Synthesis and Characterisation of NAD + Analogues
for the Cellular Imaging of Poly(ADP-Ribos)ylation
Dissertation submitted for the degree of Doctor of Natural Sciences
Presented by
Sarah Wallrodt
at the
Faculty of Science Department of Chemistry
Date of the oral examination: 31
stMarch 2017 1
stReferee: Prof. Dr. Andreas Marx
2
ndReferee: Prof. Dr. Valentin Wittmann
3
rdReferee: Prof. Dr. Elisa Ferrando-May
This work was prepared from April 2013 to October 2016 in the group of Prof. Dr. Andreas Marx (Chair of Organic and Cellular Chemistry) at the University of Konstanz. It was funded by the “Beilstein-Institut zur Förderung der Chemischen Wissenschaften” with a stipend from 2014 to 2016 and supported by the Konstanz Research School Chemical Biology.
Parts of this work are published in:
[1] Bioorthogonally Functionalized NAD+ Analogues for In-Cell Visualization of Poly(ADP- Ribose) Formation
Sarah Wallrodt, Annette Buntz, Dr. Yan Wang, Prof. Dr. Andreas Zumbusch, Prof. Dr.
Andreas Marx
DOI: 10.1002/anie.201600464
Angewandte Chemie International Edition
Volume 55, Issue 27, pages 7660-7664, June 26, 2016.
[2] Real-Time Cellular Imaging of Protein Poly(ADP-ribos)ylation
Annette Buntz*, Sarah Wallrodt*, Eva Gwosch, Michael Schmalz, Dr. Sascha Beneke, Prof. Dr. Elisa Ferrando-May, Prof. Dr. Andreas Marx, Prof. Dr. Andreas Zumbusch
* These authors contributed equally.
DOI: 10.1002/anie.201605282
Angewandte Chemie International Edition
Volume 55, Issue 37, pages 11256-11260, September 5, 2016.
Reproduced in parts with permission. Copyright © 2016 Wiley-VCH.
[3] Investigation of the action of poly(ADP-ribose)-synthesising enzymes on NAD+ analogues
Sarah Wallrodt, Edward L. Simpson, Prof. Dr. Andreas Marx DOI: 10.3762/bjoc.13.49
Beilstein Journal of Organic Chemistry Volume 13, pages 495-501, March 10, 2017.
Reproduced in parts with permission under the terms of the Creative Commons Attribution License © 2017 Wallrodt et al.; licensee Beilstein-Institut.
Acknowledgement
First, I would like to thank Prof. Dr. Andreas Marx, who gave me the opportunity, the facilities and the confidence to complete my PhD in his group. The topic was interesting, diverse and challenging. Moreover, he has always supported my ambitions and helped me to outdo myself.
Second, I wish to thank Prof. Dr. Elisa Ferrando-May and Prof. Dr. Valentin Wittmann for their support and their kind advices within my PhD committee.
Furthermore, a special thank you goes to our collaboration partners Prof. Dr. Andreas Zumbusch and Annette Buntz, Prof. Dr. Elisa Ferrando-May and Eva Gwosch as well as Dr.
Sascha Beneke and Michael Schmalz for the productive, cooperative and successful teamwork. I am especially grateful to Annette Buntz and Brunhilde Kottwitz, who introduced me to the fascinating methods in bioimaging and cell culture.
Additionally, my sincere thanks go to the ‘Beilstein-Institut zur Förderung der Chemischen Wissenschaften’ and the Konstanz Research School for Chemical Biology for its financial and ideal support. The many supported training and education courses helped me to develop not only as a scientist but also as a professional.
Another special thank you goes to the whole working group, who welcomed me, helped me and created a productive working atmosphere. In particular, I appreciated the help and advices from the ‘biologists’ Daniel Rösner, Simon Geigges, Xiaohui Zhao, Joachim Lutz and Eva Höllmüller as well as from the ‘chemists’ Kathrin Götz, Audrey Hottin, Daniela Verga and Anke Gerull.
Additionally, I wish to thank my interns and thesis students Tamara Krämer (B.Sc.), Christoph Albrecht (B.Sc.), Johannes Bayer (B.Sc.), Tanja Sack, Daniel Hammler (M.Sc.), Mareike Rapp and Edward Simpson for their help and challenging me as a teacher. In this context, I would like to thank the German Academic Exchange Service for giving me the opportunity to take part in the programme ‘Research Internships in Science and Engineering’
in 2015 and 2016.
Furthermore, I would like to acknowledge Karin Reichhardt, Petra Jakesch, Vlasta Radusevic, Anke Gerull and Meike Liebmann for supporting the daily business in the office and lab. Also, I would like to thank Kathrin, Eva and Ralf for proof-reading my thesis.
Moreover, I wish to thank Anke Friemel and Ulrich Haunz for the performed measurements and advices on NMR spectroscopy. Additionally, I thank the groups of Prof. Dr. Elke Deuerling, Prof. Dr. Martin Scheffner and Prof. Dr. Thomas Mayer for providing me with access and help to use some of their equipment for my experiments. Also, I took advantage of the kind advices and assistance from Daniela Rothöhler from the Bioimaging Center and Dr. Stefanie Bürger and Wendy Bergmann from the FlowKon Facility.
Finally, I am deeply grateful for the support of my family and friends. They have always helped me, listened to my problems and cheered me up.
Abstract
Within the last decade, research on poly(ADP-ribose) experienced a renaissance and various studies unveiled the importance of this biopolymer as a complex, functionally diverse protein modification and signalling molecule. Fundamental processes such as DNA repair and transcription are coordinated by poly(ADP-ribose) and its binding to proteins. Thus, different pathophysiological conditions and disease states, e.g. in cancer and inflammation, have been associated with this protein modification and motivated its targeting in clinical trials.
ADP-ribose chains are tightly regulated in a posttranslational manner by ADP-ribosyl transferases and hydrolases in response to external stimuli and thus cellular levels fluctuate rapidly. Built from NAD+, single or multiple ADP-ribose units are covalently attached onto arginines, glutamates, aspartates or lysines of acceptor proteins. Thereby, nicotinamide is released and mono(ADP-ribos)ylated or poly(ADP-ribos)ylated proteins are formed. Due to these dynamics and complexity, the visualisation of poly(ADP-ribos)ylation remains not only a challenging task, but would also be of high importance to understand these processes on a cellular level.
The aim of this PhD project was to explore the applicability of chemically modified NAD+ analogues for the detection of DNA damage induced poly(ADP-ribos)ylation. Thus, a perfect analogue should be an efficient substrate of ADP-ribosyl transferases in vitro and in cellula. It should be able to specifically label cellular formation of poly(ADP-ribose) in dependency of extrinsic stimuli. Ideally, the new analogue enables to monitor poly(ADP-ribos)ylation in real- time and in a dynamic fashion.
The challenge was met by synthesising NAD+ analogues that can be built into poly(ADP- ribose) by ARTD1, the major poly(ADP-ribos)ylating enzyme in DNA repair. Thus, the positions as well as the types of NAD+ modifications were investigated, and analogues substituted in adenine position 2 were found to be best-suited for this purpose. Using bioorthogonal reporter groups, the intracellular visualisation of poly(ADP-ribose) was demonstrated simultaneously in two colours, e.g. as required in time dependent or pulse- chase experiments. Moreover, a fluorophore-modified NAD+ enabled the direct monitoring of poly(ADP-ribos)ylation inside of a living cell. Thereby, a full turnover of poly(ADP-ribose) was observed after laser-induced DNA damage in real-time. Additionally, protein-specific interaction with poly(ADP-ribose) was detected in intact cells using a powerful FLIM-FRET technique and the GFP-tagged protein of interest. Finally, the substrate scopes of other poly(ADP-ribose) synthesising enzymes like ARTD2, ARTD5 and ARTD6 were explored to broaden the applicability of the developed NAD+ analogues.
In summary, the chemical biology approaches developed in here proved powerful for biological applications and the novel tools will help to elucidate PAR biology by studying the polymer in its natural environment.
Zusammenfassung
Innerhalb des letzten Jahrzehnts erlebte die Forschung über Poly(ADP-Ribose) eine Renaissance und etliche Studien deckten die Bedeutsamkeit dieses Biopolymers als komplexe, funktionell vielfältige Proteinmodifikation und Signalmolekül auf. Fundamentale Prozesse wie DNA-Reparatur und Transkription werden von Poly(ADP-Ribose) und ihrer Bindung zu Proteinen koordiniert. Daher wurden auch zahlreiche Funktionsstörungen und Krankheiten, wie zum Beispiel bei Krebs und Entzündungen, mit dieser Proteinmodifikation in Verbindung gebracht und motivierten ihre Untersuchung in klinischen Studien.
ADP-Riboseketten werden wie andere posttranslationale Modifikationen stark von ADP- Riboysltransferasen und Hydrolasen in Abhängigkeit von externen Stimuli reguliert. Daher fluktuieren zelluläre Level rapide. Einzelne oder mehrere ADP-Riboseeinheiten aus NAD+ werden kovalent auf die Arginine, Glutamate, Aspartate oder Lysine von Akzeptorproteinen übertragen. Dabei wird Nikotinamid abgespalten und mono(ADP-ribos)ylierte oder poly(ADP- ribos)ylierte Proteine gebildet. Aufgrund dieser Komplexität und der Dynamiken stellt die Visualisierung von Poly(ADP-Ribos)ylierung nicht nur eine herausfordernde Aufgabe dar, sondern ist von enormer Wichtigkeit, um diese Prozesse auf zellulärer Ebene verstehen zu können.
Das Ziel dieser Doktorarbeit bestand darin, die Anwendung von chemisch modifizierten NAD+ Analoga für die Detektion von Poly(ADP-Ribos)ylierung in Folge von DNA-Schädigung zu untersuchen. Ein geeignetes Analogon ist ein effizientes Substrat von ADP- Ribosyltransferasen sowohl in vitro als auch in cellula. Es sollte die spezifische Detektion der zellulären Bildung von Poly(ADP-Ribose) in Abhängigkeit von externen Stimuli ermöglichen.
Idealerweise verhilft das neue Analogon dazu, Poly(ADP-Ribos)ylierung in Echtzeit und dynamisch verfolgen zu können.
Die Aufgabe wurde bewältigt, indem NAD+ Analoga synthetisiert wurden, welche in Poly(ADP-Ribose) durch ARTD1 eingebaut werden. ARTD1 ist das hauptsächlich für Poly(ADP-Ribos)ylierung verantwortliche Enzym während der DNA-Reparatur. Daher wurden systematisch sowohl die Positionen als auch die Art der Modifikationen am NAD+ untersucht. Dabei wurden Analoga, die in Position 2 des Adenins substituiert sind, als bestgeeignetste Verbindungen identifiziert. Mittels bioorthogonalen Reportergruppen konnte die intrazelluläre Detektion von Poly(ADP-Ribose) simultan und in zwei Farben erreicht werden, wie es beispielsweise für zeitabhängige Experimente oder Pulse-Chase- Anwendungen erforderlich ist. Weiterhin wurde ein fluorophormodifiziertes NAD+ entwickelt, welches das direkte Beobachten von Poly(ADP-Ribos)ylierung in lebenden Zellen ermöglicht. Dabei wurde ein vollständiger Poly(ADP-Ribos)ylierungszyklus nach laser- induzierter DNA-Schädigung in Echtzeit verfolgt. Zusätzlich konnten proteinspezifische Interaktionen von Poly(ADP-Ribose) in intakten Zellen mittels einer leistungsfähigen FLIM- FRET-Technik und dem jeweiligen GFP-markierten Protein detektiert werden. Zuletzt wurde das Substratspektrum anderer Poly(ADP-Ribose) synthetisierender Enzyme wie ARTD2, ARTD5 und ARTD6 untersucht, um die Anwendbarkeit der entwickelten NAD+ Analoga zu erweitern.
Die hier vorgestellten chemisch-biologischen Lösungsansätze erwiesen sich als leistungsfähig und die neu entwickelten Hilfsmittel werden bei der Entschlüsselung der Biologie von Poly(ADP-Ribose) hilfreich sein, indem sie die Untersuchung des Polymers in seiner natürlichen Umgebung ermöglichen.
Table of Content
Acknowledgement ... 2
Abstract ... 3
Zusammenfassung ... 4
Table of Content ... 5
List of Figures ... 7
List of Tables ... 8
List of Schemes ... 8
Introduction ... 11
1 Theoretical Background ... 11
1.1 NAD+ and its Metabolites ... 11
1.2 Poly(ADP-Ribose) and its Metabolism ... 13
1.3 PAR Synthesising Enzymes and their Function ... 16
1.4 Bioorthogonal Chemistry ... 23
1.5 Fluorescence Microscopy ... 27
2 Project Task - Towards the Metabolic Labelling of PAR ... 31
Results and Discussion ... 33
3 Design of NAD+ Analogues ... 33
4 Bioorthogonal Reporter-Modified NAD+ Analogues ... 36
4.1 Synthesis of Modified NAD+ Analogues 5-8 ... 36
4.2 Investigation of Labelling Conditions ... 37
4.3 Synthesis of Dye Conjugates ... 40
4.4 Biochemical Evaluation of NAD+ Analogues 5-8 ... 42
4.5 Synthesis and Investigation of Improved NAD+ Analogues 9-10 ... 44
4.6 Visualisation of Poly(ADP-Ribos)ylation in Cells ... 47
5 Dye-Modified NAD+ Analogues ... 51
5.1 Syntheses of NAD+ Analogues 11-12 ... 51
5.2 Attempted Syntheses ... 53
5.3 Biochemical Evaluation of NAD+ Analogues 11-12 ... 54
5.4 Towards A Cellular Application ... 56
5.5 Live-Cell Monitoring of Poly(ADP-Ribos)ylation ... 58
5.6 Fluorescence Lifetime Imaging of Protein-Specific PARylation in Cells ... 60
6 Substrate Scope of PAR Synthesising Enzymes ... 64
6.1 Syntheses of Alkyne-Modified NAD+ Analogues 1-4 ... 64
6.2 Biochemical Evaluation of NAD+ Analogues 1-4 ... 65
6.3 Expanding towards Dye-Modified NAD+ Analogues 11 and 12 ... 68
7 Summary and Discussion ... 71
8 Outlook and Future Research Directions ... 73
Experimental Part ... 77
9 Chemical Synthesis ... 77
9.1 General Experimental Details ... 77
9.2 Analytical Methods and Instrumentation ... 78
9.3 Experimental Procedures ... 81
10 Biochemical Methods ... 127
10.1 Buffers ... 127
10.2 Material ... 128
10.3 Biochemical Assays ... 129
11 Cellular Methods ... 133
11.1 Cell Culture ... 133
11.2 Assays for Reporter-Modified NAD+ Analogues ... 133
11.3 Methods for Dye-Modified NAD+ Analogues ... 134
List of References ... 139
Annexes ... 147
12 Abbreviations ... 147
13 NMR Spectra ... 151
13.1 Modified Nucleosides ... 151
13.2 Modified Adenosine Monophosphates ... 164
13.3 Modified Nicotinamide Adenine Dinucleotides ... 179
13.4 Cyclooctynes and Tetrazines ... 189
13.5 Dyes... 197
13.6 Dye Conjugates ... 204
14 HPLC Chromatograms of Applied NAD+ Analogues ... 216
15 Licenses... 217
List of Figures
Figure 1. Structural highlights of NAD+. ... 11
Figure 2. Recognition sites of PAR for protein modules. ... 15
Figure 3. Structure of human ARTD1. (A) Schematic representation of human ARTD1 domains. ... 18
Figure 4. Schematic representations of PAR synthesising ARTDs. ... 20
Figure 5. Copper(I)-catalysed azide-alkyne cycloaddition (CuAAC). ... 24
Figure 6. Strain promoted azide-alkyne cycloaddition (SPAAC). ... 25
Figure 7. Tetrazine ligation (DAinv). ... 26
Figure 8. Jablonski scheme of photophysical processes absorption, non-radiative decays, fluorescence and FRET and exponential decay of fluorescence with and without FRET. ... 28
Figure 9. Scheme explaining the calculation of fluorescent lifetimes acquired in the frequency domain. ... 30
Figure 10. General considerations for the design of NAD+ analogues and structures of known derivatives. ... 33
Figure 11. Structures of natural and modified NADs 1-12 prepared and used in this thesis. ... 34
Figure 12. Structures of the selected dyes 32-34. ... 40
Figure 13. Workflow of the ADP-ribosylation assay with histone H1.2 as acceptor. ... 42
Figure 14. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 using NAD+ analogues 5-8. ... 43
Figure 15. SDS PAGE and western blot analysis of ADP-ribosylation of histone H1.2 with ARTD1 using NAD+ analogues 5-8. ... 44
Figure 16. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 using NAD+ analogues 9-10. ... 46
Figure 17. Workflow of the degradation assay with PARG. ... 47
Figure 18. SDS PAGE analysis of time-dependent degradation of ADP-ribosylated histone H1.2 with PARG using a 1:1 mixture of NAD+ and compound 9. ... 47
Figure 19. Visualisation of intracellular PAR formation using NAD+ analogues 1, 9 and 10. ... 48
Figure 20. Confocal microscopy images of unsuccessful cytochemical assay using NAD+ analogues 5 and 6. ... 49
Figure 21. Dual-labelling of PAR using 9 and 10 (1:1) with SPAAC and DAinv chemistry. 50 Figure 22. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 (A) and ARTD1 automodification (B) using NAD+ analogues 11-12. ... 55
Figure 23. SDS PAGE analysis of ADP-ribosylation of histone H1.2 with ARTD1 using NAD+ analogue 11 and natural NAD+... 56
Figure 24. Comparing methods for the uptake of NAD+ analogue 11 into HeLa cells. ... 57 Figure 25. Flow cytometric analysis of uptake efficiency and cell viability after treatment with NAD+ analogue 11. ... 58 Figure 26. Live-Cell Monitoring of PARylation upon DNA damage by microirradiation using NAD+ analogue 11. ... 59 Figure 27. NAD+ analogue 12 is not recruited to DNA damage sites. ... 60 Figure 28. SDS PAGE analysis comparing wild type ARTD1 (wt) with eGFP variant in ADP-ribosylation of histone H1.2 (A) and automodification of ARTD1 (B).. ... 61 Figure 29. Fluorescence lifetime-based read-out of protein-specific PARylation. ... 62 Figure 30. Enrichment of eGFP-fusion proteins and TMR fluorescence signals at sites of microirradiation. ... 63 Figure 31. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 (A), ARTD2 (B), ARTD5 (C) and ARTD6 (D) using NAD+ analogues 1-4. ... 66 Figure 32. SDS PAGE analyses of auto(ADP-ribos)ylation of ARTD1 (A), ARTD2 (B), ARTD5 (C) and ARTD6 (D) using NAD+ analogues 1-4. ... 67 Figure 33. SDS PAGE analyses of ADP-ribosylation of histone H1.2 with ARTD1 (A), ARTD2 (B), ARTD5 (C) and ARTD6 (D) using NAD+ analogues 11 and 12. ... 69 Figure 34. SDS PAGE analyses of auto(ADP-ribos)ylation of ARTD1 (A), ARTD2 (B), ARTD5 (C) and ARTD6 (D) using NAD+ analogues 11 and 12. ... 70 Figure 35. Chemical strategies to render NAD+ cell-permeable. ... 74 Figure 36. Possible NAD+ analogues to improve pull-down probes. ... 75
List of Tables
Table 1. Overview of ARTD family members. ... 17 Table 2. Overview of selected cycloocytnes and tetrazines used. ... 38 Table 3. Overview of reactions between azide-modified AMPs and cyclooctyne using SPAAC chemistry and between alkene-modified AMPs and tetrazines using DAinv
chemistry. ... 39 Table 4. Overview of synthesised dye-conjugates 38-49. ... 41
List of Schemes
Scheme 1. NAD+ and its metabolites... 12 Scheme 2. Nucleophilic attacks of amino acid side chains onto NAD+ and their MARylated products. ... 13 Scheme 3. Metabolism of PAR. Arrows indicate sites of enzymatic attack. ... 14 Scheme 4. General synthesis route for the preparation of modified NADs. ... 35
Scheme 5. Syntheses of 2-modified adenosines 14 and 16. ... 36
Scheme 6. Syntheses of 6-modified adenosines 18 und 19. ... 37
Scheme 7. Syntheses of modified NADs 5-8. ... 37
Scheme 8. Synthetic routes towards tetramethylrhodamine (TMR, 33). ... 41
Scheme 9. Syntheses of dye-conjugates 38-49. ... 41
Scheme 10. Structures of improved reporter groups and fluorescent dyes applied in this chapter. ... 44
Scheme 11. Synthesis of improved NAD+ analogues 9 and 10. ... 45
Scheme 12. Synthesis of TMR-NHS ester (63). ... 51
Scheme 13. Synthesis of NAD+ analogue 11. ... 52
Scheme 14. Synthesis of NAD+ analogue 12. ... 52
Scheme 15. Attempted synthesis of BODIPY®-modified NAD+ analogues. ... 53
Scheme 16. Synthesis of NAD+ analogues 2 and 4. ... 64
Scheme 17. Possible NAD+ analogue 83 and more flexible synthesis strategy. ... 73
Theoretical Background
Introduction
1 Theoretical Background
In this chapter, NAD+ and poly(ADP-ribos)ylation is put into a biological context. Therefore, the structure, function and turnover of NAD+ and poly(ADP-ribose) is explained. In addition, the structures and biological relevance of poly(ADP-ribose) synthesising enzymes are introduced and current methods for the detection of poly(ADP-ribos)ylation are discussed.
Last, the central methods of this project, namely bioorthogonal chemistry and fluorescence microscopy, are presented.
1.1 NAD
+and its Metabolites
Nicotinamide adenine dinucleotide (NAD+, Figure 1) consists of the two mononucleotides adenine monophosphate and nicotinamide monophosphate, which are linked via a phosphate anhydride bond between the two phosphate groups. Both nucleotides furthermore contain a ribose sugar, a phosphate group linked to the 5-OH of the ribose, as well as the purine base adenine or the pyridine nicotinamide, which are both connected via N-glycosidic bonds to the C1-atom of the riboses. The pyrophosphate bond and the N-glycosidic bond between nicotinamide and the ribose are of high energy. Due to the two phosphate groups, the overall negative charge of the molecule is -1.
Figure 1. Structural highlights of NAD+.
In organisms, NAD+ can be either synthesised de novo from quinolinic acid, a degradation product of tryptophan, and the vitamin niacin (nicotinic acid), or via salvage pathways involving nicotinamide, nicotinamide riboside or nicotinic acid riboside, that cycle them back into an active form.[1-2]
NAD+ holds a key position in energy metabolism due to its involvement in cellular redox reactions.[1] It functions as a cofactor and shuttles between the oxidised form NAD+ and its reduced counterpart NADH, while transferring two electrons. Apart from redox functions, NAD+ was found to be used as substrate and consumed by different enzymes that are involved in cell signalling and posttranslational protein modification.[1-2] Thus, NAD+ is now seen as a versatile biomolecule. For non-redox-related roles, enzymes make use of NAD’s
intrinsic structural features and break its highly energetic bonds[3] by the attack of different nucleophiles resulting in a variety of NAD+ metabolites (Scheme 1).
For instance, NAD+-dependent DNA ligases transfer adenosine monophosphates to the 5’- end of nicked DNA by breaking the pyrophosphate bond and thus ligate the nicked DNA ends by AMP release.[3] Moreover, several enzymes were also found that attack the N- glycosidic bond with different nucleophiles, thereby releasing nicotinamide and forming several ADP-ribose-containing molecules.
Scheme 1. NAD+ and its metabolites.
For example, ADP-ribosyl cyclases[4] generate cyclic ADP-ribose (cADPr) from NAD+ and cADPr was found to be a potent second messenger that triggers Ca2+ release from internal Ca2+ stores.[2] Instead, ADP-ribosyltransferases mediate three different types of protein posttranslational modification: protein deacetylation, mono(ADP-ribos)ylation and poly(ADP- ribos)ylation.[3] During protein deacetylation, acetylated lysine residues are converted into free lysines and nicotinamide and 2’-acetyl-ADP-ribose (OAc-ADPr) are formed. These NAD+-dependent deacetylation reactions are catalysed by the enzyme family of sirtuins[5] and were originally discovered as histone deacetylases that mediate transcriptional silencing and longevity. The complex and unique posttranslational modification called ADP-ribosylation is in the focus of this project due to its involvement in many essential cellular processes including: DNA repair, apoptosis, transcription, cell cycle progression, energy metabolism and many others.[3, 6]
Theoretical Background
1.2 Poly(ADP-Ribose) and its Metabolism
1.2.1 ADP-Ribosylation Reactions
ADP-ribosylation refers to the covalent addition of single or multiple ADP-riboses from NAD+ onto side chains of different acceptor proteins. The process was first discovered in bacterial toxins such as Corynebacterium diphtheriae toxin, which inactivates the translation elongation factor-2 by ADP-ribosylation.[6] By now, ADP-ribosylation has been identified as an important posttranslational protein modification in all eukaryotes and prokaryotes.[7] The process is catalysed by so-called ADP-ribosyltransferases (ARTs), which consist of two enzyme families:[8] the ADP-ribosyltransferases with clostridial toxin homology (ARTCs), which comprise five members in human and act as ectoenzymes outside of cells; and the ADP-ribosyltransferases with diphtheria toxin homology (ARDTs) with 18 human family members identified so far and being active inside of cells. Moreover, some sirtuin deacetylases have also been found to catalyse ADP-ribosylation.[9]
In a typical reaction, NAD+ is attacked by nucleophilic side chains of substrate proteins and ADP-ribose is covalently attached leading to mono(ADP-ribos)ylation (MARylation). A number of amino acid residues have been identified to serve as acceptors,[7] but the most common target residues were found to be arginine, glutamic or aspartic acid and lysine.[10]
Scheme 2. Nucleophilic attacks of amino acid side chains onto NAD+ and their MARylated products.
Addition on arginine residues (Arg) results in the formation of an N-glycoside. Glutamic or aspartic acid (Glu, Asp) form ester linkages, which are believed to be stabilised by a 1’-2’- acyl-shift.[11] Addition on lysine residues (Lys) results in the formation of N-glycosides, that can be stabilised towards a ketamine bond via a Schiff-base equilibrium, also known as Amadori rearrangement (Scheme 2).
Scheme 3. Metabolism of PAR. Arrows indicate sites of enzymatic attack. ADPr… ADP-ribose, ARTD… ADP- ribosyl transferases with diphtheria toxin homology, PARG… poly(ADP-ribose) glycohydrolases.
Out of the 18 family members, only some enzymes are known to use the initial ADP-ribose as a starting point for further addition of ADP-ribose molecules.[12] This process is called poly(ADP-ribos)ylation (PARylation) and is found in all eukaryotes and archaea, but not in yeast.[7] By transferring multiple units onto the 2’-OH group of the previous, adenine proximal ribose, a long and negatively charged biopolymer is formed termed poly(ADP-ribose) (PAR).
This polymer can be further modified by adding more ADP-ribose onto the 2’’-OH group of the previous unit. Branching occurs with an average frequency of one branch per 20 to 50 units of linear polymer.[13-14] This makes the biopolymer PAR the most elaborate metabolite[15]
of NAD+ comprising a highly charged diphosphate backbone as well as unique ɑ(1→2)-O- glycosidic connections between the ribose rings (Scheme 3).[8, 16] The polymer chain length is heterogeneous and can reach up to 200 to 400 ADP-ribose units in vitro,[17-18] whereas the average polymer lengths in nuclei is about 10 monomers.[19] The polymerisation process is regulated by the NAD+ concentration. It was found in vitro that auto(ADP-ribos)ylation of ARTD1 occurred around a concentration of 200 nM NAD+, whereas polymer-elongation was
Theoretical Background
found at 1 to 2 mM NAD+ and highly branched polymers were synthesised with 200 mM
NAD+.[7, 20]
Surprisingly, PAR was already discovered by Chambon et al.[21] in 1963, but its biological impact remained unclear. Recently PAR research has experienced a renaissance[22-23] and more and more insights into PAR’s importance as a posttranslational protein modification emerged.
1.2.2 Degradation of PAR
As most posttranslational modifications, PARylation is a tightly regulated process and PAR levels can fluctuate on a minute scale in response to external stimuli.[24-25] As a conclusion, the PAR metabolism includes also PAR degrading enzymes in addition to PAR-synthesising enzymes as ARTDs (Scheme 3). The major PAR degrading enzyme is poly(ADP-ribose) glycohydrolase (PARG),[26] which is able to cleave O-glycosidic PAR bonds from the attached and free polymer. Three PARG isoforms have been described, but the role and localisation of these isoforms are not yet completely understood.[6] Other PAR-degrading enzymes have been identified and characterised. These include the ARH (ADP-ribosyl hydrolase) and NUDIX (nucleoside diphosphate linked to another moiety X) families of proteins.[2] All in all, the turnover of PAR polymers by hydrolases enables temporal control over the recruitment and release of PAR-binding proteins from cellular locations, as well as transient signalling and the formation of transient sub-organellar structures in the cytoplasm and nucleus.[16]
1.2.3 Binding of PAR
Resulting from the highly dynamic and complex PARylation process, the structure and function of involved proteins is affected and regulated. The number of proteins that are modified by ARTDs is steadily increasing due to recent improvements in proteomic studies.[27] Apart from the effects mediated by covalent attachment of ADP-ribose chains, the PAR polymer is believed to interact non-covalently with over 500 proteins[28] and PAR protein binding was shown to be mediated by at least four distinct protein modules (Figure 2). [12, 16]
Figure 2. Recognition sites of PAR for protein modules.[16] PBZ… PAR-binding zinc-finger.
(1) The PAR-binding motifs (PBMs) comprise a consensus sequence of eight amino acid residues: [HKR]1-X2-X3-[AIQVY]4-[KR]5-[KR]6-[AILV]7-[FILPV]8. It binds to long and branched PAR, most probably due to the electrostatic attraction between the negatively charged polymer and the positively charged lysine and arginine residues.
(2) The PAR-binding zinc-finger (PBZ) is a 3.5 kDa domain containing a zinc-coordinated fold that promotes stacking interactions with either one or two adenine moieties of a poly(ADP-ribose) unit, and by the unique α(1→2)-O-glycosidic bond that occurs at ADP- ribose-ADP-ribose junctions.
(3) The macrodomain fold is an evolutionary conserved motif of 130 to 190 amino acids that is known to bind towards terminal ADP-ribose. It is rapidly recruited to PARylation sites and found in several proteins including the histone variant macroH2A.[29-30]
(4) The WWE domain specifically recognises the iso-ADP-ribose moiety through the conserved amino acid sequence Trp-Trp-Glu.
1.3 PAR Synthesising Enzymes and their Function
For a long time, ARTD1 was believed to be the be the only PARylating enzyme. The founding member accounts for 75-90% of cellular PAR formation upon genotoxic stress and has therefore been most extensively studied.[14] It was formerly known as poly(ADP-ribose) polymerase 1 (PARP1) and the term ‘PARP’ for these enzymes is still frequently used in the research literature.[8]
With the help of structural and computational analysis, the other ARTD family members have been identified and can be subdivided into four subfamilies on the basis of their domain structures (Table 1):[24-25] the DNA-dependent ARTDs, the tankyrases, the CCCH ARTDs, the macro-ARTDs, and some members with distinct domain structures, that do not fit into one of these classes. DNA-dependent ARTDs exhibit N-terminal DNA binding domains and their activity is regulated by binding to discontinuous DNA structures. Tankyrases comprises large ankyrin domain repeats mediating protein-protein-interactions. The CCCH ARTDs can bind to RNA with the help of their Cys-Cys-Cys-His containing zinc-fingers and macro-ARTDs contain additional macrodomain folds that help to localise them at sites of PARylation. Not all ARTD family members were found to catalyse PARylation. Indeed, only ARTD1, ARTD2, ARTD5 and ARTD6 were proven to form polymers, whereas the others exhibit MARylation activity or are either catalytically inactive or require further, yet unknown factors to trigger ADP-ribosylation. Derived from structural and computational data, ARTD3 and ARTD4 are predicted to form polymers as well, but this activity has not been confirmed yet.[12, 16]
ARTDs are involved in the regulation of many cellular processes.[6, 16, 31] By binding to DNA strand breaks, ARTDs are involved in DNA repair and recruit different DNA repair factors. As most DNA breaks occur due to genotoxic stress, ARTDs are essential for cell survival and genome maintenance. Through the modification of histones and binding to other nucleosomal proteins as well as by the modulation of RNA, ARTDs regulate chromatin compaction and have severe impact on gene expression.[31] Moreover, ARTD activity has also been found at centrosomes, microtubules and telomeres, which links PARylation with cell division and aging.
In contrast to these cytoprotective functions, ARTD overactivity can lead to pathophysiological effects, up to the point of cell death by depletion of NAD+ and ATP pools.[32] These effects are linked to a variety of disease state as found in oxidative stress, inflammatory, and metabolic diseases, neurodegenerative disorders, and heart failure.[33]
Moreover, ARTD1 expression is upregulated in various types of human cancers, and this has been linked to resistance of cancer cells to genotoxic treatment.[34-35] Thus, targeting tumour suppressor mechanisms such as DNA repair is an attractive approach in cancer therapy to sensitise tumour cells and to overcome acquired resistance. Some cancers that have mutations in the BRCA1 gene (BReast CAncer 1) rely exclusively on ARTD1-mediated DNA
Theoretical Background
repair and thus inhibiting ARTD1 can kill these cancer cells selectively. In 2014, olaparib was approved as first ARTD1 inhibitor for the monotherapeutic treatment of BRCA-mutated ovarian cancers.[33]
Because PAR synthesising enzymes are in the focus of this project, the DNA-dependent ARTD1 and ARTD2 as well as the tankyrases ARTD5 and ARTD6 are discussed in more detail.
Table 1. Overview of ARTD family members.[12, 16, 31]
Name Alternative Name Subclass Size
(aa) Subcellular
Localisation Enzymatic
Activity Structural Motifs
ARTD1 PARP1 DNA-
dependent 1014 Nuclear PARylation,
branching
WGR, zinc- fingers,
BRCT
ARTD2 PARP2 DNA-
dependent 570 Nuclear >> Cytosolic PARylation WGR
ARTD3 PARP3 DNA-
dependent 540 Nuclear > Cytosolic MARylation WGR
ARTD4 PARP4, vPARP 1724 Cytosolic (vault
particles) > Nuclear MARylation BRCT ARTD5 Tankyrase 1,
TNKS1, PARP5a Tankyrase 1327 Cytosolic >> Nuclear PARylation, OARylation
ankyrin repeat ARTD6
Tankyrase 2, TNKS2, PARP5b,
PARP6 Tankyrase 1166 Cytosolic >> Nuclear PARylation, OARylation
ankyrin repeat
ARTD7 PARP15, BAL3 macro-
ARTD 444 Cytosolic (stress
granules) MARylation Macro-
domain ARTD8 PARP14, BAL2,
CoaSt6 macro-
ARTD 1801 Cytosolic (stress
granules) > Nuclear MARylation
Macro- domain,
WWE
ARTD9 PARP9, BAL1 macro-
ARTD 854 Cytosolic >> Nuclear no activity
reported Macro- domain
ARTD10 PARP10 1025 Cytosolic >> Nuclear MARylation
ARTD11 PARP11 331 Nuclear and
Cytosolic MARylation WWE
ARTD12 PARP12, ZC3HDC1 CCCH-
ARTD 701 Cytosolic (stress
granules) >> Nuclear MARylation zinc-fingers, WWE ARTD13 PARP13, ZC3HAV1,
ZAP1
CCCH-
ARTD 902 Cytosolic (stress granules)
no activity reported
zinc-fingers, WWE
ARTD14 PARP7,
tiPARP, RM1
CCCH-
ARTD 657 Cytosolic and
Nuclear MARylation zinc-fingers, WWE
ARTD15 PARP16 630 Cytosolic MARylation
ARTD16 PARP8 854 Cytosolic MARylation
ARTD17 PARP6 322 Cytosolic MARylation
ARTD18 TPT1 unknown no activity
reported
1.3.1 ARTD1
The founding member is a nuclear, DNA-dependent ARTD and it was shown that ARTD1 activity is increased up to 500-fold[36] upon binding to short single-stranded DNA, thereby PARylating itself as main target. The process, where ARTD1 serves as its own acceptor, is called automodification.[37] Other important target proteins include tumour suppressor p53 and several histone proteins.[38] ARTD1 fulfils manifold functions inside of cells, most prominently in DNA repair.
ARTD1 is a 113-kDa-protein that displays a characteristic three-domain structure (Figure 3).[24-25, 39] The N-terminal DNA-binding domain consists of two homologous zinc- finger domains (Zn1, Zn2) and a third, distinct zinc-finger (Zn3). Crystal and NMR structures unravelled that Zn1 is important for binding and recognition of single- and double-stranded DNA lesions, while Zn2 is important for binding of single strand breaks. Zn3 plays a role for ARTD1 activation and interdomain communication. In addition, the DNA-binding domain comprises a nuclear localisation signal (NLS) and a caspase three cleavage site (Cas).[25]
The second, so-called automodification domain contains a BRCT fold. The BRCT protein interaction motif (BRCA1 C-terminus), which is found in several proteins that regulate cell- cycle checkpoints and DNA repair, is rich in lysine-residues that can be auto(ADP- ribos)ylated.
Figure 3. Structure of human ARTD1. (A) Schematic representation of human ARTD1 domains.[39] (B) Crystal and/or NMR structures of the ARTD1 domains in the absence of DNA. Shown are the NMR structures of Zn1 and Zn2 domains (PDB code 2dmj and 2cs2), the NMR structure of the Zn3 domain (PDB code 2jvn), the NMR structure of the BRCT fold (PDB code 2cok), the NMR structure of the WGR domain (PDB code 2cr9), and the crystal structure of the catalytic domain (PDB code 1a26).
Reproduced with permission.[39] (C) A model of the approximate positioning of all domains.
Reproduced with permission.[39] Structure depictions can be made using Pymol (www.pymol.org).
NLS… nuclear localization signal, Cas… Caspase cleavage site, Zn… zinc-finger, BRCT… BRCA1 C-terminus interaction motif, WGR… Trp-Gly-Arg domain, HD… helical subdomain, ART… ADP- ribosyl-transferase subdomain. Copyright © 2013 Published by Elsevier Ltd.
Theoretical Background
A WGR domain is adjacent to the catalytic domain and comprises a well-conserved region of the amino acid sequence Trp-Gly-Arg responsible for interdomain contacts.[24]
The C-terminal catalytic domain is composed of two subdomains, the helical subdomain (HD) and the ADP-ribosyl-transferase subdomain (ART). The HD consists of six ɑ-helices with connecting linkers that have regulating functions. The ART domain comprising a β-α- loop-β-α fold is the most conserved region and shares a high level of homology with bacterial toxins such as the diphtheria toxin. The ART domain is required for the binding, positioning and processing of NAD+.[24, 39]
In the absence of DNA damage, ARTD1 is believed to exist in an extended ‘beads-on-a- string’ architecture, where the catalytic domain exhibits a rigid conformation and a basal ARTD activity. Upon binding to DNA, interdomain contacts are formed, which allow the catalytic domain to become more flexible and thus being more efficient in producing PAR chains.[39] The more ADP-ribose units are added onto ARTD1, the higher the negative charge becomes. At one point, the negative charge becomes repulsive and ARTD1 and DNA dissociate from each other. This allows other repair proteins to access the DNA damage, while ARTD1 is rapidly de-PARylated by PARG and being available for the next DNA- damage induced PARylation cycles.[38]
1.3.2 ARTD2
ARTD2[40] is the family member, which is most similar to ARTD1, because they share extensive structural similarity in the catalytic domain.[41] ARTD1 and ARTD2 are known to form heterodimers, share several common nuclear binding partners and display some redundant functions in DNA repair and maintenance of genomic stability.[6] This redundancy is supported by knockout experiments in mice, where single knock-out mice are viable, but very sensitive towards genotoxic stress, while the double gene knockout is lethal in an embryonic state.[42]
Nevertheless, some evidence occurred that ARTD2 may fulfil some more specialised tasks.
For instance, it was shown that ARTD2 is selectively activated by 5’-phosphorylated DNA strand breaks and it is suggested that its activity is stimulated in response to more specific DNA repair intermediates or at particular repair stages.[43] From a structural point of view, the domain architecture outside the catalytic domain and the WGR domain differs substantially from ARTD1 (Figure 4). Instead of a complex DNA binding domain, ARTD2 contains only a short N-terminal region (NTR). The NTR comprises a nucleolar localisation sequence (NoLS) and a putative nuclear localisation signal (NLS) and was found to be involved in protein- protein interactions.[43] Previous reports[40] identified the NTR as an DNA-binding domain, but newer reports suggest that the NTR has rather contributing functions.[43] The NTR was found to be unstructured and particularly important for the binding of and activation on single stranded DNA strand breaks, whereas the WGR and catalytic domains concurrently mediate the binding to DNA damage sites.[43]
1.3.3 Tankyrases ARTD5 and ARTD6
ARTD5[44] and ARTD6[45-46], which are functionally and structurally similar, are also termed tankyrase 1 and 2. The name is derived from TRF1-interacting ankyrin-related ADP-ribose polymerase, because ARTD5 was first discovered as a regulator of telomeric DNA through its interaction with and modification of telomere repeat-binding factor 1 (TRF1).[6, 24] In contrast to ARTD1 and ARTD2, tankyrases were found to be mainly expressed in the
cytosol. Most protein interaction partners are shared by both tankyrases, which suggest that they have similar functions. Using siRNA knockdowns, it was shown that ARTD5 has essential regulatory function in mitotic segregation, whereas ARTD6 knockout mice exhibited normal telomeres, but a perturbed metabolism due to the smaller size of mice.[47] Some reports associated tankyrase-mediated PARylation activity with ubiquitin ligase activity. This suggests a link towards regulation of protein turnover and signalling pathways. A prominent target is Axin1, which is involved in Wnt signalling and thus being of high interest for cancer research.[24]
The catalytic domain of the tankyrases is located at the C-terminus. In contrast to ARTD1 and ARTD2, the tankyrases do not exhibit an ɑ-helical regulatory subdomain. Typical for the tankyrases are a series of structural modules, the ankyrin repeats (ANK), through which protein interaction is mediated. ARTD5 contains 24 of these repeats, whereas ARTD6 consists of 16 modules. Each ankyrin repeat comprises of a loop-helix-loop-helix-loop structure that is similar to the human protein ankyrin. A sterile-alpha motif (SAM) domain separates the N-terminal ankyrin repeat region and the C-terminal catalytic region of the tankyrases. Its function has not yet been determined, but it is suggested to serve as multimerisation domain. ARTD5 has an additional N-terminal region, which is missing in ARTD6. This region with presumably regulatory function is termed HPS domain, due to its low sequence complexity consisting of repeats of His, Pro and Ser.[6, 24]
Figure 4. Schematic representations of PAR synthesising ARTDs.[6, 39] NLS… nuclear localization signal, Cas… Caspase cleavage site, Zn… zinc-finger, BRCT… BRCA1 C-terminus interaction motif, WGR… Trp-Gly-Arg domain, HD… helical subdomain, ART… ADP-ribosyl-transferase subdomain, NoLS… nucleolar localisation sequence, NTR… N-terminal region, HPS… His, Pro and Ser rich domain, ANK… ankyrin, SAM… sterile-alpha motif.
Theoretical Background
1.3.4 Detection of PARylation
To understand the complex and functional processes of PARylation, it is crucial for scientists to develop tools that can detect and quantify this posttranslational modification.
However, this task remains challenging due to the following reasons.
First, PAR was found to be attached on a wide range of protein sites, such as basic and acidic protein side chains. This variability creates different chemical and enzymatic reactivity and stability of the covalent bonds formed. Furthermore, the polymer itself is rather labile due to the diphosphate backbone, which consists of highly energetic anhydride bonds and is prone to nucleophilic attacks. Moreover, the high negative charge makes the handling and purification of PAR and PARylated proteins challenging. Additionally, the polymer is very heterogeneous and consists of differently sized, linear or branched chains. Last but not least, PARylation underlies a dynamic equilibrium, where chains are attached, elongated or removed with fast kinetics and making it hard to ‘catch’ the polymer.[27]
Remarkable efforts have been undertaken to develop tools and assays for studying PARylation on a molecular level and within its natural environment. Some of these approaches are presented in the following.
One of the current standard methods for the detection of PARylation is the use of 32P- radioactively labelled NAD+. The β-decay signal obtained from isolated PAR is then proportional to the number of incorporated, labelled ADP-riboses. This can be used to determine both ARTD and PARG[48] activity and allows for the in vitro screening of enzyme inhibitors. Although successfully established, special training in dealing with radioactivity is needed and cellular applications are hampered by the lack of a spatially resolved read-out.
Another breakthrough was the development of PAR-specific antibodies by Kawamitsu et al.[49] in 1984 and the establishment of immunocytochemical protocols.[50] Detecting PAR with the help of the 10H-anti-PAR antibody and coupling to other read-outs such as chemical luminescence or fluorescence has evolved as the second standard method in PAR research.
However, this method provides only indirect measures of PARylation and might also give false-positive signals due to unspecific binding.
Moreover, several other methods have been developed to measure PARylation. Besides radioactive labels, the use of ε-NAD+ as a fluorescent analogue has also been investigated.[51] Although it was metabolised by ARTDs, fluorescence quenching was observed after incorporation into the polymer.[52] As a result, formed polymers have to be either digested in order to free the fluorescence signal or additionally immuno-stained with a specific antibody.[53] As this procedure does not offer any advantages over the standard methods, it is not applied.
In contrast to ε-NAD+, 6-biotin-NAD+ was shown to be sufficiently used as ARTD substrate and is now available in a mixture with natural NAD+ in many commercial ADP-ribosylation kits.[54-55] In addition, it was shown to be incorporated into PAR within oxidatively stressed cells.[56] Despite this breakthrough, it is not used by the PAR research community for intracellular PARylation studies. This might be due to the fact, that it bears a long linker and biotin is a rather bulky modification. Therefore, it might not be a frequently used ARTD substrate and decrease ARTD activity, when too high concentrations are applied.
Recently, a new generation of NAD+ analogues have been developed that are equipped with small, terminal alkyne-reporter groups.[57-59] These derivatives are well accepted substrates of ARTD1 and can be further modified with fluorophores and isolation tags via
copper(I)-catalysed click-reactions after incorporation. To the best of the author’s knowledge, cellular applications have not been reported yet.
Next to the development of NAD+ analogues, other approaches for measuring PARylation have been investigated. For instance, the consumption of NAD+ can be determined by either converting NAD+ chemically into a fluorescent product,[60] by applying NAD+ sensors[61] that fluoresce after binding, or by using an colorimetric substrate[62] that releases nitrophenol after incorporation.
Apart from the NAD+, the PAR polymer itself or digested ADP-riboses units can also be chemically modified. By taking advantage of the intrinsic aldehyde function of the ADP- ribose, a fluorescent analogue[63] can be formed or the aldehyde can react with hydrazine[64]- or aminooxy[11]-containing tags such as biotin or fluorophores. However, this reaction is not suitable to detect PAR attached to lysine residues and it will also label all other aldehyde functionalities being present within a biologic context such as abasic sites in DNA.
Apart from antibodies, other PAR and ADP-ribose sensors have been developed. For instance, smaller ADP-ribose binding macrodomains have been identified, which can also be applied for the detection of MARylation and free ADP-riboses.[29] Furthermore, a turn-on split- luciferase sensor was developed that assembles a functional luciferase enzyme upon binding to PAR.[65] Another approach uses a supercharged, green fluorescent protein as sensor for PARylation.[66] Here, the fluorescence of the protein is quenched by Förster resonance energy transfer (FRET), while being electrostatically bound to the PAR polymer.
Another powerful method to detect PARylation is mass spectrometry. Due to the fact, that samples get destroyed during measurement, mass spectrometric investigations rather seek to detect protein sites of PARylation, PARylation targets and interaction partners as well as changes in PARylation levels.[27] Here, scientists benefited from the application of NAD+ analogues in order to isolate PARylated proteins and to distinguish signal from background.
Recent progress has been achieved with the help of a chemical genetics approach, where orthogonal ARTD/NAD+ pairs are generated using a ‘bump-hole’-strategy.[67-70] Thus, only the mutated ARTD can use the applied NAD+ analogue as substrate and allows to read out the PARylation target proteins in an enzyme-specific manner. However, the chemical genetics approach is currently limited to in vitro applications and further analysis has to proof, if these approaches faithfully predict in vivo targets.[71]
Most of the presented detection methods can only be applied for the in vitro study of ADP- ribosylation, because they are either too unselective for biological samples or require non- physiological conditions. To date, only few methods exist for the intracellular, dynamic detection of PARylation. As already mentioned, indirect, steady-state cellular imaging can be achieved by immunostaining with PAR antibodies.[50] Moreover, dynamic studies of PARylation processes was achieved by tagging PAR synthesizing or PAR binding enzymes with fluorescent proteins, such as eGFP-ARDT1[72] or eGFP-ARTD2[43] and eGFP- macroH2A,[30] which thus give an indirect measure of PARylation.
Still, no method offers the possibility to follow PARylation directly at the level of the polymer in real-time and dynamically upon extrinsic stimuli like DNA damage induction, but would be of high interest to understand PARylation on a cellular level.[73]
Theoretical Background
1.4 Bioorthogonal Chemistry
One of the main approaches used in this project is the application of bioorthogonal chemistry to react modified ADP-ribose units with their fluorophore-containing partners, which were previously introduced by reporter-tagged NAD+ molecules into PAR chains. Thus, the concept of bioorthogonality and the most commonly used reactions are described in this chapter.
1.4.1 General Remarks
Any chemical reaction that can occur inside of living systems without interfering with native biological processes is considered a bioorthogonal reaction.[74] That means that a reaction must proceed selectively between the reporter and its counterpart without the formation of side products and without reacting with other biological function present in its surrounding, such as free thiols. Moreover, the formed conjugation bond should be stable and chemically inert towards the attack of nearby biological nucleophiles. Due to the low concentration of the species of interest and possible quick turnover, these reactions need to have fast reaction kinetics that ideally do not rely on further additives. In addition, the reaction needs to proceed under physiological conditions (37 °C and pH 6-8) and all reaction partners and product should be active, stable and non-toxic under these conditions.[75]
Huge progress has been made to develop and apply bioorthogonal chemistry for the study of biological processes. Thus, different biomolecules have been labelled with suitable reporter groups, such as amino acids, carbohydrates, nucleosides and nucleotides, as well as lipids, and other small molecules. Moreover, the spectrum and scope of bioorthogonal reactions is steadily increased, covered by numerous reviews.[74-79]
In the following the most-commonly used reactions, namely copper(I)-catalysed azide- alkyne cycloadditions (CuAAC), strain promoted azide-alkyne cycloadditions (SPAAC) and tetrazine ligations (DAinv) are introduced.
1.4.2 CuI-Catalysed Azide-Alkyne Cycloaddition
In 1963, Huisgen initially reported the 1,3-dipolar cycloaddition between azides and alkynes.[80] In 2002, Sharpless[81] and Meldal[82] independently discovered, that the addition of catalytic amounts of copper(I) (CuI) salts results in a dramatic rate enhancement. These reactions proceed readily under physiological conditions and in a biological environment to provide 1,4-disubstituted triazoles. This reaction, widely known as ‘click reaction’, is the most popular bioorthogonal reaction and alkynes as well as azides can serve as bioorthogonal reporter groups due to their small size and easy synthetic accessibility (Figure 5A).[76]
In the current model[83] of the reaction mechanism, CuI forms first a π-complex with the triple bond of the alkyne and then a second CuI atom forms a σ-bound copper-acetylide under proton release. This intermediate now coordinates the azide and forms an unusual six- membered metallacycle. This ring is contracted to a triazolyl-copper-derivative and consecutive hydrolysis yields the regioselectively formed 1,4-disubstituted triazole (Figure 5B).
To further accelerate the reaction and to stabilise the +I oxidation state, several ligands such as water soluble tris(3-hydroxypropyltriazolyl methyl)-amine (THPTA) or chelate- assisting ligands such as 2-picolyl-azide have been applied (Figure 5C). Moreover, some protocols also use CuII salts that are reduced in situ by additives such as ascorbic acid.
Using these protocols, reaction rates with second-order rate constants of 10-200 M-1·s-1 in the presence of 20-500 µM CuI are achieved.[78] The primary disadvantage of the CuAAC reaction is the cellular toxicity of the metal catalyst. It was found that CuI forms reactive oxygen species (ROS) responsible for damage and degradation of its biological environment.
Moreover, dehydroascorbate was found to be responsible for protein crosslinking and resulting in their precipitation.[78] Although more bio-friendly metal/ligand combinations were discovered and individual optimisation of the respective reaction can reduce its harmfulness,[84] the reaction is not ideal for labelling biomolecules in living cells.
Figure 5. Copper(I)-catalysed azide-alkyne cycloaddition (CuAAC). (A) CuAAC of either alkyne-tagged reporter and azide-functionalised dye or azide-tagged reporter and alkyne-functionalised dye. (B) Current model of CuAAC mechanism.[83] (C) Structures of selected ligands and additives.
1.4.3 Strain Promoted Azide-Alkyne Cycloaddition
To circumvent the need of CuI and other additives, a new concept relying on rate enhancement of the azide-alkyne cycloaddition through ring-strain was introduced in 2004 by Bertozzi and co-workers and is now termed strain-promoted or copper-free click reaction (SPAAC).[85] Here, the terminal alkyne is replaced by the cyclooctyne OCT, which releases around 18 kJ·mol-1 of ring strain, while reacting with azides. However, computational analysis revealed that cyclooctynes resemble the transition state of the 1,3-dipolar cycloaddition more likely than terminal alkynes, which leads to a decrease in distortion energy. This distortion energy, rather than strain relief, is responsible for the fast coupling of these alkynes to azides.[86-87]
During this reaction, both triazole regioisomers are formed within a concerted [4+2]
cycloaddition (Figure 6A and B). Since this discovery, a number of cyclooctyne derivatives with different improvements were reported and second order rate constants range between 10-2 and 1 M-1·s-1 (Figure 6C).[77] Rate enhancement was achieved by either attaching electron withdrawing groups at the propargylic position (MFCO, DIFO) or by further increasing the ring strain through aryl rings (DIBO, DIBAC, BARAC) or a cyclopropyl ring (BCN).[78] The most recent improvement resulted in the currently fastest SPAAC reagent:
3,3,6,6-tetramethyl-thiacyloheptyne (TMTH). Here, the ring is contracted to a seven
