UNIVERSIDAD AUTÓNOMA DE MADRID FACULTAD DE CIENCIAS
DEPARTAMENTO DE BIOLOGÍA MOLECULAR
Bacillus subtilis RadA/Sms and RecA contribute in concert to double-strand break repair and
natural transformation, and
with DisA to DNA damage tolerance
TESIS DOCTORAL
RUBÉN TORRES SÁNCHEZ
UNIVERSIDAD AUTÓNOMA DE MADRID FACULTAD DE CIENCIAS
DEPARTAMENTO DE BIOLOGÍA MOLECULAR
Bacillus subtilis RadA/Sms and RecA
contribute in concert to double-strand break repair and natural transformation, and with
DisA to DNA damage tolerance
Memoria presentada por Rubén Torres Sánchez, Licenciado en Biología, para optar al grado de Doctor en Biociencias Moleculares
por la Universidad Autónoma de Madrid
Director: Tutor:
Juan Carlos Alonso Navarro Mario Mencía Caballero
CENTRO NACIONAL DE BIOTECNOLOGÍA-CSIC
Madrid, 2019
El trabajo presentado en esta memoria ha sido realizado en el Departamento de Biotecnología Microbiana del Centro Nacional de Biotecnología (CNB), perteneciente al Consejo Superior de Investigaciones Científicas (CSIC), bajo la dirección del doctor Juan Carlos Alonso Navarro.
Rubén Torres Sánchez ha disfrutado de una beca del Programa Internacional de Becas de Doctorado “La Caixa-CNB”, convocatoria 2015, perteneciente a la Obra Social La Caixa, y una beca
“Short-Term Fellowship” (Reference 7425)
perteneciente a la European Molecular Biology
Organization.
A mi familia
Index
Index
Abstract 25
Resumen 29
Abbreviations 33
1. Introduction 37
1.1. Damage in the DNA and the DNA damage checkpoints 37
1.1.1. DNA repair mechanisms 39
1.1.2. DNA damage and replication 42
1.2. Natural transformation 44
1.3. RadA/Sms 47
1.4. DisA 51
1.4.1. The role of c-di-AMP 54
2. Objectives 59
3. Materials and Methods 63
3.1. Materials 63
3.1.1. Strains 63
3.1.2. Reagents 64
3.1.3. Media 65
3.1.4. Oligonucleotides 65
3.1.5. Plasmids 67
3.2. Methods 70
3.2.1. Cells manipulation and mutant construction 70
3.2.1.1. Preparation of E. coli competent cells 70
3.2.1.2. Preparation of B. subtilis competent cells 70
3.2.1.3. Transformation of E. coli cells 71
3.2.1.4. Transformation of B. subtilis cells 71
3.2.1.5. Bacterial transduction using bacteriophage SPP1 71
3.2.1.6. Construction of protein mutant variants 71
3.2.1.7. Construction of B. subtilis mutant strains 72
3.2.2. Chronic survival assays 73
3.2.3. Transformation assays 73
3.2.4. Protein-protein interaction in vivo 73
3.2.5. Cytological studies: Analysis of DisA dynamics 74
3.2.6. DNA manipulation 75
3.2.6.1. DNA isolation and quantification 75
3.2.6.2. DNA radiolabeling 75
3.2.6.3. Preparation of DNA substrates for biochemical assays 75
3.2.6.4. Purification of DNA substrates for biochemical assays 76
3.2.7. Protein manipulation 77
3.2.7.1. Protein overexpression 77
a) DisA 77
b) RadA/Sms 77
3.2.7.2. Protein purification 78
a) DisA 78
b) RadA/Sms 79
3.2.7.3. Protein identification 80
3.2.8. Biochemical assays 80
3.2.8.1. Protein-DNA interaction: Electrophoretic mobility shift assay (EMSA) 80
3.2.8.2. DNA helicase activity assays 80
3.2.8.3. ATPase assays 80
a) Spectrophotometer 80
b) Thin-layer chromatography (TLC) 81
3.2.8.4. DAC activity assays 81
3.2.8.5. Protein-protein interaction in vitro 82
3.2.8.6. Strand exchange reactions 82
4. Results 85
4.1. Genetic characterisation of RadA/Sms and DisA 85
4.1.1. RadA/Sms conserved domains play a crucial role in RadA/Sms-mediated
repair-by-recombination 85
4.1.2. DisA C290 impairs cell viability 88
4.1.3. RadA/Sms is required for genetic exchange 89
4.1.4. RadA/Sms mutant variants poison natural transformation 90
4.1.5. RadA/Sms is essential for natural chromosomal transformation in the recG
background 92
4.2. Biochemical characterisation of RadA 93
4.2.1. RadA/Sms and its mutant variants preferentially bind ssDNA and HJ DNA 93
4.2.2. RadA/Sms hydrolyses ATP in the absence of DNA 95
4.2.3. RadA/Sms C4 mutant variants hydrolyse ATP in a DNA-dependent manner 97
4.2.4. DisA fails to stimulate the ATPase activity of RadA/Sms 100
4.2.5. RadA/Sms unwinds forked DNA 101
4.2.6. RadA/Sms unwinds tailed DNA 102
4.3. Interplay between RadA/Sms and RecA activities 104
4.3.1. RadA/Sms physically interacts with RecA 104
4.3.2. RadA/Sms inhibits the ATPase activity of RecA 106
4.3.3. RadA/Sms unwinds the non-cognate 3´-tailed substrate in the presence of
RecA 108
4.3.4. RadA/Sms unwinds the non-cognate 5´-forked substrate in the presence of
RecA 110
4.3.5. RecA regulates RadA/Sms unwinding of a 3´-invading D-loop 113
4.3.6. RecA promotes RadA/Sms unwinding of a 5´-invading D-loop 114
4.3.7. RecA loads RadA/Sms to extend D-loop structures 116
4.3.8. RadA/Sms does not unwind HJ DNA 118
4.4. Interplay between RadA/Sms and DisA 119
4.4.1. HJ DNA suppresses DisA-mediated c-di-AMP synthesis 119
4.4.2. RadA/Sms inhibits DisA-mediated c-di-AMP synthesis 120
4.4.3. The HhH domain of DisA is crucial for its DAC activity 121
4.4.4. RadA/Sms mutations differentially regulate the DisA DAC activity 122
4.4.5. The interaction of DisA with DNA or RadA/Sms is mutually exclusive 124
4.5. Interplay between DisA and RecA activities 125
4.5.1. End resection is dispensable for DisA pausing after DNA damage 125
4.5.2. RecO and RecA are required for DisA pausing at sites of DNA damage 127
4.5.3. DisA binding to DNA is required for pausing at a damage site 128
4.5.4. DisA interacts with RecA 129
4.5.5. DisA interferes RecA·ATP nucleation onto ssDNA 131
4.5.6. DisA interferes RecA·ATP nucleation onto SsbA-ssDNA-RecO complexes 132
4.5.7. DisA does not interfere with RecA·dATP filament formation 134
4.5.8. RecA does not affect c-di-AMP synthesis 135
4.6. DisA, RadA/Sms and RecA work together for checkpoint inactivation 137
5. Discussion 143
5.1. The role of RadA/Sms in Bacillus subtilis 143
5.2. RadA/Sms works in concert with RecA 146
5.3. RadA/Sms regulates DisA-mediated c-di-AMP synthesis 152
5.4. DisA stalls at RecA-mediated DNA damage intermediates 154
6. Conclusions 161
6. Conclusiones 165
References 169
Annexes 183
Annex 1: Movies 183
Annex 2: Publications 185
List of figures
Figure 1. Damage in the DNA appears as a consequence of several factors 37
Figure 2. Induction of the SOS response in bacteria 38
Figure 3. Summary of the BER, MMR and NER mechanisms 39
Figure 4. Summary of SSG repair, HR and SDSA mechanisms in B. subtilis 41
Figure 5. Summary of fork reversal, template switching and translesion synthesis
mechanisms in B. subtilis 42
Figure 6. Summary of the natural transformation mechanisms in B. subtilis 45
Figure 7. Sequence alignment of RadA proteins 47
Figure 8. Crystal structure of S. pneumoniae RadA 50
Figure 9. Crystal structure of Thermotoga maritima DisA 52
Figure 10. Mechanistic model for the role of DisA 53
Figure 11. Subcellular localisation of DAC and PDE enzymes in B. subtilis 55
Figure 12. Metabolism and targets of c-di-AMP in osmotic stress 56
Figure 13. DNA structures used in this study 76
Figure 14. Survival of RadA/Sms mutant variants upon exposure to DNA damage 86
Figure 15. Survival of DisA mutant variants upon exposure to DNA damage 88
Figure 16. RadA/Sms preferentially binds ssDNA and HJ DNA in an ATP independent
manner 94
Figure 17. RadA/Sms hydrolyses ATP 96
Figure 18. RadA/Sms C13A or C13R preferentially hydrolyses ATP in the presence of
ssDNA 99
Figure 19. RadA/Sms physically interacts with itself and with DisA 100
Figure 20. RadA/Sms unwinds forked DNA in the 5´→3´ direction 102
Figure 21. RadA/Sms unwinds tailed duplex DNA in the 5´→3´ direction 103
Figure 22. RadA/Sms interacts with RecA 105
Figure 23. RadA/Sms inhibits RecA ATPase activity 107
Figure 24. RecA loads RadA/Sms to unwind tailed duplex DNA in the 5´→3´ direction 109
Figure 25. RecA loads RadA/Sms to unwind forked DNA in the 5´→3´ direction 110
Figure 26. Proposed models for a role of RecA in loading RadA/Sms on the 5´-forked
substrate 112
Figure 27. RadA/Sms unwinds a 3´-invading D-loop DNA 114
Figure 28. RadA/Sms unwinds a 5´-invading D-loop DNA in the presence of RecA 115
Figure 29. Proposed model for a role of RecA in loading RadA/Sms on the 5´-invading D-
loop substrate 117
Figure 30. RecA helps RadA/Sms to unwind a homologous 5´-invading D-loop substrate 118
Figure 31. RadA/Sms fails to unzip HJ DNA 118
Figure 32. DisA synthesises c-di-AMP and this synthesis is regulated by HJ DNA 119
Figure 33. DisA synthesis of c-di-AMP is regulated by RadA/Sms 120
Figure 34. Integrity of the DisA HhH domain is required for c-di-AMP synthesis 121
Figure 35. RadA/Sms mutant variants inhibit DisA-mediated c-di-AMP synthesis to
different extents 123
Figure 36. DisA synthesis of c-di-AMP is regulated by HJ DNA, dsDNA and RadA/Sms 125
Figure 37. End resection is not necessary for DisA pausing at the DNA lesion 126
Figure 38. DisA foci remain highly dynamic upon DNA damage in recO or recA cells 128
Figure 39. The DNA binding domain is required for DisA focus motion on DNA 129
Figure 40. DisA interacts with RecA 130
Figure 41. RecA-mediated ATPase activity in the presence of DisA 132
Figure 42. DisA interferes RecA·ATP nucleation onto SsbA-ssDNA-RecO complexes 133
Figure 43. RecA-mediated dATPase activity is not affected by DisA or c-di-AMP 134
Figure 44. DisA-mediated c-di-AMP synthesis is not affected by RecA·ATP 136
Figure 45. DisA, RadA/Sms and RecA work together for checkpoint inactivation 138
Figure 46. Proposed model for the role of RadA/Sms in coordination with RecA during
natural transformation and DSB repair 150
Figure 47. Proposed mechanisms for DisA in RecA-mediated DNA damage tolerance
pathways 156
List of tables
Table 1. Escherichia coli strains 63
Table 2. Bacillus subtilis strains 63
Table 3. Chemicals and compounds 64
Table 4. Media 65
Table 5. Oligonucleotides 66
Table 6. Plasmids 67
Table 7. Buffers 70
Table 8. Transformation efficiency is impaired in the radA mutant 90
Table 9. RadA/Sms mutant variants impair chromosomal and plasmid transformation 91
Table 10. RadA/Sms is essential for natural chromosomal transformation in the recG
background 93
Abstract
DNA is constantly subjected to alterations in all living cells. If unrepaired, these alterations can lead to mutations or even cell death, and this is the reason why cells have evolved diverse mechanisms that repair or tolerate the damage to survive in its presence. When cells are actively dividing, one of the major consequences of unrepaired damages is the arrest of the replication fork machinery. Bacillus subtilis RadA/Sms and DisA have been described to collaborate in the mechanisms that cope with this arrest, by coordinating the timing of cell proliferation with the DNA damage response. The mechanisms that act upon replication fork stalling include fork reversion, template switching and translesion synthesis; but when replication forks collapse, RecA-mediated homologous recombination is the preferred pathway. This DNA strand exchange mechanism performed by RecA in DNA repair shares common steps with the one occurring in natural chromosomal transformation, a crucial process for the acquisition of genetic diversity and for the restoration of mutated genes, that plays a central role in the evolution and the spread of pathogenicity traits and antibiotic resistance genes. Since both DNA repair and natural transformation are essential for bacterial survival and have crucial roles in human health, the role of RadA/Sms, DisA and RecA in these processes has been studied in this work.
RadA/Sms conserved C4 (or Zn finger domain), H1 (or Walker A domain) and KNRFG motifs are crucial for survival upon replication fork arrest and for natural transformation. DisA-mediated synthesis of the essential c-di-AMP messenger and DNA binding are crucial for survival upon replication fork stalling, but DisA is dispensable for natural transformation. RadA/Sms and its mutants in the conserved C4, H1 and KNRFG motifs bind preferentially ssDNA and HJ DNA. RadA/Sms shows ATPase activity that cannot be further stimulated by DNA substrates, while the interaction of RadA/Sms C4 motif mutants with DNA stimulates their ATPase activity. RadA/Sms and its mutant variants in the C4 motif unwind DNA in the 5´→3´ direction. RecA interacts with wt RadA/Sms, but not with RadA/Sms C4 mutants, and loads it on the DNA to promote unwinding of non-cognate substrates. This interaction is crucial to recruit RadA/Sms onto displacement loop (D-loop) DNA, and both proteins in concert facilitate D-loop extension and integration of ssDNA during chromosomal transformation. During double-strand break repair, RadA/Sms might also contribute to synthesis- dependent strand annealing rather than canonical double-strand break repair.
DisA physically interacts with RadA/Sms, that inhibits its diadenylate cyclase activity. This activity is also inhibited by HJ DNA or ssDNA, but the interaction of DisA with DNA and RadA/Sms is mutually exclusive. This could represent a mechanism to maintain c-di-AMP concentration during unperturbed growth but reduce its synthesis to signal the DNA damage as a checkpoint. DisA pausing at the site of DNA damage requires RecA and RecO activities. DisA physically interacts with RecA and modulates its ATPase and strand exchange activities to delay repair by recombination functions.
Once DNA damage is repaired, the interaction of RecA with RadA/Sms and DisA may restore c-di- AMP synthesis and inactivate the checkpoint mechanism, to resume replication and cell proliferation.
Resumen
El ADN está sometido constantemente a alteraciones. Si no son reparadas, pueden producir mutaciones o incluso la muerte celular, y por ello las células han desarrollado diversos mecanismos para reparar o tolerar el daño y sobrevivir en su presencia. Cuando las células están en división, una de las mayores consecuencias de los daños no reparados es la parada de la horquilla de replicación. Ha sido descrito que en Bacillus subtilis RadA/Sms y DisA colaboran en los mecanismos que hacen frente a esta parada, coordinando la proliferación celular con la respuesta al daño en el ADN. Cuando la horquilla de replicación pausa, los mecanismos de reversión de la horquilla, cambio de molde o síntesis translesiva actúan, pero cuando la horquilla de replicación colapsa, la recombinación homóloga mediada por RecA es la ruta preferida. Este mecanismo de intercambio de cadenas llevado a cabo por RecA durante la reparación del ADN comparte pasos con el que ocurre durante la transformación cromosómica natural, un proceso crucial para la adquisición de diversidad genética y para la restauración de genes mutados que juega un papel central en la evolución y expansion de patogenicidad y resistencia a antibióticos. Como tanto la reparación del ADN como la transformación natural son esenciales para la supervivencia bacteriana y juegan papeles cruciales en la salud humana, la función de RadA/Sms, DisA y RecA en estos procesos se ha estudiado en este trabajo.
Los motivos conservados C4 (o dominio dedo de Zn), H1 (o dominio Walker A) y KNRFG de RadA/Sms son cruciales para la supervivencia cuando la horquilla de replicación se detiene, y para la transformación natural. La síntesis del mensajero esencial c-di-AMP por DisA y su actividad de unión al ADN son cruciales para la superviviencia cuando la horquilla de replicación se pausa, pero DisA no participa en transformación natural. RadA/Sms y sus mutantes en los motivos conservados C4, H1 y KNRFG unen preferentemente ADN de cadena sencilla y estructuras HJ. RadA/Sms tiene actividad ATPasa, que no es estimulada por ADN, mientras que la de los mutantes en el motivo C4 sí lo es.
RadA/Sms y sus mutantes en el motivo C4 tienen actividad helicasa en dirección 5´→3´. RecA interacciona con RadA/Sms, pero no con sus mutantes en el motivo C4, y la carga en el DNA para promover la actividad helicasa sobre sustratos no propios. Esta interacción es crucial para reclutar a RadA/Sms en sustratos de ADN D-loop, y ambas proteínas en conjunto facilitan la extensión del D- loop y la integración de ADN de cadena sencilla en transformación cromosómica. Durante la reparación de roturas de cadena doble, RadA/Sms también podría contribuir al apareamiento de hebras dependiente de síntesis en mayor medida que a la reparación canónica de roturas de cadena doble.
DisA interacciona con RadA/Sms, que inhibe su actividad diadenilato ciclasa. Esta actividad también es inhibida por estructuras HJ o ADN de cadena sencilla, pero ambas interacciones son mutuamente excluyentes. Esto podría representar un mecanismo para mantener la concentración de c- di-AMP estable, pero reducirla para señalar la presencia de daño en el ADN. La parada de DisA en el sitio de daño en el ADN requiere la actividad de RecA y RecO. DisA interacciona con RecA y modula sus actividades ATPasa y de intercambio de cadenas para retrasar la reparación por recombinación homóloga. Una vez que el daño se ha reparado, la interacción de RecA con RadA/Sms y DisA podría restaurar la síntesis de c-di-AMP para inactivar el checkpoint y continuar la replicación y proliferación
Abbreviations
List of abbreviations
(p)ppGpp Guanosine
(penta)tetraphosphate
kDA Kilodalton
4NQO 4-nitroquinoline N-oxide KDapp Apparent dissociation constant
ADP Adenosine diphosphate LB Luria-Bertani
AP Apurinic/Apyrimidinic ldsDNA Linear double-stranded DNA
AS Ammonium sulfate MMC Mitomycin C
ATM Ataxia telangiectasia mutated MMR Mismatch repair
ATP Adenosine triphosphate MMS Methyl methanesulfonate
ATR ATM-Rad3-related MOI Multiplicity of infection
BACTH Bacterial adenylate cyclase two-hybrid
NAD Nicotinamide adenine dinucleotide oxidised
BER Base excision repair NADH Nicotinamide adenine
dinucleotide reduced
bp Base pair Nal Nalidixic acid
BSA Bovine serum albumin nc Nicked circular
c-di-AMP Cyclic di-adenosine monophosphate
NCO Non-crossover
CH Casein hydrolysate NER Nucleotide excision repair
CIP Calf intestinal phosphatase NHEJ Non-homologous end joining
CO Crossover Nm Neomycin
cssDNA Circular single-stranded DNA
nt Nucleotide
DAC Diadenylate cyclase NTP Nucleoside triphosphate
dADP Deoxyadenosine diphosphate OD Optical density DAPI Diamidino-2-phenylindole
dihydrochloride
PAGE Polyacrylamide gel electrophoresis
dATP Deoxyadenosine triphosphate PBS Phosphate-Buffered Saline
DDR DNA damage response PCR Polymerase chain reaction
DDT DNA damage tolerance PDE Phosphodiesterase
D-loop Displacement-loop PEI Polyethyleneimine
DSB Double-strand break PNK Polynucleotide kinase
dsDNA Double-stranded DNA pppApA ATP-AMP
DTT Dithiothreitol Rif Rifampicin
E Elution SDS Sodium dodecyl sulfate
EDTA Ethylenediaminetetraacetic acid
SDSA Synthesis-dependent strand annealing
EMSA Electrophoretic mobility shift assay
SEM Standard error of the mean
FT Flow-through Spc Spectinomycin
GFP Green fluorescent protein SSA Single-strand annealing HGT Horizontal gene transfer ssDNA Single-stranded DNA
HJ Holliday junction SSG Single strand gap
HR Homologous recombination TLC Thin-layer chromatography IPTG Isopropyl -D-1-
thiogalactopyranoside
TLS Translesion synthesis
IR Infrared UV Ultraviolet
kb Kilobase W Wash
kcat Catalytic rate constant wt Wild-type
Introduction
Introduction
1.1. Damage in the DNA and the DNA damage checkpoints
In all living cells DNA is constantly subjected to alterations. These alterations can affect the chemistry of this molecule when nucleotides suffer modifications, its sequence if erroneous bases are incorporated, or even its structure when breaks or nicks are originated. These modifications can arise as a consequence of multiple factors. Among them, errors introduced during inaccurate replication, DNA repair or recombination are the main source of alterations in the DNA. However, environmental factors such as radiation (UV, IR…) or products of cellular metabolism such as reactive oxygen species, represent an additional and important source of DNA damage. To cope with these DNA alterations, cells have evolved diverse mechanisms that allow them to repair the damage or even survive in its presence. Nevertheless, if these lesions are not repaired, they can be mutagenic or even lethal, representing the main source of genetic instability (e.g. cancer), although if tolerated, they are also necessary for genetic variability and species evolution (Figure 1) (reviewed in Hoeijmakers, 2001).
Figure 1. Damage in the DNA appears as a consequence of several factors
Each specific type of lesion will be repaired by a specific repair pathway. If unrepaired, DNA damage can lead to cell death, mutagenesis or DNA replication blockage in bacteria (in eukaryotes, cell-cycle arrest). Figure extracted from (Hoeijmakers, 2001).
If cells are actively dividing, one of the major consequences of unrepaired damages is the stall or collapse of the replication fork machinery, that may affect genome integrity. Thus, in order to survive, cells must activate a DNA damage response (DDR) system to bypass or repair the lesion, followed by the restart of DNA replication (reviewed in Hoeijmakers, 2001).
Among the mechanisms that have evolved to deal with DNA damage, checkpoints are the ones that sense the damage and signal their presence. These mechanisms recognise specific type of lesions, and coordinate cell response to survive, by slowing down replication and pausing the progression through the cell cycle until the damage has been repaired, inducing and repressing the expression of genes, and recruiting repair proteins to the damage site. Slowing down replication and cell proliferation gives time for the cell to remove the lesion. In eukaryotes, two DNA damage checkpoint pathways have been described: the ataxia telangiectasia mutated (ATM) that responds to double-strand breaks (DSBs), and the ATM-Rad3-related (ATR) pathway, which responds mainly to single-stranded DNA (ssDNA) regions coated by the single-stranded binding protein RPA, generated due to the uncoupling of the replicative DNA polymerase and the DNA helicase in stalled replication forks (reviewed in Zhou and Elledge, 2000). Likewise, in exponentially growing bacteria, when DNA damage does not affect the integrity of the nucleoid, replication forks stall, and ssDNA regions are generated, that activate the expression of a set of genes, including some representatives of the DNA repair system, via a RecA- and LexA-dependent mechanism (SOS response) (Figure 2) (reviewed in Maslowska et al., 2019).
Lesions that compromise nucleoid integrity, such as DSBs, cause replication fork collapse and can trigger, besides the SOS response, a more complex DNA damage response that is RecA-dependent, but LexA-independent (DSB response) (Cardenas et al., 2014).
Figure 2. Induction of the SOS response in bacteria
In the absence of DNA damage, LexA inhibits the expression of a set of genes involved in DNA repair. Upon DNA damage that stalls the replication forks, ssDNA regions are accumulated and coated by RecA. RecA then induces the self-cleavage of LexA, that in turn derepresses the expression of the SOS response genes. Figure extracted from (Maslowska et al., 2019).
1.1.1. DNA repair mechanisms
Several pathways to deal with diverse types of DNA lesions have appeared during evolution.
These mechanisms are conserved in both prokaryotic and eukaryotic cells, although differences due to the complexity of the organisms exist (such as the number or type of proteins involved). The Firmicutes (or Gram-positive with low dC+dG content) bacterium Bacillus subtilis represents a good model for the study of the DDR in prokaryotes, because it can develop a broad variety of DNA repair mechanisms. It is a soil bacterium that undergoes differentiation and development stages such as sporulation and natural competence under stress conditions. Unless stated otherwise, all genes and proteins mentioned in this work are of B. subtilis origin.
Figure 3. Summary of the BER, MMR and NER mechanisms
(A) The BER system recognises chemically altered bases, a glycosylase with the help of an AP endonuclease removes the nucleotide, and synthesis of DNA then occurs. (B) The MMR system recognises mismatched bases through the collaboration of MutS and MutL, that removes the erroneous base. The gap is further resected, and synthesis of DNA to fill the gap occurs. (C) The NER system recognises DNA distortions, as bulky adducts, that are cleaved by the action of the UvrABC proteins, generating a short ssDNA region. Then, synthesis of DNA to fill the gap occurs. Figure adapted from (Spampinato, 2017).
Three specific error-free DNA repair systems have been described for coping with lesions that do not comprise nucleoid integrity. Base excision repair (BER) removes specifically non-bulky lesions in the DNA molecule, which are originated by chemical agents that alkylate, oxidate, deaminate, etc.
the bases. In this pathway, a glycosylase detects and removes the damaged base, introducing a apurinic/apyrimidinic site (the AP site). This AP site is highly mutagenic and potentially can cause ssDNA breaks, and, to avoid that, a specific AP endonuclease removes the sugar introducing a gap, that is later filled by the action of a DNA polymerase and a DNA ligase (Figure 3A) (reviewed in
Sedgwick, 2004; Almeida and Sobol, 2007). Mismatch repair (MMR) specifically eliminates errors that are introduced during DNA synthesis, such as insertions, deletions or base mispairings. In this pathway, the protein MutS recognises the mismatch, and recruits MutL. Then, the MutSL complex interacts with the processivity factor of the DNA polymerase PolC (the clamp), that in turn activates the cryptic endonuclease activity of MutL. Activated MutL, as a part of the MutSL- clamp complex, introduces a nick in the strand at which the clamp is bound. This nick is then further resected, and finally filled by a DNA polymerase and a DNA ligase (Figure 3B) (reviewed in Lenhart et al., 2012).
This mechanism is highly conserved, except in Enterobacteria. Finally, the removal of bulky DNA adducts, thymine dimers, DNA crosslinks, etc., generated by UV light or drugs, is performed by the nucleotide excision repair (NER) pathway. When this type of lesions is recognised, endonucleolytic cleavage of the DNA molecule occurs, and a short ssDNA region containing the lesion is removed.
Then, the gap is filled by a DNA polymerase and sealed by a DNA ligase (Figure 3C) (reviewed in Reardon and Sancar, 2005).
The most lethal type of DNA damage is the generation of DSBs, that affect the integrity of the DNA molecule. Two-ended DSBs may be originated from environmental sources such as X rays, or by the action of chemical agents or antibiotics such as nalidixic acid (Nal). This type of DNA damage can be repaired via three mechanisms in B. subtilis: error-free homologous recombination (HR), and error-prone single-strand annealing (SSA) and non-homologous end joining (NHEJ). Since they are prone to introduce errors, the latter two pathways only play a role when error-free HR is impaired, or during stationary phase or spore germination, when there is no available template for HR.
Error-free HR uses an intact sister chromosome as the template to initiate recombination and re- establish replication (Figure 4A.2) (reviewed in Ayora et al., 2011; Lenhart et al., 2012; Alonso et al., 2013; Carrasco et al., 2017). In HR-mediated DSB repair, RecN in concert with PNPase is one of the first responders, recognizing the DSB and establishing a ‘repair centre’. Then, long-range end- resection takes place by the action of the AddAB complex (counterpart of Escherichia coli RecBCD) or RecJ in concert with a RecQ-like DNA helicase (RecQ and/or RecS (missing in E. coli)) and the single-stranded binding protein SsbA (counterpart of E. coli SSB), generating a 3´-ssDNA tailed duplex substrate, coated by SsbA. Later, with the help of accessory (recombinase mediators and modulators) SsbA, RecF, RecO, RecR, RecX and RecU proteins, the recombinase RecA is recruited onto the ssDNA region. RecA mediates strand exchange (its mechanism of action will be further described in the “Natural transformation” section of this Introduction), forming a displacement loop (D-loop) that allows the DNA polymerase to use the intact homologous chromosome as a template for DNA synthesis to restore genetic material lost by resection. The branch-migration translocases (RecG and RuvAB have been described at present) facilitate extension of the D-loop and the formation of a Holliday junction (HJ). Then, second end-capture by RecA produces a second HJ, and the branch migrating enzymes promote branch migration until a cognate RecU (counterpart of E. coli RuvC)
cleavage site is exposed. Then, RecU, in concert with RuvAB, catalyses the resolution of the HJs.
Depending on the orientation of the cleavage, cross-over (CO) or non-crossover (NCO) products can be generated. However, in the context of the replisome, RecGEco helps RuvCEco to cleave a HJ (Gupta et al., 2014). Alternatively, the double HJs can be dissolved by Topo III in concert with a RecQ-like helicase (RecQ or RecS), generating NCO products. If second end-capture does not occur, the intermediate is regressed in a process called synthesis-dependent strand annealing (SDSA).
Another type of DNA damage that also affects the integrity of the DNA molecule is the generation of single strand gaps (SSG) that arise during DNA replication when DNA synthesis is blocked by a lesion on one of the strands. SSGs are repaired by HR functions, in a process similar to the previously described (Figure 4A.1). Alternatively, the SSG can be prevented in the absence of HR via diverse DNA damage tolerance pathways (DDT) (see below).
During SSA, after end-resection and in the presence of regions of micro-homology, a SSA protein (RecA, RecO) may promote SSA, and an uncharacterised exonuclease removes the flaps, the gaps are filled and the ends sealed by unknown functions. In NHEJ, end-resection does not take place, and DNA ends are directly tethered by the action of the Ku protein. By this mechanism, broken ends are ligated with little or no base pairing, producing deletions or insertions (reviewed in Carrasco et al., 2017).
Figure 4. Summary of SSG repair, HR and SDSA mechanisms in B. subtilis (A.1) If a gap is left opposite to a site of DNA damage during replication, the SSG repair mechanism takes place. (A.2) DSB repair. In both cases, DNA damage is recognised by RecN in concert with the PNPase, and ends are resected by RecJ in concert with RecQ or RecS, or by AddAB. This resection leaves 3´ ssDNA overhangs, that are coated by SsbA.
Then, RecA with the help of its modulators and mediators loads onto the ssDNA, and catalyses strand invasion to form a D-loop, that is further extended by branch migration translocases. In SSG repair, the reversion of the template has been hypothesised to occur (A.1). During DSB repair, the second-end can be captured to form a second HJ, that will be resolved by the action of RecU or dissolved by the action of RecQ/RecS.
Alternatively, the structure is regressed by SDSA (A.2). Figure kindly provided by Dr. Ester Serrano.
1.1.2. DNA damage and replication
Here, the DDR has been studied in exponentially growing cells, that are actively replicating their DNA and dividing. Thus, if DNA damage is not repaired by the aforementioned mechanisms, it can induce the stalling or the collapse of the progression of the replication fork, that must be surpassed to continue replication and allow cell survival. This is the reason why cells have developed diverse mechanisms to tolerate DNA damage (DDT) when it is encountered during replication fork progression.
To study these mechanisms, as well as the mechanisms that promote DNA replication re- initiation, several synthetic agents that introduce diverse types of DNA damage have been used. The DNA damage that is caused by these chemical agents is recognised by different specific mechanisms, and can lead to either replication fork stalling or replication fork collapse if unrepaired. In this work, methyl methanesulfonate (MMS), mitomycin C (MMC), hydrogen peroxide (H2O2), 4-nitroquinoline N-oxide (4NQO) and Nal were selected as the DNA damaging agents to study the role of different proteins in the DDR of exponentially growing B. subtilis cells.
Figure 5. Summary of fork reversal, template switching and translesion synthesis mechanisms in B. subtilis (A) When the damage is on the template leading strand, RecG can mediate the reversal of the fork, to form a HJ intermediate in which synthesis of the nascent leading strand uses the nascent lagging strand as a template. Then, the fork is regressed by the action of RuvAB, and replication restarts. Alternatively, the HJ intermediate is cleaved by the action of RecU, and the product is processed via HR. (B) If the damage is on the template lagging strand, RecA or RadA/Sms has been proposed to mediate the switching of the template, and the nascent lagging strand then uses the nascent leading strand as a template for DNA synthesis. The situation is reversed, and replication is restarted. (C) In the translesion synthesis mechanism, the replicative polymerase is replaced by an error-prone translesion polymerase (PolY1 or PolY2), that introduces a nucleotide opposite to the damage site, and replication is restarted. In the three mechanisms, the damage may be removed during replication of after replication (post-
MMS introduces alkyl groups in the nitrogen atoms of purine residues, mostly at the position N7 of the guanine. The removal of this alkylated bases is predominantly performed by the BER pathway. However, if unrepaired, this type of lesion causes base mispairing (Beranek, 1990; Friedberg et al., 1995; Sedgwick, 2004). 4NQO is an UV-mimetic compound that induces the formation of bulky adducts, that are specifically repaired by the NER pathway (Kondo, 1981). If unrepaired, MMS- or 4NQO-induced lesions lead to replication fork stalling, with SSG or regressed forks (formation of a HJ) being generated. MMC produces interstrand and intrastrand DNA crosslinks, depending if the damage sites are on the opposite or on the same strand, respectively (Friedberg et al., 1995). DNA crosslinks block replication and transcription. The excision of these adducts is promoted by the NER mechanism, leading mainly to one-ended DSBs, and then promoting the collapse of replication forks (De Silva et al., 2000; Lee et al., 2006). H2O2 generally reacts with ferrous iron (Fenton reaction) to generate highly reactive hydroxyl radicals, that are also formed as by-products of aerobic respiration.
These oxidant radicals induce different types of DNA damage, including strand breaks and crosslinks.
Hydroxyl radicals react with the C8 of the guanine generating a C8-OH adduct (8-oxoG), which is regarded as the most abundant product of oxidative damage. Repair of this damage is promoted by diverse pathways, such as BER, NER or recombinational repair; the latter when one- or two-ended DSBs are produced, and leads to replication fork collapse (Henle and Linn, 1997). Lastly, Nal is a synthetic quinolone commonly used as an antibiotic. It inhibits a subunit of the DNA gyrase (topoisomerase II) and topoisomerase IV, preventing religation of DNA strands broken by DNA gyrase and the tertiary negative structure supercoiling of bacterial DNA. This leads to an accumulation of cleavage complexes, such as two-ended DSBs, and then to the collapse of the replication fork (Crumplin and Smith, 1976; Pommier et al., 2010).
To restore replication and maintain genome integrity, cells have evolved several mechanisms.
If the replication fork is stalled, three main mechanisms in which recombination functions play a major role have been described to cope with the damage, two of them are error-free, while one is error-prone (Figure 5). First, fork reversal has broadly been described as a major DDT and mechanism for DNA repair (Atkinson and McGlynn, 2009). It consists in the regression of the replication fork, with a coordinated annealing of the two newly synthesised strands into a “chicken-foot structure”, that resembles a HJ with one free end, when the replication machinery encounters a blockage in the leading strand that stalls its progression. This intermediate structure can be processed via different pathways:
after replication of the leading strand using the newly synthesised lagging strand as a template, the HJ can be reversed back to resume replication; or it can be resolved generating a one-ended DSB intermediate that will be processed by HR (Figure 5A). In B. subtilis, the branch-migration translocase RecG could mediate the reversal of the fork, another branch-migration translocase, RuvAB, regress the reversed fork, and RecU mediate the resolution generating a one-ended DSB (Gupta et al., 2014;
Bianco, 2015). If the lesion is in the lagging strand, the previous mechanism cannot operate, and
the lagging strand is changed to the newly synthesised strand by the action of a recombinase like RecA or a helicase, and then replication is resumed (Figure 5B) (reviewed in Marians, 2018). The last alternative is to bypass the damage and complete replication in a mechanism known as translesion synthesis (TLS). Although this mechanism is error-prone, it is still better than dying. Here, DNA damage bypass is promoted by specialised low-fidelity error-prone Y-type DNA polymerases, PolY1 and PolY2 in B. subtilis (counterparts of PolIVEco and PolVEco, respectively), that can incorporate a nucleotide opposite to the damaged site independently of its chemistry (Figure 5C) (reviewed in Marians, 2018). Failure to stabilise stalled replication forks can compromise nucleoid integrity, cause fork collapse and lead to lethal one-ended DSBs (Cortez, 2015).
When replication forks collapse (e.g. when one of the DNA strands is broken), they are converted in one-ended DSBs when encountered by the replisome, and HR plays a central role. If the damage is blocking the lagging strand progression, it can be also bypassed by re-priming lagging strand synthesis leaving a SSG that triggers the recruitment of RecA, activating the SOS response and the gap repair machinery (reviewed in Marians, 2018).
1.2. Natural transformation
Bacteria exchange genetic information by horizontal gene transfer (HGT), since they lack the characteristic sexual reproduction of eukaryotic organisms. HGT allows the acquisition of new genetic information that cannot be obtained through the aforementioned mutational processes, and thus it represents the major factor in the evolution of prokaryotes. HGT allows bacteria to adapt to changing environmental conditions by promoting the spread of genes that confer the ability to use new metabolic pathways, gain resistance to antibiotics or new ways of virulence, facilitates antigen variation to escape vaccine action, leads to non-clonal population structure, etc. (Kidane et al., 2012, Takeuchi et al., 2014). HGT takes place via three basic mechanisms: conjugation, transduction and natural transformation (Chen and Dubnau, 2004; Kidane et al., 2012; Takeuchi et al., 2014). Conjugation and transduction involve the transference of linear ssDNA and linear double-stranded DNA (dsDNA) between cells, respectively, mediated by episomes (plasmids, viruses, conjugative elements, etc.) (Chen et al., 2005, Kidane et al., 2012, Takeuchi et al., 2014). In contrast, natural competence is a bacterial programmed mechanism activated through a dedicated transcriptional programme that drives the synthesis of many proteins encoded in the genomes of host bacteria, and it involves the acquisition of long linear ssDNA (Chen and Dubnau, 2004, Chen et al., 2005; Takeuchi et al., 2014, Johnston et al., 2014).
B. subtilis develops the stage of natural competence when exposed to nutrient starvation, and 10-15% of the cells in the culture become competent (Chen et al., 2004; Claverys et al., 2009; Kidane et al., 2012). Natural transformation contributes to the acquisition of genetic diversity and to the restoration of mutated genes, playing a central role in the evolution and the spread of pathogenicity
traits and antibiotic resistance genes. Thus, understanding the mechanisms that underlie its processes is of major importance to deal with the spread of antibiotic resistance and pathogenic bacteria, that represent significant challenges for human health (Gogarten et al., 2002; Fraser et al., 2007).
During B. subtilis transient natural competence development, DNA replication is halted, a transcriptional program is activated, and the membrane-specific DNA uptake apparatus is transiently assembled at one of the cell poles (Chen et al., 2004). In B. subtilis, the DNA uptake apparatus binds any extracellular dsDNA, linearises it, degrades one strand, and internalises the other strand into the cytosol independently of its nucleotide sequence and polarity (Figure 6) (Chen et al., 2004; Claverys et al., 2009; Kidane et al., 2012).
The essential single-stranded binding protein SsbA and the competence-specific SsbB protein show >100-fold higher affinity than the RecA recombinase or the DprA and RecO mediators for ssDNA in vitro (Yadav et al., 2014; Carrasco et al., 2015). Therefore, it is proposed that the abundant SsbA and SsbB proteins coat the incoming linear ssDNA as soon as it leaves the entry channel and protect it from degradation.
Figure 6. Summary of the natural transformation mechanisms in B. subtilis The uptake apparatus at the cell pole binds dsDNA, linearises it and internalises one of the strands of the DNA inside the cell, that is coated by SsbA (A.1) If the DNA shares homology with the chromosomal DNA, RecA with the help of its mediators and modulators is loaded on the ssDNA, and catalyses strand invasion to form a D-loop, that is further extended and processed to integrate the DNA into the recipient chromosome through HR mechanisms. (A.2) If the DNA does not share homology with the chromosomal DNA, and it has internal homology and a replication origin, complementary strands are annealed or de novo synthesis occurs if pas sites are present on the ssDNA, to generate a dsDNA molecule. By the annealing of the ends, a plasmid is reconstituted and the transforming DNA is then stablished as a plasmid in the cell. Figure kindly provided by Dr. Ester Serrano.
In vitro, B. subtilis RecA is a ssDNA-dependent ATPase and dATPase. RecA in the ATP bound form (RecA•ATP), can filament onto ssDNA, but this RecA nucleoprotein filament cannot catalyse DNA strand exchange (Steffen et al., 2002; Manfredi et al., 2008). Furthermore, SsbA- (or SsbA and SsbB)-coated ssDNA blocks RecA•ATP nucleation and filament growth. This inhibition is surpassed with the help of mediators (RecO or the competence specific DprA), that promote RecA•ATP nucleation and filament growth onto ssDNA coated by SsbA (or SsbA and SsbB) and subsequent
activation to catalyse DNA strand exchange, in a process similar to that occurring in HR (Yadav et al., 2014; Carrasco et al., 2015). Then, RecA•ATP polymerises on the ssDNA, and nucleotide hydrolysis throughout the filament contributes to cycles of polymerisation (RecA•ATP)/dissociation (RecA•ADP) to ensure the formation of a nucleoprotein filament emanating from the entry channel towards the nucleoid (RecA threads) (Kidane et al., 2009; Yadav et al., 2014; Carrasco et al., 2015).
The cooperative RecA•ADP interaction among monomers stabilises the filament, and dissociation of RecA monomers occurs predominantly from the filament ends. The RecA filament, assembled on the internalised ssDNA coated by SsbA (or SsbA and SsbB), binds to multiple regions of one strand of the haploid recipient genome to efficiently search for a unique homologous sequence (Kidane et al., 2012).
When the incoming ssDNA shares no homology with the recipient chromosome (Figure 6A.2), but it has internal homology (e.g. oligomeric linear plasmid ssDNA) and can replicate autonomously, a duplex circular replicon is reconstituted in a RecA-independent manner in otherwise wild-type (wt) cells (plasmid transformation) (Canosi et al., 1978; de Vos et al., 1981). In this case, the RecA filaments formed on the incoming heterologous ssDNA are unproductive. To stop energy consuming RecA-promoted homology search between heterologous DNAs, and to permit the loading of single strand annealing protein(s) onto the SsbA- or SsbB-coated ssDNA, optimal for plasmid transformation, the RecA filaments are disassembled with the help of modulators (e.g. RecX, RecU) (Cardenas et al., 2012; Le et al., 2017; Serrano et al., 2018).
In contrast, when the incoming ssDNA shares homology with the recipient (Figure 6A.1), RecA•ATP, with the help of the two-component mediator (SsbA and DprA or RecO), and independently of its ATPase activity, catalyses DNA strand exchange (chromosomal transformation) (Yadav et al., 2014; Carrasco et al., 2015). Once the homologous region is found, the RecA nucleoprotein filament on the incoming ssDNA promotes strand invasion and produces metastable intermediates (D-loops), leading to the formation of heteroduplex DNA. It is likely that initial homologous pairing does not require a free end of the filament, and it can occur at any site along the RecA nucleoprotein filament; thus, a nascent three-stranded non-interwound paranemic D-loop might be formed. Finally, RecA•ATP, which is a slow motor (it hydrolyses 9 ATP/min), is able to branch migrate over long stretches of DNA. However, the average length of a RecA-mediated D-loop is 400- nucleotides (nt) (Radding et al., 1991; Bell and Kowalczykowski, 2016), but the average integration size of donor ssDNA during natural transformation is 13,000-nt (Dubnau and Cirigliano, 1972;
Fornili and Fox, 1977). Thus, it seems clear that another accessory factor (more likely a branch migration translocase) participates in this process to increase the length of the integrated DNA. The branch migration translocases RecG and RuvAB are among the candidates to mediate the extension of the D-loop, but their absence marginally impairs chromosomal and plasmid transformation (Kidane et al., 2012). Recently, it has been proposed that the proteins E. coli RadA and Streptococcus pneumoniae
RadA act as branch migration translocases, that might help RecAEco to catalyse DNA strand exchange, or RecASpn to extend a D-loop, respectively (Cooper and Lovett, 2016; Marie et al., 2017).
1.3. RadA/Sms
The radA gene was first identified in an E. coli mutant (radA100) that was sensitive to X-rays, UV-radiation and MMS (Diver et al., 1982). It was also described that the radA100 mutant was 30%
deficient in DSB repair (Sargentini and Smith, 1986). Later, it was shown that the deletion of the sms gene in E. coli sensitised cells to MMS (Neuwald et al., 1992). Last, it was observed that mutations in both radA and sms genes sensitised cells to the same DNA damaging agents, and that both genes had the same sequence, concluding that radA and sms were in fact the very same gene (Song and Sargentini, 1996).
In B. subtilis, the sms gene product, encoded in the clpC operon, shows 47% amino acid identity with E. coli RadA. The deletion of sms increased sensitivity of cells to MMS, and impaired chromosomal transformation, but not plasmid transformation (Krüger et al., 1997). In this work the sms or radA gene is referred as radA and its product in B. subtilis as RadA/Sms, while its product in other species will be referred as RadA labelled with the name of the species. It is important to highlight that the radA gene described here is not the same gene as radA from Archaea, even though they have the same name, and both encode proteins that share a similar ATPase domain to RecA. Archaeal RadA is a homolog of bacterial RecA and eukaryotic Rad51, that acts as a recombinase (Seitz et al., 1998).
Figure 7. Sequence alignment of RadA proteins
The amino acid sequence of different RadA proteins, from Thermus thermophilus (RadATth), Escherichia coli (RadAEco), Bacillus subtilis (RadA/SmsBsu) and Streptococcus pneumoniae (RadASpn), was aligned using the Clustal Omega software.
Acidic amino acids are coloured in blue, hydrophobic amino acids in red, basic amino acids in magenta and amino acids that are not included in the previous groups in green. The position of representative domains/motifs is indicated.
RadA is conserved from eubacteria to plants. B. subtilis RadA/Sms is a 458 amino acid-long protein with four well conserved domains/motifs (Figure 8a): a zinc-finger domain at the N-terminus with a CXXC-Xn-CXXC (C4) motif; an extended region with homology to RecA, with Walker A and B boxes (H domain), which is characteristic of ATPases; a highly conserved KNRFG motif, and a C- terminal region that is related to the Lon protease. In B. subtilis, S. pneumoniae and Listeria monocytogenes, the Lon-related domain active-site serine, present in E. coli and Mycobacterium tuberculosis proteins, is substituted by an alanine (Kruger et al., 1997). RadA protein sequence alignments of representatives of different bacterial groups (e.g. Gram-positive or negative, spore or non-spore forming) are shown in Figure 7, where the conserved domains regions are also indicated.
RadA/Sms shares 65% sequence identity with S. pneumoniae RadA, 47% identity with E. coli RadA, and 45% identity with T. thermophilus RadA.
Several reports have described RadA to be involved in the repair of diverse types of DNA damage. The moderate sensitivity to DNA-damaging agents of E. coli strains lacking radA is highly exacerbated in combination with mutations in ruvA, ruvB and/or recG, thus a role in the regression of stalled replication forks has been suggested, though radA cells remain recombination proficient in - Proteobacteria (Beam et al., 2002; Cooper et al., 2015). Genetic analysis in E. coli revealed that all the conserved domains of RadA (zinc finger, walker A, KNRXG and Lon protease-like) are required for DNA damage survival (Cooper et al., 2015). Moreover, the lack of radA impaired the recovery of genetic rearrangements induced by the replication fork helicase DnaB, and thus it was predicted to participate in recombination processes (Lovett, 2006). The disruption of Pseudomonas aeruginosa radA increases the mutation frequency (Wiegand et al., 2008). In -Proteobacteria, cells lacking RadA remain recombination proficient, as reported in Rhizobium etli, where the deletion of radA only slightly sensitises cells to UV-radiation, MMS or Nal, but when this deletion was combined with deletions in recG or ruvAB, the sensitivity was increased to different extents, suggesting again a role in branch migration (Martínez-Salazar et al., 2009). In R. etli, RadA is involved in gene conversion, perhaps via HJ processing functions together with RuvAB and RecG, although it has a poor efficiency (Castellanos and Romero, 2009). In bacteria of the Deinococccus-Thermus phylum, the absence of RadA confers a poor phenotype: resistance to ionizing radiation requires the D-loop processing activity of RadA in Deinococcus radiodurans (Slade et al., 2009; Kota et al., 2015), and Thermus termophilus RadA has been described to participate in the response to UV-radiation- and MMC-induced DNA damage (Inoue et al., 2017).
In bacteria of the Firmicutes phylum, RadA is also involved in the repair of diverse types of DNA damage. In S. pneumoniae, the deletion of radA sensitises cells to MMS, but not to UV-radiation (Burghout et al., 2007). In B. subtilis, deletion of radA renders exponentially growing cells sensitive to MMS, 4NQO, MMC and H2O2, and it also impairs survival of mature spores exposed to IR and revived under unperturbed conditions. In addition, deletion of radA does not increase the sensitivity to
these DNA damaging agents of the recA, recU, recG or ruvAB mutant, suggesting an epistatic interaction between these genes. Furthermore, RadA/Sms forms dynamic foci that move throughout the cell and become static in a background where branched intermediates are accumulated, such as
recU (Gándara and Alonso, 2015; Gándara et al., 2017; Raguse et al., 2017).
A RadA-like protein also contributes to DNA repair in eukaryotes. The deletion of Aspergillus nidulans radA gene sensitises cells to MMS and increases the frequency of spontaneous mutations (Seong et al., 1997). Oryza sativa and Arabidopsis thaliana RadA is expressed mainly in the meristems, where cell proliferation and DNA replication occur, and it is localised in the nucleus.
Interrupting the radA gene sensitises plant cells to MMC and UV-radiation (Ishibashi et al., 2005).
It has been also suggested a role for RadA in the SOS response in E. coli. Here, RadA seems to reduce or even inhibit the SOS response, perhaps by destabilizing RecA nucleoprotein filaments that activate this response (Massoni et al., 2012; Chen et al., 2015; Cooper et al., 2015). However, no difference in SOS induction has been observed in B. subtilis in the absence of RadA/Sms (Gándara et al., 2017)
In addition, RadA has been described to participate in the acquisition of new genetic traits.
RadA plays a role in chromosomal transformation in the natural competent bacterium S. pneumoniae, where radA expression is induced at late steps of transformation but does not participate in the exit from competence (Burghout et al., 2007; Weng et al., 2013; Marie et al., 2017). A role in chromosomal transformation for RadA/Sms has also been suggested in B. subtilis (Krüger et al., 1997; Carrasco et al., 2001). On the other hand, RadA plays no role in transformation in Gram-negative species such as Vibrio cholerae and Acinetobacter baylyi, where it is only involved in the DNA damage response. In these species, ComM has been suggested to play the same role in transformation that RadA does in Gram-positive bacteria (Nero et al., 2018).
Although some phenotypes vary among different bacterial species, taken together, a role for RadA/Sms in recombination, transformation and DNA repair, more specifically in the processing of branched intermediates, has been proposed. This role has recently been corroborated through biochemical assays. E. coli RadA is a ssDNA-dependent ATPase that can catalyse the branch migration step of the strand-exchange reaction mediated by RecA, reducing the time needed for final product formation (Cooper and Lovett, 2016). S. pneumoniae RadA is also a ssDNA-dependent ATPase that shows 5´→3´helicase activity. In addition, in this bacterium RecA interacts with and loads RadA/Sms at specific positions on the DNA, and this allows the protein to extend the D-loops that are formed during chromosomal transformation, increasing the length of the DNA that is integrated in the recipient chromosome (Marie et al., 2017). T. termophilus RadA is a DNA-stimulated ATPase, which shows sequence similarity to the ComM protein (Inoue et al., 2017).
Figure 8. Crystal structure of S.
pneumoniae RadA
(a) Domain organisation of pneumococcal RadA (453 residues long). (b) Structure of pneumococcal RadA. The C4 domain is present but not structured. The same colour code as in panel (a) has been used for the H and P domains, and the two linkers flanking the H domain. The RadA trimer in the asymmetric unit forms planar and annular hexamers 100Å wide and 75Å tall. The internal diameter of the ring is 25Å at the H domain and 15Å at the P domain. (c) The nucleotide binding site in the H domain contains the canonical Walker A and Walker B motifs. Figure extracted from (Marie et al., 2017).
S. pneumoniae RadA has been crystallised, and its structure analysed by X-ray diffraction (Marie et al., 2017). The crystal structure of the Lon-related domain of RadA in T. termophilus was also resolved (Inoue et al., 2017). RadA is found as hexamers in solution, with three protomers per each of the asymmetric units, that form an annular particle with double symmetry (Figure 8b). The central H domains form a ring, with an inner diameter of 25 Å, and a RecA-like fold with a nucleotide- binding site. This nucleotide binding site displays all the canonical residues found in the ATPase active site of RecA, but no arginine finger, which is required for nucleotide hydrolysis, is found close.
Although RadA was considered a RecA homologue due to the sequence of this H domain, its folding is similar to that of the helicase domain found in the hexameric replicative helicases of the DnaB family. The P domains also form a ring, with an inner diameter of 15Å, and their folding is structurally related to the proteolytic domain of Lon proteases, though without the catalytic residues. However, the inner diameter of the channel in DnaB is 25 Å, enough for placing dsDNA, while the inner diameter of 15Å of RadA is not sufficient for dsDNA and can only accommodate ssDNA. This domain has been described to be involved in DNA binding. The hexameric structure is primarily maintained by the interaction between adjacent P domains. The electrostatic potentials of the inner side of the rings have positive charges (Bailey et al., 2007; Inoue et al, 2017; Marie et al., 2017).
From these biochemical assays and descriptions of protein structure, it seems clear that RadA is a helicase that may perform branch migration over branched intermediates. DNA metabolic processes require the action of DNA helicases for a variety of functions, such as recombinational repair, replication or regression of stalled replication forks. Helicases are molecular motors that convert the chemical energy of nucleoside triphosphate (NTP) hydrolysis (typically ATP) into mechanical force to translocate along DNA in a directionally specific manner, and thus remodel the DNA structure. Six superfamilies have been proposed to classify these enzymes on the basis of their conserved domains or motifs (Singleton et al., 2007).
DnaB helicases belong to the superfamily 4 of helicases. Superfamily 4 includes hexameric proteins with a RecA-like core. The NTP binding site is placed in the interface between subunits, and the arginine finger is present in a neighbour subunit, thus far from the NTP binding site. It has been proposed that a conformational change is needed to activate their helicase activity, and that the DNA interacts with the positively charged amino acids in the central channel (Singleton et al., 2007).
Although the role of RadA seems to be clear in S. pneumoniae and E. coli, the genetic assays analysing the role of RadA in diverse bacteria involve this protein in different, although related, processes. Thus, understanding the role of RadA/Sms in B. subtilis will help to clarify how this protein works in DNA repair and transformation.
1.4. DisA
In B. subtilis, the radA and disA genes are localised in the same operon, the clpC operon. The structure of this operon is conserved among several bacterial species that encode both RadA and DisA, such as L. monocytogenes or Mycobacterium smegmatis (Kruger et al., 1997; Zhang et and He, 2013).
The disA gene encodes for the DisA protein, that was first found in a screening for proteins that interact with RacA. RacA is the protein that anchors the chromosome to the cell pole at the onset of sporulation, however axial filament formation was not impaired in ΔdisA cells, suggesting that DisA should have another function different to that of RacA (Ben-Yehuda et al., 2002). Later, it was observed that in the absence of DisA, cells are less efficient in producing mature spores, but enter in sporulation in the presence of Nal- or MMC-induced DNA damage. Since a mechanism to ensure chromosome integrity before asymmetric division and to prevent entry in sporulation in the presence of DNA damage exists (Ireton and Grossman, 1994), it was suggested a role for DisA in this checkpoint mechanism.
Furthermore, it was demonstrated that DisA formed a highly dynamic focus that colocalised with the nucleoid at the onset of sporulation, and paused in the presence of DNA damage. This loss of movement was predicted to induce a signal cascade that culminates with a delay in Spo0A activation and therefore a temporary block in sporulation. Thus, it was postulated that DisA dynamically scans the chromosome at the onset of sporulation, and pauses its movement in the presence of DNA damage
to give the cell time to repair the damage before entry in sporulation. This is the reason why the protein was called DNA integrity scanning A, DisA (Bejerano-Sagie et al., 2006).
When the DNA damage is not repaired upon entry into sporulation, DisA could also act as a damage checkpoint before spores return to vegetative growth (outgrowth). In fact, it was observed that DisA plays a role in arresting DNA replication during B. subtilis spore outgrowth until the chromosome is free of damage (Campos et al., 2014).
Figure 9. Crystal structure of Thermotoga maritima DisA
(A) DisA forms an octamer with central DAC domains (coloured in green) and peripheral quartets of HhH domains (coloured in orange). Both functional regions are linked by helical spine domain tetramers (coloured in blue). Four c-di-AMP moieties are bound between each of the four pairs of opposing DAC domains (coloured in magenta). (B) Section through the helical spine domain tetramer. For distinction, protomers are alternated in blue and grey. (C) Detailed view of the c-di-AMP-binding site at the DAC domain dimer interface (for distinction, one domain is shown in green, the other in grey). (D) Sequence alignment of DisA proteins from T. maritima (TmaDisA), Bacillus subtilis (BsuDisA), and Mycobacterium tuberculosis (MtuDisA) along with a structurally identified homolog from Bacillus cereus (hypothetical membrane-spanning protein COG1624; PDB code 2fb5).
Conserved residues are shaded, and the secondary structure of TmaDisA is shown on top of the alignment.
Notable functional motifs are annotated. Residues that bind the c-di-AMP or 30-deoxyATP ligands are indicated
Later, it was shown that DisA forms a large molecular complex in solution, whose molecular weight of ~370 kDa corresponds to a large assembly that was predicted to be an octamer (Witte et al., 2008). The crystal structure of Thermotoga maritima DisA, which shares 41% amino acid sequence identity with B. subtilis DisA (Figure 9), exhibited a dumbbell-shaped molecule (Witte et al., 2008).
In each monomer, three structural domains were detected: an amino-terminal globular domain, renamed diadenylate cyclase (DAC), which binds ATP and metal ions to synthetise c-di-AMP from the hydrolysis of 2 ATP molecules in the presence of divalent cations; a carboxy-terminal helical HhH domain that provides nonspecific DNA-binding activity, and a spine-liker domain that connects the DAC and HhH domains. To form the DisA octamer, two DisA tetramers interact by their DAC domains. Interestingly, the four HhH domains are arranged in both sides of the DisA octamer in a geometry that resembles the structure of the four HhH domains found in the HJ binding protein RuvA (Witte et al., 2008). In M. tuberculosis, both DAC and HhH domains were shown to be essential for c- di-AMP synthesis (Bai et al., 2012).
Branched DNAs (three- and four-way junctions) were found to specifically inhibit DisA- mediated synthesis of c-di-AMP. It was shown that DisA weakly binds ssDNA and dsDNA (Bejerano- Sagie et al., 2006; Witte et al., 2008), but it binds with apparently higher affinity HJs (Witte et al., 2008). A model was developed to explain the mechanism of DisA checkpoint control (Figure 10). A conformational change in DisA could be the basis for the allosteric regulation of the DAC domain by branched DNA binding. Higher c-di-AMP levels seem to be associated to non-damaged DNA, while lower c-di-AMP levels with damaged DNA (Witte et al., 2008). Thus, c-di-AMP emerged as a new nucleotide second messenger that revealed the integrity of the DNA at the onset of sporulation. In fact, it was observed that when c-di-AMP was ectopically added at the onset of sporulation to cells treated with Nal or MMC, they enter in sporulation even in the presence of DNA damage (Oppenheimer- Shaanan et al., 2011).
Figure 10. Mechanistic model for the role of DisA In the absence of DNA damage, DisA synthesises c-di- AMP. Recognition of branched nucleic acids, for instance, stalled replication forks or recombination intermediates, produce a fold change and might inhibit c- di-AMP synthesis, signaling the presence of damaged chromosomes. Figure extracted from (Witte et al., 2008).
The first association of RadA with DisA was suggested in M. smegmatis (Zhang and He, 2013).
In this bacterium, RadA physically interacts with DisA, modulating negatively the synthesis of c-di-
