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“Clickable” chain terminator NAD + analogs for labeling substrate proteins
of poly(ADP-ribose) polymerases
Dissertation
zur Erlangung des akademischen Grades des Doktors der Naturwissenschaften
(Dr. rer. nat.)
an der Universität Konstanz
Mathematisch-Naturwissenschaftliche Sektion Fachbereich Chemie
vorgelegt von
Magdalena Grzywa
Konstanz, August 2014
Tag der mündlichen Prüfung: 28. Januar 2015 Prüfungsvorsitz: Prof. Dr. Gerhard Müller 1. Referent: Prof. Dr. Andreas Marx 2. Referent: Prof. Dr. Valentin Wittmann
Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-0-284407
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This work was carried out between November 2008 and March 2013 in the Laboratory of Organic Chemistry and Cellular Chemistry, Department of Chemistry, University of Konstanz, under the supervision of Prof. Dr. Andreas Marx.
I would like to express my gratitude to my Doktorvater Prof. Dr. Andreas Marx, for the great opportunity he gave me to work in his group on an extremely challenging and interesting project, for giving me the opportunity of independent working, for his advices and guidance throughout my research work. Thank you for trusting me; this experience has incredibly enriched me as a scientist as well as a person.
I acknowledge the members of my thesis committee, Prof. Dr. Walentin Wittmann and Prof. Dr. Thomas Mayer, for supporting this work with productive input, discussions and ideas.
I would like to warmly thank Dr. Karl-Heinz Jung for fruitful discussions, Dr. Andreas Marquardt and Dr. Anna Śladewska-Marquardt for advices and help with the mass spectrometric analysis and possibility for independent work, Anke Gerull for the re-syntheses of compounds for the ‘carbasugar’ project, Yan Wang for discussions regarding the project and Daniel Rösner for kindly providing purified histone H1.2 and histone H2B proteins, as well as Dr.
Daniel Schneider for kindly providing purified GST-p53 protein. I would also like to thank my diligent students for the patience of going through total synthesis during their work.
I thank Dr. Norman Hard for a wonderful time without conflicts we spent in the lab, lunch and coffee time together, time of laughing as well as support during harder days and Tobias Strittmatter for creating an unforgettable atmosphere in our lab. I am thankful to all present and past members and students who created “AG Marx”, for the unique atmosphere and many unforgettable moments.
I thank my family: the late grandparents – without them I would never be who I am and to my parents for understanding, support, motivation and patience, which was invaluable.
Special warm thanks go to my husband Maciek: without your love, continuous support and patience, this work would not have been possible.
I would also like to thank to all the awesome persons I have met in Konstanz: Dr. Marta Robotta, Dr. Anna Śladewska-Marquardt, Dr. Andreas Marquardt, Dr. Norman Hardt, Dr. Olga Gutierrez, Dr. Gabriela Paraschiv, Sergiu Rusu, Dr. Camelia Vlad, Stefan Slamnoiu for the unforgettable time we spent together.
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„To, co możesz uczynić, jest tylko maleńką kroplą w ogromie oceanu, ale właśnie jest tym, co nadaje znaczenie Twojemu życiu.”
„Alles was du tun kannst, wird in Anschauung dessen, was getan werden sollte, immer nur ein Tropfen statt eines Stromes sein; aber es gibt deinem Leben den einzigen Sinn, den es haben kann, und macht es wertvoll.“
Albert Schweitzer
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9 Poly(ADP-ribose) polymerases (PARPs) play an important role in many biological processes. PARPs catalyse the transfer of multiple adenine diphosphate ribose (ADP-ribose) units from nicotinamide adenine dinucleotide (NAD+) to PARP itself, histones, p53 and many other acceptor proteins, producing poly(ADP-ribose) chains. This is a unique post-translational modification and up to now, no reports of clickable chain terminator NAD+ analogues have been published. Clickable NAD+ analogues can react with either a fluorescent dye or biotin for further visualization and purification of parylated proteins. Such NAD+ analogues with additional function of being chain terminator will help in identify proteins interacting with PARP as well as the modification sites of these proteins. Previous studies showed that dideoxy NAD+ can be a chain terminator; however, the lack of an affinity tag makes this probe inapplicable for further studies. Recently, Hong Jiang et al. have reported modified NAD+ probes for “fishing” proteins that interact with PARP. However, this method is limited due to the hampered production of PAR. In this PhD thesis, the design and efficient syntheses of novel functionalised NAD+ bearing an alkyne group on ribose part of adenosine is presented. With these modifications it is possible to investigate their influence on the formation of PAR. Moreover, the alkyne group provides an excellent possibility to label these probes fluorescently by the help of Copper-Catalyzed Azide-Alkyne Cycloaddition (CuAAC) reaction (click chemistry) with azide dyes or couple affinity tags by clicking with biotin. Using this method, histone H1.2, histon H2B, and p53 itself were labelled by a fluorescent tag and visualised on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). These results show that clickable NAD+ analogues are useful for labelling and in-gel fluorescent detection of proteins and will help to understand the biological functions of PARP.
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Zusammenfassung
Poly(ADP-ribose) Polymerasen (PARPs) spielen in vielen biologischen Prozessen eine wichtige Rolle. PARPs katalysieren den Transfer von mehreren Adenosin- diphosphoriboseeinheiten des Nicotinamidadenindinukleotid (NAD+) zu PARP, Histonen, p53 und vielen anderen Akzeptorproteinen, wobei poly(ADP-ribose)-Ketten gebildet werden. Dies ist eine spezifische posttranslationale Modifizierung und bisher wurden keine Berichte über klickbare NAD+ Analoga, die zu einem Kettenabbruch führen, veröffentlicht.
Klickbare NAD+ Analoga können entweder mit einem Fluoreszenzfarbstoff oder Biotin reagieren, um sie entweder sichtbar zu machen oder um parylierte Proteine zu reinigen. Diese NAD+ Analoga mit zusätzlichen Funktionen als Kettenabbrecher werden dazu beitragen, sowohl Proteine, die mit PARP wechselwirken, als auch deren Modifizierungsstellen zu identifizieren. Frühere Untersuchungen zeigten, dass dideoxy NAD+ zu einem Kettenabbruch führen kann. Jedoch macht es das Fehlen eines Affinitätstags unmöglich, dieses Molekül für weitere Untersuchungen einzusetzen. Kürzlich berichteten Hong Jiang et al. über modifizierte NAD+ Sonden, um Proteine, die mit PARP interagieren, zu „fischen“. Jedoch ist diese Methode aufgrund der gehemmten Produktion von PAR nur begrenzt einsetzbar. In der vorliegenden Doktorarbeit wird der Entwurf und die effiziente Synthese neuartiger funktionalisierter NAD+ Analoga, die eine Alkingruppe an der Riboseeinheit des Adenosins tragen, vorgestellt. Mit diesen Modifizierungen ist es möglich, ihren Einfluss auf die Bildung von PAR zu untersuchen. Darüber hinaus bietet die Alkingruppe die Möglichkeit, die Sonden durch kupferkatalysierte azid-alkin Cycloaddition (CuAAC) mit Azidfarbstoffen zu markieren oder Affinitätstags wie Biotin zu koppeln. Mit Hilfe dieser Methode wurden Histon H1.2, Histon H2B und p53 mit einem Fluoreszenztag markiert und auf einem Natriumdodecylsulfat- Polyacrylamidgel (SDS-PAGE) sichtbar gemacht. Diese Resultate zeigen, dass klickbare NAD+ Analoga für Markierungen und die Fluoreszenzdetektion von Proteinen in Gelen nützlich sind und dazu beitragen können, die biologischen Funktionen von PARP aufzuklären.
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1. Introduction ... 13
1.1. From DNA discovery through DNA damage to PAR ... 13
1.2. Metabolism of NAD+ ... 16
1.3. Poly(ADP-Ribose) Metabolism ... 19
1.3.1. Structure of poly (ADP-ribose) PAR ... 23
1.3.2. Poly(ADP-ribose) Glycohydrolase (PARG) ... 24
1.3.3. ADP-ribosyl protein lyase ... 26
1.4. PARP-1 ... 27
1.4.1. Biological importance of PARP-1 ... 28
1.4.2. Structure of PARP-1 ... 33
1.4.2.1. The DNA-binding domain ... 33
1.4.2.2. The automodification domain ... 34
1.4.2.3. The catalytic domain ... 35
1.5. PARP superfamily ... 36
1.6. Detection and quantification of PAR and PARP activity ... 38
1.6.1. Chromatographic methods ... 39
1.6.2. Immunodetection of PAR ... 40
1.6.3. Quantification of Cellular Poly(ADP-ribosyl)ation by Stable Isotope Dilution Mass Spectrometry ... 41
1.6.4. Mass spectrometric characterization of ADP-ribose ... 42
2. Syntheses of clickable NAD+ analogs ... 44
2.1. Introduction... 44
2.1.1. Known modified NAD+ analogues ... 44
2.1.1.1. DeoxyNAD+ ... 44
2.1.1.2. Biotinylated NAD+ ... 45
2.1.1.3. ADP-ribose-pNP as colorimetric substrate for PARP ... 47
2.1.1.4. 6-alkyne-NAD and 8-alkyne-NAD ... 47
2.1.2. Copper-catalyzed azide-alkyne cycloaddition (CuAAC) – ‘Click reaction’ ... 48
2.2. Motivation ... 52
2.3. Results and discussion ... 55
2.3.1. Synthesis of NAD+ analogues ... 55
2.3.2. Biochemical evaluation of NAD+ analogues ... 61
2.3.3. Mass spectrometric analysis of modified proteins and peptides ………68
2.4. Conclusions and Outlook ... 72
3. Towards the synthesis of a carbasugar containing PAR ... 74
3.1. Introduction... 74
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3.1.1. Known modified adenosine on 2′OH group ... 74
3.1.2. Glycosylation methods... 75
3.1. Motivation ... 78
3.2. Results and discussion ... 79
3.2.1. Synthesis of carbasugar ... 79
3.2.2. Synthesis of artificial monomeric PAR ... 85
3.3. Conclusions and outlook ... 87
4. Materials and methods ... 88
4.1. General ... 88
4.1.1. Chemicals and solvents ... 88
4.1.2. General experimental details ... 88
4.2. Synthesis of NAD+ analogues ... 89
4.3. Biochemical experiments ... 98
4.3.1. One-dimensional gel electrophoresis ... 98
4.3.1.1. SDS-PAGE gel ... 99
4.3.1.2. Tricine SDS-PAGE according to Schägger ... 100
4.3.1.3. Colloidal Coomassie staining: ... 100
4.3.2. PARylation reaction ... 101
4.3.3. Click reaction ... 102
4.4. Analysis of proteins ... 102
4.4.1. General procedure of digesting proteins in gel ... 102
4.4.2. Desalting and concentration of samples using Zip Tip pipette tips…….. ... 103
4.4.3. Desalting of the sample using Slide-A-Lyzer MINI dialysis devices. ... 104
4.5. Mass spectrometric methods ... 105
4.5.1. MALDI-TOF mass spectrometry ... 105
4.5.2. ESI-ion trap mass spectrometry ... 106
4.5.3. ESI-Orbitrap mass spectrometry ... 106
4.6. Synthesis of carbasugar with benzyl protection group ... 107
4.7. Synthesis of carbasugar with trityl protection group ... 113
4.7.1. Attempts of coupling carbasugar to adenosine ... 119
5. Abbreviations ... 127
6. Index of Figures ... 130
7. Index of Tables ... 136
8. References ... 137
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1. Introduction
1.1. From DNA discovery through DNA damage to PAR
In 1869, the Swiss chemist F. Miescher discovered the DNA [1] (Figure 1). After more than eighty years, Watson and Crick made the groundbreaking conclusion that the DNA molecule exists in the form of a three-dimensional double helix. This discovery was based on the analysis of X-ray data of Wilkins and Franklin as well as the Chargaff′s observation that the ratio of adenine to thymine and guanine to cytosine were present in fixed ratios in all studied species [1].
Figure 1 Selected evolutionary landmarks in PARP field. Pictures of scientists taken from: F. Miescher [1], A. Harden [2], F. Crick and J. D. Watson [3], P. Chambon [4], M. Miwa [5], T. Sugimura [6].
At that time, it was not known how many factors are damaging DNA, for example: UV, x-rays, ionizing radiations as well as gamma-rays, viruses, oxidants, alkylating and intercalating agents. Among others, DNA can carry the following damages: oxidation, alkylation, deamination, mismatch of bases,
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single or double strand breaks [7]. With time, more knowledge was acquired about the repair mechanisms such as: reversion repair, mismatch repair (MMR), nucleotide excision repair (NER), DNA double strand brake repair and base excision repair (BER) (Figure 2) [7, 8].
Figure 2 DNA damaging agents, consequences and repair mechanisms. DNA damaging agents cause multiple DNA lesions which are removed and repaired via specific DNA repair pathways. Abbreviations:
cis-Pt: cisplatin and MMC: mitomycin C, respectively (both DNA-crosslinking agents); (6–4) PP:
photoproduct and CPD: 6–4 cyclobutane pyrimidine dimer (both induced by UV light); BER and NER, base- and nucleotide-excision repair, respectively; HR, homologous recombination; EJ, end joining. Figure adapted from [7].
After the discovery of the DNA structure in 1953, the DNA repair involving many proteins has become one of the most interesting topics in modern biology. One of such proteins, which takes part in DNA repair, is poly(ADP-ribose) polymerase (PARP). Investigation of PARP started in the 1960s, when three research groups pioneered the discovery of a novel nucleic acid-like macromolecule, poly(ADP-ribose) PAR. The first group, P. Chambon, J. D. Weill and P. Mandel published in 1963: “We discovered that (…) nicotinamide mononucleotide treatment enhances the activity of a DNA-
15 dependent enzyme which incorporates ATP into a product which is presumably polyadenylic acid; we wish to report some properties of this enzyme which does not seem to have been described as yet” [9]. Initially, it was assumed, that the homopolymeric reaction product is poly-A, but soon there was agreement that the product is indeed PAR [10-13] whose formation is dependent on NAD+ which has already been known since 1906 [2]. This finding marked the birth of an intriguing and rapidly growing field of poly(ADP-ribosyl)ation. PAR has aroused the interest of a large number of scientists coming from a very broad range of fields of scientific research.
In 1964, R. J. Collier and A. M. Pappenheimer [14] found that NAD+ was required to inhibit the protein synthesis by diphteria toxin in a cell-free system, which led later to the discoveries and to the begin of the research about mono(ADP-ribosyl)ation transferases (MARTs) [15].
M. Miwa and T. Sugimura discovered in 1971 poly(ADP-ribose) glycohydrolase (PARG), an enzyme that cleaves the ribose-ribose bond in poly(ADP-ribose) [16]. Later on, the same authors described in detail the branched structure of poly(ADP-ribose) [16]. The enzyme responsible for the synthesis of poly(ADP-ribose) was named PARP and it was purified to homogenity by many investigators using chromatographic methods. To obtain an electrophoretically pure enzyme, single-step affinity chromatography with 3- aminobenzamide as a ligand was used. Availability of purified PARP allowed to carry out further biochemical studies [17, 18]. However, it took an additional decade to isolate the genes encoding the proteins responsible for ADP- ribosylation reactions. In the late 1980s, the gene encoding a poly(ADP-ribose) synthetase (initially named PARP, poly(ADP-ribose) synthetase, or poly(ADP- ribosyl)transferase and now named PARP-1) was isolated [15]. With the cloning of the human PARP cDNA in 1987 by three groups [19-21], the era of molecular biology and molecular genetics of poly(ADP-ribosyl)ation reactions started. The overexpression of cloned PARP cDNAs in various expression systems provided new tools to investigate the modular organization of the enzyme in three functional domains [19, 22].
Easy purification of large amounts of a recombinant chicken PARP catalytic fragment led to the X-ray determination of its structure [23]. In the 80′s,
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the involvement of poly(ADP-ribosyl)ation in DNA repair was proved. The investigations showed that PARP inhibitors such as nicotinamide, benzamide, 3-aminobenzamide prevent rejoining of DNA strand breaks enhanced cytotoxicity even when low dose of alkylating agents was used [24]. This led to the idea of combining DNA-damaging cytotoxic agents used in cancer chemotherapy with PARP inhibitors. PARP has also been shown to be activated independently of DNA strand breakage by binding to DNA structures containing nicks, gaps, cruciforms and DNA bent structures. During the 1970s and 1980s, several laboratories partially purified different enzymes associated with mono(ADP-ribosyl)ation and poly(ADP-ribosyl)ation activities. Next, target amino acids for covalent modification of proteins with PAR were identified i.e.
glutamate, aspartate, lysine, and quantitative assays to determine PAR levels in living cells were established [25]. In the 80′s, H. C. Lee and co-workers described one more type of ADP-ribosylation reaction: the cyclization of ADP- ribose, which leads to cyclic-ADP-ribose which serves as an important second messenger involved in the regulation of calcium signaling and homeostasis [15].
Recently, the crystal structure of PARP-1 [23, 26, 27] followed by a more comprehensive coverage of the additional members of the PARP family and a histone macrodomain as a PAR binding motif were discovered [28]. Mono(ADP- ribosyl)ation and poly(ADP-ribosyl)ation were postulated to be reversible posttranslational modifications of proteins, acting as regulatory mechanisms for proteins.
1.2. Metabolism of NAD
+NAD+ (Figure 3) plays an essential role in the energy metabolism. It can not only be used as a coenzyme for a large number of oxidoreduction reactions, but also it can serve as a substrate for several reactions of ADP-ribosyl transfer [29]. Furthermore, NAD+ has a great influence on ATP synthesis as well as on the balance of the cellular redox potential. Poly(ADP-ribosyl)ation has turned out to be a major NAD+-consuming process in eukaryotic cells that can interfere with other vital NAD+-dependent and cellular functions [30].
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Figure 3 Structure of nicotinamide adenine dinucleotide (NAD+). The coenzyme is built by two nucleotides:
nicotinamide (vitamin) bound via beta-glycosidic bond with ribose and adenosine connected with phosphate bond.
Cellular NAD+ derives from three precursor molecules (L-tryptophan, nicotinic acid and nicotinamide) (Figure 4). NAD+ can be synthesized de novo in a multistep synthesis which starts from L-tryptophan and goes to the following intermediates: N-formyl-kynurenine, L-kynurenine, 3-hydroxy-L-kynurenine, 3- hydroxy-anthranilic acid to quinolinic acid as intermediate for nicotinic acid mononucleotide (NaMN) which is then converted to the corresponding dinucleotides nicotinic acid adenine dinucleotide (NAAD), and to NAD+ [2, 31].
In a salvage pathway, NaMN can also be synthesized from nicotinic acid (NA) (through the Preiss-Handler pathway), nicotinamide (collectively termed niacin), or nicotinamide riboside to generate nicotinamide mononucleotide (NMN). This intermediate molecule is then converted into NAD+ by nicotinamide mononucleotide adenylyltransferases (NMNAT), which are essential enzymes in the NAD+ metabolism as they catalyze the final step in the NAD+ biosynthesis [30, 32, 33]. Cellular processes dependent on NAD+ can be either NAD+- consuming or non-consuming. The latter ones imply redox reactions catalyzed by NAD+ dehydrogenases, in which NAD+ and its reduced form NADH serve as cofactors for the generation of ATP. Moreover, NAD+ is involved in distinct signaling pathways and is mainly a substrate in four types of ADP-ribose transfer reactions: (1) ADP-ribose cyclization, which is involved in calcium signaling; (2) deacetylation of proteins by the family of sirtuins, which are involved in many cellular functions including chromatin remodeling, gene silencing and genomic stability, resulting in O-acetyl-ADP-ribose; (3)
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mono(ADP-ribosyl)ation which is associated with intracellular as well as extracellular functions in cell signaling and metabolism; and (4) poly(ADP- ribosyl)ation [15, 32, 33].
Figure 4 NAD+ metabolism. Several metabolic routes allow NAD+ synthesis from four different precursors (depicted in blue boxes). In the de novo pathway, NAD+ is synthesized from L-tryptophan, an import pathway originates with nicotinic acid (NA), and a salvage pathway uses nicotinamide (NAM) to regenerate NAD+. Na and Nam are collectively referred to as niacin, or vitamin B3. A fourth, recently discovered route incorporates nicotinamide riboside in the salvage pathway. NAD+ and its phosphorylated relative NADP (not shown) are used as cofactors in several different redox reactions that are catalyzed by NAD+ dehydrogenases as well as in ADP-ribose transfer reactions. NaDS, NAD+ synthase; NaMNAT, nicotinic acid mononucleotide adenylyltransferase; Nampt, nicotinamide phosphoribosyl transferase; NaPRTase, nicotinic acid phosphoribo- syltranferase; NMNAT, nicotinamide mononucleotide adenylyltransferase;
Nrk1, nicotinamide riboside kinase-1. Figure adapted from [33].
19 In addition to the classical role of NAD+ as a coenzyme in cellular redox reactions, it also participates in several signaling pathways [2]. Poly(ADP- ribosyl)ation is the most extensive NAD+-consuming cellular process [34]. The hydrolysis of NAD+ into ADP-ribose and nicotinamide produces a free energy of -43.3 kJ/mol [35], which is used by PARPs to catalyze the synthesis of PAR.
The cellular concentration of NAD+ is approximately 400 - 500 µM with a KM of PARP-1 for NAD+ between 20 and 80 µM [36]. However, PARP-1 activation following DNA damage can consume massive amounts of NAD+ in a dose- dependent manner [37]. The cellular level of poly(ADP-ribosyl)ation seems to be the most important determinant for the metabolism of NAD+ in cells [38]. Indeed, it was shown that cells treated with high doses of genotoxic agents such as
γ-rays, N-methyl-N′-nitro-N-nitrosoguanidine (MNNG), methyl methane-
sulfonate, N-methylnitrosurea, H2O2, peroxynitrite, bleomycin and several others, display a strong decrease in cellular NAD+ levels within 5-15 min upon the DNA-damaging insult [38]. This reduction in NAD+ levels has clearly been linked to an increased PARP activity and elevated cellular PAR levels [25, 37, 39]. DNA-damage triggered NAD+ depletion is immediately associated with ATP depletion, because NAD+ resynthesis requires two molecules of ATP per molecule of NAD+ [30]. Recently, it has been shown that NMNAT-1 functionally associates with PARP-1, thereby stimulating PARP-1 activity. This activation of PARP-1 is further dependent on the phosphorylation state of NMNAT-1, which provides not only the substrate NAD+ for poly(ADP-ribosyl)ation, but also modulates PARP-1 activity [40]. In conclusion, poly(ADP-ribosyl)ation depends on intracellular NAD+, which is also linked to various other cellular processes ranging from energy metabolism and cellular signaling to chromatin remodeling [15, 33].
1.3. Poly(ADP-Ribose) Metabolism
Poly(ADP-ribosyl)ation is a post-translational modification of nuclear acceptor proteins which is a very dynamic process as revealed by the short half- life of less than 1 min of the polymer in vivo [41]. Under physiological conditions when no DNA single and double-strand breaks can be observed, poly(ADP-
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ribosyl)ation is a very rare event. Basal levels of PAR are simultaneously very low [42-44]. DNA strand breaks can induce PARP activity leading to an increase in PAR formation by 10- to 500-fold [25, 45, 46] upon DNA damage. Under these conditions about 90 % of PAR is synthesized by PARP-1, which is at the same time the main acceptor [47] apart from other acceptor proteins involved in DNA metabolism, DNA damage signaling and chromatin architecture [29, 38, 47]. The formation of PAR is initiated by an ADP-ribosyl transfer reaction in which NAD+ is used as a substrate and its hydrolysis releases nicotinamide and covalently transfers the ADP-ribose moiety onto glutamic acid, aspartic acid, via formation of ester bond and to carboxyterminal lysine and lysine residues via ketoamide bond (Figure 5) [48, 49]. The transfer onto glutamic acid residues was controversially discussed [49, 50] however recent studies are confirming modification on glutamic acid residues [51]. The initial protein-conjugated ADP- ribosyl unit is elongated during repeated cycles of ADP-ribosyl transfer using NAD+ as a substrate, leading to the formation of the poly(ADP-ribose) biopolymer and the stoichiometric release of nicotinamide as a by-product, (Figure 6).
The poly(ADP-ribosyl)ation reaction requires three steps for PAR synthesis: initiation - transfer of a first ADP-ribose moiety from NAD+ onto the acceptor protein, elongation - successive addition of further ADP-ribose units to the formed protein-mono(ADP-ribose) adduct and branching - linear polymer chain can be branched and can be elongated. The existence of PAR in living cells is a very transient and dynamic process with a very short half-life during DNA damage and a longer half-life in unstimulated cells [38]. Different enzymes were shown to catalyze polymer hydrolysis: PARG, which is mainly hydrolyzing PAR by cleaving ribose-ribose bonds in linear and branched regions of PAR to generate free ADP-ribose, ADP-ribosylhydrolase-3 (ARH3) and ADP-ribosyl protein lyase.
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Figure 5 Connection of first ADP-ribose monomer with amino acids: (A) ketoamide bond formation with lysine, (B) ester bond formation with glutamic or aspartic acid. Figure adapted from [49].
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Figure 6 PolyADP-ribose metabolism: Initiation, elongation and branching represent anabolic reactions (depicted in green), while both types of cleavage represent catabolic reactions (depicted in red). The synthesis of poly-ADP-ribose requires three activieties: initiation or mono(ADP-ribosyl)ation on specific amino acid residue (in Figure only the ester bond is shown, which is formed when the modification is on glutamic acid or aspartic acid; for ketoamide bond formed with lysine residue see Figure 5) in the acceptor protein following by elongation of polymer and branching. The degradation requires different enzymes:
exoglycosidase and endoglycosidase PARG activietis that hydrolyze the glycosidic linkages between the ADP-ribose units, potential poly(ADP-ribosyl)protein hydrolase activities and in the last step MARH or mono(ADP-ribosyl)protein lyase activities. Figure adapted from [15, 48].
23 The release of the remaining protein-proximal ADP-ribose unit may be accomplished in a ß-elimination step by an ADP-ribosyl protein lyase or by PARG itself [52, 53]. Recently, a mammalian enzyme with poly(ADP-ribose) glycohydrolase activity called ADP-ribosylhydrolase-3 (ARH3) was described [54]. ARH3 exhibits PARG activity but it is structurally unrelated to PARG and unable to cleave the ADP-ribose moiety attached to the acceptor proteins [55- 57]. The degradation of PAR by PARG is essential for cell survival, as PARG gene disruption on exon 4 leads to massive PAR accumulation and embryo lethality [58].
1.3.1. Structure of poly (ADP-ribose) PAR
Poly(ADP-ribose) (PAR) is a heterogeneous linear or branched, highly negatively charged homopolymer of repeating ADP-ribose units (Figure 6).
Monomers are linked by unique glycosidic 1′′-2′ ribose phosphate-phosphate bonds, formed by the hydrolysis of the substrate NAD+ [12]. Branching results from the formation of glycosidic 1′′‘-2′′ ribose-ribose bonds and generally occurs after every 20 to 50 ADP-ribose unit [15, 29, 59-61]. The covalently bound ADP- ribose unit serves as a starting point for successive transfer reactions onto the protein–mono(ADP-ribosyl) adduct and subsequently onto the emerging chain of several covalently linked ADP-ribosyl residues for building linear and branched polymers by adding further ADP-ribose moieties. PAR is a complex biopolymer whose chain length can reach from 2 units in linear structures to more than 200 ADP-ribose units in multibranched molecules [29, 38, 62]. The major hydrolysis product contains one mole of adenine, two moles of ribose and two moles of phosphate per mole of nucleotide [10]. The product is susceptible to cleavage by phosphodiesterase, but not by alkaline hydrolysis. The anomeric carbon of one ADP-ribose moiety is bound to the adenosine moiety of the adjacent one via a 1′′ – 2′ glycosidic linkage which is an α anomeric linkage as it was shown by NMR. Miwa et al. first reported that the molecular weight of isolated polymers calculated from the ratio of PR-AMP to 5′′-AMP was not consistent with the sedimentation coefficient value determined by centrifugation.
This suggested aggregation of poly(ADP-ribose) chains or a branched structure,
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as in the case of polysaccharides. Identification of the nucleotide, O-α-D-ribofuranosyl (1′′′–2′)-O-α-D-ribofuranosyl-(1′′-2′)-adenosine-5′,5′′,5′′‘-
tris(phosphate), abbreviated as (PR)2-AMP, in phosphodiesterase digests of polymers confirmed the presence of branching in the polymer. The content of branching residues was estimated to range from 2 % to 3 % of the total ADP- ribose residues. Thus, more than 6.6 branching points have been identified in purified polymers of an average size of approximately 250 residues, and the number of branches per polymer increases with polymer size [10]. The constitutive level of PAR is very low in intact cells under physiological conditions; however, DNA damage triggers the synthesis of PAR. This is accompanied by a decrease in intracellular NAD+ concentrations [29, 37, 63].
Furthermore, it has been suggested that PAR chains adopt a helical secondary structure, due to the formation of hydrogen bonds, similar to those of DNA and RNA [64, 65]. A recent NMR study, however, did not show evidence for such a feature: PAR did not adopt any regular three-dimensional structure [66].
Due to the high negative charge of the polymer, this modification significantly alters the physical and biochemical properties of modified proteins, such as their DNA-binding affinity, and it is likely that such alterations will have a regulatory function concerning the interaction with other proteins.
1.3.2. Poly(ADP-ribose) Glycohydrolase (PARG)
Poly(ADP-ribose) glycohydrolase (PARG EC 3.2.1.143) is the only nuclear enzyme known to hydrolyze glycosidic linkages. While several genes are known to code for PARPs, there is only a single PARG gene known to encode an enzyme catalyzing the hydrolyzis of (ADP-ribose) polymers to free ADP-ribose [48, 67]. In 1997, a bovine complementary DNA (cDNA) encoding a protein possessing PARG activity was identified and characterized [68]. The human PARG gene has been assigned to chromosome 10q11.23 consisting of 18 exons and 17 introns which encodes for a protein of 111 kDa [69].
Three different splice variants were described giving rise to three isoforms. One large 111 kD isoform is targeted to the nucleus, whereas two smaller 102 and 99 kD isoforms are found predominantly in the cytoplasm [70]
25 respectively with exo- and endoglycosidic activities [70-73]. These observations led to the identification of a putative nuclear export signal NLS in the region of the protein coded by exon 1. The A domain of the PARG isoforms also contains other features that may be involved in the regulation of the protein, for example a caspase 3 cleavage site that is used during apoptosis [74] and has been reported to contain a putative nuclear export signal [75]. Two novel isoforms of PARG have also been identified that possess mitochondrial targeting sequences: hPARG-55/mPARG-58 may participate in the signaling of PAR from the nucleus to mitochondria [70, 73]. The presence of PARG in mitochondria is intriguing, as no definitive demonstration of PARP activity in mitochondria has been reported. Degradation of PAR proceeds in three phases: (1) endoglycosidic cleavage; (2) endoglycosidic plus exoglycosidic cleavage with processive degradation; (3) exoglycosidic cleavage with distributive degradation [71, 72, 76]. PARG is depredating branched and short polymers more slowly than long and linear ones. Moreover, the rates of hydrolysis of PAR bound to various proteins were found to be higher than those of free polymer [77]. The endoglycosidic activity of PARG plays a crucial physiological role by releasing free PAR, which is important in different cellular processes resulting in multiple physiological consequences through non-covalent protein binding and intra- as well as extra-nuclear signaling functions e.g. cell death processes [15, 76, 78].
Its products are free poly(ADP-ribose) and monomeric ADP-ribose, the latter one being a potent protein-glycating carbohydrate capable of causing protein damage. It was also shown that an ADP-ribose pyrophosphatase converts ADP- ribose to AMP and ribose 5-phosphate, thus decreasing the risk of non- enzymatic protein glycation [79]. The significance of the endoglycosidase activity of PARG is not clear, but the enzyme has the potential to generate different products depending upon its state of saturation. For example, under conditions where the enzyme is not saturated with substrate, free ADP-ribose and protein bound ADP-ribose polymers would be the expected products, but when saturated with a substrate, the enzyme would be expected to generate free ADP-ribose polymers. It has generally been assumed that PARG is unable to catalyze the removal of the protein proximal ADP-ribose residue; however, a recent reexamination of this issue indicates that PARG can catalyze the
26
removal of protein proximal ADP-ribose residues linked to protein carboxylate groups of histone H1 [80]. Recently, in Drosophila melanogaster a loss-of- function mutant has been described that lacks the conserved catalytic domain of PARG. This mutant exhibits lethality in the larval stages at the normal developmental temperature of 25 °C [81]. However, about 25% of the mutants progress to the adult stage at 29 °C, but showed progressive neurodegeneration with reduced locomotor activity and a shortened lifespan. In association with this, extensive accumulation of poly(ADP-ribose) could be detected in the central nervous system. These results suggest that poly(ADP- ribose) metabolism is required for maintaining the normal function of neuronal cells.
Recently, knockdown of PARG isoforms by stable and constitutive expression of a short hairpin RNA in HeLa cells has resulted in beneficial effects in undamaged cells, as they were protected from spontaneous single-strand breaks and telomeric abnormalities. On the other hand, irradiation of these PARG deficient cells showed centrosome amplification leading to mitotic supernumerary spindle poles and accumulated aberrant mitotic figures, which caused either polyploidy or cell death [82].
1.3.3. ADP-ribosyl protein lyase
Early studies of ADP-ribose polymer metabolism concluded that PARG was unable to remove the protein proximal ADPR residue from acceptor proteins [80]. This led two decades ago to the isolation of an ADP-ribosyl protein lyase (formerly termed ADP-ribosyl histone splitting enzyme), which removes the protein-proximal ADP-ribosyl residue linked to the acceptor protein and it was isolated from rat liver and purified by Hayaishi and co-workers [52, 83]. This enzyme is unique amongst mammalian enzymes. ADP-ribosyllyase cleaves the last ADP-ribose residue left on the acceptor by PARG and its reaction results in the dehydrated form of ADP-ribose (5′-ADP-3′′-deoxypent-2′′- enofuranose) by an enzymatic elimination [48, 52]. The substrate specificity of the ADP-ribosyl protein lyase is not very dependent on the protein portion, but it is highly specific for the mono(ADP-ribosyl) moiety [10]. Morover, it was
27 discovered that it did not split a reduced Schiff base, showing to be specific to the carboxyl ester bond of the protein. Furthermore, the enzyme cleaved a mixture of ADP-ribosyl non-histone proteins [83], but not the ADP- ribosylnicotinamide bond in NAD+ or the carboxyl ester bond in p-nitrophenyl acetate [52]. Wielckens et al. [39] showed that a removal of primary ADP-ribosyl groups from acceptor proteins was the rate-limiting step in the overall turnover of poly(ADP-ribosyl) groups in cells treated with a DNA-fragmenting agent, leading to the conclusion that ADP-ribosyl protein lyase plays a principal role in ADP-ribosylation/de(ADP-ribosyl)ation systems in mammalian cells. Although this enzyme was discovered many years ago, still little is known about its structure - function relationship and role in ADPR polymer metabolism. Recent studies showing that PARG can catalyze the removal of protein proximal ADPR residues linked to the carboxylate groups of histone H1 raised additional questions. It is possible that the property of both enzymes to catalyze the removal of these residues represents redundancy in function or that specific polymer acceptor proteins require different enzymes to catalyze the removal [80].
1.4. PARP-1
The prototypic enzyme of the PARP family is poly(ADP-ribose) polymerase-1 (PARP-1 [EC 2.4.2.30]), which is a highly conserved enzyme and has been best-studied as it accounts for 90 % of cellular PAR formation after DNA damage [29, 84]. PARP-1 is present in the nucleus (106 molecules per cell), and in centrosome [80, 85, 86]. However, the results regarding the localization of PARP-1 in mitochondria are conflicting and require further investigations [87, 88]. The human Parp1 gene is localized on chromosome 1 (1q42) [19, 89] encoding 1014 amino acids with a molecular weight of 113 kDa, consisting of several domains with distinct functions [19, 90]. PARP-1 has been identified in most eukaryotic organisms: animals, plant, fungi and protist kingdoms [91]. PARP-1 is a molecular nick sensor which is able to recognize distortions in the DNA helical backbone and binds to three and four way junctions as well as stably unpaired regions in double-stranded DNA [63, 92-
28
94]. Recent studies have demonstrated that PARP-1 may be activated not only by DNA breaks, but also by other stimuli, such as D-myo-inositol-1,4,5- triphosphate, which does not directly involve DNA damage. These studies indicate that PARP-1 might be involved in new DNA damage-independent, poly(ADP-ribosyl)ation-dependent signaling pathways. Interactions of PARP-1 with non-B DNA structures led to the catalytic activation of the enzyme in the absence of free DNA ends. DNA hairpins, cruciforms and stably unpaired regions are all effective co-activators of PARP-1 automodification and poly(ADP-ribosyl)ation of histone H1. Nevertheless, automodification of PARP-1 was reported to be mostly induced by single-strand breaks, whereas histone H1 seems to be modified preferentially when PARP-1 binds to double-strand breaks [29, 48]. PARP-1 interacts independent from the DNA sequence since it contacts ribose-phosphate backbone. PARP-1 engages the break in the DNA through hydrophobic interactions with exposed nucleobases which are a common feature of damaged DNA structures [26].
1.4.1. Biological importance of PARP-1
In response to DNA damage, PARP-1 undergoes conformational changes, leading to a 100 to 1,000 fold increase in activity to catalyze the covalent transfer of ADP-ribose moieties from NAD+ on nuclear target proteins producing PAR chains [29, 38]. Kinetic studies have revealed that PARP-1 is catalytically active as a homodimer. Furthermore, PARP-1 dimerization has been shown to be a prerequisite for its DNA-dependent stimulation [95, 96]. In vitro automodified PARP-1 may carry as many as 15 polymer molecules with an average chain length of 80 ADP-ribose subunits [59]. In the absence of DNA breaks, PARP-1 displays a very low basal enzyme activity. The main target of covalent modification is PARP-1 itself, as it catalyzes its automodification as well as many other target proteins such as p53 [97], both subunits of NF-κB [98], histones [99], CSB, DNA-topoisomerases [100] and the catalytic subunit of DNA-dependent protein kinase (DNA-PKcs) (Figure 7) [47, 98, 101-103].
Among the interaction partners of PARP-1 are also other members of the PARP-family, such as PARP-2 [104-106] and PARP-3 [107] (Figure 7).
29 Besides the covalent modification of acceptor proteins, poly(ADP-ribose) is known to interact non-covalently with a wide range of proteins. There are at least two different evolutionarily conserved poly(ADP-ribose) binding motifs that enable non-covalent binding of free or protein-bound poly(ADP-ribose) to proteins [29, 108, 109]. The specificity of such non-covalent interactions might be dependent on the degree of polymer branching or polymer chain length [78, 110]. This interaction is mediated through a sequence motif displaying a conserved pattern and it is interesting that DNA damage checkpoint proteins such as p53 and p21 possess poly(ADP-ribose) interaction domains. DNA methytransferase-1 (DNMT1) is also able to interact with PAR in a non-covalent fashion, which leads to the inhibition of DNMT1 activity [111]. Taken together, poly(ADP-ribose) can affect the function of proteins that are not direct modification targets. Automodification occurs in an intermolecular reaction, since PARP-1 acts as a catalytic dimer [95].
30
Figure 7 PARP-1 and its interaction partners with their role in genomic maintenance. ATM indicates ataxia telangiectasia mutated; Bub3, Budding uninhibited by benzimidazoles 3; Cenpa/b, centromeric protein a/b;
CSB, Cockayne syndrome type B; DEK, DEK oncogene; DNA-Polβ, DNA polymerase β; DNA-PKCS, DNA-activated protein kinase catalytic subunit; HMGB1, high mobility group box 1; Ku70/80, Ku antigens 70/80 kD subunit; MRE11, meiotic recombination 11; p21, cyclin-dependent kinase inhibitor 1A; p53, tumor protein p53; PCNA, proliferating cell nuclear antigen; TRF2, telomeric repeat binding factor 2; WRN, Werner syndrome protein; XRCC1, x-ray repair cross-complementing protein; XPA, xeroderma pigmentosum complementation group A [15, 47, 97-100, 103].
Auto-poly(ADP-ribosyl)ated PARP1 can interact with other DNA repair factors, in particular those involved in base excision repair (BER) by modifying itself and other proteins with PAR and participating in the spatial and temporal organization of the repair process through specific non-covalent interaction with other DNA repair proteins (Figure 8) [33, 112-114]. Thereby, a DNA repair complex can be formed that initiates the DNA repair machinery [115, 116].
PARP-1 binds in a specific manner to nucleosomes and modulates chromatin structure through NAD-dependent automodification, without modifying core histones or promoting the disassembly of nucleosomes [117]. It was also
31 suggested that fast and transient decondensation of the chromatin structure by poly(ADP-ribosyl)ation occurring in Aplysia californica in the absence of DNA strand breaks enables the transcriptional regulation needed to form the long- term memory in this organism [118].
Figure 8 Role of PARP in the regulation of DNA repair. Binding of PARP-1 to single strand break (SSBs) results in auto-poly(ADP)-ribosylation and increased activity. Activated PARP-1 is involved in DNA repair via three rounds: direct interaction and poly(ADP) ribosylation of XRCC1 and Polβ, leading to stimulation of BER; poly(ADP) ribosylation and activation of the 20S proteasome, leading to relaxation of the chromatin structure; and potential poly(ADP) ribosylation of DNA-repair proteins, thus modulating DNA repair [112].
In addition to its role in DNA repair [105, 112], PARP-1 plays a crucial role in many biological structures and processes such as maintenance of genomic stability [63, 112, 119] including the mediation of cellular recovery, cell cycle control [120], transcriptional regulation [121-123], the maintenance of
32
telomeres [124, 125], structure and function of vault particles, centromere function and mitotic spindle formation [38, 104, 126], regulation of proteasomal protein degradation [127], genotoxic stress, and aging, as well as elimination of overly damaged cells [128] (Figure 9).
PARP-1 cell cycle
ERK2 p53 CenpA/B Bub3
G0-G1 Mitosis
DNA repair Ligase3 XRCC1
Pol XPA
CSB
MRN Ku70
BER NER
DSB repair
DNA repair DNA repair
Caspase cleavage AIF release NAD depletion
Apoptosis Necrosis
TF (NFkB,
E2F, p53...) H1;H2B
DEK Coactivator/repressor Chromatin
decondensation
Figure 9 Cellular functions of PARP-1: PARP-1 is involved in a variety of cellular processes such as cell cycle, DNA repair which is strictly connected to aging and cancer, cell death connected with cancer, inflammation and diabetes as well as transcription connected with cancer and inflammation. PARP-1 regulates the different processes either by poly(ADP-ribosyl)ation, by direct interaction with the distinct proteins or by its metabolic impact on NAD+ metabolism.
Moreover, PARP-1 binding to non-B DNA suggests its function in the dynamics of local modulation of chromatin structure in normal cellular physiology [29].
However, PARP-1 also acts as a cell death mediator and it is involved in pathophysiological processes such as myocardial infarction, stroke, inflammatory diseases and Type I diabetes [48, 63, 129-131].
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1.4.2. Structure of PARP-1
Poly(ADP-ribose) polymerase is a multidomain protein. Early work from the Shizuta′s laboratory demonstrated that PARP can be divided into three functional domains [10] (Figure 10) representing the distinct biochemical activities and functional roles. Limited proteolysis with mild trypsin and papain of the purified enzyme showed that the N-terminal region contains a 46 kDa DNA- binding domain (DBD) (amino acid, aa1–aa373) with two zinc fingers, followed by a central 22 kDa automodification domain (aa374–aa525) and at the C- terminus a 54 kDa catalytic domain (CAT) (aa526–aa1014) containg the PARP signature which is conserved in all PARP homologues [38, 82, 132]. The three- domain structure can be further divided into specific modules A–F (not shown) [29, 133].
Figure 10 Modular structure of PARP-1. PARP-1 consists of three main domains. The DNA binding domain which comprises three zinc finger motifs (FI, FII and zinc binding domain), a nuclear localization signal (NLS) and a caspase cleavage site (C3/7). The automodification domain is located in the center with a breast cancer susceptibility protein-carboxy (C) terminus motif (BRCT). At the C-terminus there is the catalytic active domain with the NAD+ binding site with the highly conserved PARP signature.
1.4.2.1. The DNA-binding domain
The N-terminal 42 kDa DNA binding domain (DBD) contains two zinc finger domains, FI (cysteine aa21-cysteine aa56) crucial for the stimulation of PARP-1 triggered by DNA double strand breaks and ZFII (cysteine aa125- cysteine aa162) which is essential for single-strand break [133-135]. DBD has
34
also two helix-turn motifs which are located at residues ~ 200 - 220 and ~ 280 - 285, respectively. The zinc fingers found in PARP-1 coordinate Zn2+ ions with a Cys-Cys-His-Cys motif [116, 134, 136]. In addition, a third zinc binding domain (FIII) (aa216-aa366) with a distinct structure and function from ZFI and ZFII was discovered [26]. ZFIII could be necessary to relay the DNA binding signal from the first two zinc fingers to the catalytic carboxy terminus by forming an interdomain contact. Such a contact is important for DNA-dependent PARP-1 activation [137]. It was discovered that the ZFI domain of one molecule can form a dimeric break-recognition module with ZFII domain of a second molecule. This fact is permitting a cooperating pair of PARP-1 molecules to interact simultaneously at both margins of a single-strand gap of sufficient size or to interact with two separated DNA ends [138].
The DNA damage-dependent dimerization of the PARP-1-DBD fulfills the essential requirement of bringing two PARP-1 molecules into close proximity to allow trans(ADP-ribosyl)ation [138]. Furthermore, the DBD contains a bipartite nuclear localization signal (NLS, 207-217 and 221-226) [139] for the nuclear homing of PARP-1, which includes a caspase-3 and caspase-7 cleavage site (DEVD, aa210-aa213, Asp-Glu-Val-Asp), where PARP-1 is cleaved in the execution phase of apoptosis [140, 141]. As a consequence, the cleavage of PARP-1 generates two proteolytic fragments, a 24 kDa amino terminus and an 85 kDa carboxyl terminus [142]. These motifs are capable of mediating interactions between DNA and proteins and might contribute to the DNA-binding activity of PARP-1.
1.4.2.2. The automodification domain
The automodification domain is located in the central region of PARP-1 and spans residues 374 to 525. This domain is rich in glutamic acid residues, representing the major acceptor amino acid for poly(ADP-ribosy)lation, consistent with the fact that poly(ADP-ribosyl)ation occurs on such residues [50, 143]. A leucine-zipper motif was identified in the N-terminal part of the automodification domain of Drosophila melanogaster PARP, which is conserved in chicken and mammalian PARPs [144]. This motif was proposed to participate in the dimerization of the enzyme, which was demonstrated to be catalytically
35 active as a homodimer [95]. This domain is a central regulating segment comprising a breast cancer-susceptibility protein carboxy terminus motif (BRCT) which is placed between amino acids 384 - 479. The BRCT motif is common in many DNA-repair and cell-cycle proteins and allows protein-protein interactions [145]. Importantly, this domain was observed to physically associate with the N- terminal BRCT domain of XRCC-1 (x-ray repair cross-complementing protein) [115].
1.4.2.3. The catalytic domain
The 55 kDa catalytic domain is located at the C-terminus and is composed of two subdomains, the helical subdomain (HD) and the ART subdomain [137]. ART accommodates the designated PARP signature sequence containing a 50-amino acid sequence (aa526-aa1014).
The crystal structure of chicken PARP-1 was resolved by X-ray diffraction in 1996 and showed structural homology to bacterial mono(ADP-ribosyl) transferases such as the C. diphtheriae toxin or pertussis toxin [23, 146]. In close relationship to the bacterial enzymes, PARP-1 contains a ß-α-loop-ß-α- structure (residues 859 - 908) containing three essential amino acids for its activity: lysine 893 and aspartate 993 are responsible of the attachment of the first ADP-ribose moiety onto the acceptor amino acid [46], whereas glutamate 988 seems to play a role in polymer elongation, although some authors demonstrated that a PARP mutant lacking the glutamate 988 still produce oligomers [147]. In addition, tyrosine 986 is important for the branching reaction [148] which is responsible for NAD+ binding, ADP-ribose transfer and branching [22, 46, 132].
Up to now, PARP-1 mutants have been studied in terms of activity and branching frequency, displaying for instance an increase in the enzyme activity (gain of function) due to a L713F mutation [149]. In addition, several other mutations have been characterized resulting in a strong decrease in the enzyme activity, e.g. the exchange of glutamate to lysine at position 988 (E988K) converts PARP-1 into a mono(ADP-ribosyl) transferase [150]. Mutations affecting the branching reaction have also been described and demonstrated the importance of residue Tyr-986. An exchange with serine decreased the
36
branching frequency about 7-fold, whereas the mutant Y986H displayed a higher branching frequency compared to wild-type PARP-1 [150]. It was proposed that the introduced His-986 binds more tightly to the pyrophosphate moiety positioned at the acceptor site, thereby increasing the symmetry of the acceptor site [27].
The region comprises also a WGR domain, defined by conserved tryptophane (W), glycine (G) and arginine (R) residues, with uncertain function.
It is speculated that the WGR domain is implicated in the binding of nucleic acids [151].
The last PAR binding motif that was discovered was the WWE domain, which shows affinity for iso-ADP ribose, i.e. the ADP-ribose monomer [152, 153].
1.5. PARP superfamily
For a long time, PARP-1 was the best-investigated PARP protein and has been thought to be the sole enzyme responsible for the synthesis of ADPR polymers [15]. PARP-1 is with PARP-2 crucially important for living organisms since PARP-1/PARP-2 double knockout mice exhibit embryonic lethality [105].
The first evidence that structurally different PARP proteins may possess DNA- dependent poly(ADP-ribose) activities arose in 1995, with the discovery of a gene coding for a PARP-related polypeptide APP in Arabidopsis thaliana.
Development of Parp1 gene deficient mice showed that PAR could still be synthesized [84]. This discovery led to the identification of five novel poly(ADP- ribosyl)ating enzymes [154-158], indicating that PARP-1 belongs to a family of poly(ADP-ribose)polymerase. The determination of the three-dimensional structures of the catalytic domains of chicken PARP-1 and mouse PARP-2 showed that these proteins had structural homology with the active site of the bacterial ADP-ribosylating toxin from C. diphtheria. PARP-1, PARP-1b, PARP-2, PARP-3, PARP-4, PARP-5, and PARP-6a have automodification activity and most likely covalent auto(ADP-ribosyl)ation activity. The conservation in the PARP signature varies significantly among the PARP-family members. Indeed, the catalytic residue (the Glu988 residue of PARP-1) is replaced by a non-
37 conserved residue in some members (tiPARP, PARP-9/BAL1, PARP-10, PARP- 13 and PARP-16). It is still not clear whether these proteins have any activity and poly- or mono(ADP-ribosyl)ating. So far, only PARP-9/BAL1 has been reported to be inactive, whereas poly(ADP-ribosyl)ating activity has been reported for tiPARP and PARP-10 [159-161]. Up to now, eight members of the PARP superfamily have been shown to be able to catalyze poly(ADP- ribosyl)ation and possess catalytic activity for the formation of poly(ADP-ribose), i.e. PARP-1, PARP-2, PARP-3, VPARP, tankyrase-1, tankyrase-2, PARP-7, TiPARP-1, and PARP-10 [29, 33, 80]. The PARP domain is located at the C- terminus of the protein, except for PARP-4/vPARP and is adjacent to various domains that are involved in DNA or RNA binding, protein-protein interactions or cell signaling [33]. Different PARPs have distinct primary structures, subcellular localizations and cellular functions. However, they have overlapping functions between the different members. For instance, PARP-1 and PARP-2 exhibit partially redundant functions and share a couple of common interaction partners [146]. A common feature of all PARPs is that they catalyze the production of ADP-ribose using NAD+ as a substrate [33]. Up to now, only PARP-1 and PARP-2 were demonstrated to be highly stimulated in the presence of DNA damage.
PARP-family members can be divided according to the homology of their catalytic domains or established functions: Subgroup I includes DNA-dependent PARPs: PARP-1, PARP-1b, which seems to be a product of an alternative transcription initiation site within the Parp1 gene (previously described as short PARP-1), PARP-2 and PARP-3. Experimental data suggest that both PARP-1 and PARP-2 play a major role in distinct stress response pathways. Subgroup II contains a single member, PARP-4 (vault-PARP), which is the largest one of the family (192.6 kDa) and was identified as a component of the vault complex. The vault complex is a cytoplasmic ribonucleoprotein complex of unknown function associated with an untranslated vault RNA and two other highly conserved proteins, major vault protein and telomerase- associated protein-1. Tankyrase 1, tankyrase 2a, and perhaps its alternatively spliced or transcribed isoform tankyrase 2b, which are referred to as PARP-5 and PARP-6a/b respectively, belong to subgroup III. Both PARP-5 and PARP-6a were identified as
38
components of the telomeric complex. In the PARP family, also CCCH-type PARPs (PARP-12, PARP-13 and tiPARP) and macroPARPs (PARP-9/BAL1, PARP-14/BAL2/ CoaSt6 and PARP-15/BAL3) can be distinguished.
Up to now, 22 PARP gene family members [91] have been identified by highly conserved PARP β−α-loop-β−α NAD+ fold, a signature motif, which is located at the C-terminus of the enzyme [146, 162], having a 50 amino acid sequence, displaying a conservation of 100 % among vertebrates and 92 % among animal species [33, 163, 164]. Recently, a new nomenclature for mammalian ADP-ribosyltransferases was proposed according to the International Union of Biochemistry and Molecular Biology (IUBMB) as ADP- ribosyltransferases (ARTs). Since some PARP family members are still not well characterized, especially regarding the product (poly(ADP-ribose) vs.
mono(ADP-ribose)), it was proposed to name them according to the type of enzymatic reaction and omitting the prefix “poly′′ and “mono′′ [91]. Several classifications have previously been proposed [15, 146, 162], but until now they have not been adapted.
The connection between DNA damage and DNA repair with cancer, ageing as well as autoimmune, cerebrovascular and coronary artery diseases is intensively studied. Since members of the PARP family play a key role in the genome surveillance, it would not be surprising if the research in the PARPs field will provide a breakthrough in understanding the cancer development, and, maybe more futuristic, the life span of humans.
1.6. Detection and quantification of PAR and PARP activity
Identification of protein interaction partners should help understanding the PARP function in the DNA repair and cellular processes. Moreover, the quantification of PAR as well as the determination of PARP activity would be extremely useful for drug development. All methods used to determine ADP- ribose polymers in intact cells and tissues must be very sensitive and selective, since the content of ADP-ribose polymers is very low relative to other adenine- containing nuclear polymers such as DNA and RNA. In particular, the
39 quantitative analysis of basal PAR levels in vivo is a challenging task that requires specific and highly sensitive methods. While the quantification of ADP- ribose polymers is still not easy, a number of chemical and immunological methods are now available for the investigation of ADP-ribose polymer metabolism. Moreover, it is highly important to be able to determine PARP activity in vitro and in vivo, to establish high-throughput assays for screening PARP inhibitors.
1.6.1. Chromatographic methods
The first milestone to determine ADP-ribose polymer content in cultured cells and animal tissues was achieved by Jacobson et al. [25]. Dihydroxyboryl Bio-Rex (DHBB) affinity chromatography for isolating PAR was used, followed by compound derivatization and LC-based detection, which provided an accurate quantification. These resins have been used in combination with both chemical and immunological detection methods to quantify ADP-ribose polymer residues in vivo. Unfortunately, routine usage was hampered by the amount of cellular material required, radioactive labeling or derivatization of the analytes, and the overall time-consuming procedure.
Another approach allowing the quantification of total polymer residues is based on the conversion of the nucleotides to highly fluorescent 1, N6-etheno derivatives. The mixture can be separated by reverse-phase HPLC and quantified at the picomole level [165]. Other approaches were based on treating cells with [3H]adenine as a radioactive precursor to NAD+ [166]. Although leading to precise results, these techniques were prone to overestimate PAR levels, due to artificial DNA damage induced by the radioactive isotopes and subsequent collateral PARP activation.
The combination of boronate chromatography with high-resolution anion- exchange HPLC has allowed the preparation and separation of oligomers up to the 50-mer and multibranched polymers [167]. Another chromatographic method involving two-dimensional thin-layer chromatography on cellulose plates has been used to separate and quantify the three diagnostic nucletides of poly(ADP-ribose) generated by phosphodiesterase digestion: AMP, PR-AMP, and (PR)2AMP.
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1.6.2. Immunodetection of PAR
Most of the current tools to quantify PAR rely on antibodies, which are widely used in basic research as well as preclinical and clinical studies [168, 169]. Four types of antibodies have been generated: anti-poly(ADP-ribose) sera [170], anti-PR-AMP sera [171], antibodies specific for 5′AMP [172] and antibodies against an analogue of ADP-ribose [173]. The first raised antibodies against poly(ADP-ribose) did not bind poly(A) or other related nucleotides, nor yeast RNA or calf thymus DNA, but bound poly(ADP-ribose) and, to a lesser degree, ADP-ribose and PR-AMP [174]. However, a cross-reaction with double stranded RNA, poly(A)-poly(U), or poly(I)-poly(C) duplexes was observed.
Furthermore, the reactivity of these antibodies against poly(ADP-ribose) was found to be dependent on the polymer size, binding smaller polymers less efficiently than larger polymers. A radioimmunoassay involving this antibody has been used to detect poly(ADP-ribose) in vivo, but these methods were semi- quantitative [175]. For quantitative determinations, prior fractionation of the sample according to polymer size was necessary.
In the early 1980s, the first monoclonal antibodies to poly(ADP-ribose) were reported to have a greater potential for quantitative immunoassays for ADP-ribose polymers, since monoclonal antibodies recognize specific antigenic determinants [176].
Characterization of two monoclonal antibody preparations demonstrated that one of them recognized the linear structure of ADP-ribose polymers, while the second one recognized additional structures including the branched portions of the polymers. Specific antibodies have proved to be very useful for the cytological detection of ADP-ribose polymers within individual cells [10].
Antibodies highly specific for poly(ADP-ribose) can be easily visualized by coupling them to dyes such as fluorescein isothiocyanate (FITC) or by using FITC-labeled anti-IgG antibody. Cross-reactivity of the antibodies with DNA is of particular concern, because DNA is present in nuclei in much higher concentration than poly(ADP-ribose). Most antibodies do not exhibit cross- reactivity with DNA, RNA, poly(A), ADP-ribose or NAD when examined by double-immunodiffusion and membrane-binding assays [177]. However, since
