice in trisoligonucleotide synthesis.

Fig. III-23: Allyl ether side reaction on CPG solid support.

Previously unexplored was the parallel cleavage of double AOC- and double DMT-protected linkers in the synthesis of trisoligonucleotides on CPG solid support. Cleavage of the doubled amount of allyloxycarbonyl moieties should also double the generation of the allyl cations. It was unclear how the increased concentration has an effect on the allyl ether side reaction. Conversely, DMT is expected to be fully compatible on CPG solid support, since no allyl ether side reaction is possible in this case.

Three linker amidites (Fig. III-24) were introduced in the automated synthesis of a model trisoligonucleotide in order to analyse the outcome on CPG supports as part of the bachelor thesis by Michalski.[221] The linkers contained two allyloxycarbonyl-groups (“AA-Linker”), two dimethoxytrityl-groups (“DD-Linker”) and an orthogonal combination of both (“AD-Linker”).

Fig. III-24: Three linkers with altering protection-group patterns and the chosen model sequence design.

A model sequence was chosen with an equal distribution of all four nucleobases as an approximation of application-related sequences used in trisoligonucleotides. All three arms

50 III. Results and Discussion shared the same sequence for better comparison of detritylation integrals. Automated synthesis on a 3’-DMT-dC CPG solid support (500 Å; ChemGenes) employed the previously mentioned protocols (chapter III.2.1.) for orthogonal (“AD”) and semi-addressable (“AA”; “DD”) trisoligonucleotides. Both protocols are identical with the only difference in 3’- nucleoside coupling times (2x2 min instead of 1x1.5 min) to accommodate for parallel synthesis of two strands.

In the case of the orthogonal AD-linker detritylation integrals over the course of the oligonucleotide synthesis confirmed previous results (Fig. III-25) as seen in the significant drop-off in signal intensity after AOC-deprotection.

Fig. III-25: Comparison of detritylation integrals in the model trisoligonucleotide synthesis with the AA-, AD- and DD- linker.

A noticeable increase in detritylation was observed after coupling of the DD-linker indicating a parallel synthesis of two strands. Signals were not expected to show double intensity. This is explained by how the data processing of the machine works and how the internal algorithm interprets an increase in signal intensity in a non-linear fashion. As a proof the DMT-waste after deblocking was collected and absorbance was measured separately via UV-spectroscopy (Fig. III-26). Contents of DMT did indeed double after addition of the DD- linker, which indicates the synthesis of two parallel arms.

51 III. Results and Discussion

1,2 1,0817 1,0335 1,0 0,9079

0,8

0,6 0,55500,555 0,5615

0,4362 0,4

0,2

DMT-Absorbance E at 501 nm (Units) nm 501 E at DMT-Absorbance

0,0 C T G DD-Linker G T Step

Fig. III-26: Comparison of DMT-absorbance before and after DD-linker addition. Dilution factor of 25.

After introduction of the AA-linker the two AOC-groups were cleaved and therefore no detritylation signal was expected as seen in the weak signal (“1. Deprotection”). The residual signal is explained by remaining unflushed DMT-ions from the previous coupling. Afterwards, coupling efficiencies of the two parallel arms were surprisingly comparable to the DD-linker unlike the noticeable drop in the case with the AD-linker. All raw product mixtures were then analysed with denaturing polyacrylamide gel electrophoresis (Fig. III- 27).

Fig. III-27: Electrophoretic analysis of raw product mixtures and MALDI-TOF-MS (Contains background noise); 16% denat. PAGE; 1xTBE; 120 V; 2 h.

Two major bands were visible and identified by MALDI-TOF mass analysis as the desired trisoligonucleotide and the two-arm mutant. Masses were within the expected range. The two-arm mutant in case of the DD-linker is devoid of the allyl ether moiety, but the small change in mass (< 1 %) was unnoticeable on the gel. Mass analysis showed the expected shift towards lower masses.[221]

52 III. Results and Discussion

Coupling with the AD-linker led to higher amounts of the two-arm mutant than the three- arm product, which is in correlation with the detritylation profile. Deletion mutants of the third arm are visible between the two major bands. The DD-linker confirmed the expectation and yielded more full length product than DA. Synthesis of trisoligonucleotides with two identical arms can therefore successfully be accomplished with double-DMT protected linkers on commercially available reverse CPG support. No manual synthesis of solid support is therefore necessary. In the case of the AA-linker the amount of the three-arm trisoligonucleotide is also higher than the two-arm mutant in comparison to the AD-linker. Doubling the concentration of AOC-groups, and hence the allyl ether, somehow improves the yield of trisoligonucleotides instead of having a diminishing effect seemingly mitigating the allyl ether side reaction. It can only be speculated what causes this reversal in behaviour: Since the only noticeable differences between AD and AA are the number of alloc-groups and synthesis strategy, explanations must revolve around these facts. One possible explanation lies in the synthesis of AD, whereby the second arm causes a sterical hindrance in the synthesis of the subsequent third arm leading to lower yields. Another could be an alternative cleavage mechanism caused by the close proximity of two AOC-groups (Fig. III- 28).

Fig. III-28: Hypothetical cleavage mechanisms involving two alloc groups.

Either one unit of the Pd(0) species could form a theoretical bisallyl palladate or two units form a bispalladate bridged by one ligand. A mechanism with two units is statistically possible by the more then thousand-fold excess of the catalyst over alloc used in the reaction. Structurally similar compounds exist, like bis(η3-allyl)palladium,[252,253] bis[chloro(η3-allyl)palladium][254] or tris(dibenzylideneacetone)dipalladium,[255] but they are all uncharged and form stable solids. These charged species, however, are not found in

53 III. Results and Discussion literature probably due to their high activity and therefore instability. The allyl cations in these complexes are presumably less effected by the solid support and are much less likely to initiate the allyl side reaction, which caps the linker for further synthesis.

It was assumed that this highly active catalyst species is more likely to react with the trapping nucleophile, i.e. pyrrolidine base, and induce an allyl shift than the normal allyl palladium complex. Synthesis of the same trisoligonucleotide with the AA linker was repeated, but without the presence of the pyrrolidine base. Expected was a very strong drop in detritylation after AOC deprotection like in the AD-case. Detritylation integrals were then compared to the results in the presence of the base (Fig. III-29).

AA AA -Pyrrolidine 600

500

400

300

200

Detritylation Intergral [A.U.] Intergral Detritylation 100

0

C A G T A C T G G T C A T G A C

Fig. III-29: Comparison of detritylation integrals of trisoligonucleotides syntheses with the AA-linker in presence of a base Trislinker/1. Deprotection and its absence (“-Pyrrolidine”). Comparison shows reduced coupling efficiencies at the twin arms, which are now comparable to the yields of the first arm. Quantities were independently confirmed by UV- spectroscopy of the DMT-waste before and after AOC-deprotection. Unexpectedly, coupling efficiencies were still high in absence of a base. This might be seen as a clue for the existence of an alternative deprotection mechanism for which an allyl shift is unnecessary. This can be explained by the existence of alternative palladium complexes that can either stabilize the allyl cations or cause its rapid decomposition through high reactivity.

Removal of the coligand dppe in order to study its effect on the reaction and in complex formation was not possible, because the addition was necessary for reasons of catalyst solubility. No further studies were conducted.

54 III. Results and Discussion

III.2.7. Fluorous Affinity Purification Studies

Neither DMT-off nor DMT-on[256] chromatographic purification methods can be applied to unmodified trisoligonucleotides. Liquid chromatography of trisoligonucleotidyls with 9 and 15 bases per arm gave unsatisfactory separations regardless of solvents or elution gradients as discovered by Dorenbeck.[155] Longer oligonucleotides made separation increasingly more difficult. Cartridge solutions based on DMT-on solid phase extraction (SPE), like Glen-Pak by Glen Research,[245] also resulted in insufficiently resolved mixtures. This behaviour is attributed to the branched structure, which makes all branched mutants less distinguishable on solid phase separation due to similar degrees of flexibility. Preparative polyacrylamide gel electrophoresis of trisoligonucleotides was successfully established as an alternative purification method.

Pearson et al. described fluorous affinity interaction as a potent method in the purification of oligonucleotides in the form of fluorous tagging.[257,258] The final nucleoside phosphoramidite in the automated synthesis bears a perfluorated alkyl chain at its 5’-DMT group (FDMT). This fluorous modifier contains a domain rich in sp3 carbon-fluorine bonds. Fluorous-tagged oligonucleotides interact preferably with perfluorated adsorbents like perfluorooctyl- modified reverse-phase silica gel as part of F-SPE and F-HPLC purification methods.[259] The added mass of the fluorous tag also improves the separation in reverse-phase liquid chromatography (RP-HPLC). A retention of fluorinated substances on perfluorated adsorbents was observed, whereas untagged material is washed out with the aqueous solvent front. Organic solvents elute the tagged substance afterwards. Pearson et al. were able to purify long linear oligonucleotides with an F-SPE procedure between 50-100 bp with recoveries between 70-100 %. Highly fluorinated organic compounds (fluorous content of more than 60 % by weight) are both hydrophobic and lipophobic.[260,261] They are considered as fluorophilic though because of their preferred interaction with other fluorinated substances and the tendency to form their own phase. Light fluorous compounds containing less than 40 % of weight fluorine are less likely to form a separate phase. An alternative method by Gubta and Will replaces the acetate capping step in oligonucleotide synthesis with a fluorous capping step. The agent (Fig. III-30) caps every deletion sequence while the final product remains uncapped enabling separation by F-SPE. [262]

The plan was to apply fluorous tags on trisoligonucleotides, study the effect on separation and optimize a method for tag-cleavage. So far only FDMT was reported as a compatible fluorous tag in oligonucleotides.[257] FDMT can only be tagged after the synthesis of the third

55 III. Results and Discussion arm as part of the final FDMT-modified nucleoside phosphoramidite. A detritylation step in the synthesis of the third arm would cleave a FDMT-group at the end of the second arm. A trisoligonucleotide with two fluorous tags is assumed to give better separations of failure sequences than a species with only one fluorous tag. A potential linker must be stable under acidic conditions and allow for post-synthetic cleavage in aqueous solution because of the solubility of DNA. García-Echeverría and Häner described an easily available decylthioethyl- based N,N-diisopropyl phosphoramidite as a lipophilic protecting group for oligonucleotides (Fig. III-30).[263] Substitution with a perfluorous chain can potentially add fluorophilic capabilities (“F-TAG”).

Fig. III-30: Original lipophilic oligonucleotide protecting group by García-Echeverría & Häner ; Fluorous capping agent by Gupta & Will; Synthesis of a derived fluorous analogue “F-TAG”.

The 2-step synthesis started in a basic treatment of 2-mercaptoethanol 45 with sodium hydroxide in tert-butanol (Fig. III-30). Deprotonation occurs primarily at the thiol explained by the generally lower pKa-values than alcohols.[264] A substitution reaction with 1-Iodo-[1H, 1H, 2H, 2H]-perfluorodecane gave perfluorooctyl ethyl thioethyl hydroxide 47. The remaining hydroxide reacted with the Bannwarth-reagent in presence of DIPEA to yield the 2- cyanoethyl-2-perfluorooctyl ethyl thioethyl-N,N-diisopropyl phosphoramidite 48. An evaporative light scattering detector (ELSD) enabled automated chromatographic purification in cyclohexane/ethyl acetate due to weak detection in UV.

56 III. Results and Discussion

Trisoligonucleotide TF (Tab. III-10) was based on the T3 linker core with the F-TAG at the 5’- ends of the second and the third arm. Synthesis used the trisoligonucleotide protocol, whereby the F-TAG amidite was coupled twice for five minutes each at the end of the second arm and three times for five minutes each at the end of the third arm.

Sequences 5’-3’ TF ε (254 nm) Vertex (Arm 1, 2, 3 on T3 linker) [Lcm-1mol-1] FTAG-CTC CAC GTT ACG ATC TF FTAG-GTC CGA CAG TTA GTC 459861 FTAG-GTA AAC GAG GAG GTC

Tab. III-10: Sequence design and extinction coefficient of trisoligonucleotide TF. Theoretical melting point is ~55.4 °C. The study started with the chromatographic purification of the raw product. Fluorophilic interaction was tested on a FluoroFlash HPLC column (150x4.6 mm; Fluorous Technologies). This column was packed with perfluorooctylethyl-modified silica. In order to study the fluorophilic effect an unbuffered acetonitrile/water eluent (5-80 % ACN, 45 min) was used at first for separation. The fluorophilicity alone between the tags and the fluorous adsorbent was not sufficient to separate the desired double-tagged product from the other tagged mutants. F-HPLC on the FluoroFlash column showed no separation. The switch from water to a 0.1 M ammonium hydrogen carbonate buffer (AHC; pH: 8.5) was an improvement. The need for a buffer, however, necessitates a desalting step after separation. Gradients of acetonitrile in AHC-buffer (5-80% ACN) were analysed for an elution time of 45 min. It was not possible to completely separate TF from mutants on the fluorous column. The switch to reverse-phase (RP18) columns slightly narrowed the peak profiles and improved separation. A gradient of 20-30 % ACN/AHC over 30 minutes gave satisfying results. Mutants eluted after 9-15 minutes and the desired double-tagged TF had a retention time of around 27-28 minutes (Fig. III-31). MALDI-TOF-MS confirmed the expected mass of 15335 m/z. The high difference in retention times gives enough range to overload a column on a semi- preparative scale where higher loading on the column is possible.

TF with two tags showed a much more lipophilic behaviour than all of the tagged and untagged mutants. Desired product was collected, combined, desalted and resolved in water. Prior to UV-spectrophotometric quantization the sample was treated with ultrasonic waves for 20 minutes to avoid a potential decrease in solubility caused by fluorophilic association effects of TF. A yield of 32 nmol was determined after photometric analysis.

57 III. Results and Discussion

Fig. III-31: Successful separation of TF (27-30 min) in the raw product; RP-HPLC, 20-30 %, ACN/AHC (0.1 M, pH: 8.5), 30 min; 254 nm. Samples between 7 to 33 min of the previous chromatographic separation were collected. The contents were studied in denaturing gel electrophoresis (Fig. III-32). The first fraction between 7-9 minutes most likely contained the untagged trisoligonucleotide. (It is unclear, what the first three weak bands are. MALDI-analysis showed no results.) Different gradients (e.g. 5-10 %) did not allow to separate it from the untagged deletion mutants. The second and third fraction between 9 to 13 minutes contained mostly tagged material. A very predominant band is visible, which was, Fig. III-32: Gel electrophoretic study of the all presumably, the single-tagged trisoligonucleotide. noteworthy peaks of the successful TF separation; Denat. PAGE 12 %, 1x TBE, 100 V, 75 min; SYBR gold It was also not possible to isolate the single-tagged nucleic acid gel stain (post-gel). product from the mutants. The fraction at 27-29

58 III. Results and Discussion minutes showed a pure band of slightly slower electrophoretic mobility compared to the other fraction. Later fractions contained traces of the same product.

Cleavage conditions for the tag were investigated on a simpler and cheaper model compound first. Pentamer homothymidine (dT)5 was prepared under standard oligonucleotide synthesis conditions on a 1.3 µM scale for that matter. F-TAG (0.2 M; ACN) was coupled twice for five minutes after final detritylation. Preparative liquid chromatography in a 5-80 % ACN/AHC gradient (25 min) showed two species with retention times of ~4 minutes (5T) and ~11 minutes (5T-F), respectively. Products were isolated and desalted over HLB Oasis cartridges (Waters). Mass analysis via MALDI-TOF (Tab. III-11) revealed the first peak to be the homothymidine 5T and the second peak fluorous-tagged product 5T-F. The coupling efficiency of the F-TAG was ~50 %.

5T 5T-F m/z calc. 1538.2 2045.3 m/z exp. 1540.9 2047.6 yield [nmol] 175.87 179.39

Tab. III-11: Mass via MALDI and yield for 5T and 5T-F.

5T-F showed a very strong affinity to the adsorbent in F-HPLC. Fluorous material was fully retained in 100 % water (Fig. III-33).

Fig. III-33: Behaviour of 5T-F in F-HPLC on a FluoroFlash column (150x4.6 mm) with different acetonitrile/water gradients. UV-detection at 260 nm.

The addition of acetonitrile was necessary for release (8.7 min). 5T-F appeared with the solvent front (2.4 min) if a 5-80 % gradient of acetonitrile/water was directly applied. No buffer was necessary to enable fluorous interaction. This can simplify purifications in theory because no desalting step is necessary after extraction. Once the sulfide is oxidized to the

59 III. Results and Discussion sulfone the adjacent protons are acidic enough to induce β-eliminations in presence of an aqueous base. Elimination towards the phosphate groups leads to cleavage of the entire tag (Fig. III-34). The oxidation to the sulfone serves as an activation step. Fluorous-tagged sulfonyl-linkers already exist in peptide chemistry, but are considered too labile towards bases and require precautious handling.[265,266]

Fig. III-34: Mechanism of deprotection. Oxidation to a sulfone and β-elimination. The phosphate remains attached on DNA. García-Echeverría and Häner describe a reaction with a 20-fold excess of N- chlorosuccinimide in 50 mM tetraethylammoniumbromide/dioxane (3:1 v/v; pH: 7.5) for 2.5 h at room temperature and isolated the sulfone via HPLC. (Suspiciously, an agent like m-CPBA needed for oxidization to the sulfone was not mentioned.) Elimination was induced with concentrated ammonia for 16 h at room temperature and then purified again by HPLC. The need of two separate HPLC purification steps made this cleavage strategy less attractive. Alternative methods were investigated.

Sodiumperiodate was the oxidizing agent in initial setups (Tab. III-12). A 1 mM stock solution of 5T-F in ddH2O was prepared for oxidation experiments. Conversion was followed via liquid chromatography. 100 mM periodate in an overnight reaction gave a full conversion to the sulfoxide exclusively. A basic environment is needed to cleave the tag. The addition of 100 mM ammonia negatively affected the conversion rate. Similar results were found with 100 mM triethylamine. More drastic setups with 1 M periodate at 50 °C also gave a full conversion only to the sulfoxide.

60 III. Results and Discussion

Concentration of NaIO4 Temperature Time Outcome

10 mM r.t. 1.5 h 25 % sulfoxide; 75 % 5T-F

30 mM r.t 1.5 h 50 % sulfoxide; 50 % 5T-F

100 mM r.t. 20 h 100 % sulfoxide

100 mM + 100 mM NH3 r.t. 20 h 75 % sulfoxide; 25 % 5T-F

1 M 50 °C 20 h 100 % sulfoxide

Tab. III-12: Exemplary setups of the 5T-F (1 mM) oxidation with sodium periodate.

Literature examples confirmed the weak oxidizing effect, which stops on the level of the sulfoxide and instead suggested hydrogen peroxide to obtain the sulfone.[267] Oxidation setups with a 1 M H2O2 solution (~3 % w/w) resulted in high amounts of side products. In an attempt to minimize the formation of side products the reaction was conducted with 100 mM

H2O2 instead and was followed over specified time intervals via HPLC (Fig. III-35). After five minutes a new peak was detected with a one minute shorter retention time than 5T-F. Mass analysis revealed the sulfoxide again. A ~50 % conversion was reached after one hour. Full conversion was observed after a 20 hour reaction period.

61 III. Results and Discussion

Fig. III-35: Oxidation of 1 mM 5T-F with 100 mM H2O2 (~0.3 % w/w) at 50 °C. Solvent front contains reagent; RP-HPLC; 5-80 %, ACN/AHC (0.1 M, pH: 8.5), 35 min; 254 nm.

Retention Retention Time Peak 1 Peak 2 5 min 15.86 min 16.70 min 35 min 15.90 min 16.70 min

1 h 16.20 min 16.94 min 2 h 16.33 min 17.25 min

20 h 17.71 min -

Tab. III-13: Assignment of retention times. The sulfoxide of 5T-F (Peak 1) and 5T-F (Peak 2) show increased retention times explained by ongoing pump issues on the machine.

62 III. Results and Discussion

It is known from organic synthesis on the multi-gram scale that catalytic amounts of tungstate salts can increase the reactivity of hydrogen peroxide.[268–270] Treatment of 5T-F with a high excess of Na2WO4 at 55 °C for 20 hours did not cause a major decomposition of the substrate (Fig. III-36). Sodium tungstate, in fact, had mild oxidative capabilities of its own leading to a ~25 % conversion to the sulfoxide. This introduces a favourable secondary oxidation mechanism.

Na2WO4 (1 M) 55 °C, 20 h

Fig. III-36: 1 mM 5T-F in presence of 1 M Na2WO4 at 55 °C for 20 hours; Tungstate appears at solvent front; RP-HPLC; 5-80 %, ACN/AHC (0.1 M, pH: 8.5), 35 min; 254 nm.

Aqueous sodium tungstate solutions are basic (pH0.1 M ~8.5; pH1M ~10.5). It therefore does not only act as a catalyst, but also as a base that allows the immediate in-situ β-elimination of the sulfone. No traces of the sulfone intermediate were detected in all cleavage reactions.

50, 100 and 1000 mM solutions of hydrogen peroxide and sodium tungstate were mixed in 2:1 to 5:1 ratios. The different mixtures were then applied to 1 mM 5T-F. Reactions were conducted at room temperature, 40 °C and 55 °C with reaction times ranging from 1 to 3 hours and 20 hours (overnight). High concentrations of reactants resulted in side-products at high temperatures and reaction times higher than an hour. At room temperature and

63 III. Results and Discussion short reaction times only partial conversion was observed with less side products. The ratio of the reactants had no noticeable effect on the reaction.

3:1 H2O2/Na2WO4 (50 mM) 55 °C, 20 h

Fig. III-37: Successful F-TAG cleavage to 5T with 3:1 H2O2/Na2WO4 (50 mM) at 55 °C for 20 hours. No intermediate sulfone was found. Direct elimination to the final product is assumed; RP-HPLC; 5-80 %, ACN/AHC (0.1 M, pH: 8.5), 35 min; 254 nm. Higher contents of tungstate were preferred for distinct basicity. Reaction conditions with a high conversion rate as well as a low amount of side products were found with 50 mM

H2O2/Na2WO4 in a 3:1 ratio at 55 °C for 20 hours (Fig. III-37). MALDI-MS confirmed the new product as the desired untagged thymidine pentamer 5T (Fig. III-38).

64 III. Results and Discussion

Fig. III-38: MALDI-MS of 5T-F, the sulfoxide “5T-F +O” and the untagged 5T after successful cleavage. Signals of higher order are related to the addition of sodium cations caused by insufficient desalting.

These cleavage conditions were applied to a 1 mM aqueous solution of the trisoligonucleotide

TF. Treatment with 50 mM H2O2 and Na2WO4 solutions in a 3:1 ratio at 55 °C for a duration of 20 hours barely showed a turnover of TF. The switch to 1 M solutions resulted in a mixture of the double-tagged-, the single-tagged- and the desired untagged TF trisoligonucleotide. Decomposition products also started to arise (Fig. III-39). All three major peaks in chromatography were collected and their mass was confirmed via MALDI-MS (Fig. III-40). Replacement of 1 M tungstate with 1 M ammonia and 1 M sodium hydroxide[263] gave similar results. Higher concentrations and harsher conditions were not studied, due to the growing risk of decomposition. Around 4 nmol of untagged TF were obtained out of all tagged material after chromatography and desalting. At this stage no other cleavage conditions were investigated.

65 III. Results and Discussion

Fig. III-39: Chromatogram of TF after F-TAG cleavage. RP-HPLC; 10-30 %, ACN/AHC (0.1 M, pH: 8.5), 30 min.

Fig. III-40: MALDI-MS of isolated major products after cleavage attempts. Experimental masses correlate with theory.

Only ~9 % of the mass in the double-tagged TF is allocated to the fluorous tags. It has a low fluorine content and is therefore a very light fluorine compound.[260,261] The fluorous tags have

66 III. Results and Discussion probably a minor effect on the overall structure.[266] This might explain the problematic separation via fluorophilic affinity in F-HPLC. An effect on the solubility or reactivity of TF caused by the tags is also unlikely. Increased temperatures in the mix (55 °C) improve solubility of fluorous compounds.[259,271] The highly polar DNA backbone overcompensates the fluorophilicity of the given tags. An unlikely concern is an intramolecular association of both F-tags, which could potentially block the sulphur reaction sides and hinder full conversion. Ultrasonication for up to 20 minutes had no influence on applied samples. Higher temperatures most likely weaken fluorophilic interactions. Both arms should be independent to one another. No intermolecular fluorophilic association in the cleavage of 5T-F was observed.

The presence of two tags enabled liquid chromatography of trisoligonucleotides at high purity and, technically, this method of purification did yield untagged trisoligonucleotide at the end, but it was not a viable alternative to preparative gel electrophoresis at this stage. Coupling efficiency of the tags need to be optimized further. Only 32 nmol of double-tagged TF were obtained. The rest of the desired trisoligonucleotide had either one or no tag. Those species were lost in chromatographic purification and could not be separated from the remaining mutants. Better efficiency in tagging can reduce the amount of those species.

The attempts to cleave the F-TAG yielded the desired untagged product only partially. It is not clear what caused the insufficient conversion, despite the use of drastic conditions that included concentrated reactants and bases at increased temperatures. Perhaps longer reaction times are required for full conversion. Longer times were not tested, because of the imminent decomposition of the materials and time concerns.

Even if the aforementioned shortcomings were solved, this method of purification cannot be considered as a noticeable improvement over preparative electrophoresis. Though the synthesis of the tag was short it still required additional work effort. Fluorous affinity did not satisfyingly separate the double-tagged species from the tagged mutants. Buffers were required to improve separation. A subsequent desalting step is necessary. Broad peak profiles of trisoligonucleotides in HPLC led to the collection of large volumes. Evaporation overnight prior to desalting was therefore mandatory. Cleavage of the F-TAG appeared to be very slow and most likely needed reaction times longer than one day. A second HPLC step needs to be applied after cleavage to remove reagents and by-products. This was followed by another evaporation and desalting step. Preparative gel electrophoresis is faster and easier in comparison.

67 III. Results and Discussion

III.3. DNA-Tetrahedron Assembly Experiments

III.3.1. Hybridization Conditions

Aqueous stock solutions of each trisoligonucleotide were diluted with a buffer, which contained 10 mM HEPES (pH: 7.5) and 100 mM NaCl and is henceforth referred to as “HB” (hybridization buffer). The salt is necessary for oligonucleotide hybridizations to counter charge repulsion effects at the backbone. The assemblies of neither tetrahedrons[155] nor dodecahedrons[159] benefit from the addition of bivalent metal ions including Mg2+. They tend to improve the stability of smaller fragments visible in gels. The choice of buffer like the phosphate- (pH: 7.4), MES- (pH: 6.7), or TRIS-buffer (pH: 7.6) shows no effect on self- assembly studies as previous findings indicate.[152,155,159] All previous work suggested a temperature program for self-assembly studies that starts with a denaturing step, transitions to an annealing step and then reaches a final cooling step. [199] Temperature protocols were used for trisoligonucleotide tetrahedron assemblies (Tab. III-14).

TAP1 TAP2

Temperature Time Speed Time Speed

95 °C 5 min 5 min

↓ 0.3 °C/s

40 °C 5 min ↓ 0.5 °C/s

↓ 0.1 °C/s

0 °C 15 min 15 min

Tab. III-14: Trisoligonucleotide assembly protocol (TAP); TAP1 contains an annealing step set below the theoretical melting. TAP2 omits this step and goes immediately from the denaturing step to the cooling step.

The assembly of trisoligonucleotides into tetrahedral shapes starts with the denaturing step at 95 °C to prohibit specific and unspecific DNA interactions. In the first trisoligonucleotide assembly protocol TAP1 it then cools down in 0.3 °C increments per second to an annealing- temperature set 15-20 °C lower than the melting point. The original set value of 50 °C was lowered to 40 °C to improve the distinction between the melting- and the annealing- temperature. The annealing step should allow for a slow hybridization process into stable

68 III. Results and Discussion assemblies. After a specified time the cooling slowly continues towards 0 °C (0.1 °C/s) and stays for 15 minutes to ensure a quantitative assembly. Alternatively, the annealing-step can be omitted and is replaced with a direct gradient from 95 °C to 0 °C (TAP2).

Fig. III-41: A: Hybridization of T2 under TAP1: Immediate analysis (1) and after overnight storage at room temperature (2); Similarly under TAP2 conditions (3) and stored sample (4); HB; Nat. 3 % Agarose 1000, 100 V, 2 h; GelStar nucleic acid gel stain (in-gel). B: T3 set; Hybridizations in 0-7.5 mM HEPES (pH: 7.5) and in 15-75 mM NaCl; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

TAP1 and TAP2 conditions were compared with the exemplary T2 trisoligonucleotide set (Fig. III-41). The assemblies were analysed immediately (A: 1; 3) and after storage at room temperature (A: 2; 4). Both protocols led to a similar quality in hybridizations.

Buffers like HEPES alone are insufficient for successful assemblies into tetrahedrons (Fig. III-41; B). The same situation was observed with up to 10 mM sodium phosphate buffer (pH: 7.5) in a study with the T3 set. 30 mM solutions led already to sufficiently stable products, which was much lower than the suggested 100 mM solutions. Smearing below 90 bp is explained by a partial decomposition of the studied trisoligonucleotide set and is unrelated to the hybridization behaviour. For reasons of comparability to previous works the default experimental setup in this work used TAP1 conditions, 100 mM NaCl and the addition of 10 mM HEPES-buffer (pH: 7.5).

69 III. Results and Discussion

The effect of trisoligonucleotide concentration in the range of 0.3 to 3.0 µM solutions was also studied on the T2 set. In all cases a distinct band of the tetramer was visible, but with a slight increase in electrophoretic mobility with higher concentrations (Fig. III-42).

Fig. III-42: The concentration effect between 0.3 to 3.0 µM of tetramer T2; HB, TAP1; 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

III.3.2. Effect of short Alkylene Chains in T1, T2 and T3 Assembly Studies

Of all Platonic solids in Euclidian geometry the tetrahedron is the simplest conceivable object with a three-dimensional shape. In a constructional context tetrahedral scaffolds also entail a stiffness unlike higher order geometries: A dodecahedral- or cube-shaped scaffold constructed from rigid struts and flexible vertices, for instance, can collapse under the gravitational force, whereas a tetrahedral scaffold stays intact in the same situation.[272] In correspondence to Buckminster Fullers principle of tensegrity[273][274] an intrinsic tension allows the tetrahedron to maintain structural stability even if an external force is applied. The added stability might translate to tetrahedral shapes on the nanoscale.

Four trisoligonucleotides are sufficient to form a tetrahedral shape, whereas assemblies into higher geometries require a higher preparative effort. The centre of every C3h- symmetrical trisoligonucleotide is the trislinker and requires sufficient flexibility in order to position all three oligonucleotide arms into the three-dimensional shape (Fig. III-43).

70 III. Results and Discussion

Fig. III-43: Assembly of four trisoligonucleotides into a tetrahedral shape. Every colour marks complementary sequences.

The comparison of theoretical interior angles at different faces (Fig. III-44) reveals an angle of 60° for a tetrahedron, 90° for a cube and 108° for a dodecahedron. The oligonucleotide arms bend the most from a planar initial position to assemble into a tetrahedral shape. With higher geometries, trisoligonucleotides deviate less from the planar unconstrained position.

Fig. III-44: Geometrical consideration of a tetrahedron, a cube and a dodecahedron. Higher geometries lead to higher interior angles ranging from 60° to 108°.

The highest constrain for a trislinker to form a cage-like structure is found in a tetrahedral shape and therefore the most interesting scaffold to study different linker designs. The trisoligonucleotide sets T1, T2 and T3 share the same sequence design and only differ in the length of their alkylene arms (T1: methylene; T2: ethylene; T3: propylene). If a linker is flexible enough to form stable tetrahedral shapes, it is probably also applicable in higher order geometries of lesser constrain.

71 III. Results and Discussion

The quality of all trisoligonucleotide building-block monomers of the T1, T2 and T3 sets were controlled with gel electrophoresis prior to assembly studies (Fig. III-45). No noticeable side products were found. The size corresponded to ~22.5 bp and masses were confirmed with MALDI-MS. Small differences in running speeds roughly correspond to the different masses of the Fig. III-45: Comparison of T1, T2 and T3 monomers; 1.25 µM; HB; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 1 h; GelStar nucleic acid gel monomers. stain (in-gel).

Now with the purity of all trisoligonucleotides confirmed, self-assembly experiments were carried out for every possible combination of building blocks per set (Fig. III-46).

Fig. III-46: Full T1, T2 and T3 assembly gels in clockwise order beginning from top. Display of all dimer- and trimer- permutations (“12” = Monomers T.1 and T.2; “123” = Monomers T.1, T.2 and T.3. Two monomers added for reference. The last band is the final tetramer; HB, TAP1; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 75 min; GelStar nucleic acid gel stain (in-gel).

The assembly was analysed by native agarose gel electrophoresis for each set to test for errors in the sequences, namely substitution and deletion mutations, leading up to the full assembly of all four trisoligonucleotides. There are six distinguishable binary combinations leading to dimers connected at one arm (Fig. III-46, “12” to “34”), four distinguishable ternary combinations containing three double-stranded arms (“123” to “234”) and one unique quaternary combination with six double-strands. Two out of the four monomers (“1” and “4”) were added as reference. Bands show full addressability for each arm and no sequence errors. Faint higher order bands are explained by mismatching and the formation of higher aggregates, which were more noticeable in the T2 set. The trimer bands (“123” to “234”) showed an even stronger tendency to higher order mismatches most predominant in the T2

72 III. Results and Discussion set. This might be caused by slight geometric differences in the possible orientations of ethylene chains compared to methylene- and propylene chains. The bands below the trimer bands most likely belong to residual not hybridized dimer material. All mixtures assembled into tetramers (Fig. III-46; circled “T1”, “T2” and “T3”) with an expected size equivalent to 90 bp. A distinct faint band in all tetramer bands with a size of 180 bp was visible and was assigned to a cube-shaped structure (Fig. III-47). This shape is formed with two equivalents of each monomer for statistical reasons, but unlikely to occur in this specific sequence design given an entropic favouring of complexes of lower molecularity as opposed to those of high molecularity. The strong trail of high molecular aggregates in T2 made it more difficult to spot this band than in T1 and T3.

Fig. III-47: Assembly of two trisoligonucleotide sets into a cube-shaped motif via assembly of two separate tetragonal planes and subsequent dimerization, which requires a 180° rotation step along the hybridized axis. Overall, all three sets were capable to hybridize up to the tetramers according to the assembly gels. In order to prove the assembly into discrete tetrahedral objects an enzymatic digestion experiment was utilized for verification. With a highly rigid linker it becomes impossible for all the short single-strands to hybridize and fully fold into the tetrahedral

73 III. Results and Discussion shape (Fig. III-48). Mung bean nuclease is an endonuclease and preferably digests single- stranded DNA.[159] Digestion products will appear in presence of the enzyme.

Fig. III-48: A fully closed DNA-tetrahedron consists only of double-stranded DNA and is not digested by the enzyme mung bean endonuclease unlike a partially open tetrahedron or just a trimer hybrid.

In digestion experiments all tetramers of the three sets of T1, T2 and T3 were incubated with 5 units of mung bean nuclease for 10 minutes at 30 °C. (Fig. III-49). The trimer “123” of the T3 set was added as a reference to monitor the activity of the enzyme. The T1 tetramer was digested under the given conditions (T1, “+E”) and confirmed the previous results by Schorr.[202] Conversely, the T2 tetramer band stayed intact after digestion. T3 expectedly showed no digestion of its tetrahedral assembly product as established in similar studies by Cebulla.[199] Smudgy trails appeared below the tetrahedron bands, which are presumably caused by the digestion of higher molecular side products of Fig. III-49 Trimer 123 of the T3 set, tetramers T1, T2 and T3; HB, TAP1; “+E”: Incubation of sample to the left with partially folded tetramers. Based on this 5 units of mung bean nuclease for 10 min at 30 °C; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 2 h; GelStar experiment it can be concluded that ethylene nucleic acid gel stain (in-gel). chains of T2 trislinker are sufficiently flexible to allow folding into stable tetrahedrons.

74 III. Results and Discussion

III.3.3. Assembly and Digestion Experiments of TN

The commercially available tris(2-hydroxyethyl)-N,N’,N’’- isocyanurate 36 bears three ethyl chains on a six-membered heterocyclic ring and is therefore structurally similar to the T2 linker. The viability as a flexible linker core was studied in the corresponding TN trisoligonucleotide set using the same conditions and sequences as T1, T2 and T3 and assembly conditions (Fig. III-50). Gel Fig. III-50: TN assembly gel; HB, TAP1; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, analysis showed distinct 100 V, 65 min; GelStar nucleic acid gel stain (in-gel). hybridizations up to the tetramer as the prior sets.

The digestion experiment compared the trimer “123” of the TN set with the tetramer TN (Fig. III-51) with five units of mung bean nuclease at 30 °C and incubation times of 5 and 20 min. The tetramer band was stable for this duration, unlike the trimer. Analysis showed a similar behaviour to the T2 and T3 set, which is also interpreted as the formation of a closed tetrahedral shape. The isocyanurate linker is therefore flexible enough to be used as a linker in the assembly of trisoligonucleotide- based tetrahedrons like the comparable T2 trislinker. Fig. III-51: Digestion experiments with the trimer 123 of the TN set and tetramer TN; HB, TAP1; 5 U MBN at 30 °C; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 65 min; GelStar nucleic acid gel stain (in-gel).

75 III. Results and Discussion

III.3.4. Intermixing Assembly Experiments between T1, TN and T3

The same sequence design was shared by trisoligonucleotide sets with varying flexibilities at the trislinkers. This allowed to combine building blocks from different sets and study if the structural integrity of the constructs is somehow influenced by intermixing. The most interesting case was T1 with its rigid linker, incapable of assembling into tetrahedrons by itself. Experiments included a systematic intermixing of T1 trisoligonucleotides with the complementary trisoligonucleotides of more flexible sets.

Fig. III-52: Varying mixed equimolar assemblies between the T1 and the TN set; 1.25 µM; HB, TAP1; “+E” = 5 U MBN, 10 min, 30 °C; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

The first setup started with the assembly of all four trisoligonucleotides of the T1 set and zero from the TN set (Fig. III-52; “40”). T2 was omitted in these studies because of structural similarity to TN. In the next assembly mixture the trisoligonucleotide T1.1 was then replaced with the corresponding TN.1 (“31”). The other assemblies contained the mixture of two T1 with two TN trisoligonucleotides (“22”) and one T1 with three TN (“13”). The setup ended with the assembly of the pure TN set (“04”). Assembly conditions were identical to the previous experiments. Analysis of gel electrophoresis showed in the case of “31” that the addition of only one flexible trisoligonucleotide leads already to a distinct change in electrophoretic mobility. For verification, digestion experiments with mung bean nuclease (“+E”) revealed the presence of a stable tetrahedron band with partial digestion patterns (“31+E”). By increasing the TN content further the stability of the tetrahedral shape was seemingly increased even more. In a different setup T1 was mixed with T3 trisoligonucleotides to study the effect of the more flexible propyl chains (Fig. III-53). A clean tetrahedral shape with no noticeable digestion patterns was reached with the replacement of two building blocks (“22+E”) unlike the equivalent experiment with TN, which still showed minimal digestion.

76 III. Results and Discussion

Analysis suggests that the flexibility of T3 compensated the rigidity of T1 slightly better than the TN building blocks. The explanation is again given in the different flexibilities of the alkylene chain lengths. In a control experiment T1 and T3 were swapped in places to cover the influence of T1.1, which was not intermixed before. Results after the digestion were identical, but in reversed order.

Fig. III-53: Varying mixed equimolar assemblies between the T1 and the T3 set; 1.25 µM; HB, TAP1; “+E” = 5 U MBN, 10 min at 30 °C; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

Fig. III-54: Varying mixed equimolar assemblies between the TN and the T3 set; 1.25 µM; HB, TAP1; “+E” = 5 U MBN, 10 min at 30 °C; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

Based on the two previous setups, mixed assemblies of the TN and T3 led expectedly to stable intermixing tetrahedrons (Fig. III-54). Unlike the previous intermixing experiments no change in electrophoretic mobility is noticeable, because similar masses of the T3 and TN linker.

77 III. Results and Discussion

III.4. The Concept of UNO-, DOS- and TRE- Sequence Patterns

III.4.1. Introduction

All previous works on trisoligonucleotidyls and trisoligonucleotides shared two different sequence patterns: All three oligonucleotide sequences were either identical (XXX) or completely distinct (XYZ). Three unique sequences per building block enable full addressability into discrete polyhedral assemblies, whereas dendrimerization can occur when three self-complementary sequences are used. Unexplored was the middle-ground between sequence pattern, in which two arms share the same sequences and the remaining arm has a unique sequence (XXY). This XXY setup might be described as a “semi- addressable sequence pattern” in comparison to the fully addressable XYZ pattern. The twin arms (X, X) reveal an intrinsic ambiguity when being addressed by a complementary sequence, unlike the unique arm (Y). A set of XXY-trisoligonucleotides could theoretically lead to discrete and dendrimerized objects at the same time under certain conditions.

A new naming scheme for all three patterns is introduced, namely UNO, DOS and TRE. The order indicates the sum of arms per trisoligonucleotide that share the same oligonucleotide sequence (Fig. III-55).

Fig. III-55: Schematic overview of all three conceivable trisoligonucleotide sequence patters. UNO is characterized with three unique sequences, DOS with two and TRE with only one. UNO has a sequence X one time, DOS twice and TRE three times.

The T1-, T2-, T3- and TN-sets of the previous chapter are accordingly UNO-sets containing UNO-trisoligonucleotides. DOS- and TRE-trisoligonucleotides can be achieved with orthogonally protected trislinkers, but can also be synthesized with a simpler non- orthogonal linker (Fig. III-56).

78 III. Results and Discussion

Fig. III-56: The orthogonally protected trislinker on the left can generate all three possible sequence patterns, whereas the non-orthogonal linker on the right is not capable to form the UNO-pattern. Deprotection will always cleave both DMT- groups and the simultaneous synthesis of two identical arms.

III.4.2. Hybridization and Digestion of the TRE-Pattern TY

An example for the TRE-pattern, which bears three self-complementary sequences, is the trisoligonucleotide TY (chapter III.2.3.) based on the isocyanurate linker. Previous experiences with this sequence pattern include assemblies into nano-acetylene and nano- cyclobutadiene with the structurally different Ψ-Linker[152] (chapter I.5). A self- complementary design can either lead to discrete objects or dendrimerization into super aggregates depending on the hybridization conditions. It was of interest to know if a different linker core somehow changes the hybridization behaviour.

TAP3 Temperature Time Speed 95 °C 5 min ↓ 1.5 °C/s 0 °C 5 min ↓ 0.3 °C/s 35 °C 15 min ↓ 0.5 °C/s 0 °C 5 min

Tab. III-15: Trisoligonucleotide Assembly Protocol 3 (TAP3) includes a fast cooling step after denaturation to potentially avoid dendrimerization of self-complementary building blocks.

79 III. Results and Discussion

By following hybridization protocols that contain a denaturing step, an annealing step and a cooling step (like TAP1) dendrimerization (>1000 bp) occur according to Schefflers studies. Discrete bands in assemblies using the Ψ-Linker only appeared with altered protocols that included a fast cooling step to 0 °C after denaturing at 95 °C. It was then followed by heating to the annealing-temperature and subsequent cooling to 0 °C. It is assumed that kinetic control by fast cooling favours the assembly of small aggregates. A new assembly protocol TAP3 was used in the own studies that incorporated this fast cooling step (Tab. III-15). TY was assembled separately with the TAP1 and TAP3 protocol using the standard hybridization buffer (HB).

No major difference in aggregation behaviour was observed, which contradicts the previous results obtained by Scheffler based on the Ψ-Linker (Fig. III-57). TY was apparently capable to form discrete aggregates even without a fast cooling step. It is not clear how the switch from the Ψ-linker to the isocyanurate-linker had such an effect on the aggregation.

It was attempted to force super

Fig. III-57: Aggregation of TY under different assembly conditions aggregation with the addition of (TAP1 and TAP3) in hybridization buffer HB and with added 5 mol% salts (MgCl2, MnCl2, NiCl2 and ZnCl2); 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, metal-salts. In theory, it was 100 V, 70 min; GelStar nucleic acid gel stain (in-gel). expected that the bivalent alkaline earth metals (e.g. Mg2+) can favour hybridizations into discrete objects just like the monovalent alkali metals (e.g. Na+).[151,275] However, the presence of bivalent transition metal ions like Ni2+ and Zn2+ can lead to large undefined aggregates because they do not only interact with the phosphate backbone, but also with the nucleobases.[276] TY was hybridzed again under TAP 1 and TAP3 conditions, but with added 5 mol% of MgCl2, MnCl2, NiCl2 and

+ ZnCl2 to the hybridization buffer (100 mM Na ). Expected results, previous results with the Ψ-linker and studies with TY are summarized in the next table. The TAP1 and TAP3 protocols had no noticeable effect in the assembly behaviour and were neglected.

80 III. Results and Discussion

Cation Expected Results Results Ψ-linker Results TY

100 mM Na+ (HB) discrete bands discrete bands discrete bands

partial aggregation 5 mM Mg2+ discrete bands discrete bands + discrete bands

5 mM Mn2+ aggregation - discrete bands

5 mM Ni2+ aggregation discrete bands aggregation

mostly aggregation mostly aggregation 5 mM Zn2+ aggregation + weak discrete bands + weak discrete bands

Tab. III-16: Qualitative summary of the cationic effect in the assembly of a self-complementary trisoligonucleotide/-yl.

According to the old studies the addition of 5 mol% Mg2+ ions showed partial aggregation and Ni2+ gave discrete bands. In the case of TY Ni2+ did lead to aggregation and the presence of Mg2+ even improved the results as seen by the slightly decreased trailing of higher molecular aggregates. The addition of Mn2+ also improved the yields of the lowest molecular product around 50 bp compared to Mg2+. Zn2+ was close to expectation, but in both cases weak assemblies into discrete products was observed. Both old and new assembly experiments did not completely follow the expected behaviour.

In another set of experiments TY was also aggregated for 20 min at 30 °C (Fig. III-58). These simple assembly conditions still lead to discrete bands, but with a tendency to form higher molecular bands. Assemblies at double concentration (2.5 µM TY) showed besides more intense bands no other qualitative changes. No

Fig. III-58 Aggregation of TY under TAP3 conditions and distinct super aggregation was induced at for 20 min @ 30 °C at 1.25 µM (1x) and 2.50 µM (2x) concentration. “+E”= 5 U MBN, 5 min, 30 °C; 1.25 µM; double concentration. Enzymatic digestion with Nat. 3 % Agarose 1000, 1x TBE, 100 V, 120 min; GelStar nucleic acid gel stain (in-gel). mung bean nuclease (“+E”) reveals the band around 90 bp to be a fully closed tetramer. The only conceivable motif to form is the nano- cyclobutadiene as previously established.[275]

81 III. Results and Discussion

III.4.3. Hybridization and Digestion of the DOS-Pattern TD

A DOS-trisoligonucleotide has two arms with a shared sequence and one unique arm. A set of four trisoligonucleotides was prepared namely TD. TD.1/TD.2 have complementary twin arms to TD.3/TD.4, whereas TD.1 is complementary to TD.2 at the unique arm just like TD.3 to TD.4 (Tab. III-17).

Tab. III-17: Direct comparison between UNO- and DOS-trisoligonucleotide sets. Coloured letters indicate shared oligonucleotide sequences. Prime marks complementary strands. This DOS sequence design is simpler than the fully addressable UNO design bearing three unique sequences, but with potentially the same capability to assemble into a tetrahedron. Alternatively, this design might lead to dendrimerization or the formation of loop-like motifs by which two complementary DOS-trisoligonucleotides hybridize at their twin arms (Fig. III- 59; second from left).

Fig. III-59: Different conceivable motifs with two and three DOS-trisoligonucleotides.

82 III. Results and Discussion

A full assembly gel of TD was prepared in a similar fashion to the UNO sets (chapter III.3.2) for best comparison (Fig. III-60). The dimers “12” and “34” had a noticeably slower electrophoretic mobility than the other four dimer permutations. A look at band “124” showed traces of both bands simultaneously. This is Fig. III-60: TD Assembly gel and hybridization patterns; 1.25 µM; HB, TAP1; Nat. 3 % seen as evidence for the Agarose 1000, 1x TBE, 100 V, 70 min; GelStar nucleic acid gel stain (in-gel). existence of two different species. The four dimer permutations “13”, “14”, “23” and “24” contains duplicate complementary arms to form a loop-like motif, whereas the dimers “12” and “34” hybridizes only once into a branched DNA motif. The mobility of the different species on the gel is shape-dependent. Trimer permutations “123” to “234” also produced two distinct species, which in the case of band “134” coexisted. The DOS sequence design leads to either the now coined triangle-motif similar to the UNO-trimers or to the now coined swordfish-motif in which a trimer hybridizes adjacently to a loop-motif. Comparison of trimers “134” and “234” to UNO-T3 “123” and endonucleatic single- strand cleavage revealed the more mobile band to behave similarly to the UNO- trimer (Fig. III-61). Trimers of the UNO- sets can only assemble into the triangle- motif. Treatment with mung bean nuclease confirmed the shape. The digestion was still in progress on the given Fig. III-61: Comparison of UNO-T3 trimer “123”, DOS-TD “134” and “234” and full TD set; 1.25 µM; HB, TAP1; “+E”= 5 U MBN, 10 gel, but stops at the expected point. The min @ 30 °C ; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 70 min; GelStar nucleic acid gel stain (in-gel). more mobile trimeric species in DOS-“134”

83 III. Results and Discussion matches the UNO-“123” pattern. An identical motif was therefore assumed. The remaining less mobile trimeric band in DOS-“134” and the band DOS-“234” was likely the swordfish-motif in a process of elimination. The assembly of all four monomers, “TD”, was prone to form higher molecular aggregates like TRE-TY under TAP1 conditions (Fig. III-62). Repeating the full Fig. III-62: Assembly mechanisms of the triangle- and the swordfish- motif assembly experiment at tenfold in the DOS-set. concentration (12.5 µM) gave similar results and no occurrence of super aggregation.

A completely different assembly protocol was investigated in an attempt to further analyse the behaviour in hybridization of the full TD-set (Fig. III-63). This different approach started with a pre-assembly step of the UNO-T3 and DOS-TD dimers “12” and “34” under TAP1 conditions. The dimers of each set were then mixed and stirred at 25 °C. Gel electrophoretic analysis showed an almost completed conversion with minimal traces of unreacted Fig. III-63: Pre-assembly of UNO-T3 dimers (12, 34) and DOS-TD dimers (D12, D34) under TAP1 conditions and subsequent equimolar mixing at 25 dimers (T3: “0 min”). Surprisingly, °C in 5 min. intervals;. Samples were stored at 0 °C; 1.25 µM; HB; Nat. 3 % after five minutes the conversion Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel). already finished. UNO-T3 assembled into a tetrahedral shape as discussed previously. DOS- TD showed a vastly different pattern compared to the TAP1 assembly conditions with one

84 III. Results and Discussion major band around 90 bp, which was most likely nano-cyclobutadiene as the next study suggest.

A later experimental setup revealed the pre-assembly of dimers to not be a crucial step in the more directed assembly (Fig. III-64). All four monomers of the TD set were directly hybridized at 25 °C for five minutes (“Da”) and compared to the hybridization of the pre-assembled dimers (“Db”). The outcome was similar in both cases with less trailing at higher masses compared to the assembly at TAP1 conditions (“Dc”). The major band around 90 bp resisted nuclease degradation for at least ten minutes, which indicates a fully Fig. III-64: Da: Assembly of all monomers of the TD set for 5 min @ 25 °C; Db: Assembly of pre-assembled dimers D12 and D34 of the TD double-stranded motif. Nano- set for 5 min @ 25°C; Dc: Assembly of all monomers of the Td set under TAP1 conditions for reference; 1.25 µM; HB; “+E”= 5 U MBN, 10 cyclobutadiene is a very likely motif min @ 30 °C ; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel). and can be derived from the chosen DOS- sequence patterns (Fig. III-65).

Fig. III-65: Two exemplary mechanisms of assembly for nano-cyclobutadiene. The upper pathway starts with the assembly of branches and the lower with loops. Both pathways lead to the same shape.

85 III. Results and Discussion

Two mechanisms of assembly simultaneously lead to this motif with either branch- or loop- dimers at the start: The two branch-dimers connect via loops (upper path) and the two loop- dimers stack at their single-strands (lower path).

The given experiments in combination with enzymatic digestion share an insight in the behaviour of DOS-assemblies. The used sequence design allows for a more specific constructional prediction of loop-based motifs like the previously described nano- cyclobutadiene.

III.4.4. Sequence-Addressed Motif Designs via Intermixing Hybridizations

DOS-TD used a subset of the UNO oligonucleotide sequences (Tab. III-17). In analogy to chapter III.3.4. UNO- and DOS-crosslinking must therefore be possible. (Omitted were TRE- patterns because they were only studied as self-complementary building blocks not being capable to hybridize with the other available patterns.) T3 was the UNO-set of choice in all intermixing experiments. It gave slightly more pronounced bands in experiments than other UNO-sets. DOS-trisoligonucleotides contain only two unique DNA sequences and can therefore crosslink with a maximum of two UNO-trisoligonucleotides. An exception is the crosslink-dimer of T3.2 and TD.3 that is able to form a loop-motif (Fig. III-66).

Fig. III-66: Graphical visualisation of the relative arrangement of sequences in the UNO- and the DOS-set. Possible crossovers between the patterns are included.

86 III. Results and Discussion

Fig. III-67: Screening of all T3 and TD monomer pairings. Hybridization only occurs at expected pairs; 1.25 µM; HB, TAP1; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

All possible combinations of dimers between T3 and TD were screened (Fig. III-67). For instance, TD.1 only connects to T3.3/T3.4 and not to T3.1/T3.2. Experimental analysis confirmed the crosslinks for all expected combinations. A trimer band appeared as a side product whenever the D/D’ strands were involved (TD.1+T3.4; TD.2+T3.4; TD.3+T3.2; TD.4+T3.2) due to possible double hybridization at DOS-TD. The UNO- and DOS-sets were fully compatible for higher order crosslinking experiments.

UNO-trimers assembled into a triangular shape with three protruding unpaired arms by connecting each trisoligonucleotide to one another. Crosslinking with DOS- trisoligonucleotides might lead to the open canine-motif (Fig. III-68).

Fig. III-68: UNO-dimers (e.g. “12” is T3.1 + T3.2) and possible interactions with DOS-monomers to form the canine-motif.

Construction of the canine-motif started with the assembly of all UNO-dimer permutations under TAP1 conditions and the subsequent addition of an equivalent amount of viable DOS- monomers at temperatures below the theoretical melting point to avoid rehybridization. Gel electrophoresis showed varying rates of conversion to the trimer bands (Fig. III-69). The branched motif ran noticeably slower than the triangular UNO-trimers when compared to

87 III. Results and Discussion the relative position to the same DNA ladder (Fig. III-61). Double connections at the D/D’ strands caused higher molecular side-reactions.

Fig. III-69: Canine-motif between UNO-dimers “12” to “34” with equimolar DOS-monomers via hierarchical assembly. TAP1 for T3 dimers. Addition of TD monomers at 30 °C for 15 minutes; 1.25 µM; HB; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

Crosslinking also allowed for the discrete assembly of the swordfish-motif, which contained a loop-dimer and an adjacent monomer. All possible combinations were investigated and hybridized in equimolar amounts (Fig. III-70, Fig. III-71).

Fig. III-70: DOS-dimers and possible connections with monomers to construct the swordfish-motif. The monomer marked in red can possibly connect at both single-strands.

88 III. Results and Discussion

Fig. III-71: Swordfish-motif assembly. Loop-dimers mixed with suitable monomers at equimolar amounts; 1.25 µM; HB, TAP1; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

Gel electrophoretic analysis showed varying degrees of conversion just like in the canine- motif case. All trimer bands revealed noticeably different mobility on the gel in contrast to the branched trimers. One example was the addition of T3.3 to D13 and to D14. Another one was T3.2 and TD.4 to D23. It is not clear what exactly influences the variable mobility. Small conformational changes might appear to have a more pronounced effect in mobility than in other motifs. The mixing D24 and T3.1 gave uncertain results due to possible double hybridizations. The presence of two distinct bands made a clear structural assignment impossible in that case.

DOS-trisoligonucleotides can hybridize twice with a complementary UNO- trisoligonucleotide. This was demonstrated with the assembly of TD.4 with different ratios of T3.2 (Fig. III-72). The analysis clearly showed the stepwise conversion from dimer to the canine-motif by starting with the addition of 0.5 equivalents, then 1 and finally 2 equivalents of T3.2. The assembly was conducted at 0, 8 and 25 °C and under TAP1 conditions. The first three cases showed comparable results and a slightly more pronounced conversion under TAP1.

Replacement of TD.4 with TD.3 added the possibility to introduce a loop-shape by switching from sequence B’ to B (Fig. III-73). This single change had a major effect in the sequence- addressed motif design. Gel analysis showed distinctly different bands for both motifs in direct comparison. The canine-motif had a slower mobility on the gel than the swordfish- motif. The relative positioning was reversed after digestion with mung bean nuclease. The former motif had more digestible single-strands than the latter, which gave a digestion band of lower mass.

89 III. Results and Discussion

Fig. III-72: Assembly of TD.4 with altering ratios of T3.2. Conducted at 0, 8, 25 °C and under TAP1 conditions; 1.25 µM; HB; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 60 min; GelStar nucleic acid gel stain (in-gel).

Fig. III-73: Comparison between the canine-motif TD.4+2xT3.2 and the swordfish-motif TD.3+2xT3.2; 1.25 µM; HB, TAP1; “+E” = 5 U MBN, 8 min @ 30 °C; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 70 min; GelStar nucleic acid gel stain (in-gel).

90 III. Results and Discussion

Fig. III-74: Hierarchical assembly of the candy-motif (D14+T3.3+T3.1), direct assembly under TAP1 and digestion. Comparison to tetrahedral “T3” and its exemplary dimer “12” and trimer “123”; 1.25 µM; HB; “+E” = 5 U MBN, 10 min @ 30 °C; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 70 min; GelStar nucleic acid gel stain (in-gel).

Sequence-addressed motif design was also demonstrated on tetramers (Fig. III-74). The hierarchical construction started with the DOS-dimer “D14” and the stepwise crosslinks with T3.3 (“3.3”) and T3.1 (“3.1”) at 30 °C for 15 minutes to form the candy-motif (90 bp). Low temperatures minimized the risk of denaturation and unwanted rehybridization of T3.1 and T3.3 at the A/A’ arms. This method showed partial conversions to the desired bands. The switch to TAP1 assembly conditions gave very similar results. A higher excess of the used monomers might have improved the yield. The candy-motif contains four digestible single- strands unlike the fully enclosed tetrahedron “T3” as proven in the digestion experiments (“+E”).

The hybridization of DOS-TD trimer “D134” or “D234” with the UNO-T3 dimer “T23” potentially leads to pentameric motifs. Construction of cross-linked pentamers was unfortunately only detectable in traces within smudgy trails. No qualitative difference was detected by neither mixing the pre-assembled sub-units together nor by mixing all monomers directly. TAP1 conditions also did not improve the results. The motifs were presumably not stable under the given assembly conditions.

91 III. Results and Discussion

III.5. Anthracycline Intercalation

III.5.1. Qualitative Analysis

The DNA scaffold of a trisoligonucleotide-based tetrahedron was studied as a means of transport for the anthracyclines dauno- and doxorubicin (Fig. III-75). These anti-cancer agents interact with DNA by intercalation. The planar aromatic part stacks in-between the base pairs, while the daunosamine sugar is positioned at the minor groove.[277,278]

Fig. III-75: Chemical Structure of Dauno- and Doxorubicin.

In a first qualitative test a 1 mM aqueous solution of daunorubicin was mixed with increasing concentrations of all four T2 trisoligonucleotides and assembled under TAP1 conditions (Fig. III-76). The T2 tetrahedron concentration ranged between 0 and 3.1 µM. A positive charge at the ammonium residue of daunorubicin explains the inverted electrophoretic mobility.

Fig. III-76: Intercalation of 1 mM daunorubicin with up to a 3.1 µM T2 tetrahedron; HB; TAP1; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 30 min; 3 % Agarose; 100 V; 30 min; SYBR Gold stain (post-gel).

92 III. Results and Discussion

Daunorubicin disappeared in a discrete gradient with increasing tetrahedron concentrations. The band fully disappeared when mixed with the 3.1 µM tetrahedron solution. This could only be explained by an intercalation process, whereby bound anthracycline was transported by the tetrahedron.

In order to study the influence of the 3D-structure, test results were compared with a non- three-dimensional equivalent of the tetrahedron. All strands in the tetrahedron were resynthesized as a set of twelve 15mer oligonucleotides as descripted in chapter III.2. Each complementary strands of the twelve oligonucleotides were hybridized to six corresponding duplexes (AA’-FF’) under TAP1 conditions. All six dimer duplexes formed almost quantitatively, which was necessary for intercalation experiments (Fig. III-77).

Fig. III-77: Oligonucleotide set 6D. Successful hybridizations to all six double helixes. Faint upper bands attribute to bromophenol blue; Dissolved 1:5 in HB; TAP1 for dimers; 16 % native PAGE; 100 V; 100 min, UV detection.

All twelve oligonucleotide stock solutions were combined to form the oligomix “6D”. The concentration was adjusted to be identical with the tetrahedron in terms of “duplex concentration”, since both contained the same six double-strands. (A 3.125 µM solution correlates to an 18.75 µM duplex solution.)

The first loading experiment with daunorubicin was repeated with the oligomix 6D (Fig. III- 78). Gel electrophoresis revealed lesser loading efficiency than T2 with full loading only achievable by using half the amount of daunorubicin. Similar experiments with doxorubicin showed an identical behaviour in loading capabilities.

An alternative mild loading protocol was established, which allows subsequent incubation with anthracyclines with already assembled constructs (Fig. III-79). The milder conditions are also beneficial in terms of drug stability because exposure at higher temperatures might cause decomposition.[279,280] In protocol “DLP1” anthracycline and hybridized DNA were

93 III. Results and Discussion shaken for 30 min at 30 °C and then stored overnight at 8 °C. T2 and D6 were incubated with dauno- and doxorubicin and showed similar qualitative loading efficiencies as the previous experiments.

Fig. III-78: Intercalation of 1 µL and 2 µL of 1 mM daunorubicin with up to 3.1 µM 6D; TAP1; 3 % Agarose; 100 V; 20 min; GelStar nucleic acid gel stain (in-gel).

Fig. III-79: Direct qualitative comparison of the intercalation of dauno- and doxorubicin with the pre-assembled tetrahedron T2 and the oligomix 6D using alternative loading conditions. 3 % Agarose; 100 V; 20 min; GelStar nucleic acid gel stain (in- gel).

94 III. Results and Discussion

Subsequent loading was therefore as viable as simultaneous hybridization and intercalation under TAP1 conditions. Though equimolar amounts of DNA were used, spots for 6D appeared much bigger than T2. This is explained by the different compositions: 6D was as a mixture of six independent duplexes running, whereas T2 was one discrete object. All duplexes in the tetrahedron moved through the gel in a concerted fashion. Clearly visible in this direct comparison is more residual anthracycline in the loading experiment with 6D than with T2 as in the previous experiments.

III.5.2. Quantitative Analysis I

A method was developed to quantify anthracycline binding efficiencies: A set volume of 1 mM dauno- and doxorubicin was diluted with 100 mM sodium chloride in absence or presence of a set amount of T2 or 6D (~18.75 µM duplex). All samples were incubated using the DLP1 protocol and then centrifuged. A red coloured pellet was clearly visible in the case of T2, while DNA-free solutions remained homogenous (Fig. III-80). Attempts to redissolve the pellet resulted in an inhomogenous suspension and was not used for quantification.

Fig. III-80: Image of pellet formation after centrifugation of a doxorubicin-loaded T2 tetrahedron. Concentrations of each component daunorubicin (Dn), doxorubicin (Dx), tetrahedron (T2) and oligomix (6D) were measured individually by UV spectroscopy (Fig. III-81). This allowed to closely estimate the contents of each component in loaded mixtures of drug and DNA. Prepared DNA samples showed very comparable absorption maxima at 254 nm with very similar concentrations. Drugs showed a very comparable and distinct absorption at 480 nm with no interference of DNA. At 200-230 nm doxorubicin had a weaker absorption than daunorubicin indicative of the small structural difference between those two.

95 III. Results and Discussion

1,8 T2, 6D: 18.75 µM duplex in 100 mM NaCl T2 1,6 Dn, Dx: 1 mM in 100 mM NaCl 6D Dn 1,4 27 µL T2/6D + 3 µL 100 NaCl Dx 3 µL Dn/Dx + 27 µL 100 NaCl 1,2

1,0

0,8

0,6

Absorbance (units) 0,4

0,2

0,0

200 250 300 350 400 450 500 550 600 650 Wavelength (nm)

Fig. III-81: Overlaid UV spectrograms of individual components daunorubicin (Dn), doxorubin (Dx), tetrahedron (T2) and oligomix (6D) under the given setup. Calculated contents were used in the determination of loading efficiencies.

Tab. III-18: Derived concentrations and calculation of anthracycline loading ratios per duplex of T2 and 6D.

96 III. Results and Discussion

Contents of unbound anthracyclines in the supernatant of intercalated samples were quantified via UV spectroscopy at 480 nm and compared to the concentration of pure anthracycline solutions. In combination with the separately measured DNA contents it was possible to calculate how much anthracycline was loaded onto DNA (Tab. III-18). Each spectroscopic measurement was done in a triplicate fashion and averaged to determine the concentrations of the individual components and the unbound anthracyclines in the supernatants of T2 and 6D. For instance, the daunorubicin concentration in the supernatant of T2 was 0.47 mM. The concentration of free daunorubicin solution with the same dilution factor was measured as 1.4 mM. Therefore 66 % of daunorubicin (2.778 nmol) were presumably bound to the duplexes of T2. Duplex concentration was measured as 19.44 µM (0.525 nmol). The ratio between the amounts of bound daunorubicin and duplexes gave a value of 5.29. Thus, five units of daunorubicin intercalated with one duplex of T2. The same setup with doxorubicin confirmed this ratio with a value of 5.06. Ratios for the oligomix 6D averaged around a value of 3.5, which was in compliance with literature results: According to crystallographic studies by Frederick et al.,[281] anthracyclines intercalate every fourth base pair in a double-strand. A 15mer long double-strand can therefore stack either three or four intercalators depending on the loading pattern (Fig. III-82). The results found around the ratio of 3.5 were likely a statistical average of all possible loading patterns.

Fig. III-82: All loading pattern permutations of a 15mer double-strand can either lead to 3 or 4 loaded units, whereby every fourth base pair is stacked.

An entire tetrahedron was capable to intercalate 30 units (6x 5) as opposed to 21 units (6x 3.5) with 6D. This raised the question how trisoligonucleotide duplexes in tetrahedrons were capable to load more units than standard duplexes. It is assumed that each duplex still conformed to the established loading capacity of 3.5 units per 15mer duplex and additional units could only stack somewhere else in the geometry of the scaffold. A conceivable alternative is found at the vertices. More units were probably able to stack between the linker core and the 3’-nucleotides G and C (Fig. III-83). By bending the oligonucleotide arms a small cavity is formed at the linker core offering a secondary binding mechanism.

97 III. Results and Discussion

Fig. III-83: Circled areas show potential cavities for additional intercalation.

If six duplexes can hold 21 units, 9 more units need to be stored in the tetrahedron. Each of the four cavities could possibly store around two units, but one more unit would still remain. This could either be considered to be within the error of estimation or by an orientation effect of the duplexes: Though anthracyclines are much smaller than the tetrahedron and could tunnel through the scaffold, the intercalation process is more likely initiated from the outside. It is conceivable that the fixed orientation of duplexes in the tetrahedral scaffold in conjunction with the preferred angle of attack might influence the stacking pattern leaning more towards higher duplex loading. Free duplexes, on the contrary, are fully exposed from all angles for intercalation, which explains the statistical mean distribution between three and four units per duplex.

III.5.3. Quantitative Analysis II: Fluorescence Spectroscopy

An alternative method was established to validate the increased anthracycline binding capabilities of trisoligonucleotide-based tetrahedral scaffolds in comparison to linear duplexes. It is known from literature that fluorescence of anthracyclines is quenched when bound by DNA.[282] Remaining fluorescence is attributed solely to free unbound anthracyclines. This non-invasive method allowed to determine binding efficiencies of anthracyclines in homogenised samples without the need of prior centrifugation as done in the previous method (chapter III.6.1). A set amount of dauno- and doxorubicin was incubated with varying concentrations of tetrahedrons T2, T3 and the oligomix 6D as part of titration experiments to observe the quenching effect and compare binding efficiencies.

Aqueous stock solutions of T2 (5.32 µM), T3 (5.79 µM) and 6D (5.81 µM) were diluted with 100 mM sodium chloride to give a gradient of concentrations including 80, 60, 40, 30, 20 and

98 III. Results and Discussion

10 percent of the stock solutions. Concentration of each dilution was determined by UV/VIS spectroscopy and the results were plotted against the percentage of dilution in regards to the stock solution (Fig. III-84). The calibration plots showed comparable concentrations between both tetrahedrons and the oligomix across all dilutions with very good linearity.

Fig. III-84: Left: Calibration plots for T2, T3 and 6D dilutions. Stock solutions (100 %) were diluted with 100 mM sodium chloride to give 80/60/40/30/20/10 % solutions; Right: Calibration plots for Dn and Dx dilutions. Stock solutions (100 %) were diluted with ddH2O to give 50/25/10/2.5/1.0/0.5 % solutions; All measurements in 1 mM HEPES (pH: 7.5) at 25 °C. Additional concentration gradients were prepared for daunorubicin (871 µM) and doxorubicin (805 µM) including 50, 25 10, 5, 2.5, 1 and 0.5 percent dilutions in water. Concentrations were determined via UV/VIS spectroscopy. These calibration plots also showed high linearity along the gradients (Fig. III-84).

Fig. III-85: Fluorescence emission spectra of anthracycline concentration gradients. All measurements in 1 mM HEPES (pH: 7.5) at 25 °C with a dilution factor of 5/11.

All fluorescence emissions of anthracyclines were detected with an excitation wavelength of λex = 410 nm. This allowed to record the whole emission spectra between 500 to 800 nm

99 III. Results and Discussion

without interference of the water Raman band at λmax = 476 nm, which is according to literature[282] only partially fixable by baseline recordings and might distort the signals at low anthracycline concentrations. Emissions of all dauno- and doxorubicin dilutions were measured (Fig. III-85). Profiles showed two peaks at 550 and 592 nm with decreasing intensity at lowered concentrations. Emission intensity at 592 nm of each profile was then plotted against the corresponding sample concentrations (Fig. III-86).

Fig. III-86: Concentration of anthracyclines plotted against corresponding emission intensity. The right plot shows samples measured at a 1.8 times higher concentration than the left one. Boltzmann equation for sigmoidal fit: y = A2 + (A1-A2)/(1 + exp((x-x0)/dx)). All measurements in 1 mM HEPES (pH: 7.5) at 25 °C with a dilution factor of 4/55 on the left and 8/60 on the right. The left shows a linear behaviour between the fluorescence signal and anthracycline concentrations up to 400 µM solutions. The right plot shows the same samples but measured at almost double the concentration revealing a stronger asymptotic behaviour towards high concentrations.

Profiles of quenched samples were compared relatively to each other and to free anthracyclines. A given amount of anthracyclines (Dn: 3.48 nmol; Dx: 3.22 nmol) were mixed with a given volume of hybridized DNA (T2, T3 and 6D) of varying concentrations and incubated using the ”DLP1” protocol (chapter III.5.1.) The varying DNA concentrations refer back to the calibration plots (Fig. III-84). Two additional samples of anthracyclines were incubated and measured labelled as “R = 0.000”. These samples were diluted only in 100 mM sodium chloride to give profiles of fully free anthracyclines as a reference for completely unbound material.

100 III. Results and Discussion

Fig. III-87: Graphs of fluorescence emission for anthracycline quenching titration experiments. Left column: Daunorubicin quenching, Right column: Doxorubicin quenching, Top row: Intercalation with Tetrahedron T2, Middle row: Tetrahedron T3, Bottom row: Oligomix 6D; All measurements in 1 mM HEPES (pH: 7.5) at 25 °C with a dilution factor of 5/11. Signal intensity in all samples decreased with increasing amounts of DNA. Weaker signals correlate to stronger quenching of the drugs due to an increased amount of material binding to DNA. It appeared that the oligomix 6D loaded less material than the tetrahedrons T2 and T3. Another way of visualising this discrepancy was to calculate the ratio of peak areas (500- 800 nm) between the unbound anthracyclines in the mixtures and the free anthracycline (“R = 0.000”).

101 III. Results and Discussion

Fig. III-88: Binding efficiency plots. Unbound anthracycline is unquenched material in mixtures with DNA and free anthracycline is DNA-free material.

Fig. III-89: Graphs of fluorescence emission for anthracycline quenching titration experiments with only 0.75 and 0.9 equivalents of the previously used amount of anthracyclines. Point of full intercalation was reached earlier. Dilution factor of 3/10.

102 III. Results and Discussion

The percentage of bound anthracyclines was then plotted against the corresponding ratios (Fig. III-88). With the exception of one outlier (“Dn@T2, R = 0.080”) both tetrahedrons showed a very comparable quenching effect for each anthracycline, whereas the oligomix 6D displayed a noticeably smaller binding efficiency. (See Appendix for an alternative visualisation.)

It appeared that full intercalation of the tetrahedrons was reached with ratios higher than R = 0.029. Noticeable was a residual fluorescence near the baseline, which did not disappear at higher DNA concentrations. Anthracyclines bind stronger to GC base pairs than to AT.[282] An exchange equilibrium is assumed, in which as small portion of bound anthracyclines near AT base pairs leaves the DNA and is unaffected by quenching. This was confirmed with titrations that used less amounts of anthracyclines and therefore reached the point of full intercalation earlier (Fig. III-89). These measurements were also necessary to estimate the quantitative intercalation of 6D.

Based on the given data it was now possible to calculate binding efficiencies of both drugs on all three DNA carriers (Fig. III-90).

Fig. III-90: Calculated binding efficiencies. Left column gives the number of units loaded per tetrahedron and per oligomix (six duplexes). Right column gives the amount per 15mer duplex. Data with asterisk represent values calculated with hypothetical 90 % DNA solutions.

103 III. Results and Discussion

The amount of anthracycline in each sample was divided by the amount of DNA necessary to reach full quenching, which in all cases lied between 80 to 100 % of the used DNA stock solutions. Hypothetical “90 %” DNA solutions were calculated for most cases serving as a middle value between the lower and upper limit. Division by six gave the amount per duplex. These values closely resemble the findings of the other quantification method with tetrahedrons showing an increased binding efficiency over the linear oligonucleotide duplexes of 6D. Tetrahedron T2 loaded about two units more than T3. Building blocks of T3 showed stronger decomposition patterns on polyacrylamide gels than T2, which led to more defect tetrahedrons and therefore less capable of binding.

Samples of the quenching experiments were additionally studied via UV/VIS spectroscopy (Fig. III-91). Absorption increased around 260 nm with increasing concentrations of DNA. The most salient change was visible in the anthracycline absorption area between 350 to 600 nm: With increased binding a redshift from 475 nm towards 508 nm was visible explained by the π-stacking effect between the purine bases and the aromatic part of the anthracyclines. No difference between tetrahedrons and the linear oligonucleotide duplexes was observed. A shift caused by the proposed secondary binding mechanism located at the vertices of the tetrahedron is presumably too weak and/or too similar compared to the shift caused by the primary duplex binding mechanism via duplex intercalation.

104 III. Results and Discussion

Fig. III-91: UV/VIS spectroscopic analysis of samples used in the quenching experiments. Top example displays an absorption range between 190 to 600 nm and the two bottom ones show stacked plots of the distinct anthracycline absorption between 350 to 600 nm.

105 V. Experimental Section

IV. Summary and Outlook

Practical work on the field of trisoligonucleotide-based chemistry is divided into three phases: 1.) Chemical synthesis of trislinkers and solid supports; 2.) DNA synthesis, extraction and purification of trisoligonucleotides; 3.) Assembly experiments with trisoligonucleotides and structural studies of derived constructs. The first two phases are inevitable preparation steps and one ongoing effort lies in their streamlining and simplification. This work contributed to all three phases.

The previously established synthesis of the 1,3,5-tris(3-hydroxypropyl)benzene trislinker

[159] was lengthy and laborious. It was possible to simplify synthesis of this C3h-trislinker by investigating a Heck-type coupling reaction between 1,3,5-tribromobenzene and allylbenzylether, which was readily available via an ether-type reaction. This was then followed by a one-step hydrogenation and hydrogenolysis to gain the unprotected C3h- symmetrical trislinker (Fig. IV-1). The next steps followed established procedures to give the orthogonally protected phosphoramidite of this trislinker.

Fig. IV-1: Convergent synthesis over three steps to yield the unprotected 1,3,5-tris(3-hydroxypropyl)benzene trislinker core.

106 V. Experimental Section

Trislinkers based on 1,3,5-tris(3-hydroxymethyl)benzene, 1,3,5-tris(3-hydroxyethyl)benzene and the one based on 1,3,5-tris(3-hydroxypropyl)benzene (Fig. IV-2) were part of a study to systematically analyse the effect of varying side chain flexibility in trisoligonucleotide-based tetrahedral scaffolds (T1, T2, T3). The first two orthogonally protected trislinkers were available[202,203] and only the corresponding phosphoramidites needed to be prepared.

Fig. IV-2: Prepared phosphoramidites of three orthogonally protected trislinkers with varying alkyl chain lengths. DNA tetrahedra were seen as ideal molecular scaffolds to study relative flexibility effects because they possess high mechanical rigidity and structural stability. Three trislinker sets of four trisoligonucleotides were prepared. Chosen sequences were identical across all three sets for optimal comparability. (Yields of most trisoligonucleotides ranged between 60 to 165 nmol.) Self-assembly experiments followed by enzymatic digestion with mung bean nuclease (Fig. IV-3; “+E”) confirmed trisoligonucleotides based on the T3 trislinker to form stable tetrahedral scaffolds as previously described, whereas an exemplary trimer (“123”) with abundant single-strands showed digestion as expected.[159] Although the T2 trislinker was previously available it was never applied in trisoligonucleotides and studied in assembly experiments. The hybridized trisoligonucleotide set based on the T2 trislinker showed stability Fig. IV-3: Trimer 123 of the T3 set, tetramers T1, T2 towards enzymatic digestion for at least 10 to 30 and T3; HB, TAP1; “+E”: Incubation of sample to the left with 5 units of mung bean nuclease for 10 min at minutes. This work was able to demonstrate that 30 °C; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 2 h; GelStar nucleic acid gel stain (in-gel). ethyl chains in trislinkers are flexible enough to

107 V. Experimental Section fold into constrained tetrahedral scaffolds. Trisoligonucleotides based on the T2 trislinker can potentially be used in less constrained polyhedral scaffolds like octahedra or dodecahedra in the future. Also confirmed was the T1 trislinker incapable to fold into a tetrahedral scaffold, due to a high rigidity of the methyl chains as previously described.[202] These studies revealed a lower limit in DNA nanoconstruction that might be of consideration in future studies of constraint jet stable structures.

Partial stability of T1 trisoligonucleotides was found in intermixing experiments: The identical sequence design between the sets allowed cross-mixing with the more flexible and stable T3 set. Interestingly, the substitution of just one trisoligonucleotide alone showed a major increase in stability with only minimal digestion. Apparently, the more flexible T3 propyl chains are able to compensate the rigidity of T1 methyl chains for the most part and allowed the formation of a stable tetrahedron. Quantity of stable tetrahedron increased by substituting with another T3 trisoligonucleotide.

Since ethyl chains have shown enough flexibility, an entirely new generation of trislinkers was established with the commercially available 1,3,5-tris(2-hydroxyethyl)-N,N’,N’’- isocyanurate (THEIC) abolishing any synthesis up to the protection group chemistry.

Two trislinkers were prepared with THEIC (Fig. IV-4). One orthogonally protected containing two DMT-groups (TD) and an orthogonally protected one with an AOC- and a DMT-group (TN).

Fig. IV-4: Isocyanurate based trislinkers. Orthogonally protected (TN) and non-orthogonally protected (TD).

108 V. Experimental Section

Flexibility and stability of the TN trislinker in trisoligonucleotides was tested similarly to the T1- T3 sets. A set of four trisoligonucleotides with identical sequences to the previous sets were synthesised and studied in self-assembly experiments. The assembled product of all four trisoligonucleotides (Fig. IV-5, “TN”) showed no digestion in presence of the endonuclease for at least 20 minutes unlike an exemplary test trimer (“123+E”). All strands were therefore hybridized, which indicated a fully closed tetrahedral scaffold. This finding marked THEIC as a new generation of versatile C3h-symmetrial trislinkers compatible Fig. IV-5: Digestion experiments with the trimer 123 and sufficiently flexible for future use in of the TN set and tetramer TN; HB, TAP1; 5 U MBN at 30 °C; 1.25 µM; Nat. 3 % Agarose 1000, 1x TBE, 100 V, 65 trisoligonucleotide-based nanostructures. min; GelStar nucleic acid gel stain (in-gel).

With the TD trislinker a new concept of semi-addressable sequence patterns in trisoligonucleotides was investigated (Fig. IV-6). This sequence pattern is characterized by two arms with an identical sequence and one arm with a unique sequence, coined “DOS”, and lies in between the established patterns with either three unique, “UNO”, or three identical, “TRE”, sequences. Assemblies involving the DOS-pattern are closer to maximal instruction than with the TRE-pattern.

Fig. IV-6: Schematic overview of all three conceivable trisoligonucleotide sequence patters. UNO is characterized with three unique sequences, DOS with two and TRE with only one. UNO has a sequence X one time, DOS twice and TRE three times.

The behaviour of the DOS-pattern was studied in an array of hybridization experiments. A set of complementary DOS-trisoligonucleotides formed higher order aggregates, when established protocols at high temperatures were applied. Mild conditions near room

109 V. Experimental Section temperature (10 min @ 25 °C) gave a primary product, which was identified as having nano- cyclobutadiene topology. This motif was also identified in experiments with self- complementary TRE-pattern. Elaborate hybridization protocols were used in the assembly of trisoligonucleotides for two decades, which involved denaturation- and cooling steps programmed onto thermocyclers. It was successfully demonstrated that simple shaking at 25 °C for 10 minutes of the buffered building blocks is completely sufficient to assemble into tetrahedral scaffolds explained by the full sequence addressability of the UNO-pattern.

In combination with extensive cross-hybridization experiments between UNO- and DOS trisoligonucleotides different new discrete motifs were discovered and named (Fig. IV-7):

Fig. IV-7: Newly discovered discrete trisoligonucleotide-based motifs.

The introduction of the DOS-pattern enhanced the understanding of trisoligonucleotide- based self-assemblies into nanoconstructs and expanded the repertoire of conceivable motifs. In addition, synthesis of DOS-trisoligonucleotides is achievable on commercially available reverse CPG support since a TD trislinker lacks the incompatible alloc-group. This was confirmed by Michalski in studies with the T2 trislinker.[221]

This work also studied a purification method involving perfluorous tags. Fluorous affinity purification methods could not successfully separate tagged trisoligonucleotides from tagged mutants. Tagging did allow for first time to successfully isolate trisoligonucleotides via liquid chromatography. A new method was developed to efficiently cleave the sulfide- bearing fluorous tag on model linear oligonucleotides, which involved hydrogen peroxide as the oxidizing agent and sodium tungstate in a double role as catalyst and base. Application

110 V. Experimental Section of this method on tagged trisoligonucleotides, however, showed only partial conversion and patterns of decomposition. Final untagged material was then isolated via liquid chromatography. Although this method was technically successful, it was deemed not competitive to preparative gel electrophoresis due to a lengthy procedure and inefficiency.

Intercalation experiments between trisoligonucleotide-based tetrahedrons and anthracyclines (dauno- and doxorubicin) revealed improved binding efficiencies compared to a set of linear DNA duplexes of identical sequence design. This was observed in UV/VIS- and fluorescence-spectroscopy. A secondary binding mechanism is assumed by which additional units of intercalators can load at the vertices of the tetrahedron (Fig. IV-8) exploiting the fixed three-dimensional orientation of the duplexes. In the case of a 90 bp sized tetrahedron 18-24 anthracycline units are bound to the 15mer duplexes and another 8 units at the four vertices (4x2).

Fig. IV-8: Circled areas near the trislinker mark potential cavities for additional binding of intercalators via secondary loading mechanism.

Trisoligonucleotide-based tetrahedral scaffolds can potentially contribute to a new class of very small biocompatible nanoscale carriers for anti-cancer agents with an effect to improve cellular uptake and increased bioactivity in comparison to much larger origami carriers. Bio- assay experiments on cancer-cells like MCF-7 are needed for that matter. Studies on mammalian cells can give information on whether or not loaded tetrahedrons are stable enough in the cytoplasm and do not cause any toxic side effects mainly induced by the trislinker element.

111 V. Experimental Section

V. Experimental Section

V.1. Methods and Equipment V.1.1. Chromatographic Methods V.1.1.A. Thin-Layer Chromatography

Pre-cut TLC plates SIL G/UV254 on aluminium sheets by Macherey & Nagel (40x80 mm, 0.2 mm silica gel, fluorescent indicator) were used. Application of substance onto the plates was done with capillary tubes originally designed for the determination of melting points (80x0.6 mm, Marienfeld-Superior). Detection of spots was primarily done with UV-light irradiation at 254 nm (N-4 K, Benda). Staining methods included treatment with acid vapour (HCl) for compounds containing DMT or elementary iodine vapour (iodine chamber) or spraying a ninhydrin solution (for isocyanurate educt) with subsequent heating (0.3 g, 100 mL methanol).

V.1.1.B. Column Chromatography

Manual column chromatography used Silica 60 M (40-63 µm) by Macherey & Nagel. Columns were packed with a 1 cm thick layer of sand prior to the addition of the silica/eluent slurry. No sand or other buffering materials were added after applying the raw product onto the silica. Pressure was applied manually for flash chromatography with a hand pump if necessary.

Automated separation was done on the flash chromatography system Reveleris X2 by Grace. Columns came as pre-packed silica (40 µm) disposable cartridges. Depending on sample mass different sizes of cartridges were used. The flowrate of the eluent was typically set lower than theoretically possible to avoid the risk of very high pressure on the column (>30 psi; >60 psi for 120 g column).

4 g silica: 4 mg - 0.8 g sample; 20 mL/min flowrate

12 g silica: 12 mg - 2.4 g sample; 24 mL/min flowrate

40 g silica: 40 mg - 8 g sample; 30 mL/min flowrate

120 g silica: 120 mg - 24 g sample; 70 mL/min flowrate

112 V. Experimental Section

ELSD: 20 mV; Carrier: Isopropanol; Medium sensitivity slope detection

UV: 0.05 AU; 254 nm, 280 nm

Typical protocol A: Typical protocol B:

4.0 min: equilibrium 4.0 min: equilibrium

10.0 min: 5 % - 30 % CH/EE 13.0 min 1 % - 20 % DCM/MeOH

2.0 min: 30 % - 100 % CH/EE 5.0 min 20 % - 100 % DCM/MeOH

2.0 min: 100 % EE 3.0 min 100 % MeOH

5.0 min: 100 % MeOH

Acid labile products were purified in the presence 0.5 % of NEt3 added to either CH or DCM.

V.1.1.C. High-Pressure Liquid Chromatography

Kontron HPLC System

Pump: 420 (Kontron Instruments)

Mixing Chamber: M 800 (Kontron Instruments)

Autosampler: 465 (Kontron Instruments)

UV-Detector: Multi-Channel Diode Array Detector 440 (Kontron Instruments)

Software: Kroma System 2000

Column RP-HPLC: Nucleodur 100-5 (C18; 250x4 mm)

Macherey & Nagel

Column F-HPLC: FluoroFlash (FPC8; 150x4.6 mm)

Fluorous Technologies

Oligonucleotides were detected at 240, 254, 260, 280 and 300 nm.

Elution solvent: A: Acetonitrile

B: 0.1 M (NH4)HCO3 (pH: 8.5)

Gradient A: 5-80 %, ACN/AHC, 35 min

Gradient B: 20-30 %, ACN/AHC, 30 min

113 V. Experimental Section

V.1.2. Spectroscopic Methods V.1.2.A. Nuclear Magnetic Resonance Spectroscopy

NMR-Spectrometer: DPX-200 (200 MHz), Bruker

DPX-250 (250 MHz), Bruker

DRX-400 (400 MHz), Bruker

Deuterated solvents: CDCl3, DMSO-d6, D3OD (All supplied by Deutero GmbH)

Typically 20-50 mg of substance were solved in ~0.7 mL solvent.

Labels for multiplicity: s = singlet; d = doublet; t = triplet; q = quartet; p = quintet;

dd = doublet of doublet; dt = doublet of triplet; m = multiplet

br = broad signal; xJ = Coupling constant in Hz along x bonds

Shifts of deuterated solvent served as an internal standard for calibration of spectra[283,284] and tetramethylsilane (TMS) as the shift reference standard. The chemical shift is given in ppm. 1H-, 13C- and 31P-spectra were measured. The latter two were proton decoupled.

V.1.3.B. UV/VIS-Spectrophotometry

UV spectra were recorded on the photometer Cary 1E (Varian) at room temperature and pH: 7. Samples were measured in cuvettes with a 10 mm light path (quartz SUPRASIL;

Helma) and typically diluted in ddH2O. Contents of samples were determined via the Lambert-Beer law (E = εcd; E: extinction; ε: [L mol-1 cm-1]; c: [molL-1; c: [cm]). The anthracyclines doxo- and daunorubicin shared an identical extinction coefficient ε of 11500 L mol-1 cm-1 at 480 nm. For oligonucleotides the increments of the extinction coefficient ε at 254 nm were added up for each nucleotide:[249]

-1 -1 Nucleotide ε254nm increment [L mol cm ]

C 6541

T 7250

A 13200

G 13679

The absorption of the trislinker in trisoligonucleotides was not taken into account.

114 V. Experimental Section

V.1.3.C. Fluorescence Spectroscopy

Measurements were conducted on the fluorescence spectrophotometer Cary Eclipse (Varian) in conjunction with the Cary Temperature Controller (Varian). Samples were measured in the cuvette Cary 6Q (Varian). Fluorescence emission of anthracyclines was detected at an excitation wavelength of 410 nm in a 500 to 800 nm range at 25 °C. All samples were diluted in 1 mM HEPES (pH: 7.5). PMT detector voltage was set to 800 V (“high” preset). Excitation and emission slits were set to 5 nm each with an open emission filter,

V.1.3. Mass Spectrometric Methods V.1.3.A. Matrix-Assisted-Laser-Desorption-Ionization-Time-Of-Flight-MS

MALDI-TOF-MS data were collected on the autoflex mass spectrometer (Bruker Daltonics):

-6 N2-Laser LTB MNL106 (337 nm); ~25 kV acceleration voltage; 5x10 mbar vacuum; positive ion mode [M+H]+; Target: MTP 384 target plate ground steel (Bruker Daltonics); Flex Control 2.0 (Build 51); Flex Analysis 2.0 (Build 21).

Matrices:

Linear oligonucleotides: 2,4,6-trihydroxyacetophenone (THAP)

Trisoligonucleotides: 3-hydroxypicolinic acid (HPA)

80 mg of HPA or THAP were diluted in 2 mL ddH2O/ACN (1:1).

Typical sample preparation:

1.5 µL of the aqueous sample (1-10 pmol/µL) was mixed with a few beads of an ion-exchange

+ resin (NH4 ) on top of a plastic paraffin film (Parafilm; Pechiney Plastic Packaging). After ~15 seconds 1 µL of the supernatant was mixed with 3 µL of the HPA or THAP matrix on top of the parafilm. 2-4 µL of the mix was then placed on the target and allowed to dry at room temperature for at least 30 minutes. 30-100 laser shots were applied per sample at 50-75 % laser power.

V.1.3.B. Fast-Atom-Bombardment-MS

Autospec (VG Instruments); ~8 kV acceleration voltage; Cs+ bombardment; Matrices: 3- nitrobenzylalcohol, glycerin, lactic acid. Relative intensity in % is added in brackets.

115 V. Experimental Section

V.1.4. Electrophoretic Methods V.1.4.A. Buffer

1x TBE was used as the buffer, which was diluted with ddH2O from a 5x TBE stock solution. This stock solution contained 269.5 g TRIS, 137.5 g boric acid and 17.5 g EDTA (Titriplex III; pH: 8) in 5 L deionized H2O. The buffer was stored at room temperature for polyacrylamide gels and at ~8 °C for agarose gels.

V.1.4.B. Documentation

Stained gels were detected with the UV transilluminator T2201 (Sigma) and observed behind a plexiglas window. Images of stained agarose gels were recorded on the transilluminator UVT-20 M/W (Herolab) in a bioimager by Intas and controlled with the Intas GDS software.

V.1.4.C. Analytical Polyacrylamide-Gel Electrophoresis

Analytical denaturing and native PAGE were executed on the vertical electrophoresis chambers Mini-Protean 3 and Mini-Protean Tetra Cell (Bio-Rad) in conjunction with the power supplies E 865 (Consort) or EPS 3500 (Pharmacia Biotech). All gels were handcast and used 40 % PAA (19:1 acrylamide/bisacrylamide):

12 % denaturing PAGE (One gel):

4.166 g urea, 2 mL 5x TBE, 2.4 mL deionized H2O, 3 mL PAA (40 %)

16 % denaturing PAGE (One gel):

4.166 g urea, 2 mL 5x TBE, 1.9 mL deionized H2O, 4 mL PAA (40 %)

16 % native PAGE gel:

2 mL 5x TBE, 3.9 mL deionized H2O, 4 mL PAA (40 %)

The given amounts were sufficient for one gel, but typically twice of the setup was used for casting two gels simultaneously. Materials stirred in a beaker until the urea fully dissolved. A glass petri dish covered the beaker. Polymerization was induced with APS and TEMED:

Polymerization starter:

35 µL TEMED, 70 µL APS (10 %)

The mixture was filled in vertical casting plates (10.0x8.0x0.15 cm) on a stand. A 10-well comb was then attached to the gel. Successful polymerization was indicated by an exothermic

116 V. Experimental Section reaction. The polymerization finished after the gel cooled down to room temperature. Samples for native electrophoresis were usually diluted 1:10 in HB (100 mM NaCl + 10 mM HEPES, pH: 7.5) and for denaturing electrophoresis in saturated urea-solution instead. 1 µL of Gel Loading Buffer (0.05 % bromophenol blue; 40 % sucrose; 0.1 M EDTA pH: 8.0; 0.5 % SDS; Sigma) was added to native samples. Gel Loading Solution Type I (0.25 % bromophenol blue, 0.25 % xylene cyanol, 40 % sucrose; Sigma) was diluted 1:5 with 1x TBE and served as a progress marker. Electrophoresis was executed at 100-120 V for 60-100 min at room temperature without external cooling. Gels were post-stained at dark in SYBR Gold (1 µL/10 mL 1x TBE; Invitrogen) for 30-60 min at ~8 °C if needed.

V.1.4.D. Preparative Denaturing Polyacrylamide-Gel Electrophoresis and Extraction

For vertical preparative electrophoresis Protean II xi cell (Bio-Rad) was used for the purification of raw trisoligonucleotides (1.3 µmol) in conjunction with the power supplies E 865 (Consort) or EPS 3500 (Pharmacia Biotech). The cooling core was attached to the cooling unit Ultratemp 2000/Julabo F30 (Gebr. Rettberg GmbH). Denaturing polyacrylamide gels with 40 % PAA (19:1 acrylamide/bisacrylamide) were prepared manually. The material was split on two gels to avoid overloading.

12 % denaturing PAGE (Two gels):

100 g urea, 40 mL 5x TBE, 20 mL deionized H2O, 60 mL PAA (40 %)

Materials were mixed in a beaker and heated to 50 °C under stirring until all urea dissolved. A glass petri dish covered the beaker. In the meantime the glass plates and spacers (20x20x0.3 cm) were set up on a vertical casting stand. The mixture was allowed to cool down and the polymerization starter was then added.

Polymerization starter (Two gels):

200 µL TEMED, 500 µL APS (10 %)

The mixture was cast into the glass plates and 2D combs were attached on top of the gels and a comb that contained two reference wells. Polymerization was accelerated by evenly heating the outer glass surface with a heat gun (Typ 3458; Steinel). After successful polymerization (30-60 min) the gels were allowed to cool down to room temperature and placed onto the cooling core of the electrophoresis cell and into the buffer tank. Gels had a 15 min pre-run at 450 V and at a temperature of 15 °C. In the meantime 100 µL of aqueous raw samples were diluted with 300 µL of aqueous saturated urea solution and heated to 55 °C

117 V. Experimental Section for one minute with a thermomixer. 200 µL of the samples were applied to each gel. 3 µL Gel Loading Solution Type I (Sigma) was diluted 1:5 with the urea-solution and added to the outer wells as a reference. Electrophoresis was executed at 450 V for 3-4 hours at 15 °C. Trisoligonucleotides typically appeared roughly on the same level as the slower xylene cyanol dye. The gels were placed on 20x20 cm TLC silica 60 plates (F254; e.g. Macherey & Nagel) wrapped in cling film. DNA-bands were detected under UV-light.

The desired band was the slowest and all bands were slightly curved. It was extracted with a scalpel. In order to not carry over mutants only ¾ of the upper part of band was extracted. Gel extracts were placed in a 50 mL centrifuge tube and crushed manually with a spatula.

50 mL of ddH2O were added and stirred for 2 min on a vortex. The gel-water suspension was centrifuged for 15 min at 5500 rpm (Beckmann) and the supernatant was collected in a 250 mL round-bottom flask. Occasionally, the supernatant contained traces of gel, which were also collected. The process was repeated. Again 50 mL of water were added and shaken into a gel-water suspension, but was then placed on a shaker for 16 h prior to centrifuging. The collected aqueous sample (~150 mL) was evaporated into a white residue and then dissolved again in 5 mL ddH2O (~8 mL final volume). The extracts were split into three aliquots and desalted with three NAP-25 columns (Illustra; Sephadex G-25 DNA grade; GE Healthcare) using standard protocols by the manufacturer. Desalting was repeated with NAP-10 columns if necessary. The combined solutions were evaporated and aqueous stock solutions (125 µM) were prepared.

V.1.4.E. Native Agarose-Gel Electrophoresis

Agarose gel electrophoresis was executed on the horizontal system Mini-Sub Cell GT (Bio- Rad). 20 mL of 1x TBE-buffer were added to 0.6 g of 3 % Agarose 1000 (Invitrogen) and stirred until the suspension homogenized (~10 min). It was then heated in a microwave (Micromat Series; AEG) to boiling and then heated in short intervals (~10 sec) until the agarose was fully dissolved. 1 µL GelStar Nucleic Acid Stain (Lonza) was added for in-gel staining and the mixture stirred until it cooled down to 40-50 °C. The warm solution was then cast into a 7x7 or 7x10 cm tray placed in a gel caster (Sub-Cell GT UV-Transparent Mini-Gel Trays, Bio-Rad) and either an 8- or a 15-well comb was then added either on top or in the middle of the electrophoresis path. The gel was covered in aluminium foil and allowed to cool down to room temperature. The gels had a thickness of 4-7 mm depending on the added volume and chosen tray. The gels had a pre-run for 15 min at 100 V and at ~8 °C in a cooling chamber. To each sample 1 µL of Gel Loading Buffer was added (0.05 % bromophenol blue; 40 % sucrose;

118 V. Experimental Section

0.1 M EDTA pH: 8.0; 0.5 % SDS; Sigma).The addition of the GeneRuler Low Range DNA Ladder (25-700 bp; 50 µg; Life Technologies) served as an internal reference for DNA and as an external reference with the added dyes.

10x Stock DNA ladder solution:

47 µL 1x TBE; 10 µL 6x DNA Loading Dye (10 mM Tris-HCl (pH: 7.6), 0.03 % bromophenol blue, 0.03 % xylene cyanol FF, 60 % glycerol and 60 mM EDTA); 3 µL GeneRuler Low Range DNA Ladder in Storage Buffer (10 mM Tris-HCl (pH: 7.6), 1 mM EDTA).

Electrophoresis was executed at 100 V and ~8 °C in a cooling chamber for 30-120 min. Gels were post-stained in SYBR Gold (Invitrogen; 1 µL/ 10 mL 1x TBE) in the dark at ~8 °C for 30- 60 min if needed.

V.1.5. Oligonucleotide Synthesis

0.1 M nucleoside phosphoramidites A, C, G and T in acetonitrile were used in oligonucleotide synthesis. Standard protocols for detritylation, activation, oxidation and capping were applied in all cases.

Detritylation: 3 % dichloroacetic acid in dichloroethane

Activation: 0.25 M 5-benzylmercaptotetrazole in acetonitrile (trisoligonucleotides)

0.25 M 4,5-dicyanoimidazole in acetonitrile (linear oligonucleotides)

Oxidation: 0.52 g iodine, 12 mL 2,4,6-collidine, 59 mL ddH2O, 129 mL acetonitrile

Capping I: 13.2 g DMAP in 200 mL acetonitrile

Capping II: 20 mL acetic anhydride, 20 mL 2,4,6-collidine, 166 mL acetonitrile

Flush: Dichloroethane (dried over CaCl2); Acetonitrile (refluxed with CaH2 and distilled)

V.1.5.A. Linear Oligonucleotides

The synthesis of linear 3’-5’-oligonucleotides was conducted on a MerMade 6 DNA/RNA synthesizer (AME Bioscience). The 5’-solid support was LCAA-functionalized dC-, dG- and dT-CPG (500 Å; ~46 µmol/g; Glen Research) for a 1.3 µmol scale synthesis. Every 3’- nucleoside phosphoramidite (Glen Research) was coupled for 1.5 min each. The standard

119 V. Experimental Section procedures were applied to linear oligonucleotides in trityl-on mode. Fluorous tag amidites (0.2 M; ACN) were coupled twice for 5 min after a final detritylation step.

V.1.5.B. Trisoligonucleotides

The synthesis of branched oligonucleotides was done on the synthesizer Gene Assembler Plus (Pharmacia Biotech). The custom 3’-solid support was based on Primer Support 200 Amino (GE Healthcare) with 3’-end dC-, dG- and dT-starter nucleosides (~50 µmol/g) on a 1.3 µmol scale. The first arm was synthesized in 5’-3’ direction using reverse 5’-nucleoside phosphoramidites (ChemGenes). The first two 5’-amidites were coupled twice for 5 min and the following only once. DMT deprotection was done twice after each reverse nucleoside phosphoramidite with 6 % dichloroacetic acid in dichloroethane. The synthesis continued after final detritylation of the first arm by coupling of 0.2 M trislinkeramidites in acetonitrile twice for 5 min each. The direction switched to 3’-5’ with the use of 3’-nucleoside phosphoramidites (Glen Research). The synthesis continued at the second arm for orthogonally protected linkers. Each 3’-phosphoramidite coupled for 1.5 min. DMT deprotection was now done once after each nucleoside phosphoramidite with 3 % dichloroacetic acid in dichloroethane. Alloc was deprotected by flushing with a rate of 0.5 mL/min for 15 min a mixture of 17.1 mg 1,2-bis(diphenylphosphino)ethane, 24.7 mg bis(dibenzylideneacetone)palladium(0) (~4.3 mM) and 10.7 µL pyrrolidine (~13 mM) in 10 mL of ACN. The first 3’-nucleoside phosphoramidite was coupled three times for 15 min each and every subsequent one three times for 2 min. Trisoligonucleotides were typically detritylated. F-Tags (chapter III.5.2.) had a concentration of 0.2 M in acetonitrile and coupled twice for 5 min each after the second arm and three times for 5 min at the end of the third arm. In the case of double DMT protected trislinkers the second and the third arm were synthesized simultaneously. Each 3’-phosphoramidites coupled twice for 2 min.

V.1.5.C. Deblocking and Cleavage

The cartridges containing the solid supports were placed in 1.5 mL micro tubes (PP; conical bottom) and briefly centrifuged to remove residual acetonitrile. The entire cartridges were placed into a 2 mL micro tube (PP) with a gasket. 1 mL of ammonia (33 %) was added and the tube was firmly sealed with a screw cap. The tube was briefly centrifuged in order for the ammonia to fully permeate the solid support. Tubes were placed into a thermomixer and incubated typically for 16 h at 55 °C. The top was sealed with aluminium foil.

120 V. Experimental Section

V.1.5.D. Purification

After deblocking and cleavage all tubes were cooled down to room temperature by putting them into a freezer for a few minutes. The samples were centrifuged again and the supernatant ammonia solution was collected. The cartridge was placed again into a 1.5 mL tube and centrifuged to collect further solution inside the solid support and washed twice with 100 µL ddH2O each. All ammonia and water was combined and evaporated in a vacuum centrifuge (Eppendorf) overnight at 40 °C. The residues of all samples were resolved in

100 µL ddH2O. Quality was controlled with analytical denaturing PAGE. Trisoligonucleotides were then purified via preparative denaturing PAGE as previously described (V.1.4.D.). Linear DMT-on oligonucleotides were applied to Glen-Pak DNA Purification Cartridges (Glen Research) using a slightly modified standard procedure by the manufacturer: A) 2 mL of 100 mg/mL NaCl was applied instead of 1 mL; B) The Salt Wash Solution, the 2 % TFA and ddH2O were applied in volumes of 1x2 mL instead of 2x1 mL; C) Cartridges were washed with

10 mL of ddH2O and 10 mL acetonitrile for single reuse.

RP-HPLC in an ACN/AHC (pH: 7.5) buffer gradient was used to purify fluorous-tagged oligonucleotides. Samples were collected, evaporated and desalted with Oasis sample extraction cartridges HLB (1 cc, 3 cc and 20 cc; 30 µm particle size; Waters) using procedures by the manufacturer. Fluorous Affinity purification was done with Fluoro-Pak columns using standard procedures (Glen Research; Barry & Associates). Percentage of ACN in failure sequence elution was adjusted to own samples (>10 %).

V.1.5. Enzymatic Digestion of DNA Single-Strands with Mung Bean Nuclease

DNA-samples for enzymatic digestion with mung bean nuclease (MBN) were typically prepared in the following fashion:

4 µL DNA (~pmol); 5 µL ddH2O; 1 µL reaction buffer (30 mM sodium acetate (pH: 5.0), 50 mM

NaCl, 1 mM ZnCl2); 0.125 µL mung bean nuclease (40 U/µL; Roboklon GmbH) in Storage Buffer (10 mM Tris-HCl (pH: 7.5), 0.1 mM zinc acetate, 50 % (v/v) glycerol).

Samples incubated typically for 10 min at 30 °C on a thermocycler. Incubation times ranged from 5 to 30 min. The samples were immediately transferred to the cooling chamber (~8 °C), mixed with 1 µL of Gel Loading Buffer (0.05 % bromophenol blue; 40 % sucrose; 0.1 M EDTA pH: 8.0; 0.5 % SDS; Sigma) and placed as soon as possible onto the gel.

121 V. Experimental Section

V1.7. Further Equipment and Software

Pipettes: 2.5/10/20/100/1000 µL, Eppendorf

Calibrated with defined masses of water. (ρ = 1 g/mL)

Pipette tips: 10 (opaque)/100 (yellow)/1000 (blue) µL, PP, Sarstedt

Micro tubes: 0.5/1.0/2.0 mL, PP, Sarstedt

Centrifuge tubes; 15/50 mL, PP, Sarstedt

Vortex: Vortex Genie 2, Scientific Industry

Centrifuge: >2 mL: GS15R + S4180 rotor, Beckmann

<2 mL: 5415 C, Eppendorf

Shaker: THYS 2, MLW

Ultrasonic bath: Sonorex Super RK 255 H, Bandelin

Thermocycler: MiniCycler, PTC 150, MJ Research

Thermomixer: Thermomixer comfort, Eppendorf pH meter: SevenGo, Mettler-Toledo GmbH

Weighing scales: Faust (<310 g);

Ohaus (<110 g);

Sartorius (<210 g; <2 kg)

Diaphragm vacuum pump: Vacuubrand

Rotary vane vacuum pump: Trivac, AEG

ABM Greiffenberger Antriebstechnik GmbH

R 28, Vacuubrand

Freeze dryer: Alpha 1-2, CHRIST

Dry-ice evaporator: Rotavapor-RE, Büchi (Used for Py, DMF)

Ice machine: Ziegra Eismaschine GmbH

Melting point determination: Melting Point M-565 + Sample Loader M-569, Büchi

122 V. Experimental Section

Refractive Index determination: Refractometer: 84570, Carl Zeiss; Cooler: Ecoline Staredition 003 + E100, Lauda

Camera: Digital IXUS 90 IS, Canon

Xperia Z2, Sony

ChemBioDraw Ultra 14.0 PerkinElmer, Inc.

MestReNova 9.01 Mestrelab Research S.L.

Origin Pro 2015 Sr2 OriginLab Corporation

DNASequenceGenerator 1.01b U.Feldkamp, H. Rauhe

PrimerSelect 3.11 DNASTAR, Inc.

Paint.NET 4.0.6 dotPDN LLC

WinSCP 5.1.2 M. Prikryl

Inchi 1.04 IUPAC; InChI Trust

Word/Powerpoint/Excel 2013 Microsoft Corp.

The primary font used is “Alte DIN 1451 Mittelschrift”.

123 V. Experimental Section

V.2. Chemicals

Dry ice, liquid nitrogen and technical solvents were supplied by the “Chemikalienlager” at the faculty of chemistry of Ruhr-Universität Bochum. Quality grades of chemicals: “for synthesis” and “for analysis” (p.a.)

V.2.1. Solvents

Acetonitrile (ACN) p.a.: J.T. Baker

Benzene p.a.: VWR Chemicals,

Chloroform p.a.: J.T. Baker, Fisher Chemical

Cyclohexane (CH) Technical grade: Distilled prior to use.

p.a.: J.T. Baker

Dichloromethane (DCM) Technical grade: Distilled prior to use.

p.a.: Fisher Chemical, J.T. Baker

abs.: p.a. grade was stored for at least three days over molecular sieve (3 Å; 8-12 mesh) under argon- atmosphere; J.T. Baker

Diethylether abs.: (BHT stabilizer); Sigma-Aldrich

N,N-dimethylformamide p.a.: Riedel-de Haën

abs.: p.a. grade was refluxed overnight with calcium hydride under argon-atmosphere and distilled.

1,4-Dioxane p.a.: VWR Chemicals

Ethanol p.a.: Sigma-Aldrich

Ethyl acetate (EE) Technical grade: Distilled prior to use.

p.a.: Fisher Chemical, J.T. Baker n-Hexane Technical grade: Distilled prior to use.

p.a.; VWR Chemicals

Methanol p.a.: VWR Chemicals

124 V. Experimental Section n-Pentane p.a.: J.T. Baker

Pyridine (Py) p.a.: VWR Chemicals

abs.: p.a. grade was refluxed overnight with calcium hydride under argon-atmosphere and distilled.

Tetrahydrofuran (THF) p.a.: VWR Chemicals

abs.: p.a. grade was refluxed with sodium for at least two days until colour change of benzophenon.

ddH2O Deionized water was flowing through a quartz boiler and attached to the distillation unit Muldestor, Wagner & Munz

V.2.2. Educts, Reagents and Other Chemicals

Acetic acid, glacial Sigma-Aldrich

Allyl alcohol Riedel-de Haën

Allyl chloroformate Acros Organics

Ammonium chloride J.T. Baker

Ammonium hydrogen carbonate (AHC) J.T. Baker

Benzyl chloride Acros Organics, Merck n-Butyllithium (2.5 M; Hexane) Aldrich

Calcium hydride Acros Organics

Diethyl malonate Sigma-Aldrich

N,N-Diisopropylethylamine Fluka

4,4’-Dimethoxytrityl chloride Fluka

Hydrogen Alphagaz GmbH

Hydrogen chloride J.T. Baker

Iodine Sigma-Aldrich

125 V. Experimental Section

1-Iodo-[1H, 1H, 2H, 2H]-perfluorodecane Apollo Scientific

Kieselgur (Celite 545) Sigma-Aldrich

Lithium alanate Merck

Magnesium sulfate monohydrate Sigma-Aldrich

Mercaptoethanol Merck

Mesitylene Sigma-Aldrich

Methyl acrylate Sigma-Aldrich

Morpholin Acros Organics

Palladium(II)acetate ABCR

Palladium/charcoal (10 %) Fluka

Paraformaldehyde Merck

Potassium hydroxide J.T. Baker

Potassium tert-butoxide Aldrich

Sodium chloride J.T. Baker

Sodium hydride (60 % dispersion) Acros Organics

Sodium hydroxide VWG Chemicals

Sulfur Otto Fischar GmbH

Sulfuric acid Sigma-Aldrich tert-Butanol J.T. Baker

THEIC Aldrich trans-4-Methoxy-3-buten-2-one Aldrich

1,3,5-Tribromobenzene ABCR

Triethylamine Sigma-Aldrich

Triphenylphosphine Acros Organics

Tri-(o-tolyl)phosphine ABCR

126 V. Experimental Section

V.2.3. Automated Oligonucleotide Synthesis

5-benzylmercaptotetrazole emp Biotech GmbH

1,2-Bis(diphenylphosphino)ethane Acros

Bis(dibezylideneacetone)palladium(0) Acros

Dichloroacetic acid Merck

Dichloroethane Merck, Roth

4-Dimethylaminopyridine Acros Organics

2,4,6-Collidine Acros Organics

2,6-Lutidine Sigma-Aldrich

3’-O-DMT-nucleoside-phosphoramidite ChemGenes

5’-O-DMT-nucleoside-phosphoramidite Glen Research

Pyrrolidine Acros

Primer Support 200 Amino GE Healthcare

Standard CPG 500 Å Glen Research

Reverse CPG 500 Å ChemGenes

1H-Tetrazole Fluka

V.2.4. Biomolecular Reagents

Agarose 1000, Ultra-Pure Invitrogen

Ammonia (25 % and 33 %) J.T. Baker

Ammonium hydrogen carbonate AppliChem, J.T. Baker

Ammonium persulfate Merck

Boric acid Sigma

Daunorubicin hydrochloride (USP testing grade) Sigma-Aldrich

DNA-Ladder GeneRuler Life Technologies

Doxorubicin hydrochloride (for fluorescence) Sigma-Aldrich

127 V. Experimental Section

DOWEX Ion-Exchange resin 50WX8-200 Sigma-Aldrich

EDTA (Titriplex III) Merck

Gel Loading Buffer Sigma

Gel Loading Solution Type I Sigma

GelStar Nucleic Acid Stain Lonza

HEPES Sigma

Hydrogen peroxide (~30 %) Alfa Aeser

3-Hydroxypicolinic acid Fluka

Magnesium chloride Merck

Manganese(II) chloride tetrahydrate Merck

Mung bean nuclease Roboklon GmbH

Nickel(II) chloride hexahydrate Aldrich

PAA (40 %), acrylamide/bisacylamide 19:1 Roth

Sodium chloride p.a. VWR Chemicals, Fluka Analytics

Sodium metaperiodate Sigma-Aldrich

Sodium tungstate dihydrate Acros Organics

SYBR Gold Nucleic Acid Stain Invitrogen

TEMED Sigma

Trifluoroacetic acid Merck

Tris/Trizma Sigma

2,4,6-Trihydroxyacetophenon Fluka

Urea AppliChem, J.T. Baker, Sigma-Aldrich

Zinc(II) chloride J.T. Baker

128 V. Experimental Section

V.3. Organic Synthesis

V.3.01. AOC-DMT-PNO-T1 (05)

1-([2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy] methyl)-3-(allyloxycarboxyl methyl)- 5-(dimethoxytrityloxy methyl) benzene

C43H51N2O7P M = 754.86 g/mol

850 mg (1.53 mmol) 1,3-bis(2-allyloxycarboxyl methyl)-5-(2-dimethoxytrityloxy methyl) benzene was diluted with 20 mL DCMabs in an atmosphere of argon. 534 µL (396.19 mg, 3.07 mmol) of DIPEA was added. To the stirring solution 684 µL (379.41 mg, 3.07 mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h (TLC: CH/EE, 2:1, Rf: 0.53). The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE, 2:1, +1 %

NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 0.50 g; 0.66 mmol; 43 % of theory

1 H-NMR (250 MHz; CDCl3): δ [ppm] = 7.46-6.69 (m, 16 H, ArH); 5.95-5.75 (m, 1H, CH=CH2);

5.31-5.10 (m, 2H, CH=CH2); 5.21 (s, 2H, CH2OAOC); 4.75-4.60 (m, 2H, ArCH2OP); 4.60-4.55 (m,

CH2CH=CH2); 4.05 (s, 2H, CH2ODMT); 3.86-3.69 (m, 2H, CH2CH2CN); 3.65 (s, 6H, OCH3); 3.62-

3.40 (m, 2H, CH2CN); 2.57-2.46 (m, 2H, NCH); 1.20-1.00 (m, 12H, CHCH3)

13 C-NMR (63 MHz; CDCl3): δ [ppm] = 158.52 (2C, PhO, C-1’); 154.94 (1C, OCO2); 144.99 (1C, Ph, C- 1’’); 139.96, 139.70, 139.59 (3C, Ar, C-1,-3,-5); 136.20, 135.30 (2C, PhO, C-1’); 131.56 (1C,

CH2CH=CH2); 130.08 (4C, PhO, C-2’,-6’); 128.19, 127.86, 126.79 (4C, Ph, C-2’’,-5’’,-3’’,-6’’); 125.87,

125.73, 125.59 (3C, Ar, C-2,-4,-6); 118.93 (1C, CH2CH=CH2); 117.61 (1C, CN); 113.17 (4C, PhO, C-3’,-

5’); 86.51 (1C, CAr3); 69.55 (1C, CH2CH=CH2); 68.57 (1C, CH2OAOC);65.38, 65.31 (1C, CH2CH2CN);

129 V. Experimental Section

65.09 (1C, CH2ODMT); 58.43 (1C, ArCH2OP); 55.23 (2C, OCH3); 43.33, 43.13 (2C, CHCH3); 24.68,

24.61, 24.57 (4C, CHCH3); 20.42, 20.31 (1C, CH2CN)

31 P-NMR (101 MHz; DMSO-d6); δ [ppm] = 148.88 (s, 1P, P)

V.3.02. 1,3,5-Triacetybenzene (12)

C12H12O3 M = 204.23 g/mol

15.3 mL (15.0 g, 0.135 mol) trans-4-methoxy-3-buten-2-one, 120 mL EtOH and 30 mL deionized water stirred at room temperature. 0.90 mL (0.94 g; 15.72 mmol) glacial acetic acid was added via a syringe and then the mixture refluxed for 2 d. The solution turned orange and needles precipitated. It was then allowed to cool down to room temperature and then cooled overnight at -20 °C. The needles were filtered off while cold, washed with 3x 50 mL of pre-cooled Et2Oabs and dried at high vacuum. The crude solid was recrystallized once with EtOH (~200 mL). Pale yellow needles were obtained.

Yield: 6.16 g; 30.16 mmol; 67 % of theory

1 H-NMR (400 MHz; CDCl3): δ [ppm] = 8.67 (s, 3H, ArH); 2.69 (s, 9H, CH3)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 196.52 (3C, CO); 137.88 (3C, Ar, C-1,-3,-5); 131.65 (3C, Ar,

C-2,-4,-6); 26.76 (3C, CH3)

Melting point: 160-164 °C (Lit.[216] 162-165 °C)

130 V. Experimental Section

V.3.03. 2,2',2''-(Benzene-1,3,5-triyl)triacetic acid (08)

C12H12O6 M = 252.22 g/mol

6.00 g (29.38 mmol) 1,3,5-triacetylbenzene 12, 17.97 mL (17.80 g; 0.20 mol) morpholine and 6.54 g (0.20 mol) sulphur were refluxed together for 16 h. The black suspension was then poured into ice-cooled deionized water and the brown solid was collected by filtering. To the solid 23 mL of each deionized water, conc. H2SO4 and glacial acetic acid were added and refluxed for 16 h. The black suspension was turned basic with solid NaOH and stirred for 2 h under reflux. At room temperature the suspension was filtered off and the filtrate was turned acidic again with conc. HCl. The solution was extracted with 3x 100 mL of diethylether and the combined organic phases were dried over MgSO4, filtered and evaporated. The yellow to brown residue was recrystallized with glacial acetic acid to yield a beige powder.

Yield: 5.63 g; 22.33 mmol; 76 % of theory

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 12.12 (s br, 3H, CO2H); 7.05 (s, 3H, ArH); 3.53 (s, 6H,

CH2)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 172.48 (3C, CO2); 134.89 (3C, Ar, C-1,-3,-5); 128.55 (3C,

Ar, C-2,-4,-6); 40.53 (3C, CH2)

Melting point: 205-208 °C (Lit.[219] 197-204 °C; 214-216 °C)

131 V. Experimental Section

V.3.04. 2,2',2''-(Benzene-1,3,5-triyl)triacetic acid (09)

C15H18O6 M = 294.30 g/mol

10.13 g (40.16 mmol) 08 were dissolved in 155 mL methanol (p.a.). 16 mL trimethyl orthoformate and 4.2 mL conc. sulfuric acid were added and the mixture refluxed for 17 h at 80 °C. It was then neutralized with 25 % NaOH (aq.) and methanol was evaporated. The remaining aqueous solution was extracted three times with ethyl acetate, the combined organic phases were dried with MgSO4 and the solvent evaporated to yield a brownish-red oil.

Yield: 8.60 g; 29.22 mmol; 73 % of theory

1 H-NMR (200 MHz; CDCl3): δ [ppm] = 7.11 (s, 3H, ArH); 3.68 (s, 6H, CH2); 3.65 (s, 9H, CH3)

13 C-NMR (50 MHz; CDCl3): δ [ppm] = 171.42 (3C, CO2); 134.52 (3C, Ar, C-1,-3,-5); 128.80 (3C, Ar,

C-2,-4,-6); 51.62 (3C, CH3); 39.87 (m, 3C, CH2)

V.3.05. 1,3,5-Trishydroxyethylbenzene (10)

C12H18O3 M = 210.27 g/mol

8.00 g (27.18 mmol) of 09 were dissolved in 21 mL THFabs and added dropwise to a stirring ice-cooled suspension of 3.70 g (97.5 mmol) LiAlH4 in 85 mL THFabs. After full addition the mixture stirred for 2.5 h at room temperature and then for 30 min under reflux. 6.4 mL ddH2O, then 6.4 mL 15 % NaOH (aq,) and finally 19 mL ddH2O were added slowly and the cloudy mixture was filtered over a Büchner funnel. The yellowish solid was washed several times with warm THF and filtered again until the yellow colour disappeared. All organic

132 V. Experimental Section

phases were combined, dried with MgSO4, filtered and the solvent was evaporated. The residue was purified via column chromatography on silica (DCM/MeOH, 2-7 % MeOH). The faint yellow oil was dried in high vacuum to give a white solid.

Yield: 4.50 g; 21.40 mmol; 79 % of theory

1 3 H-NMR (200 MHz; CDCl3): δ [ppm] = 6.90 (s, 3H, ArH); 4.63 (s br, OH); 3.61 (t, J=7.2 Hz, 6H,

3 CH2O); 2.68 (t, J=7.2Hz, 6H, ArCH2)

13 C-NMR (50 MHz; CDCl3): δ [ppm] = 138.98 (3C, Ar, C-1,-3,-5); 127.03 (3C, Ar, C-2,-4,-6); 62.26

(3C, CH2O); 39.03 (m, 3C, ArCH2)

V.3.06. AOC-AOC-T2 (18)

1,3-Bis(2-allyloxycarboxyl ethyl)-5-(2-hydroxy ethyl) benzene

C20H26O7 M = 378.42 g/mol

1.00 g (4.76 mmol) of 10 were dissolved in 5 mL THFabs and 769 µL (9.5 mmol) Pyabs. The solution was cooled to 0 °C and 0.96 mL (1.09 g, 9.0 mmol, 1.9 eq.) allyl chloroformate diluted in 15 mL THFabs was added dropwise under stirring. The solution stirred for another 3 h at room temperature. Solvent was evaporated and the crude product was purified via column chromatography on silica (CH/EE; 2:1). A clear oil was obtained.

Yield: 446 mg; 1.18 mmol; 25 % of theory

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 6.99 (m, 3H, ArH); 6.01-5.84 (m, 2H, CH=CH2); 5.40-

3 5.20 (m, 4H, CH=CH2); 4.56 (m, 1H, OH); 4.64-4.56 (m, 2H, CH2CH=CH2); 3.61 (t, J=6.7 Hz, 2H,

3 3 CH2OH); 3.39 (s, 4H, CH2OAOC); 2.89 (t, J=6.9 Hz, 4H, CH2CH2OAOC): 2.70 (t, J=7.2Hz, 2H,

CH2CH2OH)

13 C-NMR (50 MHz; DMSO-d6): δ [ppm] = 154.25 (2C, OCO2); 139.69, 137.73 (3C, Ar, C-1,-3,-5);

132.16 (1C, CH2CH=CH2); 127.59, 126.59 (3C, Ar, C-2,-4,-6); 118.25 (2C, CH2CH=CH2); 67.88 (2C,

133 V. Experimental Section

CH2CH=CH2); 67.69 (2C, CH2OAOC); 62.11 (1C, CH2OH); 38.92 (1C, CH2CH2OH); 34.14 (2C,

CH2CH2OAOC)

V.3.07. AOC-AOC-PNO-T2 (19)

1-(2-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy] ethyl)-3,5-bis-(2-allyloxycarboxyl ethyl) benzene

C29H43N2O8P M = 578.64 g/mol

394 mg (1.04 mmol) of 18 were diluted with 15 mL DCMabs in an atmosphere of argon. 291 µL (215.92 mg, 1.67 mmol) of DIPEA were added. To the stirring solution 373 µL (395.38 mg, 1.67 mmol; 1.6 eq.) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h. The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE, 2:1, +0.75 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 298 mg; 0.52 mmol; 50 % of theory

1 H-NMR (250 MHz; CDCl3): δ [ppm] = 6.91- 6.94 (m, 3H, ArH); 5.97-5.75 (m, 2H, CH=CH2); 5.33-

3 5.13 (m, 4H, CH=CH2); 4.60-4.47 (m, 4H, CH2CH=CH2); 4.25 (t, J=7.3 Hz, 4H, CH2OAOC);3.83-

3.63 (m, 4H, ArCH2CH2OP + CH2CH2CN); 3.59-3.43 (m, 2H, NCH); 2.95-2.73 (m, 6H,

3 CH2CH2OAOC + CH2CH2OP); 2.51 (t, J=6.5 Hz, 2H, CH2CN); 1.23-1.03 (m, 12H, CHCH3)

13 C-NMR (63 MHz; CDCl3): δ [ppm] = 154.90 (2C, OCO2); 139.30, 137.49 (3C, Ar, C-1,-3,-5); 131.59

(2C, CH2CH=CH2); 128.08, 127.52 (3C, Ar, C-2,-4,-6); 118.86 (2C, CH2CH=CH2); 117.64 (1C, CN);

68.38 (2C, CH2CH=CH2); 68.30 (2C, CH2OAOC); 64.40, 64.12 (1C, CH2CH2CN); 58.50, 58.20 (1C,

ArCH2CH2OP); 43.17, 42.98 (2C, CHCH3); 37.66, 37.55, (1C, ArCH2CH2OP); 34.95 (1C,

CH2CH2OAOC); 24.57, 24.54, 24.55, 24.46 (4C, CHCH3); 20.24, 20.34 (1C, CH2CN)

31 P-NMR (101 MHz, CDCl3); δ [ppm] = 147.59 (s, 1P, P)

134 V. Experimental Section

V.3.08. DMT-DMT-T2 (16)

1,3-Bis(2-dimethoxytrityloxy ethyl)-5-(2-hydroxy ethyl) benzene

C54H54O7 M = 815.02 g/mol

1.00 g (4.76 mmol) of 1,3,5-trishydroxyethylbenzene 10 were dissolved in 15 mL Pyabs and stirred in an atmosphere of argon. A solution of 2.42 g (7.14 mmol, 1.6 eq.) 4,4’- dimethoxytrityl chloride in 40 mL Pyabs was added dropwise under vigorous stirring at room temperature over a period of 30 min and then stirred for 16 h at room temperature. The reaction was then quenched with 10 mL MeOH, while stirring at room temperature for 10 min. Solvent was condensed and crude product was purified via column chromatography on silica (DCM + 0.5 % NEt3). The product was a yellow, viscous oil.

Yield: 764 mg; 0.94 mmol; 20 % of theory

1 3 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 7.32-6.71 (m, 29H, ArH); 4.66 (t, J=5.2 Hz, 1H, OH);

3 3 3.71 (s, 12H, OCH3); 3.59 (q, J=7.2 Hz, 2H, CH2OH); 3.15 (t, J=6.2 Hz, 4H; CH2ODMT); 2.86-2.63

(m, 6H, CH2CH2OH + CH2CH2ODMT)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 157.92 (4C, PhO, C-1’); 145.13 (2C, Ph, C-1’’); 138.98, 138.75 (3C, Ar, C-1,-3,-5); 135.87 (4C, PhO, C-4’); 129.56 (8C, PhO, C-2’,-6’); 128.90, 127.66, 127.62 (8C, Ph, C-2’’,-5’’,-3’’,-6’’); 127.33, 126.47, 123.87 (3C, Ar, C-2,-4,-6); 113.01, 112.73 (4C, PhO, C-

3’,-5’); 85.36 (1C, CAr3); 64.22 (1C, CH2ODMT); 62.39 (1C, CH2OH); 55.93 (2C, OCH3); 35.94 (1C,

CH2CH2ODMT); 39.06 (in DMSO signal, 1C, CH2CH2OH);

135 V. Experimental Section

V.3.09. DMT-DMT-PNO-T2 (17)

1-(2-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy] ethyl)-3,5-bis-(2-dimethoxytrityl oxy ethyl) benzene

C63H71N2O8P M = 1015.24 g/mol

764 mg (0.94 mmol) of 16 were diluted with 15 mL DCMabs in an atmosphere of argon. 261 µL (215.92 mg, 1.67 mmol) of DIPEA was added. To the stirring solution 335 µL (355.10 mg, 1.50 mmol, 1.6 eq.) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h. The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE, 2:1, +0.75 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 580 mg; 0.57 mmol; 61 % of theory

1 H-NMR (250 MHz; CDCl3): δ [ppm] = 7.36-6.70 (m, 16H, ArH); 3.67 (s, 6H, OCH3); 3.66-3.55 (m,

3 4H, ArCH2CH2OP + CH2CH2CN); 3.54-3.41 (m, 2H, NCH); 3.17 (t, J=6.9 Hz, 2H, CH2ODMT); 2.82-

2.67 (m, 8H, CH2CH2OH + CH2CH2OP + CH2CH2DMT); 2.36 (tt, J=6.5, 0.9 Hz, 2H, CH2CN); 1.08 (d,

3 3 J=6.8 Hz, 6H, CHCH3); 1.02 (d, J=6.8 Hz, 6H, CHCH3)

13 C-NMR (63 MHz; CDCl3): δ [ppm] = 158.33 (2C, PhO, C-1’); 145.26 (2C, Ph, C-1’’); 139.32, 138.16 (3C, Ar, C-1,-3,-5); 136.52 (4C, PhO, C-4’); 129.94 (8C, PhO, C-2’,-6’); 129.14, 128.15, 127.84, 127.77 (8C, Ph, C-2’’,-5’’,-3’’,-6’’); 127.72, 127.69, 126.65 (3C, Ar, C-2,-4,-6); 117.62 (1C, CN), 113.18, 112.99

(4C, PhO, C-3’,-5’); 85.97 (1C, CAr3); 64.76 (1C, CH2ODMT); 58.49 (1C, CH2CH2CN); 58.19 (1C,

ArCH2CH2OP); 55.17 (2C, OCH3); 43.18, 42.89 (2C, CHCH3); 37.85, 37.77 (1C, ArCH2CH2OP); 36.63

(1C, CH2CH2ODMT); 24.68, 24.58, 25,57, 24.47 (4C, CHCH3); 20.12, 20.22 (1C, CH2CN)

31 P-NMR (101 MHz, CDCl3); δ [ppm] = 147.49 (s, 1P, P)

136 V. Experimental Section

V.3.10. DMT-T2 (13)

1-(2-Dimethoxytrityloxy ethyl)-3,5-bis(2-hydroxy ethyl) benzene

C33H36O5 M = 512.65 g/mol

1.50 g (7.13 mmol) of 1,3,5-trishydroxyethylbenzene 10 were dissolved in 22 mL Pyabs and stirred in an atmosphere of argon. A solution of 1.93 g (5.70 mmol, 0.8 eq.) 4,4’- dimethoxytrityl chloride in 60 mL Pyabs was added dropwise under vigorous stirring at room temperature over a period of 30 min and then stirred for 16 h at room temperature. The reaction was then quenched with 10 mL MeOH, while stirring at room temperature for 10 min. Solvent was condensed and crude product was purified via column chromatography on silica (DCM/MeOH, 0-5 % MeOH + 0.5 % NEt3). The product was a yellow, viscous oil.

Yield: 1.80 g; 3.51 mmol; 50 % of theory

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 7.34-6.76 (m, 16H, ArH); 4.60 (s br, 1H, OH); 3.74 (s,

3 3 6H, OCH3); 3.55-3.38 (q, J=7.2 Hz, 2H, CH2OH); 3.13 (t, J=6.6 Hz, 4H; CH2ODMT); 2.80-2.60 (in

NEt3, m, 6H, CH2CH2OH + CH2CH2ODMT)

137 V. Experimental Section

V.3.11. AOC-DMT-T2 (14)

1-(3-Allyloxycarboxyl ethyl)-3-(3-dimethoxytrityloxy ethyll)-5-(2-hydroxy ethyl) benzene

C37H40O7 M = 596.72 g/mol

1.82 g (3.55 mmol) of 13 were dissolved in 5 mL THFabs and 316 µL (3.92 mmol) Pyabs. The solution was cooled to 0 °C and 0.34 mL (386 mg, 3.20 mmol; 0.9 eq.) allyl chloroformate diluted in 10 mL THFabs was added dropwise under stirring. The solution then stirred for 16 h at room temperature. Solvent was evaporated and the crude product was purified via column chromatography on silica (DCM/MeOH; 0.5-5 % MeOH + 0.5 % NEt3). A clear viscous oil was obtained.

Yield: 538 mg; 0.90 mmol; 25 % of theory

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 7.37-6.75 (m, 16H, ArH); 6.01-5.75 (m, 1H,

3 CH=CH2);5.39-5.15 (m, 2H, CH=CH2); 4.66 (t, J=5.2 Hz, 1H, OH); 4.62-4.52 (m, 2H, CH2CH=CH2);

3.74, 3.71 (s, 6H, OCH3); 3.60 (q, J=7.0, 6.5 Hz, 1H, CH2OH); 3.42-3.37 (water impurity, m, 2H,

3 CH2OAOC); 3.15 (t, J=6.5 Hz, 2H; CH2ODMT); 2.82-2.64 (m, 6H, ArCH2)

13 C-NMR (50 MHz; DMSO-d6): δ [ppm] = 157.92 (1C, PhO, C-1’); 154.24 (1C, OCO2); 145.08 (1C, Ph,

C-1’’); 139.34, 139.16, 138.77 (3C, Ar, C-1,-3,-5); 136.98 (2C, PhO, C-4’); 132.13 (2C, CH2CH=CH2); 129.56 (3C, Ar, C-2,-4,-6); 127.69, 127.61 (4C, Ph, C-2’’,-5’’,-3’’,-6’’); 127.18, 126.50 (8C, PhO, C-

2’,-6’); 118.23 (2C, CH2CH=CH2); 113.03 (4C, PhO, C-3’,-5’); 85.34 (1C, CAr3); 67.95 (2C,

CH2CH=CH2); 67.67 (1C, CH2OAOC); 64.21 (1C, CH2ODMT); 62.26 (1C, CH2OH); 54.95 (2C, OCH3);

39.06 (in DMSO signal, 1C, CH2CH2OH); 35.83 (1C, CH2CH2ODMT); 34.70 (1C, CH2CH2OAOC);

138 V. Experimental Section

V.3.12. AOC-DMT-PNO-T2 (15)

1-(2-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy] ethyl)-3-(2-allyloxycarboxyl ethyl)- 5-(2-dimethoxytrityloxy ethyl) benzene

C46H57N2O7P M = 796.94 g/mol

500 mg (0.84 mmol) of 13 was diluted with 15 mL DCMabs in an atmosphere of argon. 365 µL (270.75 mg, 2.09 mmol) of DIPEA was added. To the stirring solution 468 µL (495.80 mg, 2.09 mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h (TLC: CH/EE, 2:1, Rf: 0.6). The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE, 2:1,

+0.75 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 382 mg; 0.48 mmol; 57 % of theory

1 H-NMR (250 MHz; DMSO-d6): δ [ppm] = 7.36-6.70 (m, 16H, ArH); 6.00-5.79 (m, 1H, CH=CH2);

5.37-5.17 (m, 2H, CH=CH2); 4.62-4.51 (m, 2H, CH2CH=CH2); 4.35-4.20 (m, 2H, CH2OAOC); 3.73

(s, 6H, OCH3); 3.73-3.61 (m, 4H, ArCH2CH2OP + CH2CH2CN); 3.60-3.41 (m, 2H, NCH); 3.17-3.09

(m, 2H, CH2ODMT); 2.94-2.63 (m, 8H, CH2CH2OAOC + CH2CH2OP + CH2CH2DMT + CH2CN); 1.21-

0.89 (m, 12H, CHCH3)

13 C-NMR (63 MHz; CDCl3): δ [ppm] = 158.26 (1C, PhO, C-1’); 154.80 (1C, OCO2); 145.14 (1C, Ph, C- 1’’); 139.77, 139.22, 138.67 (3C, Ar, C-1,-3,-5); 136.91, 136.40 (2C, PhO, C-4’); 131.55 (2C,

CH2CH=CH2); 129.89 (3C, Ar, C-2,-4,-6); 128.31, 128.05, 127.75 (4C, Ph, C-2’’,-5’’,-3’’,-6’’); 129.94

(8C, PhO, C-2’,-6’); 118.73 (2C, CH2CH=CH2); 117.64 (1C, CN); 112.96 (4C, PhO, C-3’,-5’); 85.86 (1C,

CAr3); 68.33, 68.26 (2C, CH2CH=CH2); 64.63(1C, CH2OAOC); 64.50, 64.22 (1C, CH2ODMT); 60.28

(1C, CH2CH2CN); 58.41, 58.11 (1C, ArCH2CH2OP); 55.11, 55.09 (2C, OCH3); 43.09, 42.89 (1C,

ArCH2CH2OP); 37.56 (2C, CHCH3); 36.50 (1C, CH2CH2ODMT); 34.90 (1C, CH2CH2OAOC); 24.59,

24.49, 24.39 (4C, CHCH3); 20.22, 20.12 (1C, CH2CN)

31 P-NMR (101 MHz, DMSO-d6); δ [ppm] = 146.51 (s, 1P, P)

139 V. Experimental Section

V.3.13. 1,3,5-Tris(E- methyl acryoyl) benzene (28)

C18H18O6 M = 330.34 g/mol

50 g (0.16 mol) of 1,3,5-tribromobenzene were dissolved in 170 mL ACN (p.a.) and stirred for a few minutes under argon-atmosphere. 1.06 g (4.72 mmol; 3 mol%) Pd(OAc)2, 2.71 g (8.90 mmol; 6 mol%) tri-(o-tolyl)phosphine and 125 mL NEt3 were added to the suspension. The mixture was refluxed until a clear dark-brown solution appeared. 46 mL (508.19 mmol) methyl acrylate were added quickly dropwise to the refluxing solution. After a few hours needles formed in the solution. After one day (TLC: DCM Rf: 0.47) another 2 mL of methyl acrylate and 200 mg of the catalysts and the ligand were added. The reaction continued for 16 h, was cooled to room temperature and stirred for 10 min after the addition of 100 mL deionized water. The grey precipitate was filtered off with a Büchner funnel and washed with

200 mL of ddH2O and 2x 50 mL n-hexane. The crude product was dissolved in 500 mL dioxane and boiled to reflux under stirring. The solution had a faint greenish colour. The solution was treated with charcoal for 10 min and the still hot solution was then filtered off over a glass fiber filter. A solid precipitated quickly while cooling and the mixture was further cooled in an ice bath for 5 min. The product was then rinsed with each 30 mL of cold DCM and n-hexane. The faint greyish solid was dried overnight at high vacuum and was pure enough for further synthesis.

Yield: 41.83 g; 126.66 mmol; 80 % of theory

1 3 H-NMR (400 MHz; CDCl3): δ [ppm] = 7.69 (d, J=16.1 Hz, 3H, ArCH); 7.65 (s, 3H, ArH); 6.51 (d,

3 J=16.0 Hz, 3H, CHC=O); 3.84 (s, 9H, OCH3)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 166.80 (3C, CO2); 143.01 (3C, C=CHCO2); 135.86 (3C, Ar, C-

1,-3,-5); 128.86 (3C, Ar, C-2,-4,-6); 119.56 (3C, C=CHCO2); 51.86 (3C, CH3)

Melting point: 212-215 °C (Lit.[222] 210-211 °C)

140 V. Experimental Section

V.3.14. 1,3,5-Tris(methyl propyloyl) benzene (24)

C18H24O6 M = 336.38 g/mol

7.0 g (21.20 mmol) of 28 and 0.5 g of 10 % Pd/C were carefully dissolved in 200 mL THFabs inside a 0.5 L Parr bottle and placed in a Parr shaker. (Appendix contains technical instructions.) The black suspension was purged twice with argon and hydrogen (40 psi). Hydrogenation was carried out at room temperature under mechanical shaking. The absorption of hydrogen stopped after 15 min, but the reaction proceeded for another 15 min and the system was purged with argon. The catalyst was filtered off with a glass fiber filter and the solvent was evaporated giving a clear greyish oil. 41.71 g of combined crude product (41.75 g theoretical) was distilled in vacuo (b.p. 180-190 °C/~0.5 mbar) using a heating mantle and graphite as the heating medium. A clear colourless oil was collected.

Yield: 39.63 g; 117.81 mmol; 95 % of theory

1 3 H-NMR (200 MHz; CDCl3): δ [ppm] = 6.80 (s, 3H, ArH); 3.59 (s, 9H, OCH3); 2.82 + 2.52 (t, J=7.5

Hz, 6H, CH2)

13 C-NMR (50 MHz; CDCl3): δ [ppm] = 173.2 (3C, CO2); 140.99 (3C, Ar, C-1,-3,-5); 126.22 (3C, Ar,

C-2,-4,-6); 51.56 (3C, CH3); 35.65 (3C, ArCH2CH2); 30.80 (3C, ArCH2)

141 V. Experimental Section

V.3.15. Allyl benzyl ether (31)

C10H12O M = 148.21 g/mol

10 g (178.24 mmol) KOH were dissolved in 43.73 mL (37.17 g; 0.64 mol) allyl alcohol. It was cooled down to ~10 °C with an ice bath and 14.73 mL (16.20 g; 127.98 mmol) benzyl chloride was added dropwise to the stirred solution. The reaction mixture was then heated to 60 °C for 1 h (TLC: DCM; Rf: 0.47). The precipitate was filtered off and washed with 20 mL DCM. The combined organic phases were evaporated and the liquid residue was distilled in vacuo (b.p. 100-108 °C/~35 mbar) to give a clear colourless liquid.

Yield: 14.84 mL; 14.23 g; 96.01 mmol; 75 % of theory

1 3 H-NMR (400 MHz; CDCl3): δ [ppm] = 7.45-7.27 (m, 5H, ArH); 6.01 (ddt, J=17.3, 10.4, 5.6 Hz,

3 4 3 1H, OCH2CH=CH2); 5.37 (dq, J=17.3 Hz, J=1.7 Hz, 1H, trans-CH=CH2); 5.26 (dq, J=10.4 Hz,

4 3 4 J=1.4 Hz, 1H, cis-CH=CH2); 4.58 (s, 2H, ArCH2); 4.09 (dt, J=5.7 Hz, J=1.4 Hz, 2H CH2CH=CH2)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 138.36 (1C, Ar, C-1); 134.80 (1C, OCH2C=CH2); 128.39 (2C,

Ar, C-3,-5); 127.73 (2C, Ar, C-2,-6); 127.59 (1C, Ar, C-4); 117.06 (1C, OCH2C=CH2); 72.14 (1C,

OCH2C=CH2); 71.16 (1C, ArCH2O)

[285] Refractive index: nD21 = 1.506 (Lit. nD20 = 1.507)

142 V. Experimental Section

V.3.16. 1E,3E,5Z-1,3,5-Tris(benzyl oxy isoallyl) benzene (32)

C36H36O3 M = 516.68 g/mol

10 g (31.77 mmol) of 1,3,5-tribromobenzene were dissolved in 30 mL ACN (p.a.) and stirred under an argon atmosphere. 214 mg (0.95 mmol; 3 mol%) Pd(OAc)2, 0.5 g (1.91 mmol; 6 mol%)

PPh3 and 25 mL NEt3 were added. The brown suspension was stirred and refluxed until the solution gave a clear dark-brown solution. 17.18 mL (16.48 g; 0.11 mol) of allyl benzyl ether 31 were added dropwise to the refluxing solution. The reaction stirred for 16 h (TLC: CH/EE; 9:1; Rf: 0.4+0.5). The mixture cooled to room temperature, filtered off through a Büchner funnel and rinsed with 50 mL of acetonitrile. The combined organic phases were evaporated and dissolved in 200 mL DCM. The organic phase was treated with 20 mL each of deionized water, 10 % HCl and again with water and finally washed with brine. The organic phase was dried with magnesium sulfate, filtered and evaporated. The brownish oil was purified via column chromatography on silica (CH/EE; 95:5).

Yield: 6.07 g; 11.75 mmol; 37 % of theory

1 H-NMR (400 MHz; CDCl3): δ [ppm] = 7.30-7.15 (m, 15H, Bn ArH); 6.82-6.71 (m, 3H, ArH); 6.33 (ddt, 3J=12.6, 4J=2.7, 1.4 Hz, 1H, cis-ArCH=CH); 6.06-5.99 (m, 2H, trans-ArCH=CH); 4.94 (dt,

3 J=12.6, 7.6 Hz, 1H, cis-ArCH=CH): 4.74-4.71 (m, 4H, trans-ArCH2); 4.66-4.62 (m, 2H, cis-

3 ArCH2); 4.52 (td, J=7.4, 6.1 Hz, 1H, trans-ArCH=CH); 3.37-3.30 (m, 4H, trans-CH=CH-CH2);

3.15-3.07 (m, 2H, cis-CH=CH-CH2)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 145.04, 144.86 (3C, Ar, C-1’); 141.63 (3C, Ar, C-1,-3,-5);

137.78 (3C, C=CHCO2); 128.50, 128.51 (6C, Ar, C-3’,-5’); 127.86, 127.88 (3C, OCH2C=CH2); 127.57

(3C, Ar, C-4’); 127.37 (6C, Ar, C-2’,-6’); 125.81, 125.87 (3C, Ar, C-2,-4,-6); 73.71 (3C, OCH2C=CH2);

71.17 (3C, ArCH2O);

FAB-MS: 516.1 (50); 181.0 (90); 91.0 (100) [M]

143 V. Experimental Section

V.3.17. 1,3,5-Tris(3-hydroxy propyl) benzene (25)

C15H24O3 M = 252.35 g/mol

Method A:

To 17.15 g (0.46 mol) LiAlH4 300 mL of THFabs were added in an argon atmosphere. The stirring solution was cooled down with an ice bath and a solution of 39.00 g (115.94 mmol) 24 in

250 mL THFabs was added dropwise. After full addition of the educt the reaction continued for 1.5 h at room temperature and another 1.5 h at reflux. The reaction was quenched by adding dropwise 20 mL of deionized water, 20 mL of 15 % NaOH and 60 mL of deionized water. The solid was filtered off and washed several times with warm THF. The combined organic phases were evaporated leaving a yellowish wax-like residue and then coevaporated in THF.

The crude viscous oil (~29 g) was recrystallized by adding as little THFabs as possible at room temperature until full solvation. n-Pentane was added dropwise until a steady cloudiness in the solution appeared. This mixture was cooled to -20 °C for at least 2 h. The product precipitated as colourless pellets. A Büchner funnel and an appropriate filter were pre- cooled on dry ice and the solid was then filtered off quickly. It was then quickly washed twice with each 50 mL of a 1:3 THF/pentane solution, which was pre-cooled with liquid nitrogen. The wet solid needs to stay cool to avoid transformation into an oil near room temperature. The solid was then dried over the funnel and then dried at high vacuum overnight. The solid remains then as a colourless powder.

Alternatively the crude product can be purified by column chromatography (DCM/MeOH, 100:1 - 10:1).

Yield: 21.94 g; 86.94 mmol; 75 % of theory

Method B:

2.00 g (3.87 mmol) of 32 and 0.3 g of 10 % Pd/C were carefully dissolved in 70 mL THF p.a. inside a 0.5 L bottle and placed in a shaker. (Appendix contains technical instruction for the

144 V. Experimental Section

Parr apparatus) The black suspension was purged twice with argon and hydrogen (40 psi). Hydrogenation was carried out at room temperature under mechanical shaking. The reaction was carried out for 16 h. The system was then purged with argon and the catalyst was filtered off over a 1 cm thick layer of kieselgur. It was flushed with 20 mL MeOH and the combined organic phases were evaporated giving a clear greyish oil. The residue was purified via column chromatography on silica (DCM/MeOH 100:1 - 10:1).

Yield: 0.89 g; 3.53 mmol; 91 % of theory

1 3 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 6.82 (s, 3H, ArH); 4.42 (t, J=5.12 Hz, 3H, OH); 3.47-

3 3.38 (m, 6H, CH2O); 2.61-2.47 (m, 6H, ArCH2); 1.70 (p, J=7.01 Hz, 6H, CCH2C)

13 C-NMR (50 MHz; DMSO-d6): δ [ppm] = 141.86 (3C, Ar, C-1,-3,-5); 125.63 (3C, Ar, C-2,-4,-6);

60.16 (3C, CH2O; 34.35 (3C, ArCH3CH2); 31.58 (3C, ArCH2)

Melting point: 48-52 °C (Lit.[222] 49-51 °C)

V.3.18. AOC-T3 (33)

1-(3-Allyloxycarboxyl propyl)-3,5-bis(3-hydroxy propyl) benzene

C19H28O5 M = 336.43 g/mol

18.00 g (71.33 mmol) 25, 90 mL THFabs and 4.61 mL (4.51 g, 57.05 mmol) pyridineabs were stirred in an atmosphere of argon. The mixture was cooled to 0 °C and a solution of 6.25 mL

(7.09 g, 58.85 mmol) allyl chloroformate in 45 mL THFabs was added dropwise, while keeping the temperature <5 °C. The mix turned cloudy in the process. The reaction continued for another 5 h at low temperature and another 30 min at room temperature. Py*HCl was filtered off and the filtrate was evaporated. The residue was chromatographed on silica (Automated: DCM/MeOH, 1 %-20 % over 20 min; Manual alternative: CH/EE; 3:1). The product was a viscous oil.

Yield: 9.03 g; 26.84 mmol; 47 % of theory

145 V. Experimental Section

1 3 H-NMR (400 MHz; DMSO-d6): δ [ppm] = 6.85-6.81 (m, 3H, ArH); 5.99-5.88 (ddt, J=17.3, 10.7,

3 4 3 5.5 Hz, 1H, CH=CH2); 5.33 (dq, J=17.3 Hz, J=1.6 Hz, 1H, trans-CH=CH2); 5.25 (dq, J=10.5 Hz,

4 3 4 3 J=1.4 Hz, 1H, cis-CH=CH2); 4.59 (dt, J=5.7 Hz, J=1.5 Hz, 2H, CH2CH=CH2); 4.45 (t, J=5.1 Hz,

3 2H, OH); 4.08 (t, J=6.5 Hz, 2H, CH2OAOC); 3.45-3.40 (m, 4H, CH2OH); 2.60-2.52 (m, 6H, ArCH2);

3 3 2.03-1.82 (pm, J=14.2 Hz, 2H, CH2CH2OAOC); 1.78-1.60 (pm, J=14.2 Hz, 4H, CH2CH2OH)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 154.34 (1C, OCO2); 142.05, 140.66 (3C, Ar, C-1,-3,-5);

132.21 (1C, CH2CH=CH2); 125.98, 125.60 (3C, Ar, C-2,-4,-6); 118.21 (1C, CH2CH=CH2); 67.66 (1C,

CH2CH=CH2); 66.99 (1C, CH2OAOC); 60.16 (1C, CH2OH); 34.30 (2C, CH2CH2OH); 31.55 (2C,

CH2CH2CH2OH); 31.10 (1C, CH2CH2CH2OAOC); 29.77 (1C, CH2CH2OAOC)

V.3.19. AOC-DMT-T3 (34)

1-(3-Allyloxycarboxyl propyl)-3-(3-dimethoxytrityloxy propyl)-5-(2-hydroxy propyl) benzene

C40H46O7 M = 638.80 g/mol

9.00 g (26.75 mmol) of 33 were dissolved in 60 mL of pyridineabs and the solution was stirred at room temperature in an atmosphere of argon. A solution of 7.50 g (22.13 mmol) 4,4’- dimethoxytrityl chloride in 50 mL Pyabs was added dropwise over a period of 2 h at room temperature and stirring continued for 16 h. Solvent was removed and the residue was purified via column chromatography on silica (Automated: CH/EE, +0.5 % NEt3, 5 %-30 % over

10 min; Manual alternative: CH/EE; 3:1, +0.5 % NEt3).

Yield: 6.49 g; 10.16 mmol; 38 % of theory

1 H-NMR (400 MHz; DMSO-d6): δ [ppm] = 7.43-7.15 (m, 9 H, ArH); 6.93-6.73 (m, 7 H, ArH); 5.99-

3 4 3 5.87 (m, 1H, CH=CH2); 5.33 (dq, J=17.3 Hz, J=1.6 Hz, 1H, trans-CH=CH2); 5.24 (dq, J=10.5 Hz,

4 3 4 3 J=1.4 Hz, 1H, cis-CH=CH2); 4.59 (dt, J=5.5 Hz, J=1.5 Hz, CH2CH=CH2); 4.41 (t, J=4.8 Hz, 1H,

3 3 4 OH); 4.07 (t, J=5.5 Hz, 2H, CH2OAOC); 3.74 (s, 6H, OCH3); 3.40 (td, J=6.3 Hz, J=4.3 Hz, 2H,

146 V. Experimental Section

3 CH2OH); 2.99 (t, J=6.3 Hz, 2H, CH2ODMT); 2.48-2.63 (m, 8H, ArCH2 + CH2CN); 1.90-1.77 (m, 4H,

CH2CH2OAOC + CH2CH2ODMT); 1.61-1.72 (m, 2H, CH2CH2OH)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 157.94 (2C, PhO, C-1’); 154.33 (1C, OCO2); 145.21 (1C, Ph, C-1’’); 142.05, 141.56, 140.62 (3C, Ar, C-1,-3,-5); 136.04 (2C, PhO, C-4’); 132.23 (1C,

CH2CH=CH2); 129.54 (4C, PhO, C-2’,-6’); 126.48, 127.63, 127.69 (4C, Ph, C-2’’,-5’’,-3’’,-6’’); 125.59,

125.66, 125.99 (3C, Ar, C-2,-4,-6); 118.19 (1C, CH2CH=CH2); 113.05 (4C, PhO, C-3’,-5’); 85.18 (1C,

CAr3); 67.65 (1C, CH2CH=CH2); 66.97 (1C, CH2OAOC); 62.06 (1C, CH2ODMT); 60.16 (1C, CH2OH);

54.96 (2C, OCH3); 34.30 (1C, CH2CH2OH); 31.77 (1C, CH2CH2CH2ODMT); 31.56 (1C,

CH2CH2CH2OAOC); 31.00 (1C, CH2CH2CH2OH); 29.78 (1C, CH2CH2ODMT); 26.31 (1C, CH2CH2OAOC)

V.3.20. AOC-DMT-PNO-T3 (35)

1-(3-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy]propyl)-3-(3-allyloxycarboxyl propyl)-5-(3-dimethoxytrityloxy propyl) benzene

C49H63N2O8P M = 839.02 g/mol

512 mg (0.80 mmol) of 34 was diluted with 12 mL DCMabs in an atmosphere of argon. 297 µL (207.19 mg, 1.60 mmol) of DIPEA was added. To the stirring solution 385 µL (379.41 mg, 1.66 mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture was stirred at room temperature for 2 h (TLC: CH/EE, 2:1 +1 % NEt3, Rf: 0.59). The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica

(CH/EE, 2:1, +1 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 403.49 mg; 0.48 mmol; 60 % of theory

147 V. Experimental Section

1 H-NMR (400 MHz; CDCl3): δ [ppm] = 7.42-7.08 (m, 9 H, ArH); 6.77-6.71 (m, 7 H, ArH); 5.95-

3 4 5.81 (m, 1H, CH=CH2); 5.35-5.16 (m, 2H, CH=CH2); 4.56 (dt, J=5.79 Hz, J=1.39 Hz, CH2CH=CH2);

3 4.08 (t, J=6.51 Hz, 2H, CH2OAOC); 3.72 (s, 6H, OCH3); 3.86-3.68 (m, 2H, CH2CH2CH2OP); 3.65-

3 3.47 (m, 4H, NCH + CH2CH2CN); 3.03 (t, J=6.32 Hz, 2H, CH2ODMT); 2.62-2.48 (m, 8H, ArCH2 +

CH2CN); 1.94-1.77 (m, 6H, ArCH2CH2); 1.16-1.05 (m, 12H, CHCH3)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 158.35 (2C, PhO, C-1’); 155.05 (1C, OCO2); 145.37 (1C, Ph,

C-1’’); 142.55, 141.92, 140.96 (3C, Ar, C-1,-3,-5); 136.71 (2C, PhO, C-4’); 131.68 (1C, CH2CH=CH2); 130.02 (4C, PhO, C-2’,-6’); 128.23, 127.67, 126.57 (4C, Ph, C-2’’,-5’’,-3’’,-6’’); 126.35, 126.05, 125.95

(3C, Ar, C-2,-4,-6); 118.83 (1C, CH2CH=CH2); 117.59 (1C, CN); 112.98 (4C, PhO, C-3’,-5’); 85.75 (1C,

CAr3); 68.34 (1C, CH2CH=CH2); 67.51 (1C, CH2OAOC); 63.15, 62.98 (1C, CH2CH2CN); 62.88 (1C,

CH2ODMT); 58.43, 58.24 (1C, CH2CH2CH2OP); 55.18 (2C, OCH3); 43.12, 43.00 (2C, CHCH3); 32.92,

32.86 (1C, CH2CH2CH2OP); 32.60 (1C, CH2CH2CH2ODMT); 31.89 (1C, CH2CH2CH2OAOC); 31.84 (1C,

CH2CH2OP); 30.34 (1C, CH2CH2ODMT); 29.68 (1C, CH2CH2OAOC); 24.68, 24.61, 24.45 (4C, CHCH3);

20.40, 20.33 (1C, CH2CN)

31 P-NMR (162 MHz, CDCl3); δ [ppm] = 147.48 (s, 1P, P)

V.3.21. AOC-TN

1-(2-Allyloxycarboxyl ethyl)-3,5-bis(2-hydroxy ethyl) isocyanurate

C13H19N3O8 M = 345.31 g/mol

3.90 g (14.9 mmol) 1,3,5-tris(2-hydroxyethyl)isocyanurate were dissolved in 90 mL Pyabs and then cooled down to 0 °C. 1.34 mL (1.51 g, 12.51 mmol) allyl chloroformate were added slowly over a period of 1 h with a syringe. Stirring was continued at room temperature for 4 h. Solvent was condensed and the residue was purified via column chromatography on silica

(DCM/MeOH, 95:5, Rf: 0.55).

Yield: 2.86 g; 8.28 mmol; 55 % of theory

148 V. Experimental Section

1 3 H-NMR (400 MHz; DMSO-d6): δ [ppm] = 5.92 (ddt, J=17.3, 10.7, 5.5 Hz, 1H, CH=CH2); 5.33 (dq,

3 4 3 4 J=17.2 Hz, J=1.6 Hz, 1H, trans-CH=CH2); 5.25 (dq, J=10.5 Hz, J=1.4 Hz, 1H, cis-CH=CH2); 4.76

3 3 4 3 (t, J=6.1 Hz, 2H, OH); 4.59 (dt, J=5.5 Hz, J=1.5 Hz, 2H, CH2CH=CH2); 4.27 (t, J=5.4 Hz, 2H,

3 3 3 CH2OAOC); 4.07 (t, J=5.4 Hz, 2H, CH2CH2OAOC); 3.84 (t, J=6.4 Hz, CH2CH2OH); 3.53 (q, J=6.3

Hz, 4H, CH2CH2OH)

13 C-NMR (101 MHz; CDCl3): δ [ppm] = 154.17 (1C, OCO2); 149.01, 148.90 (3C, NC=O); 132.05 (1C,

CH2CH=CH2); 118.22 (1C, CH2CH=CH2); 67.90 (1C, CH2CH=CH2); 64.33 (1C, CH2OAOC); 57.43 (2C,

CH2OH); 44.30 (2C, CH2CH2OH); 40.97 (1C, CH2CH2OAOC)

V.3.22. DMT-TN (37)

1-(2-Dimethoxytrityloxy ethyl)-3,5-bis(2-hydroxy ethyl) isocyanurate

C30H33N3O8 M = 563.61 g/mol

5.00 g (19.1 mmol) 1,3,5-tris(2-hydroxyethyl)isocyanurate were dissolved in 20 mL 1:1

DMFabs/Pyabs and the solution stirred in an atmosphere of argon. A solution of 5.19 g (15.3 mmol) 4,4’-dimethoxytrityl chloride in 30 mL Pyabs was added dropwise under vigorous stirring at room temperature over a period of 1 h. The mixture stirred for another 30 min at room temperature and for 16 h at 60 °C (TLC: DCM/MeOH, 95:5, Rf: 0.2; HCl stain). The reaction was quenched with 10 mL MeOH and stirring at room temperature for 10 min. The solvent was condensed and the yellow suspension was purified via column chromatography on silica (DCM/MeOH, 100:1 + 0.5% NEt3). The yellow foam was evaporated twice with 10 mL p.a. DCM and dried in high vacuum.

Yield: 3.3 g; 4.8 mmol; 30 % of theory

149 V. Experimental Section

1 3 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 7.42-6.75 (m, 13H, ArH); 4.81 (t, J=6.01 Hz, 2H, OH);

3 3 4.02 (t, J=6.3 Hz, 2H, CH2ODMT); 3.82 (t, J=6.3 Hz, 4H, CH2OH); 3.75 (s, 6H, CH3); 3.52 (q,

3 3 J=6.3 Hz, 4H, CH2CH2OH); 3.24 (t, J=5.3 Hz, 2H, CH2CH2ODMT)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 157.64 (2C, PhO, C-1’); 149.20, 148.16 (3C, NC=O); 144.43 (1C, Ph, C-1’’); 135.69 (2C, PhO, C-4’); 129.15 (4C, PhO, C-3’,-5’); 128.52 (2C, Ph, C-2’’,-6’’);

127.33 (2C, Ph, C-3’’,-5’’); 126.20 (1C, Ph, C-4’’); 112.68 (2C, PhO, C-2’,-6’); 85.11 (1C, CAr3); 59.38

(1C, CH2ODMT); 57.07 (2C, CH2OH); 54.58 (2C, OCH3); 43.75, 43.21 (2C, CH2CH2OH); 41.60 (1C,

CH2CH2ODMT);

Melting point: 129-136 °C (contains traces of triethylamine)

V.3.23. AOC-DMT-TN (38)

1-(2-Allyloxycarboxyl ethyl)-3-(2-dimethoxytrityloxy ethyl)-5-(2-hydroxy ethyl) isocyanurate

C34H37N3O10 M = 647.68 g/mol

To 3.00 g (5.25 mmol) 37, 7 mL THFabs and 0.51 mL (0.51 g, 6.45 mmol) Pyabs stirred in an atmosphere of argon and cooled to 0 °C, a solution of 0.45 mL (0.51 g, 4.2 mmol) allyl chloroformate in 45 mL THFabs was added dropwise under vigorous stirring over a period of 1 h. The mixture stirred at room temperature for 4 h after the addition was completed (TLC:

DCM/MeOH; 95:5; Rf: 0.5; HCl stain). The reaction was quenched with 7 mL MeOH and stirring at room temperature for 10 min. The solid was filtered off and the filtrate was evaporated. The residue was purified via column chromatography on silica DCM/MeOH, 200:1 + 0.5%

NEt3). The product was evaporated with 10 mL p.a. DCM and dried in high vacuum.

Yield: 1.04 g; 0.15 mmol; 30 % of theory

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] = 7.42-6.75 (m, 13H, ArH); 6.00-5.72 (m, 1H,

3 4 3 OCH2CH=CH2); 5.37-5.15 (dm, J=1.6 Hz; J=1.1 Hz, 2H, OCH2CH=CH2); 4.81 (t, J=6.0 Hz, 1H,

150 V. Experimental Section

3 3 OH); 4.60 (dt, J=4.6, 1.4 Hz, 2H, OCH2CH=CH2); 4.28 (t, J=5.3 Hz, 2H, CH2CH2OAOC); 4.08 (t,

3 J=5.3 Hz, 2H, CH2CH2OAOC); 3.78-3.54 (m, 4H, CH2CH2OH + CH2CH2OH); 3.72 (s, 6H, CH3); 3.78

3 3 (t, J=6.3 Hz, 2H, CH2ODMT); 3.22 (t, J=5.3 Hz, 2H, CH2CH2ODMT)

1 H-NMR (400 MHz; DMSO-d6): δ [ppm] = 7.37-7.07 (m, 9H, ArH); 6.90-6.78 (m, 4H, ArH); 5.84

3 (dddd, J=22.3, 17.1, 10.8, 5.5 Hz, 1H, CH=CH2); 5.34-5.14 (m, 2H, CH=CH2); 4.53 + 4.49 (dt,

3 3 J=5.6, 1.6 Hz, 2H, OCH2CH=CH2); 4.23 + 4.19 (t, J=5.35 Hz, 2H, CH2CH2OAOC); 4.05-3.95 (m,

3 2H, CH2CH2OAOC); 3.92 (t, J=6.3 Hz, 2H, CH2ODMT); 3.75-3.72 (m, 4H, CH2CH2OH +

3 CH2CH2OH); 3.71 (s, 6H, CH3); 3.21 (t, J=5.9 Hz, 2H, CH2CH2ODMT)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 158.00 (2C, PhO, C-1’); 154.10, 154.03 (1C, OCO2); 148.54,

148.51 (3C, NC=O); 144.76 (1C, Ph, C-1’’); 135.45 (2C, PhO, C-4’); 131.99, 131.95 (1C, CH2CH=CH2); 129.49, 128.88 (4C, PhO, C-3’,-5’); 127.72, 127.69, 127.61 (2C, Ph, C-2’’,-6’’); 127.51, 127.46, 127.35

(2C, Ph, C-3’’,-5’’); 126.59, 126.20 (1C, Ph, C-4’’); 118.20, 118.13 (1C, CH2CH=CH2); 113.07, 113.04

(2C, PhO, C-2’,-6’); 85.49 (1C, CAr3); 67.87, 67.81 (1C, CH2CH=CH2); 64.22 (1C, CH2OAOC); 59.66,

59.64 (1C, CH2ODMT); 57.68 (1C, CH2OH); 54.97, 54.94 (2C, OCH3); 43.05 (1C, CH2CH2OH); 42.03

(1C, CH2CH2ODMT); 41.04 (1C, CH2CH2OAOC)

V.3.24. AOC-DMT-PNO-TN (39)

1-(2-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy] ethyl)-3-(2-allyloxycarboxyl ethyl)- 5-(2-dimethoxytrityloxy ethyl) isocyanurate

C43H54N5O11P M = 847.90 g/mol

1.05 g (1.62 mmol) of 38 was diluted with 20 mL DCMabs in an atmosphere of argon. 565 µL (419.06 mg, 3.24 mmol) of DIPEA was added. To the stirring solution 724 µL (767.40 mg, 3.24

151 V. Experimental Section mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h (TLC: CH/EE, 2:1 +0.5 % NEt3). The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE,

2:1, +0.5 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 0.62 g; 0.73 mmol; 45 % of theory

1 H-NMR (250 MHz; DMSO-d6): δ [ppm] = 7.34-7.04 (m, 9 H, ArH); 6.78-6.67 (m, 4 H, ArH);

3 4 5.90-5.72 (m, 1H, CH=CH2); 5.31-5.12 (m, 2H, CH=CH2); 4.51 (dt, J=5.7 Hz; J=1.4 Hz, 2H,

CH2CH=CH2); 4.28-4.19 (m, 2H, CH2OAOC); 4.14-3.93 (m, 6H, CH2ODMT + CH2CH2OAOC +

3 4 ArCH2CH2OP); 3.71 (s, 6H, OCH3); 3.77-3.63 (m, 4H, CH2OPOCH2); 3.48 (dp, J=10.3 Hz; J=6.8

3 3 Hz, 2H, NCH); 3.34 (t, J=11.7 Hz, 2H, CH2CH2ODMT); 2.49 (t, J=5.7 Hz, 2H, CH2CN); 1.09 (d,

3 3 J=6.8 Hz, 6H, CHCH3); 1.04 (d, J=6.8 Hz, 6H, CHCH3)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 158.45 (2C, PhO, C-1’); 154.71 (1C, OCO2); 148.95, 148.75,

148.54 (3C, NC=O); 144.74 (1C, Ph, C-1’’); 135.93 (2C, PhO, C-4’); 131.53 (1C, CH2CH=CH2); 129.96 (4C, PhO, C-3’,-5’); 128.06 (2C, Ph, C-2’’,-6’’); 127.78 (2C, Ph, C-3’’,-5’’); 126.73 (1C, Ph, C-4’’);

118.76 (1C, CH2CH=CH2); 117.63 (1C, CN); 113.07 (2C, PhO, C-2’,-6’); 86.20 (1C, CAr3); 68.95 (1C,

CH2CH=CH2); 64.37 (1C, CH2OAOC); 60.11 (1C, CH2ODMT); 59.71, 59.40 (1C, CH2CH2CN); 58.70,

58.40 (1C, NCH2CH2OP); 55.14 (2C, OCH3); 43.43, 43.33 (1C, NCH2CH2OP); 43.19, 42.98 (2C, CHCH3;

42.90 (1C, CH2CH2ODMT); 41.49 (1C, CH2CH2OAOC); 24.64, 24.53, 24.43 (4C, CHCH3); 20.32, 20.23

(1C, CH2CN)

31 P-NMR (101 MHz; CDCl3); δ [ppm] = 148.36 (s, 1P, P)

152 V. Experimental Section

V.3.25. DMT-DMT-PNO-TN (40)

1,3-Bis(2-dimethoxytrityloxy ethyl)-5-(2-hydroxy ethyl) isocyanurate

C51H51N3O10 M = 865.98 g/mol

5.00 g (19.1 mmol) 1,3,5-tris(2-hydroxyethyl)isocyanurate were dissolved in 20 mL 1:1

DMFabs/Pyabs and the solution stirred in an atmosphere of argon. A solution of 12.32 g (36.37 mmol) 4,4’-dimethoxytrityl chloride in 40 mL Pyabs was added dropwise under vigorous stirring at room temperature over a period of 1.5 h. The mixture stirred for another 30 min at room temperature and for 16 h at 60 °C (TLC: CH/EE, 1:1, Rf: ~0.3, HCl stain). The reaction was quenched with 10 mL MeOH and stirring at room temperature for 10 min. Py*HCl was filtered off and the filtrate was condensed. The residue was purified via column chromatography on silica (CH/EE, 1:1 + 0.5% NEt3). The faint yellow foam was coevaporated twice with 10 mL p.a. DCM and dried in high vacuum.

Yield: 5.47 g; 6.32 mmol; 33 % of theory

1 H-NMR (400 MHz; DMSO-d6): δ [ppm] = 7.35-7.21 (m, 18H, ArH); 6.85-6.76 (m, 8H, ArH); 4.76

3 3 3 (t, J=5.9 Hz, 1H, OH); 3.95 (m, J=5.9 Hz, 4H, CH2ODMT); 3.77-3.72 (m, J=6.3 Hz, 2H, CH2OH);

3 3 3.71 (s, 6H, CH3); 3.47 (q, J=6.5 Hz, 2H, CH2CH2OH); 3.21 (t, J=5.9 Hz, 4H, CH2CH2ODMT)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 158.00 (4C, PhO, C-1’); 148.52, 148.41 (3C, NC=O); 144.79 (2C, Ph, C-1’’); 135.44 (4C, PhO, C-4’); 129.51 (8C, PhO, C-3’,-5’); 127.69 (4C, Ph, C-2’’,-6’’);

127.47 (4C, Ph, C-3’’,-5’’); 126.56 (2C, Ph, C-4’’); 113.04 (4C, PhO, C-2’,-6’); 85.47 (2C, CAr3); 59.74

(1C, CH2ODMT); 57.43 (1C, CH2OH); 54.94 (2C, OCH3); 44.12 (1C, CH2CH2OH); 41.96 (2C,

CH2CH2ODMT)

Melting point: 148-168 °C (contains triethylamine)

153 V. Experimental Section

V.3.26. DMT-DMT-PNO-TN (41)

1-(2-[2-Cyanoethoxy-diisopropylamino-phosphanyl-oxy]ethyl)-3,5-bis-(2- dimethoxytrityloxy ethyl) isocyanurate

C60H68N5O11P M = 1066.20 g/mol

1.00 g (1.15 mmol) of 40 was diluted with 20 mL DCMabs in an atmosphere of argon. 402 µL (298.50 mg, 2.31 mmol) of DIPEA was added. To the stirring solution 516 µL (564.63 mg, 2.31 mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 2 h (TLC: CH/EE, 2:1 +0.5 % NEt3). The solvent was evaporated (flush with argon only) and the residue was purified via column chromatography on silica (CH/EE,

2:1, +0.5 % NEt3). A clear viscous oil was obtained, which was stored in an atmosphere of argon.

Yield: 0.54 g; mmol; 44 % of theory

1 H-NMR (250 MHz; DMSO-d6): δ [ppm] = 7.34-7.03 (m, 18 H, ArH); 7.72-6.63 (m, 8 H, ArH);

4.05-3.86 (m, 6H, CH2ODMT + ArCH2CH2OP); 3.68 (s, 12H, OCH3); 3.73-3.59 (m, 4H,

3 4 3 CH2OPOCH2); 3.46 (dp, J=10.0 Hz, J=6.7 Hz, 2H, NCH); 3.28 (t, J=5.6 Hz, 2H, CH2CH2ODMT);

3 3 3 2.42 (t, J=6.5 Hz, 2H, CH2CN); 1.07 (d, J=6.8 Hz, 6H, CHCH3); 1.01 (d, J=6.8 Hz, 6H, CHCH3)

13 C-NMR (63 MHz; DMSO-d6): δ [ppm] = 158.42 (4C, PhO, C-1’); 148.64 (3C, NC=O); 144.80 (1C, Ph, C-1’’); 135.93 (4C, PhO, C-4’); 129.96 (4C, PhO, C-3’,-5’); 128.03 (2C, Ph, C-2’’,-6’’); 127.78 (2C,

Ph, C-3’’,-5’’); 126.71 (1C, Ph, C-4’’); 118.76 (1C, CN); 113.05 (2C, PhO, C-2’,-6’); 86.17 (1C, CAr3);

60.22 (1C, CH2ODMT); 59.68 (1C, CH2CH2CN); 59.45 (1C, NCH2CH2OP); 55.15 (2C, OCH3); 43.55 (1C,

NCH2CH2OP); 43.19 (2C, CHCH3); 42.44 (1C, CH2CH2ODMT); 24.54, 24.42 (4C, CHCH3); 20.16 (1C,

CH2CN)

31 P-NMR (101 MHz; CDCl3); δ [ppm] = 148.49 (s, 1P, P)

154 V. Experimental Section

V.3.27. Perfluorooctyl ethylthioethyl hydroxide (47)

C12H9F17OS M = 524.24 g/mol

3.41 g (4.40 mL, 46.00 mmol) tert-butanol, 0.42 g (10.45 mmol) NaOH and 733 µL (0.82 g, 10.45 mmol) mercaptoethanol were refluxed for 20 min in an atmosphere of argon. 6.00 g (10.45 mmol) perfluorooctyl ethyl iodide was added over a period of 30 min and then refluxed for another 3h. Solvent was evaporated. The residue was suspended in chloroform and solid materials were filtered off. The organic phase was evaporated. The colourless solid was dried in high vacuum and was sufficiently pure for further synthesis.

Yield: 4.11 g; 7.84 mmol; 75 % of theory

1 3 H-NMR (200 MHz; CDCl3): δ [ppm] = 3.71 (t, J=11.7 Hz, 2H, CH2O); 2.77-2.62 (m, 4H, CH2SCH2);

2.49-2.18 (m, CH2CF2); 1.81 (s, 1H, OH)

13 C-NMR (101 MHz; DMSO-d6): δ [ppm] = 60.19 (1C, CH2OH); 33.48 (1C, SCH2CH2O); 30.98, 30.76,

30.54 (1C, CH2CF2); 21.43 (1C, SCH2CH2CF2);

Melting point: 68-72 °C (Lit.[286] 71-72 °C)

V.3.28. F-TAG amidite (48)

2-Cyanoethyl-2-perfluorooctylethylthioethyl-N,N-diisopropylphosphoramidite

C21H26F17N2O2PS M = 724.46 g/mol

1.00 g (1.91 mmol) of 47 was diluted with 25 mL DCMabs in an atmosphere of argon. 498 µL (677.21 mg, 2.86 mmol) of DIPEA was added. To the stirring solution 638 µL (677.21 mg, 2.86 mmol) Bannwarth-reagent was added slowly via a syringe. The reaction mixture stirred at room temperature for 1.5 h. The solution was washed once with 25 mL deionized water, the organic phase was dried with MgSO4 and then evaporated (flush only with argon). The raw product was purified via column chromatography on silica (Automated: CH/EE, +0.75 % NEt3,

155 V. Experimental Section

5 %-30 % over 10 min). A clear colourless oil was obtained, which was stored in an atmosphere of argon.

Yield: 520 mg; 0.72 mmol; 38 % of theory

1 3 H-NMR (250 MHz; CDCl3): δ [ppm] = 3.90-3.64 (m, 4H, CH2OPOCH2); 3.62-3.45 (dp, J=10.2

4 3 Hz, J=6.8 Hz, 2H, NCH); 2.80-2.67 (m, 4H, CH2SCH2); 2.57 (t, J=6.3 Hz, 2H, CH2CN); 2.46-2.19

3 3 (m, 2H, CH2CF2); 1.13 (d, J=2.1 Hz, 6H, CHCH3); 1.11 (d, J=2.2 Hz, 6H, CHCH3)

13 C-NMR (63 MHz; CDCl3): δ [ppm] = 117.44 (1C, CN); 63.10, 62.83 (1C, SCH2CH2OP); 58.54, 58.24

(1C, CH2CH2CN); 43.15, 42,95 (2C, CH(CH3)2); 32.44, 32.36 (1C, OCH2CH2S); 31.51, 31.30, 31.08 (1C,

CH2CF2); 24.49, 24.42, 24.37, 24.30 (4C, CHCH3); 23.06, 22.99, 22.92 (1C, SCH2CH2CF2); 20.30,

20.19 (1C, CH2CN)

31 P-NMR (101 MHz; CDCl3); δ [ppm] = 148.36 (s, 1P, P)

156 VI. References

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167 VII. Appendix

VII. Appendix

VII.1 Glossary

°C degrees Celsius

A adenine; ampere

α alpha abs absolute

Ac acetyl

ACN acetonitrile

AFM atomic force microscopy

AHC ammonium hydrogen carbonate

AOC allyloxycarbonyl

APS ammonium persulfate aq. aqueous

Ar aryl

B nucleobase in oligonucleotides

B’ protected nucleobase in oligonucleotides

β beta

BHT butylated hydroxytoluene

5-BMT 5-benzylmercapto-1H-tetrazole

Bn benzyl bp basepair(s) b.p. boiling point

Br bromine

168 VII. Appendix

C cytosine cal. calorie(s)

Cl chloride conc. concentration

CPG controlled pore glass d 2’-deoxyrobose in oligonucleotides; days; doublet

δ chemical shift

ΔG Gibbs free energy dba dibenzylideneacetone

DBPO dibenzoylperoxide

DCM dichloromethane denat. denaturing (electrophoresis)

ddH2O double distilled water dest. distilled

DIPEA N,N-Diisopropylethylamine; Hünig's base

DLP drug loading protocol

DMAP N,N-4-dimethylaminopyridine

DMF N,N-dimethylformamide

DMSO dimethylsulfoxide

DMT 4,4’-dimethoxytrityl

DNA deoxyribonucleic acid

DTT 1,4-dithio-DL-threitol ds double-stranded

EDTA ethylenediaminetetraacetic acid

ε extinction (or absorption) coefficient

169 VII. Appendix eq. equivalent

Fig. figure g gram

G guanine h hour

HB hybridization buffer

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

HPA 3-Hydroxypicolinic acid

HPLC high performance liquid chromatography

HBTU O-(benzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium hexafluorophosphate

I iodine

InChI IUPAC International Chemical Identifier k kilo kc/m kilocalories per mole

L litre m milli; multiplet; metre

M molar; mol/L; molecular mass

µ micro

MALDI-TOF matrix assisted laser desorption ionisation time of flight

MBN mung bean nuclease

Me methyl

MeOH methanol min minutes mol mole: Avogadro constant: ~6.0221 ∙ 1023 units

MPPS macroporous polystyrene

170 VII. Appendix

MS mass spectrometry n amount of substance; nano nat. native (electrophoresis)

NBS N-bromosuccinimide nm nanometre

NMR nuclear magnetic resonance

Nt nucleotide

π pi p pico; quintet p.a. purum analyticum

PAA acrylamide/bisacrylamide 19:1 stock solution

PAGE polyacrylamide gel electrophoresis prep. preparative (electrophoresis)

Pd palladium

Pd(OAc)2 palladium(II)acetate

Ph phenyl

+ pH -log10[H3O ]

PP polypropylene

PPh3 triphenylphosin ppm parts per million

Ψ psi

PBS phosphate buffered saline

PNO 2-cyanoethyl-N,N-diisopropyl-phosphoramidite

Py pyridine

PySSPy 2,2’-dipyridyl-disulfide

171 VII. Appendix q quadruplet/quartet

RNA ribonucleic acid

RP reversed phase (columns in HPLC) rpm rounds per minute r.t. room temperature s second; singlet sat. saturated

SDS sodium dodecyl sulfate ss single-stranded m.p. melting point t triplet

T thymine

Tab. table

TAP trisoligonucleotide assembly protocol

TBDMS tert.-butyl-dimethylsilyl

TEM transmission electron microscopy

TEA triethylamine

TEMED N,N,N’,N’-tetramethylethylenediamine

THAP 2,4,6-trihydroxyacetophenone

THEIC 1,3,5-tris(2-hydroxyethyl)-N,N’,N’’-isocyanurate

THF tetrahydrofuran

THP 3-hydroxypicolinic acid

TLC thin layer chromatography

Tm melting temperature of DNA

TMS trimethylsilyl

172 VII. Appendix

TOTP tri(o-tolyl)phosphine

Trityl Triphenylmethyl

U units of enzyme

UV ultraviolet photometric range between ~10 to ~380 nm

V volt

VIS visible photometric range between~380 to ~750 nm

173 VII. Appendix

VII.2 Parr Hydrogenation Apparatus German Instruction Manual

Written by Dr. Wolf Matthias Pankau

174 VII. Appendix

VII.3 Full Sequence Pool

DNASequenceGenerator; Nearest Neighbour Model; SantaLucia[247] parameters;

Concentration= 1.25 µM; 110 mM NaCl; set Tm = 58.3-58.4 °C; No Fraying;

Longest Longest Most stable No. Sequence 5’-3’ GC % T [°C] m hairpin palindrome dimer

0 CCTCCTGTCTCATCC 0.60 58.33 2 2 2 1 CAGGGTACATCCGAC 0.60 58.36 2 4 4 2 GCTATTTGCTTGCAG 0.47 58.35 3 4 4 3 GGTTCTGCATGATGC 0.53 58.37 4 4 4 4 GCCGAATTAACCGAC 0.53 58.30 2 4 4 5 CCGCCTTCATACTAG 0.53 58.31 2 4 4 6 CCACTATGGTCGTCC 0.60 58.36 3 2 3 7 CGCTACTTTTAGGGG 0.53 58.33 3 2 3 8 GTGACCAAATCGTCG 0.53 58.32 3 2 3 9 CACGAACACTACCCC 0.60 58.39 1 2 2 10 CACGTTGGAAACGTC 0.53 58.35 5 4 5 11 CGCTTATGAGTCCTC 0.53 58.40 3 2 3 12 GAATGCGAGAGGTAG 0.53 58.40 1 2 2 13 GAAGTCCCGGTATGG 0.60 58.35 2 4 4 14 CTCACCCGTACCAAC 0.60 58.39 2 4 4 15 CATCTAAGGCTACCG 0.53 58.31 2 2 2 16 GGTGTGTTAGGTCGG 0.60 58.39 1 2 2 17 GATGACCTCAGAGGG 0.60 58.33 4 2 4 18 CGAAGCATAGACTCC 0.53 58.40 2 2 2 19 GCGTCTACATATCCG 0.53 58.40 2 4 4 20 CTGAGTTCCCACTCC 0.60 58.36 4 0 4 21 CATTCAACGACGGAC 0.53 58.32 2 2 2 22 GAGGAGACTTTGGGG 0.60 58.35 2 0 2 23 CTCTGTGTATAGGCG 0.53 58.32 2 4 4 24 GGACGGTGGGTTTAG 0.60 58.38 2 2 2

175 VII. Appendix

Longest Longest Most stable No. Sequence 5’-3’ GC % T [°C] m hairpin palindrome dimer

25 GTTTGCATTGGGAGC 0.53 58.39 2 4 4 26 GTAGCAATCTCCGAG 0.53 58.40 3 2 3 27 GAAAGCGGTGTCTTC 0.53 58.31 3 2 3 28 CCGGTCAGGGAATAC 0.60 58.35 2 4 4 29 GCCCCTCATTTTTGC 0.53 58.37 2 2 2 30 GGCAGGTGCAATTTC 0.53 58.39 3 4 4 31 CATGTATTGCGCTTG 0.47 58.36 2 4 4 32 GGTAGAGTTGCCTCC 0.60 58.35 3 2 3 33 CACAGGGGTCGATAC 0.60 58.36 2 4 4 34 GGCTTGTGCCATTTC 0.53 58.39 3 2 3 35 CGGCCCTTTAGTAAG 0.53 58.33 3 4 4 36 CGAGCAACCTATCTC 0.53 58.40 3 2 3 37 CGCCTGGACTAATAG 0.53 58.31 3 2 3 38 GTGGAGGGACGTATG 0.60 58.36 2 4 4 39 CAGAAGTTCTCCCCC 0.60 58.35 4 0 4 40 CGGCCATAACTAGAG 0.53 58.31 2 4 4 41 GGAACGAAGAAAGGC 0.53 58.30 1 2 2 42 GTAACGGAACAACGC 0.53 58.35 2 2 2 43 CCGAAGTGTAACCCC 0.60 58.38 2 2 2 44 GACCCTTATATCGCG 0.53 58.38 2 4 4 45 CTGAGATGATCTCGG 0.53 58.37 5 4 5 46 CAGATCCAGAGACCC 0.60 58.33 2 4 4 47 GAACGGTACTGTGGG 0.60 58.39 2 4 4 48 CGTACTACTGCCAAG 0.53 58.36 3 4 4 49 CGCCAGATACTCTTC 0.53 58.40 3 2 3

No. Sequence 5’-3’ GC % T [°C] Longest Longest Most stable m hairpin palindrome dimer

50 GAAGCGTATCCTAGC 0.53 58.39 2 4 4 51 CAGGATCGGAAAGAG 0.53 58.39 2 4 4 52 GGTATTACTCGCCCC 0.60 58.34 3 2 3

176 VII. Appendix

53 CGGAGTTTACCACCC 0.60 58.38 2 2 2 54 GACACAACCGTAGGG 0.60 58.39 2 2 2 55 GGCTCAATGTTTGGC 0.53 58.39 3 2 3 56 GCCGGATTATCGTAG 0.53 58.38 3 4 4 57 GCCGGTGATACCTAC 0.60 58.35 3 4 4 58 CCGCAGATTCTACTC 0.53 58.40 3 2 3 59 CCCAAGCTCTCTTAG 0.53 58.31 3 4 4 60 GACTAGCAGCACTAG 0.53 58.33 4 4 4 61 GTTTTCTCTCACGCC 0.53 58.31 2 2 2 62 GGGCTGATTCATTGC 0.53 58.36 3 2 3 63 GTACGCGGATCAAAC 0.53 58.31 2 4 4 64 CTGTCCCAGACCTTC 0.60 58.36 3 0 3 65 GCGACTCTCCAATAG 0.53 58.40 2 2 2 66 CCAGGGCATTTGTTG 0.53 58.31 2 2 2 67 GATCACGAGCCTAAG 0.53 58.40 2 4 4 68 CACGGTCTTATCCCC 0.60 58.35 2 2 2 69 CTCATGTGTGGCATG 0.53 58.30 4 4 4 70 GTACACGGAGGATGG 0.60 58.36 2 4 4 71 CTATAACGAGGCCAG 0.53 58.31 2 4 4 72 CTAGCTGCCTAGAAC 0.53 58.31 4 4 4 73 CCAGTGTCCTTCCTC 0.60 58.36 2 0 2 74 GAATATACGGCTCCG 0.53 58.38 3 4 4

No. Sequence 5’-3’ GC % T [°C] Longest Longest Most stable m hairpin palindrome dimer

75 CCTTGGACCACGTAC 0.60 58.39 3 4 4 76 GGGTGCGGTAAGTAC 0.60 58.38 3 4 4 77 GCAACATGGATGCTC 0.53 58.37 3 4 4 78 CAATGGTTGGCTGTG 0.53 58.32 3 2 3 79 CTACTATCGGCACTG 0.53 58.32 2 2 2 80 CATTCCGGTTTCGAC 0.53 58.31 2 4 4 81 CCTGTATGTCTAGCG 0.53 58.32 2 4 4

177 VII. Appendix

82 CCACCAGTCTGACTC 0.60 58.38 4 0 4 83 GTCCGAACGTAATGC 0.53 58.31 2 4 4 84 CAGTCCGCTCTAATC 0.53 58.40 2 2 2 85 GACAACAGTTCGAGC 0.53 58.33 3 4 4 86 CGACATTCGTCAACC 0.53 58.32 3 2 3 87 CGTAGCTTTACTGGG 0.53 58.34 3 4 4 88 GCAGACGAGTGAAAC 0.53 58.33 2 2 2 89 GGAAATACGACACGC 0.53 58.31 2 2 2 90 GGCTTCAGGTCCTAC 0.60 58.35 3 2 3 91 CCTGGGTAATCGGTC 0.60 58.35 2 2 2 92 CTAAGTTGTAGGCCG 0.53 58.34 3 4 4 93 CGGGGGTGTAGTTTC 0.60 58.38 1 2 2 94 GCCAATTCTTTGCCC 0.53 58.37 3 4 4 95 GAACCCGCGAATAAC 0.53 58.30 2 4 4 96 GACAGAAACAGCGAC 0.53 58.33 1 2 2 97 GTTGAGGCGATCTAG 0.53 58.40 2 4 4 98 CGATGGAGCTAACTC 0.53 58.40 3 4 4 99 GTTCGCTCCTTTGTC 0.53 58.31 1 2 2

178 VII. Appendix

VII.4 Dimer Energies PrimerSelect 3.11: Energies are calculated according to Breslauer et al.[248] (Nearest- Neighbour model); c= 1.25 µM; c(Salt): 110 mM

Comparison between the self pair dimer energies of the old set and the new set:

179 VII. Appendix

Ten most staple mismatch pair dimer energies out of 427 to the left and ten most stable self dimers out of 37 to the right:

180 VII. Appendix

All hairpin formation energies:

181 VII. Appendix

VII.5 Full MALDI-TOF-MS Data Set

182 VII. Appendix

183 VII. Appendix

184 VII. Appendix

185 VII. Appendix

186 VII. Appendix

VII.6 Alternative Visualisation of Anthracycline Binding Efficiencies

Alternative visualisation based on data presented in the main text. Samples of similar DNA concentrations were overlaid. 30, 60 and 80 % are dilutions of the 100 % DNA stock solutions. The strong red profiles are from oligomix 6D and show a stronger fluorescence than the tetrahedrons T2 and T3.

187 VII. Appendix

VII.7 NMR-Spectra V.3.01. AOC-DMT-PNO-T1 (05)

1 H-NMR (250 MHz; CDCl3): CH

InChI=1S/C43H51N2O7P/c1-8-13-42(46)49-29-34-26-35(28-36(27-34)31-52-53(51-25-12-24-44)45(32(2)3)33(4)5)30-50-43(37- 14-10-9-11-15-37,38-16-20-40(47-6)21-17-38)39-18-22-41(48-7)23-19-39/h8-11,14-23,26-28,32-33H,1,12-13,25,29-31H2,2-7H3

188 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

189 VII. Appendix

31 P-NMR (101 MHz; CDCl3);

190 VII. Appendix

V.3.02. 1,3,5-Triacetybenzene (12)

1 H-NMR (400 MHz; CDCl3):

InChI=1S/C12H12O3/c1-7(13)10-4-11(8(2)14)6-12(5-10)9(3)15/h4-6H,1-3H3

191 VII. Appendix

13 C-NMR (101 MHz; CDCl3):

192 VII. Appendix

V.3.03. 2,2',2''-(Benzene-1,3,5-triyl)triacetic acid (08)

1 H-NMR (200 MHz; DMSO-d6):

InChI=1S/C12H12O6/c13-10(14)4-7-1-8(5-11(15)16)3-9(2-7)6-12(17)18/h1-3H,4-6H2,(H,13,14)(H,15,16)(H,17,18)

193 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

194 VII. Appendix

V.3.04. 2,2',2''-(Benzene-1,3,5-triyl)triacetic acid (09)

1 H-NMR (200 MHz; CDCl3):

InChI=1S/C15H18O6/c1-19-13(16)7-10-4-11(8-14(17)20-2)6-12(5-10)9-15(18)21-3/h4-6H,7-9H2,1-3H3

195 VII. Appendix

13 C-NMR (50 MHz; CDCl3):

196 VII. Appendix

V.3.05. 1,3,5-Trishydroxyethylbenzene (10)

H-NMR (200 MHz; DMSO-d6):

InChI=1S/C12H18O3/c13-4-1-10-7-11(2-5-14)9-12(8-10)3-6-15/h7-9,13-15H,1-6H2

197 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

198 VII. Appendix

V.3.06. AOC-AOC-T2 (18)

1 H-NMR (200 MHz; DMSO-d6):

water

InChI=1S/C20H26O7/c1-3-9-24-19(22)26-11-6-17-13-16(5-8-21)14-18(15-17)7-12-27-20(23)25-10-4-2/h3-4,13-15,21H,1-2,5-12H2

199 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

200 VII. Appendix

V.3.07. AOC-AOC-PNO-T2 (19)

1 H-NMR (250 MHz; CDCl3):

InChI=1S/C29H43N2O8P/c1-7-14-34-28(32)36-17-10-25-20-26(11-18-37-29(33)35-15-8-2)22-27(21-25)12-19-39-40(38-16-9- 13-30)31(23(3)4)24(5)6/h7-8,20-24H,1-2,9-12,14-19H2,3-6H3

201 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

202 VII. Appendix

31 P-NMR (101 MHz; CDCl3):

203 VII. Appendix

V.3.08. DMT-DMT-T2 (16)

1 H-NMR (200 MHz; DMSO-d6):

InChI=1S/C54H54O7/c1-56-49-23-15-45(16-24-49)53(43-11-7-5-8-12-43,46-17-25-50(57-2)26-18-46)60-35-32-41-37-40(31- 34-55)38-42(39-41)33-36-61-54(44-13-9-6-10-14-44,47-19-27-51(58-3)28-20-47)48-21-29-52(59-4)30-22-48/h5-30,37- 39,55H,31-36H2,1-4H3

204 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

205 VII. Appendix

V.3.09. DMT-DMT-PNO-T2 (17)

1 H-NMR (250 MHz; CDCl3):

DCM CH

InChI=1S/C63H71N2O8P/c1-47(2)65(48(3)4)74(72-40-15-39-64)73-43-38-51-45-49(36-41-70-62(52-16-11-9-12-17-52,54-20- 28-58(66-5)29-21-54)55-22-30-59(67-6)31-23-55)44-50(46-51)37-42-71-63(53-18-13-10-14-19-53,56-24-32-60(68-7)33-25- 56)57-26-34-61(69-8)35-27-57/h9-14,16-35,44-48H,15,36-38,40-43H2,1-8H3

206 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

207 VII. Appendix

31 P-NMR (101 MHz, CDCl3):

208 VII. Appendix

V.3.10. DMT-T2 (13)

1 H-NMR (200 MHz; DMSO-d6): signal + signal +

impurity NEt3

InChI=1S/C33H36O5/c1-36-31-12-8-29(9-13-31)33(28-6-4-3-5-7-28,30-10-14-32(37-2)15-11-30)38-21-18-27-23-25(16-19- 34)22-26(24-27)17-20-35/h3-15,22-24,34-35H,16-21H2,1-2H3

209 VII. Appendix

V.3.11. AOC-DMT-T2 (14)

1 H-NMR (200 MHz; DMSO-d6): signal + water

InChI=1S/C37H40O7/c1-4-22-42-36(39)43-23-19-29-25-28(18-21-38)26-30(27-29)20-24-44-37(31-8-6-5-7-9-31,32-10-14- 34(40-2)15-11-32)33-12-16-35(41-3)17-13-33/h4-17,25-27,38H,1,18-24H2,2-3H3

210 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

211 VII. Appendix

V.3.12. AOC-DMT-PNO-T2 (15)

1 H-NMR (250 MHz; DMSO-d6): CH

InChI=1S/C46H57N2O8P/c1-8-27-52-45(49)53-29-23-37-32-38(34-39(33-37)25-31-56-57(55-28-12-26- 47)48(35(2)3)36(4)5)24-30-54-46(40-13-10-9-11-14-40,41-15-19-43(50-6)20-16-41)42-17-21-44(51-7)22-18-42/h8-11,13-22,32- 36H,1,12,23-25,27-31H2,2-7H3

212 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

213 VII. Appendix

31 P-NMR (101 MHz, CDCl3);

214 VII. Appendix

V.3.13. 1,3,5-Tris(E- methyl acryl) benzene (28)

1 H-NMR (400 MHz; CDCl3):

InChI=1S/C18H18O6/c1-22-16(19)7-4-13-10-14(5-8-17(20)23-2)12-15(11-13)6-9-18(21)24-3/h4-12H,1-3H3/b7-4+,8-5+,9-6+

215 VII. Appendix

13 C-NMR (101 MHz; CDCl3):

216 VII. Appendix

V.3.14. 1,3,5-Tris(methyl ester propyl) benzene (24)

1 H-NMR (200 MHz; CDCl3):

InChI=1S/C18H24O6/c1-22-16(19)7-4-13-10-14(5-8-17(20)23-2)12-15(11-13)6-9-18(21)24-3/h10-12H,4-9H2,1-3H3

217 VII. Appendix

13 C-NMR (50 MHz; CDCl3):

218 VII. Appendix

V.3.15. Allyl benzyl ether (31)

1 H-NMR (400 MHz; CDCl3):

InChI=1S/C10H12O/c1-2-8-11-9-10-6-4-3-5-7-10/h2-7H,1,8-9H2

219 VII. Appendix

13 C-NMR (101 MHz; CDCl3):

220 VII. Appendix

V.3.16. 1E,3E,5Z-1,3,5-Tris(benzyl oxy isoallyl) benzene (32)

1 H-NMR (400 MHz; CDCl3):

InChI=1S/C36H36O3/c1-4-13-31(14-5-1)28-37-22-10-19-34-25-35(20-11-23-38-29-32-15-6-2-7-16-32)27-36(26-34)21-12-24- 39-30-33-17-8-3-9-18-33/h1-21,25-27H,22-24,28-30H2/b19-10-,20-11+,21-12+

221 VII. Appendix

13 C-NMR (101 MHz; CDCl3):

222 VII. Appendix

V.3.17. 1,3,5-Tris(3-hydroxy propyl) benzene (25)

1 H-NMR (200 MHz; DMSO-d6):

InChI=1S/C15H24O3/c16-7-1-4-13-10-14(5-2-8-17)12-15(11-13)6-3-9-18/h10-12,16-18H,1-9H2

223 VII. Appendix

13 C-NMR (50 MHz; DMSO-d6):

224 VII. Appendix

V.3.18. AOC-T3 (33)

1 H-NMR (400 MHz; DMSO-d6):

InChI=1S/C19H28O5/c1-2-11-23-19(22)24-12-5-8-18-14-16(6-3-9-20)13-17(15-18)7-4-10-21/h2,13-15,20-21H,1,3-12H2

225 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

226 VII. Appendix

V.3.19. AOC-DMT-T3 (34)

1 H-NMR (400 MHz; DMSO-d6): acetone

InChI=1S/C40H46O7/c1-4-25-45-39(42)46-26-9-12-32-28-31(11-8-24-41)29-33(30-32)13-10-27-47-40(34-14-6-5-7-15-34,35- 16-20-37(43-2)21-17-35)36-18-22-38(44-3)23-19-36/h4-7,14-23,28-30,41H,1,8-13,24-27H2,2-3H3

227 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

228 VII. Appendix

V.3.20. AOC-DMT-PNO-T3 (35)

1 H-NMR (400 MHz; CDCl3): EE

InChI=1S/C49H63N2O8P/c1-8-30-55-48(52)56-31-12-16-40-35-41(37-42(36-40)18-14-33-58-60(59-34-15-29- 50)51(38(2)3)39(4)5)17-13-32-57-49(43-19-10-9-11-20-43,44-21-25-46(53-6)26-22-44)45-23-27-47(54-7)28-24-45/h8-11,19- 28,35-39H,1,12-18,30-34H2,2-7H3

229 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

230 VII. Appendix

31 P-NMR (101 MHz, CDCl3):

231 VII. Appendix

V.3.21. AOC-TN

1 H-NMR (400 MHz; DMSO-d6):

InChI=1S/C13H19N3O8/c1-2-8-23-13(22)24-9-5-16-11(20)14(3-6-17)10(19)15(4-7-18)12(16)21/h2,17-18H,1,3-9H2

232 VII. Appendix

13 C-NMR (101 MHz; CDCl3):

233 VII. Appendix

V.3.22. DMT-TN (37)

1 H-NMR (200 MHz; DMSO-d6): δ [ppm] =

InChI=1S/C30H33N3O8/c1-39-25-12-8-23(9-13-25)30(22-6-4-3-5-7-22,24-10-14-26(40-2)15-11-24)41-21-18-33-28(37)31(16-19- 34)27(36)32(17-20-35)29(33)38/h3-15,34-35H,16-21H2,1-2H3

234 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

235 VII. Appendix

V.3.23. AOC-DMT-TN (38)

1 H-NMR (200 MHz; DMSO-d6): water

InChI=1S/C34H37N3O10/c1-4-22-45-33(42)46-23-19-36-30(39)35(18-21-38)31(40)37(32(36)41)20-24-47-34(25-8-6-5-7-9- 25,26-10-14-28(43-2)15-11-26)27-12-16-29(44-3)17-13-27/h4-17,38H,1,18-24H2,2-3H3

236 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

237 VII. Appendix

V.3.24. AOC-DMT-PNO-TN (39)

1 H-NMR (250 MHz; DMSO-d6): CH

InChI=1S/C43H54N5O11P/c1-8-27-55-42(52)56-29-24-45-39(49)46(41(51)47(40(45)50)26-31-59-60(58-28-12-23- 44)48(32(2)3)33(4)5)25-30-57-43(34-13-10-9-11-14-34,35-15-19-37(53-6)20-16-35)36-17-21-38(54-7)22-18-36/h8-11,13-22,32- 33H,1,12,24-31H2,2-7H3

238 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

239 VII. Appendix

31 P-NMR (101 MHz; CDCl3);

240 VII. Appendix

V.3.25. DMT-DMT-PNO-TN (40)

1 H-NMR (400 MHz; DMSO-d6): water

InChI=1S/C51H51N3O10/c1-59-43-23-15-39(16-24-43)50(37-11-7-5-8-12-37,40-17-25-44(60-2)26-18-40)63-35-32-53- 47(56)52(31-34-55)48(57)54(49(53)58)33-36-64-51(38-13-9-6-10-14-38,41-19-27-45(61-3)28-20-41)42-21-29-46(62-4)30-22- 42/h5-30,55H,31-36H2,1-4H3

241 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

242 VII. Appendix

V.3.26. DMT-DMT-PNO-TN (41)

1 H-NMR (250 MHz; DMSO-d6): DCM

InChI=1S/C60H68N5O11P/c1-44(2)65(45(3)4)77(75-40-15-36-61)76-43-39-64-57(67)62(37-41-73-59(46-16-11-9-12-17-46,48- 20-28-52(69-5)29-21-48)49-22-30-53(70-6)31-23-49)56(66)63(58(64)68)38-42-74-60(47-18-13-10-14-19-47,50-24-32-54(71- 7)33-25-50)51-26-34-55(72-8)35-27-51/h9-14,16-35,44-45H,15,37-43H2,1-8H3

243 VII. Appendix

13 C-NMR (63 MHz; DMSO-d6):

244 VII. Appendix

31 P-NMR (101 MHz; CDCl3);

245 VII. Appendix

V.3.27. Perfluorooctyl ethylthioethyl hydroxide (47)

1 H-NMR (200 MHz; CDCl3):

InChI=1S/C12H9F17OS/c13-5(14,1-3-31-4-2-30)6(15,16)7(17,18)8(19,20)9(21,22)10(23,24)11(25,26)12(27,28)29/h30H,1-4H2

246 VII. Appendix

13 C-NMR (101 MHz; DMSO-d6):

247 VII. Appendix

V.3.28. F-TAG amidite (48)

1 H-NMR (250 MHz; CDCl3):

InChI=1S/C21H26F17N2O2PS/c1-12(2)40(13(3)4)43(41-8-5-7-39)42-9-11-44-10-6- 14(22,23)15(24,25)16(26,27)17(28,29)18(30,31)19(32,33)20(34,35)21(36,37)38/h12-13H,5-6,8-11H2,1-4H3

248 VII. Appendix

13 C-NMR (63 MHz; CDCl3):

249 VII. Appendix

31 P-NMR (101 MHz; CDCl3);

250 VII. Appendix

VII. Curriculum Vitae

 personal details

Name: Christos Panagiotidis date of birth: 19.08.1985 birthplace: Wuppertal, NRW marital status: single citizenship: german/greek

 education and academic activities

2012 - 2016 Ph.D. in bioorganic chemistry at Ruhr-Universität Bochum

Title: “Sequence-Addressed Assemblies of Trisoligonucleotides into Nanoscale Motifs and Structural Studies” 2009 - 2011 Master of Science in chemistry at Bergische Universität Wuppertal

Title: „Versuche zur Synthese von α-phosphonylierten Imidazolidinonen“

2005 - 2009 Bachelor of Science in chemistry at Bergische Universität Wuppertal

Title: „Versuche zur Synthese von neuen spirocyclischen Imidazolidinonen“

1992 - 2005 Abitur at the Carl-Duisberg Gymnasium in Wuppertal

 additional practical experience

2009 Student assistant in the field of organic chemistry at Bergische Universität Wuppertal

 publications

. Christos Panagiotidis, Stephanie Kath-Schorr, Günter von Kiedrowski, “Flexibility of C3h- Symmetrical Linkers in Trisoligonucleotide-based Tetrahedral Scaffolds”, ChemBioChem 2016, 17(3), 254-259 . Book publication of master thesis at AV Akademikerverlag, 2014 (ISBN-13: 978-3-639-63091-6)

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