DNA binding by the Rev1 BRCT region : implications for biological and structural function
Groote, F.H. de
Citation
Groote, F. H. de. (2011, February 9). DNA binding by the Rev1 BRCT region : implications for biological and structural function. Retrieved from https://hdl.handle.net/1887/16451
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9
Chapter I
General Introduction
General Introduction
10
General introduction to eukaryotic DNA replication and DNA damage response.
In order to divide, every eukaryotic cell has to progress through the cell cycle consisting of different, strictly controlled stages or phases. In the G1 phase new proteins are synthesized and new organelles are formed in order to prepare the cell for division and duplication of its DNA. During the synthesis (S)-phase, the DNA is replicated with great precision. In order to do so, multiple specialized proteins are involved. Once this phase is successfully completed, the cell progresses into a second growth phase (G2), where the cell prepares for the final phase, the mitotic (M)-phase. In the M-phase, the chromosomes are separated into two identical sets and are delivered to the daughter cells.
Of all phases in cell division, the S-phase is the most critical. Incorrect duplication of DNA leads to mutations and will eventually result in cell-death or cancer. The discovery of the duplex structure of DNA by Watson and Crick gave insight into how new DNA is synthesized [Watson and Crick, 1953]. DNA is ‘replicated’ by using each strand of the duplex as a template to generate a new, complimentary strand. DNA replication is initiated by the recognition of specific DNA sequences by the six- protein Origin Recognition Complex (ORC) [Bell and Stillman, 1992; Diffley and Cocker, 1992] (Figure 1.1A step I). To prevent uncontrolled DNA replication these replication origins are ‘licensed’ by binding of several cell cycle-checkpoint proteins and the Mini-Chromosome Maintenance proteins (MCM). To initiate DNA replication, a fraction of the licensed origins are ‘fired’, while some remain dormant (Figure 1.1A, step II). The MCM proteins unwind the DNA, and create two separate branches of single stranded DNA. The fork-shaped structure that is formed is usually referred to as the replication fork [Donaldson and Blow, 1999; Ishimi, 1997] (Figure 1.1A, step III). At the replication fork, the unwound strands serve as templates for DNA synthesis. Due to the chemical asymmetry of the DNA backbone, DNA can only
Chapter I
11
(Figure 1.1) DNA replication on undamaged and damaged DNA. (A) Binding of the Origin Recognition Protein (ORC) licenses the replication origin (step I), followed by helicase activity to unwind the DNA. Only a fraction of origins will actually be utilized while most remain dormant (step II). Because of the asymmetry of DNA, replication is continuous on the leading strand and discontinuous on the lagging strand (Step III). (B) Discontinuous DNA replication on the lagging strand in more detail. While the synthesis of DNA on the leading strand procedes in synchrony with the DNA helicase, the single stranded DNA on the lagging strand is coated with RPA until a new Okazaki fragment is formed (step I). Loading of PCNA by RFC mediates the switch from polα to polδ. Polδ extends from the Okazaki fragment to the 5’
primer terminus of the previous Okazaki fragment and displaces RPA (step II). “Idling” by polδ: once the PCNA-Polδ complex encounters the previous Okazaki fragment it cycles between a state where 2-3 nucleotides of the RNA primer are displaced which subsequently are degraded by Fen1, and a state of a ligatable nick, whereby the 3’→5’ exonuclease domain removes the newly incorporated nucleotides (step III). Once the RNA is removed, DNA ligase is able to form an ester bond between the 5’ and the 3’ end of the nick (step IV). (C) DNA lesions and fork stalling (step I). If a lesion is positioned on the leading strand, the stalling of polε will sequester DNA helicase and stall the replication fork, while a lesion on the lagging strand does not stall the replication fork. As a result the leading strand is uncoupled and replication is continued downstream, leaving a single stranded gap (step II). Most DNA is replicated before the DNA damage is repaired. In the case of severe DNA damage, dormant origins might be initialized to complete replication (step III).
General Introduction
12
be replicated in the 5->3’ direction. Since the DNA strands pair in an anti-parallel orientation, only one of the daughter strands is replicated continuously, parallel to the direction of the replication fork (leading strand). The other daughter strand is replicated in a direction opposite to replication fork movement (lagging strand), as discontinuous small fragments, called Okazaki fragments, which are subsequently joined (Figure 1.1A, step III) [Okazaki et al. 1968].
The actual synthesis of DNA is initiated on the template DNA by the polymerase alpha (polα)-primase complex. The primase subunit of polα synthesizes a short RNA primer from which the polymerase subunit of polα is able to synthesize DNA. DNA synthesis by polα is tightly regulated by Replication Factor C (RFC). Once polα has progressed approximately 20 base-pairs, it is displaced by RFC which binds at the 3’end and initiates the ‘polymerase switch’. This switch is initiated by the binding of the toroidally shaped, homotrimeric protein called the proliferating cell nuclear antigen (PCNA) to RFC. Binding to RFC opens PCNA where upon the DNA strands pass into the central cavity. PCNA then releases RFC, snaps shut and is said to be loaded onto the DNA. The topological entrapment of PCNA on closed DNA, without site specific binding, results in a binding mechanism which has been likened to a sliding clamp. The high affinity of PCNA for the replicative polymerases delta (polδ) and epsilon (polε) causes their recruitment to the replication fork. Loading of PCNA by RFC occurs in an ATP-dependent manner. Upon the loading of PCNA, ATP is hydrolyzed by RFC and causes RFC to dissociate from the DNA [Krishna et al. 1994;
Majka et al. 2004]. Recently it has been confirmed that the eukaryotic replicative system utilizes polδ solely for lagging strand synthesis and polε for leading strand synthesis [Nick McElhinny et al. 2008; Chilkova et al. 2007].
While DNA synthesis on both the leading and the lagging strand is initiated in a similar manner, it progresses differently. PCNA/Polε follow closely behind the MCM- helicase on the leading strand, while DNA synthesis on the lagging strand requires
Chapter I
13
re-initialization of Okazaki fragments every 200 nucleotides. This requires accurate coordination between the two polymerases and the regulation of the replication speed. Meanwhile the single-stranded (ssDNA) part of the lagging strand is protected by binding of replication protein A (RPA), which has a high affinity for single stranded DNA (Figure 1.1B, step I) [Glover and McHenry 2001; Conti et al.
2007]. Once a new Okazaki fragment is initialized, polδ progresses towards the 5’
end of the previous Okazaki fragment (Figure 1.1B, stepII). Completion (maturation) of the Okazaki fragment by joining the fragment to the growing daughter strand requires the coordinated activity of both polδ and Flap Endonuclease 1 (Fen1). Polδ displaces 2-3 nucleotides of the RNA primer by the synthesis of DNA followed by the cleavage of the displaced strand by Fen1. To prevent extensive strand displacement, polδ uses a control mechanism called idling. Newly synthesized nucleotides are removed by the 3’->5’ exonuclease activity of polδ, allowing the DNA substrate to be successively cycled between a state of a ligatable nick, and the state of a displaced strand, which can be degraded by Fen1 (Figure 1.1B, step III). This process is tightly regulated by DNA ligase, which cannot process an RNA-DNA junction. Successful nick closure by DNA ligase occurs as early as a few nucleotides past the RNA-DNA junction (Figure 1.1B, step IV) [Garg et al. 2004; Cho et al. 2009; Rossi et al. 2008;
Maga et al. 2001].
One of the most severe challenges a cell faces in order to carry out its function or to divide is the accumulation of DNA damage, which can be induced by either endogenous or environmental factors. The vast majority of DNA damage consists of chemical modification of the bases, which affects the overall structure of DNA, and is caused by damaging agents such as reactive oxygen species, aromatic compounds, UV radiation or radiation at higher frequencies.
General Introduction
14
There are six main types of damage lesions that can occur: 1) Hydrolysis of bases, which leads to apyrimidinic/apurinic (AP) or abasic sites [Lindahl and Andersson, 1972]. 2) Alkylation of bases by a wide variety of electrophilic chemicals [Horstfall et al. 1990]. 3) Oxidation of bases by, for instance, free radicals, of which 8-Oxo- 2’deoxyguanosine (8-oxo-dG) is the most extensively studied lesion [Cooke et al.
2003]. 4) Bulky polycyclic aromatic compounds such as benzo[a]pyrenes, which usually require oxidation by cytochrome p450 to become highly carcinogenic [Wood et al. 1999]. 5) Intrastrand crosslinks such as cyclobutane pyrimidine dimers (CPDs) that are formed under stimulation of UV radiation [Patterson and Davies, 2006]. 6) Interstrand crosslinks which are formed by chemicals such as the frequently prescribed anti-cancer drug cisplatinum [Kartalou and Essigmann, 2001].
Most DNA damage is repaired before the cell cycle progresses into S-phase by the major pathways that include: Nucleotide Incision Repair (NIR), Nucleotide Excision Repair (NER) and Base Excision Repair (BER) (reviews: [Nouspikel, 2009; Dalhus et al.
2009]. However, it is still possible that lesions persist as the cell begins DNA replication. Since the basis of the high fidelity of polδ and polε lies in the processing of exact Watson:Crick base pairs, they cannot replicate past these lesions. This creates a situation that is different for the leading and lagging strand (Figure 1.1C-I).
In the situation of the leading strand, strands can be uncoupled [Cob et al. 2003], leaving a single stranded gap [Lopes et al. 2006] (Figure 1.1C-II left). In the situation of the lagging strand the lesion has a smaller effect on the rate of progress of the replication fork. It has been shown that UV-induced lesions do not inhibit replication fork progression but only leave a small gap in the lagging strand [Svoboda et al.
1995; Pagès et al. 2008]. While polδ stalls at the lesion, the helicase and leading strand polymerase continue and a new Okazaki fragment is initiated downstream of the lesion (Figure 1.1C-II right). Alternatively, dormant origins that are positioned in between stalled forks can be activated [Blow and Ge, 2009]. As a result of either of
Chapter I
15
these mechanisms, most of the chromosomal DNA is replicated during S-phase.
However, under specific conditions single stranded gaps are left behind which are repaired at the end of S phase or in early G2 phase [Daigaku et al. 2010, Karras and Jentsch, 2010]. (Figure 1.1C-III).
DNA Translesion Synthesis
The impairment of the replication fork by the accumulation of DNA damage is a serious threat to dividing cells. Any replication fork that is stalled and left unattended could lead to double stranded DNA breaks or other genomic instability.
To circumvent the lesions that cause the stalling of the replication fork, cells have developed a specialized mechanism of DNA synthesis that is able to synthesize past the site of damage (referred to as bypass repair or synthesis). The first evidence of a bypass mechanism was discovered in the early 1960s and was later defined as the SOS repair pathway [Radman 1975]. The later discovery of specialized polymerases that can replicate past lesions resulted in a renaming of this mechanism to DNA Translesion Synthesis (TLS) [Caillet-Fauquet et al. 1977; Villani et al. 1978, Lawrence and Christenen, 1982, Strauss 1985].
After their discovery in E. Coli, these specialized polymerases where found to be conserved throughout the prokaryotic and eukaryotic kingdoms [Lawrence and Christensen, 1982]. From prokaryotic studies the initial idea arose that TLS occurs in two steps. First, replication is stopped at the point of the DNA damage, leaving a gap of single stranded DNA. Second, specialized polymerases are able to bypass the site of the lesion by incorporating nucleotides opposite the damage. These polymerases cannot ‘read’ a damaged base properly and therefore have a high chance of introducing mutations [Bridges and Woodgate, 1985]. While it has long been thought that most TLS events occur during S-phase, while the DNA is being replicated, recent data have shown that in yeast, TLS can be uncoupled from normal
General Introduction
16
replication and can take place at the end of S-phase, as was initially proposed [Daigaku et al. 2010, Karras and Jentsch, 2010].
In eukaryotes, the mechanism of TLS can be divided into five different steps:
1) The encountering of a lesion by the replicative polymerase and the steric exclusion of the lesion from the polymerase active site which causes the fork to stall.
Despite any possible pairing of the damaged template base with the incoming nucleotide, it is the geometric shape of the base-pair that determines whether the dNTP can be successfully incorporated. Any deviation from Watson:Crick base
(Figure 1.2) Mechanism of TLS. (A) The high fidelity DNA polymerase stalls when it encounters a lesion. (B) Ubiquitination of PCNA by Rad6/Rad18 increases the affinity of TLS polymerases for PCNA and recruits these to the lesion. (C) The TLS polymerase bypasses the lesion by inserting a nucleotide opposite the damage. Possibly, the PCNA ring works like a tool belt by rotating each polymerase in place to try to insert a nucleotide. Alternatively, polymerases are recruited sequentially to the lesion to attempt the bypass of the lesion. (D) Once the lesion is bypassed, the TLS polymerases or another polymerase will extend from the damage. (E) To avoid mutations induced by the low fidelity TLS polymerase, USP1 removes the ubiquitin from PCNA to switch to the normal replication system.
Chapter I
17
pair geometry decreases the ability of the polymerase to incorporate a nucleotide (Figure 1.2A) [Dzantiev et al. 2001].
2) The first polymerase switch, which is initiated by the ubiquitination of all three subunits of PCNA by Rad6/Rad18 [Kannouche et al. 2004]. Ubiquitination of PCNA decreases the affinity of PCNA for polδ or polε and causes the polymerase to release the DNA [Haracska et al. 2006]. Simultaneously, the affinity of PCNA for a low fidelity TLS polymerase is increased, resulting in the recruitment of the TLS polymerases to the site of damage [Hoege et al, 2002; Ulrich and Jentsch 2000; Watson et al. 2008].
To safeguard the replication of DNA by the error-prone TLS polymerases, specialized deubiquitinating enzymes negatively regulate the monoubiquitination of PCNA.
When the DNA damage is induced, these enzymes are degraded. The Ubiquitin Specific Protease 1 USP1, for instance, undergoes autocleavage which is stimulated by UV-radiation. (Figure 1.2B) [D’Andrea and Pellman, 1998; Huang et al. 2006].
3) The actual bypass of the lesion by TLS polymerases. PCNA acts as a platform for the TLS polymerases, which only shortly remain at the lesion to attempt to bypass the lesion by inserting a nucleotide opposite the damage. A possible, attractive model describes the binding of the TLS polymerases to all three subunits of PCNA, which can freely rotate in order to provide quick access to the 3’ primer terminus.
For this reason PCNA is sometimes referred to as a ‘tool-belt’, facilitating the quick cycling between a subset of tools in order to circumvent the damage (Figure 1.2C) [Solovjeva et al. 2005; Pagès and Fuchs 2002]. The elucidated structure of the PCNA- DNA-Lig1 complex contradicts this model, since binding of ligase 1 to PCNA occupies more than one open position on the PCNA ring [Mayanagi et al. 2009]. However, this might be different in the situation of TLS polymerases bound to PCNA. An alternative model suggests the sequential recruitment of TLS polymerases. In this model one polymerase is recruited to the lesion, and is substituted by another TLS polymerase if it cannot bypass the lesion [Ohmori et al. 2009].
General Introduction
18
4) Extension from the DNA lesion, which might require different polymerases depending on the nature of the DNA lesion. For instance, the complex chemistry of bulky adducts or pyrimidine dimers requires multiple different polymerase steps, before the final polymerase can synthesize DNA from the 3’end opposite of the DNA lesion (Figure 1.2D) [Shachar et al. 2008, Ziv et al. 2009].
5) To avoid spontaneously induced mutations by the low fidelity polymerase, a second polymerase switch occurs back to the high fidelity polymerases, polδ or polε.
The nature of what induces this polymerase switch remains poorly understood.
Possibly, the reappearance of deubiquitinating enzymes results in the deubiquitination of PCNA and increases the affinity of PCNA for polδ or polε (Figure 1.2E) [Huang et al. 2006].
The eukaryotic sliding clamps.
The sliding clamp PCNA performs a critical, central role in facilitating and regulating correct DNA replication. A number of protein complexes have been found in eukaryotes that bear a structural relationship to PCNA. These alternative sliding clamps include a complex of the proteins Hus1, Rad1 and Rad9, which is often referred to as the 9-1-1 complex. Similarly to PCNA, 9-1-1 is also loaded onto DNA by a multi-protein complex. This complex shares 4 subunits with RFC but also contains a unique, 9-1-1 specific subunit. In contrast to PCNA, however, the recruitment of the 9-1-1-complex to DNA is induced by DNA damage. This section will briefly discuss both sliding clamps and their role in DNA damage tolerance.
PCNA
In E. Coli the β subunit of DNA polymerase III was found to completely encircle the DNA and bind it in a sequence independent manner. The observation that it was able to slide across the length of DNA led to the name ‘sliding clamp’ [O’Donnel et
Chapter I
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al. 1996; Kong et al. 1992]. Its eukaryotic homologue, Proliferating Cell Nuclear Antigen (PCNA) is a homotrimeric complex that shares many structural features with the β subunit of polymerase II [Krishna et al. 1994; Freudenthal et al. 2009] (Figure 1.3A), and acts as a scaffold for all polymerases involved in DNA replication and repair. Binding of the DNA polymerase to PCNA tethers the polymerase to the site of synthesis thereby stimulating the incorporation of nucleotides and the processivity of the polymerase [Johnson et al. 1999; Maga et al. 2001]. Furthermore, PCNA also mediates an interaction with the sister chromatid, securing correct separation and distribution amongst the daughter cells [Moldovan et al. 2006].
PCNA is post-translationally modified as a result of induced DNA damage. The nature of the modification governs which of the different modes of the DNA tolerance
(Figure 1.3) Structure of the Eukaryotic sliding clamp PCNA and the DNA damage checkpoint clamp 9-1-1. The toroidal, trimeric structure is clearly visible in the ribbon diagram of both complexes. (A) PCNA can be modified at different positions by, respectively ubiquitination at K164 and sumoylation at K164 and K127. The region that stimulates the TLS polymerases is indicated. While only one modification is indicated, all three subunits can be simultaneously modified (adapted from [Majka and Burgers, 2004]. (B) Hydrophobic grooves on all three subunits of 9-1-1 can interact with Fen1 [Sohn et al. 2009]. Rad9 can also be phosphorylated to stimulate DNA binding [Sohn et al. 2009], while Rad1 can be ubiquitinated by an unknown mechanism [Dore et al. 2009].
General Introduction
20
pathway are utilized [Stelter and Ulrich, 2003]. Covalent linkage of ubiquitin to Lysine 164 by Rad6/Rad18 (Ubiquitination) leads to the recruitment of the TLS polymerases to the replication fork. As with polδ and polε, PCNA stimulates nucleotide incorporation by the TLS polymerases. However, this does not require the interaction with ubiquitin but rather depends on a PCNA binding motif which is conserved in all polymerases [Wood et al. 2007; Parker et al. 2007; Freudenthal et al. 2010].
In addition to PCNA monoubiquitination, polyubiquitination may occur via the ligation of one ubiquitin monomer to another through Lysine 63 by the MMS2- UBC13 pathway. Polyubiquitination causes a shift away from TLS to an error-free DNA tolerance pathway, DNA damage avoidance, where the sister strand is used as a template to bypass the lesion [Ulrich and Jentsch 2000; Haracska et al. 2004; Chiu et al. 2006; Bi et al. 2006; Papouli et al. 2006]. PCNA can also be ubiquitinated independently of Rad18 suggesting there are other regulatory proteins that perform this role as well [Simpson et al. 2006].
Alternatively, the small ubiquitin-related modifier (SUMO) can be covalently linked to Lysine 164 or Lysine 127 of PCNA by the SUMO ligase SIZ1 (sumoylation) [Hoege et al. 2002]. While ubiquitination is directly connected to different DNA damage tolerance pathways and mediates the different protein-protein interactions involving PCNA, sumoylation affects DNA damage tolerance differently by inhibition of PCNA dependent lesion bypass pathways [Hoege et al. 2002]. Sumoylation recruits Srs2 to the replication fork, a helicase that prevents unwanted recombination events after replication fork stalling, and inhibits the binding of TLS polymerases to PCNA [Haracska et al. 2004; Pfander et al. 2005].
Chapter I
21 The 9-1-1 complex
Genetic studies revealed that the protein products of the genes Rad1, Rad9 and Hus1 are important mediators of cell cycle checkpoint signaling in both yeast and humans [Volkmer and Karnitz, 1999; Bermudez et al. 2003]. Biochemical studies later revealed that these proteins behaved similarly to the eukaryotic sliding clamp PCNA [Bermudez et al. 2003]. Like PCNA, these proteins form a ring-shaped trimeric complex that encircles double stranded DNA (Figure 1.3B). However, there are a number of important differences between PCNA and the 9-1-1 complex. For example, the Rad9 subunit must be phosphorylated in order to form a stable complex with DNA [Sohn et al. 2009; Delacroix et al. 2007]. Moreover, unlike PCNA, which is loaded at the 3’ terminus, 9-1-1 is loaded at the 5’ terminus by an RFC-like complex [Majka et al. 2006]. Once loaded, the 9-1-1 complex influences/regulates a number of important protein-protein interactions at the 5’ terminus of a primer [Xu et al. 2009]. For instance, all three subunits of the 9-1-1 complex can possibly interact with Fen1, which is required for the repair of post replicative gaps [Sohn et al. 2009]. The interaction between 9-1-1 and a protein kinase called Mec1 links the replication/repair of DNA with cell cycle progression [Majka et al. 2006]. Like PCNA, the 9-1-1 complex can be ubiquitinated (on the Rad1 subunit), creating a binding platform for DNA polymerases that contain a ubiquitin binding motif (UBM), such as Rev1 [Dore et al. 2009].
The TLS polymerases
In eukaryotes DNA translesion synthesis is carried out by a combination of the specialized Y-family of polymerases and polymerase zeta (polζ). The Y family inbudding yeast consists of Rev1 and Rad30 (also known as polymerase eta (polη)) while in higher eukaryotes it contains the additional polymerase kappa (polκ) and polymerase iota (polι) members (reviewed by [Guo et al. 2009]). Rev1, the largest
General Introduction
22
(Figure 1.4). Domain composition of the four Eukaryotic members of the Y-family of polymerases. Rev1 is the only member that has a BRCT domain. The C-terminal part of Rev1 interacts with the other three family members which each have a Rev1 interacting domain.
Rev1 and polι contain a ubiquitin binding motif, while polη and polκ have a ubiquitin binding zinc finger (adapted from [Guo et al. 2009]).
member of the Y-family [Larimer et al. 1989], contains an N-terminal BRCT domain, a polymerase domain, a ubiquitin binding motif (UBM) and a C-terminal polymerase interacting domain (PID) (Figure 1.4A). The other three family members lack the BRCT region of Rev1 but share a similar composition consisting of: an N-terminal polymerasedomain, in the case of polκ two UBMs and in the case of polι and polη two ubiquitin binding zinc fingers (UBZs), and a Rev1 interacting domain (RID) (Figure 1.4B-D) [Ohashi et al. 2009]. The UBZs of polη and polι perform a similar role to the UBMs of polι and Rev1 in endowing binding to ubiquitinated PCNA. The RID domains interact with the PID of Rev1. This C-terminal PID consists of roughly 100 amino acids and binds the RID of other Y-family members in addition to polζ [Guo et al. 2003]. In yeast however, the PID has been shown to only bind polζ [D’Souza et al.
2008].
The DNA synthetic activity of all known polymerases requires three distinctive domains: the thumb, fingers and palm domains (Figure 1.5A) [Sawaya et al. 1994].
Chapter I
23
(Figure 1.5) Structures of the replicative and TLS DNA polymerases. (A) Polδ. the various domains are color coded as follows: fingers (green), palm (cyan), thumb (purple), and the exonuclease domain (brown). The Y-family polymerases: Rev1 (B), Polκ (C), Polη (D) and Polι (E). All four polymerases have an extra PAD domain (shown in green). Rev1 and Polκ possess an extra N-terminal extension, respectively the N-Digit and the N-clasp (shown in blue). (F) Bacterial DNA polymerase II, which is an ortholog of Rev3, the catalytic subunit of polζ,.
General Introduction
24
The palm positions the template DNA appropriately in the active site. The fingers domain directs the incoming nucleotide to the active site and orientates the pyrophosphate group towards the 3’OH of the primer terminus. The palm contains the active site including the bound metal ions, typically Mg2+, that catalyze the condensation reaction. Proper Watson-Crick base pairing creates a short distance between the template base and the incoming nucleotide to facilitate incorporation [Swan et al. 2009, Steitz, 1999]. Mismatched and damaged base pairs alter this distance resulting in a dramatic reduction in the rate of catalysis (Figure 1.5A) [Rothwell and Waksman, 2005].
The Y-family of polymerases all share an extra motif that is lacking in all other polymerases, the Polymerase Associated Domain (PAD) (Figure 1.5). Furthermore, Y family polymerases lack an exonuclease domain and the ability to proofread. Despite their structural similarity, each Y family polymerase performs a unique role in TLS.
Rev1 is the only polymerase that specifically inserts a cytidine opposite a damaged base and is therefore often referred to as a dCMP transferase [Nelson et al. 1996].
Structurally, Rev1 binds DNA differently compared to the other polymerases (Figure 1.5B) [Nair et al. 2005]. Extra amino acid sequences form a so-called N-digit that connects the PAD and palm domains thereby completely encircling the DNA.
Compared to the other TLS polymerases, the PAD of Rev1 is significantly larger which facilitates bypass synthesis opposite bulky guanine adducts. Interestingly there is a difference between the Rev1 PAD domain in humans and yeast [Swan et al. 2009].
While in yeast the catalytic core is exposed to the solvent, the catalytic core of human Rev1 forms a completely closed hydrophobic pocket that stabilizes the interaction with the damaged nucleotide. Polκ is also capable of bypassing bulky guanine lesions [Choi et al. 2006]. As with Rev1, polκ contains an extension at the N- terminus, the N-Clasp, which encircles the DNA. However, in contrast to Rev1, the N- Clasp is dispensable for polymerase activity [Uljon et al. 2004]. Polη is primarily
Chapter I
25
required for the bypass of cyclobutane pyrimidine dimers (CPDs) generated by exposure to UV light in both yeast and humans. The gene was first characterized as Rad30 in S. cerevisiae by its homology to DINB and REV1 [McDonald et al. 1997;
1999, Johnson et al. 1999]. Patients having a mutation in the gene coding for polη develop the skin disease xeroderma pigmentosum [Masutani et al. 1999]. Polη also efficiently bypasses abasic sites [Zhao et al. 2004]. The structure of polη was the first Y-family polymerase to be solved. The structure indicates that the fingers domain is reduced in size, which allows polη to accommodate both nucleotides of a CPD at the active site [Trincao et al. 2001]. The openness of the active site is a critical feature for the binding of CPDs which distinguishes it from other polymerases [Biertümpfel et al. 2010; Silverstein et al. 2010] (Figure 1.5C). Polι, identified as Rad30b, is an ortholog of polη that has no known prokaryotic ortholog [McDonald et al. 1999, 2001]. Similarly to polη, polι is able to bypass CPDs and lesions caused by oxidative stress, such as 8-oxo-dG [Vaisman et al. 2002, 2003; Zhang et al. 2001]. However, the in vivo role of polι remains unclear since it is dispensable for UV-induced mutagenesis in human cells [Wang et al. 2007]. The ability of polι to bypass lesions that are normally bypassed by polη could be a reason why XPV patients have higher rate of induced skin cancer, since they have lost the more reliable bypass mechanism [Donny Clark et al. 2009]. A feature that distinguishes polι from other polymerases is the more efficient bypass of damaged purines compared to the bypass of damaged pyrimidines [Nair et al. 2005; Zhang et al 2000; Jain et al. 2009].
In addition to the Y-family of polymerases, there is one other polymerase that participates in TLS. Polζ, which consists of two subunits called Rev3 and Rev7 [Lawrence and Hinkle, 1996; Torpey et al. 1994], was initially discovered as a polymerase required for induced mutagenesis, but that was not essential for replicative processes [Morrison et al. 1989]. Polζ has been shown to be able to bypass a wide variety of lesions, despite the absence of a PAD domain, which is an
General Introduction
26
essential domain for the Y-family polymerases to compensate for the altered geometric shape of a damaged base-pair [Yuan et al. 200; Johnson et al. 2000].
Furthermore Polζ plays a central role in coordinating the repair of post replicative gaps [Jansen et al. 2009].
Despite the lack of a 3D structure of Rev3, the catalytic subunit of polζ, there is structural data available that gives insight into its capability to bypass and extend from DNA lesions. Bacterial polymerase II (polII), an ortholog of Rev3, lacks essential carboxylates in the exonuclease domain which disable the proofreading ability.
Sequence alignment of polII and Rev3 shows that Rev3 also lacks these essential residues to perform proofreading. Furthermore, PolII possesses an N-palm linker, which could enlarge the binding cavity to fit damaged lesions and therefore increase the spectrum of lesions that can be bypassed. Rev3 also contains such an extension, which is predicted to be even longer, enhancing its capability to bypass damaged lesions [Wang et al. 2009].
While the in vitro capability of each of the TLS polymerases to bypass various types of DNA damage has been well characterized, little is known about the in vivo selection of a given polymerase upon damage encounter. For instance, Polζ can substitute for polη in the bypass of CPDs, while polδ can substitute for Rev1 in the bypass of abasic sites [Gibbs et al 2005]. Additionally, the in vitro bypass of benzo[a]pyrene adducts requires the coordination of three different polymerases, polζ, polη and Rev1 [Zhao et al. 2006]. The question remains, which of these polymerases performs the bypass synthesis in vivo? While the tool belt model seems to be logical, the model that posits sequential recruitment of TLS polymerases could explain several unanswered questions about the regulation of TLS [Ohmori et al.
2009].
Chapter I
27 Role of Rev1 in DNA Translesion synthesis.
REV1 was initially identified as a gene required for DNA Translesion Synthesis in yeast [Lawrence and Christensen, 1976], where it encodes a dCMP transferase.
While the existence of the REV1 gene is restricted to the eukaryotic kingdom, it is closely related to the prokaryotic umuc genes [Nelson et al. 1996; Lin et al. 1999]. In yeast, dCMP transferase activity of the polymerase domain is mainly required for the bypass of abasic sites, while the bypass of other lesions, such as CPDs, depends on other TLS polymerases [Nelson et al. 2000; Otsuka et al. 2002]. In higher eukaryotes, however, dCMP transferase activity does not contribute significantly to the bypass of abasic sites [Ross et al. 2005]. However, this activity is required for somatic hypermutation in the immune system [Jansen et al. 2006]. Furthermore, Rev1 from humans and yeast is capable of bypassing guanine lesions, such as 8-oxoguanine and benzo[a]pyrene diolepoxide (BPDE) in vitro [Harackska et al. 2002; Zhang et al.
2002].
The polymerase dependent role of Rev1 remains limited to the incorporation of a dCMP opposite the DNA lesion (Figure 1.6A). Although it is capable of utilizing other nucleotides, it has a strong preference for dCTP [Piao et al. 2010, Masuda and Kamiya 2002]. In addition, Rev1 is able to incorporate the oxidized form of guanine efficiently, contributing to mutagenesis induced by oxidative processes [Satou et al.
2009]. The X-ray crystal structure of Rev1 revealed that it is unique among all known DNA polymerases in that it utilizes a protein template, rather than a DNA template, to select the incoming base [Nair et al. 2005]. This fact explains the constitutive preference for dCTP independent of the sequence of the parental strand [Nair et al.
2005]. In addition to the polymerase domain, the UBMs also contribute to the polymerase dependent role of Rev1 in lesion bypass by binding to ubiquitinated PCNA [Guo et al. 2006]. However, the BRCT domain and C-terminal polymerase
General Introduction
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(Figure 1.6) Different roles for Rev1 in DNA translesion synthesis. (A) The polymerase- dependent role where Rev1 bypasses the lesion by insertion of a dCTP. (B) The polymerase independent role illustrating by interaction with a different Y-family polymerase, for example Polη, which inserts a nucleotide dependent on the nature of the lesion. (C) Extension from the lesion by polymerase ζ. Interaction of Rev1 with Rev7 facilitates the correct complex formation, and keeps polymerase ζ in frame with the template.
interaction domain are dispensable for the polymerase dependent role of Rev1 during TLS [Ross et al. 2006].
Despite the limited contribution of the dCMP transferase activity of Rev1 to the bypass of lesions in vivo, Rev1 still seems to be required for TLS. Studies in which the dCMP transferase activity of Rev1 is rendered non functional by mutation have shown that the presence of otherwise intact Rev1 increases the viability of cells that have sustained DNA damage [Gibbs et al. 2000; Otsuka et al. 2005; Simpson and Sale, 2003]. This polymerase-independent role of Rev1 requires interaction with and stimulation of the other TLS polymerases (Figure 1.6B [Otsuka et al. 2005, Ohashi et al. 2009]. The binding of Rev1 to the other Y-family polymerasess and polζ is conserved across species although the exact mechanism has diverged. In mammalian cells the C-Terminal domain plays an important role. In yeast however, this interaction also occurs via the PAD domain [Achyara et al. 2005; Kosarek et al.
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2008].The interaction of polη with Rev1 is the best characterized. Here the activity of polη requires complex formation with Rev1, rather than binding to PCNA via the UBMs [McDonald et al. 1997; Tissier et al. 2004; Achayra et al. 2007; Akagi et al.
2009]. In addition to the C-terminal polymerase binding domain, the BRCT domain is also required for the polymerase independent role of Rev1. Although an intact N- terminus of Rev1 is required to stimulate the synthetic activity of TLS polymerases, a molecular basis for this stimulation remains elusive [Nelson et al. 2000; Jansen et al.
2005; Otsuka et al. 2005].
Most lesion bypass performed by TLS polymerases, including Rev1, requires polζ activity to extend past the site of damage (Figure 1.6C) [Washington et al. 2004; Nair et al. 2008; Haracksa et al. 2001, Yang et al. 2009]. In return, Polζ requires the interaction with Rev1 in order to function. This interaction stimulates the enzymatic activity of both proteins in vitro and in vivo in yeast [Murakumo et al. 2001; Guo et al. 2004; Okada et al. 2005]. In humans, complex formation of Rev1 and polζ is also required for TLS, however, the catalytic activity of Rev1 is not stimulated by this interaction [Masuda et al. 2003]. Not only is Rev1 responsible for recruiting polζ to the replication fork, it also orientates the polymerase correctly at the 3’primer end, avoiding the introduction of a possible frameshift mutation [Baynton et al. 1999;
Szuts et al. 2008].
In both yeast and higher eukaryotes, interactions between Rev1 and Rev7 are dependent on the minimal C-terminal fragment of Rev1 [D’Souza et al. 2008, Guo et al. 2003]. Furthermore, the hypothetical structure of the yeast Rev1-Rev7 complex reveals a possible interaction between Rev7 and the PAD domain of Rev1 [Acharya et al. 2005]. However, this interaction remains speculative for the human Rev1-Rev7 complex [Masuda et al. 2003; Hara et al. 2010]. Upon binding of Rev3, Rev7 undergoes several structural changes that result in formation of a binding interface for Rev1. Recruitment of polζ to the 3’ terminus displaces the previously active TLS
General Introduction
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polymerase and enables extension past the lesion [Hara et al. 2010]. Most interestingly, recent studies have shown that Rev1 can also bind polδ , independent of PCNA. Possibly this interaction also plays a role in the switch back to high fidelity DNA synthesis by polδ once polymerase ζ has extended a short length of DNA from the lesion [Acharya et al 2009].
Surprisingly, the majority of Rev1 expression in yeast does not occur in S-phase as would be expected, but rather occurs in G2/M phase [Waters and Walker, 2006].
Furthermore, Rev1 expression is dramatically increased upon high levels of UV exposure, which might suggest that the DNA damage response pathways requiring Rev1 are only induced by the presence of exceptionally high levels of DNA damage and that the mutagenic nature of these pathways is avoided under physiological conditions [Waters and Walker, 2006]. However, in human cells, Rev1 accumulates as a result of both induced and spontaneous DNA damage, suggesting that here the pathways requiring Rev1 are more generally involved in the response to DNA damage [Akagi et al. 2009]. It seems therefore that Rev1 have evolved somewhat different functions in yeast and higher eukaryotes. In higher eukaryotes, the polymerase independent role of Rev1 could be essential for organizing TLS.
Although the polymerase dependent role of Rev1 is relatively minor in TLS, the polymerase independent roles of Rev1 are of more importance. This is also reflected by the observation that Rev1 performs separate organizing roles in both ubiquitin dependent and independent TLS [McDonald et al. 1997; Edmunds et al. 2008]. For example, it has recently been shown that Rev1 cooperates with helicases belonging to the Werner Syndrome family (WRN). This research shows that the primary pathway to bypass a lesion depends on Rev1 and WRN in a ubiquitin independent manner. The secondary pathway involves ubiquitin dependent TLS, and is also dependent on Rev1. However, this pathway is only followed if the primary pathway fails or under conditions of heavily induced DNA damage [Philips and Sale, 2010].
Chapter I
31 Interstrand Cross Link repair by Rev1
Interstrand crosslinks (ICL), which are caused by electrophylic chemicals such as cisplatinum or nitrogen mustard, are one of the most persistent types of DNA damage. The damage caused by these chemicals usually triggers apoptosis, which is the main reason why they are used as anti-cancer drugs. In some cases ICLs can be correctly repaired allowing the cell to survive. In G0 and in G1 phase, the main pathway to repair these lesions is NER. However, the majority of these lesions are repaired in S/G2 phase, in a replication dependent manner (Reviewed [Ho and Scharer, 2010]). The key proteins involved are Rev1, polζ and the Fanconi Anemia proteins [Shen et al. 2006; Sarkar et al. 2006; Knipscheer et al. 2009]. The Fanconi Anemia core complex is able to recruit a variety of DNA repair proteins, including Rev1, and performs an organizing role in replication coupled ICL repair [Niedzwiedz et al. 2004; Knipscheer et al. 2008 ].
The mechanism of replication-coupled ICL repair was elegantly demonstrated in Xenopus egg extracts [Raschle et al. 2008]. In this cell free system, both the leading and the lagging strands stall at a distance of roughly 20 nucleotides from the ICL (Figure 1.7A). Subsequently, slow replication is observed on the leading strand advancing precisely to the base next to the lesion (Figure 1.7B). On one of the strands, incisions are made on both sides of the lesion by a member of the Fanconi family, the Fanconi Associated Nuclease 1 (FAN1) (figure 1.7C) [Huang and D’Andrea 2010], resulting in the separation of the sister chromatids, which are held in place by the Fanconi core complex. The separated strands are rejoined by HR in a later phase (not shown in this figure). Next, Rev1 bypasses the lesion by inserting a dCMP opposite the damage (Figure 1.7D), followed by extension past the lesion by polζ (Figure 1.7E). The polymerase dependent role of Rev1 in this pathway, however, can be substituted by polκ [Minko et al. 2008]. However, Rev1 likely performs a polymerase independent role in this process as well [Hlavin et al. 2006].
General Introduction
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(Figure 1.7) Mechanism of replication-coupled repair of inter strand crosslinks. (A) The DNA is completely replicated from both sides until the polymerase advancing towards the ICL stalls roughly 20 nucleotides from the lesion. (B) The DNA polymerase advances slowly to the lesion. (C) Incisions are made separating the two sister strands. (D) Rev1 inserts a dCTP opposite of the damage. (E) Polζ extends the DNA replication and fills the gap (modified from [Räschle et al. 2008]).
Interestingly, the recruitment of Rev1 by the Fanconi core complex does not require the UBMs of Rev1, but is dependent on the BRCT domain [Mirchandani et al. 2008].
The polymerase-independent role could be related to the Rev1 BRCT domain, however, the exact nature of the involvement of the Rev1 BRCT domain remains unknown.
Biological relevance of the Rev1 BRCT domain.
The BRCT domain is one of the most puzzling aspects of Rev1. The BRCT domain was initially reported to be required for the polymerase independent role of Rev1 in
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yeast cells [Nelson et al. 2000], but the role of the BRCT domain remains unclear in higher eukaryotes. Here the data describing the biological role of the Rev1 BRCT domain are reviewed.
The Rev1 BRCT domain has proven to be dispensable for efficient DNA damage tolerance in chicken DT40 cells, however, in these experiments the over expression of Rev1 could compensate for the deletion of the BRCT domain [Ross et al. 2005].
Furthermore, mouse cells with a targeted deletion of the BRCT domain show a threefold reduction in UV-induced mutation frequency [Jansen et al. 2005]. It was further noted that the deletion of the BRCT domain resulted in a reduction by approximately 50% in the number of replication foci found in S-phase. Additional deletion of the UBMs of Rev1 completely abolishes replication focus localization altogether [Guo et al. 2006]. This data suggests that the Rev1 BRCT domain is not an essential, but more a kinetic factor in the polymerase-independent role of Rev1 in TLS. Possibly, the functional interaction between Rev1 and the polymerase which bypasses the damage is not altered when the BRCT domain is deleted, this interaction occurs at a lower frequency due to the less efficient localization of Rev1 to the site of damage. Mouse embryonic stem cells containing a targeted deletion of the BRCT domain display delayed progression through late S-phase, while cells that are completely deficient for Rev1 do not complete S-phase at all [Jansen et al. 2009].
This data suggests again that the BRCT domain is not essential but rather is required for efficient lesion bypass in late S-phase and that it might be significant for DNA damage tolerance under more physiological conditions.
The BRCT domain has been reported to mediate the interaction between PCNA and Rev1 [Guo et al. 2006] and between Rev1 and Rev7 [D’Souza and Walker, 2006].
However, a direct interaction remains questionable, since common binding to either DNA or a protein might govern this interaction as well. Furthermore, other domains also govern this interaction in both yeast and humans [Wood et al. 2007; Masuda et
General Introduction
34
al. 2003]. Interestingly, substitution of the BRCT domain of Dbf4, a regulatory protein required for Cdc7K kinase activity in yeast, with the yeast Rev1 BRCT domain does not alter the viability of yeast cells suggesting both BRCT domains have a common binding partner [Harkins et al. 2009].
The BRCT domain has also been observed to be significant for non-TLS roles.
Surprisingly, Rev1 has recently been connected with stabilization of trinucleotide repeats in yeast. Here, Rev1 reduces the contraction and expansion of CAG-CTG repeats, which solely depends on the BRCT domain [Collins et al. 2007]. While these contractions are mainly repaired by post replication repair, and more specifically, large loop repair, it is possible an interaction of the BRCT domain with DNA or a protein might govern the activity of Rev1 in this process [Daee et al. 2007; Sommer et al. 2008].
It has been observed that Rev1 is recruited to sites of double stranded DNA breaks and that this recruitment is independent of PCNA ubiquitination. This recruitment however, does appear to be dependent on the BRCT domain [Hirano et al. 2009, Hirano and Sugimoto 2006]. Rev1 found at the site of the double stranded break is associated with polymerase ζ in a Mec1 dependent manner [Hirano and Sugimoto, 2006; Kolas and Durocher, 2006]. As with TLS, modification of PCNA governs different pathways with respect to the repair of double stranded breaks [Moertl et al. 2008]. It is possible that Rev1 could perform a similar organizing role in the repair of double stranded breaks as in TLS and that this role is dependent on the BRCT domain.
BRCT domains.
BRCT domains were first identified as the C-terminal domain of the Breast Cancer type 1 Susceptibility protein (BRCA1), where it consists of a tandem repeat of roughly 90 amino acids [Koonin et al. 1996]. Nowadays, these domains represent a
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broad superfamily that is directly or indirectly associated with DNA repair, replication, or the regulation of cell-cycle checkpoints. However, the sequence similarity across the BRCT superfamily is rather low [Bork et al. 1997; Callebaut and Mornon, 1997]. BRCT domains can occur in proteins as single, tandem or multiple repeats, where they mainly mediate protein-protein interaction [Derbyshire et al.
2002; Williams et al 2001]. Several studies indicate that BRCT domains mediate
(Figure 1.8) Different structures involving BRCT domains that mediate protein-protein interactions. (A) The BRCT domains of XRCC1 form a symmetric homodimer, involving the α1 helix. (B) In the tandem repeat of 53BP1, the inter- domain linker performs a central role in protein-protein interaction, as well as the α3-helix of the N-terminal BRCT motif. (C-D) Phosphopeptide binding by the tandem repeats of BRCA1 mediate its interaction with BACH1.
Two factors are important: Binding of the phosphorylated serine by residues in the N-terminal BRCT motif (blue) (C). Residues located on the α2 helix of the C-terminal BRCT motif, the α1 and α3 helices of the N-terminal BRCT motif and the inter-linker domain (blue) bind specifically to the phenylalanine at the +3 position from the phosphorylated serine in the peptide (D).
General Introduction
36
protein-protein interactions by either BRCT-BRCT or BRCT-non-BRCT pairing [Sibanda et al. 2001; Yu et al 2003; Sum et al. 2002, Liu et al. 1994]. However, these interactions are structurally different.
All BRCT domains share a conserved overall protein fold: a core of four β strands, which are flanked by two α helices on one side, and a third α helix on the other side [Williams et al. 2001; Zhang et al. 1998; Joo et al. 2002; Krishnann et al. 2001] . In the known structures of BRCT dimers from isolated BRCT domains, the α1 helix and the N-terminus mediate the dimerization (Figure 1.8A) [Zhang et al. 1998], while in the tandem repeats of 53BP1, a protein that binds p53 to enhance it transcriptional activation, the inter-linker domain performs a central role in mediating the protein- protein interaction (Figure 1.8B) [Derbyshire et al. 2002; Joo et al. 2002]. The structures of tandem BRCT domains indicate that dimerization is asymmetric, occurring in a head-to-tail fashion. The binding interface consists of the α2 helix of the N-terminal BRCT domain interacting with the α1 and α3 helices of the C-terminal domain [Glover et al. 2004].
Studies have indicated that tandem BRCT domains from a variety of proteins specifically recognize unstructured peptides containing a phosphorylated serine and a specific amino acid at the +3 position from the serine. Binding of tthis pSer-x-x-y motif (where x can be any amino acid, and y indicates the specifically recognized amino acid) was first identified for the tandem repeats of BRCA1. and MDC1 [Rodriguez et al. 2003; Yu et al. 2003; Lee et al. 2005]. Interestingly, several isolated BRCT domains were also found to bind the phosphoserine motif, including the BRCT domain of yeast Rev1 [Yu et al. 2003]. However, the subsequent crystal structures clearly demonstrate that recognition of the phosphorylated serine motif requires both BRCT domains of a tandem repeat. Binding of the phosphate solely depends on amino acids within the N-terminal BRCT domain (Figure 1.8C). However, recognition of the phenylalanine at the +3 position of the peptide requires amino acids in the N-
Chapter I
37
terminal BRCT, the linker and the C-terminal BRCT [Botuyan et al. 2004; Campbell et al. 2010; Lee et al. 2010;] (Figure 1.8D). Therefore, either the isolated BRCT domains bind phosphoserine peptides using a different mechanism or the interpretation of the original data is inconclusive.
In addition to protein-protein interactions, a limited number of BRCT domains are able to bind DNA. The BRCT domain of the bacterial NAD+ dependent ligase for instance, binds DNA specifically, where it is required to bind the single stranded nick
(Figure 1.9) DNA binding by the Rfc1 BRCT region. (A) Binding of the 5’ phosphate by the BRCT domain. (B) Binding of the double stranded DNA by the 28 amino acids N-terminal to the BRCT domain. Here the α1’ helix is located in the major groove of the DNA. (C) Residues involved in phosphate binding (blue), and residues involved in double stranded DNA binding (D).
General Introduction
38
in DNA. Deletion of the BRCT domain from the ligase severely reduces the ability to form the phosphodiester bond [Feng et al. 2004; Jeon et al. 2004; Wilkinson et al.
2005]. The BRCT domain of the large subunit of RFC, Rfc1, is also able to bind DNA [Kobayashi et al. 2006]. Not only does this require a functional BRCT domain, it also requires approximately 28 additional amino acids N-terminal to the BRCT domain. In the elucidated ternary complex, binding of the 5’ phosphate appears to be mainly mediated by the BRCT domain itself, which is similar to the N-terminal BRCT domain of BRCA1 (Figure 1.9A). However, the sequences N-terminal to the BRCT domain form an α-helix (α1’) that lies within the major groove of the DNA [Kobayashi et al.
2010] (Figure 1.9B). Several key residues dictate the specificity for a 5’phosphate and the double stranded DNA. The conserved residues T415, G416 and K458 interact with the oxygen atoms of the 5’ phosphate. Residues K375, Y382 and R388 of α1’
interact with the DNA backbone in the major groove. Residues R423 and N440 interact with respectively the base and the backbone phosphate of the single stranded DNA. (Figure 1.9C-D) [Kobayashi et al. 2010].
Rev1 BRCT as a potential DNA binding domain.
Although the amino acid sequence similarity of the BRCT domain of Rev1 to the BRCT domain of Rfc1 is not outstanding, both domains are structurally very similar (Figure 1.10) [Thompson et al. 1994]. Secondary structure prediction of Rev1 revealed an α-helix N-terminal to the BRCT region which is analogous to α1’ of Rfc1 (Figure 1.10A) [McGuffin et al. 2000]. In Rfc1 this α-helix is essential for binding of DNA [Kobayashi et al 2006]. In order to clarify the requirement for residues outside of the BRCT domain for the biological function, these are referred to as BRCT regions. While the Rev1 BRCT has already been reported to bind DNA [Kobayashi et al. 2006], it is unknown whether it is mechanistically similar to DNA binding by Rfc1
Chapter I
39
(Figure 1.10) Comparison of the mouse Rev1 BRCT region with the RFC BRCT region. (A) Alignment of the Rfc1 and Rev1 BRCT sequences from multiple species by ClustalW. Only the mouse Rev1 and human Rfc1 are shown. Indicated below the sequence is the secondary structure. Indicated by a Dashed boundary is the predicted secondary structure using the PPsired algorithm. Structures of respectively the Rfc1 BRCT region (B), the Rev1 BRCT region (C) and the structural superposition of the two structures (D).
and the bacterial ligase. Considering its biological relevance, further determination of the nature of the DNA binding properties of the Rev1 BRCT region is warranted in order to identify its potential role in the biological function of Rev1.
Outline of this thesis.
This thesis reports the results of studies primarily aimed at investigating a potential interaction between the BRCT region of Rev1 and DNA. In chapter II, binding of Rev1 to DNA is demonstrated and the nature of the DNA structure that is required for binding is determined. Several different yeast and mouse proteins were generated
General Introduction
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to investigate the amino acid requirements for DNA binding. Furthermore the binding kinetics are analyzed. Chapter III describes the results of point mutagenesis studies on DNA binding by the Rev1 BRCT region. Here, residues that perform key roles in binding of the DNA and recognizing structural features of the substrate are identified. Using the RFC-DNA complex as blueprint, a model structure of the Rev1 BRCT-DNA complex is obtained. In Chapter IV DNA binding by the full length Rev1 protein is compared to the various BRCT region constructs. Using N-terminally truncated proteins but otherwise intact, full length Rev1, both 3’ terminus and 5’
terminus binding has been found and characterized. Chapter V describes the determination of the cellular function of the RFC BRCT region. By generating several yeast strains that contain knockouts of different genes connected to DNA repair or replication, different pathways are explored in which the RFC BRCT region might function.
Chapter I
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