Regulatory network built by Cytochrome c with Histone Chaperones in response to DNA damage

A mi familia Agradecimientos Sirvan estas líneas para agradecer a todas aquellas personas que han contribuido con su esfuerzo y conocimiento al desarrollo de esta tesis doctoral. Primero, destacar el trabajo de mis directores de tesis, Miguel Ángel De la Rosa e Irene Díaz Moreno, a quienes realmente agradezco el esfuerzo y tesón que han invertido en formarme como científico a lo largo de esta etapa de formación predoctoral. A Miguel Ángel agradecer el acogerme en su grupo de investigación y saber guiarme a tiempo para poder realizar el trabajo que aquí presento. Asimismo, agradecerle los consejos que siempre me ha dado a lo largo del desarrollo de esta tesis doctoral, frutos de sus años de experiencia como científico y docente. A Irene, por ver creer en mi potencial como científico y animarme a continuar hasta llegar a donde hoy me encuentro. Así mismo, por su incansable esfuerzo a la hora de hacer de mí un científico mejor, ayudándome a ser más crítico con aquello que hago, a razonar los resultados obtenidos y saber adaptarme a la situación. Además, por todas las horas que ha dedicado a mi trabajo, que no han sido pocas, tanto en el imán como en el escritorio y en el laboratorio. Agradezco también las fuentes de financiación que han permitido que pueda realizar esta tesis doctoral: - Ministerio de Educación, Cultura y Deportes, por la beca de Formación del Profesorado Universitario (FPU013/04373). - Ministerio de Economía y Competitividad — Proyectos BFU2012-31670, BFU2015-71017-P y PGC2018-096049-BI00. - Junta de Andalucía — Incentivos al grupo de investigación (BIO-198). - Fundación Ramón Areces — Proyecto 2015/212. - cicCartuja — Plataforma de Interacciones Biomoleculares (BIP). - Sociedad Española de Bioquímica y Biología Molecular (SEBBM); Grupo de Resonancia Magnética Nuclear de la Real Sociedad Española de Química (GERMN, RESQ) – Bolsas de viaje para la asistencia a cursos y congresos. Cabe agradecer a los componentes del grupo Biointeractómica por el apoyo científico y personal que me han brindado a lo largo del desarrollo de esta Tesis Doctoral. A Antonio Díaz, por introducirme en el mundo de las dinámicas y saber infundirme su pasión por ellas. Además, por tener siempre tiempo para cualquier cosa que necesitase y ser una fuente inagotable de conocimiento. A Katiuska González, por todo ese esfuerzo y tiempo que me ha dedicado, así como los consejos para realizar buenas prácticas científicas y la eterna paciencia que ha tenido conmigo durante el desarrollo y la escritura de esta tesis. A Alejandra Guerra, por los buenos consejos que siempre me ha dado, y por estar ahí en las buenas y en las malas. A Sofía Muñoz, por esos cafés a media mañana y el apoyo incondicional que siempre me ha brindado. A Carlos Elena, por estar durante este periodo a mi lado en el laboratorio y hacer que esta tesis haya sido más llevadera, con charlas y discusiones de diferentes índoles, con toques a veces científicos. A Alejandro Velázquez, por soportarme y enseñarme a ser un poco, que no mucho, más ordenado y organizado, además de estar siempre a nuestro lado. A Gonzalo Pérez, por mostrarme que, aunque la meta sea difícil, siempre habrá un matraz de esperanza para producir un poco más de aquello de lo que nunca es suficiente. A Miguel Ángel Casado, que aunque haya estado poco tiempo durante el desarrollo de mi tesis, me ha enseñado que nunca hay que deja de ser uno mismo y no avergonzarse de aquello que te hace especial. A todos aquellos que han vivido de cerca las diversas fases de mi formación científica y me han acompañado en esta etapa de crecimiento personal y científico, tales como Blas, Isa, Rafa y Jonathan. A Seamus, por las correcciones que han hecho que esta tesis sea un poco más correcta y británica. A Inés Rodríguez, por su franqueza y momentos graciosos en el laboratorio. A Abraham Muñiz, por su apoyo a la hora de trabajar delante del ordenador y siempre tener tiempo para ayudarme con los scripts. A continuación, agradezco a los siguientes colaboradores que han contribuido a mejorar los resultados expuestos en esta tesis doctoral: - Al Dr. Abelardo López Rivas (Centro Andaluz de Biología Molecular y Medicina Regenerativa, Sevilla) por los consejos en el campo de la biología celular y ofrecerme su laboratorio para realizar experimentos cuando lo he necesitado. - Al Dr. Serio Gil Caballero (Serveis Tècnis de Recerca, Universitat de Girona) por sus consejos en el campo de la Resonancia Magnética Nuclear y su ayuda en la puesta a punto de dichos experimentos. Agradecer a mis padres, hermanas y familia el estar siempre ahí cuando más lo he necesitado, en los momentos de alegría y en los momentos donde más abajo he estado. Sin su apoyo no habría podido conseguir estar aquí. A Andrea, que, aun habiendo llegado en la última etapa de esta tesis doctoral, me ha apoyado incondicionalmente, dándome ánimos incluso en esos momentos que no los merecía. A María, por estar ahí siempre desde que nos hicimos amigos en aquella sala de estudio de biología y saber aguantarme durante la etapa de la carrera, máster y esta tesis doctoral. También, por estar en mis momentos más bajos y apoyarme siempre. A Guille, por esas discusiones sobre líneas temporales, videojuegos y cosas en general de la vida, además de estar siempre ahí cuando lo he necesitado. A Javi, por ser mi amigo y confidente desde el instituto, y por esas conversaciones acerca de chicas, móviles o cualquier cosa que siempre me han animado. A la Wowtherhood, por esos cafés y ser la muestra viviente que, si se quiere, se puede y que los hilos de la amistad no son precisamente los de coser. A Javi, Cris, Manu, Álvaro y Aurora, por no dejarme de lado y ofrecerme siempre su amistad. A F.C. por esos momentos de diversión y apoyo que siempre me ha brindado. Y por último, a usted, por dedicar su preciado tiempo a leer estos agradecimientos y esta tesis. CONTENTS - Abbreviations 13 - Publications and merits of the PhD Candidate 23 - Summary 31 - Chapter 1: Introduction 35 1. General overview of the DNA damage response 1.1. DNA damage inducers 1.1.1. Endogenous inducers 1.1.2. Exogenous inducers 1.2. DNA damage response upon double-strand break formation 1.2.1. Double-strand break sensors 1.2.2. Signal transducer kinases 1.2.3. Effector kinases and mediators 1.2.4. Effector factors 1.3. Fine tuning of the DNA damage response by Protein Phosphatase 2A 1.4. Role of histone chaperones in the DNA damage response 1.5. Liquid demixing process induced by a Prion-Like Domain from RNA Binding Proteins during the DNA damage response 1.6. DSB repair mechanisms 2. Introduction to histone chaperones herein studied 2.1. Acidic leucine-rich Nuclear Phosphoprotein 32 protein family 2.2. SET/Template Activating Factor-I 2.3. NAP-1 Related Protein 1 3. Deciphering the role of cytochrome c in the DNA damage response through its interaction with histone chaperones 3.1. General features of cytochrome c 3.2. Dual role of cytochrome c in cell 3.3. Post-translational modifications of cytochrome c 3.4. Novel targets of cytochrome c in humans and plants under DNA damage Francisco Rivero-Rodríguez 16 ATRIP ATM and ATR Interacting Protein BD Brownian Dynamics B.E. Binding Event BER Base Excision Repair BiGGER Biomolecular Complex Generation with Global Evaluation and Ranking BiP1/2 Luminal Binding Protein 1/2 BRCA1 BReast CAncer type 1 BSA Bovine Serum Albumin C3 Complement 3 CAF-1 Chromatin Assembly Factor 1 CAMELSPIN Cross-relaxation Appropiate for Minimolecules Emulated by Locked SPINs CARD Caspase Activation and Recruitment Domains CAT Catalases Cc Cytochrome c CD Circular Dichroism Cdk1/2 Cycline-Dependent Kinase 1/2 Cdk5 Cyclin-dependent protein kinase 5 CHK1/2 Checkpoint kinase 1/2 CI Complex I or NADH dehydrogenase CIF Complement Indirect immunoFluorescence CII Complex II or succinate dehydrogenase CIII Complex III or cytochrome bc1 complex CIV Complex IV or cytochrome c oxidase COX Cytochrome c OXidase CPT CamPtoThecin CRM1 ChRomosomal Maintenance 1 Abbreviations 17 CSP Chemical-Shift Perturbation CtIP Ctbp-Interacting Protein CV Complex V or ATP synthase DAPI 4’-6’-DiAmidino-2-PhenylIndole dihydrochloride DBSCAN Density-Based SCAN DDR DNA Damage Response DIF Direct ImmunoFluorescence DMA DiMethyl Adipimate DNA-PK DNA-dependent Protein Kinase DSB Double-Strand Break DTT DiThioThreitol EDTA EthyleneDiamineTetraAcetic acid EGTA EGTAzic acid or Ethylene Glycol Tetraacetic Acid eIF2 eukaryotic translation Initiation Factor 2A eIF2 eukaryotic translation Initiation Factor 2 ETC Electron Transport Chain EWS ErWin Sarcoma FACT FAcilitates Chromatin Transcription FBS Fetal Bovine Serum FUS FUsed in Sarcoma GARP Globally-optimized, Alternating-phase, Rectangular Pulses GFP Green Fluorescence Protein GPU Graphics Processing Unit GSK-3 Glycogen-Synthase Kinase-3 HCh Histone Chaperone HEK293T Human Embryonic Kidney 293 Transfectable Francisco Rivero-Rodríguez 18 HIGD2A Hypoxia Induced Gene 1 Domain family member 2A HIRA HIstone Regulator A Hist Histone mixture hnRNPC1/C2 heterogeneous nuclear RiboNucleoProtein C1/C2 hnRNPUL heterogeneus nuclear RiboNucleoProtein U-Like HOHAHA HOmonucear HArtmann-HAhn HR Homologous Recombination HRP HorseRadish Peroxidase HSPA5 Heat Shock 70 kDa Protein 5 HSQC Heteronuclear Single-Quantum Correlation HuR Human antigen R Hus1 Checkpoint protein Hus1 I1PP2A Inhibitor 1 of Protein Phosphatase 2A I2PP2A Inhibitor 2 of Protein Phosphatase 2A I3PP2A Inhibitor 3 of Protein Phosphatase 2A IIF Indirect ImmunoFluorescence INEPT Insensitive Nuclei Enhanced by Polarization Transfer INHAT INhibitor of Histone AcetylTransferase IPTG IsoPropyl--D-1-ThioGalactopyranoside IR Ionizing Radiation ITC Isothermal Titration Calorimetry KD Equilibrium dissociation constant KLF5 Krüpper-Like transcription Factor 5 koff Dissociation kinetic constant kon Association kinetic constant LB Luria-Bertani Abbreviations 19 LCAR Low Complexity Acidic Region LLPS Liquid-Liquid Phase Separation LRR Leucine-Rich Region LSPT Liquid-to-Solid Phase Transition Lw Linewidth MBP Maltose Binding Protein MD Molecular Dynamics MDC1 Mediator of DNA damage Checkpoint protein 1 Mdm2 Murine doble minute 2 MdmX Murine doble minute X MNase Micrococcal Nuclease Mre11 Meiotic recombination 11 n Stoichiometry NAP Nucleosome Assembly Protein Nbs1 Nijmegen breakage syndrome protein 1 NES Nuclear Export Signal NFT Intraneuronal NeuroFibrillary Tangles NHEJ NonHomologous End-Joining NLS Nuclear Localization Signal NMR Nuclear Magnetic Resonance NOE Nuclear Overhauser Effect NOESY Nuclear Overhauser Effect SpectoscopY NRP1/2 NAP-1 Related Protein 1/2 OPC OPtimal 3-Charge, 4-point rigid water model PARP Poly-ADP Ribose Polymerase PBS Phosphate-Buffered Saline PCD Programmed Cell Death Francisco Rivero-Rodríguez 20 PCR Polymerase Chain Reaction PKA cAMP-dependent Protein Kinase PLD Prion-Like Domain PME Particule Mesh Ewald PMSF PhenylMethylSulfonyl Fluoride pNP p-NitroPhenol pNPP p-NitroPhenyl Phosphate PP1 Protein Phosphatase 1 PP2A Protein Phosphatase 2A PP4 Protein Phosphatase 4 PP6 Protein Phosphatase 6 PTM Post-Translational Modification PTP Permeability Transition Pore PVDF PolyVinyliDene Fluoride Rad1 DNA repair protein Rad1 Rad50 DNA repair protein Rad50 Rad9 DNA repair protein Rad9 RBM14 RNA-Binding Motif protein 14 RBP RNA-Binding Protein REMD Replica Exchange Molecular Dynamics RG Gyration Radius RMSD Root Mean Square Deviation RMSF Root Media Square deviation of atomic position (Fluctuation) RNS Reactive Nitrogen Species ROESY Rotating frame Overhauser Effect SpectroscopY ROS Reactive Oxygen Species Abbreviations 21 RPA Replication Protein A rREMD Restrained Replica Exchange Molecular Dynamics SAS Surface Area exposed to Solvent SD Standard Deviation SDS-PAGE Sodium Dodecyl-Sulfate PolyAcrylamide Gel Electrophoresis SET/TAF-I SET/Template Activating Factor-I SOD SuperOxide Dismutase SPR Surface Plasmon Resonance SSB Single-Strand Break STRAP Serine-Threonine kinase Receptor-Associated Protein TAF15 TATA-binding protein-Associated Factor 15 TOCSY TOtal Correlation SpectroscopY TopoI Topoisomerase I TPBS Tween 20 Phosphate Buffer Saline VDAC Voltage-Dependent Anion Channel VMD Visual Molecular Dynamics Wip1 Wild-type p53-induced phophatase 1 WT Wild-Type XLF XRCC44 Like Factor XRCC44 X-Ray Cross-Complementation group 4 G Gibbs energy H Binding Enthalpy S Binding Entropy AVG Averaged Chemical-Shift perturbation H2AX Phorphorylated histone H2AX Francisco Rivero-Rodríguez 22 c Correlation time 23 Publications and merits of the PhD candidate Publications and merits of the PhD candidate 25 SCIENTIFIC PUBLICATIONS Published manuscripts 1. González-Arzola, K., Díaz-Quintana, A., Rivero-Rodríguez, F., Velázquez-Campoy, A., De la Rosa, M.A. & Díaz-Moreno, I. (2017) Histone chaperone activity of Arabidopsis thaliana NRP1 is blocked by cytochrome c. Nucleic Acids Res., 45: 2150-2165. (Chapter 6). 2. García-Mauriño, S.M., Rivero-Rodríguez, F., Velázquez-Cruz, A., Hernández-Vellisca, M., Díaz-Quintana, A., De la Rosa, M.A. & Díaz-Moreno, I. (2017) RNA binding protein regulation and cross-talk in the control of AU-rich mRNA fate. Front. Mol. Biosci., 4: 71-80. 3. García-Mauriño, S.M., Díaz-Quintana, A., Rivero-Rodríguez, F., Cruz-Gallardo, I., Grüttner, C., Hernández-Vellisca, M. & Díaz-Moreno, I. (2018) A putative RNA binding protein from Plasmodium vivax apicoplast. FEBS Open Bio, 8: 177-188. (Appendix 1). Manuscripts in preparation 1. Rivero-Rodríguez, F., et al., De la Rosa, M.A. & Díaz-Moreno, I. Cytochrome c binds to the unstructured acidic domain of ANP32B regulating its PP2A inhibitory activity. 2. Rivero-Rodríguez, F., et al., De la Rosa, M.A. & Díaz-Moreno, I. Modulation of ANP32B function and cellular localization by means of its Thr244 phosphorylation and cytochrome c binding. Summary 33 The maintenance of genome integrity is a critical process for cell life. To face DNA lesions, living beings have developed a complex network called DNA damage response (DDR), which encompasses an intricate phosphorylation signalling cascade, enabling a quick DNA damage repair. To control this signalling cascade and to avoid an illegitimate activation, this network is tightly regulated by protein phosphatases, such as Protein Phosphatase 2A (PP2A). Moreover, histone chaperones assist the DNA repair mechanisms altering chromatin dynamics through their nucleosome assembly activity. Recent studies have reported that the mitochondrial respiratory protein cytochrome c is promptly translocated into the cell nucleus upon DNA damage, where it interacts with several histone chaperones. Within this context, the role of nuclear cytochrome c has been partially disclosed. Here, we have delved into the functional consequences of the novel DNA damage-induced complex formed by cytochrome c with the histone chaperones ANP32A and ANP32B. Strikingly, the heme protein impairs the PP2A inhibitory effect exerted by the histone chaperones ANP32A, ANP32B and SET/TAF-I. Furthermore, biophysical and structural approaches have been used to characterize the cytochrome c complex with ANP32A and ANP32B and to unveil the molecular mechanism underlying the cytoplasmic accumulation of ANP32B upon its phosphorylation. In addition, we have extended our knowledge on the histone chaperone regulation in the DDR by studying the Arabidopsis thaliana complex between cytochrome c and NRP1, a SET/TAF-I analogous. Herein we propose a novel standpoint about DDR regulation, in which cytochrome c exerts a modulating role on such signalling cascade through its interaction with histone chaperones, both by tuning their nucleosome assembly and PP2A inhibitory activity. 35 Chapter 1: Introduction “The fate of destruction is also the joy of rebirth” Seele, NGE, Hideaki Anno Introduction 37 1. General overview of the DNA damage response Living beings are constantly exposed to endogenous and exogenous agents that can alter the integrity of their genetic material. These agents produce lesions in DNA which can hinder essential processes such as DNA replication and transcription. If these lesions are not mitigated or properly repaired, the genetic material can suffer mutations or genomic aberrations, which threaten cell viability and can lead to cancer or genetic diseases (Jackson and Bartek, 2009). Proper repair of these lesions is crucial for cell survival. Therefore, cells have developed specific mechanisms —collectively known as the DNA Damage Response (DDR)— which identify damage and subsequently repair it via several pathways (Rouse and Jackson, 2002; Harrison and Haber, 2006; Harper and Elledge, 2007). Although different stimuli can trigger several DDR signalling pathways, they all follow a common scheme: i) sensor proteins detect DNA damage and begin the signalling pathway; ii) transducer kinases translate the signal from the sensor proteins and propagate the signalling cascade; iii) mediator proteins coordinate and regulate DDR factors; iv) effector kinases amplify the DDR signal and activate effector factors; v) effector factors regulate the transcription of DDR-involved genes and control the cell cycle until the damage is mended (Jackson and Bartek, 2009; Burgess and Misteli, 2015). 1.1. DNA damage inducers According to the origin of DNA damage inducers, they can be classified as; endogenous, if they are produced inside the cell, and exogenous, if they originate from the extra-cellular environment. Francisco Rivero-Rodríguez 38 1.1.1. Endogenous inducers Among the endogenous agents which cause DNA lesions, Reactive Oxygen Species (ROS) are the biggest culprits. ROS are one of the most common causes of DNA damage in cells, and can initiate tumorigenesis (Iida et al., 2001; Li et al., 2002). ROS include superoxide anion (O2-·), hydrogen peroxide (H2O2), hydroxyl radicals (·OH) and singlet oxygen (1O2; De Bont and van Larebeke, 2004). ROS are generated in a wide variety of cellular processes. However, mitochondrial respiration is considered the major source of ROS in cell (Muller, 2000; Li et al., 2016). During mitochondrial respiration, O2-· is produced as a byproduct of the electron transport chain (ETC) when a single electron is transferred to O2 (Chance et al., 1979). O2-· stimulates the production of different ROS through various other routes. O2-· can be detoxified by superoxide dismutases. These enzymes cause two O2-· molecules to react with a H+, generating H2O2 as a byproduct; which can be detoxified to give H2O by catalases and peroxidases (Fridovich, 1997). Additionally, O2-· can reduce Fe3+ to Fe2+ in enzymes containing iron-sulfur cofactors, thereby inactivating the enzyme and releasing Fe2+ in the process. This Fe2+ reduces H2O2 and generates ·OH in a process known as the Haber-Weiss reaction (Koppenol, 2001). Alternatively, O2-· can be protonated, generating HO2· radicals (De Grey, 2002). Reactive Nitrogen Species (RNS), such as peroxynitrite (ONOO-) are generated as a byproduct of O2-· reacting with nitric oxide (NO·, Stamler, 1992). Furthermore, ONOO- may form secondary radicals which are also harmful to the cell (Wang et al., 2018; Figure 1.1.1). In high-energy situations, such as exercise, the electron transfer rate in mitochondria is increased to meet energetic demands (Conley, Introduction 39 2016). This may generate more ROS by ETC components (Hirst et al., 2008; Bleier and Dröse, 2013; Grivennikova et al., 2017). To avoid this, ETC components can rearrange themselves into more effective macrostructures known as respiratory supercomplexes. These molecular ensembles make electron flow be rapidly adjusted in response to stress conditions inside the cell and allow cells to fulfil their energy requirements, while maintaining low levels of ROS and high efficiency in electron transfer (Genova and Lenaz, 2014; Louro and Díaz-Moreno, 2015). Some ROS and RNS, such as O2-·, H2O2 and NO· are signalling molecules involved in cellular communication (D’Autreaux and Toledano, 2007). Their abundance is controlled by anti-oxidant mechanisms to prevent cellular damage (Birben et al., 2012). Additionally, ROS promote lipid peroxidation, which deteriorates cell membranes. For instance, cardiolipin peroxidation has a negative impact on mitochondrial function as it causes a marked reduction in the activity of ETC respiratory complexes I, III and IV (Paradies et al., 2000; 2001; 2002). Moreover, cardiolipin oxidation by cytochrome c has been described as a critical event in the initiation of mitochondrial-mediated apoptosis (Iverson and Orrenius, 2004; Gonzalvez and Gottlieb, 2007). However, some studies point out that these peroxidated lipids (which are very short-lasting) may act as secondary messengers of oxidative stress (Barrera et al., 2008). Furthermore, ROS and RNS can interact with DNA to generate aberrant nucleotides, such as 8-Oxo-2’-deoxyguanosine (8-oxo-dG). These altered nucleotides pair incorrectly with their complementary bases resulting in altered mRNA transcripts or point mutations in DNA during transcription or replication. For example, 8-oxo-dG may pair with adenine instead of cytosine, thereby generating a GC to TA transversion (De Bont and van Larebeke, 2004). These oxidized nucleotides must be Francisco Rivero-Rodríguez 40 promptly repaired to maintain genome and proteome integrity. In this way, cells use specific systems —namely Base Excision Repair (BER)— to remove damaged bases (Svilar et al., 2011). However, if these nucleotides are not properly repaired, Single-Strand Breaks (SSBs) and Double-Strand Breaks (DSBs) can occur during transcription (Woodbine et al., 2011). Figure 1.1.1. Scheme of the ROS derived from O2-·. Superoxide anion O2-· initiates the production of different ROS in cell. For instance, superoxide dismutase enzymes (SOD) catalyze the formation of hydrogen peroxide (H2O2) using O2-· as a substrate. Later, H2O2 can be detoxified by catalases (CAT) or peroxidases (PX) to water. Additionally, H2O2 can undergo the Haber-Weiss reaction to generate hydroxyl radicals (·OH) in a process mediated by ironsulfur cofactors (4Fe-4S), which can be stimulated by O2-·. Moreover, O2-· can be protonated and generate HO2· radicals. O2-· can also react with nitric oxide (NO·), generating peroxynitrite (ONOO-), which can form secondary RNS radicals. Adapted from Wang et al., 2018. 1.1.2 Exogenous inducers Besides endogenous inducers, cells are also exposed to a variety of environmental stimuli capable of provoking DNA damage. These exogenous inducers can either be chemical substances (such as genotoxic agents used in medical treatments), contaminants present in the media, or ionizing radiation (IR; Jackson and Bartek, 2009). Introduction 41 There are several kinds of chemical agents used in medical treatments which cause DNA damage. For example, alkylating and methylating agents form adducts with DNA bases, producing mismatches during transcription and replication, as described before (Kondo et al., 2010). Other groups of anticancer drugs, such as epirubicin and doxorubicin, disrupt the balance of ROS:antioxidants by increasing the endogenous levels of ROS. Both molecules possess an anthracycline structure which leads to ROS production in cell (Minotti et al., 2004). Additionally, other drugs, such as camptothecin (CPT) and its derivatives, promote SSBs and DSBs in DNA. These drugs are inhibitors of topoisomerases I or II, and mainly affect cells in S-phase of the cell cycle (Del Bino et al., 1991). CPT has a planar ring structure that can be interspersed in the double helix of DNA, establishing hydrogen bonds with both DNA and topoisomerase I (Topo I). In fact, when Topo I encounters a CPT molecule, a tertiary complex is formed between Topo I, CPT and DNA. These complexes inhibit the re-joining activity of Topo I, but are not lethal alone. Their lethal effect is achieved when, during transcription or replication, RNA polymerase or the replication fork collides with the Topo I-CPT-DNA tertiary complex, in an event known as the collisional model (Liu et al., 2000). On one hand, if the elongating RNA polymerase collides with the Topo I-CPT-DNA complex transcription is halted and a SSB produced at the position of the Topo I- CPT-DNA complex (Wu and Liu, 1997; Figure 1.1.2.A). On the other hand, if the replication fork encounters a Topo I-CPT-DNA complex during S-phase, the replication process will stop and a DSB will be formed in one of the replicating strands of the DNA (Hsiang et al., 1989; Tsao et al., 1993; Figure 1.1.2.B). Beside drugs, IR including X-rays, gamma rays,  and  particles and neutrons can produce different types of DNA damage: base Francisco Rivero-Rodríguez 48 phosphorylated by CHK2, which promotes binding to 14-3-3 proteins and subsequent degradation (LeBron et al., 2006). Additionally, p53 is phosphorylated by the aforementioned kinases at residues Ser15, Ser20 and Ser36, which prevent Mdm2 interacting with p53, thus evading p53 ubiquitination and degradation (Shieh et al., 1997; 2000; Appella and Anderson, 2001). The transcription factor E2F1 is also regulated by ATM and CHK2 phosphorylation. These phosphorylation events increase E2F1 stability. This results in an increase in the transcript levels of genes related to DDR repair mechanisms and apoptosis (Lin et al., 2001; Stevens et al., 2005). 1.3 Fine tuning of the DNA damage response by Protein Phosphatase 2A The DDR is a combination of several signalling pathways based on alternative phosphorylation reactions, as described in the previous section. These phosphorylation events exponentially amplify the DDR signal, allowing it to be fine-tuned, thus evading illegitimate activation. Phosphatases play a key role in controlling the DDR signalling network at different levels (Virshup and Shenolikar, 2009). Two major families of phosphatases regulate this process: i) the serine/threonine phosphoprotein phosphatases, including Protein Phosphatase 1, 2A, 4 and 6 (PP1, PP2A, PP4 and PP6, respectively); ii) the metal-dependent protein phosphatases, such as PP2C and Wild-type p53-induced phosphatase 1 (Wip1; Freeman and Monteiro, 2010). Among these, PP2A stands out as it is ubiquitously expressed and contributes almost 1% of the total protein content in mammalian cells (Ruediger et al., 1991). PP2A regulates several cellular processes Introduction 49 including glycolysis, lipid metabolism and the synthetic pathway of catecholamines (Tung et al., 1985). Additionally, PP2A participates in other important cellular processes such as cell cycle regulation, DNA replication, transcription, mRNA translation, cell proliferation, signal transduction and apoptosis (Ronne et al., 1991; Glenn and Eckhart, 1992; Alberts et al., 1993; Schönthal, 2001). Notably, PP2A activity can be downregulated by the following histones chaperones: i) Acidic leucine-rich Nuclear Phosphoprotein 32 family member A (ANP32A) and ii) SET/Template Activating Factor-I (SET/TAF-I), a.k.a. I1PP2A and I2PP2A, respectively. Both histone chaperones are responsible for catalysing the hyperphosphorylation of tau protein in Alzheimer’s Disease (AD; Chen et al., 2008; Arnaud et al., 2011). PP2A is a trimeric enzyme composed of three subunits: i) a catalytic subunit (PP2A-C), which is responsible for enzymatic activity; ii) a regulatory subunit (PP2A-B), which coordinates substrate specificity and localization of the enzyme; iii) a scaffold subunit (PP2A-A), which acts as a platform for PP2A-B and PP2A-C (Figure 1.1.4; Seshacharyulu et al., 2013). PP2A possesses high variability in its constituent subunits, allowing the enzyme to adapt its activity depending on the type of substrate and cellular needs. For instance, PP2A-A and PP2A-C have two different isoforms, A, A, C and C, respectively, which are predominantly expressed in different tissues and cellular compartments (Khew-Goodall and Hemmings, 1988; Hemmings et al., 1990). Furthermore, the PP2A-B subunit, considered the master regulator of PP2A, has 26 different isoforms divided in four different B subfamilies that modulate PP2A targets and enzyme subcellular localization (Zolnierowicz et al., 1994). Another mechanism by which PP2A activity can be regulated is through the PTMs of different subunits. These PTMs include phosphorylation and methylation (Seshacharyulu et al., 2013). Francisco Rivero-Rodríguez 50 PP2A catalyses the dephosphorylation of several proteins involved in different stages of the DDR (Figure 1.1.5). For example, PP2A dephosphorylates RPA, in a process required for the effective repair of DSBs (Feng et al., 2009). Additionally, PP2A dephosphorylates the transducer kinase ATM, diminishing its activity and ability to bind the MRN complex, and DNA-PK and Ku70/Ku80 subunits, thereby increasing the affinity of DNA-PK for Ku proteins, which in turn promotes DNA repair (Goodarzi et al., 2004; Wang et al., 2009). Figure 1.1.4. Tridimensional structure of the heterotrimeric PP2A enzyme. PP2A scaffold subunit (PP2A-A), represented in orange, acts as a platform for the regulatory (PP2A-B) and the catalytic (PP2A-C) subunits, represented in blue and red respectively. PDB code: 2IAE (Cho and Xu, 2007). Effector kinases CHK1 and CHK2 are also regulated by PP2A. Dephosphorylation of these kinases ceases DNA damage signal transmission (Goodarzi et al., 2004; Petersen et al., 2006; Freeman et al., 2010). It has also been described that PP2A-mediated -H2AX dephosphorylation is a signal which initiates DNA repair machinery after a DSB forms (Chowdhury et al., 2005). Some data has been published describing the dephosphorylation of Ser15 of p53 (Messner et al. 2006), Introduction 51 but further studies are needed to explore the functional consequences of p53 dephosphorylation by PP2A. Altogether, the role of PP2A in the fine tuning of DDR has been clearly stated. The prominent role played by the protein in DDR regulation has prompted the idea that it may act as a DDR switch. This makes the enzyme a plausible drug target for the treatment of several diseases including cancer (Ruvolo et al., 2016; O’Connor et al., 2018). Figure 1.1.5. Regulatory role of PP2A during the DNA damage response. Proteins involved in the DDR signalling pathway are represented using the same colour scheme as Figure 1.1.3 with connections in grey. Dephosphorylation events carried out by PP2A, coloured in light brown, are represented in red. Those which promote DNA repair mechanisms are represented by red arrows; whereas those impacting the DDR pathway are represented by red blunt head arrows. Red-dashed blunt head arrows correspond to the aforementioned dephosphorylation of Ser15 from p53 by PP2A. 1.4 Role of histone chaperones in the DNA damage response During DDR events, chromatin dynamics play an essential role by facilitating the entry of DNA repair factors into the damaged site. As Francisco Rivero-Rodríguez 52 previously mentioned, phosphorylation of the histone H2AX induces PTMs of different histones via the recruitment of histone modifiers (Polo and Jackson, 2011; Gospodinov and Herceg, 2013). Histone PTMs, such as acetylation and ubiquitination, are related to DDR-induced chromatin remodelling. These PTMs increase the mobility of histones and eviction rates from nucleosomes. Furthermore, histone PTMs destabilize chromatin structure by recruiting ATP-dependent remodelers to DNA foci. These enzymes disrupt contacts between DNA and histones, resulting in histone eviction from damaged sites (Wang et al., 2006; Ikura et al., 2007; Lan et al., 2012). Apart from ATP-dependent remodelers, a set of proteins known as histone chaperones also participate in chromatin remodeling. Histone chaperones assist in the incorporation of newly synthesized histones into DNA, thus restoring chromatin integrity after DNA has been repaired. For example, Chromatin Assembly Factor 1 (CAF-1) and Anti- Silencing Factor 1 (ASF1) regulate chromatin dynamics by initiating the de novo incorporation of histone H3 variants during DNA damage repair (Mello et al., 2002; Polo et al., 2006). Additionally, HIstone Regulator A (HIRA) also promotes nucleosome assembly by depositing histone H3 variants (H3.1 and H3.3) after DSBs have been fixed. Following this, transcription is re-established (Adam et al., 2013). Furthermore, H2A- H2B turnover is controlled by the histone chaperones FAcilitates Chromatin Transcription (FACT), Aprataxin and PNPK-Like Factor (APLF), which allow the deposition of newly synthesized H2A and H2B histones after DNA has been repaired. Moreover, FACT facilitates H2A/H2AX exchange upon DSBs formation (Mehrotra et al., 2011; Dinant et al., 2013). Besides assembling nucleosomes, it has also been speculated that some histone chaperones can evict nucleosomes from damaged Introduction 53 foci. For instance, nucleolin and ASF1 act by removing histones from nucleosomes at DSB foci (Goth et al., 2007; Goldstein et al., 2013). ASF1 functions by binding the H3-H4 dimer occluding the tetramerization interface, thus evicting the two histones as dimers from the nucleosome (Natsume et al., 2007). Taken together these findings suggest a more extensive role for histone chaperones in the DDR whereby they play a dual role in: i) assisting histone eviction from nucleosomes and facilitating DNA repair mechanisms; ii) restoring nucleosomes with newly synthesized histones when the damage is repaired (Polo, 2016). 1.5 Liquid demixing process induced by a Prion-Like Domain from RNA Binding Proteins during the DNA damage response RNA Binding Proteins (RBP) participate in DDR repair mechanisms (Dutertre and Vagner, 2017). For example, heterogeneous nuclear RiboNucleoProtein U-Like (hnRNPUL) proteins 1 and 2 have been identified as members of the MRN complex during the reparation of DSBs. Where, hnRNPUL promotes DNA-end resection and recruits DNA helicases (Hong et al., 2013). Strikingly, recent studies have hypothesized that RBPs could also participate in the DDR by triggering a Liquid-Liquid Phase Separation (LLPS) at DSB sites. LLPS, a.k.a. liquid demixing, generates a membrane-less environment with different physical properties than the surrounding nucleoplasm, reorganising the soluble nuclear space around the damage (Kai et al., 2016). The molecular mechanism underlying LLPS formation at DNA damage foci is still poorly understood. Some reports hypothesize that DNA damage-induced LLPS formation depends on electrostatic Francisco Rivero-Rodríguez 54 interactions between Poly-ADP Ribose Polymerase (PARP) and Prion- Like Domains (PLDs) of RBPs (Pears et al., 2012; Kai et al., 2016). Among PLD-containing RBPs, FUsed in Sarcoma (FUS), ErWin Sarcoma (EWS), TATA-binding protein-Associated Factor 15 (TAF15) and RNA-Binding Motif protein 14 (RBM14) are localized at DNA damage foci (Altmeyer et al., 2015; Hennig et al., 2015). PLDs are low complexity regions, mainly composed of uncharged polar amino acids and glycine residues. PLDs also contain Arg-Gly-Gly (RGG) repeats, which could interact electrostatically with PARP. Additionally, recent research points out that LLPS are stabilized by cation-π interactions between Arg and Tyr residues in PLDs. Such interactions reduce the number of contacts between PLD regions and increase the affinity of PLD for RBP, thus stabilizing the LLPS (Monahan et al., 2017; Murray et al., 2017; Wang et al., 2018). By this mechanism, DNA damage concentrates PLD-containing RBPs around PARP-triggering LLPS (Figure 1.1.6). The functional consequences of DNA damage-induced LLPS remains unveiled. It has been proposed that LLPS generates a platform by which repair machinery including PARP and the MRN complex are recruited to begin DDR signalling (Kai et al., 2016). Therefore, further studies are needed to accurately determine the function of LLPS under DSB-induced DNA damage. LLPS increases the risk of FUS pathological aggregation, in a process known as the Liquid-to-Solid Phase Transition (LSPT). The LLPS-to-LSPT transition occurs naturally due to age-related processes. However, mutations in FUS accelerate LLPS-to-LSPT transition and are associated with Amyotrophic Lateral Sclerosis (ALS) and other agerelated diseases (Woulfe et al., 2010; Deng et al., 2014; Patel et al., 2015). Introduction 55 Figure 1.1.6. Scheme of liquid demixing formation catalysed by RNA Binding Protein-Prion Like Domanins at DNA damaged foci. Upon the formation of DSBs (A), PARP, coloured in blue, is recruited to the DNA foci. PARP synthesizes chains of ADP-ribose, represented as blue lines (B). This process recruits PLD containing RBPs, such as FUS, coloured in light orange, which interact with PARP. Moreover, histone H2AX is phosphorylated — H2AX, represented in light green — to mark the DNA damaged foci (C). Additional PLD-RBPs, such as RBM14, coloured in salmon, or TAF15, in magenta, are recruited to the DSB site to generate phase separation, represented as the cyan oval (D). This phenomenon creates a unique environment required to recruit the MRN complex and repair mechanisms (E). Adapted from Kai et al., 2016. 1.6 Double Strand Break repair mechanisms After DSBs are induced, cells initiate repair mechanisms to mitigate DNA damage. There are two main mechanisms which are activated upon the formation of DSBs, namely homologous recombination (HR) and non-homologous end-joining (NHEJ). During HR, the sister chromatid is used as a template in the S and G2 phases of the cell cycle. This prevents recombination occurring Francisco Rivero-Rodríguez 56 between non-identical sequences and increases the chances of proper reparation. As previously described, the MRN complex is recruited to DNA foci upon the formation of DSBs. To initiate this process, CtBP- interacting protein (CtIP) and other exonucleases are recruited in a MRN-dependent manner to foci (Limbo et al., 2007). These proteins then generate a 3’- single strand DNA, whose tail is coated with RPA (Wyman and Kanaar, 2006). Later, BRCA proteins catalyse the exchange of RPA for Rad51, which forms a nucleoprotein filament. Rad51 and other proteins promote the sister chromatid strand invasion, which is used as a template to repair the damage (Pardo et al., 2009). NHEJ employs a different mechanism, which can be used at any moment of the cell cycle, as it does not rely on the sister chromatid to repair the damage. When a DSB is caused, Ku70 and Ku80 proteins bind at the damaged site. There, the Ku heterodimer recruits DNA-PK and promotes autophosphorylation (Ciccia and Elledge, 2010). Then, DNA-PK recruits either Artemis nuclease or polymerases if the ends need to be trimmed or filled, respectively. Immediately after the damage is repaired, the ligation complex composed of DNA ligase IV, X-Ray Cross-complementation group 4 (XRCC44) and XRCC44 Like Factor (XLF) joins the strands (Lieber, 2010). Interestingly, there are alternative mechanisms to NHEJ, in which ATM kinase or PARP participate. The process is very similar to the one described above, although the proteins involved in signalling and the repair machinery differ. However, this alternative form of NHEJ tends to commit more errors than canonical NHEJ, especially deletions and chromosomal translocations. Notably, this type of NHEJ has been found to be more active in cancer cell lines, where the levels of DNA-PK are lower and ATM or PARP are overexpressed (Rassool and Tomkinson, 2010). Introduction 57 2. Introduction to histone chaperones herein studied 2.1 Acidic leucine-rich Nuclear Phosphoprotein 32 protein family. The Acidic leucine-rich Nuclear Phosphoprotein 32 protein family (ANP32) is composed of evolutionary conserved histone chaperone proteins present in different organisms of the animal kingdom, with homologous proteins in plants and protists (Matilla and Radrizzani, 2005). ANP32 proteins range in size from 220 to 290 amino acidic residues, and contain a N-terminal structured domain constituted by four Leucine-Rich Regions (LRRs), along with a C-terminal Low Complexity Acidic Region (LCAR; Figure 1.2.1.A). LRRs are structural motifs composed of a -strand connected by a variable loop to an -helix or an extended region. The -strand contains a preserved Leu-X-X-Leu-X- Leu-X-X-Asn-X-Leu motif, where X can be any amino acid. The association of four LRRs results in its four -strands being oriented in a parallel fashion, yielding a concave groove responsible for binding histones (Matilla and Radrizzani, 2005; Tochio et al., 2010). Additionally, the convex side of the LRR domain acts as a platform for protein-protein interactions between ANP32 members and ChRomosomal Maintenance 1 (CRM1) or PP2A (Figure 1.2.1.B; Brennan et al., 2000; Chen et al., 2008). The ANP32 LCAR C-domain is an unstructured region, mainly composed of stretches of negatively charged amino acids, such as Asp and Glu residues, with no remarkable patterns (Matilla and Radrizzani, 2005). The accumulation of negative charges prevents the formation of any secondary or tertiary structure elements, thus allowing the LCAR to remain flexible and extended in solution, as described for other proteins Francisco Rivero-Rodríguez 64 helices and two extended loops. Additionally, Cc houses a heme group, composed of an iron atom coordinated by a porphyrin ring. The heme group is linked to the protein by the c-type consensus sequence Cys-X- X-Cys-His and is essential for the correct folding of Cc; stabilising both hydrophobic and aromatic interactions (Figure 1.3.1. left panel). Although the heme group is located within a hydrophobic pocket, it is partially exposed to the solvent, explaining how Cc is able to participate in redox reactions (Figure 1.3.1. right panel; Rajagopal et al., 2013; Imai et al., 2016). Figure 1.3.1. Structure of human cytochrome c. Left panel — Ribbon representation of Cc globular structure. The porphyrin ring of the heme group is coloured in green and the iron atom, in orange. The axial ligands of the heme group, His18 and Met80, are included. The four -helices are named 1 to 4. Right panel — Cc surface representation showing that the heme group, coloured in green, is partially exposed to the solvent. PDB code 3ZCF. The iron atom from the heme group provides Cc with two different redox states, Fe2+ and Fe3+, which are essential for electron transfer. The iron atom, apart from being coordinated by the porphyrin ring, is also bound to the peptide chain by two axial ligands, His18 and Met80 residues in human Cc (Rajagopal et al., 2013). Introduction 65 3.2 Dual role of cytochrome c in cell Cc is involved in two opposing functions depending on the status of the cell. During homeostasis, Cc is localized in the mitochondrial intermembrane space where it acts as an electron carrier between complex III and IV of the ETC, and is therefore an integral part of cellular energy metabolism (Díaz-Moreno et al., 2011a). The ETC is composed of four different transmembrane protein complexes: complex I (CI; NADH dehydrogenase), complex II (CII; succinate dehydrogenase), complex III (CIII; cytochrome bc1 complex) and complex IV (CIV; cytochrome c oxidase). The main function of the ETC is to generate an electrochemical proton gradient, which is later used by ATP synthase (complex V; CV) to synthetize Adenosine TriPhosphate (ATP). To do so, the ETC oxidizes reduced substrates generated in cellular metabolism, namely NADH and FADH2, and transfers the electrons through the complexes to the final acceptor, molecular oxygen (O2; Lenaz and Genova, 2010; Papa et al., 2012). As described in section 1.1, ETC components can rearrange into macro-structures known as supercomplexes (Genova and Lenaz, 2014; Louro and Díaz-Moreno, 2015). The organization and dynamics of ETC components is a hot topic. Three different models have been proposed: i) the fluid model, in which ETC components independently diffuse through the membrane (Hackenbrock et al., 1986); ii) the solid model, which proposes that ETC complexes are stably associated in supercomplexes, facilitating electron flow (Chance and Williams, 1955; Lapuente-Brun et al., 2013); iii) the plasticity model, which considers that both the fluid and solid model are not exclusive, but a balanced distribution between free respiratory components and supercomplexes adapting the efficiency of the ETC according to cellular need (Acín- Pérez and Enriquez, 2014). These supercomplexes decrease the diffusion distance for electron carriers such as Cc, thus increasing Francisco Rivero-Rodríguez 66 electron transfer efficiency and reducing ROS production (Chaban et al., 2014; Genova and Lenaz, 2014). The stoichiometry of these associations depends on cell type, tissue-specific isoforms and cellular conditions (Fukuda et al., 2007; Ikeda et al., 2013; Genova and Lenaz, 2014; Sun et al., 2016). The mechanisms by which supercomplexes are assembled remains unresolved. However, recent studies have identified certain molecules capable of facilitating their assembly. One such factor is the Hypoxia Induced Gene 1 Domain family member 2A (HIGD2A), which under hypoxic conditions mediates the formation of the CI/2·CIII/CIV supercomplex (Chen et al., 2012). Importantly, the lipid composition of the mitochondrial membrane can influence supercomplex assembly. For example, lack of cardiolipin impacts the stability of CIV and its ability to form supercomplexes (McKenzie et al., 2006). Additionally, phosphatidylethanolamide depletion favours the formation of large supercomplexes (Böttinger et al., 2012). On the other hand, under apoptotic stimuli, Cc is released into the cytosol where it has a central role in programmed cell death. During death signalling, Cc can be released into the cytoplasm by two different mechanisms: i) mediated by Bcl-2 protein family members (Zhang et al., 2017); ii) through opening the Voltage-Dependent Anion Channel (VDAC), which is a component of the Permeability Transition Pore (PTP) (Trindade et al., 2016). Cytosolic Cc interacts with the WD40 domains of Apoptotic Protease Activating Factor-1 (APAF-1), leading to conformational changes in APAF-1, which expose its oligomerization domains (Acehan et al., 2002). Seven subunits of both APAF-1 and Cc then assemble and recruit caspase-9 through the APAF-1 Caspase Activation and Recruitment Domains (CARD), thereby forming the apoptosome (Rield et al., 2005). This complex is capable of cleaving Introduction 67 caspase-3 to activate the caspase cascade and trigger apoptosis. (Cheng et al., 2016). 3.3 Post-translational modifications of cytochrome c Cc, like many other cell proteins, is subject to PTMs which alter or tune its function. For instance, residues Tyr46, Tyr67 and Tyr74 can be nitrated (Cassina et al., 2000; MacMillan-Crow et al., 2001; Abriata et al., 2009; Díaz-Moreno et al., 2011b) and residues Thr28 and Ser47 can be phosphorylated (Zhao et al., 2011). Moreover, Cc contains residues which can be either nitrated or phosphorylated, such as Tyr48 and Tyr97 (Lee et al, 2006; Yu et al., 2008; Abriata et al., 2009; Guerra- Castellano et al., 2018). PTMs regulate the ability of Cc to function as an electron donor in the mitochondria and an apoptotic inducer in the cytoplasm. For example, nitration of Tyr46 and Tyr48 residues triggers the assembly of a non-functional apoptosome (García-Heredia et al., 2012). Moreover, nitration at Tyr74 blocks Cc-mediated caspase activation (García- Heredia et al., 2010). The functional consequences of Cc phosphorylation at Thr28 is a matter of debate. Guerra-Castellano et al. described in 2016 that mimicking Thr28 phosphorylation by a Thr-by- Asp substitution resulted in increased Cc oxidase (COX) activity using commercial CIV. However, both pThr28 Cc and a Thr-by-Glu phosphomimetic mutant showed a decrease in COX activity using CIV obtained from bovine tissue that preserves PTMs (Yu et al., 2008; Mahapatra et al., 2017). Phosphorylation of Cc and CIV could act synergistically to inhibit COX activity (Pérez-Mejías et al., 2019). Importantly, phosphomimicking the Ser47 residue increased COX activity and decreased Cc-mediated caspase-3 activation (Guerra- Castellano et al., 2016). Moreover, phosphorylation at Tyr48 elicited Francisco Rivero-Rodríguez 68 both a decrease in COX activity and caspase-3 activation (Pecina et al., 2010; Guerra-Castellano et al., 2015; Moreno-Beltrán et al., 2017) Additionally, a Tyr97 phosphomimetic mutant provided a cytoprotective effect by decreasing ROS production (through an increase in the electron transfer rate to CIV) and lowering the rate of caspase-3 activation (Guerra-Castellano et al., 2018). 3.4 Novel targets of cytochrome c in humans and plants under DNA damage Previous proteomic studies unveiled novel extra-mitochondrial partners for Cc under DNA damage conditions in both humans and plants (Martínez-Fábregas et al., 2013; 2014a; 2014b). These novel partners are involved in different pro-survival functions such as protein synthesis and folding, cellular metabolism, protection against oxidative stress, chromatin remodelling and transcriptional regulation. These functions fall outside the well-known role of Cc, raising the question whether it could regulate pro-survival processes under DNA damage stimuli. In humans, several cytosolic proteins have been described as Cc partners during DNA damage. Among these proteins, 14-3-3, Serine-Threonine kinase Receptor-Associated Protein (STRAP), Heat Shock 70 kDa protein 5 (HSPA5) and eukaryotic translation initiation factor 2A (eIF2) stand out as proteins involved in pro-survival pathways such as apoptosis regulation, cellular signalling, protein folding and transcription regulation, respectively (Kouroku et al., 2006; Macario and Conway de Macario, 2007; Kim et al., 2012; Manoharan et al., 2018). Moreover, the interaction between Cc and 14-3-3 can regulate the apoptotic caspase-cascade by blocking the inhibitory action of 14-3-3 towards APAF-1 (Elena-Real et al., 2018). Additionally, Cc is also Introduction 69 translocated into the nucleus upon DNA damage (Nur-E-Kamal et al., 2004; Zhao et al., 2010; González-Arzola et al., 2015; Nolin et al., 2016). This translocation allows Cc to interact with the histone chaperones SET/TAF-I and ANP32B, as well as heterogeneous nuclear ribonucleoproteins C1/C2 (hnRNPC1/C2; Lee et al., 2005; Muto et al., 2007; Tochio et al., 2010). Remarkably, recent studies have characterized the functional consequences of the Cc:SET/TAF-I interaction. This DNA damage-induced interaction impairs the nucleosome assembly activity of SET/TAF-I which leads reduced nucleosome formation. This phenomenon could modulate the DDR by halting DNA compaction (González-Arzola et al., 2015). In plants, Cc release from the mitochondria also takes place during DNA damage. Nevertheless, protease activation is not apoptosome-mediated, since no APAF-1 homologue has been identified in plants (Reape and McCabe, 2008). However, the aforementioned proteomic study found that Cc interacts with several novel cytosolic and nuclear pro-survival partners when DNA damage is present (Martínez- Fábregas et al., 2013). Among others, luminal binding proteins BiP1 and BiP2, the eukaryotic initiation factor 2 (eIF2) and NRP1, involved in endoplasmic reticulum stress, protein synthesis and nucleosome assembly respectively, were identified as Cc partners (Suragani et al., 2005; Zhu et al., 2006; Valente et al., 2009). Interestingly, Cc interaction studies with NRP1 revealed that the heme protein impairs the nucleosome assembly activity of NRP1, in a similar fashion as described for SET/TAF-I (González-Arzola et al., 2015; 2017). In addition, plant BiP1/BiP2, eIF2 and NRP1 are functionally analogous to human HSPA5, eIF2 and SET, respectively. Several homologous components of the DDR machinery have been found in plant genomes (Mannuss et al., 2012). In fact, Francisco Rivero-Rodríguez 70 homologues of both MRN and 9-1-1 sensor complexes have been described in Arabidopsis thaliana (Gallego and White, 2001; Heitzeberg et al., 2004; Akutsu et al., 2007). Moreover, both ATM and ATR kinases have plant homologues, namely AtATM and AtATR (García et al., 2003; Culligan et al., 2004). Notably, phosphorylation of the histone H2AX is also a hallmark of DSB-induced DNA damage in plants (Friesner et al., 2005; Amiard et al., 2010). The protein mediator BRCA1 and the effector factor Cdc25 have equivalent proteins in plants, namely AtBRCA1 and Cdc25s (Lafarge and Montané, 2003; Spadafora et al. 2011). Components of HR and NHEJ are also conserved in plants, suggesting that animals and plants share similar repair mechanisms (Hays, 2002; Kimura and Sakaguchi, 2006). Altogether, these findings suggest that there may be Cc-centred interactions conserved in human and plants in the context of: i) programmed cell death signalling pathways (Martínez-Fábregas et al., 2014b) and ii) DDR regulatory networks (a new proposal first presented in this thesis). 3.5 Cytochrome c-mediated regulation of the DNA damage response by tuning PP2A activity Based on our recent results concerning Cc-mediated inhibition of SET/TAF-I and NRP1 activity (González-Arzola et al., 2015; 2017), we propose that Cc modulates a novel regulatory DDR pathway. The fact that SET/TAF-I and ANP32A inhibit the DDR regulator PP2A (Li et al., 1995; Chowdhury et al., 2005) raises the question, can nuclear Cc regulate DDR signalling by releasing PP2A from histone chaperones? We hypothesized that, at the onset of DSBs, LLPS promotes the accumulation of histone chaperones (HCh) at the damaged area, thereby sequestering PP2A and blocking the dephosphorylation of Introduction 71 H2AX, ATM, DNA-PK, CHK1 and CHK2. In fact, H2AX, along with RPA, is kept in its phosphorylation state by ATM and DNA-PK. This is a hallmark of damaged DNA foci; phosphorylated H2AX acts as an alarm to signal DNA damage sites (Binz et al., 2004; Stiff et al., 2004). Additionally, the DDR signalling pathway activates DDR downstream factors, such as p53. This DDR activation initiates the transcription of protection mechanisms required by cells to mitigate DSBs (Figure 1.3.2.A). During DSB signalling, Cc is translocated into the nucleus (Nur- E-Kamal et al., 2004; Zhao et al., 2010; González-Arzola et al., 2015; Nolin et al., 2016) where it binds HCh and dissociates the HCh:PP2A complex. This dissociation event would take place in a scenario in which the affinity of HCh towards PP2A and Cc are very similar, as González- Arzola et al. described for SET/TAF-I:Cc and SET/TAF-I:histone complexes (2015), or different binding stoichiometries. Nevertheless, binding affinities for HCh:PP2A complexes are still unknown and further research is needed on this topic. Dissociation of the HCh:PP2A complex would increase the fraction of active PP2A in the damaged area, thereby increasing the rate of dephosphorylated H2AX and RPA proteins, either directly or through the inhibition of ATM and DNA-PK kinases (Goodarzi et al., 2004; Chowdhury et al., 2005; Feng et al., 2009; Wang et al., 2009). Once a DSB is signalled, the alarm needs to be switched off (dephosphorylation of H2AX) so repair mechanisms can act. Moreover, Cc binding HCh would hinder their nucleosome assembly activity (Díaz-Moreno et al., 2018). As a result, repair mechanism can access the naked DNA at the damaged site. Additionally, PP2A activity would also tune the phosphorylation rates of other proteins involved in the DDR signalling cascade, while damage is being repaired (Figure 1.3.2.B). Francisco Rivero-Rodríguez 72 Figure 1.3.2. Regulatory role of cytochrome c in the DNA damage response by tuning PP2A activity. (A) DSBs activate the DDR signalling pathway, allowing p53-mediated transcription of genes involved in protection mechanisms. Around DSB foci, a liquid-liquid phase transition is established in which the HCh concentration is substantially high. This “excess” of HCh can sequester PP2A, inhibiting dephosphorylation of DDR-involving kinases, such as ATM and DNA-PK, and their targets, H2AX and RPA. Phosphorylated H2AX acts as an alarm recruiting the repair mechanisms onto the DSB foci. (B) After a short period of DSB signalling, a small amount of Cc is translocated to the nucleus. Cc interacts with HCh releasing PP2A from the HCh:PP2A complex. Free PP2A dephosphorylates H2AX and RPA, in a step essential for the repair mechanisms to mend the damage. Additionally, PP2A activity tunes the DDR by dephosphorylating ATM, DNA-PK and CHK1/2 kinases. Moreover, the inhibition of ATM and DNA-PK also decreases the levels of phosphorylated H2AX and RPA. In addition, DDR downstream elements are controlled by PP2A-mediated regulation of DDR upstream elements. Furthermore, the sequestration of HCh by Cc blocks histone turnover, and therefore nucleosome formation. Hence, damaged DNA remains naked being accessible to repair machinery (Díaz-Moreno et al., 2018). (C) If DSBs persist and they are not properly mitigated, p53 translocates to the cytosol to initiate the Cc-mediated mitochondrial intrinsic apoptotic pathway. In the nucleus, Cc can completely sequester the pool of HCh, thereby releasing PP2A. The dephosphorylation activity of PP2A fully abolishes the DDR signalling pathway. Moreover, Cc completely blocks nucleosome assembly and disassembly, thus halting DNA repair (Díaz-Moreno et al., 2018). Introduction 73 If DSBs persist or even accumulate due to deficient repair, the damage may reach a point at which it cannot be repaired. At this stage p53 translocates into the cytoplasm. There, p53 activates the mitochondrial intrinsic apoptotic pathway, releasing Cc into the cytoplasm (Endo et al., 2006; Muñoz-Pinedo et al., 2012). Extramitochondrial Cc triggers apoptosome assembly and caspase cascade activation (Cheng et al., 2016). In the nucleus, the excess of Cc means that it can sequester most of the HCh pools with two major functional consequences: i) a complete halt to histone turnover due to the inhibition of nucleosome assembly activity by HCh (Díaz-Moreno et al., 2018); ii) The release of PP2A from HCh:PP2A complexes, which in turn, completely blocks the DDR and repair mechanisms. Indeed, it does not make sense to continue signalling DNA damage in cells that have already decided to die (Figure 1.3.2.C). Francisco Rivero-Rodríguez 80 nitrophenol (pNP), a product that loses a proton and is converted into pnitrophenolate under alkaline conditions. The p-nitrophenolate absorbs light at 405/410 nm, and can be easily quantified in a spectrophotometer (Figure 2.2). The amount of p-nitrophenolate generated positively correlates with PP2A activity. Figure 2.2. Schematic representation of PP2A enzymatic activity. The enzymatic activity of PP2A is measured using a spectrophotometric assay. In this assay, PP2A dephosphorylates the substrate pNPP to give pNP. Under alkaline conditions, pNP loses a proton and is converted into p-nitrophenolate, which absorbs light at 405/410 nm and can be quantified spectrophotometrically. The amount of p-nitrophenolate generated can be correlated with PP2A activity. PP2A, as previously explained in chapter 1, is a heterotrimeric enzyme composed of three subunits: scaffold, regulatory and catalytic. The commercially available PP2A enzyme lacks the regulatory subunit. This subunit modulates PP2A substrate specificity and regulates its function (Zolnierowicz et al., 1994). To properly study PP2A activity and its inhibition, it is necessary to obtain the intact form of the enzyme from mammalian cell extracts. Methodology 81 Additionally, these types of assays allow us to test enzyme inhibitors within in cell environments. If the inhibitor is a protein, it can be overexpressed by transfecting cells with a plasmid encoding the inhibitor. Afterwards, PP2A activity from cell extracts can be measured using the same methodology as described above. 3. Circular Dichroism Circular Dichroism (CD) is a phenomenum based on the interaction of polarized light with chiral molecules. Due to the chiral nature of amino acids, CD is a powerful technique in studying the structural composition of proteins in terms of secondary (Greenfield, 2006a) and tertiary structure elements (Gasymov et al., 2014). CD spectroscopy is also used to study thermal stability in proteins, as well as folding processes and how point mutations affect the overall structure of the protein (Greenfield, 2006b). Circular polarized light is divided into two main components. Left circularly polarized light (L component), which has an anti-clockwise turn, and right circularly polarized light, which turns clockwise (R component). Spectropolarimeters measure the difference in absorbance between the L and R components. If L and R components are absorbed equally, the resulting radiation will be the same as the incident source (Figure 2.3.A). In contrast, if L or R components are absorbed differently, the resulting radiation will possess elliptical polarization (Figure 2.3.B). In proteins, this phenomenon can occur when peptide bonds are placed in an asymmetric environment, as a consequence of protein tridimensional structure (Kelly et al., 2005). Secondary structure in proteins can be studied using Far-UV CD spectroscopy. Each element of secondary structure possesses a characteristic CD profile, as they differentially absorb the L or R Francisco Rivero-Rodríguez 82 components of the polarized light (Figure 2.3.C; Greenfield and Fasman, 1969). Computational analysis of the CD spectrum of a protein can estimate the fraction of the protein which adopts an -helix, -strand or random coil conformation. However, CD approaches cannot determine the exact number or position in the folding of each component. Figure 2.3. Circular dichroism and protein secondary structure determination. (A) When polarized light passes through a sample which absorbs the L and R components equally the resulting radiation is the same as the incident one. (B) However, if the sample absorbs the L and R components differently, the resulting radiation possesses elliptical polarization (Adapted from Kelly et al., 2005). (C) Each element of secondary structure has a particular CD profile. A protein with predominantly an -helix structure would show a similar pattern as the orange line. However, a protein with predominant -sheets would present a profile close to the blue line. Finally, disordered proteins would present a profile like the green line. 4. Isothermal Titration Calorimetry Isothermal Titration Calorimetry (ITC) is one of the most common calorimetry assays used to study the thermodynamic parameters of a reaction. ITC is a powerful technique capable of determining the Gibbs Methodology 83 energy (G), the binding enthalpy (H) and the binding entropy (S) of protein-protein and protein-ligand interactions (Velázquez-Campoy et al., 2004). In this technique, protein-protein interactions with mM to nM equilibrium dissociation constant (KD) values can be measured without labelling or being attached to a surface, which may interfere with the binding process. A standard ITC experiment consists of injecting defined amounts of a reagent, placed in a syringe, at regular intervals of time into a calorimetric cell, containing a second reagent. These injections lead the system to a sequence of binding equilibrium states. The system monitors the heat uptake or release upon the inter-molecular interactions after each injection, thus detecting changes in the thermodynamic potentials (G, H and S) of the reaction. Figure 2.4. ITC in protein-protein interaction studies. Left panel — Scheme of a standard ITC experiment, in which a protein contained in the cell is titrated with a second present in the syringe. Right panel — The upper panel of the graph shows the thermogram obtained in an ITC experiment: a sequence of peaks corresponding to the injections performed. The lower panel of the graph corresponds to the integration of the thermogram, also known as binding isotherms. With these integrations, thermodynamic parameters, such as KD, H and n, can be inferred. Francisco Rivero-Rodríguez 84 For data analysis, the thermogram is integrated to obtain the reaction heat and plotted as a function of concentration (binding isotherm). Different mathematical binding models can be used to determine thermodynamic parameters. Commonly, independent binding sites or cooperative phenomena are used to determine the KD, the enthalpy contribution (H) and the stoichiometry of the reaction (n) in a precise manner (Figure 2.4, right panel. Freire et al., 2009). G and S values can be derived from the KD and H values. 5. Surface Plasmon Resonance Surface Plasmon Resonance (SPR) is a biophysical technique which allows the measurement of biomolecular interactions in real-time, thus providing kinetic information about the interaction (kon and koff). The binding equilibrium constant (KA and KD) can be inferred from these parameters (Roos et al., 1998). SPR is based on the detection of changes in the refractive index on the surface of metals placed on a biochip, in contact with an aqueous solution (Kurihara and Suzuki, 2002). This biochip is composed of a thin metallic layer on top of a dielectric material, such as glass (which forms a prism). The metallic layer —usually gold— is composed of cations surrounded by a cloud of delocalized electrons (conduction band). These delocalized electrons behave in a similar way to plasma. The electronic cloud of the metal can interact with an incident photon if their frequencies match, this is known as plasmon resonance. If plasmon resonance occurs, the metal absorbs the photons from the incident light from a plane polarized (p-polarized) light source and the energy is transferred to the electrons, generating the surface plasmons. These surface plasmons are surface electromagnetic waves confined to the metal-dielectric interphase (Homola et al., 1999). Methodology 85 The excited plasmons in the metal-dielectric interface produce a similar electric field which is extended to both metal surfaces. This electric field is known as evanescent wave and its amplitude decreases exponentially as it moves away from the metal-dielectric interface towards the aqueous solution (Figure 2.5.1 left panel; Mao et al., 2011). The evanescent wave can excite plasmons in the metal-solution interface, which are very sensitive to any changes in the refractive index of the aqueous solution. Hence the formation or breakdown of molecular complexes changes the refractive index of the solution within the propagation range of the evanescent wave. This phenomenon produces a proportional change in the plasmon wave, thereby altering the angle of incidence required to create the resonance phenomenon. This change is measured and subtracted from the initial angle and is represented as the SPR signal (Figure 2.5.1 right panel; Nguyen et al. 2015). Figure 2.5.1. Plasmon resonance generation. When a p-polarized light incises over a metallic-dielectric surface with a certain angle (), a surface plasmon is generated. Then, an electric field is produced, known as evanescent wave, whose intensity decreases as it moves away from the surface. This evanescent wave can excite plasmons on the opposite side of the metaldielectric interface, where the metal-aqueous solution is located (left panel). Surface plasmons are very sensitive to changes in the solution, so if there is any molecule attached to the surface (for example due to a complex formation) the angle of incidence required to form the surface plasmons varies (right panel). Francisco Rivero-Rodríguez 86 A typical SPR experiment consists of several phases which are repeated for each analyte injection. First, there is a phase in which there is no analyte running on the surface of the biochip, known as the baseline phase (Figure 2.5.2.A). During the baseline phase, there are no changes in the refractive index, so the response is zero and is always taken as the baseline of the experiment. When the analyte is injected, there is an initial phase in which the analyte binds with the molecules attached to the surface, this is known as the association phase (Figure 2.5.2.B). During the association phase the refraction index changes due to complex formation, therefore altering the angle of the p-polarized light. This variation of angle causes the light to incise into a different position (II-I) in the detector. This change is monitored in real-time and specific software subtracts the alteration from the initial incident angle. This phase is represented as an increase in the response of the sensogram, from which kon is calculated. Afterwards, when all the binding sites are occupied by the analyte, there is a plateau known as the saturation phase (Figure 2.5.2.C), in which the change in the incident angle is at its maximum. During this phase, all the possible binding sites are occupied, so the response is maximal (Rmax). Finally, when analyte injections end, the surface is in contact with the buffer, causing the bounded analyte to dissociate from the molecules attached to the surface during a fourth phase known as the dissociation phase (Figure 2.5.2.D). During this phase, buffer without analyte flows over the biochip surface and the complexes formed then dissociate. Thus, the plasmon curve decreases and the angle returns to the initial position. Data obtained from this phase is used to calculate the koff of the complex. Methodology 87 Figure 2.5.2. SPR experiment workflow. (A) Before an analyte injection, there is a baseline phase in which there is no analyte running over the biochip surface. Therefore, there is no change in the incident angle (middle panel) and no response in the sensogram (right panel). (B) Then, when an analyte is injected, the binding sites begin to be occupied, in a process known as the association phase. The incident angle changes due to complex formation (middle panel). This change is subtracted from the initial baseline and represented in the sensogram (right panel). Kinetic association rates (kon) can be calculated with the data obtained in this phase. (C) During the saturation phase all binding sites are occupied. Here the change in the incident angle is maximum (middle panel) and the response is the greatest possible (Rmax, right panel). (D) When the injection of the analyte ends, buffer flows over the biochip surface, dissociating the complexes and removing the analyte from the surface. This is known as the dissociation phase; here the incident angle returns to the original position as the analyte is being removed (middle panel) and the signal in the sensogram decreases (right panel). From this information, the koff can be calculated. (E) Finally, the surface is cleaned to remove all possible analyte bound to the surface. This is known as the regeneration phase. Francisco Rivero-Rodríguez 88 Additionally, if the analyte has not been completely removed from the surface, there is a regeneration phase (Figure 2.5.2.E), in which the surface is cleaned with an agent (such as NaCl, NaOH or HCl) that breaks the interaction completely, thereby recovering the initial surface state. 6. Nuclear Magnetic Resonance Solution Nuclear Magnetic Resonance (NMR) spectroscopy is a powerful technique used in structural, functional and dynamics studies of biological molecules, such as sugars, nucleic acids and proteins (Kwan et al., 2011). NMR is a versatile tool, which allows the use of different experiment combinations to obtain certain information about a desired molecule, such as composition, nuclei proximity and tertiary structure (Marion, 2013). 1D 1H NMR spectra is one of the most sensitive NMR experiments, in which hydrogen atoms of a given molecule are detected to identify any compounds present in a sample. However, the complexity of biological molecules demands a greater number of dimensions to be utilized in NMR spectra. In fact, 1D 1H NMR spectra of proteins displays significant signal overlapping. Only atoms in specific chemical environments can be unambiguously identified and analyzed, such as the -methyl proton of the heme axial ligand Met80 in reduced human Cc. The Met80 residue, which coordinates the iron atom, is impacted by the shielding phenomenon due to the electron density of the porphyrin ring. This results in that the Met80--CH3 resonance shifts upfield (lower ppms, -3.2 ppm) in reduced Cc. Hence, the Met80--CH3 resonance at -3.2 ppm in the 1D 1H NMR spectrum becomes an exceptional probe to test if Cc interacts with Methodology 89 a target, since its shift or broadening can be monitored upon complex formation (for further explanation see section 6.2.1). This property has been used to study the competition between Cc and histones for binding with a histone chaperone; either SET/TAF- I or NRP1 (González-Arzola et al., 2015; 2017). First, a 1D 1H NMR spectrum of free reduced Cc is acquired, and the intensity and linewidth of the Met80--CH3 signal is used as a reference (Figure 2.6.1.A). Then, a histone chaperone (HCh) is added to the Cc solution, which results in Cc:HCh complex formation. This complex shows longer diffusional correlation time, leading to a fast signal relaxation. This produces a linewidth broadening in the 1H-Met80--CH3 signal beyond detection limits (Figure 2.6.1.B). Additionally, there is an intermediate exchange rate effect on the 1H-Met80--CH3 signal upon complex formation. Next, a histone mixture (Hist) is added to the solution. These histones compete with Cc for HCh-binding, thereby dissociating the Cc:HCh complex and releasing free Cc. As the number of histones rises, a higher fraction of free Cc is present in the solution in respect to the Cc:HCh complex, allowing the recovery of the 1H Met80--CH3 Cc resonance (Figure 2.6.1.C). To better identify protein resonances, 2D NMR spectra can be employed. These experiments relay on the coupling, both spatial and/or scalar, between two nuclei (Aue et al., 1976). Depending on the nuclei used in these types of experiments, 2D NMR spectra can be divided into homonuclear, where both nuclei in the direct and indirect dimensions are the same, or heteronuclear, in which the indirect dimension contains a different nucleus to that used in the direct dimension. Francisco Rivero-Rodríguez 96 In ROESY experiments, magnetization is directed to the x-axis and a spin lock pulse is applied along the y-axis – instead of using the cross-relaxation effect from the z-axis. This spin lock is achieved by a low-power continuous wave and is used during the mixing time of the experiment, so nuclear spins are unable to precess. After removing the spin lock field, acquisition takes place. Following this procedure, transversal cross relaxation can be observed during the mixing time (Figure 2.6.4.B; Bothner-By et al., 1984). 6.2 Heteronuclear 2D NMR Spectra 6.2.1 [1H-15N] Heteronuclear Single-Quantum Correlation Spectra Homonuclear 2D NMR experiments are useful for determining the structure of small biomolecules. However, for larger molecules such as proteins, the amount of hydrogen atoms makes identifying individual signals in 2D [1H-1H] NMR spectra a laborious task due to signal overlapping. One way in which this complexity can be reduced is the use of 2D [1H-15N] Heteronuclear Single-Quantum Correlation (HSQC) spectra. These experiments exhibit cross-peaks corresponding to the N-H scalar correlations that are present in the amide groups of the protein backbone. To carry out these experiments, a protein must be isotopically labelled with 15N, as the 14N isotope NMR signals are significantly broadened by quadrupolar interactions to the extent that they are not observable on a high-resolution NMR spectrometer. During the acquisition of a [1H-15N] HSQC spectrum, magnetization is transferred from the direct dimension (proton) to the indirect dimension (nitrogen) using an Insensitive Nuclei Enhanced by Polarization Transfer (INEPT) pulse train. Then, the magnetization Methodology 97 evolves in this dimension for a variable period of time, which allows the chemical-shift resolution in the indirect dimension to be obtained. Afterwards, magnetization is transferred to the direct dimension using a reverse-INEPT pulse train and acquired while decoupling in the indirect dimension (Figure 2.6.5; Bodenhausen et al., 1980). Figure 2.6.5. 2D [1H-15N] HSQC pulse sequence. Scheme of a HSQC pulse sequence, namely hsqcph. First, an INEPT pulse train (red-dashed square) is used to transfer the magnetization from 1H to 15N. Then, the antiphase 15N magnetization evolves during a variable period of time (d0). Afterwards, a reverse-INEPT pulse train (blue-dashed square) is used to convert 15N magnetization into in-phase 1H magnetization. Finally, the magnetization is acquired in 1H while a broadband decoupling sequence (GARP, Globallyoptimized, Alternating-phase, Rectangular Pulses) is applied in the 15N dimension. [1H-15N] HSQC spectra are used to monitor protein-ligand or protein-protein interactions, and to determine the interaction surface and dissociation constants. HSQC spectra provide a fingerprint of the labelled protein, in which each cross-peak corresponds to a residue. Upon the addition of a non-labelled partner such as a ligand, nucleic acid or other protein, perturbations in the fingerprint signals of the labelled protein can be followed. These perturbations affect those residues in direct contact with the unlabelled partner. However, Francisco Rivero-Rodríguez 98 structural and conformational changes can take place upon binding, thus effecting amide signals placed far away from the interaction surface. Among these perturbations, two were analyzed in this Thesis: chemical-shift perturbations (CSP) and specific signal linewidth broadening (Lw) (Figure 2.6.6). Figure 2.6.6. 2D [1H-15N] HSQC perturbations used to map protein interaction surfaces. (A) Overlap of a 2D [1H-15N] HSQC spectra of a free protein (red) and in complex with a partner (blue). (B) Detailed view of a CSP in a NH group with a fast-exchange rate on the NMR time scale. (C) Broadening of two amide signals upon complex formation. (D) Perturbations can be mapped on the protein surface to identify the interaction area. CSPs are coloured in a blue gradient. Signals with substantial and specific broadening (Lw) are coloured in magenta. Unassigned residues are in gray and the heme group in green. CSPs provide reliable and valuable information about interactions, due to the high sensitivity of chemical-shifts towards changes in the environment (Williamson, 2013). Furthermore, the Methodology 99 analysis of differences in chemical-shifts between free and bound states provides thermodynamic information, as well as association and dissociation time scales of the binding event (Crowley and Ubbink, 2003; Prudêncio and Ubbink, 2004). The exchange rate of a complex can be defined by comparing the differences in resonances between free and complex-bound protein. Slow exchange rates show two separated resonances for the same residue, one of them corresponds to the free state and the another represents the bound state. In fast exchange, only one well-defined resonance is observed, which is the average between the free and bound state. Finally, intermediate exchange rates result in the coalescence of the resonance lines (Williamson, 2013). Beyond CSPs, Lw information can be extracted from resonances appearing in 2D [1H-15N] HSQC experiments. In a protein-protein or protein-ligand interaction, the c of the labelled protein increases when compared with the c of the free state. This change correlates with an increase in the transverse relaxation rate, leading to meaningful changes in the Lw of the signals. Likewise, specific signal line broadening can be perceived in complexes with low-to-intermediate binding kinetics on the NMR timescale. The differences in Lw between the free state and complex can be analyzed and mapped onto the structure of the labelled protein (Madl and Sattler, 2012). 7. In silico calculations 7.1 Molecular Dynamics The use of Molecular Dynamics (MD) calculations has grown in the last few years due to the increase in computing power. MD facilitates the analysis of a wide variety of structural processes, such as conformational changes, protein folding and structural rearrangements upon ligand binding. Furthermore, these studies can be performed Francisco Rivero-Rodríguez 100 within the range of femtoseconds to milliseconds at an atomic level (Hollingsworth and Dror, 2018). MD calculations use Newton’s equations to predict the motion of atoms in a certain molecule. For MD calculations, atoms are treated as point masses using force-fields, such as AMBER (Assisted Model Building with Energy Refinement). This is necessary to reduce the computational cost of the simulation (Weiner and Kollman, 1981). The calculation is subjected to precise and controlled conditions, such as pressure, temperature, ionic strength and protonation state. Then, Newton’s laws of motion are used to calculate the position and velocity of each individual atom for every fraction of time. After each calculation step, the position and velocity of the atoms are updated and used in the next cycle until the end of the simulation is reached. Moreover, the system evaluates every atomic interaction taking place in each frame in order to determine the acceleration of each atom for the following frame. Due to this approximation, large systems, such as lipidic layers containing proteins or large macromolecule complexes, or millisecond scale simulations tend to take a long time to perform. However, the increased performance of Graphics Processing Units (GPUs) and their relative low cost have made it easier for researchers to run these kinds of calculations. MD tools such as AMBER have allowed the code to work with multi-GPUs configurations for large systems or lengthy simulations (Salomon-Ferrer et al., 2013). 7.2 Replica Exchange Molecular Dynamics Replica Exchange Molecular Dynamics (REMD) is a special algorithm developed to perform protein-folding MD calculations. The Methodology 101 algorithm allows the exchange independent replicas of the same structure between MD running in parallel, which only differ in a certain parameter, such as temperature (Sugita and Okamoto, 1999). This replica exchange allows better sampling of conformational space, as it overcomes energy barriers between minimal local free energy. During a REMD, different parallel trajectories are computed for a given initial structure. The number of replicas needed increases according to the square root of the number of atoms in the molecule. Each MD is set up at a certain temperature within a non-linear gradient, ensuring energy overlap between replicas. Once the REMD starts, the replica computations communicate regularly between each other and attempt a replica exchange if their free energies are similar. The replica exchange is performed between the closest temperature trajectories in a pairwise mode. If there is an odd number of replicas, one of the trajectory temperatures is not exchanging (Figure 2.7.1, left panel). Figure 2.7.1. REMD calculations algorithm explanation and data derived. Left panel — At the beginning of a REMD calculation, several MDs are set up in parallel within a non-linear temperature gradient. At certain periods, the parallel MDs intercommunicate (shown as black-dashed lines) allowing a replica exchange (represented by red-dashed arrows). These replica exchanges are performed pairwise and when the energy levels of each replica overlap. Right panel — REMD calculations allow the stochastic exploration of a higher number of folding pathways (represented by the black arrows) as the changes in temperature avoid the structures to get stuck in a local free energy well. Francisco Rivero-Rodríguez 102 Accordingly, if the temperature of a trajectory increases upon a replica exchange, the free energy of the system also increases. This allows a structure locked in a local minimum to explore higher energy levels. Conversely, if the temperature decreases, the free energy of the system decreases. In this case, the probability of capturing the structure in a minimum free energy well increases. This methodology allows fast scouting of the conformational space of a protein with proper statistical parameters (Figure 2.7.1 right panel). Therefore, this algorithm increases the probability that a trajectory samples the global minimum of free energy. 7.3 Brownian Dynamics In silico calculations used in MD enable the precise determination of atomic motions. However, in protein-ligand or proteinprotein interactions faster approximations are available. Indeed, complex formation heavily depends on diffusional and rotational movements. In fact, to compute the binding effect in all-atom simulations, high computational time cost and brute-force simulations are requirements. Moreover, to obtain reliable association kinetic rates, it is mandatory to explore a high number of molecular encounters (Martinez et al., 2015). Rigid-body Brownian Dynamics (BD) computations are an alternative to MD. In these calculations, the system is simplified using rigid-body models and a limited number of point effective electric charges fitting the electrostatic potential previously computed by the Poisson-Boltzmann equation and all-atom partial charges (Gabdoulline and Wade, 1997). This method reduces the total number of moving particles allowing quicker calculation times. In addition, solvation is included by implicit approaches in the simulation (Martinez et al., 2015). Methodology 103 Furthermore, desolvation effects are also implicitly included in the simulation At the beginning of BD calculations, the target protein is fixed in the centre of a sphere with a determined radius (such as 200 Å), while a probe is placed on the surface of the sphere. The probe is then allowed to move and approach the target by Brownian diffusional motions, while the approximation trajectory is recorded (Figure 2.7.2, left panel). If the probe forms an encounter complex with the target, the collision is monitored and stored as a center of mass for the probe located over the surface of the target. Moreover, the system registers vectors, both translational and rotational, which can be used to calculate whole coordinates for the probe along a trajectory. Complex formation depends on the surface charges and the shape of the electrostatic grid of the target and the probe. Figure 2.7.2. Brownian Dynamics workflow algorithm. At the beginning of a BD calculation, the target protein is placed in the centre of a sphere, and the probe on the surface. Then, the probe moves towards the target by Brownian diffusion (left panel). When the encounter complex is formed, the centre of mass is stored, and the trajectory towards the second sphere is recorded (middle panel). Finally, in the following trajectory, the probe is randomly set on the surface of the initial sphere (right panel). Every trajectory of the probe is monitored until it reaches the surface of a second sphere with a radius of 300 Å (Figure 2.7.3, middle panel), whether a collision takes place or not. In the next trajectory, the Francisco Rivero-Rodríguez 104 position of the probe is randomized over the first sphere to allow a new cycle to begin (Figure 2.7.2 right panel). 105 Chapter 3: Objectives Francisco Rivero-Rodríguez 112 release of the latter from mitochondria upon DNA damage (Martínez- Fábregas et al., 2014a). ANP32B shuttles from the nucleus to the cytosol when is phosphorylated at residue Thr244 by casein kinase II (Fries et al., 2007; Chemnitz et al., 2009). Thr244 is adjacent to the Nuclear Localization Signal (NLS) at the C-end LCAR. The ANP32B NLS is composed of the consensus sequence Lys-Arg-Lys-Arg and is located between residues 239 and 242 (Fries et al., 2007; Marfori et al., 2012). The Nuclear Export Sequence (NES) of ANP32B resides within a stretch of residues ranging from position 109 to 120 (Fries et al., 2007). In ANP32B nucleocytoplasm translocation, the histone chaperone travels alongside the RNA binding protein HUman antigen R (HuR) and viral mRNAs (Brennan et al., 2000; Bodem et al., 2011; Bauer et al., 2014). However, how phosphorylation of Thr244 regulates ANP32B cellular localization and its binding mode for the abovementioned partners remains elusive. Within this frame, it is remarkable the previously mentioned proteomic study revealing the interaction of Cc with SET/TAF-I and 14- 3-3, other chaperones than ANP32B (Martínez-Fábregas et al., 2014a; Martínez-Fábregas et al., 2014b). Strikingly, the Cc:SET/TAF-I interaction heavily impairs nucleosome assembly activity (González- Arzola et al., 2015) while the Cc:14-3-3 complex releases APAF-1 from 14-3-3-mediated inhibition, driving cells into apoptosis (Elena-Real et al., 2018). In the current study, we explore the molecular recognition between Cc and ANP32B, and how this regulates both the well-known histone binding activity of ANP32B and the herein identified function as a novel PP2A inhibitor. We will also study the structural basis of cytoplasmic ANP32B accumulation upon Thr244 phosphorylation, as Regulation of the known and the unknown functions of ANP32B 113 well as its impact on ANP32B binding with Cc which, in turn, may modulate histone chaperone function. 4.2. Materials and methods Cell cultures Heltog and HEK293T cells were cultured in Dulbecco’s modified Eagle’s medium (Sigma Aldrich) supplemented with 10% heatinactivated Fetal Bovine Serum (FBS), 2 mM L-glutamine, 100 U/mL penicillin, 100 g/mL streptomycin, and maintained at 37 ºC in a humidified atmosphere supplemented with 5% of CO2. In subcellular localization assays, 300,000 Heltog cells were grown over coverslips placed in 3.5 mm wells for 24 h before camptothecin (CPT) treatment. DNA damage was induced by adding 20 M CPT to the media for 1 or 4 h. In transfection assays, 2.5 x 106 HEK293T cells were cultured in 150 mm plates in complete media. After three days of growth, the old media was removed by aspiration and fresh media was added prior to the transfection protocol. Immunofluorescence assays After CPT treatment, cells were rinsed in PBS pre-warmed at 37 ºC to remove traces of media. Afterwards, cells were fixed by immersing the coverslips in pre-cooled absolute methanol at -20 ºC for 5 min, followed by a washing step in a PBS-0.1% Tween 20 solution. Then, the coverslips were incubated for 1 h with a solution of primary antibodies against GFP (humanized antibody, Institut Curie, catalog number A-R-H#11) and ANP32B (Abcam, catalog number ab4224) prepared in a PBS-0.1% Tween 20-3% Bovine Serum Albumin (BSA) solution at 1/25 and 1/200 dilutions for anti-GFP and anti- Francisco Rivero-Rodríguez 114 ANP32B, respectively. Afterwards, the coverslips were washed 3 times with a PBS-0.1% Tween 20 solution. Then, a mixture of secondary antihuman DyLight 488 (Jackson Immunoresearch, catalog number 709- 485-149) and anti-goat DyLight 549 (Jackson Immunoresearch, catalog number 705-505-147) antibodies (prepared in a PBS-0.1% Tween 20- 3% BSA solution at 1/50 and 1/500 dilutions) were added to the coverslips, and incubated for 45 min. This was followed by 3 washing steps in PBS-0.1% Tween 20. Nuclear staining was performed by incubating the coverslips with 4’-6’-DiAmidino-2-PhenylIndole dihydrochloride (DAPI) at a final concentration of 1 g/mL for 5 min. After repeating the washing steps, the coverslips were dried by incubating them in absolute ethanol for 2 min. Afterwards, the coverslips were mounted into glass slides using N-propyl gallate, to preserve the fluorescence, and sealed with nail polish. Images were obtained in a Zeiss LSM 7 DUO confocal microscope using a 63x oil objective. Transfection assays HEK293T cells were transfected with calcium phosphate as previously described (Kignston et al. 2003). After 48 h, cells were harvested from the plate surface using a scraper and washed with 5 mL of cold PBS solution to remove traces of media. After centrifuging the samples for 5 min at 1,500 rpm, the supernatant was removed, and cells were washed with 5 mL of cold PBS solution. Then, the supernatant was discarded by centrifugation and cells were resuspended in a specific lysis buffer for each assay. Pull-down assays Transfected HEK293T cells were resuspended in a lysis buffer containing 10 mM Tricine-NaOH pH 8.5, 1 mM PhenylMethylSulfonyl Fluoride (PMSF) and cOmplete protease inhibitors (Roche) to perform Cc pull-downs; or 25 mM Tris-HCl pH 8.0, 50 mM NaCl, 20 mM Regulation of the known and the unknown functions of ANP32B 115 Imidazole, 5% glycerol, 1% NP-40, 1 mM PMSF and cOmplete protease inhibitors for the PP2A pull-downs. Cells were then lysed by sonication (10 s, 10% amplitude, on ice). Afterwards, cell debris was removed by 15 min of centrifugation at 13,300 rpm at 4 ºC. In Cc pull-down assays, clean cell extracts were incubated with 100 g of recombinant Cc for 16 h at 4 ºC in batch. Afterwards, cell extracts were incubated with carboxymethyl cellulose matrix (Whatman), previously equilibrated with lysis buffer for 30 min at 4 ºC in batch. To remove any non-specific binding, the matrix was washed 3 times with 1 mL of washing buffer containing 10 mM Tricine-NaOH pH 8.5, 20 mM NaCl, 1 mM PMSF and cOmplete protease inhibitors. Then, Cc:ANP32B complexes were eluted from the matrix with elution buffer containing 10 mM Tricine-NaOH pH 8.5, 360 mM NaCl, 1 mM PMSF and cOmplete protease inhibitors. Results were assessed by Western- Blot using antibodies against c-myc clone 4A6 (EMD Millipore, catalog number #05-724) and Cc (obtained by immunizing male rabbits with recombinant Cc). In PP2A pull-down assays, cell extracts were incubated with a Ni-NTA matrix (Generon) for 16 h at 4 ºC in batch. Afterwards, the matrix was cleaned by washing 3 times with lysis buffer. PP2A:ANP32B complexes were eluted by adding an elution buffer containing 25 mM Tris-HCl pH 8.0, 50 mM NaCl, 500 mM Imidazole, 1 mM PMSF and cOmplete protease inhibitors. Results were assessed by Western-Blot, using antibodies against c-myc clone 4A6 (EMD Millipore, catalog number #05-724) and the catalytic subunit of PP2A clone 1D6 (EMD Millipore, catalog number #05-421). In control assays, cell extracts from HEK293T cells transfected with an empty vector were processed in the same way as the experimental samples to discard the possibility of non-specific binding. Francisco Rivero-Rodríguez 116 Histone competition assays 1D 1H NMR spectra were recorded in a 700 MHz Bruker-Avance III with a cryoprobe to monitor the signal of Met80--CH3 in a sample of 13 M unlabelled reduced Cc, either free or in the presence of 6.5 M ANP32B1-251 WT or ANP32B1-251 T244E and calf thymus histones (10- 40 g, Sigma Aldrich). To discard any non-specific interactions, a 1D 1H NMR spectrum of 13 M reduced 14N Cc was acquired in the presence of either 40 g of BSA or 40 g of calf thymus histones. All measurements were prepared in 3 mm NMR tubes containing samples with a final volume of 0.2 mL. These samples were prepared in a 10 mM sodium phosphate buffer pH 7.4, 10% D2O with 65 M sodium ascorbate to maintain Cc in its reduced redox state. Spectra were acquired and processed using TopSpin 3.5pl7 NMR software (Bruker) and represented using OriginLab 2018. Nucleosome assembly assays The nucleosome assembly activity of ANP32B1-251 species were tested using a micrococcal nuclease digestion assay, as described by González-Arzola et al., 2017. The pBlueScript SK (-) was purified by alkaline lysis as previously described by González-Arzola et al. 2015. 1 g of pBlueScript II SK (-) plasmid was relaxed with 2.5 U of Topoisomerase I from wheat germ (Promega) in a final volume of 50 L of TopoI buffer, containing 10 mM Tris-HCl pH 8.0, 50 mM NaCl, 3.5 mM MgCl2, 1 mM DiThioThreitol (DTT), 2 mM Adenosine TriPhosphate (ATP) and 0.1 mg/mL BSA at 37 ºC for 3 h. 4 g of Core Histones HeLa (EMD Millipore) were incubated with 6 g of ANP32B1-251 WT for 3 h at 37 ºC in 50 L of assembly buffer, containing 10 mM Tris-HCl pH 8.0, 50 mM NaCl, 2 mM MgCl2, 1 mM DTT, 2 mM ATP and 0.1 mg/mL BSA. Cc was also incubated together with the histone-chaperone mixture using the same conditions as before. TopoI-relaxed pBlueScript SK II Regulation of the known and the unknown functions of ANP32B 117 plasmid was combined with the histone-chaperone mixture, with and without Cc, for 3 h at 37 ºC. Afterwards, 5 mM CaCl2 was added to the mixture before the addition of MNase to improve its activity. The sample was then incubated with 7.5 U of MNase for 30 s at 37 ºC. MNase digestion was halted by adding stop buffer, containing 20 mM EthyleneDiamineTetraAcetic acid (EDTA) pH 8.0, 1 % sodium dodecyl sulfate (SDS) and 0.2 mg/mL of Proteinase K and incubated for 30 min at 37 ºC. Digested DNA was extracted using FavorPrep GEL/PCR Purification Mini Kit (Favorgen). Purified DNA was subjected to electrophoresis in 2% agarose gel, run in Tris-borate-EDTA buffer for 1 h at 80 V and visualized by ethidium bromide staining. Protein Phosphatase 2A assays Relative PP2A activity was measured as previously described (Arnaud et al. 2011) with minor modifications. HEK293T cells transfected with pcDNA 3.1 plasmids were resuspended in lysis buffer containing 10 mM Tris-HCl pH 7.4, 50 mM NaCl, 1 mM egtazic acid (EGTA) and 1 mM EDTA. Afterwards, cells were lysed by sonication (10 s, 10% amplitude, on ice). Cells debris was removed by centrifugation for 15 min at 13,300 rpm. 120 g of cell extract was incubated with increasing concentrations of recombinant Cc in reaction buffer (50 mM Tris-HCl pH 7.4; 0.1 mM CaCl2; 2.5 mM NiCl2), containing 1 mg/mL p-nitrophenyl phosphate (Sigma Aldrich) as an enzyme substrate for 15 min at 37 ºC. Reactions were stopped by adding 13% K2HPO4. Activity was monitored by measuring the absorbance at 410 nm in a plate reader spectrophotometer (BIO-RAD, model 680). Data was normalised using cell extracts transfected with an empty vector but the same concentrations of Cc to discard any impact Cc may have on PP2A Francisco Rivero-Rodríguez 118 activity. Each experimental data point was the mean ± SD value of three independent measurements. Western-Blot assays Proteins were separated by Sodium Dodecyl Sulphate- PolyAcrylamide Gel Electrophoresis (SDS-PAGE) (15% acrylamide) and transferred onto PolyVinyliDene Fluoride (PVDF) membranes (EMD Millipore) using a Mini Trans-Blot electrophoretic transfer cell (Bio-Rad). Membranes were blocked with 5% fat-free dry milk in PBS-0.1% Tween 20 (TPBS). Immunoblotting was performed with specific primary antibodies against targeted proteins. HRP-conjugated secondary antibodies were used for detection depending on the primary antibody host (Goat IgG, Dako, ref. P0449; Mouse IgG, Sigma Aldrich, ref. A9044; Rabbit IgG, Sigma Aldrich, ref. A0545). The immunoreactive bands were detected using Amersham ECL Plus Western Blotting Detection Reagents (GE Healthcare Life Sciences). DNA constructs The DNA coding for full-length ANP32B (ANP32B1-251) was inserted into the pET28a(+) bacterial expression vector with a N- terminal 6xHis-tag. DNA for cloning was obtained via polymerase chain reaction (PCR) using cDNA (Geneservice, UK). The oligonucleotides used in the PCR were 5’-AGCCATATGATGGACATGAAGAGGAGG-3’ and 5’-GTGGTGCTCGAGTCAATCATCTTCTCC-3’. ANP32B N-end LRR domain (ANP32B1-161) from pET28a(+) was obtained by mutagenic PCR for pET28a(+)-ANP32B1-251 using 5’-GATGCCGAGTAAGATGGT- GTG-3’ and 5’-CACACCATCTTACTCGGCATC-3’ as forward and reverse primers, respectively. ANP32B1-251 and ANP32B1-161 constructs were cloned into the mammalian expression vector pcDNA 3.1 using the In-Fusion HD Cloning kit (Clonetech) following the manufacturer’s instructions. Regulation of the known and the unknown functions of ANP32B 119 Primers used to linearize the pcDNA 3.1 were 5’-GAGCAGAAACTCA- TCTCTGAA-3’ as a forward primer and 5’-GGATCCGAGCTCGGTAC- CAAG-3’ as a reverse primer. The ANP32B inserts were amplified using the primers 5’-ACCGAGCTCGGATCCATGGACATGAAGAGGAGGA- TC-3’ and 5’-GATGAGTTTCTGCTCATCATCTTCTCCTTCATCATC-3’ for ANP32B1-251; and 5’-ACCGAGCTCGGATCCATGGACATGAAGAG- GAGGATC-3’ and 5’-GATGAGTTTCTGCTCCTCGGCATCTGAGTCA- GGTGC-3’ for ANP32B1-161. Maltose Binding Protein (MBP) was cloned into a pcDNA 3.1 plasmid using In-Fusion HD Cloning Kit (Clonetech) following the manufacturer’s instructions. pcDNA 3.1 was linearized using the primers described above. The MBP insert was amplified using the following primers: 5’-ACCGAGCTCGGATCCATGGCACACCATCACC-ACCAT- 3’ forward primer and 5’-GATGAGTTTCTGCTCCGGGCCCT- GAAACAGAACTTC-3’ reverse primer. The LCAR domain (ANP32B162- 251) constructs were cloned at the C-end of pcDNA 3.1 - MBP. Primers used to linearize the pcDNA 3.1- MBP were 5’-GAGCAGAAACTCATC- TCTGAA-3’ forward primer and 5’-CGGGCCCTGAAACAGAACTTC-3’ reverse primer. ANP32B162-251 inserts were amplified using the forward primer 5’-CTGTTTCAGGGCCCGGTGGATGGTGTGGATGAAGAG–3’ and the reverse primer 5’-GATGAGTTTCTGCTCATCATCTTCTCCTT- CATCATC–3’. A T244E mutant in both pET28a(+) and pcDNA 3.1 plasmids was obtained by mutagenic PCR from pET28a(+)-ANP32B1-251 or pcDNA 3.1-ANP32B1-251, using 5’-GTGAAAAGAGGAAGAGAGAAGAAGATG- ATGAAGGAGAAGATG-3’ and 5’-CATCTTCTCCTTCATCATCTTCTT- CTCTCTTCCTCTTTTCAC-3’ as forward and reverse primers. Francisco Rivero-Rodríguez 120 DNA coding for Cc was cloned into the pBTR1 vector (Olteanu et al. 2003) which also encoded the yeast heme lyase to guarantee proper protein folding. Protein expression and purification Cc was expressed in E.coli BL21 (DE3) cells as previously described (Moreno-Beltrán et al., 2015). After electroporation of the BL21 cells, clones containing the plasmid encoding Cc were selected by growing them for 16 h in LB-agar plates supplemented with 100 g/mL of ampicillin. A single clone was selected and grown at 37 ºC in 25 mL of Luria-Bertani (LB) medium. 2.5 mL of pre-culture was used to inoculate 2.5 L of LB medium and incubated at 30 ºC under agitation for 24 h. Cells were then harvested by centrifugation (6,000 rpm for 10 min) and resuspended in 1.5 mM borate buffer pH 8.5. Lysis was performed by sonication for 4 min, and the soluble protein separated from cellular debris by centrifugation (20,000 rpm for 30 min). For NMR measurements, 15N-labeled Cc was produced in M9 minimal media with 15NH4Cl as a nitrogen source. Cc purification was carried out through ionic chromatography with a carboxymethyl cellulose matrix (Whatman). Fractions containing pure Cc were dialyzed and concentrated in a 20 mL Pierce Protein Concentrator PES 3.0 M.W.C.O. (Thermo Scientific) until the desired protein concentration was achieved. The expression of ANP32B1-251 Wild-Type (WT) and T244E mutant recombinant proteins was performed in E.coli BL21 (DE3) pLysS cells, while ANP32B1-161 was expressed in E.coli BL21 (DE3). Precultures were inoculated with a single clone selected from a LB-agar plate supplemented with 100 g/mL kanamycin and 50 g/mL chloramphenicol or 100 g/mL kanamycin from a transformation with the pET28a(+)-ANP32B1-251 species or pET28a(+)-ANP32B1-161 plasmid, respectively. Pre-cultures were grown at 37 ºC for 16 h. Then, 2 L of LB Regulation of the known and the unknown functions of ANP32B 121 media was inoculated at a ratio of 1:1000 with the abovementioned precultures. Cultures were grown at 37 ºC until the O.D.600nm reached 0.4- 0.6, at which point the expression of the protein was induced by adding 1 mM IsoPropyl--D-1-ThioGalactopyranoside (IPTG). Afterwards, bacteria expressing ANP32B1-251 species were grown at 16 ºC for 16 h, whereas those expressing ANP32B1-161 were grown at 37 ºC. Cells were harvested by centrifugation (6,000 rpm for 10 min) and resuspended in lysis buffer containing 20 mM Tris-HCl pH 8.0, 100 mM NaCl, 1 mM DTT, 1 mM PMSF, 0.2 mg/mL lysozyme, 0.02 mg/mL DNase and cOmplete protease inhibitors (Roche). Cells were ruptured by sonication (cycles of 30 s at 40% of amplitude, 60 s of rest, 3 min total time, on ice). Afterwards, cellular debris was separated by centrifugation at 20,000 rpm for 30 min. The supernatant was loaded onto a Ni-NTA matrix (Generon), previously equilibrated with the lysis buffer, and incubated for 1 h in batch at 4 ºC. Recombinant proteins were eluted using a noncontinuous imidazole gradient and purity was checked by SDS-PAGE. Fractions containing the purified ANP32B proteins were merged and dialyzed for 24 h at 4 ºC against a lysis buffer without imidazole. For ANP32B1-251 proteins, impurities were removed by an additional purification step using a Superdex 200 10/300 GL (GE Healthcare) and a 10 mM sodium phosphate buffer pH 7.4 as a running buffer at a 0.5 mL/min flow rate. Protein purity was later checked by SDS-PAGE and concentrated. Circular dichroism All circular dichroism (CD) spectra were recorded using a Jasco J-815 spectropolarimeter equipped with a Peltier temperature-control system. CD intensities are presented in terms of molar ellipticity [θmolar] using molar protein concentrations. Secondary structure analysis was carried out by recording far-UV CD spectra (190–250 nm) at 25 °C in a 1-mm quartz cuvette. Samples contained 15 M of the ANP32B232-251 Francisco Rivero-Rodríguez 128 under CPT-induced DNA damage, chimeric Cc-GFP translocates into the nucleus where it colocalizes with endogenous ANP32B (Figure 4.3.1). Figure 4.3.1. CPT-induced nuclear translocation of cytochrome c and its colocalization with ANP32B. Subcellular localization of chimeric Cc-GFP (green channel) and ANP32B (red channel) in Heltog cells under homeostatic conditions (control) and CPT-induced DNA damage after 1 h and 4 h. Nuclei are stained with DAPI. Merge 1 corresponds to the overlapped images of Cc and DAPI, whereas merge 2 represents the image that combines Cc and ANP32B channels. Regulation of the known and the unknown functions of ANP32B 129 In addition, we performed pull-down assays to corroborate the physical interaction between Cc and ANP32B. Interestingly, it has been described that ANP32B can shuttle between the nucleus and the cytoplasm upon Thr244 phosphorylation by casein kinase 2 (Fries et al., 2007; Chemnitz et al., 2009). Hence, we decided to test: i) the Cc:ANP32B interaction and ii) whether such complex is regulated by phosphorylation of the histone chaperone. To do so, we designed a phosphomimetic mutant by performing a Thr244-by-Glu substitution in a mammalian expression vector containing ANP32B1-251 wild-type (WT; pcDNA 3.1-ANP32B1-251 WT). Therefore, we performed pull-down assays using cell extracts obtained from HEK293T transfected cells with either ANP32B1-251 WT or an ANP32B1-251 T244E mutant. Recombinant Cc was used as a bait to catch ANP32B1-251 species. Cc:ANP32B1-251 species complexes were co-purified using a carboxymethyl cellulose matrix. Afterwards, analysis by Western-Blot demonstrated that ANP32B1-251 WT interacted with Cc, whereas the Cc:ANP32B1-251 T244E interaction was barely detected (Figure 4.3.2). Figure 4.3.2. Pull-downs between ANP32B1-251 species and cytochrome c. Upper panel — Western-blot against cmyc tag, encoded at the C-end of ANP32B1-251 proteins, after performing the pull-down assays. Middle panel — Western-blot against Cc, demonstrating that recombinant Cc was captured in the carboxymethyl cellulose in all cases. Lower panel — Western-blot against cmyc tag as loading and transfection control. Francisco Rivero-Rodríguez 130 Altogether, this data suggests that Cc interacts with nonphosphorylated ANP32B in the nucleus. Cytochrome c competes with histones for ANP32B binding ANP32B is described as a histone chaperone, and its ability to bind histones has already been tested (Tochio et al. 2010). Hence, to determine whether Cc could impair this function, histone competition assays were performed by acquiring 1D 1H NMR spectra of the reduced Cc in the presence of ANP32B1-251 WT or ANP32B1-251 T244E mutant at increasing histone concentrations. In these titration experiments, the addition of either ANP32B1-251 WT (Figure 4.3.3.A) or ANP32B1-251 T244E (Figure 4.3.3.B) to a Cc sample at a 1:0.5 (Cc:ANP32B) ratio resulted in strong broadening of the 1H signal from the -methyl of the Cc iron axial ligand Met80, which indicates binding between ANP32B species and the hemeprotein. Later, titration of histones onto the Cc:ANP32B mixture gradually restored the initial signal intensity. This data hints that ANP32B proteins can bind histones, preventing contact between ANP32B and Cc. This recovery was more efficient in the titrations using the phosphomimetic T244E mutant. This agrees with the fact that ANP32B1-251 T244E has a substantially lower affinity for Cc, as observed in the pull-down assays. Indeed, the higher affinity of the Cc:ANP32B1-251 WT complex may hamper the binding of ANP32B1-251 WT to histones. Additionally, the presence of more negative charges in the T244E mutant may increase its affinity towards histones, due to their basic net charge. As controls, either BSA or histones were added to free Cc. The line-width of Met80--CH3 of the Cc remained unchanged, suggesting that Cc does not interact with either BSA or histones (Figure 4.3.3.C). Regulation of the known and the unknown functions of ANP32B 131 Figure 4.3.3. Cytochrome c competes with histones for binding to ANP32B1-251 WT and ANP32B1-251 T244E without functional consequences on ANP32B nucleosome assembly activity. Detailed view of the 1D 1H NMR spectra monitoring the Met80--CH3 signal of reduced Cc (black) in the presence of ANP32B1-251 WT (A) or ANP32B1-251 T244E (B) (red traces) and upon adding increasing concentrations of calf thymus histones (blue traces). (C) Details of 1D 1H NMR spectra of reduced Cc either free (black) or in the presence of BSA (orange) or histones (brown). (D) MNAse assay was performed by mixing relaxed plasmid with either histones (lane 4); histones and ANP32B1-251 WT (lane 5) or a mixture of histones, ANP32B1-251 WT and increasing amounts of Cc (lanes 6-8). Lane 2 and 3 correspond to the free plasmid and the free plasmid treated with MNase, respectively. DNA fragments corresponding to one or two nucleosomes are marked on the right. Next, we tested whether Cc could inhibit the ability of ANP32B to incorporate nucleosomes into DNA. First, ANP32B1-251 WT nucleosome assembly activity was corroborated (Figure 4.3.3.D, lane 5; Tochio et al., 2010). Surprisingly, increasing concentrations of Cc negligibly affected ANP32B nucleosome assembly in vitro (Figure 4.3.3.D, lanes 6-8). These results suggest that Cc competes with histones for binding both ANP32B species. However, this competition does not Francisco Rivero-Rodríguez 132 affect ANP32B nucleosome assembly activity, which relays on its N- terminal structured LRR rather than on the LCAR (Tochio et al., 2010). In fact, the role of the LCAR in enhancing the affinity of ANP32B towards histones is upregulated by phosphorylation of Thr244 in ANP32B. Cytochrome c blocks ANP32B, a new Protein Phosphatase 2A inhibitor The LRR domain of ANP32B shares an 81% sequence identity and 92% homology with the N-end structured region of ANP32A (a.k.a. I1PP2A or pp32). Reilly et al. published findings that the two proteins can exchange functions during brain development (Reilly et al. 2011). In addition, ANP32A has been described as an inhibitor of the Protein Phosphatase 2A (PP2A; Li et al., 1995; 1996; Chen et al., 2008). Hence, we hypothesized that ANP32B could also inhibit PP2A activity, and this inhibition could be regulated through its interaction with Cc. Therefore, we tested the ability of PP2A to bind both ANP32B1-251 WT and ANP32B1-251 T244E by pull-down assays using PP2A obtained from HEK293T cell extracts as bait. Strikingly, we detected the interaction of both ANP32B1-251 WT and the T244E mutant with PP2A in total cell extracts (Figure 4.3.4). Figure 4.3.4. ANP32B1-251 WT and its phosphomimetic T244E mutant interact with PP2A. Upper panel — Western-blot against c-myc tag, encoded at the C-end of ANP32B1-251 proteins, after performing pull-down assays. Middle panel — Western-blot against the catalytic subunit of PP2A, demonstrating that the enzyme was captured within the Ni-NTA sepharose matrix. Lower panel — Western-blot against c-myc tag as loading and transfection control. Regulation of the known and the unknown functions of ANP32B 133 To begin, we tested the ability of ANP32B to hinder PP2A activity and the role that Cc could play in regulating this inhibition. Notably, overexpression of ANP32B1-251 WT decreased PP2A activity up to ca. 60% as measured in control cells. This finding suggests the histone chaperone inhibits PP2A phosphatase activity (Figure 4.3.5.A), in a similar manner to ANP32A. Interestingly, subsequent additions of Cc led to an almost full recovery of PP2A activity. When overexpressing the phosphomimetic mutant ANP32B1-251 T244E, the degree of inhibition was lower, reaching ca. 70% of total PP2A activity as compared with control extracts (Figure 4.3.5.B). Figure 4.3.5. Cytochrome c activates PP2A by sequestering its novel inhibitor ANP32B. Relative PP2A enzymatic activity of HEK293T cell extracts transfected with either empty pcDNA 3.1, pcDNA 3.1-ANP32B1-251 WT-c-myc (A) or pcDNA 3.1-ANP32B1-251 T244E-c-myc (B) upon the addition of increasing amounts of recombinant Cc. Data shown are the mean ± S.D. of three independent experiments. Detection by Western-blot of transfected ANP32B species in cell extracts shown in panels A and B, by using anti-c-myc antibody, are included. Francisco Rivero-Rodríguez 134 Altogether, these results suggest that PP2A recognises both ANP32B species and Cc blocks PP2A binding to ANP32B, whether the histone chaperone is phosphorylated or not. To determine which domain of ANP32B is responsible for PP2A inhibition, we performed PP2A enzymatic activity assays by overexpressing either the N-terminal LRR domain (ANP32B1-161) of ANP32B or its LCAR region (MBP-ANP32B162-251). The LCAR region of ANP32B was cloned at the C-end of MBP to protect the unstructured acidic region from degradation by proteases. Surprisingly, overexpression of ANP32B1-161 inhibited PP2A activity, as did ANP32B1- 251. This agrees with previous data described for the ANP32A LRR domain (Chen et al. 2008; Figure 4.3.6.A). However, such PP2A inhibition was barely affected by addition of Cc, suggesting that: i) the LRR domain of ANP32B is responsible for PP2A inhibition; and ii) Cc is unable to recover PP2A activity after inhibition by LRR-mediated ANP32B. In addition, when the LCARs of ANP32B WT (Figure 4.3.6.B) and T244E (Figure 4.3.6.C) were overexpressed, there was negligible effect on PP2A activity (neither inhibition by ANP32B nor recovery by Cc). In conclusion, the presence of the LCAR slightly enhances PP2A inhibition in a ANP32B1-251-dependent manner. It is plausible that Cc recognises the LCAR of ANP32B (despite its own inability to affect phosphatase activity), inducing the release of PP2A from ANP32B or hampering the binding between both proteins. The novel PP2A inhibitor, which could be referred to as I3PP2A, is regulated by phosphorylation at its LCAR. In fact, the phosphomimetic ANP32B mutant blocks PP2A activity with reduced efficiency, possibly due to a weaker ANP32B1-251 T244E:PP2A interaction or a shorten lifetime of this complex. Similarly, Cc may recover PP2A activity by Regulation of the known and the unknown functions of ANP32B 135 sequestering ANP32 proteins with different dynamics. Suggesting Cc may bind ANP32B1-251 WT and the T244E mutant with different affinities, corroborating the results inferred from the histone competition and pulldown assays. Figure 4.3.6. Role of LRR and LCAR domains of ANP32B in regulating PP2A activity, along with the effect of cytochrome c. Relative PP2A activity of HEK293T cell extracts transfected with either empty pcDNA 3.1, pcDNA 3.1- ANP32B1-161-c-myc (A), pcDNA 3.1-MBP-ANP32B162-251-WT-c-myc (B) or pcDNA 3.1-MBP- ANP32B162-251-T244E-c-myc (C) upon the addition of increasing concentrations of recombinant Cc. Data shown are the mean ± S.D. of three independent experiments. Western-blots of transfected ANP32B species, using anti-c-myc antibody, corresponding to cell extracts shown in panels A-C, are included. Francisco Rivero-Rodríguez 136 ANP32B low complexity acidic region leads its binding to the heme surroundings of cytochrome c Results herein indicate that both ANP32B1-251 and ANP32B1-161 inhibit PP2A phosphatase activity. This inhibition was only sensitive to Cc addition when the ANP32B1-251 species acted as inhibitors. Furthermore, mimicking ANP32B phosphorylation altered its sensitivity to Cc. Therefore, we tested the ability of Cc to bind to different domains of ANP32B and its phosphomimic mutant by Nuclear Magnetic Resonance (NMR) titration assays, specifically 2D [1H-15N] HSQC experiments. Spectra recorded with both ANP32B1-251 WT (Figure 4.3.7.A) and the ANP32B1-251 T244E mutant (Figure 4.3.7.B) showed chemical shift perturbations (AVG; Figure 4.3.7.C and D) of specific amide signals. Residue-specific line-width broadening (Lw; Figure 4.3.7.E and F) were also evident at a 1:1 Cc:ANP32B1-251 ratio. Such behaviour is consistent with an intermediate chemical exchange regime, with kinetics in the range of 10 ms, in agreement with the transitory nature of these complexes and a dissociation constant (KD) within the micromolar range. However, Cc amide signals showed no significant chemical changes in titrations carried out between reduced Cc and ANP32B1-161 (Figure 4.3.8), even at 1:10 ratios (Cc:ANP32B1-161). Thus, Cc must recognize the LCAR of the histone chaperone ANP32B. To determine the region on the surface of Cc which interacts with ANP32B1-251 complexes, we mapped AVG, as well as broadening onto the Cc surface. Notably, both ANP32B1-251 WT (Figure 4.3.9, upper panel) and ANP32B1-251 T244E (Figure 4.3.9, lower panel) explored similar surface patches of Cc. In both complexes, the residues with large chemical-shift perturbations and significant broadening appeared near the Cc heme-crevice. These residues included Gln16, Cys17, Thr49, Regulation of the known and the unknown functions of ANP32B 137 Ile81 and Val83. Furthermore, residues with the largest AVG values were Lys5, Lys8 and Glu89 in both titrations. Figure 4.3.7. Cytochrome c amide signals show specific chemical-shift perturbations and broadening upon binding with ANP32B1-251 species. Detailed view of the superimposed [1H-15N] 2D HSQC spectra of 15N-labeled Cc upon titration with increasing concentrations of ANP32B1-251 WT (A) and ANP32B1-251 T244E (B). Colour guide for ANP32B1-251 concentration is shown in the panels. Resonance shown corresponds to Cc glutamate 89 amide group. (C-D) Plot of averaged chemical-shift perturbations (AVG) for 15N-labeled Cc titrated with ANP32B1-251 WT (C) or ANP32B1-251 T244E (D) at 1:1 molar ratio as a function of residue number. The colour bars represent the AVG categories, as follows: insignificant <0.025 ppm (blue), small 0.025–0.050 ppm (yellow), medium 0.050–0.075 ppm (orange) and large >0.075 ppm (red). (E-F) Profile of amide line-width broadening (1H1/2) of 15N-labeled Cc titrated with ANP32B1-251 WT (E) or ANP32B1-251 T244E (F) at 1:1 molar ratio as a function of residue number. The threshold was set for those residues with a broadening higher than the average plus two standard deviations, 40.2 Hz and 27.0 Hz for ANP32B1-251 WT and ANP32B1-251 T244E respectively. Francisco Rivero-Rodríguez 144 structure elements and favours an extended conformation of the LCAR, as described for similar protein regions (Uchikoga et al. 2005). Nevertheless, the C-terminal region of the LCAR contains the Nuclear Localization Signal (NLS) which is composed of positively-charged residues (239Lys-Arg-Lys-Arg242 stretch). The positive charge of the NLS can be partially shielded by the surrounding negative charges of the LCAR. As previously mentioned, casein kinase II-mediated phosphorylation of Thr244 from ANP32B1-251 leads to the accumulation of the histone chaperone in the cytoplasm (Fries et al., 2007; Chemnitz et al. 2009). Since the residue Thr244 is adjacent to the NLS of ANP32B1-251, we hypothesize that the functional differences between ANP32B1-251 WT and the T244E mutant in terms of binding to histones and PP2A inhibition can be explained by structural changes in the area close to the phosphorylation site. The PTM may somehow alter the C- terminal conformation, hence modulating cellular localization, function and regulation of ANP32B. To characterize the LCAR comprising the NLS and its adjacent residues, two synthetic peptides — one corresponding to the WT form (ANP32B232-251 WT) and another to the phosphomimic mutant (ANP32B232-251 T244E) — were obtained. The specific sequences were EEGGKGEKRKRETDDEGEDD for ANP32B232-251 WT and EEGGKGEKRKREEDDEGEDD for ANP32B232-251 T244E, in which the NLS is underlined and the position 244 is in bold. CD spectra of both ANP32B232-251 WT and T244E species indicated that they were fully unstructured or, at least, they underwent a highly dynamic folding-unfolding exchange (Figure 4.3.12.A). To gain further insight into the conformations of both peptides, we acquired 1D 1H NMR spectra (Figure 4.3.12.B). Contrary to what was expected for unfolded states, amide signals spread in the range of 7-9 ppm instead of collapsing at ca. 8 ppm in the random coil region, suggesting a non- Regulation of the known and the unknown functions of ANP32B 145 homogeneous chemical environment for all the amide groups of the peptides. The tridimensional structure of these peptides was inferred from 2D [1H-1H] TOCSY, NOESY and ROESY NMR spectra. These spectra allowed us to obtain intramolecular distances between the protons of the peptides, which can be used as distance restraints. We identified the spin systems in the TOCSY spectrum. Notably some of the signals were split in two different ones with different intensities, which suggested a slow conformational exchange within the NMR timescale. Analysis of H signals according to the chemical-shift index (H index) pointed to the presence of -helix elements involving residues Lys8-Lys10 and Asp14-Asp20 in the ANP32B232-251 WT peptide (Figure 4.3.12.C, left panel). For the ANP32B232-251 T244E peptide the -helix extends from Lys8 to Asp20 (Figure 4.3.12.C, right panel). Interestingly, during NOESY cross-peak assignment, we could detect several NHi–NHi+1 correlations in both peptides, namely: Gly6– Glu7, Arg11–Glu12, Asp14–Asp15, Asp15–Glu16 and Asp19–Asp20 in ANP32B232-251 WT and Gly6–Glu7, Arg11–Glu12, Glu12–Glu13, Asp14– Asp15 and Asp19–Asp20 in ANP32B232-251 T244E. Moreover, in the ANP32B232-251 T244E peptide, we found H–NH cross-correlation peaks involving the Thr-by-Glu substitution, namely: Lys10 H–Glu13 HN and Glu13 H–Asp15 HN, characteristic of -helix motifs. This NMR data reveals a possible transient -helix at the C-end of the ANP32B LCAR, undetected in the CD spectra because it is averaged to an unfolded state. Although the -helix is present in both WT and T244E peptides, its length and lifetime are longer in the mutant. The LCAR -helix could lead to the concealment of the NLS in phosphorylated ANP32B, thus increasing its concentration in the cytoplasm, adding a new layer of regulation to its novel functions. Francisco Rivero-Rodríguez 146 Figure 4.3.12. Structural analysis of ANP32B232-251 WT and T244E peptides. Far UV CD spectra (A), 1D 1H NMR spectra (B) and H chemicalshift index (H index; C) of ANP32B232-251 WT (left panels) and ANP32B232-251 T244E (right panels) peptides. Evaluating NOE intensities, we estimated distance restraints among different hydrogen atoms of the peptides. For this purpose, we used the NOE signal intensity from the NH-H cross-correlation of Gly4 and its NH-H distance as a reference. Such distance restraints were included in Restrained Replica Exchange Molecular Dynamics (rREMD) calculations, in which the structure of parallel molecular dynamic calculations —each set up at a non-linear temperature gradient— periodically exchange to achieve extended conformation sampling. After 20 ns of rREMD, we extracted the lowest energy structures. The results Regulation of the known and the unknown functions of ANP32B 147 showed that both ANP32B232-251 WT and ANP32B232-251 T244E peptides had a compact structure (Figure 4.3.13). Clustering analysis revealed six groups of conformations for the WT peptide and nine for the T244E peptide. Representative structures showed differences in RMSD and gyration radius (RG) compared to the lowest energy structure, but with common features within their cluster group (Figure 4.3.14.A and 4.3.14.B). Notably for the T244E peptide, representative structures of certain clusters exhibited a short -helix (Figure 4.3.14.C). Figure 4.3.13. Structural models of ANP32B232-251 WT and T244E peptides. Lowest energy models of ANP32B232-251 WT (A) and ANP32B232-251 T244E (B) peptides derived from rREMD simulations including distance restraints. The electrostatic surface potential is represented on the structure surface following the kBT colour levels represented on the right. N- and C-ends of the peptides are indicated with bold N and C letters, respectively. Thr244 phosphorylation masks the NLS, thus retaining ANP32B in the cytoplasm Aside from the secondary structure elements formed in the ANP32B232-251 constructs, we also investigated, at the atomic level, differences between WT and T244E peptides. Francisco Rivero-Rodríguez 148 Figure 4.3.14. rREMD cluster analysis of ANP32B232-251 peptides. Clustering analysis reveals 6 different structural clusters for the ANP32B232-251 WT peptide simulation (A) and 9 for the ANP32B232-251 T244E peptide simulation (B). For each representation, cluster 1 is coloured in red; cluster 2, in orange; cluster 3, in gold; cluster 4, in green; cluster 5, in cyan; cluster 6, in blue; cluster 7, in purple; cluster 8 in magenta and cluster 9, in brown. (C) Representative structures of ANP32B232-251 T244E clusters following the same colour scheme as in B. In fact, the orientation of Lys and Arg sidechains in the NLS showed substantial differences among ANP32B232-251 species. In the ANP32B232-251 WT peptide, Lys8, Lys10 and Arg11 sidechains were located on the same plane with respect to the peptide backbone (Figure 4.3.15.A, left panel). However, in the ANP32B232-251 T244E peptide the Regulation of the known and the unknown functions of ANP32B 149 Lys10 sidechain was orientated to the opposite side to the backbone, close to Arg9 (Figure 4.3.15.A, right panel). Strikingly, the rearrangement of sidechains in the ANP32B232-251 WT peptide is similar to those previously described for the consensus NLS KRKR motif (Marfori et al., 2012). Next, we measured the distances between the nitrogen atoms in the sidechains of representative structures of the clusters for both ANP32B232-251 peptides. The statistical analysis showed that the clusters of each peptide shared similar patterns. Moreover, the representative structures of each WT cluster exhibited clear differences with those of the mutant clusters (Figure 4.3.15.B). To dig deeper into the reasons for such differences, we compared each individual distance from all the clusters of both peptides. Notably, the distances between the sidechains of Lys8 and Arg11 were larger in the WT than in the T244E mutant. On the other hand, the recorded distances between Lys10 and Arg11 were larger in the T244E mutant than in the WT (Figure 4.3.15.C), consistent with different sidechain orientations in the ANP32B232-251 species. Additionally, we analysed the surface area exposed to the solvent (SAS) of those residues comprising the NLS sequence in both WT and T244E peptides in the six lowest temperature rREMD calculations, which differ in temperature from 214.2 to 302.0 K (Figure 4.3.16). This temperature range was chosen as higher temperatures could result in unfolded structures due to the energy of the system. The results showed that Lys8 and Lys10 made only partial contacts with the solvent in both ANP32B232-251 species, although the degree of solvent exclusion around Lys10 is substantially larger in the T244E mutant. Moreover, we performed 1 s-MDs of ANP32B1-251 WT and T244E proteins that included rREMD-derived restraints. New SAS analysis of Francisco Rivero-Rodríguez 150 NLS residues in the context of ANP32B full-length proteins agreed with the results inferred from rREMD on the peptides (Figure 4.3.16, black shapes). These findings highlight –within the frame of the full-length construct– that NLS SAS distribution is comparable to those obtained with ANP32B232-251 peptides. Figure 4.3.15. The sidechains of ANP32B NLS rearrange upon Thr244 phosphorylation. Representation of the sidechains involved in the NLS of ANP32B232-251 WT (A, left panel) and ANP32B232-251 T244E (A, right panel). (B) Statistical analysis of the differences in distances of the nitrogen atoms from NLS sidechains between ANP32B232-251 WT and ANP32B232-251 T244E. (C) Individual distance comparison of the NLS nitrogen atoms between the ANP32B232-251 WT and T244E clusters. Altogether, our data suggests that upon Thr244 phosphorylation, ANP32B NLS adopts a transient -helix in which sidechains of the NLS residues alter their orientation and solvent accessibility. This could prevent the recognition of the NLS by nuclear import mechanisms, Regulation of the known and the unknown functions of ANP32B 151 explaining at a molecular level why phosphorylated ANP32B is preferentially retained in the cytoplasm. Figure 4.3.16. The phosphorylation of Thr244 masks the ANP32B NLS. SAS analysis of residues comprising the NLS in the context of ANP32B232-251 peptides (coloured shapes) or ANP32B1-251 full-length proteins (black shapes). For each positively charged amino acid, data of ANP32B WT constructs are represented in the upper panels, whereas data of ANP32B T244E constructs are in the lower panels. Such data corresponds to rREMD calculations at different replica temperature: replica 1 (214.2 K) is represented in magenta; replica 2 (229.4 K), in red; replica 3 (245.8 K), in orange; replica 4 (263.2 K), in golden; replica 5 (281.9 K), in yellow; replica 6 (302.0 K), in green. Vertical axis at 0 delineates the surface exposed (positive SAS values) and hidden (negative SAS values) from the solvent. Francisco Rivero-Rodríguez 152 Cytochrome c recognizes the NLS of ANP32B LCAR To test whether the NLS region is recognized by Cc, we performed NMR titration experiments between reduced Cc and ANP32B232-251 WT or T244E peptides. Chemical-shift perturbation data was fitted to a 1:1 binding model in which the KD values were 68.0 ± 1.2 and 11.9 ± 0.8 M for WT and T244E, respectively (Figure 4.3.17). Maps of the Cc surface in which AVG resulting from the interaction with both ANP32B232-251 WT and T244E (Figure 4.3.17) showed that the Cc:ANP32B232-251 peptide interfaces were similar to those involving the ANP32B full-length species. The main difference was the lack of broadening in the titrations involving ANP32B peptides because of the small increase in the correlation time (c) for Cc upon binding to these peptides. Figure 4.3.17. Binding curves resulting from titrations between cytochrome c with either ANP32B232-251 WT or T244E. Curves representing the best global fit of 1H chemical-shifts experienced by several Cc amide signals upon interacting with WT (A) and T244E (B) peptides in a 1:1 binding model. ITC experiments were also performed to study the thermodynamic parameters of both Cc:ANP32B232-251 WT and T244E complexes. Data confirmed that the two peptides could bind Cc with KD values of 146.4 M for ANP32B232-251 WT and 96.9 M for ANP32B232- Regulation of the known and the unknown functions of ANP32B 153 251 T244E (Figure 4.3.19), after imposing a 1:1 stoichiometry (Table 4.3.3). We performed similar experiments at pH 7.4 intending to compare them with ANP32B1-251 species data. However, due to enthalpy-entropy compensation, heat per injection values were close to baseline levels and we could not analyse the data. Figure 4.3.18. Surface of cytochrome c interacting with ANP32B232-251 WT (upper panels) and T244E (lower panels) peptides. The AVG categories are as follows: insignificant <0.010 ppm (blue), small 0.010–0.025 ppm (yellow), medium 0.025–0.050 ppm (orange) and large >0.050 ppm (red). Prolines and unassigned residues are coloured in grey; the heme group, in green. Cc surface is rotated 180° around vertical axes in each view. There is an apparent discrepancy between KD values calculated from complexes involving either the ANP32B full-length protein or Francisco Rivero-Rodríguez 256 breaks. In addition, NRP1 interacts with cytochrome c using its histone-binding domain. This explains that cytochrome c competes with histones for binding to NRP1, with subsequent inhibition of the nucleosome assembly activity. These findings reveal a novel regulation of the DNA damage response in plants. 7. A cell-free expression system has been successfully used to solubilize an apicoplast RNA Binding Protein from Plasmodium vivax. The protein interacts preferentially with RNA containing U- rich stretches; and the interaction is highly dependent on ionic strength. The lack of hydrophobic residues in the 3-strand and the presence of a positive patch comprising the 2-strand and 3-helix suggest and atypical RNA binding platform. 257 Chapter 8: References “Humans forget their foolishness and repeat their mistakes. 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Doi: 10.1002/2211-5463.12351 PhD candidate contribution: Performed the cell-free expression of the RNA binding protein from Plasmodium vivax apicoplast. Wrote the paper and he was involved in planning the experiments and in the discussion of results. “Never underestimate the ability of the human animal to adapt to its environment” Misato Katsuragi, NGE, Hideaki Anno A putative RNA binding protein from Plasmodium vivax apicoplast 313 Abstract Malaria is caused by Apicomplexa protozoans from the Plasmodium genus entering the bloodstream of humans and animals through the bite of the female mosquitoes. The annotation of the Plasmodium vivax genome revealed a putative RNA binding protein (apiRBP) that was predicted to be trafficked into the apicoplast, a plastid organelle unique to Apicomplexa protozoans. Although a 3D structural model of the apiRBP corresponds to a noncanonical RNA recognition motif with an additional C-terminal α-helix (3), preliminary protein production trials were nevertheless unsuccessful. Theoretical solvation analysis of the apiRBP model highlighted an exposed hydrophobic region clustering 3. Hence, we used a C-terminal GFP-fused chimera to stabilize the highly insoluble apiRBP and determined its ability to bind U-rich stretches of RNA. The affinity of apiRBP toward such RNAs is highly dependent on ionic strength, suggesting that the apiRBP-RNA complex is driven by electrostatic interactions. Altogether, apiRBP represents an attractive tool for apicoplast transcriptional studies and for antimalarial drug design. Supplemental Material A putative RNA Binding Protein from Plasmodium vivax Apicoplast Sofía M. García-Mauriño, Antonio Díaz-Quintana, Francisco Rivero- Rodríguez, Isabel Cruz-Gallardo, Christian Grüttner, Marian Hernández-Vellisca and Irene Díaz-Moreno Supplemental Figure 1. Solubilization tests of His-apiRBP synthesized in cell free extracts. His-apiRBP (molecular weight of 13.3 kDa) was expressed by cell-free protein synthesis. Expression was carried out either in the absence (lane 1 and 8) or in the presence of different detergents (lanes 2-7 and 9-14). Specifically, the following detergents to a final concentration (w/v) were added: Brij 35 at 1 % (lanes 2 and 9); Brij 35 at 2.5 % (lanes 3 and 10); n-Dodecyl -D- maltoside (DDM) at 0.08 % (lanes 4 and 11); DDM at 0.4 % (lanes 5 and 12); Brij 58 at 0.1 % (lanes 6 and 13) and Brij 58 at 0.5 % (lanes 7 and 14). The resulting material was then centrifuged and separated into supernatant and pellet fractions and run in a SDS-PAGE gel. After protein transference to a PVDF membrane, Western Blot analysis of the anti-His-tag was performed. M: molecular weight marker (kDa). Supplemental Figure 2. Protein purification, stability and identification of apiRBP-GFP-His. A. SDS-PAGE gel of the full length chimera apiRBP-GFP-His (molecular weight of 39.9 kDa) after purification by FPLC (0 h) and after 24 h at room temperature (24 h). B and C. apiRBP-GFP-His peptides identified by mass spectrometry after trypsin digestion. B shows the expected and the experimental mass of the identified peptides (errors are comprehended +/- 0.5 Da) while C localizes the peptides in the chimera sequence (in grey) where apiRBP amino acids are colored in blue and GFP sequence in green. Supplemental Figure 3. Control experiments of the apiRBP-GFP- His binding to RNA by Isothermal Titration Calorimetry. Left, 10- mer U-rich RNA (1 M, in the calorimetric cell) was titrated with GFP- His (10 M, in the syringe). Right, ITC buffer was titrated with 10 M apiRBP-GFP-His. Each plot shows the heat released versus time in the upper panel with a thermogram of the integrated peak intensities plotted against the molar ratio of the complex in the lower panel. Both experiments were conducted in 20 mM Tris (pH 7.4) at 25 ºC. Supplemental Figure 4. apiRBP-GFP structural model. Model of apiRBP (in blue) with the GFP-tag (in green) obtained with Robetta (http://robetta.bakerlab.org). Each view is rotated 180° around the horizontal axis of the molecule.