The Functional Interplay of DEK and PARP-1/2 in the Replication Stress
Response
Dissertation
zur Erlangung des akademischen Grades des Doktors der Naturwissenschaften
(Doctor rerum naturalium)
an der
Mathematisch-Naturwissenschaftliche Sektion Fachbereich Biologie
vorgelegt von
Magdalena Ganz
Tag der mündlichen Prüfung: 28.06.2016
1. Referentin: Prof. (apl.) Dr. Elisa Ferrando-May 2. Referent: Prof. Dr. Alexander Bürkle
Konstanzer Online-Publikations-System (KOPS) URL: http://nbn-resolving.de/urn:nbn:de:bsz:352-0-348315
Zuerst möchte ich mich bei Frau Prof. Dr. Elisa Ferrando-May bedanken. Herzlichen Dank für die Aufnahme in die Arbeitsgruppe, die Möglichkeit an einem sehr spannenden Thema zu arbei- ten, sowie für die gute fachliche und auch persönliche Betreuung während dieser Zeit.
Als Mitgliedern meines Thesis Komitees danke ich Herrn Prof. Dr. Alexander Bürkle und Herrn Prof. Dr. Andreas Marx für ihr Interesse an meiner Arbeit, für die fachliche Unterstützung und wertvolle Anregungen, sowie für die Erlaubnis Gerätschaften aus ihren Laboren zu nutzen.
Herrn Prof. Dr. Alexander Bürkle und Herrn Prof. Dr. Christof Hauck danke ich für ihre Bereit- schaft, in meinem Prüfungskomitee mitzuwirken.
Auch möchte ich herzlichst allen andern Personen danken, die durch ihre Unterstützung zum Gelingen dieser Arbeit beigetragen haben: Dr. Ferdinand Kappes und Malte Prell von der Uni- versität Aachen danke ich für den super Austausch, für die Hilfe bei fachlichen Fragen, für die Weitergabe von Protokollen sowie die freundliche Bereitstellung von Plasmiden, Proteinen und Zellen. Meinen Kollegen von der AG Bürkle, allen voran Jan, Aswin, Sebastian und Arthur, dan- ke ich für ihre theoretische und praktische Unterstützung in der Biochemie, die Versorgung mit Materialien und für ihre offenen Ohren für jede noch so kleine PARP Frage. Dr. Benjamin Hanf danke ich für die Bereitstellung seiner PARP-1/2 siRNA Sequenzen und den dazugehörigen Protokollen. Christoph Paone vom Lehrstuhl Prof. Dr. Christof Hauck danke ich für die Bereit- stellung der BL21(DE3) Bakterien. Dr. Tanja Waldmann danke ich für die Bereitstellung von H3K4me4 Antikörpern und für ihre Infos zur DEK Biochemie.
Im Laufe meiner Promotion hatte ich die Freude, die Abschlussarbeiten von sehr motivierten Studenten zu betreuen. Janine, Christina, Lydia, Veronika, Josefine und Leo danke ich herzlich für ihr Interesse an meinem Thema und für ihre gute Arbeit.
Ein ganz herzliches Dankeschön geht an Kathrin, Chris und Nadine. Danke, dass ihr euch die Zeit genommen habt, Teile dieser Arbeit Korrektur zu lesen.
Ein besonderer Dank geht an meine Arbeitsgruppe: Ihr seid die besten Kollegen die man sich wünschen kann. Vielen Dank für die super Stimmung auf der Arbeit -die maßgeblich dazu bei- getragen hat, dass ich die letzten vier Jahre immer gerne im Labor gewesen bin-, eure Freund- schaft, die vielen Kaffeepausen, Kuchen, Blödeleien, Diskussionen über Gott und die Welt, Ausflüge, Feiern, Spieleabende, Gin-Verkostungen und so viele andere schöne Momente. Ein ganz großer Dank auch noch mal an Eva, Dani und Nadine: Danke für alle Hilfestellungen, Ge- spräche und offenen Arme in den letzten Jahren.
Ein herzlicher Dank geht auch an meine Kollegen von der AG Bürkle. Danke für die Aufnahme in euren Kreis, die vielen Späße, Grillabende, Feiern und Kinobesuche, die die Promotionszeit so bunt gemacht haben.
Ein großer Dank geht an meine Familie. Auch wenn ihr so manches Mal in den letzten vier Jah- ren nur noch ungläubig die Köpfe schütteln konntet, habt ihr mich doch immer auf meinem Weg bekräftigt und unterstützt. Danke dafür, ihr seid die Besten.
Mein letzter, aber auch größter Dank geht an Jan. Danke dass du (nicht nur) in den letzten vier Jahren alle Höhen und Tiefen mit mir durchfeiert und durchlitten hast.
A Deutzmann, M Ganz, F Schönenberger, J Vervoorts, F Kappes, E Ferrando-May, (2015): The human oncoprotein and chromatin architectural factor DEK counteracts DNA replication stress. Oncogene ; 2015 34(32):4270-7. doi: 10.1038/onc.2014.346.
A Breckner, M Ganz, D Marcellin, J Richter, N Gerwin, M Rausch (2013): Effect of Calstabin1 depletion on calcium transients and energy utilization in muscle fibers and treatment opportunities with RyR1 stabilizers.
PLoS One. 2013 Nov 26;8(11):e81277. doi: 10.1371/journal.pone.0081277.
C Strasser, P Grote, K Schäuble, M Ganz, E Ferrando-May (2012): Regulation of nu- clear envelope permeability in cell death and survival Nucleus ; 2012 Nov- Dec; 3(6):
540-51. doi: 10.4161/nucl.21982.
L Rank, S Veith, E Gwosch, J Demgenski, M Ganz, M Jongmans, C Vogel, A Fisch- bach, S Bürger, A Stier, JM Fischer, C Renner, M Schmalz, S Beneke, M Groettrup, R Kuiper, A Bürkle, E Ferrando-May, A Mangerich: Structure-function relationships of natural and artificial PARP1 variants in reconstituted HeLa PARP1knock-out cells.
Submitted in Nucleic Acids Research.
Poster Presentations
• KoRS-CB Retreat – Gültstein, Germany 2015
• SFB969 Review – Konstanz, Germany 2015
• Two days of Proteostasis in Konstanz – Konstanz, Germany 2014
• DNA Replication as a Source of DNA Damage, Fusion Conferences – El Jadida, Morocco 2014
• KoRS-CB Retreat – Bad Herrenalb, Germany 2014
• KoRS-CB Retreat – Gültstein, Germany 2013
• Two days of Proteostasis in Konstanz – Konstanz, Germany 2012
• KoRS-CB Retreat – Gültstein, Germany 2012
• 6th PARP Regio Meeting 2012 – Aachen, Germany 2012
• RRR (Recombination, Replication and Repair) Meeting – Zurich, Switzerland 2015
• SFB969 Retreat – Bräunlingen, Germany 2013
• KoRS- CB Student Seminar – Konstanz, Germany 2013/2014
Fellowships
• 2012-2016: Fellow of the Konstanz Research School Chemical Biology (KoRS-CB)
• 2012-2015: Member of the Sonderforschungsbereich 969 (SFB969)
Attended Courses of the Graduate School Chemical Biology (KoRS-CB)
• Gene Expression and Protein Purification Strategies.
• Biomedicine: Flow Cytometry and Fluorescence Activated Cell Sorting.
• Proteomics
• Advanced Bioimaging
• Understanding Peer-reviewed Publishing
Additional Courses attended at the University of Konstanz
• ‘Laboratory Animal Sciences’ after guidelines of FELASA-Cat. B
• Introduction into Safety Difficulties of Genetic Engineering (Approved training ac- cording to the German § 15 GenTSV)
1 Zusammenfassung ... 1
2 Summary ... 3
3 Introduction ... 5
3.1 Overview over Eukaryotic DNA Replication ... 5
3.2 DNA Replication Stress ... 6
3.2.1 Sources of DNA Replication Stress ... 6
3.2.2 Checkpoint Signaling at Challenged Replication Forks... 9
3.2.3 Pathways of Replication Fork Rescue and Restart ... 11
3.2.3.1 Direct Restart of Replication at Stalled Forks ... 12
3.2.3.2 DNA Damage Tolerance Pathways at Impaired Replication Forks (DDT) ... 12
3.2.3.3 Replication Fork Reversal as an Emerging Way to Rescue Impaired Replication Forks .. ... 13
3.2.3.4 Rescue of Stalled Replication Forks Using the Fanconia Anemia Pathway ... 15
3.2.4 Replication Stress and Cancer ... 16
3.3 The Human Oncoprotein DEK ... 17
3.3.1 Structure of DEK ... 18
3.3.2 Posttranslational Modifications of DEK ... 19
3.4 Molecular Functions of DEK ... 20
3.4.1 DEK Functions as Chromatin Architectural Factor and Histone Chaperone ... 20
3.4.2 DEK’s Function in Transcriptional Regulation and mRNA Processing ... 22
3.4.3 DEK Impacts on DNA Repair, Replication and Genomic Stability ... 23
3.4.4 The Implication of DEK in Cancer ... 24
3.5 Poly(ADP-ribose) Polymerases and Poly(ADP-ribosyl)ation ... 26
3.5.1 Poly(ADP-ribosyl)ation ... 26
3.5.2 PAR-Writers: PARP-1 and PARP-2 ... 28
3.5.2.1 Structure of Poly(ADP-ribose) Polymerase-1 (PARP-1) ... 29
3.5.2.2 Structure of Poly(ADP-ribose) Polymerase-2 (PARP-2) ... 30
3.5.3 Non-Covalent Modification of Proteins with PAR ... 30
3.5.4 PARP’s Involvement in DNA Repair, Chromatin Modulation, Replication and the Replication Stress Response ... 31
4 Aim ... 35
5 Material and Methods... 36
5.1 Material ... 36
5.1.1 Bacterial Strains and Cell Lines ... 36
5.1.1.1 Bacterial Strains... 36
5.1.1.2 Eukaryotic Cell Lines ... 36
5.1.2 Plasmids and DNA ... 36
5.1.4 Sequences of shRNAs, siRNAs and TALENs ... 37
5.1.5 Primer Sequences ... 37
5.1.6 Culture Media and Supplements ... 37
5.1.7 Antibiotics ... 37
5.1.8 Thymidine Analogues ... 38
5.1.9 Transfection Reagents ... 38
5.1.10 Dyes ... 38
5.1.11 Antibodies ... 38
5.1.12 Mounting Media ... 39
5.1.13 Loading Dyes ... 39
5.1.14 PCR Reagents ... 39
5.1.15 Ladders and Standards ... 39
5.1.16 Kits ... 39
5.1.17 Chemicals ... 39
5.1.18 Equipment ... 41
5.2 Methods ... 43
5.2.1 Cell Culture and Cell–based Assays ... 43
5.2.1.1 Cell Culture ... 43
5.2.1.2 Culturing and Plating of Cells ... 43
5.2.1.3 Cryoconservation of Cells ... 43
5.2.1.4 Thawing of Cells ... 43
5.2.1.5 Generation of U2-OS DEK Knockdown Cells ... 43
5.2.1.6 Establishment of a U2-OS DEK Knockout Cell Line ... 44
5.2.1.7 Downregulation of Cellular PARP-1 Levels Using siRNA ... 44
5.2.1.8 Transfection of U2-OS Cells Using Lipofectamine 3000 ... 45
5.2.1.9 Flow Cytometry and Cell Sorting ... 45
5.2.1.10 Clonogenic Survival Assay ... 45
5.2.1.11 Micronucleus Assay ... 45
5.2.1.12 Sister Chromatid Exchange Assay ... 46
5.2.1.13 DNA Fiber Assay ... 46
5.2.2 Immunocytochemistry and Microscopy ... 47
5.2.2.1 Immunofluorescence Detection of DEK, 53BP1, Cyclin A, FancD2, γH2AX, PARP-1 and α- Tubulin ... 47
5.2.2.2 Immunofluorescence Detection of RPA70 ... 48
5.2.2.3 Immunofluorescence Detection of Rad51 ... 49
5.2.2.4 Immunofluorescence Detection of G-Quadruplexes ... 49
5.2.2.5 Immunofluorescence Detection of Incorporated EdU in Replicating Cells ... 50
5.2.2.6 Confocal Laser Scanning Microscopy ... 50
5.2.2.7 Determination of DNA Lesion Foci ... 50
5.2.2.8 Immunofluorescence Detection of H3K9me3 and H3K4me3 ... 50
5.2.2.9 Epi-fluorescence Microscopy ... 51
5.2.3 Molecular Biology and Biochemistry ... 51
5.2.3.1 Isolation of Genomic DNA from Eukaryotic Cells ... 51
5.2.3.2 Amplification of the DEK Genomic Sequence Using PCR ... 51
5.2.3.3 Agarose Gel Electrophoresis ... 51
5.2.3.4 Quick Change Site-directed Mutagenesis ... 52
5.2.3.6 Expression and Purification of Recombinant GST-Tagged Proteins from E.Coli ... 52
5.2.3.7 Hot SDS Lysis ... 53
5.2.3.8 Cold NP-40 Lysis (Westernblot Analysis of PCNA modification) ... 53
5.2.3.9 Determination of Protein Yield ... 53
5.2.3.10 SDS-PAGE and Western Blotting ... 54
5.2.3.11 In-vitro Synthesis of Poly(ADP-ribose)... 55
5.2.3.12 PAR Overlay Assay ... 55
5.2.3.13 Electromobility Shift Assay (EMSA) ... 56
6 Results ... 57
6.1 Establishment of DEK Knockout Cell Lines Using TALEN Technology ... 57
6.1.1 DEK Knockout Cell Lines Do Not Display a Uniform Phenotype ... 59
6.2 The Choice of PARP Inhibitor is Crucial for the Analysis of Replication Stress- Associated DNA Damage ... 61
6.2.1 AZD-2281 Induces High Levels of 53BP1 Foci in a Time-Dependent Manner ... 61
6.2.2 ABT-888 Does Not Induce Enhanced 53BP1 Foci Formation ... 65
6.3 Analysis of the Functional Role of DEK and PARP-1/2 in Response to DNA Replication Stress ... 67
6.3.1 DEK and PARP-1 Are Spatially Separated in Interphase Nuclei ... 67
6.3.2 The Localization of DEK with Respect to Sites of DNA Damage Is Not Regulated by PAR ... 68
6.3.3 DEK and PARP-1/2 Activity Influence Fork Progression after Replication Stress Induction . 69 6.3.4 Stalled Replication Forks Are Reactivated with Different Efficiency, Depending on PARP-1/2 Activity and DEK ... 73
6.3.5 DEK and PARP-1/2 Activity Influence the Formation of DNA Double Strand Breaks ... 75
6.3.6 DEK and PAR Impact on the Accumulation of Rad51, FancD2 and RPA Foci Arising from Perturbed DNA Replication ... 78
6.3.7 PARP-1 is Important for the Formation of 53BP1-OPT Domains in APH-Treated G1-Phase Cells ... 83
6.3.8 The Influence of DEK and PARP-1/2 Activity on Genomic Stability and Translesion Synthesis ... 84
6.3.9 DEK Level Influence the Formation of G4-DNA ... 87
6.4 Biochemical Analysis of DEK’s Non-Covalent Modification with PAR ... 88
6.4.1 Basic Amino Acids in DEK’s Central PBM Are Important for PAR-Binding ... 88
6.4.2 The DEK_PAR Mut Protein Does Not Rescue RPA Foci Formation ... 91
7 Discussion ... 93
7.1 Analysis of the TALEN-Mediated DEK Knockout ... 93
7.2 PARP-1/2 Inhibitors and Their Influence on DNA Damage and Replication Stress . 95 7.3 The Impact of DEK on the Cellular Replication Stress Response ... 96
7.3.1 DEK’s Impact on Replication Fork Progression in Response to DNA Replication Stress ... 97
7.3.2 DEK’s Effect on the Formation of DNA Lesion Foci in Cells ... 98
7.3.3 The Effect of DEK on Translesion Synthesis ... 102
7.3.4 The Formation of G4-DNA is Modulated by DEK Expression Levels ... 103
7.4.1 Subcellular Localization of DEK and PARP-1 ... 104
7.4.2 The Influence of DEK and PARP-1/2 Activity on Fork Progression, Restart of Stalled Replication Forks and Genomic Instability ... 106
7.4.3 The Influence of DEK and PARP-1/2 on the Formation of DNA Damage Foci ... 109
7.4.4 The Influence of PARP-1 on the Formation of 53BP1-OPT Domains ... 112
7.4.5 The Influence of DEK and PAR on Translesion Synthesis ... 113
7.5 Analysis DEK PAR-Binding Mutants ... 113
7.6 Conclusion ... 116
8 Bibliography... 117
9 Abbreviations ... 138
10 Index ... 142
Figures ... 142
Tables ... 143
11 Record of Contributions ... 144
1 Zusammenfassung
Das Onkoprotein DEK ist in den meisten Eukaryoten hoch konserviert und ist ein abundanter Bestandteil des Chromatins. Die bisher wichtigste identifizierte Funktion von DEK ist seine topologische Aktivität, durch die es die Chromatinstruktur nachhaltig beeinflussen kann. In vitro bindet DEK bevorzugt an nicht-kanonische DNA Strukturen (z.B. Vier-Wege Gabelungen), welche in Zellen auftreten können wenn die Replikation oder DNA Reparatur gestört ist. DEK wird in einer Vielzahl von Neoplasmen überex- primiert und es konnte gezeigt werden, dass die Überexpression von DEK die schädli- chen Folgen von Replikationsstress minimiert und so zur Transformation von Zellen beitragen könnte.
Viele Aminosäuren innerhalb der DEK-Sequenz werden posttranslational modifiziert und DEK interagiert sowohl kovalent als auch nicht-kovalent mit Poly(ADP-ribose) (PAR). Diese Modifikationen können sowohl die Funktion, als auch die Lokalisation von DEK beeinflussen. Ebenso wie DEK kann auch Poly(ADP-ribose) Polymerase-1 (PARP-1) mit nicht-kanonischen DNA Strukturen interagieren und wird zudem durch diese aktiviert. Verschiedene Studien zeigen, dass die Aktivierung von PARP-1 einen wichtigen funktionellen Schritt in der zellulären Reaktion auf Replikationsstress dar- stellt.
Erste Replikationsstress Experimente ließen vermuten, dass die Funktion von DEK durch PARP-1/2 moduliert wird. Ziel dieser Arbeit war es daher, die Funktion von DEK im Replikationsstress in Abhängigkeit der Aktivität von PARP-1/2 zu beleuchten und die nicht-kovalente Modifizierung von DEK mit PAR biochemisch näher zu charakteri- sieren.
Im Zuge dieser Arbeit konnte durch immunfluoreszenz-basierte Replikationsanalysen gezeigt werden, dass DEK das Voranschreiten von Replikationsgabeln unter verschie- denen Replikationsstress-Bedingungen unterstützt. Immunzytochemische Charakteri- sierungen ergaben, dass DEK die Anhäufung von DNA Läsionen und die Akkumulation von Reparaturmarkern während topologischem Stress vermindert und womöglich an der Regulation von Replikationsintermediaten beteiligt ist. Zudem unterdrückt die Ex- pression von DEK die Akkumulation von G4-DNA Sekundärstrukturen in hyperprolifera- tiven Zellen. In Zellen mit verringertem DEK Level führte die Inhibition von PARP-1/2 zu auffälligen Veränderungen in der DEK-abhängigen zellulären Antwort auf Replikati- onsstress: Replikationsgabeln wurden beschleunigt, die Menge an auftretenden DSBs verringert und die Akkumulation von Gabel-modulierenden Faktoren verstärkt. Zudem lassen Western-Blot Analysen vermuten, dass sowohl DEK als auch PARP-1/2 die Initiation der Transläsionssynthese unterstützen. In Zuge dieser Arbeit ergab eine Eva- luation von PARP-1/2 Inhibitoren, dass Inhibitor-spezifische off-target Effekte die fokale Akkumulation von Stress-assoziierten Reparaturmarkern auslösen können und damit die Evaluation von Replikationsstress-spezifischen Effekten erschweren.
Schlussendlich konnte durch die biochemische Analyse von DEK-Peptiden gezeigt werden, dass basische Aminosäuren in DEKs zentralem PAR-Bindemotiv die Interakti-
on von DEK mit PAR ermöglichen und in vitro einen großen Anteil der DEK- spezifischen Interaktion mit PAR vermitteln.
Zusammenfassend erweitert diese Arbeit die Erkenntnisse über die Funktion von DEK in der zellulären Reaktion auf Replikationsstress und gibt erste Hinweise für eine Regu- lation von DNA Sekundärstrukturen in Krebszellen durch DEK. Die Experimente in die- ser Arbeit zeigen zudem, dass es einen funktionalen Zusammenhang zwischen DEK Expression und PARP-1/2 Aktivität gibt, welcher sich in der zellulären Antwort auf DNA Torsionsstress manifestiert.
2 Summary
DEK is a widely expressed, non-histone chromosomal protein that displays high affinity for recombinogenic and distorted DNA structures and regulates chromatin architecture by influencing DNA topology and folding. Depletion of DEK results in the accumulation of DNA damage induced by various genotoxic agents, including DNA replication inhibi- tors. Recently, research unraveled a novel link between DEK and the response to DNA replication stress, proposing a role for this protein in the protection from replication- associated DNA damage in hyperproliferative cells.
DEK is strongly posttranslationally modified by phosphorylation, acetylation and cova- lent as well as non-covalent poly(ADP-ribosyl)ation (PARylation). These posttransla- tional modifications collectively play a crucial role in regulating DEK’s affinity to DNA, its subcellular localization and the choice of protein interaction partners. Similar to DEK, Poly(ADP-ribose) Polymerase-1 (PARP-1) has been reported to bind to cruciform DNA structures and has recently been shown to be essential for the recovery from fork stalling under conditions of mild DNA replication stress. In combination with preliminary results, showing that PARP-1/2 activity modulates DEK’s function in response to mild DNA replication stress, these findings suggested a corporate role for DEK and PARP- 1/2 in the handling of DNA replication problems.
This study aimed at understanding the biological consequences of a concerted role for DEK and PARP-1/2 in the context of the replication stress response. Furthermore, for a better understanding of how PARP-1/2 interacts with DEK, the non-covalent modifica- tion of DEK with poly(ADP-ribose) (PAR) was closer characterized.
Fiber assays performed in this thesis revealed that DEK facilitates replication fork pro- gression in response to different genotoxic agents. Using immunocytochemical ap- proaches, DEK was shown to attenuate the accumulation of CPT-induced DNA lesions, which could be related to the regulation of replication intermediates. Additionally, it was demonstrated that DEK levels influence the formation of G4-DNA in hyperproliferative cells. In comparison to DEK depletion alone, the concomitant inhibition of PARP-1/2 activity was shown to severely alter the DEK-dependent cellular response to CPT- induced replication stress, leading to an acceleration of replication forks, limitation of DSBs as well as an increased accumulation of lesion markers, which are possibly as- sociated with fork remodeling and/or protection of nascent DNA. Furthermore, western blot experiments imply that DEK as well as PARP-1/2 activity might support translesion synthesis. In this context, the evaluation of PARP-1/2 inhibitors revealed that inhibitor- specific off-target effects can induce the recruitment of DNA lesion marker into nuclear foci, thereby challenging the analysis of the cellular response to replication stress.
Finally, by analyzing the non-covalent modification of DEK with PAR on a biochemical level, basic amino acids were identified to mediate this posttranslational modification within DEK’s central PAR-binding motif. Furthermore, these amino acids were shown to mediate a great amount of the overall PAR-binding affinity of DEK in vitro.
In sum, data obtained in the course of this thesis extend the previously reported obser- vations about DEK’s function in the response to DNA replication stress and provide first
evidence for a DEK-dependent regulation of DNA secondary structures in cells. Fur- thermore, experiments indicate that active PARP-1/2 is an important modulator, influ- encing the DEK-dependent cellular response to DNA replication stress, pointing to- wards a functional regulation between these two proteins.
3 Introduction
3.1 Overview over Eukaryotic DNA Replication
DNA replication is a fundamental process, allowing duplication of DNA and passing on of genetic information onto daughter cells. The duplication of the genome requires a well regulated network of cellular factors forming and coordinating the replication ma- chinery, which is called “replisome”. Replication is accomplished at a mean speed of about 3000 bp/min and is highly precise, preventing the establishment of mutations. In general, the DNA double helix needs to be unwound and separated into two single pa- rental strands. These serve as replication templates and allow the formation of the typi- cal three-way junction replication fork. This is a prerequisite to allow the binding of DNA polymerases, which execute DNA synthesis in 5’-3’ direction via the pairing and incor- poration of nucleotides at the template strand. As DNA replication at the same time synthesizes two new DNA strands with antiparallel orientation, DNA synthesis is con- tinuous on one strand (leading strand) and discontinuous on the other, forming okazaki fragments that have to be subsequently ligated by Ligase I (lagging strand) (Leman and Noguchi, 2013).
Replication is restricted to the S-phase of the cell cycle and DNA is only copied once, preventing detrimental gene amplifications. Cell cycle progression is regulated by pro- teins of the CDK family (cyclin-dependent kinases), which are activated by defined cy- clin proteins whose expression is tightly regulated in time. The association of CDK4/Cyclin D during G1-phase leads to the activation of transcription factor E2F, the production of replication- and cell cycle-associated proteins and the initiation of origin licensing. The following association of CDK2 with Cyclin E initiates the G1/S-phase transition and induces origin firing. Subsequently, CDK2 associates with Cyclin A, driv- ing S-phase progression, release of pre-replication complex proteins, loading of PCNA and activation of ligase activity (Coverley et al., 2002; Suryadinata et al., 2010)
DNA synthesis is initiated at well-defined places within the DNA, which are called “rep- lication origins”. At this places the replication factories are pre-assembled, a process referred to as “licensing”. This allows origins to be fired upon cellular signaling and S-phase entry. The core of the helicase, which is required for the unwinding of the pa- rental DNA strands, is the MCM2-7 complex (mini-chromosome maintenance proteins).
The recruitment and loading of the MCM2-7 double hexamer requires a number of co- factors of the ORC-CDC6 complex. Together these proteins form the pre-replication complex (pre-RC) (Evrin et al., 2009; Leman and Noguchi, 2013; Remus et al., 2009).
The activation of MCM2-7 helicase and subsequent origin firing requires the CDK- dependent recruitment of additional factors, such as Cdc45 and the GINS complex, which stabilize MCM2-7 at DNA. Together the proteins resemble the pre-initiation com- plex and the active replicative helicase. Cdc45 is also required for the recruitment of replicative DNA polymerases α and δ. Furthermore, Cdc45 and GINS are involved in helicase/ polymerases coupling, keeping their activity in close proximity. This is sup- posed to be accomplished by regulating helicase activity and providing a physical link between helicase and polymerase (Costa et al., 2011; Leman and Noguchi, 2013;
Mimura and Takisawa, 1998). Upon assembly, the active helicase starts to unwind DNA, exposing stretches of ssDNA (single-stranded DNA). These are coated by single strand-binding protein RPA (replication protein A), which supports the binding and ac- tivity of polymerase α/primase and initiation of replication. The polymerase α/primase complex is able to initiate DNA synthesis de novo as the primase synthesizes a short ribonucleotide patch, providing a 3’-OH end that can be extended by Polymerase α (Hübscher et al., 2002; Leman and Noguchi, 2013; Walter and Newport, 2000). Full genome replication is accomplished by polymerase δ on the lagging strand and poly- merase ε on the leading strand, substituting polymerase α after replication initiation, as they possess higher processivity. Polymerases are supported by the cofactor PCNA (proliferating-cell-nuclear-antigen), which strengthens the interaction of polymerases with DNA and stabilizes the replication complex. The loading of PCNA onto DNA at primer template junctions requires the help of clamp loaders by the RFC-complex (rep- lication factor C), which is the canonical loading complex during replication and also initiates polymerase switching (Hübscher et al., 2002; Leman and Noguchi, 2013;
Maga et al., 2000; Mossi et al., 2000). Due to different posttranslational modifications, PCNA was found to coordinate a variety of different processes at the replication fork, also controlling the DNA damage response during DNA replication (Zhang et al., 2011a).
3.2 DNA Replication Stress
Albeit a lot of research is focusing on the cellular response to replication stress the term itself is not yet definitely defined. It is generally used to describe any circumstanc- es that lead to a slowdown or even stall of an active replication fork, thereby impacting DNA synthesis. Replication stress can be induced by a variety of endogenous as well as exogenous sources that form barriers to normal replication fork progression. As ac- curate and timely genome duplication is important for the maintenance of genome sta- bility and cell survival, replication stress is a challenge for cells and requires a coordi- nated cellular response.
3.2.1 Sources of DNA Replication Stress
The most recognized source for replication stress is the encounter of unrepaired DNA lesions by the replisome, prohibiting further progression of the replication fork. This includes, but is not limited to, DNA nicks, bulky adducts and interstrand crosslinks (Zeman and Cimprich, 2014). A further trigger for replication stress can be the miss- incorporation of ribonucleotides, which stall the replicative polymerases and are detri- mental for cell survival when not removed (Nick McElhinny et al., 2010).
Faithful replication requires a great number of different molecules. When these are limited replication progression is slowed down, leading to the induction of replication stress. This applies to components of the replication machinery, histones, and dNTPs.
Hydroxyurea (HU) is an inhibitor of the ribonucleotide reductase, which is required for the reduction of ribonucleotides to deoxyribonucleotides. HU treatment in eukaryotes leads to a partial depletion of nucleotide pools and to an immediately reduced or im-
paired replication elongation and changes in origin firing (Alvino et al., 2007; Ge and Blow, 2010; Poli et al., 2012; Zellweger et al., 2015).
Aphidicolin (APH) is an inhibitor of replicative polymerases. APH treatment was shown to slow down replication and to induce DNA damage at special DNA regions (Glover et al., 1984). Within the genome there are various sites at which the progression of repli- cation forks is challenged, especially upon reduction of replication efficiency. These regions are called fragile sites. Common fragile sites (CFS) preferentially harbor A-T- rich repetitive DNA sequences that can form extensive secondary structures and are replicated in late S-phase. Due to low numbers of origins and late replication timing, a slowdown or stall of forks is a great challenge within these regions and promotes the formation of under-replicated DNA. This is converted into double strand breaks (DSBs) during chromosome condensation in subsequent mitosis. DSBs are proposed to be induced by mitotic nucleases, trying to resolve DNA damage prior to segregation- dependent chromosomal breakage (Franchitto, 2013; Glover et al., 1984; Naim et al., 2013; Ying et al., 2013).
Transcription and replication in general are carefully spatially and temporally separated (Wei et al., 1998). However, there is emerging evidence that transcription machineries can collide with replication machineries, slowing down replication and inducing the for- mation of DSBs by so far unknown reasons. This especially happens in early replicat- ing regions, where transcription is not fully terminated until onset of S-phase and which are located in close proximity of replication origins. Therefore, these regions are termed early replicating fragile sites (ERFS). The probability of collision is enhanced when the transcription process is slowed down (Mortusewicz et al., 2013). Besides the risk of collision, simultaneous transcription and replication can induce torsional stress. Espe- cially upon head-on encounters, positive supercoiling ahead of both machineries is a great topological impediment. This stress is reported to be further enhanced by the linkage of transcripts and their matrices to nuclear pores (Bermejo et al., 2011;
Bermejo et al., 2012). When a replication fork encounters a transcription site, the nas- cent RNA can re-hybridize to the open DNA strand within the replication bubble. This risk is especially enhanced when mRNA (messenger RNA) processing is impaired. Re- hybridization leads to the formation of a three-stranded nucleic acid structure, which is called R-loop, and comprises a DNA:RNA hybrid and a displaced a single stranded DNA. R-loop formation was shown to be favored by negatively supercoiled DNA accu- mulations behind the transcription site. R-loops were associated with various cellular processes, such as transcription termination. The persistence of these structures is a challenge to genomic integrity and was shown to induce replication fork slowing in bac- teria as well as human cells (Bermejo et al., 2012; Gan et al., 2011; Santos-Pereira and Aguilera, 2015).
Also the formation of other DNA secondary structures was proposed to challenge repli- cation fork progression. G-quadruplexes (G4-DNA) can form as intra- or intermolecular folds at G-rich sequences. Guanines anneal to stacks of G-tetrads and are stabilized via Hoogsteen base paring (Rhodes and Lipps, 2015). In vitro, a multitude of sequenc- es were identified to potentially fold into G4-DNA. These sequences preferentially colo- calize with functional genomic regions, such as telomeres or replication origins (Lipps and Rhodes, 2009). In vivo, G4-DNA structures were identified using immunocyto-
chemistry and immunoprecipitation and G4-DNA has been shown to possess important regulatory functions in replication, gene expression and telomere maintenance. How- ever, G-quadruplexes are also proposed to form at stretches of ssDNA and need to be removed in order to assure faithful replication fork progression (Henderson et al., 2014;
Huppert and Balasubramanian, 2007; Rhodes and Lipps, 2015).
DNA replication is closely linked to topological changes within the DNA. As DNA is ro- tating, supercoils are generated directly in front of the fork due to DNA over-winding (positive supercoils) as well as behind the fork by under-winding (negative supercoils).
Additionally, freshly synthesized sister chromatids can intertwine due to polymer- ase/helicase rotation forming precatenates, but releasing positive stress ahead of the fork. However, accumulation of positive supercoils leads to a block of replication fork progression and unresolved intertwining prevents the separation of sister chromatids during mitosis and induces genomic instability (Postow et al., 2001; Schalbetter et al., 2015). Relaxation of this torsional constrains is achieved by topoisomerases, which nick DNA and allow rotation of DNA strands. During replication, topoisomerase I (Topo I) is proposed to mainly act in front of the fork while topoisomerase II (Topo II) is sup- posed to mainly act behind the fork and resolves precatenates. However, this classifi- cation is not exclusive. An inhibition of topoisomerases via camptothecin (CPT) or etoposide (ETO), inhibiting Topo I or Topo II, respectively, can be used for the extrinsic induction of replication stress. These agents trap topoisomerases on cleaved DNA, which can be converted into DNA DSBs during replication. Topo I was also shown to be important to limit the formation of R-loops. This is likely due to the ability of topoiso- merases to relieve negative supercoils, which can facilitate re-annealing between the nascent transcript and template DNA strand, or defects in mRNA processing (McClendon et al., 2005; Pommier, 2006; Tuduri et al., 2009).
Figure 3.1 Sources of DNA replication stress
Various obstacles can challenge replication fork progression and consequently induce replication stress.
These include repetitive sequences, DNA secondary structures, DNA lesions, transcription complexes, DNA:RNA hybrids, miss-incorporation of ribonucleotides, as well as limitations in dNTP supply or polymer- ase fidelity. Replication stress can be chemically induced by aphidicolin (APH), camptothecin (CPT), etoposide (ETO) or hydroxyurea (HU). FS: fragile site. Figure is based on (Zeman and Cimprich, 2014).
Just recently, the overexpression or constitutive activation of oncogenes was reported as a source for replication stress, as shown for Cyclin E and c-Myc. Activation of these leads to deregulation of origin firing, exhaustion of cellular nucleotide pools, impaired replication fork progression and the formation of under-replicated DNA (Dominguez- Sola et al., 2007; Ekholm-Reed et al., 2004; Jones et al., 2013; Srinivasan et al., 2013).
Also a deregulation of origin firing in non-cancerous cells was shown to lead to an ex- haustion of dNTPs (Ge et al., 2007).
3.2.2 Checkpoint Signaling at Challenged Replication Forks
To limit the tremendous danger of replication stress and to deal with the constant chal- lenges of replication-encountered DNA lesions, a tightly regulated cellular DNA dam- age response (DDR) is required. This coordinates cell cycle checkpoints as well as lesion repair and determines cellular fate. However, the underlying molecular mecha- nisms of this response are only poorly understood.
During the last years new methods (isolation of proteins on nascent DNA [iPOND], nascent chromatin capture analysis [NCC]) were developed to identify proteins that function at normal and disturbed replication forks (Alabert et al., 2014; Sirbu et al., 2011).
There is consensus among different publications that most sources of exogenous repli- cation stress (including treatment with CPT, HU and APH) lead to the formation of ssDNA, mainly due to helicase-polymerase uncoupling. The extending stretches of ssDNA are bound by RPA and are believed to serve as activator for the cellular stress response, which is mediated via ataxia telangiectasia mutated (ATM) and, more promi- nently in replication stress, ATM- and Rad3-related (ATR) kinases (Allen et al., 2011;
Byun et al., 2005; Zellweger et al., 2015). Single-strand bound RPA is recognized by the ATR-interacting protein (ATRIP), which is associated with ATR. This way the checkpoint kinase is targeted to sites of ssDNA, leading to its auto-activation (Liu et al., 2011; Zou and Elledge, 2003). Subsequently, RPA recruits Rad17, which in turn loads the 9-1-1 clamp onto DNA (Delacroix et al., 2007; Zou et al., 2003). As in normal repli- cation, the clamp is required for the further recruitment and binding of proteins to DNA.
9-1-1 interacts with TOPBP1 (topoisomerase II binding protein 1), targeting it to the replication fork (Delacroix et al., 2007). TOPBP1 acts as an additional activator of ATR and enhances its substrate recognition (Kumagai et al., 2006; Liu et al., 2011). Interest- ingly, ssDNA was also found to accumulate in unstressed early S-phase cells, implying that during this stage of replication cells encounter high amounts of intrinsic replication stress (Buisson et al., 2015).
ATR phosphorylates a wide range of downstream proteins. This includes the phos- phorylation of histone H2AX (γH2AX), which is a general marker for DNA lesions within cells. The phosphorylation allows for the recruitment of MDC1 which acts as a scaffold, recruiting further ATR molecules and extends H2AX phosphorylation over long dis- tances (Ward and Chen, 2001). The modification of H2AX already occurs upon transi- ent stalling of replication forks (e.g. via HU or CPT) and in the absence of DNA double strand breaks. With prolonged fork stalling, the modification spreads in a time depend- ent manner over great parts of the genome (Sirbu et al., 2011; Zellweger et al., 2015).
Moreover, MDC1 is important for the recruitment of additional repair proteins, such as 53BP1 (p53-binding protein 1), to sites of DNA lesions and the formation of DNA dam- age foci (Stewart et al., 2003). A further target of ATR is RPA, which are also targeted by other kinases like DNA-PK (DNA-dependent protein kinase). These phosphoryla- tions are believed to contribute to the stabilization of stalled forks, cell cycle inhibition and preparation of fork restart. Phospho-RPA (S33) was shown to stimulate DNA syn- thesis under conditions of HU-induced replication stress, alleviating consequences of replication stress and increasing cell survival in late S-phase (Liu et al., 2012b; Murphy et al., 2014; Vassin et al., 2009). RPA also forms an interaction site for Timeless/Tipin, which in turn recruit claspin. Phosphorylation of claspin by ATR provides an interaction site with Chk1 (checkpoint kinase 1), thereby bringing it in close proximity to ATR. This results in the phosphorylation and activation of Chk1, the main target of ATR and regu- lator of the intra S-phase cell cycle checkpoint. The interaction of Timeless/Tipin and Chk1 with the ATR complex is only transient, allowing phosphorylation of a high num- ber of Chk1 proteins. Activation of Chk1 leads to the inhibition of cell cycle progression, which is mainly mediated via the inhibition of CDK-cyclin complexes (Ghosal and Chen, 2013; Jeong et al., 2003; Kemp et al., 2010). Recent research shows that active Chk1 regulates origin firing by redirecting replication to dormant origins of existing replication factories and preventing the activation of new replication machineries, thereby minimiz- ing the formation of under-replicated DNA at regions of HU-stalled forks (Ge and Blow, 2010). Simultaneously, active Chk1 prevents the exhaustion RPA molecules, thereby limiting the formation of ssDNA, which can be converted into DSBs (Toledo et al., 2013). All of this is supposed to contribute to fork stability, prevention of fork collapses as well as the formation of DNA damage and further allows time for lesion repair. Chk1 activity was also linked to the activation of lesion bypass repair, leading to a stress- dependent stabilization of CDC7/ASK and recruitment of Rad18 to DNA (Yamada et al., 2013).
Recent research extended the picture of a strictly cascade-like regulation of ATR and Chk1, proposing a role for both proteins in the tolerance of endogenous replication stress. In unperturbed replication, ATR was shown to limit the formation of ssDNA in highly replicative, early S-phase cells via down-regulation of origin firing and enhance- ment of nucleotide synthesis via Chk1/CDK2 regulation. This saves cells from acquiring large amounts of DNA replication stress and undergoing replication catastrophe. Mid and late S-phase cells with lower replicative potential were shown to be insensitive to a loss of ATR function, as they downregulate origin firing in a DNA-PK/Chk1-dependent rescue backup mechanism before the accumulation of dangerous amounts of ssDNA (Buisson et al., 2015).
The role of ATM in the cellular response to replication stress is controversially dis- cussed. There is evidence that ATM signaling is also important during replication stress conditions, especially for the processing of collapsed forks, which require homologous recombination (HR)-dependent restart. The MRN complex (Mre11, Rad50 and Nbs1) is the major sensor of DNA DSBs and facilitates their repair. MRN in concert with other proteins, like 53BP1, catalyzes the activation of ATM. This leads to the initiation of cell cycle checkpoints and mediates the repair of DSBs. Homology-directed repair requires MRN-dependent initial resection of DNA ends. This is supported by Exo1, BLM and
DNA2, which perform bulk resection. Emerging ssDNA is bound by RPA, which cross- activates ATR-dependent checkpoint signaling. This allows the potentiation of check- point responses and loading of Rad51 filaments, which mediate sister strand invasion and HR repair (Stracker and Petrini, 2011; Trenz et al., 2006). Just recently, ATM has been specifically related to the repair of blocked DSBs as occurring after treatment with etoposide (Álvarez-Quilón et al., 2014). Analysis of impaired replication forks revealed that DSB repair proteins, except Rad51, remain absent from HU-stalled replication forks until 4 hours after treatment. This is followed by a great accumulation of Rad51 and other repair proteins and is supposed to mark fork collapse. The late accumulation of Rad51 was shown to require MRN, while the early did not; implicating that late load- ing is dependent on fork resection (Sirbu et al., 2011). Also, ATM/DNA-PK activity was shown to be important for γH2AX formation and spreading after persistent stalling and subsequent collapse of replication forks by HU (Sirbu et al., 2011). ATM has also been related with fragile site stability. Upon expression of fragile sites, ATM was shown to be recruited to emerging DSBs and to be involved in the activation of Chk1. Here, ATM was shown to act in parallel to ATR activation (Ozeri-Galai et al., 2007). It is proposed that under-replicated DNA from fragile sites is converted into strand breaks during mi- tosis. These lesion sites are sequestered in G1-phase and shielded by 53BP1-OPT domains. The formation of these bodies was shown to be dependent on ATM (Harrigan et al., 2011; Lukas et al., 2011).
Albeit the established pathways of checkpoint activation, it was shown that treatment with mild doses of various genotoxic agents, accompanied by fork slowing and accu- mulation of ssDNA, do not generally lead to an activation of ATR and ATM. This un- couples checkpoint kinase activity from the accumulation of stress-associated damage markers. Analyzing γH2AX positive cells for ssDNA markers revealed only low colocal- ization, further uncoupling ATR signaling from ssDNA accumulation (Zellweger et al., 2015). Reversed forks expose open DNA ends and therefore display potential for ATM activation. Interestingly, single studies report the activation of ATM in response to low- dose CPT and HU upon fork reversal in the absence of DSBs. However, not all agents inducing fork reversal were associated with ATM /ATR activity, uncoupling the for- mation of regressed arms and accumulation of DNA damage markers from checkpoint signaling (Ray Chaudhuri et al., 2012; Zellweger et al., 2015). It still remains elusive which structural features at stalled forks induce checkpoint signaling. Also, forks could be bound and processed by different factors that induce checkpoint signaling as shown for DNA2 and FBH1 (Fugger et al., 2015; Thangavel et al., 2015).
3.2.3 Pathways of Replication Fork Rescue and Restart
Upon encounter of replication stress, replication forks can stall. In this situation forks are arrested but maintain their replicative potential and are able to resume replication as soon as the stress is removed. Upon extensive or prolonged stress replication forks can collapse. This means that forks lose their replication competence as the replisome is either non-functional or destabilized, losing its association with the replication fork.
The stabilization and rescue of forks as well as resumption of replication is essential for cell survival and genome maintenance, as prolonged replication stress and unresolved DNA lesions can lead to genomic alterations that promote the development of diseases
and cancer (Zeman and Cimprich, 2014). However, when replication stress cannot be released there are several pathways to restart replication nevertheless, to prevent ma- jor genetic instability.
3.2.3.1 Direct Restart of Replication at Stalled Forks
Replication is initiated at licensed origins that are the starting points of replication forks.
However, at beginning of S-phase much more licensed origins are available as re- quired for normal replication. Most origins are only fired when the progression of repli- cation is challenged. The activation of dormant origins allows restart of replication downstream of a stalling lesion as well as in response to APH- and HU-induced fork slowing, preventing the formation of under-replicated DNA (Alver et al., 2014; Ge et al., 2007; Ibarra et al., 2008; Woodward et al., 2006).
Re-priming describes a process during which replication skips the region of damaged DNA and is de-novo re-initiated downstream of the blocking lesion, leading to the for- mation of ssDNA gaps. Re-priming is often related to the encounter of UV-induced le- sions, but also to HU treatment, and strictly requires a primase activity as shown for PrimPol in human cell lines. DNA gaps are filled using either specialized bypass poly- merases of the Rad6/Rad18 pathway or a recombination-dependent pathway mediated by Rad51 (Elvers et al., 2011; Karras and Jentsch, 2010; Mourón et al., 2013).
3.2.3.2 DNA Damage Tolerance Pathways at Impaired Replication Forks (DDT)
DNA damage tolerance pathways allow cells to tolerate lesions encountered by the ongoing replication fork. This is accomplished via bypassing of the lesion without ex- tensive fork stalling and a repair of lesions via post replication repair (PRR). The two major pathways of DDT are translesion synthesis (TLS) and template switching (TS).
During TLS, replicative polymerases are exchanged by less specific TLS-polymerases.
This pathway is supposed to be error-prone. For TS, the undamaged sister chromatid is used as template to replicate over the lesion-containing site and is considered error- free (Ghosal and Chen, 2013).
The key regulator of DDT pathways is PCNA, which is posttranslationally modified with ubiquitin upon lesion encounter. Mono-ubiquitination of PCNA induces the recruitment of TLS-polymerases via their ubiquitin-interaction domain, while poly-ubiquitination has been associated with TS (Ghosal and Chen, 2013; Zhang et al., 2011a). The E3- ubiquitin ligase Rad18 was found to be the major mono-ubiquitinating enzyme of PCNA upon HU- and UV-induced DNA damages. The process of poly-ubiquitination is still unclear, but is suggested to be as well mediated by Rad6/Rad18 and additional E3-/E2-ligases. The recruitment of the Rad6/Rad18 complex and subsequent ubiquiti- nation of PCNA was reported to be induced via the formation of RPA-coated ssDNA due to polymerase/helicase uncoupling and Rad18 was shown to directly interact with DNA, RPA and PCNA (Davies et al., 2008; Ghosal and Chen, 2013; Masuda et al., 2012; Watanabe et al., 2004).
In TLS, mono-ubiquitination of PCNA induces the switch from replicative polymerases to TLS-polymerases, which mainly belong to the Y-family of polymerases (Pol η, Pol ι, Pol κ, Rev1). These contain a larger active center and are able to replicate over le- sions. However, they are less productive and exhibit no proof-reading ability. For each
type of lesion there are specific polymerases, exhibiting different error-rates. After rep- lication over the lesion site the synthesized DNA patch is predominantly extended by TLS-polymerases of the B-family (Pol ζ), prior to re-loading of replicative polymerases.
This allows the lesion to escape proof-reading function. Unloading of TLS-polymerases is supposed to be accomplished via PCNA de-ubiquitination, which is mediated by USP1 after UV-lesions, but probably not after chemically-induced damage (Bienko et al., 2005; Ghosal and Chen, 2013; Huang et al., 2006; Kannouche et al., 2004). Mono- ubiquitination of PCNA was also found in response to HU-induced replication stress and was shown to persist over long time periods, even after removal of stress. In this context Pol η was shown to be recruited to replication forks, promoting replication pro- gression and preventing fork break down. However, this was also associated with in- duction of apoptosis at G1/S-transition (Brown et al., 2009; de Feraudy et al., 2007).
Earlier studies showed that TLS-polymerases may also be linked to HRR repair as Rad51 was found to induce the recruitment of Pol η for D-loop extension (McIlwraith et al., 2005).
3.2.3.3 Replication Fork Reversal as an Emerging Way to Rescue Impaired Replication Forks
A newly recognized cellular response to replication stress is the frequent formation of reversed forks. The term ‘replication fork reversal’ describes a process that includes the rewinding of parental DNA strands as well as the annealing of newly synthesized strands, leading to the formation of a 4-way junction. This structure is also referred to as ‘chickenfoot’ (Neelsen and Lopes, 2015).
The formation of these replication intermediates has already been proposed 40 years ago. However, for a long time reversed forks in eukaryotes were only visualized in checkpoint-deficient yeast, questioning their relevance for mammalian replication (Higgins, 1976; Lopes et al., 2001). Just recently, fork slowing and replication fork re- versal were established as a conserved, transient response to a variety of replication stress inducers, including CPT, ETO, HU and APH, in eukaryotes. The accumulation of reversed forks was shown in cancerous as well as non-cancerous cells and at concen- trations of genotoxic agents that do not induce the formation of DSBs (Ray Chaudhuri et al., 2012; Zellweger et al., 2015). Replication fork reversal allows cells to stably pause replication forks and limit genomic instability, due to fork uncoupling, accumula- tion of ssDNA and fork fragility. Fork reversal places a lesion back into a DNA double strand and allows time for excision repair to prevent duplication of damaged or discon- tinuous DNA. Alternatively, it allows time for the activation of damage bypass mecha- nisms (Neelsen and Lopes, 2015). For mild CPT-induced replication stress fork rever- sal was shown to be an important tolerance mechanism, limiting the formation of dou- ble strand breaks (Ray Chaudhuri et al., 2012).
Ongoing research is investigating molecular determinants required for the formation of reversed forks. Rad51, an established HR component, was shown to be a constitutive part of replicating chromatin (Hashimoto et al., 2010; Zellweger et al., 2015). Further- more, Rad51 was found to be enriched at challenged replication forks after CPT- and HU-induced replication stress in the absence of DSBs. This enrichment is mediated via accumulated RPA on ssDNA, a known target for Rad51 binding. However, Rad51 load-
ing is supposed to require additional loading factors, which are supposed to belong to Fanconi Anemia (FA)- or HR-families (Neelsen and Lopes, 2015; Sirbu et al., 2013;
Zellweger et al., 2015). For fork reversal, Rad51 loading and strand invasion was found to be fundamental, driving the conversion of uncoupled, ssDNA-containing forks into reversed structures by parental strand re-annealing (Zellweger et al., 2015). Fork re- gression was shown to be supported by FBH1 helicase activity (F-box DNA helicase 1) after HU treatment. This support is probably mediated by displacement of the lagging strand (Fugger et al., 2015). Furthermore, there is first evidence that FBH1 regulates Rad51 function via ubiquitination (Chu et al., 2015).
Figure 3.2 Replication fork reversal and restart of reversed forks.
Stalled forks can be transiently paused by replication fork reversal. ssDNA, emerging due to hel- icase/polymerase uncoupling, is bound by RPA. Rad51 partially replaces RPA and mediates the reversal of forks by parental strand annealing. FBH1 supports this process by lagging-strand displacement. For the restart of reversed forks different pathways were reported. RecQ1 was shown to bind to reversed forks and mediate their progression via branch migration. RecQ1 activity is inhibited by PARP-1 activity, pre- venting a pre-mature restart of reversed forks. Also unwinding and nucleolytic regression of reversed arms by DNA2 and WRN was reported. It is supposed that resected arms can recruit branch migration factors that mediate the restart of the reversed fork and have yet not been described further. Interestingly, DNA2 activity can be inhibited by RecQ1. It is also possible that reversed forks are restarted by HR-repair.
ssDNA regions on regressed arms promote the binding of Rad51, which can mediate homology- dependent short tract strand invasion and recombination-dependent restart of forks. Figure is based on (Neelsen and Lopes, 2015).
Restart of reversed replication forks requires the restoration of a typical three-way fork.
This was found to be dependent on RecQ1 branch migration activity, which is regulated by poly(ADP-ribose) (PAR) in response to different replication inhibitors (Berti et al., 2013; Ray Chaudhuri et al., 2012; Zellweger et al., 2015). However, it remains unclear how Poly(ADP-ribose) Polymerase-1 (PARP-1) is activated by genotoxic agents with different cellular modes of action. Additionally, an alternative pathway of replication fork restart was discovered, which is mediated via the functional interaction of DNA2 (DNA replication ATP-dependent helicase/nuclease) and WRN (Werner syndrome ATP- dependent helicase). DNA2 was shown to degrade regressed forks with a 5’-3’ activity.
This is supposed to be supported by WRN, detaching the newly synthesized strands.
Yet unknown branch migration factors, distinct from RecQ1, are proposed to recognize the resected fork to mediate their restart. Interestingly, RecQ1 was found to be able to inhibit DNA2 activity (Thangavel et al., 2015). Although having found evidence for the- se mechanisms of fork restart, a recombination-dependent restart of reversed forks by Rad51 is not excluded. Loading of Rad51 at regressed arms is supposed to protect the reversed forks and furthermore to promote short tract homology-dependent repair (Petermann et al., 2010; Thangavel et al., 2015; Zellweger et al., 2015).
There is emerging evidence that fork reversal is also a conserved response to endoge- nous replication stress. In unchallenged cells a small but consistent amount of reversed forks can be detected already (Ray Chaudhuri et al., 2015; Zellweger et al., 2015). Fur- thermore, the inhibition of Poly(ADP-ribose) Glycohydrolase (PARG), and therefore interference with PAR metabolism, leads to the slowdown of replication forks and the accumulation of reversed forks in unstressed cells. This indicates that fork reversal is tightly regulated by PAR and might be a frequent event at endogenous lesions or repli- cation obstacles (Ray Chaudhuri et al., 2015). Also depletion of RecQ1 and DNA2 in unperturbed cells was shown to induce the accumulation of reversed forks, supporting the notion that reversal is a conserved response to intrinsic replication challenges (Thangavel et al., 2015; Zellweger et al., 2015). Additionally, the formation of reversed forks is proposed to occur upon encounter of repetitive DNA sequences, which form extensive secondary structures, linking fork reversal to endogenous difficult-to-replicate structures (Follonier et al., 2013).
3.2.3.4 Rescue of Stalled Replication Forks Using the Fanconia Anemia Pathway Fanconia anemia (FA) is a genetic bone marrow disease, linked to increased sensitivity to interstrand crosslinks and cancer susceptibility. The FA network consists of a large protein cluster and the FA pathway is mainly supposed to be required for the removal of interstrand crosslinks (ICLs) (Walden and Deans, 2014). ICLs are recognized by the FancM (Fanconi anemia complementation group M) anchor complex, which can also identify other DNA structures, including 3-way and 4-way junctions. When activated, this complex induces the formation of the Fanc-core complex, which resembles an E3-ubiquitin ligase complex. This complex mono-ubiquitinates the FancI/FancD2 het- erodimer (ID complex), which is the central component of the FA pathway, leading to its re-localization to damage sites. Furthermore, FancM activity was related to the acti- vation of ATR-mediated checkpoint signaling. Mono-ubiquitination of FancI/FancD2 leads to the recruitment of downstream endonucleases (FancP/FancQ) and repair fac- tors (FancD1, FancN, FancO) that coordinate the incision at both sites of the ICL. As now the lesion is only tethered to one DNA strand it can be bypassed by TLS. The re- sulting DSB from the incision procedure is supposed to be repaired by Rad51-mediated HRR (Kim and D'Andrea, 2012; Knipscheer et al., 2009; Long et al., 2011; Schwab et al., 2010; Walden and Deans, 2014).
The central event of FA pathway is the mono-ubiquitination of FancD2, which is con- sidered to be a marker for network activation. Just recently, it has been shown that Rad6/Rad18 is also able to initiate the activation of the ID complex via PCNA mono- ubiquitination, which recruits FancL to lesion sites (Geng et al., 2010). Furthermore, ubiquitination of FancD2 was shown to mediate an interaction between FancD2 and
Pol η. This interaction targets TLS polymerases to ICL lesion sites, linking FancD2 ac- tivity closer to PRR (Fu et al., 2013). Monoubiquitination of FancD2 and its recruitment to damage sites has also been detected in response to CPT-induced replication stress in human cancer and primary cells. Interestingly, here the modification of FancD2 was shown to be dependent on Rad18 but was independent from PCNA mono- ubiquitination and PRR (Palle and Vaziri, 2011).
Additionally, FancD2 monoubiquitination has also been detected in the cellular re- sponse to HU- and APH-induced replication stress, which do not induce physical DNA lesions (Howlett et al., 2005). In response to APH, FancD2 was shown to localize to gaps and under-replicated DNA at chromosomal fragile sites and to increase genomic stability during chromosome segregation in concert with BLM (Chan et al., 2009; Naim and Rosselli, 2009). Just recently, also a role for FancD2 in PRR has been proposed.
Upon HU-treatment unmodified FancD2 was found to form a complex with Rad18 and Rad51, influencing PCNA monoubiquitination and subsequent chromatin localization of Pol η in response to HU-induced fork collapse. This finding also revealed a further non- canonical function of Rad51, independent of HR repair (Chen et al., 2015).
As many proteins of the FA pathway are involved in HR, a crosstalk between these two pathways is evident. In line with this, BRCA2 (FancD1) and PALB2 (FancN) are both required for Rad51 loading onto ssDNA overhangs. Furthermore, Schlacher and col- leagues showed that monoubiquitinated FancD2 in concert with BRCA2 and Rad51 epistatically stabilize stalled replication forks, protecting them from nucleolytic degrada- tion and fork collapse after HU-induced fork stalling (Schlacher et al., 2011; Schlacher et al., 2012). Just recently, Yang and colleagues extended this findings by showing that monoubiquitinated FancD2 and Rad18 are required to protect nascent replication tracts from CPT-induced DSBs formation and degradation via REV1, as well as for Rad51 stabilization (Yang et al., 2015). Furthermore, a role for FA proteins in transcription- induced replication fork stalling and resolution of DNA:RNA hybrids was revealed. Hy- brid-induced fork stalling was shown to lead to the activation of FancD2 and stabiliza- tion of stalled forks. The removal of R-loops is proposed to be achieved via the trans- locase activity of FancM (Schwab et al., 2015).
All of this highlights the multifunctionality of DNA repair proteins. Besides being associ- ated to special repair pathways, in which their function is essential, there is emerging evidence that a great crosstalk between all pathways exists, underlining the complexity of the cellular response to replication stress.
3.2.4 Replication Stress and Cancer
Cancer cells transform to gain highly proliferative potential, genetic instability and es- cape from apoptosis. This is accompanied by mutations in repair pathways, cell cycle regulators and checkpoint genes. Therefore, cancer cells are highly exposed to replica- tion stress. It has been shown that the activation of oncogenes (e.g. Cyclin E, c-Myc, Ras) is coupled with a shortage of nucleotides, impairment of replication fork progres- sion, accumulation of DNA damage and de-regulation of origin usage. All of this leads to the formation of under-replicated DNA, which is converted into DNA damage during mitosis. Especially chromosomal fragile sites have been associated with the accumula-
tion of oncogene-induced DNA lesions and translocations. Interestingly, fragile sites often encode tumor suppressor genes. However, the exact mechanisms how replica- tion stress promotes cancer progression still remain elusive (Bester et al., 2011; Hills and Diffley, 2014; Jones et al., 2013; Miron et al., 2015; Ozeri-Galai et al., 2012;
Srinivasan et al., 2013).
Macheret and colleagues propose oncogene-induced replication stress to be an im- portant driving force in early cancer development. Mutations in genes that regulate cell growth induce aberrant proliferation, which challenges cellular replication machineries and induces replication stress. This situation is supposed to be encountered in nearly all early cancer cells. Enhanced replication stress leads to the establishment of DNA damage and mutations that further drive cancer development. DNA lesions, however, also activate the DNA damage response driving apoptosis of precancerous cells, thereby forming an important anti-cancer barrier. However, this is believed to put selec- tive pressure on mutations in corresponding DDR genes, allowing cells to escape apoptosis induction. This might allow the transformation of precancerous lesions into invasive cancers (Macheret and Halazonetis, 2015).
Precancerous cells need to tightly regulate their response to replication stress, since the gain of mutations is favorable but can be deleterious when occurring at too high levels, favoring cell death. In many early cancers levels of ATR and Chk1 were shown to be upregulated. Furthermore, the cell cycle regulator Chk1 was shown to be under the direct transcriptional control of c-Myc or E2F, both classified as oncogenes. An ac- tivation of these genes therefore also leads to an overexpression of Chk1. Despite be- ing counterintuitive on a first sight, high levels of these proteins might ensure survival of early cancer cells (Lecona and Fernandez-Capetillo, 2014). Recent research by Neelsen and co-workers associates oncogene activation with the slowdown of replica- tion fork progression and the formation of reversed forks, probably induced by topologi- cal stress due to replication deregulation. If these replication intermediates cannot be removed, e.g. due to checkpoint impairment, nucleolytic processing during mitosis leads to the accumulation of DSBs, driving genotoxicity of oncogenes (Neelsen et al., 2013).
3.3 The Human Oncoprotein DEK
The DEK protein was firstly discovered in 1992 in a patient suffering from a specific subtype of acute myeloid leukemia. In this patient the 3′ part of the Can gene from chromosome 9q34 was found to be fused to the 5′ end of the DEK gene on chromo- some 6p23 (6;9)(p23;q34) leading to the expression of a DEK-CAN fusion protein. After the patient in whom it was firstly discovered the until then unknown protein component was termed DEK (Derrek K.) (von Lindern et al., 1992a).
DEK is an abundant, non-histone, chromatin-associated protein and was found to be involved in different important cellular processes like transcription, mRNA processing, replication and repair. However, DEK exhibits no known enzymatic activity. It is the only protein of its kind and so far no paralogues have been identified. DEK is con- served in all higher eukaryotes but is absent in yeast and C. elegans. Between species, the DEK protein varies in overall length but all variants contain highly conserved C- and
