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Role of Polynucleotide Phosohorylase during double-strand recognition in B. subtilis

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Universidad Autónoma de Madrid.

Facultad de Ciencias

Departamento de Biología Molecular

Role of Polynucleotide Phosphorylase during double-strand break

recognition in B. subtilis.

Tesis doctoral- Paula P. Cardenas M

Noviembre, 2010

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Memoria presentada por Paula Cárdenas M para optar al grado de Doctor en Ciencias por la Universidad Autónoma de Madrid.

El trabajo ha sido realizado en el Centro Nacional de

Biotecnología (CNB-CSIC) bajo la dirección del Prof. Juan Carlos

Alonso Navarro y la tutela de la Dra. Silvia Ayora Hirsch

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RESUMEN

RecN purificada de Bacillus subtilis con un 98% de pureza parece poseer una actividad exonucleasa en ADN de cadena sencilla (ADNcs), con polaridad 3’5’

dependiente de Mn2+. En este trabajo se encontró que la enzima polinucleótido fosforilasa (PNPasa), encargada de regular el metabolismo de ARN, y que se encontraba presente en aproximadamente un 0.2% de la muestra de RecN purificada, era la responsable de tal actividad. Cuando RecN era purificada de células con una mutación nula en el gen pnpA (ΔpnpA, codificante para PNPasa) la actividad nucleasa desaparecía. Sin embargo, el mismo patrón de degradación se obtenía cuando PNPasa purificada se agregaba a la preparacion de RecN purificada de células ΔpnpA.

En el presente trabajo se demuestra que PNPasa es capaz de degradar ADNcs en presencia de Mn2+ y de bajas concentraciones de fosfato inorgánico (Pi). El procesamiento de ADNcs llevado a cabo por PNPasa es distributivo, se encuentra regulado por la presencia de ATP ó dATP, y es inhibido por la presencia de Mg2+ ó de niveles altos de Pi (< 100 µM). Por el contrario, PNPasa puede degradar ARN en presencia de Mg2+ y niveles altos de Pi (10 - 30 mM), lo que sugiere que las actividades degradativas llevadas a cabo por PNPasa en ARN y en ADNcs, ocurren por mecanismos mutualmente exclusivos.

Además, en este estudio se demostró in vitro que en presencia de Mn2+ PNPasa tambien era capaz de polimerizar sin necesidad de molde en los extremos 3’-OH de un ADNcs empleando dNDPs. Como estos resultados se obtuvieron después de 5 décadas de investigación en esta enzima, se pensó que podían ser algo exclusivo de B. subtilis. Para comprobar lo anterior se purificó PNPasa de Escherichia coli, como representante de un género evolutivamente distante, y se encontró que las actividades nucleasa y polimerasa estaban muy conservadas en ambos géneros de bacterias. También se observó que en ambas enzimas las variaciones en las concentraciones de dNDPs ó Pi podrian estimular la degradación ó polimerización de ADNcs, mientras que las de (d)ATP inhibían ambas actividades. Adicionalmente se encontró que las proteínas RecN y RecA, podían modular las actividades de la enzima PNPasa, a través de la estimulación de la reacción de polimerización ó la de degradación, respectivamente.

Análisis genéticos demostraron que las células con una mutación nula en pnpA (ΔpnpA) eran más sensibles al tratamiento con H2O2 al compararlas con la cepa wt.

También se encontró que la mutación ΔpnpA no era epistática con ΔrecA, ni con funciones que procesan los extremos a los que se une RecA (addA, ΔrecJ), ni tampoco con funciones que procesan los intermediarios de recombinación catalizados por RecA (ΔrecU, ΔrecG); sin embargo, la mutación ΔpnpA sí era epistática con ΔrecN y con Δku. Lo anterior parece indicar que PNPasa estaría involucrada en la fase temprana de reparación de rupturas de doble cadena (RDC) antes de que se decida la vía de reparación, ya sea por recombinación homóloga (RH), ó por unión de extremos no homólogos (UENH). Estudios de microscopía de fluorescencia, indican que PNPasa debe ser una de las primeras proteínas en unirse a los extremos de ADNcs, ya que era necesaria para la formación adecuada de un único foco de RecN (primera proteína que reconoce el lugar del daño) por nucleoide luego de la inducción de RDC.

Los datos presentados en éste trabajo indican que PNPasa estaría involucrada en diversas vías del metabolismo de los ácidos nucleicos. Los resultados obtenidos sugieren que en ausencia de RH, la actividad degradativa de PNPasa puede estar contribuyendo a la reparación de la RDC, mediante la generación del sustrato necesario para la unión de la proteína Ku vía UENH.

 

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ABSTRACT

RecN purified from Bacillus subtilis, with 98% purity seems to have a Mn2+- dependent exonuclease activity, specific for single stranded (ss) DNA, with 3’  5’

polarity. In this work, it was found that polynucleotide phosphorylase (PNPase) enzyme, which carries out the regulation of RNA metabolism, and that was present in ~ 0.2% of purified RecN, was responsible of this nuclease activity. When RecN was purified from cells with a null mutation in pnpA gene (ΔpnpA, coding for PNPase), the nuclease activity disappeared. Furthermore, a similar degradation pattern was obtained when pure PNPase was added to RecN purified from a ΔpnpA strain. In this work is shown that PNPase is able to degrade ssDNA in the presence of Mn2+ and low concentrations of inorganic phosphate (Pi). The ssDNA processing carried out by PNPase is distributive, and is allosterically inhibited by the presence of ATP or dATP. PNPase was unable to degrade ssDNA when Mn2+ was replaced by Mg2+ or when Pi concentration exceeded 100 µM.

On the other hand, PNPase can degrade RNA in the presence of Mg2+ and high concentrations of Pi (10- 30 mM), which suggests that degradative activities carriedout by PNPase in RNA and ssDNA, occurr by mutually exclusive mechanisms. In this study, was also shown that in the presence of Mn2+, PNPase was able to polymerise in a template-independent mannerthessDNA 3’ -OH end using dNDPs. Since these activities were not detected during the 5 decades of research in this enzyme, it was thought that this could be specific for B. subtilis PNPase. To test this hypothesis E. coli PNPase was purified, as a representative of an evolutivelly divergent genera from B. subtilis. It was found that both activities, nuclease and polymerase, were conserved in both bacteria genus. The activity of both enzymes was directed towards degradation or polymerisation of ssDNA, through the manipulation of Pi or dNDPs concentrations, while (d) ATP was identified as an allosteric inhibitor of both activities. In addition, it was also found that RecN and RecA proteins were able to modulate the activities of PNPase, through the stimulation of polymerase or nuclease activity, respectively.

Genetic analysis showed that a null pnpA mutation (ΔpnpA) makes cells more sensitive to H2O2 treatment when compared with the wt strain. It was also found, that a ΔpnpA mutation was not epistatic with ΔrecA, neither with functions that process the DNA ends that will bind to RecA (addA, ΔrecJ) or with functions that process the recombination intermediates generated by RecA (ΔrecU, ΔrecG); however, ΔpnpA mutation was epistatic with ΔrecN and Δku. These data suggest that PNPase could be involved in an early stage of DSB repair, before the repair pathway has been chosen whether homologous recombination or non-homologous end-joining.

Fluorescence microscopy studies showed that PNPase, must be one of the first proteins that binds to single-strand (ss) DNA ends, because it is needed for the adequate formation of a discrete RecN (first protein that recognizes the site of damage) focus per nucleoid upon induction of DSBs.

The data showed in this work suggest that PNPase could be involved in various nucleic acid metabolic pathways. The obtained results suggest that in absence of HR, PNPase degrading activity might contribute to repair by NHEJ, generating the ssDNA substrate for Ku binding.

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ABREVIATURAS

ADNcs: ADN cadena sencilla ADNcd: ADN cadena doble CR: Centro de reparación

MEC: Mantenimiento de estructura cromosómica PNPasa: Polinucleótido fosforilasa

RDC: Ruptura de doble cadena RH: Recombinación homóloga

UENH: Unión de extremos no homólogos PNPasa: Polinucleótido fosforilasa

 

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ABBREVIATIONS

ATM: Ataxia telangiectasia mutated BRCT: Breast cancer carboxi terminal BER: Base excision repair

Cm: Chloramphenicol

DAPI: 4’,6-diamino-2-phenylindole DDR:DNA damage response

dNDPs: Deoxynucleotides diphosphate dNTPs: Deoxynucleotides triphosphate DSB: Double strand breaks

DTT: Dithiothreitol

HR: homologous recombination Htg-PNP-: His-tagged PNP MMC: Mitomycin C

MMR: Mismatch repair

MMS: Methyl methanesulfonate MRN(X): Mre11-Rad50-Nbs1(Xrs2) NDPs: Nucleosides diphosphate NER: Nucleotide excision repair NHEJ: Nonhomologous end-joining Ni-NTA Nickel matrix

NTPs: Nucleotides triphosphate

PAGE: polyacrylamide gel electrophoresis PEI: Poliethylenimine

PfkA: Phosphofructokinase

PNPase: Polynucleotide phosphorylase RecN-Htg: RecN-His tagged

RC: Repair center

ROS: Reactive oxygen species SDS: Sodium dodecyl sulfate

SMC: Structural maintenance of chromosomes Spc: Spectinomycin

SSG: Single-strand gaps 4NQO: 4-Nitroquinoline-1-oxide

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INDEX

INTRODUCCION 1

INTRODUCTION 3

1. Mechanisms of DNA repair 3

1.1 NHEJ 4

1.2 DSB response in bacteria 5

1.2.1. Homologous recombination in Bacillus subtilis 5 1.2.1.1. Break recognition and initial response to DNA damage in

B. subtilis 7

1.2.1.2. DNA end-processing in B. subtilis 8

1.3 DSB response in eukaryotes 9

2. Structural maintenance of chromosomes (SMC) proteins 9

2.1. Architecture of SMC proteins 9

2.1.2. Effect of ATP binding and hydrolysis in SMC conformation 11

2.2 B. subtilis SMC-like proteins 11

2.2.1. B. subtilis RecN 11

2.2.2 B. subtilis SbcC 13

2.2.3. B. subtilis SbcE 14

2.2.4. Comparative analysis of Bacillus subtilis SMC proteins

involved in DNA repair 14

2.3 Rad50 protein 15

2.4. The Rad50-Mre11-Nbs1 (Xrs2) complex 16

2.4.1. Cellular localization of the MRN complex after DSB in vivo 16

2.4.2. Structural and biochemical analysis 17

2.4.3. The CtIp/Ctp1/Sae2 protein 19

2.4.4. Functions of the MRN(X) complex 19

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OBJETIVOS 21

OBJECTIVES 22

MATERIALES Y METODOS 23

MATERIALS AND METHODS 24

1. Materials and Methods 24

1.1. Strains 25

1.2. Reagents and materials 25

1.3 Buffers 27

2. Methods 27

2.1. Cells manipulation 27

2.1.1. Competent cells production 27

2.1.2. Bacterial transformation 27

2.2. DNA manipulation 28

2.2.1. DNA isolation and quantification 28

2.2.2. Plasmids 28

2.2.3. Radioactive DNA labelling 29

2.3. Purification and analysis of proteins 29

2.3.1. Overexpression of proteins 29

2.3.2. Purification of proteins 30

2.3.2.1. Purification of B. subtilis RecN 30

2.3.2.2. Purification of B. subtilis RecN-His (RecN-HtgBsu) 30 2.3.2.3. Purification of B. subtilis His-PNPase (Htg-PNPaseBsu) 30

2.3.3. Identification of proteins 31

2.3.4. Molecular mass determination 31

2.3.5. Preparative two-dimensional (2D) polyacrylamide

gel electrophoresis (PAGE) 31

2.4. Biochemical assays 32

2.4.1. Measurement of protein-DNA interactions 32

2.4.1.1. Measurement of nuclease and polymerase activities 32

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2.5. In vivo assays 32

2.5.1. Viability assays 32

2.5.2. Fluorescence microscopy analysis 32

2.5.2.1. Analysis of chromosomal segregation in ΔpnpA mutants 32 2.5.2.2. Effect of a ΔpnpA mutation in the localization of RecN

after damage induction 33

2.5.3. Western-blot analysis of RecN and RecA expression 33

RESULTADOS 34

RESULTS 35

Chapter 1. Characterization of PNPase nuclease activity 35 1.1. Characterization of the nuclease activity exhibited by purified

RecN protein 35

1.1.1. Purified RecN shows a Mn2+ -dependent exonuclease activity 35 1.1.2. The nuclease activity is not associated with RecN itself 36 1.1.3. The exonuclease activity has 3’  5’ polarity and is

Mn2+ -dependent 37

1.1.4. PNPase is responsible of the 3’ 5’ ssDNA exonuclease activity 37 1.2. PNPase is able to bind and degrade ssDNA with high affinity 38 1.3. The end processing carried out by PNPase is distributive 41 1.4. Effect of Pi and divalent cations on ssDNA exonuclease activity

of PNPase 42

1.5. Effect of Mn2+ and Mg2+ on PNPase exoribonuclease activity 44 Chapter 2. Characterization of PNPase polymerase activity 45 2.1. PNPase polymerization on ssDNA60 in the presence of rNTP/dNTP

and Pi 45

2.2. PNPase polymerization is markedly increased with the addition

of dNDPs 46

Chapter 3. Comparison between B. subtilis and E. coli PNPases 47 3.1. Bacterial PNPase shows Mn2+ -dependent phosphorolysis 47

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3.2 Bacterial PNPase shows template independent dNDP polymerization

of ssDNA 49

3.3 Bacterial PNPase can elongate dsDNA 3’-ends 51

Chapter 4. In vivo studies of B. subtilis PNPase 53

4.1. PNPase could be needed for DNA repair 53

4.2. Mechanism of PNPase effect on DNA repair 54

4.3. Epistasis analysis of ΔpnpA mutants 56

4.3.1. A Δku mutation is epistatic with pnpA 61

Chapter 5. Modulators of PNPase nuclease and polymerase activity 62

5.1. PNPase is modulated by rATP and dATP 62

5.2. PNPase is among the first proteins that interact with DNA ends 63

5.3. RecN increases PNPase polymerase activity 64

5.4. Modulator effect of RecA on PNPase activities 65

5.4.1. RecA stimulates PNPase nuclease activity 66

DISCUSION 68

DISCUSSION 72

1. PNPase is involved in nucleic acid metabolism 72

1.1. PNPase is involved in RNA processing 72

1.2. PNPase is involved in DNA processing 74

1.3. PNPase is required for DSB repair 74

2. PNPase activities are regulated by Pi, metal ions (Mg2+/ Mn2+)

and d/rNTPs 75

2.1. PNPase requires Mn2+ and low Pi for its activity on ssDNA 75 2.2. PNPase requires Mg2+ and high Pi for its activity on RNA 75

3. Proposed role for PNPase in DSB repair 76

4. DNA DSB repair pathways 78

4.1. DSB recognition and basal processing is conserved 78 5. Are RecN “contaminants” genuine components of the

repairsome? 80

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CONCLUSIONES 81

CONCLUSIONS 83

BIBLIOGRAPHY 84

ANNEXE 93

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INDEX OF FIGURES

Figure 1. Model for generation and repair of two types of DSBs. 4 Figure 2. Model for NHEJ in bacteria and eukaryotes. 5 Figure 3. Model of homologous recombination in B. subtilis. 6 Figure 4. Temporal order of protein assembly at a DSB in

B. subtilis cells. 7

Figure 5. Architecture of SMC proteins. 10

Figure 6. Secondary structure prediction and motifs of SMC and

SMC-like proteins. 10

Figure 7. Characteristics of RecNBsu protein. 12

Figure 8. Domain architecture of Rad50. 15

Figure 9. Domain structure of Mre11. 17

Figure 10. Structure of Nbs1/Xrs2. 18

Figure 11. Nuclease activity present in the purified RecN sample. 35 Figure 12. Analysis of the proteins present in the RecN sample prior

purification by FPLC. 36

Figure 13. The 3’-to-5’ ssDNA exonuclease activity is due to PNPase. 37 Figure 14. Binding and degradation of ssDNA by increasing

concentrations of PNPase.

38

Figure 15. PNPase promotes limited degradation of ssDNA. 39 Figure 16. PNPase nuclease activity needs a 3’OH end. 40 Figure 17. PNPase activity on ssDNA is distributive. 41 Figure 18. Effect of Mg2+ and Pi in the degradative activity

of PNPase. 42

Figure 19. Effect of Pi concentrations in the exonuclease activity

of PNPase 43

Figure 20. Mn2+ inhibits the exoribonuclease activity of PNPase. 44 Figure 21. B. subtilis PNPase is able to polymerize in presence

of rNTP or dNTP. 45

Figure 22. Polymerization of B. subtilis PNPase is optimal in the

presence of dNDPs. 46

Figure 23. Mn2+-dependent phosphorolysis and template-independent synthesis of ssDNA catalyzed by PNPase with oligo (dA)

substrate. 47

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Figure 24. Mn2+-dependent phosphorolysis and template-independent synthesis of ssDNA catalyzed by PNPase with oligo

A substrate. 48

Figure 25. Inhibition of phosphorolysis and polymerization by

high concentrations of Pi and dADP, respectively. 49 Figure 26. Phosphorolysis and/or template-independent synthesis

of ssDNA in the presence of Mg2+ and Feions. 50 Figure 27. PNPase catalyzes template-independent polymerization

of 3’ tailed duplex DNA. 51

Figure 28. PNPase polymerization on dsDNA with 3’-protruding,

blunt or 3’- recessed ends. 52

Figure 29. Survival of ΔpnpA cells exposed to a chronic dose of H2O2,

MMS, 4NQO or MMC. 53

Figure 30. Chromosomal segregation in wt and mutant B. subtilis cells. 55 Figure 31. Survival of strains exposed to an acute dose of MMS

or H2O2. 58

Figure 32. Survival of strains exposed to a chronic dose of H2O2,

MMS or MMC. 59

Figure 33. Survival of the null ku mutation in the ΔpnpA context. 61 Figure 34. PNPase activities are modulated by ATP and dATP. 62 Figure 35. PNPase promotes assembly of a discrete RecN

focus per nucleoid. 63

Figure 36. PNPase polymerase activity is modulated by RecN. 64 Figure 37. RecA interaction with ssDNA•RecN•ATP•Mg2+ networks. 65 Figure 38. RecA stimulates PNPase degradative activity. 66 Figure 39. Effect of RecA on PNPase exoribonuclease activity. 67 Figure 40. Structure of S. antibioticus PNPase 73 Figure 41. Possible mechanism of action of RecN in PNPase polymerase

activity 77

Figure 41. Proposed model for the role of the bacterial RecN-PNPase

complex in DSB repair 79

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INTRODUCCION

La estabilidad del material genómico es crucial para la estabilidad de la información genética y la supervivencia celular. Por esta razón muchas funciones celulares se encargan de garantizar que la replicación, la reparación, la recombinación y la segregación cromosómica ocurran con alto grado de fidelidad. Los daños en el ADN pueden ocasionar alteraciones a nivel de replicación, detener la progresión del ciclo celular, para finalmente desencadenar una respuesta transcripcional que incremente las opciones de supervivencia. Estos daños son mayoritariamente reparados por procesos dedicados especificamente a tales actividades, como son: reparación mediante la excisión de bases (base excision repair, BER), excisión de los segmentos de ANDcs que llevan el daño (nucleotide excision repair, NER), eliminación de bases erroneamente incorporadas (mismatch repair, MMR) (Friedberg, 2006 #2). Cuando estos sistemas específicos fallan, la reparación vía RH o UENH es la última opción. Los daños en el ADN pueden causar frenos en el progreso de las horquillas de replicación dando lugar a la formación de zonas de ADNcs sin que haya discontinuidad en la otra hebra (single strand gap). También se puede provocar el colapso de la horquilla de replicación cuando se producen cortes en una de las hebras del ADN, lo que origina un ADN de cadena doble (ADNcd) cortado (one-ended double strand break). Aquellos agentes que producen cortes en el ADN, en ausencia de replicación, dan lugar a cortes en ambas hebras del ADN (two-ended double strand breaks). Estos últimos pueden ser reparados por RH ó UENH, mientras que los primeros sólo por RH. La reparación vía RH estrictamente necesita un ADNcd homólogo para poder llevar a cabo la reparación, sin embargo, en ausencia de una copia de ADNcd homóloga la reparación sólo puede llevarse a cabo vía UENH. La reparación por UENH religa directamente los dos extremos del ADN roto, y puede generar la pérdida ó ganancia de algunas bases en el punto de sellado (Pitcher et al, 2007).

En eucariotes las primeras proteínas reclutadas al ADN dañado se encuentran formando el complejo Mre11, Rad50 y Xrs2 (en levaduras) ó Nbs1 (en mamíferos), MRX(N) (Lisby et al, 2004). La reparación por RH en B. subtilis se inicia con el reconocimiento de una RDC por parte de RecN, lo que ocasiona su re-distribución a nivel celular, pasando de una localización difusa a formar un foco claramente distinguible en el lugar en el que ha ocurrido la ruptura (Kidane et al, 2004). Posteriormente, los extremos 3’ del ADN son procesados por la exonucleasa RecJ, junto con una helicasa de la familia RecQ (RecQ ó RecS), ó por el complejo nucleasa-helicasa AddAB (contraparte del complejo RecBCD de E. coli ó AdnAB de Mycobacterium tuberculosis) (Sanchez et al, 2007). Luego, SsbA (homóloga de SSB de E. coli) se une a los extremos 3’ del ADNcs mientras que RecN lo hace a los extremos 3’-OH (Sanchez et al, 2008). RecN facilita la aproximación de los extremos de ADN, y promueve la formación de un centro de reparación (CR) (Kidane et al, 2004; Sanchez et al, 2008). Posteriormente, RecO, RecR, RecA, y más tarde, RecF y RecU co-localizan con RecN en el CR (Kidane et al, 2004). RecA, con ayuda de las proteínas mediadoras (RecOR), desplaza a SsbA del ADNcs y polimeriza sobre él.

RecA al unirse al ADNcs da lugar a un filamento nucleoproteico dextrógiro dinámico, que se corresponde con la forma activa de RecA (1 RecA cada 3 nt, ó 6 RecA por cada vuelta de hélice). RecA activada cataliza la invasión de la cadena de ADNcs en el ADNcd homólogo intacto, dando lugar a la formación de un intermediario de recombinación, denominado bucle D (D-loop). En un segundo paso, se genera una estructura de Holliday, sobre la que se cargan las translocasas RecG ó RuvAB. RuvAB promueve el cargado de la resolvasa RecU (ortólogo de RuvC de E. coli), que junto con RuvAB ó con RecG, resuelven la estructura de Holliday, que es finalmente sellada por una ligasa.

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Introduction

Análisis in vitro indican que tanto la proteína eucariota Rad50, como la proteína bacteriana RecN, pertenecientes a la superfamilia de proteínas de mantenimiento de la estructura cromosómica (MEC), se encargan de unir y atar extremos de ADN (Moreno- Herrero et al, 2005; Sanchez et al, 2008). Además, ambas proteínas se encuentran involucradas en los estadios iniciales de la recombinación homóloga lo que implica su participación en la identificación del daño en el ADN, y el posterior reclutamiento de otras proteínas que participarán en su reparación. Sabiendo que en eucariotas la proteína Rad50 se encuentra formando un complejo con la proteína Mre11, cuya función es degradar los extremos 3’ del ADN, en una reacción dependiente de Mn2+ y regulada por ATP (Paull & Gellert, 1998; Trujillo et al, 1998), es lógico pensar que siendo homólogas las funciones de RecN y de Rad50, RecN debería estar asociada con una nucleasa que se encargara del procesamiento inicial del ADN. La búsqueda de tal función, su identificación y su caracterizacion genética, citológica y bioquímica fueron objetivos de la presente tesis doctoral.

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INTRODUCTION

1. Mechanisms of DNA repair

The faithful replication and maintenance of the genome(s) are of primary importance for all living organisms. However, free radicals generated during essential metabolic processes or exposure to natural or man-made mutagens can damage the DNA. DNA damages can be repaired by several dedicated error-free pathways (e.g., base excision repair [BER], nucleotide excision repair [NER], mismatch repair [MMR], etc.). Base excision repair (BER): repairs damage to a single base caused by oxidation, alkylation, hydrolisis or deamination, etc. The damaged base is removed by a DNA glycosylase, the AP endonuclease, which recognizes the missing base, and cuts the phosphodiester bond. DNA polymerase resynthesise the single-stranded gap and DNA ligase performs the final sealing step.

Nucleotide excision repair (NER): recognizes bulky distortions in the shape of the DNA double helix as purine adducts, thymine dimers, and 6-4-photoproducts that can be generated by several sources including chemicals (as 4-Nitroquinoline-1-oxide [4NQO], ultraviolet [UV] light, mitomycin C [MMC] etc. Recognition of these distortions leads to the removal of a short single-stranded DNA segment that includes the lesion, creating a single-stranded gap in the DNA, which is finally filled by a DNA polymerase, which uses the undamaged strand as a template. DNA ligase performs the final sealing step.

Mismatch repair (MMR): recognizes erroneous insertions, deletions and miss- incorporation of bases that can occurr during DNA replication. The MMR machinery distinguishes the newly synthesised from the template (parental) strand by a poorly characterized mechanism. The deformitiy caused by the mismatch is recognized, and a short single-stranded DNA segment that includes the mismatch is excised. The removal process involves more than just the mismatched nucleotide itself. A few or up to thousands of base pairs of the newly synthesised DNA strand can be removed. DNA polymerase is used to re-fill the gap and DNA ligase performs the final sealing step.

When those specific systems fails and the damage remains in the ADN, the replication machinery stops, and one-ended DSB, that will be repaired exclusively by HR, will be generated. Certain type of DNA damages will directly generate two-ended double strand breaks (DSBs),which are potentially catastrophic lesions (See Figure 1), if unrepaired they will lead to loss of genetic information and cell death (Pierce et al, 2001; So et al, 2004). The two-ended DSBs will be repaired by homologous recombination or non- homologous end-joining(NHEJ).

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Introduction

Figure 1. Model for generation and repair of two types of DSBs. One-ended DSBs can arise spontaneulsy during S phase of the cell cycle, or when a replication fork collides with lesions caused by MMS and UV. These DSBs may be repaired accurately by HR. On the other hand, two-ended DSBs can arise spontaneusly throghout the cell cycle, or by exposure to DSB inducing agents, like X-rays, or treatment with I-SceI nuclease. These DSB can be repaired by HR or NHEJ.

One-ended DSBs are repaired by HR, whereas, two-ended DSBs are repaired by HR or NHEJ. The major difference between these two pathways is that HR utilizes and intact DNA sequence homologous to the DSB site to guide the repair event and accurately restore the DNA structure and sequence. In contrast, NHEJ requires minimal base- pairing at the junction. Then, the two DNA ends are brought together and ligated directly in an error-prone mechanism that is capable of producing genetic alterations ranging from the loss or addition of several nucleotides to chromosome translocations.

1.1 NHEJ

NHEJ was first discovered in eukaryotes. This DSB repair pathway is also involved during normal physiological processes such as V(D)J joining of immunoglobulin genes. In mammalian cells ~ 50 to 70% of the DSBs created by I-SceI endonuclease can be repaired by NHEJ with the remaining fraction being repaired by HR (Liang et al, 1998). It has been proposed that repair pathway choice is regulated by cell cycle phase. For example, if mammalian cells are in the G1 phase of the cell cycle the NHEJ pathway is preferentially used (Frank-Vaillant & Marcand, 2002); whereas HR is preferred during the S/G2 phase, when the homologous template for HR is available (the sister chromatid is only present in this phase of the cell cycle) (Johnson & Jasin, 2001). Despite a cell cycle preference, HR and NHEJ can nevertheless be coupled for the repair of a single DSB in mammalian cells (Richardson & Jasin, 2000), indicating that the two repair pathways are not completely restricted to different cell cycle phases and that other factors could be influencing pathway choice.

In eukaryotes, NHEJ proceeds in a stepwise manner beginning with limited end- processing by the MRE11/RAD50/NBS1 (MRN) complex, end-binding by heterodimeric Ku, comprising the Ku70 and Ku80 subunits, and recruitment of the DNA-dependent protein kinase catalytic subunit (DNAPKcs), forming the trimeric DNA-PK holoenzyme.

Once bound to broken ends, DNA-PK is activated and it phosphorylates itself and other targets including RPA, WRN, and Artemis. In the final step, DNA ligase IV, with its binding partners XRCC4 and XLF, seals the break (Figure 2).

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Figure 2. Model for NHEJ in bacteria and eukaryotes. DNA damage generates DSB.

In higher organisms, broken DNA ends are bound by a heterodimeric complex of Ku70 and Ku80, which also recruit the catalytic subunit of protein kinase (DNA PKcs). In bacteria the Ku-like protein YkoV, binds as a homodimer to the DNA end. In both eukaryote and bacteria the Ku-like protein diffuses along DNA away from the break, enabling the loading of further Ku-like complexes. Next, the DNA ends associate, through interactions bteween Ku-like molecules bound to different DNA ends. The DNA end-joining enzymes, DNA ligase IV in higher organisms and YkoU in bacteria are then recruited by Ku and YkoV, and the broken ends are re-joined (Hiom, 2003) .

In B. subtilis and in other microorganisms a functional bacterial NHEJ repair apparatus has been identified and characterised (Bowater & Doherty, 2006; Pitcher et al, 2007;

Shuman & Glickman, 2007). Here the DSBs created by random breaks or by site-specific incision through HO endonuclease are mainly repaired by HR, but in their absence or in the absence of a DNA remplate NHEJ can repair the two-ended DSBs (Ayora et al, 2010;

Bowater & Doherty, 2006; Pitcher et al, 2007). Unlike eukaryotes, B. subtilis has just a single Ku-like protein (also termed YkoV), homologue of the mammalian heterodimer Ku70/Ku80, and a protein with limited homology to Ligase IV (LigD, also termed YkoU) protein. The mechanism of action can be seen in (Figure 2).

1.2. DSB response in bacteria

1.2.1. Homologous recombination in B. subtilis

Using genetic, cytological and biochemical approaches the HR pathway was subdivided into five general steps:

A. Recognition of the break site and initial response to DNA damage.

B. End-processing at the break (generation of ssDNA) and DSB “coordination”.

C. Loading of the strand exchange protein RecA onto ssDNA.

D. Strand exchange between broken and non-broken sister chromosome, branch migration and resolution.

E. Replication fork re-start and chromosomal segregation.

Genetic studies have helped to classify the B. subtilis recombination genes, except for recA, within seven different epistatic groups (α, β, γ, δ ε, ζ and η groups). RecA plays a central role, hence null recA mutant (ΔrecA) cells are severely affected by DNA damaging agents. Mutations in genes classified within the α (recF, recO and recR), δ (recN), ε (recU, ruvA and ruvB genes) and η (recG) epistatic groups markedly affect the viability of cells exposed to DNA damaging agents, whereas mutations in genes

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Introduction

classified within the β (addA and addB), γ (recH and recP) and ζ (recS, recQ and recJ) epistatic groups slightly reduce the viability of cells exposed to DNA damaging agents (Fernandez et al, 2000).

These functions act in one or several of these general steps, with RecA being the central player of the recombination reaction. In stage A, RecN a protein of the δ group is required, whereas stage B is mainly performed by proteins included within the β (AddAB) and ζ (RecJ-RecQ or RecS) epistatic groups. The stage C can be performed by proteins included within the α (RecFOR) or β (AddAB) groups. Stage D is mainly performed by RecA, and is positively and negatively regulated by several RecA modulators (RecF, RecO, RecR, RecX and RecU). The final steps E and F include proteins classified within the ε (RuvAB-RecU) and η (RecG) epistatic groups. (See Figure 3).

The recA, recF, recG, recJ, recN, recO, recQ, recR, ruvA and ruvB genes have their counterpart in E. coli in genes with identical name. The addAB and recU genes, which have their counterpart in E. coli recBCD (recBCDEco) and ruvCEco genes respectively, have a different name, because they are mechanistically different from their E. coli counterparts (Ayora et al, 2010; Sanchez et al, 2007). The recS gene has no counterpart in E. coli and the genes linked with recH and recP mutations are unknown.

Figure 3. Model of homologous recombination in B. subtilis. A. After the introduction of a DSB RecN recognizes the damaged ends. If the ends already have a ssDNA region, the single strand binding protein (SsbA) might bind to them.

B. The 5’-ends are resected by AddAB or recJ in concert with a RecQ-like enzyme (RecQ or RecS) with SsbA binding to the ssDNA tails that result from the resection.

RecN tethers the DNA ends and promotes loading of the RecA mediators.

C. RecO alone or the (RecN)-RecR-RecO complex could promote the disassembly of the SsbA protein and the loading of recA onto ssDNA. RecA-polymerized (RecA threads) onto SsbA coated ssDNA, promotes the search for a homologous template, and promotes DNA-strand invasion. RecA modulators (RecF, RecX, PcrA, RecU, RecO, RecR, RecF), control RecA by modulating its filament extension.

D. The invading strand provides the template for primosome loading. Then the replicase primes DNA synthesis using the intact homologous chromosome as template to restore the lost genetic information.

E. Capture of the second DNA derived form the other end of the DSB, led to a double HJ, that can be migrated via RecG or RuvAB.

To understand the temporal order of events during HR a study of the localisation of different recombination and repair proteins in living cells, after the induction of DNA damage, was performed (see Figure 3). Recombination proteins were fused either at their 5´-end or 3´-end with a gene coding for a fluorescent protein (gfp, cfp, yfp, etc.), and the fused gene was used to replace the wt gene in its natural location. From those, only fusions in recA, recF, recG, recO, recQ, recN, recR, and recU genes were informative (Kidane & Graumann, 2005; Kidane et al, 2004; Mascarenhas et al, 2002; Mascarenhas

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et al, 2005). Under all conditions, the fused protein was fully functional, as judged by the ability of cells expressing only the fusion to survive MMC treatment (Kidane et al, 2004;

Sanchez et al, 2005). Figure 4, shows the obtained results.

Figure 4. Temporal order of protein assembly at a DSB in B. subtilis cells. A. RecN recognizes a DSB and assembles onto DNA, forming discrete repair foci (RC) 15 min upon induction of a DSB (time 0). B. The 5’ resection of the DNA ends, either by the AddAB complex or by a RecQ-like enzyme (RecQ/RecS) in concert with RecJ, leads to the recruitment of multiple DSB towards a discrete RecN-promoted RC (15-30 min). After DNA ends resection, SsbA should bind and protect ssDNA. C. 30-45 min after DSB induction RecO, RecR and RecA are recruited to RecN-promoted RC. At a latter stage (60 min), RecF and RecX are recruited to the RC. E. At the same time, during stages C and D other repair processes [e.g. SbcC-dependent repair and replisome assembly via PriA, DnaD, DnaB, DnaIC] take place. D. RecU recruitment was visualized 90 min after the damage. F.

RipX/CodV tyrosine recombinase resolves the dimeric chromosomes, and resumption of cell proliferation takes place at 180 min after damage induction.

1.2.1.1. Break recognition and initial response to DNA damage in B. subtilis

This work will focus on the poorly characterized initial step of HR during DSB repair. The protein(s) involved in DNA damage recognition and end processing, necessary to start with the HR will be reviewed. By cytological studies it has been shown that B. subtilis exponentially growing cells, in minimal medium, show a low basal level of RecN-YFP fluorescence diffused throughout the cell (Kidane et al, 2004) . However, 15 min after the induction of a DSB, generated either by the exposure to ionising radiation, the addition of nalidixic acid or MMC, or by the action of a site-specific HO-endonuclease, RecN re- localizes, from a diffuse distribution, to form a discrete focus or RC per nucleoid in wt cells (Kidane et al, 2004; Krishnamurthy et al, 2010; Shintomi & Hirano, 2007). This foci formation was dose-independent (a single focus per nucleoid in cells treated with low or high doses of MMC) and takes place in the majority of the cells (> 75 % of total cells) (Kidane et al, 2004).

In untreated, exponentially growing wt cells, there is a low basal level of fluorescent focus. However, RecN foci were present in several exponentially growing recombination mutants, as in ~ 35% of ΔrecA cells, in ~ 5% of ΔrecU cells, or in ~ 2% of addAB ΔrecJ triple mutant cells not treated with any DNA damaging agent (Kidane et al, 2004;

Sanchez et al, 2006). These findings suggest that sites of DNA damage accumulate in the absence of RecA, RecU or AddAB and RecJ during normal growth.

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Introduction

RecO, RecR or RecA are simultaneously recruited and re-localized with RecN 30 – 45 min after induction of DSBs. RecF is recruited to the RecN-RecOR-RecA multiprotein complex 60 – 90 min after induction of DSBs, these data show that RCs are formed sequentially in response to DNA damage in B. subtilis (Figure 4). RecN forms discrete foci, but RecA fails to form foci in the absence of end processing (addA ΔrecJ cells), suggesting that RecN acts prior end-processing and RecA after it. In the absence of RecN, RecO forms aberrant and patchy foci, indicating that RecN is an early detector of DSBs and is important for the proper formation of discrete RCs. Recently it was shown that SsbA physically interacts with RecO (Manfredi et al, 2008; Manfredi et al, 2010), raising the posibility that SsbA bound to the ssDNA might recruit RecO towards RecN- mediated focus. RecN, RecO and RecR have been shown to form DSB-induced foci in the absence of RecA protein (Kidane et al, 2004)and RecN, RecO and RecA foci are formed in the absence of RecF, but RecF fails to form foci in the absence of RecO (Kidane & Graumann, 2005; Kidane et al, 2004). This is consistent with biochemical and biophysical analyses leading to conclude that RecN, which is the first protein recruited to the DSB, appears to organise the assembly of a single network of protein-protein and protein-ssDNA complexes, and may even recruit several DSBs into one single RC (Kidane et al, 2004; Sanchez et al, 2008; Sanchez et al, 2006). There are also few RCs formed in ΔrecN cells, suggesting that an “unknown factor(s)” might partially organise the RCs in the absence of RecN.

1.2.1.2. DNA end-processing in B. subtilis

In order for RecA to mediate HR, the DSB ends must be modified leaving a 3´-termined ssDNA to which RecA can bind (Figure 3). In B. subtilis cells, the DNA ends are processed by redundant avenues consisting of nucleases and DNA helicases: the AddAB enzyme, which comprises one set of helicase motifs and two distinct nuclease activities, unwinds and degrades both DNA strands during translocation. Upon recognition of a short sequence χ site, 5'-AGCGG-3'), AddAB responds by attenuating the 3' to 5' nuclease to allow the generation of a 3'-terminated ssDNA starting at χ (Dillingham &

Kowalczykowski, 2008; Yeeles & Dillingham, 2010). A second pathway involves one of the RecQ-like (RecQ or RecS) DNA helicases, in concert with the putative RecJ ssDNA exonuclease, to generate a 3´-ssDNA overhang. A third pathway involves the RecD DNA helicase, in concert with the putative RecJ ssDNA exonuclease, to generate a 3´-ssDNA overhang. This is congruent with genetic studies revealing that the double mutant addAB ΔrecJ has a synergistic effect, with survival after DNA damage induction reduced to the levels seen with ΔrecA cells (Sanchez et al, 2006).

Cytological studies have revealed that in the absence of AddAB and RecJ, which are the

“major” exonucleases, RecN formed two to four foci instead of one focus per nucleoid (Kidane & Graumann, 2005; Sanchez et al, 2006). This finding suggests that: i) AddAB and RecJ nucleases play a redundant role in vivo, ii) AddAB and/or RecJ are necessary for the formation of a RecN RC, and iii) concomitantly with end-processing, RecN binds to the ssDNA tail of the duplex molecule at the DSB, protects the 3´-OH end and is likely to facilitate the tethering of these DNA ends together to form mainly one discrete focus or RC. (Kidane & Graumann, 2005; Kidane et al, 2004; Sanchez & Alonso, 2005) It seems therefore, that 3’-end processing takes place between the damage recognition by RecN protein and before RecA loading (Figure 3).

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1.3. DSB recognition and initial response to DNA damage in eukaryotes

The mammalian Mre11/Rad50/Nbs1 (MRN) or budding yeast Mre11/Rad50/Xrs2 (MRX) complexes, appear to be the earliest sensors of DSBs, by directly binding to DNA ends(Lisby & Rothstein, 2005). Basal processing of the ends is performed by the human MRN complex in concert with CtlP, or MRX in concert with Sae2 allowing latter recruitment of ATM to the DSB site (Mimitou & Symington, 2009)..

The response mechanism(s) to DSBs in eukaryotes is centred on kinases, like ataxiatelangiectasia mutated (ATM), ataxiatelangiectasia and Rad3 related (ATR), and DNA-PK (DNA-dependent protein kinase) in mammals, or Mec1 and Tel1 in budding yeast. These protein kinases trigger cell cycle arrest following DNA damage, therefore allowing DNA repair to take place (Kurz & Lees-Miller, 2004).

2. Structural maintenance of chromosomes (SMC) proteins

2.1. Architecture of SMC proteins

In all living organisms a speciallized SMC protein(s) recognize(s) and tethers the DNA ends (e.g., Rad50, SbcC, RecN, SbcE). SMC are usually large polypeptides, 1000 to 1300 amino acid long, with a characteristic domain organization. Two nucleotide-binding motifs, known as the Walker A and Walker B motifs, at the N-terminal and C-terminal domains, respectively, separated by a long coiled-coil region. In the dimeric form the coiled-coil regions are connected by a non-helical sequence. An SMC monomer can fold back on itself through antiparallel coiled-coil interactions, creating an ATP-binding ‘head’

domain at one end, and a hinge domain in the other (Figure 5A). Two monomers can associate with each other at the hinge domain to adopt different conformations, like open-V, closed-V and ring-like molecules (Figure 5B and 5D).

Structural studies had demonstrated that ATP binding to the SMC head domains allows the formation of a nucleotide sandwich dimer. In these structures ATP binds to a pocket formed by the Walker A and Walker B motifs from one SMC. Mutational analysis of key residues in B. subtilis SMC protein support the idea that head-head engagement is essential for ATP hydrolisis; a mutation in the Walker A motifs blocks ATP binding, whereas a mutation in the Walker B motif allows ATP binding but blocks head-head engagement and ATP hydrolysis (Hirano et al, 2001).

Almost all bacterial genomes encode a single smc gene whose product forms a homodimer and its functional form associates with two non-SMC regulatory subunits (ScpA-ScpB) to form a functional condensin complex. In eukaryotes, however, there are at least six different SMC proteins that form heterodimers in specific combinations.

SMC1-SMC3 is the core of the cohesin complex that mediates sister-chromatid cohesion during mitosis, and is loaded onto DNA during replication (Harvey et al, 2002). The SMC2-SMC4 complex is part of the condensin complex, and functions in the condensation of chromosomes during mitosis (Losada & Hirano, 2001). Whereas, the SMC5-SMC6 complex forms a third complex that has been implicated in DNA repair and checkpoint responses. Each of the heterodimers associates with a distinct set of non- SMC regulatory subunits to form the functional complex (Figure 5C).

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Introduction

Figure 5. Architecture of SMC proteins. A. Basic architecture of SMC protein dimer. Each subunit self-folds by antiparallel coiled-coil interactions (indicated by arrows) to form a hinge domain at one-end and an ATP-binding head at the other. Interactions through the hinge domain of two subunits allow dimerization and produce a V-shaped molecule. B. Electron micrographs of B. subtilis SMC homodimers show variety of conformations. Scale 50 nm. C.

SMC-protein complexes in bacteria and eukaryotes. D. Electronic microscopy photographs of human condensin (left panels) and cohesins (right panels) (Hirano, 2006).

Although the aminoacid sequence of the Walker A and Walker B motifs are highly conserved, the hinge region is very characteristic for each class of SMC protein. For example, the canonical SMC protein has a glycine rich motif in this region, while the members of the Rad50-SbcC sub-family have a cysteine rich motif (Hopfner et al, 2002;

Mascarenhas et al, 2006). On the other hand, RecN posses a leucyne rich motif (Sanchez et al, 2008), whereas, SbcE hinge motif is poorly defined (Figure 6)

Figure 6. Secondary structure prediction and motifs of SMC and SMC-like proteins.

The secondary structure of B. subtilis SMC-like proteins (SMC, RecN, SbcC and SbcE) and its comparison with E. coli SbcC and human Rad50 were made using the program COILS.

The Walker A and B motifs, and the CXXC and GGXXXGG motifs needed for dimerization of Rad50, SbcC, EcoSbcC and SMC proteins are shown. The putative dimerization (Leu rich) domain of RecN is also indicated (Sanchez et al, 2008). Unlike the head-head interaction, that requires ATP binding and hydrolysis, the hinge-hinge interaction is very strong and occurrs independently of ATP. It has been shown that mutations of glycine residues in B.

subtilis SMC, destabilized the dimerization of the monomers (Hirano et al, 2001).

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2.1.2. Effect of ATP binding and hydrolysis in SMC conformation

The enzymology of SMC proteins has been studied using B. subtilis SMC as the model system. A SMC homodimer can interact with both ssDNA and dsDNA, displaying an intrinsic DNA-independent ATPase activity as well as a DNA-stimulated ATPase activity (Hirano et al, 2001; Hirano & Hirano, 1998). When hinge mediated dimerization is disrupted, the resulting monomer loses its capabililty to interact with any type of DNA (Hirano & Hirano, 2002). Also, it seems that ATP has little effect on DNA binding by SMC, however, when a mutation that slows ATP hydrolysis and stabilizes head-head engagement is introduced, ATP-stimulated DNA binding becomes detectable. So it is probable that the ATPase cycle of SMC proteins has an important role in their dynamic interaction with DNA by modulating the cycle of head-head engagement and disengagement (Hirano & Hirano, 2002; Hirano, 2006).

2.2. B. subtilis SMC-like proteins 2.2.1. B. subtilis RecN

The recN gene is widespread in free-living bacterial species. The only known absences are in phylum Chlamydiae and in bacteria of the Mollicutes Class (Rocha et al, 2005).

Mutations in the recN gene genes of E. coli (Sargentini & Smith, 1986)

,  

B. subtilis (Alonso et al, 1993; Sanchez et al, 2006), Helicobacter pylori (Wang & Maier, 2008), Neisseria gonorrhoeae (Skaar et al, 2002), and Deinococcus radiodurans (Narumi et al, 1999), confered DNA damage repair defects, which indicates that the protein develops a key role in DNA repair processes.

B. subtilis RecN is a 64.4 kDa protein, with a structural organization that resembles SMC proteins, composed by an SMC head domain and a coiled-coil region that is much shorter than that of SMC or Rad50/SbcC proteins (Figure 6). The central region of RecN contains around 50 amino acids, and has no homology with any of the hinge regions reported for SMC or SMC-like proteins, because it is formed by leucine zippers motifs, instead of glycine rich regions or CXXC motifs. It has been hypothesised, that this region must correspond to RecN dimerization domain, although it has not been tested experimentally. However, it has been demonstrated that RecN in solution is able to multimerise in homo-octamers that can complex with ssDNA molecules (Kidane et al, 2004; Sanchez et al, 2008).

RecN is involved in the initial steps of homologous recombination as has been shown by fluorescence microscopy analysis. In vivo, RecN accumulates at defined DSBs 15-30 min after DNA damage induction, being the first protein recruited to the site of lesion, and forming one defined foci per nucleoid (Figure 7A). However, about 3 hours after the DSB induction, the RecN foci disappear and cells resume growth, which indicates that RecN accumulates transiently at the DSB. It was also demonstrated that RecN is necessary for the adequate organization of the repair centers, because its presence is a pre-requisite to allow the loading of further repair proteins like RecO and RecF (Kidane et al, 2004). So, it has been proposed that RecN could be acting as an initiator of RCs that accumulates at DSB sites and induces or maintains a DNA topology at the DSB that facilitates loading of RecO/F and thus RecA. RecN also could be organizing DSBs in large repair centers based on its ability to form multimers and high-molecular-weight complexes after induction of DSBs (Kidane et al, 2004; Sanchez et al, 2008), in a similar way to SMC proteins that condense DNA or hold sister chromosomes together by embracing DNA with their arms, and also influence DNA topology (Hirano, 2006).

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Introduction

Figure 7. Characteristics of RecNBsu protein. A. Fluorescence microscopy studies of RecN-YFP protein in exponentially growing wt cells, before (left panel) and 60 min after (right panel) the addition of 50 ng/ml MMC.B. DNA binding by RecN. [γ32P] labelled ssDNA60 (1 nM), was incubated with RecN (0.4, 0.8, 1.7, 3.7, 15, 30 and 60 nM) for 30 min at 37°C in buffer C in the absence (lanes 1–9) or presence of 1 mM ADP (lanes 10–18) or 1 mM ATP (lanes 19–27). RecN was not added to the controls in lane 1, 10 and 19. In lanes 10 and 19, (+) denotes that identical results were observed in the presence or absence of a nucleotide cofactor. C. RecN binds ssDNA tails in a duplex molecule. Linear DNA (150 nM) with short ssDNA tails and RecN (10 nM) were incubated for 10 min at 37°C and deposited on freshly cut mica. D. Images of “rosette-like structures”. Linear DNA (150nM in nt) with short ssDNA tails and RecN (10 nM) were incubated for 10 min at 37°C, then ATP (1 mM) was added and incubated for 20 min at 37°C. Scale bar 100 nm. E. RecA-promoted DNA strand invasion in presence of RecN and ATP. Linear DNA (150 nM) with short ssDNA tails and 10 nM RecN were pre-incubated for 10 min at 37°C, then 1 mM ATP was added and incubated for 10 min at 37°C. Homologous circular plasmid DNA (150 nM) and RecA protein (40 nM) were added to the pre-formed ssDNA•RecN•ATP•Mg2+ complex, incubated for another 10 min at 37°C, and deposited on freshly cut mica. Scale bar = 500 nm (inset picture, bar = 100 nm).

RecN as member of the SMC family has an intrinsic ATPase activity, however there is controversy about the ATPase activity of RecN purified from different microorganisms.

Recent biochemical studies revealed that although Deinococcus radiodurans RecN (RecNDra) and B. subtilis RecN (RecNBsu) hydrolyze ATP with similar turnover rates, and that such rates are stimulated 4- to 6-fold by DNA, the RecNBsu ATPase activity is

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stimulated by ssDNA, whereas RecNDra ATPase is increased by the presence of dsDNA (Reyes et al, 2010; Sanchez & Alonso, 2005). On the other side, it has been demonstrated that H. influenzae and A. aeolicus RecN have a weak ATPase activity (Grove et al, 2009), which is not stimulated by the addition of ssDNA or dsDNA. In contrast, RecN from Haemophilus influenzae, Aquifex aeolicus or Bacteriodes fragilis fails to bind DNA. RecNBsu can bindssDNA larger than 30-nt in length, and the complexes formed can vary dependending on the presence or absence of nucleotide cofactor. In the absence of ATP or ADP, a CI is complex formed, whereas, larger complexes, named CII and CIII, are formed when RecN and ssDNA are incubated together with ATP, ADP, dATP or ATPγS (Figure 7B).

AFM studies demonstrate that RecNBsu was able to bind to a duplex DNA with 3’ ssDNA extensions of about 150-450 nt (Figure 7C). It was also found that the addition to ssDNA- RecN of ATP-Mg2+, ADP-Mg2+ or ATPγS-Mg2+ favoured the formation of “rosette like structures”, that could correspond to RecN tethering of ssDNA ends (Figure 7D) (Sanchez et al, 2008). This same tethering activity has been reported for RecNDra with dsDNA, instead of ssDNA (Reyes et al, 2010).

AFM showed that when a pre-formed ssDNA•RecN•ATP•Mg2+ complex was incubated with a homologous circular DNA and sub-saturating RecA concentration, DNA strand invasion by RecA was promoted with more than 12-fold higher efficiency when compared to the yield of joint molecules in the absence of RecN (Figure 7E). Based on this result, it was proposed that RecA interacts with RecN and allows a conformational change of RecN, that favous the strand-exchange reaction and consequently facilitating the RecA search for homology.

2.2.2. Bacillus subtilis SbcC

This protein shares some similar features with Rad50 from eukaryotic cells because both of them have a zinc-bridge motif, and are complexed with a nuclease (Mre11 for Rad50, and SbcD for SbcC). B. subtilis SbcC plays an important role during the repair of inter- strand crosslinks caused by MMC treatment, and also during the repair of DSBs caused by γ irradiation, or mainly those originated by collapsed replication forks. Genetic and epistasis analysis showed that the deletion of sbcC led to a considerable sensitivity to DNA damage; and that SbcC is epistatic with RecA, but not with RecN or with AddAB, which suggests that SbcC acts on DNA repair via homologous recombination (Shrivastav et al), with a different function from RecN or AddAB.

SbcC transiently assembles into discrete subcellular centers on the nucleoids 60 min after the induction of inter-strand crosslinks and DSBs occurring at the replication forks.

SbcC assembled most frequently at the replication machinery, suggesting that SbcC acts on DNA damage, which can block replication (e.g. cross links or base adducts), or lead to fork collapse (e.g. when forks run into ssDNA gaps) (Mascarenhas et al, 2006).

The available information suggest that there is a clear functional distinction between SbcC and RecN, because RecN is never associated with the replication machinery and appears to act at the DNA repair centers away from the replication machinery, while SbcC appears to assemble at the DNA damages that occur at the replication factory.

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Introduction

2.2.3. Bacillus subtilis SbcE (SbcC2,YhaN)

This protein is similar to SbcC, but lacks a zinc-bridge motif and has instead a central domain that does not show any similarity with the reported SMC hinge domain. The sbcE gene (yhaN) is downstream of yhaO that codes for SbcF (a SbcD-like protein). Genetic analysis had shown that loss of SbcE renders cells sensitive to MMC treatment (Krishnamurthy et al, 2010).

SbcE is recruited to the DNA uptake machinery during competence. In competent cells SbcE moves between the cell poles, contrasting with RecA protein that remains at the pole containing the DNA uptake machinery (Krishnamurthy et al, 2010)

2.2.4. Comparative analysis of Bacillus subtilis SMC proteins involved in DNA repair

There are several differences between the three SMC-like proteins of B. subtilis involved in DNA repair:

1. SbcC exclusively assembles at the replication machinery in response to DNA damage, whereas RecN assembles at breaks occurring away from the replication forks (Mascarenhas et al, 2006).

2. SbcE can assemble at the replication machinery in some cells, while in most cells, SbcE accumulations are clearly located at different sites on the nucleoid, away from the replication forks. The SbcE foci formation resembles the pattern of RecN foci, with one single focus per nucleoid after DNA damage induction (Krishnamurthy et al, 2010).

3. SbcE accumulations were present in a considerable number of exponentially growing cells (∼10%), whereas SbcC and RecN foci are DNA damage induced. It is not unreasonable to assume that DNA damage and/or collapsing of replication forks are generally occurring in this group of growing cells, and that SbcE is employed to deal with these normal situations. Thus, SbcE appears to be a constitutive system that deals with a low number of DSBs or DNA damages occurring during growth, while SbcC and RecN systems are turned on in response to a dramatic increase in DNA damage (Kidane et al, 2004; Krishnamurthy et al, 2010; Mascarenhas et al, 2006).

4. Dual visualization of SbcE and of RecN showed that cells generally induce (or contain) an SbcE accumulation or a RecN accumulation is response to DNA damage, but rarely contain assemblies of both proteins, supporting the idea that both factors can induce a distinct avenue to DNA repair via homologous recombination, and that cells use either one of these, and rarely both (Krishnamurthy et al, 2010).

Genetic data support the idea that two avenues exist towards loading of RecA, namely one involving RecN, and a further one involving SbcE, because loss of both RecN and SbcE resulted in an exacerbation of the single gene losses. Therefore, although RecN and SbcE act at an early step in DSB repair, they are not epistatic, and SbcE somewhat complements for the loss of RecN during induction of DSBs.In a similar way, the deletion of both, sbcC and sbcE also increases the severity of the single mutations, suggesting that both proteins are necessary for an adequate response to DNA damages (Krishnamurthy et al, 2010).

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2.3 Rad50 Protein

Is the most characterized and studied SMC protein. Rad50 is a very conserved protein and its homologs can be found in archeal, fission and budding yeast as well as in higher metazoans (Hopfner et al, 2000; Williams et al, 2007). Rad50 shares the general structure of SMC proteins (Figure 8). Rad50 consists of a bipartite N-and C- terminal, ATP binding and ATPase domains, respectively, at the ends of a 150-600 Å long coiled- coil region, for eukaryote homologs. In S. cerevisiae, replacement of a conserved lysine residue within the Walker A motif with alanine, glutamate or arginine results in the same DNA damage sensitivity as well as DNA repair (Chen et al, 2005), and meiosis defects observed in rad50 null mutant.

Outside the Walker A and B motifs, the central region of Rad50 is composed of a large coiled-coil structure that can fols back on itself, via a “hinge” region (Hopfner et al, 2001).

This intramolecular interaction allows both Walker motifs to directly interact formig a globular ATP binding and hydrolysis motif. Furthermore, co-expression and electron microscopy revealed that Mre11 binds to the coiled-coil ner the ATPase domain. The

“hinge” region contains a zinc hook composed of the CXXC motif, that allows Rad50 molecules to dimerize (Hopfner et al, 2001). These results suggest an architectural role for the Rad50 coiled-coils in forming metal-mediated bridging complexes between two DNA-binding heads (Figure 8).

Figure 8. Domain architecture of Rad50. A. Walker A and B motifs are located at both ends of the protein (red squares). Mre11 interacts with Rad50 in a region near to the Walker motifs (yellow rectangles). The center of Rad50 is formed by two coliled.coil regios (dark green) that are linked by a “hinge” containing a Zinc-hook (light green rectangle). B.

Schematic dimerisation of Rad50, where Rad50 proteins folds back on itself via a “hinge”

region resulting in the formation of a long flexible arm by the coiled-coil regions. Rad 50 molecules can interact with each other, forming dimers by using the Zn-hook. The interaction between two Rad50 proteins occurs via the “ hinge” regions in the presence of a Zn2+ ion.

On the opposite end of the Zinc hook, a globular head is formed by the Walker A and B motifs and Mre11 interacting domains (Rupnik et al, 2010).

Referencias

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