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Dynamics of nucleotide excision repair complex assembly and disassembly in vivo

Luijsterburg, M.S.

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Citation for published version (APA): Luijsterburg, M. S. (2008). Dynamics of nucleotide excision repair complex assembly and disassembly in vivo.

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Dynamics of Nucleotide Excision Repair Complex Assembly and Disassembly In Vivo

Martijn S. Luijsterburg 2008 DYNAMICS OF NUCLEOTIDE EXCISION REPAIR COMPLEX ASSEMBLY AND DISASSEMBLY IN VIVO

ACADEMISCH PROEFSCHRIFT

ter verkrijging van de graad van doctor aan de Universiteit van Amsterdam op gezag van de Rector Magnificus prof. dr. D.C. van den Boom ten overstaan van een door het college voor promoties ingestelde commissie, in het openbaar te verdedigen in de Agnietenkapel op vrijdag 12 september 2008, te 12:00 uur

door

Martijn Sebastiaan Luijsterburg

geboren te Utrecht Promotiecommissie

Promotor: Prof. dr. R. van Driel overige leden: Prof. dr. L.H.F. Mullenders Prof. dr. T. Höfer Prof. dr. T.W. Gadella Prof. dr. N. Nanninga Dr. W. Vermeulen Dr. N.P. Dantuma Dr. J. Goedhart

Faculteit der Natuurwetenschappen, Wiskunde en Informatica

The research desribed in this thesis was financially supported by the Netherlands organisation for scientific research (NWO)

2 Table of contents

Chapter 1 5 DNA Repair Systems in Mammals

Chapter 2 19 Nucleotide Excision Repair in Mammals

Chapter 3 45 Recruitment of the Nucleotide Excision Repair Endonuclease XPG to Sites of UV-induced DNA Damage Depends on Functional TFIIH

Chapter 4 63 Dynamic In Vivo Interaction of DDB2 E3 Ubiquitin Ligase with UV-damaged DNA is Independent of Damage-Recognition Protein XPC

Chapter 5 83 Spatial Organization of Nucleotide Excision Repair Proteins After UV-induced DNA Damage in the Human Cell Nucleus

Chapter 6 97 Choreography of Nucleotide Excision Repair Unraveled by Live-Cell Imaging and Kinetic Modeling

Chapter 7 123 Heterochromatin Protein 1 is Involved in the DNA Damage Response

Chapter 8 139 Assembly of Multi-Protein Complexes that Control Genome Function

Bibliography 151

Summary 177

Samenvatting 180

Curriculum vitae 183

3 4 Chapter 1 DNA Repair Systems in Mammals

Martijn S. Luijsterburg and Roel van Driel

5 Chapter 1

Introduction A series of elegant experiments in the 1950s demonstrated that DNA (deoxyribonucleic acid) is the carrier of genetic information (Hershey and Chase, 1952). This heritable information is crucial for cellular identity and is therefore faithfully replicated and passed on to daughter cells. The building blocks of DNA are the nucleotides that contain four nitrogenous bases (A,T,C and G) and the sequence of these bases encodes the recipes of all processes in the cell. The structure of DNA is a double helix, in which two complementary DNA strands are kept together by hydrogen bonds between the bases on opposite strands (Watson and Crick, 1953). Despite the fact that DNA is a rather stable molecule, it is not inert and can be compromised in many ways during the lifetime of an organism. The intrinsic instability of DNA can lead to spontaneous hydrolysis of nucleotides resulting in the loss of bases (abasic sites). Moreover, errors made during DNA replication and reactive compounds formed during cellular metabolism (such as oxygen radicals) pose an intracellular threat to the integrity of DNA. For example, more than 100 types of oxidative modifications have been identified in DNA as the result of cellular metabolism (Hoeijmakers, 2001). In addition to endogenous sources of DNA damage, various external factors such as ionizing radiation (IR) and ultra-violet (UV) light can also damage DNA. Ionizing radiation can result in the formation of single-strand and double-strand breaks, whereas UV light induces cross-links between adjacent bases in the same DNA strand (intrastrand cross-links). Various genotoxic chemicals present in nutrition or the environment link themselves directly to the DNA and form bulky adducts or cross-links between two different DNA strands (interstrand cross-links). An example of a highly cytotoxic cross-linking agent is psoralen, which has two reactive sites that can form adducts with two nucleotide bases simultaneously. It is estimated that each hour in every cell between 100-1000 oxidative lesions and ~50 double strand breaks are produced either spontaneously or by cellular metabolism (Klungland et al., 1999; Vilenchik and Knudson, 2003). In addition, solar UV light is responsible for the formation of ~30.000 DNA lesions per cell per hour. It is not surprising that cells have evolved sophisticated systems to cope with DNA damage. Repair of lesions in the genomic DNA and more specifically by the nucleotide excision repair machinery is the main topic of this thesis. First, the effects of the presence of DNA lesions on cellular processes such as replication and transcription will be discussed. Then, an overview will be given of the versatile repair pathways utilized by mammalian cells. Finally, nucleotide excision repair will be discussed in more detail.

Cellular responses to DNA damage The presence of DNA lesions interferes with correct functioning of transcription and replication, which involves the translocation of polymerases along the DNA. Lesions in the DNA physically obstruct polymerases and damages in actively transcribed genes result in inhibition of gene expression due to stalling of RNA polymerase II (RNAPII)

6 DNA repair systems in mammals

(Brueckner et al., 2007; Moné et al., 2001). Persistent transcription stress arising from blockage of RNAP II at lesions constitutes an efficient trigger for p53-dependent apoptosis (Ljungman and Zhang, 1996). This could be an efficient strategy to eliminate cells that might potentially form tumors. Rather than transcription elongation blockage due to stalling of RNAPII at lesions, transcription initiation also appears to be repressed upon damage induction (Rockx et al., 2000). Similarly to transcription, DNA lesions interfere with replication by DNA polymerase δ (lagging strand) or pol ε (leading strand) and possibly by DNA pol α (which is the primase during DNA replication). A set of specialized DNA polymerases (denoted pol ζ to pol κ) temporarily takes over from stalled replicative polymerases to overcome replication stress by DNA lesions (Friedberg et al., 2002). These special polymerases have more flexible base-pairing properties that allow DNA synthesis past DNA lesions (Lehmann et al., 2007). However, this occurs at a high error rate (100- to 1000-fold lower fidelity than replicative polymerases) and translesion DNA synthesis is responsible for the majority of damage-induced point mutations. Nevertheless, TLS is still important for genome stability, which is illustrated by the skin cancer-prone phenotype of inherited defects in the TLS polymerase η (Masutani et al., 1999). For correct functioning of cells it is of vital importance to counteract the deleterious effects of DNA damage since unrepaired lesions can lead to mutations, which can eventually result in the formation of tumors. Moreover, the persistence of DNA damage in the genome is believed to contribute to aging (Hasty et al., 2003; Hoeijmakers, 2007). Cells have evolved sophisticated repair pathways to remove a plethora of different DNA injuries from the genome. These different DNA repair pathways will be briefly discussed in the next section.

DNA repair pathways Cells utilize an impressive arsenal of DNA repair enzymes to remove a variety of lesions from their genome. No single repair pathway can cope with all kinds of lesions and cells have evolved several different repair pathways that are highly conserved throughout the kingdoms of life. Damages that affect only one of the two DNA strands are removed by excision repair, in which the injury (usually with a flanking DNA region) is removed and the resulting gap is re-synthesized using the complementary DNA strand as a template. Oxidative lesions (usually of endogenous origin) are mainly removed by base excision repair (BER), whereas nucleotide excision repair (NER) removes a variety of helix-distorting lesions including UV-induced damage and bulky adducts. Excision repair is in principal error-free. Damages that affect both DNA strand, such as double-strand breaks (DSB) are more difficult to repair than damages that affect only one of the DNA strands. Cells can either use the sister-chromatid that is present after DNA replication as a template to direct the error-free repair of DSBs (this pathway is know as homologous recombination; HR) or cells can fuse the broken ends together in the absence of a second copy in an error-prone mechanism termed

7 Chapter 1 non-homologous end-joining (NHEJ). Repair of interstrand cross-links (i.e. between complementary DNA strands) requires factors of the Fanconi anemia (FA) pathway. The precise mechanism of the FA pathway is currently unknown. Certain lesions can be converted to other types of lesions that might be more amenable for repair. For example, interstrand cross-links are often converted into DSBs that can be removed by HR or NHEJ. The following sections will briefly address the main characteristics of the different repair pathways. Nucleotide excision repair, which is the main topic of this thesis, will be discussed in more detail.

Base excision repair Base excision repair removes a variety of base alterations induced by spontaneous hydrolysis of bases, oxidative events, ionizing radiation or alkylating agents. More than 100 different types of oxidative lesions have been identified in mammalian cells (Hoeijmakers, 2001) and examples of frequently occurring highly mutagenic lesions are 8-oxoguanine and O6-methylguanine (Lindahl and Wood, 1999). These lesions can form non-Watson-Crick base pairs (e.g. A opposite to 8-oxyG and T opposite to O6-methylguanine), resulting in the misincorporation of nucleotides during replication leading to mutations. Some oxidized pyrimidines (such as cytosine and thymine glycol) are poorly mutagenic but strongly inhibit replicative DNA polymerases making these lesion highly cytotoxic (Aller et al., 2007). Moreover, the faulty incorporation of uracil (i.e. deaminated cytosine) in the DNA alters the binding affinities of DNA binding proteins and complete substitution of T with U in cellular DNA is lethal (Lindahl and Wood, 1999). In order to remove oxidative lesion from the genome, mammalian cells employ a variety of DNA glycosylases that each act on a different subset of base alterations (see Fig 1). Generally, DNA glycosylases flip out the altered base to accommodate it in an internal cavity of the protein (Krokan et al., 1997). Inside the protein, the bond between the sugar and the altered base (i.e. the glycosidic bond) is incised resulting in an apurinic/apyrimidinic (AP) site. Examples of DNA glycosylases are OGG1 (incision of 8-oxyG opposite to a C), NTH1 (incision of thymine/cytosine glycol, which requires NER endonuclease XPG as a co-factor (Klungland et al., 1999)) and UNG1 (incision of uracil). Inactivation of individual glycosylases in mice has no obvious phenotype, suggesting partial redundancy between different glycosylases (Parsons and Elder, 2003; Wilson and Thompson, 1997). The subsequent steps during BER involve the binding of an endonuclease with affinity for AP sites (called APE/HAP1), which makes an incision at the 5’ side of the AP site opening up the sugar-phosphate backbone creating a normal 3’-hydroxyl group and a 5’-deoxyribose-phosphate residue (i.e. a baseless sugar) that needs to be removed to complete repair (Hoeijmakers, 2001; Lindahl and Wood, 1999). The major BER pathway is the so called short-patch repair, in which DNA pol β incorporates a single nucleotide and removes the 5’-terminal baseless sugar residue using its intrinsic lyase activity (Allinson et al., 2001; Matsumoto and Kim, 1995). The resulting nick is sealed

8 DNA repair systems in mammals by XRCC1-Ligase III (left pathway in Fig 1). Alternatively, during long-patch BER pathway a stretch of 2-8 nucleotides is incorporated by DNA pol δ/ε that replaces the original stand. This BER pathway requires proliferating cell nuclear antigen (PCNA) to stimulate pol δ/ε and the endonuclease FEN1 to incise and release the displaced DNA strand. Finally, the resulting nick is sealed by DNA ligase I (right pathway Fig 1). The choice between short- and long patch BER probably depends on the type of lesion that BER is faced with and the cell cycle phase (Fortini and Dogliotti, 2007). The importance of BER is underscored by the fact that inactivation of core BER proteins (e.g. XRCC1, pol β, Lig 1 and APE1) results in embryonic lethality in mice (Wilson and Thompson, 1997). In agreement, no disorders caused by inherited defects in BER have been identified in humans.

Mismatch repair Replication of the genomic DNA by the mammalian DNA polymerases α, δ and ε is not error-free (1 error every ~1.106 nucleotides) and base-base mismatches as well as small insertion or deletion mispairs are often produced by replication errors (Kunkel, 2004). Moreover, uncorrected base alterations (such as 8-oxyG) contribute to misincorporation of nucleotides during replication. There can be 8 different types of mismatches in genomic DNA (AA, TT, CC, GG, AG, TC, CA and GT), which can lead to stable mutations if left uncorrected. The mismatch repair (MMR) machinery increases the fidelity of DNA replication with several orders of magnitude by recognizing and correcting all possible base-base mismatches and small insertions/ deletions. The MMR pathway in Escherichia coli has been well characterized and provides an excellent model for MMR in mammals (Kunkel and Erie, 2005). The general MMR mechanism comprises the following steps: i) mismatch recognition, ii) discrimination between the parental (i.e. correct) and daughter (i.e. erroneous) strand, iii) excision of an ssDNA fragment and iv) gap-filling by DNA pol and sealing the nick by DNA ligase. Recognition of mismatches in bacteria is carried out by the MutS and MutL proteins (that will be discussed in more detail below). Discrimination between parental and daughter strands is then directed by DNA methylation that is carried out by deoxyadenine methylase (DAM), which methylates adenine residues in a GATC context (Jiricny, 2006). The DAM enzyme lags behind the replication fork by ~2 minutes, which means that newly replicated daughter strands are transiently unmethylated. The MutS and MutL proteins use this short time window to incise the erroneous daughter strand (in E. coli this also requires the MutH endonuclease, which is absent in mammals) (Jiricny, 2006). Human MMR can be reconstituted in vitro using the following proteins: MutSα, MutSβ, MutLα, replication factor C (RF-C), replication protein A (RPA), PCNA and DNA pol δ. In addition, Lig 1, HMG-B and exonuclease I (EXO1) are needed (Dzantiev et al., 2004; Zhang et al., 2005). However, it is unknown how the mammalian MMR machinery targets the newly synthesized DNA strand for repair since adenine methylation is absent in eukaryotes. In mammals, strand

9 Chapter 1

discrimination may be directed by interactions between MMR proteins and the nearby replication machinery, since MutSα and MutSß directly interact with PCNA(Hoeijmakers, 2001; Masih et al., 2007; Umar et al., 1996). Alternatively, it has been suggested that strand discrimination during MMR is directed by strand breaks in close vicinity of the mismatch and indeed substrates used for in vitro MMR assays can only be processed when containing nicks (Dzantiev et al., 2004; Zhang et al., 2005). The following working model for MMR in mammals has been proposed (see Fig 2). MMR of base- base mismatches is initiated by the MutSα dimer (composed of Msh2 and Msh6) and recognition of insertion/deletions mispairs is carried out by MutSβ (composed of Fig 1. Model for the mechanism of oxidized base damage repair by base excision repair. Msh2 and Msh3) (Jiricny, 2006). Adapted from (Hoeijmakers, 2001) Inactivation of Msh2 in mice completely inactivates MMR and results in a severe cancer-prone phenotype (de Wind et al.,

Fig 2. Model for the mechanism of base-base mismatch repair by mismatch repair. Adapted from (Dzantiev et al., 2004; Jiricny, 2006; Zhang et al., 2005)

10 DNA repair systems in mammals

1995). The MutSα homodimer encircles DNA and can move along the DNA contour as a sliding clamp. Once bound to a mismatch, MutSα recruits MutLα (another sliding clamp, composed of Mlh1 and Pms2) and the MutS-MutL complex dissociates from the mismatch and translocates in either direction from the mismatch until it encounters a DNA strand break (Gradia et al., 1999). Similar to Msh2, mice lacking functional Mlh1 are very cancer-prone, illustrating the importance of functional MMR (Prolla et al., 1998). At a strand break, RF-C is bound at the 5’ side of the nick and recruits PCNA at the 3’ side. MutS-MutL clamps that diffuse upstream of the mismatch will encounter RF-C (that is bound 5’ of the nick), which is displaced by the MutS-MutL clamp followed by recruitment of the exonuclease EXO1. A ssDNA fragment of ~150 nucleotides towards and past the mismatch is subsequently degraded by EXO1 in a 5’-3’ orientation and the resulting single-stranded gap is stabilized by RPA (Jiricny, 2006). The exonuclease activity of EXO1 is inhibited by MutLα and RPA when the mismatch is removed (Zhang et al., 2005). Finally, pol δ is loaded by PCNA (still bound at the 3’-side of the original nick) and fills in the gap. Repair is completed by DNA lig I that seals the nick. However, MutS-MutL clamps could also diffuse downstream of the mismatch, where PCNA is encountered at the 3’ side of the nick. EXO1 is also recruited here and degrades a ssDNA fragment (past the mismatch) in a 3’-5’ orientation, while RF-C bound at the 5’ side of the nick prevents degradation in the 5’-3’ direction (i.e. away from the mismatch) (Jiricny, 2006). It is surprising that EXO1 can catalyze 3’-’5 strand degradation in vitro, since it is a 5’-3’ exonuclease (Tran et al., 2004). Inactivation of EXO1 is mice result in a cancer-prone phenotype but does not completely abolish MMR, suggesting that other exonucleases might play a role during MMR in vivo (Wei et al., 2003). Indeed, the 3’-5’ proofreading activity of pol δ/ε have been implicated in MMR as well as the 3’-5’ exonuclease activity of DSB repair protein MRE11 (Tran et al., 1999; Vo et al., 2005). Alternatively, the endonuclease activity of MutLα was suggested to cleave distal from the mismatch creating a 5’ terminus that allows 5’-3’-directed degradation by EXO1 (Kadyrov et al., 2006). After removal of the mismatch by the 3’-5’ exonuclease activity, the resulting gap is subsequently filled by pol δ and sealed by Lig 1 to complete the repair process. The importance of MMR in human carcinogenesis is illustrated by the finding that ~50% of the hereditary colon cancers carry mutation in genes encoding mismatch repair proteins (Hoeijmakers, 2001). Moreover, instability of dinucleotide repeats in the human genome that are caused by defects in MMR is often associated with cancer.

Homologous recombination-mediated repair Breaks in both strands of the DNA (double strand breaks; DSB) are the most dangerous threat to genomic integrity. At the DSB, genetic information is lost that cannot be recovered from the same DNA molecule. DSBs are induced by ionizing radiation, several anti-tumor agents (e.g. cisplatin) or during replication of cyclobutane pyrimidine dimers (CPD) (Garinis et al., 2005), single-stranded breaks or oxidative

11 Chapter 1 lesions. Mammalian cells utilize homologous recombination (HR) to direct the error- free repair of DSBs. Homologous sequences are present on the homologous chromosome as well as on the sister chromatid that is present after DNA replication. HR-directed repair is mainly active in S and G2 (i.e. when a sister chromatid is present) indicating that recombination-directed repair between the homologous chromosomes is rare. In fact, recombination between homologous chromosomes is potentially dangerous since it may lead to homozygosity for recessive mutations. Mammalian HR- directed repair comprises the following steps (also see Fig. 3): i) recognition of the DSB and processing of the DNA ends to create single-stranded regions, ii) transfer to the homologous sequence and strand invasion of the single-stranded donor DNA into the double-stranded target sequence, iii) DNA repair synthesis using the intact DNA sequence as a template (and the processed single-stranded DNA ends as a primer) to direct re-synthesis of the damaged strand and iv) separation of the recombined molecules by resolution of the crossed DNA strands and ligation of the nicks (Wyman and Kanaar, 2006; Wyman et al., 2004). The cellular response to DSB involves arrest of the cell cycle, which is mediated by the checkpoint kinases ATM and ATR, providing time to repair the damaged chromosomes (McGowan and Russell, 2004). Moreover, an early cellular response to DSB is the phosphorylation of histone variant H2A.X but since inactivation of H2A.X in mice does not lead to a repair-deficient phenotype it might be non-essential for repair (Bassing et al., 2002; Celeste et al., 2002). Finally, cells respond to DSBs by assembling repair proteins that mediate recombination between the damaged and non-damaged homologues. The first protein complex detected at DSBs is the MRE11, RAD50 and NBS1 (MRN) complex that is believed to beinvolved in keeping the broken DNA ends together (Lisby et al., 2004; Wyman and Kanaar, 2006). The RAD50 homodimer consist of a globular ATPase domain (where two MRE11 proteins and presumably two NBS1 molecules are also located) and two ~50 nm long flexible coiled-coils arms that can interact at their apex (forming an closed ring due to intra-molecular interactions) (de Jager et al., 2001; van Noort et al., 2003). MRN molecules bind preferentially at DNA ends with their globular domain and this binding triggers a conformational change in the coiled-coil arms that prevents interactions between the arms within the same MRN molecule and promotes interactions between different DNA-bound MRN complexes, thereby tethering different DNA molecules together as a molecular glue (Moreno-Herrero et al., 2005). The subsequent step in HR involves creating free 3’-ends on both sides of the break, which would require a 5’-3’ nuclease activity (Hoeijmakers, 2001). Although the MRN complex is indicated in this step, it is unlikely that the exonuclease activity of MRE11 (3’-5’) is responsible, since this would create overhangs with the wrong polarity (Wyman and Kanaar, 2006). The created 3’ overhangs are stabilized by ssDNA-binding protein RPA, which subsequently facilitates (in cooperation with RAD52 and BRCA2) the assembly of a RAD51 nucleoprotein filament on the ssDNA ends. The RAD51 filament then facilitates the exchange between the ssDNA donor sequence and the

12 DNA repair systems in mammals homologous dsDNA target sequence thought an unknown mechanism. Searching for the homologous sequence is possibly mediated by structural maintenance of chromosomes (SMC) proteins (like cohesins) (Cost and Cozzarelli, 2006; Hirano, 2006; Strom and Sjogren, 2005) and also requires RAD54, which is a SWI/SNF ATP-dependent chromatin remodeling factor (Heyer et al., 2006). The DNA structure that is formed after strand invasion and joint molecule formation has crossed DNA strands (a holliday junction) that need be resolved before the DNA molecules can be separated. Structure-specific nucleases (or resolvases) are required to catalyze the cleavage of the crossed DNA strands. In mammalian cells, the XRCC3 and RAD51C resolvases have been implicated in HR-directed repair and mutations in RAD51C lead to a loss of resolvase activity in human cells (Liu et al., 2004; Liu et al., 2007). However, crossed DNA strands might also be resolved topologically by the combined action of a helicase (from the RecG family, like the WRN or BLM helicases) and a topoisomerase (Heyer et al., 2003; Wyman and Kanaar, 2006). The importance of HR-directed DSB repair is highlighted by several highly cancer-prone human disorders such as ataxia telangiectasia (AT; caused by defects in the ATM and ATR kinases), Nijmegen breakage syndrome (caused by defects in NBS1) and predisposition to breast cancer by mutations in the BRCA genes. In addition, mutation in genes encoding helicases of the RecG family causes the cancer-prone disorders Werner syndrome (defective WRN helicase) and Bloom syndrome (defective BLM helicase) (Hoeijmakers, 2001). Inactivation of HR proteins (e.g. RAD51, RAD50, BRCA1, and BRCA2) in mice results in gross chromosomal rearrangements and is often lethal during early embryogenesis (Brugmans et al., 2007; van Gent et al., 2001).

Non-homologous end joining The dominant pathway to remove DSBs from mammalian cells is the error-prone non- homologous end-joining (NHEJ) pathway. During NHEJ, broken chromosomes ends are simply re-joined by ligation, which does not require the presence of a sister chromatid and NHEJ can therefore operate irrespective of the cell cycle phase (see Fig 4 for a model). NEHJ plays an important role during DNA rearrangements in the immune system that depend on the targeted formation of DSBs, such as rejoining of V (variable), D (diversity) and J (joining) segments to produce functional immuno- globulin (Ig) genes in B lymphocytes. Recognition of DSBs by NHEJ is initiated by the ring-shaped heterodimeric KU protein (consisting of KU70 and KU80) that has high affinity for DNA ends. The binding of KU to DNA ends is highly dynamic and does not require any other NHEJ protein (Mari et al., 2006). KU subsequently recruits the catalytic subunit of the DNA-dependent protein kinase (DNA-PKcs), which phosphorylates several target proteins including p53, KU and DNA-PKcs itself (Mari et al., 2006). Although the relevance of phosphorylation of KU and p53 are currently not understood, autophosphorylation of DNA-PKcs is essential for functional NHEJ and the phosporylation status of DNA-PKcs regulates the stability of its binding to

13 Chapter 1

Fig 3. Model for the mechanism of double-strand break repair by homologous recombination. Adapted from (Hoeijmakers, 2001; Wyman and Kanaar, 2006)

DNA ends (Uematsu et al., 2007). The KU and DNA-PKcs complex bound to DNA termini can juxtapose the two DNA ends in a synaptic complex, ensuring that the DNA ends are in close proximity. There is some evidence that the MRE11, RAD50 and NBS1 (MRN) complex is involved in NHEJ although its exact role is remains unclear. Processing of the DNA ends might be required before joining and the Fig 4. Model for the mechanism of double-strand 5’-3’ exonuclease Artmemis has break repair by non-homologous end-joining. been implicated in this step. Loss of Adapted from (Wyman and Kanaar, 2006) functional Artemis can lead to the severe combined immunodeficiency syndrome (SCID) in humans and inactivation of Artemis in mice results in the complete absence of T- and B cells (Moshous et al., 2001; Rooney et al., 2002). Finally, ligation of the DNA ends is catalyzed by the XRCC4-lig IV complex, which is stabilized by the Cernunnos protein (also called XLF) (Ahnesorg et al., 2006; Buck et al., 2006). The recruitment of XRCC4-lig IV to DSBs is mediated by interactions between KU and XRCC4 and does not require DNA-PKcs in vivo. However, efficient ligation of the DNA ends is blocked in the absence of DNA-PKcs autophosporylation, possibly

14 DNA repair systems in mammals because unphosphorylated DNA-PKcs has a more stable interaction with DNA ends (Uematsu et al., 2007). Inactivation of NHEJ factors in mice results in a severe sensitivity to ionizing radiation and a defective immune system caused by the inability to carry out V(D)J recombination. Moreover, deletion of XRCC4 or Lig IV is lethal during early embryogenesis.

Contributions of HR and NHEJ to DSB repair The two major repair pathways that deal with DSBs (HR and NHEJ) are highly conserved between bacteria, yeast and mammals. However, the dominant pathway to repair DSBs in bacteria and yeast (S. cerevisiae) is HR-directed repair whereas NHEJ contributes more significantly to DSB removal in mammals. The difference in DNA organization between mammals and yeast may account for the difference in relative contributions of HR and NHEJ to the repair of DSBs. The human genome consists of 3.109 base-pairs that are organized as chromosomes that each occupies a distinct space in the cell nucleus called chromosome territories (Bolzer et al., 2005; Cremer and Cremer, 2001). The homologous chromosomes in mammals are not often is close proximity to each other and the folding of the genome into a higher-order chromatin structure makes the search for the homologous sequence extremely difficult. This is why NHEJ is solely responsible for the removal of DSB if cells are in G1. If DSBs arise during replication, the cell can use HR since the sister-chromatids are in close proximity to each other. However, after replication the sister-chromatids are packaged into chromatin and separated, making the search for homologous sequences and thus HR-mediated repair difficult again. In fact, NHEJ accounts for the repair of ~60% of the exogenously induced DSBs in mouse cells (Liang et al., 1998). Conversely, HR- directed repair is operational in yeast cells even during G1 and the contribution of NHEJ to DSB removal is only minor. In yeast, the search for homologous sequences is apparently more efficient than in mammals. Chicken cells lacking functional RAD54 (HR) and KU70 (NHEJ) proteins are more sensitive to ionizing radiation than single RAD54 or KU70 mutant cells, illustrating that both pathways contribute to DSB removal in mammals (Sonoda et al., 2006). Genetic evidence suggests that NHEJ is slightly suppressed during late S and G2 in a PARP-dependent manner, presumably to favor error-free HR-mediated repair of DSBs. It is, however, not exactly clear how the choice between HR and NHEJ is regulated in mammalian cells.

Fanconi Anemia repair Certain chemicals compounds can link two complementary DNA strands resulting in an interstrand cross-link (ICL). If left unrepaired a single ICL is lethal because this type of lesion inhibits strand separation and thereby efficiently blocks transcription and replication (Magana-Schwencke et al., 1982). Examples of chemicals that produce ICLs are cisplatin and mitomycin C that are successfully used as chemotherapeutical agents. In addition, by-products of lipid peroxidation (e.g. malondialdehyde) also

15 Chapter 1 contribute to the endogenous production of ICLs (Niedernhofer et al., 2003). Mammalian cells have developed a sophisticated mechanism to deal with ICLs known as the Fanconi anemia pathway (FA). Individuals suffering from the X-linked hereditary disorder Fanconi anemia (FA) are characterized by progressive bone marrow failure and cancer susceptibility (often acute leukemia) (Kennedy and D'Andrea, 2005). Cells derived from FA patients display chromosomal instability and extreme sensitivity to crosslinking agents. Twelve FA complementation groups have been identified to date (FANCA, B, C, D1, D2, E, F, G, I, J, L, M) and the FANC gene products operate in a common biochemical pathway that is not yet understood. About ~60% of the FA patients belong to complementation group A. It is currently believed that the FA pathway is directly involved in the repair and bypass of ICLs (Niedernhofer et al., 2005). Interestingly, the FANCD1 gene product is identical to the BRCA2 protein, which is involved in homologous recombination (HR) (Howlett et al., 2002). In addition, defects in the nucleotide excision repair endonuclease ERCC1-XPF causes extreme sensitivity to crosslinking agents suggesting a role in ICL repair (Larminat and Bohr, 1994; Li et al., 1999). A speculative model for FA-mediated repair during replication was recently suggested and comprises the following steps (also see Fig 5) (Niedernhofer et al., 2005): i) recognition of the ICL, which is likely to be mediated by stalling of pol δ/ε at lesions during replicating, ii) incision on one side of the ICL resulting in a DSB, iii) unwinding around the incised ICL by a helicase, iv) second

Fig 5. Model for the mechanism of interstrand cross-link repair by the fanconi anemia pathway. Adapted from (Niedernhofer et al., 2005)

16 DNA repair systems in mammals incision (called unhooking) at the other side of the ICL in the same strand resulting in a gap that in filled in by TLS polymerases (since the opposite strand still contains the excised ICL) v) removal of the residual damage in the opposite strand by NER (or spontaneous hydrolysis followed by BER) restoring the integrity of the DNA template, vi) HR-mediated repair of the DSB that was created by the first incision using the intact template followed by reinitiating of replication. Although FA repair protects against cytotoxicity of ICLs, it is very error-prone and highly mutagenic. It is likely that FA repair is initiated by phosphorylation of FANCD2 by the ATR kinase, which is activated by replication blocks (Andreassen et al., 2004). In addition, FANCD2 is mono-ubiquitylated in S-phase by the E3 ligase FANCL (an event that also requires the FANCI gene product), which triggers its association with chromatin (Niedernhofer et al., 2005). This leads to the recruitment of the FA core complex containing almost all FANC proteins, including the FANCJ and FANCM helicases that are thought to unwind the DNA around the ICL. The endonucleases Mus81-Eme1 and ERCC1-XPF have been implicated in the multi-strand incisions needed to release the ICL from one DNA strand (which also requires RPA) and both nucleases were shown to be required to convert ICLs into DSBs allowing repair by homologous recombination (Hanada et al., 2007; Hanada et al., 2006; Niedernhofer et al., 2004). Finally, FANCD2 interacts with HR proteins including BRCA1, BRCA2/FANCD1 and RAD51 that mediate sister- chromatid exchange (Taniguchi et al., 2002). The FA core complex also interacts with the BLM helicase that is needed for the resolution of recombination intermediates such as holliday junctions (Meetei et al., 2003). The FA pathway is switched off during late S phase by the ubiquitin-specific protease 1 (USP1), which deubiquitylates FANCD2 (Nijman et al., 2005). It is expected that many more proteins are involved in FA repair, including nucleases, TLS polymerases and chromatin remodeling complexes. Although we are beginning to understand how FA contributes to the removal and bypass of ICLs during replication, it is not clear how the transcription machinery deals with ICLs that block elongation.

17 18 Chapter 2 Nucleotide Excision Repair in Mammals

Martijn S. Luijsterburg and Roel van Driel

19 Chapter 2

Introduction Nucleotide excision repair (NER) is a highly versatile DNA repair pathway of which the molecular mechanism has been conserved from bacteria to man. NER removes a variety of structurally unrelated lesions from the genome such as the UV-induced pyrimidine-pyrimidone (6-4) photoproducts (6-4 PP) and cyclobutane pyrimidine dimers (CPD; see Fig 1). Placental mammals fully rely on NER for the removal of UV-induced damages from their genome. In addition, NER removes bulky adducts formed by polycyclic aromatic hydrocarbons (such as benzopyrene that is present in cigarette smoke and car exhaust fumes), intrastrand cisplatin adducts and small (~10 kDa) protein-DNA cross-links (Nakano et al., 2007). An important molecular determinant for efficient lesion recognition by NER is the degree of double helix distortion inflicted by the lesion. For instance, (6-4) PPs are more helix-distorting than CPD and as a result the repair rate of 6-4 PP is ~5 times higher than that of CPDs (Bohr et al., 1985; Kim and Choi, 1995; Mitchell and Nairn, 1989). A hallmark of NER is the removal of lesions as part of a single-stranded DNA fragment that is usually ~30 nucleotides in size. Mammalian NER involves ~30 gene products and comprises the following steps (see Fig 2): i) damage recognition, ii) DNA unwinding and lesion demarcation, iii) dual incision around the lesion, iv) release of a single-stranded DNA fragment containing the DNA injury and v) repair synthesis and ligation of the nick (for a more detailed model see Fig 3) (Aboussekhra et al., 1995; de Laat et al., 1999).

Two mechanistically distinct repair pathways and associated syndromes Two mechanistically distinct NER pathways exist: a transcription-coupled repair pathway (TCR) that removes lesions exclusively from actively transcribed genes and a global genome repair pathway (GGR), which removes lesions all throughout the genome (Mellon et al., 1987). Three human disorders are connected to inherited defects in NER: Xeroderma pigmentosum (XP), Cockayne syndrome (CS) and trichothiodystrophy (TTD) (de Boer and Hoeijmakers, 2000; Lehmann, 2003). XP is the classic NER-deficiency syndrome and XP patients suffer from extreme photosensitivity and high susceptibility to skin-cancer. The hallmarks of XP are a dry parchment skin and pigmentation abnormalities in sun-exposed areas (de Boer and Hoeijmakers, 2000). Nine XP complementation groups have been identified (XP-A, B, C, D, E, F, G) and defects in the TLS polymerase η lead to XP-V (XP variant) (Flejter et al., 1992; Keeney et al., 1994; Legerski and Peterson, 1992; Masutani et al., 1999; Scherly et al., 1993; Sijbers et al., 1996a; Tanaka et al., 1990; Weeda et al., 1990). Moreover, the first human patient with an inherited defect in the repair protein ERCC1 (which was the first human repair gene to be cloned) was recently described representing a novel XP complementation group (Jaspers et al., 2007; Westerveld et al., 1984). The products of all the XP genes are involved in both NER sub-pathways except XPC and XPE, which are only required for GGR (van Hoffen et al., 1995). Two CS complementation groups have been identified (CS-A, CS-B) and cells from both groups are defective in

20 NER in mammals

TCR but not GGR, illustrating that the CS proteins are dedicated to TCR (Henning et al., 1995; Troelstra et al., 1992). Patients suffering from CS display neurological, developmental and aging-related problems and are photosensitive although not to the same extend as XP patients. Moreover, CS patients do not have an increased risk of developing skin-cancer (de Boer and Hoeijmakers, 2000; Lehmann, 2003). Some patients from the XP-B, XP-D and XP-G complementation groups also have features of CS (Broughton et al., 1995; Weeda et al., 1990). Defects in TCR alone do not Fig 1. UV-induced photoproducts. The upper image explain the severity of the CS shows a CPD formed by double bonds (between the phenotype and it is the current view 5-5 and 6-6 carbon atoms) between two adjacent that besides a NER-deficiency thymidines. The lower image shows a 6-4 PP formed by the linkage of the 6-4 carbon atoms between a disorder, CS is also a transcription thymidine and cytosine disorder. This explains why combined XP/CS is only found in XP-B, XP-D and XP-G patients since besides NER, the corresponding gene products are also involved in transcription (Drapkin et al., 1994). XPB and XPD are the helicase subunits of general transcription factor IIH (TFIIH) and mutations that lead to combined XP/CS were shown to impair the transcription function of TFIIH (Coin et al., 1999). In addition, XPG was shown to stabilize TFIIH, which is required to activate transcription (Ito et al., 2007). One TTD complementation group has been identified (TTD-A) and the TTD gene product was shown to be the tenth subunit of the TFIIH protein

Fig 2. Schematic overview complex that also contains the XPB of the mechanism of NER. and XPD subunits (Giglia-Mari et al., 2004). In agreement, some XP-B and

21 Chapter 2

XP-D patients have features of TTD (Coin et al., 1998; Weeda et al., 1997b). Patients suffering from TTD display brittle hair, unusual facial features and show severe clinician manifestation reminiscent of CS. Although the TFIIH complex from TTD cells is fully functional, the cellular levels of TFIIH are extremely low implying TTDA in maintaining normal levels of TFIIH (Giglia-Mari et al., 2004; Vermeulen et al., 2000).

Molecular mechanism of NER After the identification of the different XP complementation groups, NER was successfully reconstituted in vitro using the following proteins: XPC, TFIIH, XPA, XPG, RPA, ERCC1 and XPF, which are sufficient to perform dual incision. In addition, the in vitro reaction required RF-C, PCNA, pol ε and DNA lig 1 for repair synthesis (Aboussekhra et al., 1995). Although this represents the minimal number of proteins to perform NER on naked DNA in vitro, additional factors are required for in vivo NER on chromatinized DNA templates. The main characteristics of the mammalian NER factors will be summarized in this section. Special attention will be paid to our understanding of NER in vivo and recent insights into the dynamic properties of core repair factors will be integrated into a model for the NER reaction in the living cell (Essers et al., 2005; Hoogstraten et al., 2002; Houtsmuller et al., 1999; Luijsterburg et al., 2007; Moné et al., 2004; Rademakers et al., 2003; van den Boom et al., 2004; Volker et al., 2001; Zotter et al., 2006). Finally, we will consider which chromatin remodelling events might be required to facilitate NER in living cells.

Damage recognition in GGR: XPC and DDB2 Global genome repair (GGR) in mammalian cells is initiated by XPC, which forms a complex with HR23B (the human homologue of RAD23) and CEN2 (Araki et al., 2001; Masutani et al., 1994; Sugasawa et al., 1998; Volker et al., 2001). Recruitment of NER proteins TFIIH, XPA, XPG and ERCC1-XPF to sites of UV-induced DNA lesions is abolished in XP-C deficient cells, demonstrating that in vivo GGR complex assembly is strictly XPC-dependent (Volker et al., 2001). Rather than binding to lesions directly, XPC binds to the accessible non-damaged DNA opposite to a DNA injury, which allows the recognition of a broad spectrum of lesions that are structurally unrelated (see Fig 3) (Sugasawa and Hanaoka, 2007). Recently, it was demonstrated that two aromatic residues in the evolutionary conserved part of the XPC polypeptide (Trp 690 and Phe 733) mediate an aversion to associate with damaged nucleotides and a preference to bind single-stranded DNA regions, driving preferential recruitment of XPC to DNA lesions (Buterin et al., 2005; Maillard et al., 2007). Interestingly, similar ssDNA-binding motifs are found in RPAand BRCA2/FANCD1 (Maillard et al., 2007). The crystal-structure of the yeast XPC homologue RAD4 bound to DNA containing a single CPD indeed confirms that damage recognition is mediated by binding to non- damaged DNA opposite to a lesion (Min and Pavletich, 2007). The role of HR23B

22 NER in mammals appears to be stabilization of the XPC protein and loss of both HR23A and HR23B results in a decrease in cellular XPC levels (Ng et al., 2003). Live cell FRET measurements suggest that HR23B dissociates when XPC binds to damaged DNA, indicating that HR23B is not directly involved in repair (S. Bergink, unpublished). The precise role of CEN2 during NER is still unclear but it appears that CEN2 considerably stimulates XPC (Nishi et al., 2005). Mobility studies on GFP-tagged XPC suggest that the majority of XPC molecules (>90%) transiently interact non-specifically with genomic DNA (Hoogstraten, 2003; Politi et al., 2005). It is unclear whether this low affinity binding to genomic DNA contributes to the detection of lesion in the genome. Although XPC has a high affinity for 6-4 PPs, its binding to CPDs is rather inefficient and efficient recognition and repair of CPDs requires the damaged-DNA binding protein 2 (DDB2) (Tang et al., 2000). Moreover, DDB2 significantly stimulates the repair of 6-4 PPs particularly at low UV doses (Hwang et al., 1999; Moser et al., 2005; Tang et al., 2000). Mutations in DDB2 cause the XP-E phenotype and cells from these patients (as well as rodent cells that also lack functional DDB2) are deficient in CPD repair (Nichols et al., 2000; Rapic-Otrin et al., 2003). DDB2 contains WD40 repeats and is incorporated into a functional CUL4A-based E3 ubiquitin ligase through its interaction with DDB1 (Groisman et al., 2003; He et al., 2006). Live cell imaging revealed that DDB2, CUL4A and DDB1 are rapidly recruited to UV-lesions with similar association kinetics (Chapter 4) (Luijsterburg et al., 2007). Moreover, DDB2 was shown to associate with and dissociate from UV-induced lesions independently of XPC and it was demonstrated that binding to lesions depends on the Arg 273 residue, which is located within theWD40 domain (Chapter 4) (Luijsterburg et al., 2007). The E3 ubiquitin ligase activity of the DDB2 complex is transiently activated by UV irradiation and is specifically directed to chromatin at damaged sites (Groisman et al., 2003). Several substrates for ubiquitylation were identified, including XPC, DDB2 itself and histones H2A, H3 and H4 (Groisman et al., 2003; Kapetanaki et al., 2006; Sugasawa et al., 2005; Wang et al., 2006a). The ubiquitylation of XPC increases its affinity for DNA (both damaged and non-damaged) whereas ubiquitylated DDB2 is targeted for degradation (Rapic-Otrin et al., 2002; Sugasawa et al., 2005). Besides ubiquitylation, XPC was recently demonstrated to be modified by SUMO in an XPA- dependent manner, which was suggested to stabilize XPC (Wang et al., 2005). The ubiquitylation of H3 and H4 by the DDB2 complex was shown to weaken the interaction between histones and DNA thereby facilitating access to repair proteins in chromatin (Wang et al., 2006a). Indeed, knock-down of DDB2, CUL4A or DDB1 interferes with lesion recognition by XPC in vivo (El-Mahdy et al., 2006; Li et al., 2006). Ubiquitylation of H2A by the DDB2 complex is still under debate since different laboratories have reported conflicting results (Bergink et al., 2006; Kapetanaki et al., 2006; Wang et al., 2006a). The precise interaction between XPC and DDB2 at DNA lesions is still unclear. Live cell imaging suggests that the bulk of DDB2 interacts with lesions independently of XPC and that there is little interaction between the two

23 Chapter 2 recognition proteins in vivo (Chapter 4) (Luijsterburg et al., 2007). On the other hand, XPC and DDB2 were shown to interact in solution and the DDB2-mediated ubiquitylation of XPC suggests a direct although most likely transient interaction in vivo (Sugasawa, 2006; Sugasawa et al., 2005). Both possibilities are not mutually exclusive and eventually it is a matter of quantity. The cellular levels and the fraction of DDB2 molecules that binds to UV-lesions are significantly higher than those of XPC, suggesting that many more DDB2 molecules are bound to chromatin than XPC molecules after UV irradiation (Araujo et al., 2001; Hoogstraten, 2003; Keeney et al., 1993; Luijsterburg et al., 2007). However, this does not exclude the possibility that both damage sensors directly interact at some possibly poorly recognized lesions. A recent study revealed that XPC in which Trp 690 was substituted for Ser failed to accumulate at sites of UV-lesions in similar amounts as wild-type XPC (Yasuda et al., 2007). Surprisingly, a small fraction of this mutant XPC polypeptide did accumulate upon UV irradiation in a DDB2-dependent manner (Yasuda et al., 2007). Conversely, live-cell imaging of GFP-tagged XPC (with a Trp 690 to Ser mutation) did not reveal any immobilization of the mutated XPC protein upon UV irradiation (Hoogstraten et al. Submitted). Thus, the precise interplay between these two damage-recognition proteins and the mechanism underlying lesion detection in chromatin remains to be elucidated.

Damage recognition in TCR: RNAPII, CSA and CSB The two NER pathways GGR and TCR differ in the way lesions are detected providing an explanation why XPC is dispensable for TCR. However, it is believed that TCR and GGR funnel into a common molecular mechanism that involves the core NER proteins TFIIH, RPA, XPG, XPA and ERCC1-XPF once lesions have been detected. The contribution of TCR to the total cellular NER activity following UV irradiation is ~1%, which explains why TCR-specific accumulation of NER proteins (for example XPA binding in XP-C cells) can not be detected in intact cells but requires more sensitive techniques such as chromatin immunoprecipitation (Fousteri et al., 2006; Rademakers et al., 2003; Volker et al., 2001). During transcription elongation, RNA polymerase II is stalled at a variety of lesions that efficiently block translocation along the DNA (Brueckner and Cramer, 2007; Brueckner et al., 2007; Cline et al., 2004). The arrest of RNAPII at lesions is the molecular basis for damage recognition during TCR (Laine and Egly, 2006). The CSA and CSB proteins are TCR-dedicated repair factors and the interplay between these factors was recently elucidated (Fousteri et al., 2006; Groisman et al., 2006). CSB is an ATP-dependent chromatin remodelling factor of the SWI2/SNF2 family that interacts with histones and remodels nucleosomes (Citterio et al., 2000). Recruitment of CSB to lesions is mediated by a direct interaction with lesion-stalled hyperphosporylated RNAPII (RNAPIIo), which is not displaced during TCR complex assembly (Fousteri et al., 2006; van den Boom et al., 2004). Upon binding to stalled RNAPIIo, CSB functions as a coupling-factor that mediates the

24 NER in mammals recruitment of subsequent repair factors TFIIH, XPG, RPAand ERCC1-XPF (Fousteri et al., 2006). Live cell imaging revealed that GFP-tagged CSB transiently interacts with the transcription machinery and that the presence of DNA damage stabilizes this interaction suggesting that CSB monitors the progression of transcription elongation by continuously probing elongation complexes (van den Boom et al., 2004). The CSA protein contains WD40 repeats and is part of a CUL4A-based E3 ubiquitin ligase (through interaction with DDB1) that has a similar complex composition as the DDB2 containing E3 ligase (Groisman et al., 2003; He et al., 2006). Recruitment of CSA is CSB-dependent and results in the ubiquitylation and degradation of CSB by the proteosome system (Fousteri et al., 2006; Groisman et al., 2006). The break-down of CSB is a rather late event (~3 hrs) during TCR and was shown to be important for the recovery of transcription following UV irradiation. Although CSA is dispensable for the recruitment of pre-incision proteins such as the nucleases XPG and ERCC1-XPF, it is required to recruit repair synthesis factors like PCNA, RFC and pol δ (M. Fousteri, unpublished). This raises the question whether dual incision takes place in CS-A cells. The interaction between the two CS proteins is generally thought to be very transient and GFP-tagged CSA protein is not detected at sites of UV-induced DNA damage, whereas GFP-CSB does accumulate after UV irradiation (V. van den Boom, unpublished) (Licht et al., 2003; van den Boom et al., 2004). The rate at which TCR removes lesions from actively transcribed genes (e.g. the DHFR gene) is generally higher than that at which GGR performs repair and TCR is therefore considered to be faster than GGR (Bohr et al., 1985; Mellon et al., 1987). At the cellular level, TCR appears to be slower than the bulk of GGR since participation of GFP-CSB can be detected up to 16 hrs after UV irradiation, which is considerably longer than the time core NER factors such as XPC and XPA are immobilized following irradiation (Chapter 6). The explanation for this paradox is that TCR removes 6-4 PPs and CPDs with equal efficiencies from transcribed genes whereas GGR mainly removes 6-4 PPs from the genome since CPDs are poorly recognized (van Hoffen et al., 1995). Thus, the long (i.e. up to 16 hrs) immobilization of CSB likely reflects the assembly of TCR complexes at CPDs in genes that are not frequently transcribed.

NER helicase: TFIIH Upon recognition of lesions by GGR and TCR, the NER reaction funnels into a common molecular mechanism that involves the recruitment of TFIIH. In GGR, XPC recruits TFIIH via direct protein-protein interactions, whereas CSB mediates recruitment of TFIIH to TCR complexes (Araujo et al., 2001; Fousteri et al., 2006; Yokoi et al., 2000). The ten-subunit TFIIH complex, which is composed of a seven-subunit core (XPB/p89, XPD/p80, TTD/p8, p34, p44, p52, p62) and a three-subunit kinase complex (Cdk7, cyclin H and MAT1) is ~700 kDa in size and plays key roles in RNAPI and RNAPII transcription as well as in NER (Drapkin et al., 1994; Giglia-Mari et al., 2004; Iben et al., 2002). The cyclin-activating kinase (CAK) complex of TFIIH is dispensable

25 Chapter 2 for in vitro NER but the CAK subunits are recruited to UV-induced lesions in vivo, suggesting a possible role in NER (Hoogstraten, 2003). The XPB subunit of TFIIH is an ATP-dependent helicase that mediates unwinding of promoter DNA in a 3’-5’ orientation during transcription initiation (Douziech et al., 2000). Only six families suffering from XP-B have been described world-wide of which four families display the XP/CS phenotype and the remaining two either XP or XP/TTD (Oh et al., 2006; Vermeulen et al., 1994; Weeda et al., 1997b). These families show only a limited number of mutations in XPB probably reflecting the limited range of alteration in this vital protein that is compatible with life. The XPD subunit of TFIIH is a 5’-3’ ATP- dependent helicase that is required for strand separation during NER, which requires the ATPase but not the helicase activity of XPB (Coin et al., 2007; Winkler et al., 2000). The ATPase activity of XPB is stimulated by p52, whereas the helicase activity of XPD is activated by p44, illustrating that these subunits are not merely structural components of TFIIH (Coin et al., 2007). Patients with XP-D are more frequent than XP-B patients and display clinical characteristics varying from classic XP to XP/CS and XP/TTD. It is generally thought that the XP phenotype in XP-B and XP-D patients is the result of defective NER, whereas the CS/TTD characteristics are caused by transcription defects due to malfunctioning TFIIH (Coin et al., 1999; Dubaele et al., 2003). Open complex formation during NER critically depends on functional TFIIH and it appears that a two-step mechanism is employed in which TFIIH mediates the initial opening (8-10 nucleotides) after which RPA, XPA and XPG bind to obtain full opening of ~30 nucleotides around the lesion (see Fig 3) (Evans et al., 1997a; Evans et al., 1997b; Mu et al., 1997). In the presence of lesions, XPAspecifically stimulates the ATPase activity of TFIIH whilst RPA and XPG stabilize the partially unwound repair intermediate thereby contributing to full opening around the lesion (Evans et al., 1997b; Winkler et al., 2001). Live cell imaging of GFP-tagged XPB, which is incorporated into functional TFIIH complexes revealed that TFIIH dynamically shuttles between RNAPI and RNAPII transcription and NER (Hoogstraten et al., 2002). Mobility measurements showed that TFIIH is transiently involved in RNAPI (2-10s) and RNAPII (~25s) transcription and shuttles between transcription and NER where it is immobilized for maximally ~4 min (Hoogstraten et al., 2002). The involvement of TFIIH in NER did not affect its engagement in transcription showing that NER does not affect transcription in trans (Hoogstraten et al., 2002; Moné et al., 2001). Interestingly, analysis of GFP-tagged TTDA in living cells revealed a dynamic interaction between TFIIH and TTDA, which is stabilized by the presence of bona fide NER lesions (Giglia-Mari et al., 2006). Introduction of lesions that trigger an abortive NER response (such as photo-activation of intercalated actinomycin D) did cause accumulation of XPC and TFIIH but failed to stabilize the interaction between TFIIH and TTDA, possibly reflecting a kinetic proofreading mechanism to gain higher substrate specificity during NER (Giglia-Mari et al., 2006). Both biochemical and in vivo data reveal that TTDA is a NER-dedicated subunit of TFIIH that stimulates the ATPase

26 NER in mammals activity of XPB, which is essential for open complex formation. The stimulation of the ATPase activity of XPB by TTDA is probably mediated through the p52 subunit of TFIIH that interacts with both TTDA and XPB (Coin et al., 2007). In addition, TTDA is essential to maintain the cellular concentration of TFIIH suggesting that this small subunit (8kDa) is required for stabilization of the ~700 kDa TFIIH complex (Coin et al., 2006; Vermeulen et al., 2000).

RPA and XPA: a dual probe sensor to detect structural DNA alterations Replication protein A (RPA) is an abundant single-stranded DNA-binding protein that plays key roles in replication, recombination and several DNA repair pathways in mammalian cells (Fanning et al., 2006; Wold, 1997). RPA consist of three subunits (RPA70, 32 and 14) and its affinity for ssDNA is ~3 orders of magnitude higher than the affinity for dsDNA (Kim et al., 1992). RPAcontains four ssDNA-binding domains, three of which are present in RPA70 (tethered to each other through flexible linkers) and the fourth is found in RPA32 (Fanning et al., 2006). Two modes of binding have been identified; RPA associates with a minimal region of ~10 nucleotides and then elongates to cover an optimal region of ~30 nucleotides (Blackwell and Borowiec, 1994; Kim et al., 1992). The latter mode of binding is ~100-fold more stable and the two-step binding of RPA probably underlies its capacity to unwind DNA (Blackwell and Borowiec, 1994). The tethering of several weak ssDNA-bindings domains results in cooperative association of RPA to ssDNA and increases its overall affinity for ssDNA with ~2 orders of magnitude (Arunkumar et al., 2003). During NER, RPA associates with the undamaged DNA strand and is indispensable for open complex formation, suggesting that RPA binds to a partially unwound structure that is opened by TFIIH (~8-10 nucleotides) after which it elongates to bind to a ~30 nucleotideregion acting as a wedge to separate the DNA strands around a lesion (de Laat et al., 1998b; Evans et al., 1997b; Hermanson-Miller and Turchi, 2002; Mu et al., 1997). RPA interacts with several core NER proteins, including XPA, XPG and ERCC1-XPF and the incision activity of both NER endonucleases is significantly stimulates by RPA(de Laat et al., 1998b; He et al., 1995; Matsunaga et al., 1996). In vitro experiment revealed that RPA binds to damaged DNA with a defined polarity; initial RPA binding to 8-10 nucleotides occurs at the 5’ side of the lesion after which RPAstretches along the DNA in the 3’ direction (de Laat et al., 1998b; Kolpashchikov et al., 2001). This defined DNA-binding polarity of RPA correctly positions the NER enodnuclease XPG and ERCC1-XPF on the DNA. The 3’-oriented side of RPA specifically interacts with and stimulates the 5’ incision by endonuclease ERCC1-XPF at the ssDNA-dsDNA junction without interacting with XPG. Conversely, the 5’-oriented side of RPAmediates stable binding of 3’ endonuclease XPG but inhibits ERCC1-XPF incisions (see Fig 3) (de Laat et al., 1998b; He et al., 1995; Matsunaga et al., 1996). However, binding of XPG to RPA is not sufficient to stimulate the 3’ incision (de Laat et al., 1998b). It was previously suggested that RPA and XPA form a stable complex in cell nuclei, but

27 Chapter 2 measurements on GFP-tagged XPA protein argue against this notion (He et al., 1995; Matsunaga et al., 1996; Rademakers et al., 2003). Moreover, the binding of RPA to UV lesion in intact cells was shown to be independent of XPA, suggesting that RPA- XPAinter-actions only occur on damaged DNA (Rademakers et al., 2003). In contrast, recent ChIP experiments revealed co-precipitation of RPA with XPA in non-damaged cells (Moser et al., 2007), suggesting that XPA and RPA transiently interact in cell nuclei. Besides the pre-incision stage of NER, RPAalso plays a crucial role in the DNA repair synthesis step that requires RF-C, PCNA and DNA pol ε in vitro (Shivji et al., 1995). Analysis of the assembly and disassembly of repair proteins on immobilized damaged-DNA templates in vitro revealed that RPA remains bound after dual incision and initiates the assembly of DNA synthesis factors such as PCNA (Riedl et al., 2003). This notion is supported by recent in vivo experiments demonstrating that RPA redistribution upon UV irradiation is significantly slowed down in the presence of DNA synthesis inhibitors reflecting a role in DNA repair synthesis (J. Moser, unpublished). Cells expressing GFP-tagged RPA34 have been established and it would be of particular interest to study the kinetics of RPA involvement in NER (Solomon et al., 2004). XPAis a small (36 kDa) DNA-binding zinc metalloprotein that has a central role in NER complex assembly and is essential for GGR and TCR. Although XPA originally was proposed to be the initial damage recognition protein, it is now well established that GGR is initiated by XPC (Sugasawa et al., 1998; Volker et al., 2001; Wakasugi and Sancar, 1999). In agreement, live-cell imaging revealed that recruitment of GFP-tagged XPA is strictly XPC-dependent (Rademakers et al., 2003). Evidence for dimerization of XPAhas been presented but it is at present unclear if XPAassociates with repair complexes as a monomer or a dimer (Yang et al., 2002). The N-terminus of XPA (residues 1-97) binds to RPA34 and ERCC1, whereas the C-terminus interacts with TFIIH (residues 226-273) (Li et al., 1995a; Park et al., 1995). The central XPA domain contains the minimal DNA binding domain (residues 98-219) and interacts with RPA70 (Ikegami et al., 1998; Kuraoka et al., 1996; Li et al., 1995a). These many interactions with other core NER proteins suggest a central role for XPA in NER complex assembly. Structural analysis revealed that XPA binds preferentially to bent DNA duplexes, which is characteristic of an architectural protein and not consistent with a direct participation in damage recognition (Camenisch et al., 2006; Camenisch et al., 2007). The DNA binding domain of XPA consist of a concave surface that contains seven positively charged residues (mainly lysines) that interact with the negatively charged DNA backbone by electrostatic interactions (Buchko et al., 1998). Sharply bent DNA has an increased negative electrostatic potential caused by the shortened distance between adjacent phosphate residues, which drives association of XPA to kinked DNA duplexes (Camenisch et al., 2007). Interestingly, both XPC and DDB2 were show to introduce kinks in the DNA upon binding thereby providing a bent substrate for XPA (Fujiwara et al., 1999; Janicijevic et al., 2003). The binding of XPA to damaged DNA is greatly enhanced by RPA and the interaction between RPA,

28 NER in mammals which is bound to the undamaged strand and XPAthat associates with the kinked DNA duplex allows the simultaneous detection of base-pair and backbone distortions contributing to the lesion specificity of NER (Missura et al., 2001; Wang et al., 2000). In this way, the XPA-RPA complex forms a composite sensor that can detect the two major structural alterations of DNA lesions (i.e. partial unwinding and DNA bending) at the same time reflecting a damage verification step before dual incision. In addition, XPA was shown to inhibit DNA unwinding by RPA possibly to avoid excessive opening of the DNA around the lesion (Missura et al., 2001; Patrick and Turchi, 2002). Cells deficient in XPA are completely devoid of incision activity, suggesting that XPA plays an important role in incision coordination. The recruitment of 5’ endonuclease ERCC1-XPF to NER complexes is mediated by a small stretch of consecutive glycine residues in XPA that bind a V-shaped hydrophobic groove in the central domain of ERCC1, providing an explanation for the lack of 5’ incisions in XP-A cells (Li et al., 1995b; Tsodikov et al., 2007; Volker et al., 2001). Although XPG recruitment was suggested to depend on XPAin vitro, XPG is recruited to UV lesions in XPA-deficient cells demonstrating that XPA is not required for XPG association in vivo (Riedl et al., 2003; Volker et al., 2001). However, incision by XPG depends on XPA since XP-A cells are devoid of 3’ incisions (Evans et al., 1997a). Activation of dual incision by XPA is likely to be mediated through the XPA-dependent and lesion-specific stimulation of ATP hydrolysis by the TFIIH helicase subunits, which is essential for dual incision (Mu et al., 1996; Winkler et al., 2001).

Dual incision: XPG and ERCC1-XPF The XPG protein is a 133 kDa structure-specific endonuclease of the FEN1 family that mediates the 3’ incision during NER (O'Donovan et al., 1994a). XPG specifically incises DNA at the 3’ side of the junction between single-stranded DNA and double- stranded DNA (Hohl et al., 2003; O'Donovan et al., 1994a; O'Donovan et al., 1994b). During NER, XPG cleaves the damage-containing strand about 2-8 nucleotides from the 3’ side of the lesion. Like FEN1, XPG contains two highly conserved N- and I- nuclease motifs (amino acids 1-95 and 766-873, respectively) that juxtapose and form the catalytically active core of the protein (Constantinou et al., 1999). The N- and I- regions of XPG are separated by a region of ~600 amino acids (designated as the spacer region; residues 95 - 766) that does not contain any structural motifs (Scherly et al., 1993). XPG has several interactions with RPA and TFIIH (XPB, XPD, p62 and p44) and the N-terminal part of the spacer region was demonstrated to mediate the recruitment of XPG to repair complexes (Dunand-Sauthier et al., 2005; Iyer et al., 1996; Thorel et al., 2004). In a complementary study, the recruitment of XPG was shown to depend on functional TFIIH, suggesting that TFIIH recruits XPG to NER complexes through its spacer region (Chapter 4) (Zotter et al., 2006). Moreover, the association of XPG-GFP in living cells was ~3-fold slower at 27°C than at 37°C in contrast to the association kinetics of XPC and TFIIH, which were not affected at lower temperature

29 Chapter 2

(Moné et al., 2004; Zotter et al., 2006). Similarly, the recruitment of ERCC1-XPF was also slowed down at lower temperature, suggesting that the association of both endonucleases is influenced by an enzymatic reaction, possibly unwinding of the DNA around a lesion that requires TFIIH (Chapter 4) (Moné et al., 2004; Zotter et al., 2006). Interestingly, several TFIIH mutants that are defective in open complex formation displayed slower binding of ERCC1-XPF and XPG, suggesting that binding of XPG and ERCC1-XPF requires a partially unwound repair intermediate but that recruitment of the endonucleases as such does not depend on open complex formation (Oh et al., 2007; Zotter et al., 2006). Besides its catalytic function, XPG also has a structural role in NER since the 5’ incision requires the presence of XPG even if catalytically inactive (Wakasugi et al., 1997). Some XP-G individuals that produce severely truncated XPG proteins display symptoms of CS, suggesting that XPG has an additional function in transcription (Lalle et al., 2002). In support of this notion, it was recently suggested that XPG and TFIIH form a stable complex in cells and mutations in XPG that abolish the interaction between the two protein complexes were shown to cause dissociation of the CAK complex and XPD from the TFIIH core complex (Araujo et al., 2001; Ito et al., 2007). Loss of CAK from TFIIH subsequently results in defective phosphorylation and transactivation of transcription factors, such as estrogen receptor α (Chen et al., 2000; Ito et al., 2007). Thus, XPG is required for stabilization of TFIIH, providing an explanation for how defects in XPG can cause a transcription syndrome such as Cockayne syndrome (Friedberg and Wood, 2007). In addition, XPG was suggested to bind stalled RNAPII in cooperation with CSB, which was suggested to be important for initiation of TCR (Sarker et al., 2005). Although it is possible that transient interactions with XPG stabilize TFIIH, live cell imaging shows that XPG and TFIIH are distinct complexes in vivo based on the following observations: i) The mobility of TFIIH-GFP is temperature sensitive consistent with a role in transcription, whereas XPG-GFP mobility is temperature insensitive, ii) transcription inhibitors such as 5,6- dichloro-1-D-ribofuranosyl benzimid azole (DRB) severely slow down the mobility of TFIIH-GFP while that of XPG-GFP is not affected and iii) the kinetics of incorporation of these two proteins into the NER complex are different (t1/2 of 110s for TFIIH and 200s for XPG), demonstrating that XPG and TFIIH are indeed distinct protein complexes in vivo and only interact on damaged DNA (Chapter 4) (Hoogstraten et al., 2002; Zotter et al., 2006). A direct interaction was found between XPG and PCNA, which could possibly reflect a mechanistic linkage between the incision and repair synthesis stage of NER (Gary et al., 1997). However, XPG associates with DNA at the 3’ side of the lesion, while loading of DNA pol δ/ε by PCNA occurs at the 5’ side of the lesion (i.e. where ERCC1-XPF is bound), thus it appears that PCNA and XPG operate on opposite side of the repair patch.

The second NER endonuclease is a hetereodimer that consists of ERCC1 (33 kDa) and XPF (103 kDa), which cleaves DNA at the 5’ side of the junction between single-

30 NER in mammals stranded and double-stranded DNA. During NER, ERCC1-XPF incises the DNA between 15-24 nucleotides away from the 5’ side of the lesion (de Laat et al., 1998a; Matsunaga et al., 1995; Sijbers et al., 1996a). Although the XPF subunit contains the nuclease domain that cleaves the DNA, ERCC1 is indispensable for nuclease activity. The ERCC1 and XPF proteins share significant structural similarities and were suggested to have a common origin (Gaillard and Wood, 2001). In contrast to XPF, the ERCC1 polypeptide contains an inactivate nuclease domain and is incapable of cleaving DNA (Gaillard and Wood, 2001). The stability of both ERCC1 and XPF depends on complex formation, suggesting that these proteins stabilize each other in cells (Sijbers et al., 1996b; Yagi et al., 1997). Stable ERCC1-XPF complex formation depends on the C-termini of ERCC1 (residues 224-297) and XPF (residues 814-905) that form a double helix-hairpin-helix (HhH) motif and the ERCC1 HhH-motif only folds properly in the presence of XPF (de Laat et al., 1998c; Tripsianes et al., 2005). The HhH-motif of ERCC1 is involved in binding to ssDNA and its recruitment to repair complexes is mediated via specific interactions with a small region of XPA that binds a V-shaped hydrophobic groove in the central domain of ERCC1 containing mainly basic and aromatic residues (Tsodikov et al., 2005; Tsodikov et al., 2007). Evidence has been presented that the unstructured ERCC1-binding domain and the DNA-binding domain of XPA associate, rendering the ERCC1-binding domain unavailable for XPA-ERCC1 interactions. Binding of XPA to DNA lesions disrupts this intra-molecular interaction and triggers binding of ERCC1 to XPA, providing a structural basis for the damage-specific association between these two repair proteins (Buchko et al., 2001). Unlike, XPG, ERCC1-XPF does not appear to have an architectural role in NER complex assembly since it is dispensable for full open complex formation and appears to be required only for its catalytic activity (Evans et al., 1997b). The incision activity of ERCC1-XPF is stimulated by direct interactions with RPAand phosphorylation of a serine residue in XPB (S751) was shown to prevent ERCC1-XPF-mediated incision (Coin et al., 2004; de Laat et al., 1998b). Thus, it appears that de-phosphorylation of XPB S751 is required to incise 5’ of the lesion, reflecting an additional role for TFIIH in incision coordination (Coin et al., 2004). The two incisions during NER appear to be coupled. The 5’ incision by ERCC1-XPF follows the XPG-mediated 3’ incision resulting in excision of a single-stranded DNA fragment containing the lesion (Mu et al., 1996). ERCC1 is generally assumed to be the last factor to join the repair complex based on the in vitro finding that ERCC1- XPF can be added to a preformed incision complex containing all other factors to make the 5’ incision (Evans et al., 1997b; Mu et al., 1996; Mu et al., 1997). Interestingly, the in vivo binding rates of ERCC1-GFP and TFIIH-GFP are similar showing that pre- incision factors bind to the nascent NER complex with almost the same rate (Moné et al., 2004). Moreover, inefficient ERCC1-XPF recruitment is observed in several NER deficient cell lines raising doubts about whether there really is a strict order of NER factor assembly (Oh et al., 2007). Besides in NER, ERCC1-XPF is involved in several

31 Chapter 2 other processes such as homologous recombination, repair of interstrand cross-links and telomere maintenance (Niedernhofer et al., 2004; Sargent et al., 2000; Sargent et al., 1997; Zhu et al., 2003). This explains why ERCC1- and XPF-deficient cells are highly sensitive to crosslinking agents in contrast to other NER-deficient cells (Larmi- nat and Bohr, 1994; Li et al., 1999). In agreement, inactivation of ERCC1 or XPF in mice results in a more severe phenotype compared to other NER-deficient mice, which involves sensitivity to UV, developmental delay and early death (Friedberg and Meira, 2006; Tian et al., 2004; Weeda et al., 1997a). The only ERCC1-deficient patient described to date displayed a dramatic phenotype reminiscent of ERCC1- deficient mice and died at the age of 13 months (Jaspers et al., 2007). Some XP-F individuals (mainly Japanese) have been described, which displayed a relatively mild phenotype compared to XPF-deficient mice. The lack of developmental problems in these patients was attributed to considerable residual XPF activity (Sijbers et al., 1996a; Sijbers et al., 1998). A unique XP-F patient with a severe phenotype reminiscent of ERCC1- and XPF-deficient mice was recently described displaying signs of accelerated aging (progeria-like syndrome) and sensitivity to crosslinking agents (Niedernhofer et al., 2006).

Repair synthesis machinery: RPA, RF-C, PCNA, DNA pol δ/ε and DNA Ligase III During NER, dual incision is followed by repair synthesis in which the generated ssDNA gap is filled in by replication proteins. The dual incision and repair synthesis stages of NER can be separated in vitro and the only shared factor appears to be RPA. Consistent with this notion, isolation of repair intermediates in vitro showed that RPA remains associated with the excised DNA upon dual incision (Riedl et al., 2003). In addition to RPA, repair synthesis requires RF-C, PCNA, pol ε and Lig I in vitro (Aboussekhra et al., 1995; Shivji et al., 1992; Shivji et al., 1995). Indeed, both endogenous and GFP-tagged RF-C and PCNA accumulate after UV irradiation in living cells, which is consistent with repair synthesis by DNA pol δ/ε (R. Overmeer & A. Gourdin, unpublished; Essers et al., 2005; Moser et al., 2007). PCNA is a homotri- meric sliding clamp that encircles the DNA and acts as a processivity factor for pol δ/ε by tethering these enzymes to the DNA (Maga and Hubscher, 2003; Podust et al., 1998). The ATP-dependent binding of PCNA is mediated by the clamp loader RF-C. Binding of both protein complexes requires dual incision, which is consistent with the idea that a free 3’-OH group (that is formed upon ERCC1-XPF incision) is required for binding of PCNA to DNA (Balajee et al., 1998; Essers et al., 2005). In fact, binding of PCNA and subsequent repair synthesis is observed in XP-G cells in which a catalytically inactive XPG protein is expressed that allows uncoupled 5’ incisions by ERCC1-XPF. This demonstrates that a free 5’-end is sufficient to initiate repair synthesis (Wim Vermeulen, personal communication). The dissociation of GFP-PCNA is significantly delayed in the presence of DNA synthesis inhibitors consistent with a role in repair synthesis (Chapter 6). Interestingly, we provide evidence in chapter 6

32 NER in mammals that the binding GFP-XPA, but not XPC-GFP or ERCC1-GFP is stabilized by the presence of repair synthesis, suggesting that XPA remains associated with RPA during this late stage of NER (Chapter 6). In agreement, chromatin immuno-precipitation experiments using antibodies against repair synthesis factors also pulled down XPA, but not XPC or TFIIH, supporting a role for XPA in repair synthesis (L. Mullenders, personal communication) (Moser et al., 2007). Recent in vivo experiments showed that predominantly DNA pol δ but not pol β is recruited to repair patches upon UV irradiation in replicating and quiescent cells (Moser et al., 2007). The final step in NER is the ligation of the 5’ end of the newly synthesized DNA to the original sequence. Although Lig I is sufficient for sealing nicks during in vitro repair synthesis, recent in vivo studies revealed that the dominant ligase involved in NER is XRCC1-Lig IIIα, which is essential for removal of lesions and rejoining of nicks in chromosomal DNA (Moser et al., 2007). In addition to XRCC1-Lig III, recruitment of Lig I together with pol ε is also observed in replicating cells that are in late G1 or S phase (Moser et al., 2007). These findings reveal that during the cell cycle, cells have differential requirements of ligases and polymerases in repair synthesis.

Assembly and disassembly of repair complexes: a model for GGR In the following section we propose a model for global genome repair based on the characteristics of the repair proteins described above.

Damage recognition UV-induced DNA lesions are recognized by the XPC complex in global genome repair (Sugasawa et al., 1998; Volker et al., 2001). It is thought that a significant fraction (~90%) of the XPC protein pool continuously associates with and dissociates from chromatin non-specifically (Hoogstraten et al., 2003). It is unclear if this low affinity binding to DNA contributes to the detection of helix distorting lesions. Two conserved aromatic residues in the XPC polypeptide confer an aversion for damaged-DNA and an ssDNA binding domain targets XPC to the undamaged strand (Maillard et al., 2007). The binding of XPC result in slight opening of the DNA surrounding a lesion (~6 nt) (Tapias et al., 2004), which facilitates the binding of subsequent factors. In fact, locally pre-melted lesions are efficiently repaired without the need for XPC suggesting that the main function of this protein is to increase the single-strandedness of a DNA injury (Mu et al., 1997). Besides XPC, mammalian cells encode a second damage-recognition protein: DDB2, which is dispensable for in vitro reconstituted NER (Aboussekhra et al., 1995). However, DDB2 is essential for CPD repair in cells and considerable accelerates the repair of 6-4 PPs at low UV fluencies and is likely to mediate repair in a chromatin context (Hwang et al., 1999; Moser et al., 2005). We proposed that DDB2 creates a local chromatin environment around lesions that facilitates the assembly of repair complexes (Luijsterburg et al., 2007). In support of this idea, it was demonstrated that DDB2 mediates the ubiquitylation of histones H3 and H4 resulting in

33 Chapter 2 destabilization of nucleosomes (Wang et al., 2006a). Moreover, the interaction between DDB2 and the p300 acetyltransfersase might also contribute to more efficient repair in chromatin (Datta et al., 2001). In addition to facilitation of repair by modifying chromatin, it was also suggested that DDB2 directly recruits XPC to lesions via protein-protein interactions (Sugasawa et al., 2005; Yasuda et al., 2007). It is likely that several other chromatin-modifying activities are involved in repair. These will be discussed in the last section.

Open complex formation Upon lesion recognition, TFIIH binds to damaged DNA via an interaction with XPC (Yokoi et al., 2000). It can not be excluded that RPA binds before or together with TFIIH since accumulation of RPA is observed in XP-B/CS cells (Rademakers et al., 2003). It might be that the small ssDNA bubble created by XPC is sufficient for RPA binding. TFIIH contains the ATP-dependent XPD helicase that unwinds a stretch of ~10 nucleotides following stimulation by XPB (Coin et al., 2007). In addition to TFIIH and XPC, full opening of ~30 nucleotides requires RPA and XPA and it appears that XPG but not ERCC1-XPF stabilizes opened pre-incision complexes (Evans et al., 1997b). In addition to its helicase activity, TFIIH appears to have a structural role in NER complex formation (Mu and Sancar, 1997; Mu et al., 1997). XPA and TTDA stimulate the ATPase activity of TFIIH in the presence of lesions, which is required for full opening around the lesion (Coin et al., 2006; Giglia-Mari et al., 2006; Winkler et al., 2001). The open complex is formed asymmetrically around an injury with the 3’ border closer to the lesions (2-8 nucleotides) than the 5’ border (12-24 nucleotides) (Evans et al., 1997b). RPAbinds to the undamaged strand and correctly positions XPG at the 3’ side and ERCC1-XPF at the 5’ side (de Laat et al., 1998b). The 5’-oriented side of RPA contains a strong ssDNA-binding motif and this side of RPA interacts with XPG bound at the 3’ side of the lesions, which defines the 3’ boundary of the NER complex ~10 nucleotides from lesion. Subsequently, RPA elongates to bind ~30 nucleotides and its 5’-oriented side interacts with ERCC1-XPF, which defines the 5’ boundary ~20 nucleotides from the lesion. Measurements on the in vivo assembly rates of XPC-GFP, XPG-GFP and GFP-XPA show that these factors associate with the nascent repair complex with similar binding rates of about 15-20 molecules/s (Chapter 6). Similarly, TFIIH-GFP and ERCC1-GFP have been shown to have assembly rates in the same range, suggesting a scenario in which NER complex assembly is very rapid once XPC associates with a lesion (Moné et al., 2004). It has been suggested that the XPC dissociation rate increases (~ 3-fold) after helical unwinding by TFIIH or by binding of XPG to repair complexes in vitro (Riedl et al., 2003; Tapias et al., 2004). We provide evidence that the XPC dissociation rate is affected by all pre-incision proteins (TFIIH, XPA, XPG, ERCC1, XPF; Chapter 6), consistent with a scenario in which XPC remains associated until a full pre-incision complex is assembled or possibly until dual incision is performed.

34 NER in mammals

Dual incision Following full open complex formation, incisions on both sides of the lesion are catalyzed by the structure-specific endonucleases ERCC1-XPF at the 5’ side and XPG at the 3’ side. The DNA is incised asymmetrically around the lesion consistent with the borders of the open complex. The two incisions are closely coupled even though the XPG-mediated incision precedes incision by ERCC1-XPF (Evans et al., 1997a; Mu et al., 1996). Cleavage by ERCC1-XPF requires the presence but not the catalytic activity of XPG, suggesting that XPG has a structural role in creating a suitable substrate for the 5’ incision (Wakasugi et al., 1997). Incision coordination is regulated by RPA and TFIIH. The 5’ incision by ERCC1-XPF is stimulated by the 3’ oriented side of RPA bound to the opposite strand and requires de-phosphorylation of XPB at serine 751, whereas the 3’ XPG-mediated incision is likely to be stimulated by TFIIH (Coin et al., 2004; de Laat et al., 1998b; Wakasugi et al., 1997). The activity of both nucleases is inhibited by TFIIH in the absence of ATP, suggesting that ATP hydrolysis is required to incise the DNA (Winkler et al., 2001). It has been suggested that most pre-incision factors are released simultaneously following dual incision with the exception of RPA. However, measurements of the off-rates of repair proteins in vivo suggest non- simultaneous release for some repair factors. For instance, XPC dissociation (t1/2 ~ 25s) is significantly faster than that of XPG (t1/2 ~ 50s) and XPA, whereas XPA (t1/2 ~ 75s) is released considerably slower than XPG (Chapter 6) (Luijsterburg et al., 2007; Rademakers et al., 2003; Zotter et al., 2006). The dissociation kinetics of XPG, TFIIH and ERCC1 are similar (t1/2 ~ 50s) (Hoogstraten et al., 2002; Luijsterburg et al., 2007; Zotter et al., 2006). We provide evidence in Chapter 6 that XPA binding is stabilized by repair synthesis, suggesting that XPA has affinity for a repair intermediate formed after dual incision. This seems to be supported by ChIP experiments on late repair factors such as XRCCI (Moser et al., 2007). On the other hand, repair synthesis does not result in strand displacement beyond the repair patch at the 3’ side. This might be mediated by the interaction between PCNA and XPG, but could also be coordinated by RPA (Gary et al., 1997). In vitro evidence indicates that XPG is not released following dual incision but might require additional factors. Whether this is true in vivo remains to be elucidated (Riedl et al., 2003).

Repair synthesis Finally, the repair synthesis factors RF-C, PCNA, RPA and DNA pol δ fill in the resulting gap, which is subsequently sealed by XRCC1-Lig III (Moser et al., 2007; Riedl et al., 2003). The dissociation of GFP-PCNA from sites of damage is significantly slower (t1/2 ~ 225s) than that of other repair factors. Moreover, steady-state levels of bound PCNA are high up to 9 hrs following irradiation when pre-incision proteins such as XPC or XPA are not detected at repair sites any longer (Chapter 6) (Essers et al., 2005). This suggests that PCNA has affinity for a repair intermediate to which pre-

35 Chapter 2 incision proteins do not bind. Similarly, GFP-tagged RPA is also detected at repair sites up to 9 hrs following irradiation, suggesting it may bind to the same repair intermediate that PCNA binds to (Gourdin and Luijsterburg, unpublished). The nature of this repair intermediate remains to be elucidated. This raises questions about what triggers the release of PCNA and other repair synthesis factors following gap-synthesis. It would be of particular interest to study the dynamics of GFP-tagged repair synthesis factors (RPA, RF-C) as well as pol δ and Lig III. In addition, ChIP experiments on late and early factors are very powerful and provide molecular insight into the type of repair complexes formed during NER in vivo (Fousteri et al., 2006; Moser et al., 2007).

Chromatin remodelling and post-repair chromatin restoration A major obstacle for NER in eukaryotic cells is the fact that nuclear DNA is packaged as chromatin. This has several implications for repair. In general, chromatin is considered repressive to processes associated with DNA and the efficiency of repair in a chromatin context is decreased to about 10% of that on naked DNA (Hara et al., 2000; Ura et al., 2001; Wang et al., 1991). To overcome this obstacle, cells employ a number Fig 3. Cartoon model for NER of chromatin remodel ing activities to create a more permissive environment for repair. In addition, following removal of DNA injuries, cells need to restore the original chromatin structure to maintain the epigenetic information, which requires additional factors that are recruited to sites of repair. Such a model in which chromatin rearrangements precede and follow DNA repair itself, has been put forward as the access-repair-restore model that has proven very useful for understanding how NER operates on a chromatin template (Green and Almouzni, 2002; Smerdon, 1991). In this section, we will analyze which chromatin remodeling activities

36 NER in mammals are or might be involved in repair of UV-induced DNA lesions. First, the organization of chromatin in the cell nucleus is discussed.

Chromatin: from nucleosome to chromatin fiber The nucleosome is the basic unit of chromatin and consists of ~150 bp of DNA tightly wrapped around a protein octamer. The octamer, in turn, comprises two of each of the highly conserved histone proteins H2A, H2B, H3 and H4 that all have a characteristic histone-fold made up of 3 hydrophobic α-helices (Luger et al., 1997; Sandman and Reeve, 2006). Two H3-H4 dimers form a tetramer through interactions between the histone-folds (between H3-H3’) and two H2A-H2B dimers associate with this tetramer to form an octamer (with interactions between H2B-H4). The histone octamer interact with DNA by inserting arginine residues into the minor groove every helical turn. One nucleosome is found approximately every 200 bp of DNA throughout the genome (Luger et al., 1997). Text books often depict a hierarchical organization of chromatin, in which nucleosomal arrays coil up to form the 30 nm fiber that is subsequently stabilized by linker H1 and folded into a higher order structure that eventually generates interphase chromosomes and finally mitotic chromosomes (Varga-Weisz and Becker, 2006). In reality, however, very little is known about the organization of chromatin beyond the nucleosomal array, although we are beginning to understand some of its organizing principles. The emerging view is that chromosomes are confined to discrete territories (~2 µm in diameter) that show little intermingling with chromatin from neighboring territories (Cremer and Cremer, 2001). The chromatin fiber within territories is organized in loops that may be attached at their base to a structural network of scaffold proteins (Cremer and Cremer, 2001). DNA sequences that bind this nuclear scaffold as well as specific scaffold-binding proteins have been identified (Cai et al., 2006; Galande et al., 2007; Nickerson, 2001). Scaffold-binding proteins play an import role in organizing chromatin loops that have been suggested to be ~100 kb in size that fold into rosette like-structures of up to several Mb (Galande et al., 2007; Goetze et al., 2007). One example is SATB1, which is a scaffold-binding protein that folds the MHC class 1 locus in distinct chromatin loops in T cells. This folding regulates the expression of cytokine genes from the locus (Cai et al., 2006).The precise nature of the nuclear scaffold or matrix is highly controversial. Chromatin loops containing active genes from different positions along the same chromosome or even from different chromosomes have been suggested to come together in transcription factories (de Laat and Grosveld, 2007; Fraser and Bickmore, 2007). Each chromosome territory contains several irregularly-shaped dense chromatin domains that are about 0.1-0.5 µm in size and are surrounded by inter-chromatin space that contains little or no chromatin. It has been suggested that the surface of dense chromatin domains constitute a functional barrier that allows single proteins to but not larger protein complexes above a certain threshold to pass (Cremer and Cremer, 2001). However, chromatin domains were shown to be readily accessible to macromolecules in the range of several hundred kDa

37 Chapter 2 arguing against this notion (Verschure et al., 2003). At the interface between the dense chromatin domains and the inter-chromatin space is the perichromatin region that constitutes a shell (~ 80 nm) of partly decondensed chromatin that loops outward into the inter-chromatin space (Fakan and van Driel, 2007). Interestingly, the major chromatin-associated processes such as replication and transcription occur predominantly in the perichromatin region and also polycomb-mediated gene silencing occurs locally at the periphery of condensed chromatin (Cmarko et al., 1999; Cmarko et al., 2003; Jaunin et al., 2000). In contrast to replication and transcription, little is known about the spatial organization of NER in the human interphase nucleus. In Chapter 5, we provide evidence that the NER proteins XPC and XPA accumulate specifically in the perichromatin region after UV irradiation, suggesting that this is the main site of NER complex assembly. In addition, we show that UV-damaged chromatin undergoes considerable large-scale decondensation and we propose that this facilitates the relocation of DNA lesions from within dense chromatin domains to the perichromatin region (Chapter 5). Condensation of UV-damaged chromatin domains is observed 1 hr after UV irradiation, suggesting a transient decondensation of chromatin domains after UV irradiation (Chapter 5). Recent studies demonstrated decondensation of chromatin upon induction of double-strand breaks and UV-induced DNA damage, suggesting that chromatin relaxation might be a more general response to DNA damage (Carrier et al., 1999; Kruhlak et al., 2006; Murga et al., 2007; Rubbi and Milner, 2003; Ziv et al., 2006). Several mechanisms may contribute to these large- scale chromatin changes, including covalent modifications of histone tails and chromatin remodeling by specialized remodeling complexes. These mechanisms will be discussed below.

Histone modifications and NER Numerous covalent modifications of core histones have been identified that are tightly linked to gene activity and repair. These modifications, that are collectively described as “the histone code”, include acetylation, methylation, phosphorylation and ubiquitylation (Jenuwein and Allis, 2001; Turner, 2002). Acetylation and ubiquitylation of histones have both been associated with NER and will be discussed below. Although phosphorylation of the histone variant H2A.X at S139 (γH2A.X) is generally considered to be a marker for DSBs, H2A.X is also phosphorylated in response to UV irradiation (Hanasoge and Ljungman, 2007; Marti et al., 2006). This phosphorylation is mediated by the ATR kinase and depends on NER if cells are in G1 (Marti et al., 2006; Stiff et al., 2006). It is likely that γH2A.X has a function in signaling to checkpoint proteins. Several studies have demonstrated that histones H3 and H4 are hyper-acetylated in human cells following UV irradiation, resulting in higher repair rates (Brand et al., 2001; Ramanathan and Smerdon, 1986). Acetylation of histones is generally associated with a more open and active chromatin structure, for instance transcriptionally active genes are often acetylated. Enzymes known as histone

38 NER in mammals acetyltransferses (HAT), of which p300/CBP and GCN5 are well known examples, mediate the acetylation of N-terminal histones tails. Histone acetylation is directly linked to damage recognition in GGR and TCR since p300/CBP directly interacts with DDB1 and DDB2 and this HAT is recruited to DNA damage by CSB (Datta et al., 2001; Fousteri et al., 2006; Rapic-Otrin et al., 2002). Another study provided additional evidence for a role of histone acetylation in repair, by showing that local UV irradiation caused a p53-dependent global chromatin relaxation mediated by p300 (Rubbi and Milner, 2003). Interestingly, the TFTC and STAGA protein complexes, which both have HAT activity (since one of the subunits is GCN5), have been suggested to have affinity for UV lesions (Brand et al., 2001; Martinez et al., 2001). Both the TFTC and STAGA complexes contain a DDB1 homologue (called SAP130) and STAGA also associates with DDB2 (Brand et al., 2001; Martinez et al., 2001). Therefore, it is likely that binding of both complexes to UV lesions is mediated by DDB2. In addition to recruiting HATs, DDB2 is part of an E3 ubiquitin (ub) ligase that ubiquitylates histones H3 and H4 following UV irradiation. This subsequently results in weakened interaction between histones and DNA, which is likely to facilitate the recruitment of NER proteins (Wang et al., 2006a). In contrast to H3 and H4 ubiquitylation, conjugation of H2A (at lysine 119) with ubiquitin is generally associated with a closed and inactive chromatin state and gene silencing (Jason et al., 2002; Wang et al., 2004). In this respect, the NER-dependent and ATR-dependent mono-ubiquitylation of H2A at Lys119 (uH2A), which was shown to be mediated by the Ring2 ub ligase is of particular interest (Bergink et al., 2006). Although Ring 2 is part of the polycomb- repressive protein complex 1 (PRC1), it is not known whether Ring2 is recruited to UV lesion as part of PRC1 or as part of another Ring2-containing complex. Interestingly, recent ChIP experiments revealed enrichment of tri-methylated H3 at lysine 27 (H3K27Me; L. Mullenders, personal communication) and this histone modification is recognized by polycomb proteins containing a chromodomain (CD) such as Cbx7 that is part of PRC1 (Jones et al., 2000). It is feasible that recruitment of Ring2 and subsequent ubiquitylation of H2A is triggered by the presence of H3K27me. More recently, the E3 ub ligase RNF8 was demonstrated to mediate poly-ubiquitylation of H2A in response to DSBs (Mailand et al., 2007) and UV-induced lesions (W. Vermeulen, personal communication). This suggest a two-step mechanism in which Ring2-mediated mono-ubiquitylation of H2A following UV irradiation precedes poly-ubiquitylation by RNF8. Interestingly, an enzyme that mediates the de-ubiquitylation of H2A (called USP3) was recently identified, which is expected to also catalyze the removal of NER-induced uH2A (Nicassio et al., 2007). The precise role of H3K27me and uH2A during or following NER remain to be elucidated.

Chromatin modifying proteins and NER Several non-chromosomal proteins have the ability to modify chromatin (either directly by for instance bending it or by recruiting an enzymatic activity) or to bind to specific

39 Chapter 2 histone modifications (i.e. “read” the histone code). Well known examples are chromo- and bromo-domains, which specifically bind to methylated and acetylated histones, respectively (de la Cruz et al., 2005; Jones et al., 2000). Other examples are proteins that mediate DNA methylation (such as the Dnmt family) and factors that specifically bind to methylated DNA (such as MeCP2) or proteins that methylate histones (such as the SUV and SET families) and factors that bind methylated histones (such as HP1 and PcG proteins) (Bestor, 2000; Kouzarides, 2002). Some of these chromatin modifiers have been linked to NER and these will be discussed in this section. Several acidic proteins have been suggested to compete with DNA for binding to histones thereby modulating the accessibility of nucleosomal DNA. An example of such a protein is GADD45, which is a small (19 kDa) acidic stress-responsive protein that binds to and increases the accessibility of UV-irradiated and acetylated nucleosomes in vitro (Carrier et al., 1999). It appears that GADD45 preferentially interacts with altered chromatin structures including nucleosomes containing CPDs without the need for specific repair proteins. Whether GADD45 binds to UV damaged-chromatin independently of NER in living cells remains to be elucidated. Inactivation of GADD45 in mice or knock-down of GADD45 expression in human cells results in considerably reduced repair rates of both 6-4 PPs and CPDs suggesting a role in facilitating NER in chromatin (Smith et al., 2000; Smith et al., 1996). Interestingly, GADD45 was shown to interact with tumour suppressor proteins of the ING1 family (inhibitor of growth 1) that share many functions with p53 and contain a conserved PHD motif (Cheung et al., 2001). Two particular isoforms of ING1, designated ING1b and ING2, were shown to increase the accessibility of chromatin to NER proteins by inducing H4 acetylation and chromatin relaxation (Kuo et al., 2007; Wang et al., 2006b). In addition, over- expression of INGb significantly enhanced NER in vivo, which also required p53 (Cheung et al., 2001). Thus, the combined action of p53, GADD45 and ING proteins may increase the accessibility of UV-lesions in nucleosomal DNA. Several studies have provided evidence for the involvement of HMG proteins, which often contain acidic tails, in the repair of UV lesions. HMG proteins are abundant architectural proteins that bend DNA up to 100° and destabilize higher chromatin structures. The HMG1 protein (belonging to the HMG-B family) was shown to have affinity for UV- damaged nucleosomes in vitro, although the significance of this observation is currently not known (Pasheva et al., 1998). Better understood is the role of HMG-N, which was demonstrated to be recruited to TCR complexes by CSB (Fousteri et al., 2006). No recruitment of HMG-N was found in cells deficient in CSB, showing that this protein is exclusively involved in TCR and not GGR (Fousteri et al., 2006). Indeed, HMG-N-deficient cells show decreased rates of CPDs removal in actively transcribed genes but not in the genome overall (Birger et al., 2003). Several studies have shown that HMG proteins directly compete for binding sites with histone H1 that bind to linker DNA and fold chromatin fibres into more compact structures (known as chromatosomes) (Brown et al., 2006; Catez et al., 2002; Catez et al., 2004). This would

40 NER in mammals provide a mechanism as to how HMG proteins destabilize higher-order chromatin structures and it was shown that association of HMG proteins with nucleosomes assembled on promoters upon transcription activation resulted in the loss of H1 (Ju et al., 2006). Interestingly, partial depletion of H1 from mouse cells was shown to strengthen the DNA damage response and in particular activation of ATR during DBS repair (Murga et al., 2007). Given that ATR is also activated upon UV irradiation (Stiff et al., 2006), it is feasible that the loss of H1 that might be triggered by binding of HMG proteins also potentiates the DNA damage response following UV irradiation. Recently, evidence was provided that Dnmt1 (a maintenance DNA methyl-transferase) but not Dnmt3a and 3b (which are de novo Dnmt’s) are recruited to sites of laser- induced DNA damage after sensitisation with BrdU (Mortusewicz et al., 2005). However, the types of DNA injuries that are produced using this method are ill-defined and several repair pathways were shown to be activated simultaneously (Dinant et al., 2007; Williams et al., 2007). We have recently demonstrated that GFP-tagged Dnmt1 is recruited to sites of UV-induced DNA damage as well as HP1α, β and γ (Chapter 7). Moreover, we show that the binding of the HP1 proteins depends on the chromoshadow-domain (CSD) but not on the chromo-domain (CD). The CSD domain (once dimerized with a second CSD) interacts with several nuclear proteins that contain a short hydrophobic motif (PxVxL), whereas the CS binds methylated H3 at lysine 9 (Jones et al., 2000; Thiru et al., 2004). The fact that HP1 recruitment to UV-lesions depends on the CSD suggests that HP1 is recruited to DNA lesion by a protein already present at the site of damage. This may be mediated by direct interaction between HP1 and RNA polymerase II (Mateescu et al., 2008). Interestingly, HP1 accumulation is observed in cells deficient in any of the known damage-recognition proteins involved in NER (i.e. XPC, DDB2 and CSB) and appears to be repair-independent (Chapter 7). The precise mechanism by which HP1 and Dnmt1 proteins are recruited to UV lesion remains elusive and requires further studies. HP1 and Dnmt1 may play a role in inhibition of transcription, which occurs in UV-damaged cells independently of functional NER (Moné et al., 2001). Chromatin assembly factor (CAF-1), which also binds to HP1, was shown to be recruited in an NER-dependent manner to sites of UV- induced DNA damage in living cells (Green and Almouzni, 2003). This is of particular interest since CAF-1 and HP1 are known to interact during replication (Quivy et al., 2004). However, binding of CAF-1 depends on functional NER in contrast to that of HP1 raising the question whether these proteins interact during DNA repair (Green and Almouzni, 2003). In agreement with the role of CAF-1 as chromatin assembly factor, it was demonstrated that histone H3.1 was assembled de novo at repair sites reflecting a chromatin restoration step following repair (Polo et al., 2006). It would be of interest to investigate whether histone variants, such as H2A.Z or H3.3 are deposited following repair. In conclusion, it appear that chromatin modifiers have a key role in modulating the accessibility of nucleosomal DNA to the repair machinery as well as in the restoration of the chromatin state following repair, consistent with the proposed

41 Chapter 2 access-repair-restore model (Green and Almouzni, 2002).

ATP-dependent nucleosome remodelling factors and NER A number of large multi-protein complexes (200 kDa – 2 MDa) can enzymatically modulate chromatin structure at the expense of ATP hydrolysis (Becker and Horz, 2002; Varga-Weisz and Becker, 2006). These nucleosome remodelling factors contain an ATPase of the SWI2/SNF2 (Switch/Sucrose non-fermentable) family, which is reminiscent of a DNA helicase. In addition, nucleosome remodelling factors contain targeting and regulatory subunits (Varga-Weisz and Becker, 2006). Examples of nucleosome remodelling factors are CHRAC, BRAHMA and INO80 and several of these protein complexes have been linked to NER. A direct link between nucleosome remodelling and repair was provided by the finding that CSB, which is a key TCR factor, is also an ATPase of the SWI2/SNF2 family that directly binds to histones (probably at their tails) and remodels nucleosomes in vitro (Citterio et al., 2000). Inactivation of the ATPase domain of CSB abolished its function in repair and it was suggested that CSB has a more general role in chromatin remodelling and regulation of genes involved in cell growth and the stress response that is not restricted to repair (Newman et al., 2006; Selzer et al., 2002). Additional evidence suggesting a role for ATP-dependent remodelling factors in repair comes from in vitro studies, showing that ACF1 (ATP-utilizing chromatin assembly and remodelling factor) facilitates dual incision in di-nucleosomes, especially in the linker region (Ura et al., 2001). The bromodomain-containing ACF1 is the largest subunit of the human CHRAC complex (chromatin accessibility complex) that contains the ATP-ase subunit ISWI (imitation switch) and two histon-folds proteins (Eberharter et al., 2001; Poot et al., 2000; Varga- Weisz et al., 1997). Mammalian cells contain two ISWI proteins designated SNF2H and SNF2L and ACF1 was found to predominantly associate with SNF2H (Poot et al., 2000). Nucleosome remodelling as well as sliding by ISWI was shown to be considerably stimulated by ACF1 and it was demonstrated that the tails of H4 are required for nucleosome sliding (Eberharter et al., 2001). Recently, GFP-tagged ACF1 was shown to be recruited to sites of UV-induced DNA damage in human cells, providing evidence for an in vivo role of ACF1, and possibly CHRAC, in NER (Chapter 7). It would be of interest to determine if ISWI and the closely related ACF1 homologue WSTF (William syndrome transcription factor) (Poot et al., 2004) are also recruited to UV lesions. In addition to ACF1 (containing ISWI), also SWI/SNF was shown modulate the accessibility of UV-damaged mono-nucleosomes in vitro (Gaillard et al., 2003; Hara and Sancar, 2002; Hara and Sancar, 2003). In fact, NER proteins XPC, XPA and RPA were found to stimulate the remodelling activity of SWI/SNF to allow access to lesion in the nucleosome core (Hara and Sancar, 2002). It is interesting to note that the homologues of XPC-HR23B in yeast (called RAD4-RAD23) directly interact with a SWI/SNF remodelling complex and this interaction was shown to be enhanced following UV irradiation (Gong et al., 2006). Mammalian cells have two

42 NER in mammals closely related SNF2 proteins, known as BRM (Brahma) and BRG1 (Brahma related gene 1) that are part of different large multi-protein complexes in vivo. In agreement with a putative role for SWI/SNF during repair, we found that YFP-tagged BRG1 accumulates at sites of UV-induced DNA damage in human fibroblasts suggesting a role in modulating lesion accessibility during or prior to repair (Chapter 7). Another type of remodelling complex containing the ATPase INO80 has been intimately linked to DSB repair in yeast and it was shown that recruitment of INO80 to DSBs required phopsphorylation of H2A (Morrison et al., 2004; van Attikum et al., 2004). Mammalian cells also contain INO80-containing complexes and it is anticipated that these nucleosome remodelling factors are involved in facilitating NER in chromatin. Altogether, several ATP-dependent remodelling complexes, containing SWI/SNF (such as BRG1) or ISWI (such as ACF1) have been directly linked with mammalian NER and more proteins are likely to be identified in the years to come. It would be of particular interest to study how NER is affected by the depletion (e.g. knock-down) or inactivation of these nucleosome remodelling proteins.

Final considerations and outline of this thesis Cells respond to UV damage by activating several pathways. One of these includes assembly of a multi-protein repair complex on damaged chromatin that mediates removal of the damaged DNA. Although all core repair factors are known, we are only beginning to understand the complex choreography of pre-incision NER factors during repair in living cells. In Chapter 3 binding of NER endonuclease XPG is shown to depend on functional THIIH. Moreover, TFIIH and XPG are demonstrated to be distinct protein complexes, which interact only on damaged DNA. Chapter 6 provide a detailed quantitative analysis of all pre-incision NER proteins and shows that pre- incision factors exchange rapidly at sites of repair under steady-state repair conditions. Using mathematical modelling, we reveal that NER factors dissociate and rebind during the repair process. This in contrast to previous models in which pre-incision proteins were proposed to irreversibly bind until repair was completed. We propose a model in which irreversible enzymatic reactions produce repair intermediates (such as unwound or incised DNA) to which pre-incision NER proteins bind with different affinities. Such as model can fully explain the kinetic behaviour of NER factors in living cells. Chapter 5 provides insight into the spatial organization of NER and shows that NER pre-incision complexes are mainly assembled at the surface of compact subchromosomal chromatin domains. Moreover, we show that UV-damaged chromatin undergoes large-scale decondensation and propose that damaged DNA inside compact chromatin domains is relocated to the surface of these domains. In Chapter 4 the interplay between damage-recognition proteins DDB2 and XPC is studied. DDB2 is shown to bind UV-lesions independently of XPC and we propose that DDB2 prepares UV-damaged chromatin for the assembly of NER complexes. In contrast to detailed knowledge on the kinetics of pre-incision factors, we are only beginning to uncover the

43 Chapter 2 kinetic behaviour of repair synthesis NER proteins. In Chapter 6, we show that PCNA binding reaches a plateau several hours after UV irradiation when pre-incision proteins are no longer bound to repair sites. RPA, which has roles in pre-incision NER and repair synthesis, displays similar kinetic behaviour (Gourdin and Vermeulen, unpublished). These results reveal that repair synthesis factors display kinetic behaviour that is remarkably different from pre-incision NER proteins. It underscores that little is understood of the events that occur after dual incision. Both RPAand PCNA exchange at repair sites, showing that both proteins dissociate and (re)bind at time-points when pre-incision proteins are no longer bound. Clearly, these repair synthesis proteins have affinity for repair intermediates that are present several hours after UV irradiation. The nature of these repair intermediates remains elusive. It may be that repair synthesis proteins bind to DNA that is re-synthesized but on which nucleosomes are not yet assembled. In support of this, it was shown that the protein assembling H3.1 after NER (CAF-1), can be detected at repair sites 6 hrs following UV irradiation (Polo et al., 2006). It would be of interest to study the kinetic behaviour of repair synthesis factor such as RF-C, XRCCI-LigIII and DNA pol δ. This information could be used to build a complete mathematical model for the NER process, which provides comprehensive insight into how NER operates on chromatin in living cells. In Chapter 7 the HP1 proteins, which are versatile epigenetic regulators are demonstrated to bind to UV-damaged sites independently of any of the know NER factors. Similarly, DNA methtyltranferase 1 (Dnmt1) and chromatin remodelling complexes ACF1 and BRG1 are shown to bind to UV-damaged sites in repair-deficient cells. This suggest these proteins are involved in a DNA damage response pathway, which is independent of NER. We propose that HP1 proteins may regulate repression of transcription following UV irradiation. Chapter 8 is a perspective in which the mechanisms to assemble chromatin-associated complexes that carry out genome- controlling functions such as DNA repair, transcription and replication are discussed.

44 Chapter 3 Recruitment of the Nucleotide Excision Repair Endonuclease XPG to Sites of UV-induced DNA Damage Depends on Functional TFIIH

Angelika Zotter, Martijn S. Luijsterburg, Daniël O. Warmerdam, Shehu Ibrahim, Alex Nigg, Wiggert A. van Cappellen, Jan H. J. Hoeijmakers, Roel van Driel, Wim Vermeulen and Adriaan B. Houtsmuller

Adapted from Mol. Cell. Biol. 23, 8868-79, December 2006

45 Chapter 3

Abstract The endonuclease XPG is an indispensable core protein of the nucleotide excision repair (NER) machinery. XPG cleaves the DNA strand at the 3’ side of the DNA damage. Binding of XPG stabilizes the NER pre-incision complex and is essential for the 5’ incision by the ERCC1/XPF endonuclease. We have studied the dynamic role of XPG in its different cellular functions in vivo. We have created mammalian cell lines that lack functional endogenous XPG and stably express EGFP-tagged XPG. Live cell imaging shows that in undamaged cells XPG-EGFP is uniformly distributed throughout the cell nucleus, diffuses freely and is not stably associated with other nuclear proteins. XPG is recruited to UV-damaged DNA with a half-time of 200 s and dissociates under steady-state repair conditions with a half-time of 50 s. Recruitment to repair complexes requires functional TFIIH, although some TFIIH mutants allow slow XPG recruitment. Remarkably, binding of XPG to damaged DNA does not require the DDB2 protein, which is thought to enhance damage recognition by NER factor XPC. Together, our data present a comprehensive view of the in vivo behaviour of a protein that is involved in a complex chromatin associated process.

Introduction Nucleotide excision repair (NER) is a versatile DNA repair mechanism that removes different types of helix-distorting damage from the genome, including UV light- induced DNA damage, such as cyclobutane pyrimidine dimers (CPD) and 6-4 photo products (6-4 PP) (Friedberg, 2001; Hoeijmakers, 2001). The severe clinical features of three photo-hypersensitive hereditary NER disorders underscore its biological importance: the cancer-prone syndrome Xeroderma pigmentosum (XP), the neuro- developmental conditions Cockayne syndrome (CS) and trichothiodystrophy (TTD) (Lehmann, 2003). The multi-step NER process requires the coordinated actions of at least 25 polypeptides (Friedberg, 2005). The general modus operandi for NER comprises the following steps: i) recognition of DNA damage, ii) unwinding around the lesion, iii) dual incision on both sides of the damage, iv) removal of the excised oligonucleotide, and v) filling the generated gap by DNA polymerase and ligase (de Laat et al., 1999). Two different modes of NER exist, i.e. transcription-coupled NER (TC-NER) and global genome NER (GG-NER) (Hanawalt, 2000). TC-NER removes lesions exclusively from the transcribed strand of active genes, whereas GG-NER repairs damage at any other position in the genome. GG-NER protects against damage- induced mutagenesis and can thus be considered a cancer-preventing process, whereas TC-NER primarily promotes cellular survival and therefore may counteract aging (Mitchell et al., 2003). The damage sensor for 6-4 PP in GG-NER is the heterotrimeric XPC/HR23B/centrin2 complex (Sugasawa et al., 1998; Volker et al., 2001). In addition, the UV-damaged DNA binding protein (UV-DDB) assists XPC in the recognition of CPD (Fitch et al., 2003; Wakasugi et al., 2002) and facilitates 6-4 PP repair (Moser et

46 XPG dynamics al., 2005). In TC-NER lesions are detected by stalled elongating RNA polymerase II (van den Boom et al., 2004). After lesion-detection the two NER sub-pathways funnel into a common mechanism. Damage sensing is followed by the recruitment of the ten- subunit TFIIH complex (Giglia-Mari et al., 2004), which utilizes its helicase components XPB and XPD to locally unwind the DNA around the lesion. The structure-specific endonuclease XPG subsequently binds and promotes formation of an open DNA complex around the lesion (Evans et al., 1997). The next proteins that bind to the repair complex are the single-stranded DNA (ssDNA) binding replication protein A (RPA) and the damage verification factor XPA, which play an important role in the correct positioning of the 3’ endonuclease XPG and the 5’ endonuclease ERCC1/XPF (de Laat et al., 1998). After dual incision a stretch of ~30 nucleotides ssDNA containing the damage is released, after which the replication factors RPA, PCNA and DNA polymerase δ/ε fill in the resulting gap (Shivji et al., 1995). In the last step the newly synthesized DNA is sealed by DNA ligase I and the original chromatin structure is restored by chromatin assembly factor I (CAF I) (Green and Almouzni, 2003).

In vitro studies have resulted in a number of models for the assembly of the NER complex onto damaged DNA, proposing a completely pre-assembled holo-complex (Svejstrup et al., 1995), a partly pre-assembled NER complex (Guzder et al., 1996; Guzder et al., 1997; Guzder et al., 1999; Habraken et al., 1996), and the sequential assembly of individual NER factors, assuming conflicting assembly sequences (Riedl et al., 2003; Sugasawa et al., 1998; Wakasugi and Sancar, 1998; Wakasugi and Sancar, 1999). In vivo assembly studies on locally damaged nuclei support a sequential assembly scenario (Volker et al., 2001). We have previously studied the in vivo kinetics of the NER components ERCC1/XPF (Houtsmuller et al., 1999; Moné et al., 2004), TFIIH (Hoogstraten et al., 2002; Moné et al., 2004), XPA (Rademakers et al., 2003), XPC (Politi et al., 2005), and CSB (van den Boom et al., 2004). Together, these studies culminated to a model in which NER factors move freely throughout the nucleus and are incorporated one-by-one into repair complexes after the induction of DNA damage. However, the above-mentioned studies could not unambiguously identify the precise role of XPG, including at what stage the protein is incorporated in the emerging NER complex. Therefore, we have carried out a comprehensive in vivo analysis of the behaviour of XPG in DNA repair.

Results

Generation of cell lines stably expressing functional XPG-EGFP To study the nuclear distribution and dynamics of XPG in living cells we tagged the protein with enhanced green fluorescent protein (EGFP). EGFP was fused to the carboxy-terminus of human XPG (Fig. 1A), resulting in an XPG-EGFP fusion protein, which was stably expressed in XPG-deficient human fibroblasts (XPCS1RO-SV) and

47 Chapter 3 in Chinese Hamster Ovary cells (UV135). Fluorescently tagged XPG is predominantly located in the nucleus of both cell types in which it is uniformly distributed, with nucleoli being less populated (Fig. 1B). These observations are in accordance with earlier findings for fixed cells (Dunand-Sauthier et al., 2005; Thorel et al., 2004; Volker et al., 2001). Immuno-blot analysis of whole cell extracts of both cell types, using anti- XPG antibodies, showed that XPG-EGFP migrates in SDS-PAGE with a mobility corresponding to the expected size of the full-length fusion protein (~180 kDa, Fig. 1C; (O'Donovan et al., 1994)). Labeling with anti-EGFP antibodies did not reveal the presence of any other EGFP-containing polypeptides in the crude extracts (data not shown). This implies that all microscopy-based studies in this paper truly reflect the behaviour of XPG-EGFP. The Western blot in Fig. 1C indicates that XPG-EGFP is expressed at about the same level as endogenous XPG in wild type (HeLa) cells. Importantly, XPG-EGFP was able to restore normal UV-sensitivity of XP-G cells (Fig. 1D), showing that the fusion protein is functional in NER when expressed at physiologically levels.

Figure 1 Expression and functionality of XPG-EGFP. (A) Schematic representation of the XPG-EGFP-His(9)- HA fusion gene with the N-terminal and C-terminal nuclease domains (N and C, respectively) and different interaction domains indicated. I, Internal domain; PBD, PCNA binding domain; NLS, probable nuclear localization signal; aa, amino acids. (B) Localization of the XPG fusion protein in human fibroblasts (left two images, showing the fluorescence signal and an overlay of fluorescence and phase contrast) and CHO cells (right two images). XPG-EGFP is present mainly in the nucleus, except in the nucleoli. (C) Immuno-blot, probed with monoclonal anti-XPG, of 40 µg of whole-cell extract from HeLa (lane 2), human XPCS1RO-Sv (XP-G) expressing XPG-EGFP (lane 3), CHO (UV135) cells expressing XPG-EGFP (lane 4) and untransfected XPCS1RO-Sv (lane 5). The molecular mass of protein markers is indicated in kilodaltons (kDa). EGFP-tagged XPG migrates slower than endogenous XPG (upper and lower arrow, respectively). No XPG protein was detected in the human fibroblasts in which the severely truncated XPG-mRNA is probably highly unstable or not recognized. Chinese hamster XPG cannot be detected with our anti-XPG serum. Loading control: PCNA (arrowhead), asterisk indicates cross-reacting non-specific band only present in human cell extracts. (D) UV-survival of repair-proficient human MRC5 cells (wild type; light blue line), XPCS1RO cells (violet line), XPCS1RO cells stably expressing XPG-EGFP (clone 5cM; brown line), wild type CHO cells (AA8; dark blue line), XPG-deficient CHO cells (UV135; purple line), and UV135 cells expressing XPG-EGFP (clone 129; yellow line). The transfected cell lines show a correction of UV sensitivity to the wild type level.

48 XPG dynamics

Mobility of XPG-EGFP in the nucleus XPG has been reported to interact with other DNA repair proteins and with transcription factors and might therefore be part of a larger protein complex (Sarker et al., 2005). To investigate whether XPG moves freely through the nucleoplasm, is part of a larger complex, or is bound to immobile nuclear structures, we used FRAP (Fluorescence Recovery after Photo- bleaching; Fig. 2A). XPG-EGFP molecules in a specific nuclear region are bleached by a short light pulse, followed by monitoring the kinetics and extent of recovery of fluorescence due to the diffusion of non- bleached XPG-EGFP molecules into the bleached area. In non-UV irradiated living cells, monitored at 37°C, essentially all XPG-EGFP was mobile. The same redistribution kinetics were found for XPG-EGFP in CHO cells (Fig. 2B) and in human fibroblasts (Fig. 2C), indicating that XPG-EGFP mobility is independent of cell type and organism. Curve fitting shows that the effective diffusion coefficients (Deff) of XPG-EGFP in CHO cells and human fibroblasts are very similar, i.e. 6.1 ± 1.5 µm2/s and 4.0 ± 0.8 µm2/s, respectively (Fig. 2B and 2C). The expected Deff for a protein of ~180 kDa is between 5 and 6 µm2/s. Using combined FLIP and FRAP (Hoogstraten et al., 2002) we showed that the mobility of

Figure 2 FRAP analysis of XPG mobility. (A) Example of FRAP analysis to determine effective diffusion coefficients in non UV-irradiated cells. A strip (red rectangle) spanning the nucleus containing EGFP- tagged protein is bleached at high laser intensity. Subsequently, fluorescence recovery after photobleaching is monitored in the strip. (B-C) Graphical representation of FRAP analysis of EGFP-XPG in non-UV irradiated CHO cells (B) and human fibroblasts (C), the mean relative fluorescence (flu. after bleaching/flu. before bleaching) is plotted against the indicated time in seconds. Red lines: experimental data; green lines: simulated curves; blue lines at the bottom of each graph represent residuals which are a measure for the quality of the fits. (D) Simultaneous FLIP/FRAP analysis of XPG mobility in the nucleus of CHO cells. A small area at one pole of the nucleus is bleached for 1 s, subsequently fluorescence is monitored in time in the bleached (FRAP) and unbleached area (FLIP). The difference in EGFP intensity between the two areas after the bleach pulse is plotted on a log scale as a function of time. Light blue line, XPG redistribution at 37°C; dark blue line, XPG redistribution at 27°C; purple line, XPG redistribution in UV irradiated cells at 37°C; violet line, XPG redistribution in UV irradiated cells at 27°C. Experiments on UV- irradiated cells were performed between 10 min and 30 min after global UV-C irradiation.

49 Chapter 3

XPG is the same at 27°C and 37°C (Fig 2D). Together, this indicates that XPG in undamaged cells is not part of a larger complex, e.g. with TFIIH as has been suggested elsewhere (Araujo et al., 2001), and does not interact with immobile nuclear components, such as chromatin.

In vivo assembly of XPG-EGFP into the NER complex Analysis of the kinetics of de novo NER complex assembly in vivo has shown that incorporation of TFIIH and ERCC1/XPF into the pre-incision complex is not diffusion- limited (Moné et al., 2004). Association of the ERCC1/XPF incision factor depended on the presence of functional TFIIH (Moné et al., 2004). To determine how XPG-EGFP is incorporated into the repair complex in vivo, we analyzed its recruitment kinetics in nuclei immediately after local UV irradiation (Fig 3A). XPG-EGFP accumulation in the damaged area reached a plateau after about 10 min (Fig 3B). This plateau reflects a pseudo steady-state in which DNA repair takes place at a constant rate and the number of XPG molecules that are incorporated into repair complexes per unit time equals the number of molecules that are released from repair complexes. The rate of incorporation of XPG-EGFP was the same in CHO cells and in human fibroblasts, with a t1/2 of ~200s for both cell types (Fig 3B). This shows that the absence of the DDB2 subunit of the UV-DDB protein in CHO cells has no effect on the kinetics of

Figure 3 Accumulation of XPG-EGFP after local UV-DNA damage. (A) Time-lapse series of XPG-EGFP expressed in CHO UV135 cells prior and immediately after UV-C irradiation (100 J/m2). After taking pre-irradiation images cells were irradiated for 39s (lightning arrow), subsequently images were taken with 20s interval. (B) Incorporation kinetics of XPG-EGFP in CHO cells (UV135 – green line, n=5), CHO cells transfected with DDB2-mCherry (UV135 + DDB2 – red line, n=5) and human fibroblasts (XPCS1RO – blue line, n=5) at UV- damaged areas after 100 J/m2 UV-C. The local relative accumulation of XPG-EGFP was measured versus time. (C) Incorporation kinetics of XPG- EGFP in UV135 cells at 37°C (red line, n=5) and 27°C (blue line, n=5). The local accumulation of XPG-EGFP was measured and plotted as a percentage of the total EGFP fluorescence of the cell nucleus (37°C n=11, 27°C n=20) versus time after start of UV irradiation. Error bars represent SD between different experiments.

50 XPG dynamics incorporation of XPG in NER complexes that assemble on UV-damaged DNA. To investigate the role of DDB2 more directly, we transiently transfected CHO cells that stably express XPG-EGFP with DDB2 fused to the red fluorescent protein mCherry (Shaner et al., 2004). Subsequently, binding of XPG-EGFP was measured in cells that also expressed DDB2-mCherry. The rate of incorporation of XPG-EGFP was the same in transfected and non-transfected cells, i.e. with and without expression of DDB2 (Fig 4). Our experiments show that DDB2 does not changes the incorporation kinetics of XPG into the NER pre-incision complex.

Binding of XPG depends on the presence of functional TFIIH The rate of binding of XPG-EGFP to the emerging NER complex is temperature-dependent. The initial rate of XPG-EGFP incorporation at 37°C is about 1.5% per min and at 27°C about 0.5% per min (Fig 3C). Analogous to previous studies of the in vivo kinetics of incorporation of ERCC1/XPF (Moné et al., 2004) it is likely that the temperature- dependent step in NER complex formation is the DNA unwinding process, catalyzed by the helicase activity of TFIIH (Schaeffer et al., 1993). Previous studies, using fixed cells and in vitro NER assembly, were not conclusive about the question whether XPG incorporation required functional TFIIH (Dunand- Sauthier et al., 2005; Riedl et al., 2003; Thorel et al., 2004) .

Figure 4. Accumulation of XPG-EGFP after local UV irradiation in cells transfected with DDB2- mCherry. (A) Time-lapse series of XPG-EGFP expressed in CHO UV135 cells transiently transfected with DDB2-mCherry prior and immediately after UV-C irradiation (100 J/m2). Left column: XPG-EGFP signal, middle column: DDB2-mCherry signal, right column: Merge of green and red signal. Co-localization of both signals appears as yellow. The time after UV irradiation is indicated in seconds in each frame (B) Quantification of incorporation kinetics of XPG-EGFP in non-transfected hamster cells (green line, n=5) and incorporation kinetics of XPG-EGFP in hamster cells transfected with DDB2- mCherry (blue line, n=5) after 100 J/m2 UV-C. The red line shows de binding kinetics of DDB2-mCherry in cells also expressing XPG-EGFP (n=2). Error bars represent SD between different experiments.

51 Chapter 3

Figure 5 Accumulation of XPC and XPG after local UV-damage in human wild-type cells (C5RO) and various TFIIH mutants: 10 min (columns 1 and 2) and 30 min after UV-irradiation (columns 3 and 4). Columns 1 and 3, immunofluorescence labeling with anti-XPC antibody (green); columns 2 and 4, labeling with anti-XPG antibody (red).

Figure 6 FRAP analysis of UV-treated and untreated cells to visualize DNA damage- dependent immobilization of XPG-EGFP. (A, B) FRAP recovery curves (normalized to pre-bleach intensity set to one) in CHO cells and human fibroblasts, respectively. Black curves: XPG-EGFP recovery in untreated cells (as a reference); grey curves: recovery in UV irradiated cells. (C, D) FRAP recovery curves (normalized to maximum recovery post-bleach set to one) of XPG-EGFP in CHO cells and human fibroblasts, respectively. Black curves: XPG recovery in untreated cells (as a reference); grey curves: recovery in UV irradiated cells. (E, F) Immobilized XPG-EGFP fractions in CHO cells and human fibroblasts, respectively, in response to different UV doses. Light blue and dark blue bars depict measurements in cells cultured at 27ºC, after 8 and 16J/m2 UV, respectively. Error bars show the standard error of the mean.

52 XPG dynamics

To determine whether XPG binding depends on functional TFIIH, we measured UV-induced accumulation of XPG (using a specific antibody) at different time points after local UV-irradiation in various cell lines mutated in the TFIIH helicases XPB or XPD (Table 1) and in wild-type cells. Accumulation of XPG, measured 10 min after local UV-induced DNA damage, was strongly reduced in all TFIIH mutants tested, in comparison to wild-type cells. In contrast, XPC, which binds to DNA damage before TFIIH, accumulated normally in all mutants (Fig. 5). However, some XPG accumulation was observed in all mutant TFIIH cell lines 30 min after local UV damage, except in TTD cells (Fig. 5). These results show that functional TFIIH is required for normal XPG binding. However, several TFIIH mutants still support what seems a slow or limited XPG incorporation in the NER complex.

XPG-EGFP is transiently immobilized by UV-damaged DNA In addition to the binding (i.e. pre steady-state) kinetics of XPG-EGFP into the nascent NER complex, we have analyzed the steady-state kinetics and the nuclear redistribution after UV damage. Different UV-C doses have been employed, i.e. 2, 4, 8, and 16 J/m2. Whole cell exposure to UV-light induces a uniform distribution of DNA lesions in the nucleus. This did not result in a detectable redistribution of XPG-EGFP, in contrast to the study by Park et al. that reported the formation of XPG foci upon UV irradiation in fixed cells (Park et al., 1996). However, these XPG foci could be the result of the fixation procedure. FRAP analyses performed between 10 and 45 min after UV- treatment (when incorporation of XPG is in steady-state), showed incomplete fluorescence recovery in human fibroblasts and in CHO cells within the time frame of 8 seconds after photobleaching (Fig. 6A and 6B, compare to untreated cells). This indicates that maximally 20-30% of the XPG-EGFP molecules are immobile. The mobility of unbound XPG-EGFP did not change upon UV irradiation, as shown by the recovery plots of Fig. 6C and 6D. This demonstrates that the XPG molecules that do not participate in NER have the same mobility and therefore the same molecular size before and after UV irradiation. Similar to other NER factors (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003) we conclude that the observed immobilization of XPG-EGFP reflects its incorporation in the NER complex.

Despite exhibiting similar total cellular fluorescence intensities (representing the expression levels of XPG-EGFP), human fibroblasts showed less XPG-EGFP immobilization at a NER-saturating UV dose than CHO cells, i.e. 20% and 30%, respectively (Fig. 6E and 6F). Furthermore, a quantitatively different response to UV- dose was observed for both cell types. While in human cells a larger fraction of the XPG-EGFP molecules was immobilized at low UV-doses (saturating at 8 J/m2) CHO cells show a significant increase in immobilized XPG-EGFP up to 16 J/m2 (Fig. 6E and 6F). This shows that the fraction of immobilized XPG-EGFP of the total cellular amount of XPG is different in both cell types. This is probably due to the fact that the

53 Chapter 3 concentration of other NER factors is different. If cells were cultured at 27°C instead of at 37°C, a significantly larger fraction of XPG-EGFP was immobilized at the same UV dose in both cell lines (Fig. 6E and 6F), indicating that at any given time point more XPG-EGFP molecules participate in NER events. Combined FRAP and FLIP at 27°C instead of 37°C, confirmed a decrease in mobility of XPG-EGFP after UV-irradiation (Fig 2D). This implies that the dissociation Figure 7. FLIP analysis of locally UV-damaged of XPG-EGFP from the NER complex areas in the nucleus. (A) A strip opposite a locally (presumably after dual incision) is damaged area in the nucleus is bleached and the temperature-dependent, resulting in a redistribution of bleached and unbleached XPG- EGFP is monitored in time. (B) FLIP curve of the slower dissociation rate at lower locally damaged nucleus. The relative difference temperature and thus increased between redistribution in the damaged versus the immobilization. In addition to a role in non-damaged area is plotted in time. Error bars depict the standard error of the mean. NER, XPG has been shown to be involved in base excision repair of oxidative DNA lesions in vitro (Klungland et al., 1999). To investigate a possible role of XPG in BER in vivo we treated cells with ionizing radiation or paraquat. Both procedures induce oxidative lesions that are removed by BER. After treatment with these agents, we did not observed increased immobilization of XPG-EGFP (data not shown), suggesting that XPG does not play a major role in BER. However, we cannot rule out that the number of lesions introduced by these procedures is too low to detect changes in XPG-EGFP immobilization or that the kinetics of XPG in BER are different and do not result in detectable immobilization of XPG. Besides a role in BER, it has been suggested that XPG associates with transcription bubbles containing stalled RNAPII molecules together with TFIIH and CSB (Sarker et al., 2005). In addition, the S. cerevisiae XPG homolog Rad 2 has been shown to be required for RNAPII activity (Lee et al., 2002). We previously showed that the mobility of TFIIH, which is involved in RNAPI and RNAPII activity, is affected by treatment with transcription inhibitors (e.g. 5,6-dichloro-1-D-ribofuranosyl benzimid azole or DRB (Hoogstraten et al., 2002; Iben et al., 2002)). However, we did not observe any effect on XPG-EGFP mobility after treatment with DRB (data not shown) and were thus not able to confirm a role of XPG in transcription bubbles in vivo (Sarker et al., 2005). It cannot be excluded that the interaction of XPG with transcription bubbles is too transient or involves only a very small fraction of molecules escaping detection.

54 XPG dynamics

The residence time of XPG-EGFP in the NER complex is in the order of minutes To determine the dissociation kinetics of XPG from NER complexes we applied a FRAP variant on locally damaged cells. Briefly, an elongated area distant from the local damage is bleached. Subsequently, the fluorescence redistribution is monitored (Fig. 7A). The time required to re-establish the pre-irradiation distribution of XPG- EGFP is a measure for the koff of XPG molecules from NER complexes. A new equilibrium between bleached and non-bleached molecules was reached within 3 to 4 minutes with a t1/2 of 50s. (Fig. 7B),reflecting the dissociation rate of XPG from NER complexes. This value is similar to the measured dissociation kinetics of other components of the NER complex (Hoogstraten et al., 2002; Houtsmuller et al., 1999).

Discussion

The endonuclease XPG is a multi-functional nuclear protein. It plays a central role in nucleotide excision repair (NER) of helix-distorting DNA damage and is thought to be involved in transcription, transcription-coupled repair of non-helix distorting DNA lesions and in base excision repair (BER) (Klungland et al., 1999; Lee et al., 2002; Sarker et al., 2005). In the NER complex XPG carries out the incision at the 3’ side of the damage and stabilises the protein complex on the locally unwound DNA (de Laat et al., 1999). In this study we present a comprehensive analysis of the dynamic behaviour of XPG inside the nuclei of living CHO cells and human fibroblasts that do not contain functional endogenous XPG and that stably express human XPG-EGFP at levels similar to endogenous XPG in wild type cells (Fig. 1). Since these cells show normal UV-resistance, the XPG-EGFP is fully functional.

XPG only interacts with NER components on damaged DNA Evidence has been presented that XPG associates with TFIIH in vitro, among others by co-immunoprecipitation studies (Araujo et al., 2001; Mu et al., 1995). Our FRAP measurements on cells expressing functional XPG-EGFP show that in undamaged living cells the protein is not associated with other nuclear proteins, because the observed apparent diffusion constant (5 ± 1 µm2/s) is what is expected for a monomeric XPG protein (180 kDa). Importantly, also after UV damage the XPG molecules that are not engaged in DNA repair show the same in vivo mobility, indicating that XPG only interacts with other nuclear components if it binds to NER complexes that assemble onto damaged DNA (Fig 6). This is supported by the finding that other NER proteins show apparent diffusion rates that are different from what is observed here for XPG-EGFP (Giglia-Mari et al., 2006; Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003). Moreover, the mobility of TFIIH is different at 27°C and 37°C, whereas that of XPG-EGFP is temperature independent (Fig. 2). Also the kinetics of incorporation of these two proteins into the NER complex is different (t1/2 of 200 s and 110 s, respectively (Fig. 3 and (Moné et al., 2004))). The observation that

55 Chapter 3 endonuclease XPG does not associate with nuclear components except in the DNA damage-induced NER complex, supports the notion that this protein is only involved in DNA repair.

The dynamics of XPG engagement in NER After UV-induced DNA damage XPG-EGFP is incorporated into the NER complex with a t1/2 of incorporation of about 200 s at 37°C (Fig 3). CHO cells and human fibroblasts show the same assembly rate. This rate of incorporation is somewhat lower than that of XPC and TFIIH (t1/2 is 100 and 120 s, respectively (Moné et al., 2004; Politi et al., 2005)) and significantly lower than that of 5’ endonuclease ERCC1/XPF (t1/2 is 65 s (Moné et al., 2004)). A quantitative model has been proposed that is able to at least partly explain these differences in rates of incorporation (Politi et al., 2005). After about 5 min, XPG incorporation into NER complexes reaches a steady-state. FRAP experiments show that under these conditions XPG continuously exchanges between the bound and free diffusing state. Under these conditions, the protein dissociates rapidly within 3-4 min with a t1/2 of 50s (Fig. 7). This is similar to what has been found for TFIIH and ERCC1/XPF (t1/2 ~ 50s) and faster than dissociation of XPA (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003). Interestingly, XPC dissociation is considerably faster (t1/2 ~ 25s), probably because it leaves the NER complex before repair is complete (Hoogstraten and Vermeulen, unpublished results, (Politi et al., 2005)). Under steady-state condition, at the highest UV doses used here (16 J/m2), maximally 30% of the XPG-EGFP molecules are associated with a NER complex and therefore engaged in NER (Fig. 6). These results support a model in which the pre-incision NER complex assembles from its individuals components on a time scale of minutes, remains intact for 3 to 4 min (maybe except for XPC) during which the actual repair takes place and subsequently dissociates allowing its components to reassemble on another lesion.

Although the dynamic behaviour of XPG is largely the same in CHO cells and in human fibroblasts, a difference is observed in the degree of XPG immobilization under steady state conditions at high UV dose (16 J/m2). In CHO cells an almost two-fold larger fraction of the XPG-EGFP molecules becomes engaged in NER than observed for human fibroblasts (at 37°C 20% and 30%, respectively; Fig 6). The simplest explanation is that the expression levels of XPG and/or other NER proteins differ in the two cell types. Alternatively, the endogenous truncated, non-functional XPG protein that is present in the human fibroblasts may compete with the functioning of XPG- EGFP, resulting in a lower immobilized fraction. The XPG mutation in CHO UV135 cells is unknown but can be considered a null mutation, since XPG mRNA can hardly be detected in these cells (MacInnes et al., 1993).

56 XPG dynamics

DDB2 (p48) does not affect the rate of XPG incorporation kinetics The kinetics of incorporation of XPG into the NER complex and its dissociation rate from NER complexes are the same in CHO cells and in human fibroblasts (Fig. 4, data not shown). This is remarkable, since CHO cells lack functional DDB2 (p48), which is a subunit of the UV-DDB complex that is thought to enhance the association of the damage-recognition protein XPC with DNA lesions, in particular pyrimidine dimers (Moser et al., 2005; Sugasawa et al., 2005; Tang et al., 2000). Since XPC binding precedes incorporation of XPG into the NER complex, it was expected that XPG binding in CHO cells would be slower that in human fibroblasts, which contain endogenous DDB2. Expression of DDB2 in CHO cells did not result in accelerated binding of XPG to UV-induced DNA damage (Fig. 4). These results indicate that UV-DDB does not significantly increase the rate of binding of XPG to UV-damaged DNA.

Recruitment of XPG requires functional TFIIH Previous experiments showed that the incorporation of core NER factors into the NER complex occurs in a specific sequence (Rademakers et al., 2003; Volker et al., 2001). However, the precise timing of XPG incorporation could not be established unambiguously. Here we show that incorporation of XPG into the NER complex is temperature-dependent (Fig 3). The same has been found for ERCC1/XPF, whereas binding of TFIIH and XPC are temperature-independent (Moné et al., 2004; M.S. Luijsterburg and R. Van Driel, unpublished data). This was interpreted that binding of ERCC1 requires an enzyme activity, i.e. the helicase activity of the TFIIH subunits XPB and XPD (Moné et al., 2004). Therefore, our data suggests that also XPG binding requires TFIIH helicase activity, indicating that TFIIH binding must precede XPG incorporation. Studies in cell lines that have a mutated TFIIH gene show that impairment of TFIIH function severely affects XPG incorporation into the NER complex (Fig. 5, Table 1).

Comparison of the effect of different TFIIH mutation indicates which parts of the TFIIH molecule are important for XPG binding. XP/TTD cells (Stefanini et al., 1993b; Vermeulen et al., 2000; Vermeulen et al., 2001) that carry a C-terminal R722W substitution in XPD (Stefanini et al., 1993a), exhibit the most severe reduction of XPG recruitment. This suggests that XPD plays an important role in the recruitment of XPG to sites of UV damage in vivo. Recent findings demonstrate that the stability of the TFIIH complex is severely affected in TTD cells (D. Hoogstraten and W. Vermeulen, unpublished data; Botta et al., 2002; Giglia-Mari et al., 2004). Therefore, it is conceivable that impaired recruitment of XPG in TTD cells is due to the compromised stability, rather than a direct interaction of XPG with the C-terminus of XPD. A recent study demonstrated that phosphorylation of S751 of XPB controls the 5’ incision by ERCC1/XPF, whereas the 3’ incision by XPG is unaffected (Coin et al., 2004).

57 Chapter 3

Accordingly, our experiments show that XPG binding is only moderately affected in the XPB mutant, which has a truncated C-terminal domain, lacking the serine 751 residue (Fig. 5). This suggests that another part of the TFIIH complex controls the 3’ incision by XPG, possibly the N-terminal PH-fold of p62, which has been shown to interact directly with XPG (Gervais et al., 2004). Nonetheless, our results unambiguously show that stable recruitment of XPG to the pre-incision complex depends on functional TFIIH.

Summarising, our results unfold a consistent and simple picture for the dynamic behaviour of XPG in living CHO cells and human fibroblasts. The protein diffuses freely as a monomer, not showing any prominent interactions other than the nascent NER complex that is formed in UV-damaged cells after binding of XPC and TFIIH. The in vivo dynamics of the XPG protein are remarkably similar in human cells and Chinese hamster cells, showing that major differences in genetic background hardly affect XPG behaviour.

Materials and Methods

Cell lines. Cell lines used in this study were simian virus 40 (SV-40)-immortalized human fibroblasts MRC5 (NER- proficient), XPCS1RO (XPG-deficient); HeLa cells (NER-proficient), CHO AA8 (NER-proficient), CHO UV135 (XPG-deficient), 3T3 cells (inducible EGFP expression) and the primary human fibroblasts C5RO (NER-proficient), XPCS1BA (XPB-deficient), XP131MA (XPB-deficient), XP6BE (XPD-deficient), XPCS2 (XPD-deficient) and TTD1BEL (XPD-deficient). All cell lines were cultured in a 1:1 mixture of DMEM/Ham’s F10 medium containing Ultra-Glutamine (Cambrex Corporation, New Jersey, USA) supplemented with antibiotics and 10% FCS at 37°C in an atmosphere of 5% CO2.

Generation of cells expressing XPG-EGFP and construction of DDB2-mCherry. EGFP-tagged XPG was generated by in-frame ligation of full length human XPG cDNA into a EGFP N1 vector (Clontech Laboratories, California, USA)). This resulted in a fusion gene under control of a cytome- galovirus promoter encoding an XPG-EGFP hybrid polypeptide. The fusion gene was expressed in the XPG- deficient CHO cell line UV135 and the XPG-deficient SV40-transformed human fibroblast cell line, XPCS1RO-SV (Ellison et al., 1998). After subsequent rounds of selection on the presence of the neomycin resistance gene (by G418-resistance selection) and UV-irradiation (to select for functional XPG expression) stable expressing clones were isolated for each of the cell types. The pDDB2-eYFP (Moser et al., 2005) plasmid was digested with AgeI and BsrGI in order to replace the eYFP for mCherry (Shaner et al., 2004) to yield pDDB2-mCherry

Immunoblot analysis and UV survival. Cell extracts were generated by sonication, separated by sodium dodecyl sulfate polyacrylamide gel (8%) electrophoresis and transferred to nitrocellulose membranes. Expression of the fusion protein was analyzed by immunoblotting with a mouse monoclonal anti-XPG antibody (IB5 T.C., 1:100 in PBS/0.05% Tween-20; a gift from Dr. J.M. Egly), followed by a secondary antibody (goat anti-mouse conjugated with alkaline phosphatase (Biosource International, California, USA) and detection using bromo-4-chloro-3-indolyl phosphate (BCIP) and nitro blue tetrazolium (NBT). As loading control, mouse monoclonal anti-PCNA antibody (Dako, Glostrup, Denmark) at a dilution of 1:1000 was used. For UV-survival experiments, cells were exposed to different UV doses 2 days after seeding. Survival was determined 3 days after UV-irradiation by measuring cell proliferation with the aid of [3H] thymidine pulse-labeling at 37°C, as described previously

58 XPG dynamics

(Hamel et al., 1996).

Immunofluorescence. Cells were grown on 24 mm glass coverslips and fixed with 3% paraformaldehyde in PBS with 0.3% Triton X-100 for 20 min at room temperature (RT). Coverslips were washed three times for 10 min with phosphate-buffered saline (PBS) containing 0.1% Triton X-100, and were subsequently incubated for 1 h with PBS containing 1% BSA. Cells were incubated at RT with the primary antibody for 1.5 - 2h in a moist chamber. Subsequently, coverslips were washed three times for 10 min with PBS-Triton X-100 and 5 min with PBS 1% BSA. Incubation with the secondary antibody was for 30 min – 1 h at RT (dark chamber) followed by extensive washing with PBS 1% BSA and finally PBS. Samples were embedded in Vectashield (Vector Laboratories, California, USA) mounting medium containing 0.1 mg of DAPI (4’-6’-diamidino- 2-phenylindole) per ml. Primary antibodies used for immunolabeling were: mouse monoclonal antibody against XPG (8H7, Lab Vision Fremont, California, united States, 1:2000), and affinity-purified rabbit monoclonal antibody against XPC (Ng et al., 2003). Secondary antibody: cy3-conjugated goat anti-mouse antiserum and FITC-conjugated anti-rabbit antiserum (Both from Jackson ImmunoResearch Laboratories, West Grove, Pennsylvania, USA). All antibodies were diluted in PBS containing 0.15% glycine and 0.5% bovine serum albumin. Fluorescence microscopy images were obtained with a Leica DMRBE microscope (Leica Microsystems, Wetzlar, Germany) equipped with epifluorescence optics, a PL-FLUOTAR 100x, 1.3- numerical aperture oil immersion lens and a Hamamatsu (Hamamatsu Photonics, Hamamatsu City, Japan) dual mode cooled CCD camera.

Confocal imaging. Digital images of EGFP-expressing living cells were obtained using a Zeiss LSM 410 microscope equipped with a 60-mW Ar laser (488 nm) and a 40x, 1.3-numerical aperture oil immersion lens and a Zeiss LSM 510 equipped with a 60-mWAr laser (488 nm) and a 40x, 1.2-n.a., or 63x Planapochromat, n.a. 1.4, oil immersion lens (Zeiss, Oberkochen, Germany). Both microscopes were equipped with an objective heater. Unless stated otherwise, living cells were examined at 37°C.

UV irradiation. For induction of global UV DNA damage, cultured cells on coverslips were rinsed with PBS and irradiated with a Phillips TUV lamp (254 nm) at a dose rate of ~0.8 J/m2/s. To induce local UV-damage, cells were UV- irradiated through an polycarbonate filter (Millipore Billerica, Massachusetts, USA) with pores of 5 µm diameter, as described previously (Moné et al., 2001; Volker et al., 2001). At indicated time points after filter removal the cells were either microscopically examined, or fixed with 2% paraformaldehyde and further processed for immuno-histochemistry as described above. For kinetic measurements on locally UV- damaged cells that express XPG-EGFP were grown to confluency in glass bottom dishes (MatTek, Ashland, Massachusetts, USA). Local UV-irradiation was performed as described previously (Moné et al., 2004). Briefly, a petridish was filled with microscopy medium (137 mM NaCl, 5,4 mM KCl, 1,8 mM CaCl2, 0,8 mM MgSO4, 20 mM D-glucose and 20 mM HEPES) and a small piece of Alcian blue-coated filter (5 µm pores) was sunk onto the cells. A glass ring was carefully placed on top of the filter after which the petridish was sealed with a lid containing a quartz window. The cells were transferred to an Zeiss Axiovert 200 M microscope with a 37o incubator and an objective heater, to ensure the appropriate temperature for this live cell experiment. Subsequently, irradiation was performed using a homemade box, containing four UV lamps (Philips TUV 9W PL-S) above the microscope stage. The UV dose rate was measured to be 3 W.m-2 at 254 nm. Cells were irradiated for 39 s, resulting in a UV dose of 100 J.m-2

FRAP and fluorescence loss in photobleaching (FLIP). For all experiments cells were seeded onto 24 mm glass coverslips three days prior to the experiments. FRAP experiments. Using a Zeiss LSM510 METAconfocal microscope, equipped with a 60 mW Argon laser and a 40x oil immersion lens (1.2 n.a.), mobility measurements were performed by FRAP analysis at high time resolution (Strip-FRAP; 29, modified). A strip spanning the nucleus was photo-bleached for 20 ms at 100% laser intensity (120-160µW, argon laser at 488nm). Recovery of fluorescence within the strip was monitored with 20 ms intervals at low laser intensity (450-750nW) to avoid photobleaching by the probe beam. Measurements were performed at 37˚C, using a heated stage with feedback temperature control. Raw data were corrected for background and fluctuations in the monitoring laser power.

59 Chapter 3

Fluorescence Loss in Photobleaching (FLIP). To determine the residence time of XPG at locally UV-irradiated areas local UV irradiation was applied as described above. Using a Zeiss LSM 410 microscope equipped with a 60 mW Argon laser (488 nm) and a 40x, 1.3 n.a. oil immersion lens, a strip was bleached (at 100% 488 nm) for 5 s near the edge of the nucleus opposite to the local damage site. Redistribution of fluorescence was monitored over time (at 488 nm). Evaluation was performed by comparing the loss of fluorescence of bound protein over time (in local damaged area) versus non-bound protein (outside the damaged area) of EGFP-tagged protein.

Combined FLIP-FRAP analysis. Using a Zeiss LSM510 META confocal microscope, equipped with a 60- mW Argon laser and a 40x oil immersion lens (1.2 n.a.), a 2 µm (30-pixel) strip spanning the cell nucleus at one pole was bleached for 1s at a laser power of 120-160 µW. Redistribution of fluorescence throughout the nucleus was recorded at low laser power (1.6-1.9 µW), keeping monitoring bleaching to a minimum (<5 %). We compared the difference between the fluorescence in the bleached and the unbleached area (at a distance of 150 pixels = 10.2 µm) of the nucleus, and plotted the fluorescence values against time. Unless otherwise specified, measurements were performed at 37˚C, using a heated stage with feedback temperature control. At least 9 independent measurements were averaged to form a single mobility curve. Redistribution of fluorescence was corrected for lateral cell movement. Rotating cells, or cells moving out of focus were excluded from evaluation.

FRAP analysis. For analysis of FRAP data, FRAP curves were normalized to pre-bleach values and the best fitting curve (least squares) was picked from a large set of computer simulated FRAP curves in which three parameters representing mobility properties were varied: diffusion rate (ranging from 0.04 to 25 µm2/s), immobile fraction (0, 10, 20, 30, 40, 50 %) and time spent in immobile states, ranging from very short residence times (0.02, 0.04, 0.08, …, 1 s) to relatively long residence times (2, 4, 8, 16, 32, 64, 128, ∞ s). Monte Carlo computer simulations used to generate FRAP curves were based on a model of diffusion in an ellipsoid volume representing the cell nucleus, and simple binding kinetics representing binding to immobile elements in the cell nucleus. Simulations were performed at unit time steps corresponding to the experimental sample rate of 21 ms. Diffusion was simulated by each step deriving novel positions M(x+ dx, y+ dy, z+ dz) for all mobile molecules M(x, y, z), where dx = G(r1), dy = G(r2) and dz= G(r3), ri is a random number (0 ≤ ri ≤ 1) chosen from a uniform distribution, and G(ri) is an inversed cumulative Gaussian distribution with µ = 0 and σ2 = 6Dt, where D is the diffusion coefficient and t is time measured in unit time steps. Immobilization was based on simple binding kinetics described by: kon/koff = Fimm / (1 - Fimm), where Fimm is the relative number of immobile molecules. The chance for each particle to become immobilized (representing chromatin-binding) was defined as Pimmobilise = kon = koff .Fimm / (1 - Fimm), where koff = 1/ timm, and timm is the average time spent in immobile complexes measured in unit time steps; the chance to release was Pmobilise = koff = 1/ timm. The FRAP procedure was simulated on the basis of an experimentally derived 3D laser intensity profile providing a chance based on 3D position for each molecule to get bleached during simulation of the bleach pulse.

Assembly at local damaged sites. For analyzing the dynamics of NER complex assembly cells were kept on an Axiovert 200M microscope stage at the appropriate temperature, using a temperature-controlled microscope chamber. The objective (Zeiss Apochromat 100X) was temperature-controlled with an objective heater. One image was taken to determine the position and the GFP fluorescence intensity of the cells (monochromator at 470 nm, bandwidth 20 nm). A reflection image of the filter was obtained moving up in the z-direction. Images of the cells and the filter were overlaid to determine which nuclei are located under a filter pore. The distance between cells and filter was measured with a Piezo electrical element and had to be less than 7 µm to obtain a well-defined damaged area. CHO cells were transiently transfected with DDB2-mCherry with lipofectamine 2000 (Invi- trogen, Breda, the Netherlands) according to the manufacturer's instructions. The cells were irradiated (100 J.m-2) and images were collected at 20 s interval for 30 minutes to allow EGFP accumulation in the locally damaged area to reach a plateau level. DDB2-mCherry accumulation was monitored with 550 ± 20 nm. The accumulation of XPG-EGFP at sites of local damage was quantified with Object-Image software (Vischer et al., 1999). A macro was written to determine the centre of gravity of the fluorescent spot at every time point. Next, the average fluorescence intensity was measured in a region of 20 µm2 around the centre of gravity for every time point. The average intensity of the entire nucleus, except the damaged area, was also measured, representing the unbound protein pool. The EGFP signal in the undamaged area was subtracted from

60 XPG dynamics

the damaged area. The resulting value represents the NER-related bound protein fraction. All values were corrected for photo bleaching. Time courses were normalized with respect to the plateau level. Start of the UV irradiation was defined as t=0. Assembly curves were normalized to 1 or to the bound fraction calculated by the equation: Bound (%) = (Ispot-Ioutspot)* pixelsspot / (Inucleus-Ibackground)* pixelsnucleus, where Ispot and Ioutspot are the average pixel intensities inside the damaged spot and outside the spot, respectively. Inucleus is the average pixel intensity of the nucleus, including the spot and Ibackground is the average pixel intensity outside the cell.

Table 1 Human cell-lines used for local damage induction

Cell strain TFIIH mutation Affected function Syndrome Reference

C5RO none wild type

(Vermeulen et al., XPCS1BA F99S in XPB unknown mild XP/CS 1994) frame-shift 742 in (Evans et al., XP131MA helicase activity XP/CS XPB 1997) (Protic-Sabljic et XP6BE R638W in XPD p44 interaction XP al., 1986) (Vermeulen et al., XPCS2 G602D in XPD helicase activity XP 1991) (Taylor et al., TTD1BEL R722W in XPD p44 interaction TTD 1997)

Acknowledgement This work was supported by grants of the Netherlands Organisation for Scientific Research (NOW): NWO-CW 700.98.302 (AZ); ZonMW 912-03-012 (MSL), 917-46- 364 (WV) and 901-01-229. The DDB2-EYFP plasmid was kindly provided by Dr. L.H. Mullenders and the mCherry cDNA was kindly provided by R.Y. Tsien. The authors would like to thank A. Theil and N. Wijgers for technical assistance, N.O.E. Vischer (Center for advanced microscopy (CAM)/UvA) for valuable assistance with data analysis. Dr. J. Goedhart (CAM) for critical reading of the manuscript and Drs. E.M.M. Manders and T.W.J. Gadella (CAM) for support.

61 62 Chapter 4 Dynamic In Vivo Interaction of DDB2 E3 Ubiquitin Ligase with UV-damaged DNA is Independent of Damage-Recognition Protein XPC

Martijn S. Luijsterburg , Joachim Goedhart, Jill Moser, Hanneke Kool, Bart Geverts, Adriaan B. Houtsmuller, Leon H.F. Mullenders, Wim Vermeulen and Roel van Driel

Adapted from J. Cell. Sc. 120, 2706-16, August 2007

63 Chapter 4

Abstract Damaged DNA-binding protein 2 (DDB2) has a high affinity for UV-damaged DNA and has been implicated in the initial steps of global genome nucleotide excision repair (NER) in mammals. DDB2 binds to CUL4A and forms an E3 ubiquitin ligase. In this study, we have analyzed the properties of DDB2 and CUL4A in vivo. The majority of DDB2 and CUL4A diffuse in the nucleus with a diffusion rate consistent with a high molecular weight complex. Essentially all DDB2 binds to UV-induced DNA damage, where each molecule resides for ~2 minutes. After the induction of DNA damage, DDB2 is proteolytically degraded with a half-life that is two orders of magnitude larger than its residence time on a DNA lesion. This indicates that binding to damaged DNA is not the primary trigger for DDB2 breakdown. The bulk of DDB2 binds to and dissociates from DNA lesions independently of damage recognition protein XPC. Moreover, the DDB2-containing E3 ubiquitin ligase is bound to many more damaged sites than XPC, suggesting that there is little physical interaction between the two proteins. We propose a scenario in which DDB2 prepares UV-damaged chromatin for assembly of the NER complex.

Introduction Genome integrity is continuously challenged by various sources that potentially damage DNA. Short-wavelength UV light introduces DNA injuries, such as 6-4 photoproducts (6-4 PP) and cyclobutane pyrimidine dimers (CPD), both which can interfere with transcription and replication. The mammalian cell utilizes nucleotide excision repair to remove UV-induced DNA damage and various other bulky DNA lesions from the genome (de Laat et al., 1999; Hoeijmakers, 2001). Two different modes of NER exist: transcription-coupled NER (TC-NER) and global genome NER (GG-NER). TC-NER removes lesions from the transcribed strand of active genes, whereas GG-NER repairs damage at any other position in the genome (de Laat et al., 1999). More than 20 gene products are involved in mammalian NER that employs a dual incision mechanism to remove lesions (Aboussekhra et al., 1995). A crucial step in the initiation of GG-NER is the detection of DNA lesions. Although the mechanism of damage detection in chromatin is not well understood, various studies have identified the XPC-RAD23B-CEN2 complex as the principal initiator of GG-NER (Araki et al., 2001; Sugasawa et al., 1998; Sugasawa et al., 2001; Volker et al., 2001). Detection of CPDs by XPC is inefficient and it has been suggested that the UV- damaged DNA binding (UV-DDB) complex plays a crucial role in CPD detection and repair. UV-DDB consists of DDB1 (p127) and DDB2 (p48) (Dualan et al., 1995; Takao et al., 1993), and mutations in the DDB2 subunit are responsible for the XP-E phenotype (Nichols et al., 2000; Rapic-Otrin et al., 2003). Cells lacking functional DDB2 such as rodent and human XP-E cells, have a deficiency in the removal of CPDs and delayed removal of 6-4 PPs, particularly at low UV doses (Hwang et al., 1999; Moser et al., 2005; Tang et al., 2000). Transfection of DDB2 cDNA in hamster cells

64 DDB2 dynamics restores the binding activity of UV-DDB to damaged DNA, showing that DDB2 rather than DDB1 is responsible for binding to damaged DNA (Hwang et al., 1999; Li et al., 2006; Tang et al., 2000). The complete inactivation of DDB2 expression in mice results in a significant increase in the formation of malignant tumors (Yoon et al., 2005), while over-expression of DDB2 protects mice from the carcinogenic effects of UV-B irradiation (Alekseev et al., 2005), illustrating the tumor-preventing property of DDB2.

Recently, UV-DDB was implicated in 6-4 PP repair in vivo (Alekseev et al., 2005; Moser et al., 2005), which supports results of several in vitro assays that demonstrated a higher affinity of UV-DDB for 6-4 PPs compared to CPDs (Fujiwara et al., 1999; Reardon et al., 1993; Treiber et al., 1992). UV-DDB has a 500,000-fold preference for damaged over non-damaged DNA (Hwang and Chu, 1993), which is significantly higher than the selectivity of XPC for damaged DNA (3,000-fold preference) (Sugasawa et al., 1998). UV-DDB appears to bind to DNA lesions in the absence of functional XPC protein (Fitch et al., 2003; Wakasugi et al., 2001), in contrast to the NER components XPA, TFIIH, XPG and ERCC1-XPF, which all require XPC for binding to damaged DNA (Volker et al., 2001). Although much less efficiently, XPC binds to 6-4 PPs in XP-E cells, indicating that UV-DDB is not an absolute requisite for XPC binding to this particular type of lesion (Moser et al., 2005). DDB2 is crucial for the recognition and repair of CPDs and over-expression of DDB2 leads to enhanced recruitment of XPC to CPDs (Fitch et al., 2003). These observations strengthen the hypothesis that UV-DDB facilitates and/or stabilizes the binding of XPC to UV- induced photoproducts.

Biochemical evidence suggests that UV-DDB is part of an E3 ub ligase complex that contains Cullin 4A (CUL4A) (Shiyanov et al., 1999b) and the COP9 signalosome, which is a negative regulator of the ub ligase activity of the E3 DDB2 complex (Grois- man et al., 2003). The ub ligase activity of the E3 DDB2 complex is transiently activated by UV irradiation and several substrates for ubiquitination have been identified, including DDB2 itself, XPC and histones H2A, H3 and H4 (Groisman et al., 2003; Kapetanaki et al., 2006; Sugasawa et al., 2005; Wang et al., 2006). It is currently believed that DDB2 recruits XPC to lesions after which both DDB2 and XPC are ubiquitinated (Sugasawa et al., 2005). Ubiquitinated DDB2 is proteolytically degraded (Chen et al., 2001; Nag et al., 2001; Rapic-Otrin et al., 2002), whereas the reversible ubiquitination of XPC increases its affinity for DNA (Sugasawa et al., 2005). To investigate the interplay between DDB2 and XPC in vivo, we have analyzed the properties of fluorescently labeled DDB2 in wild-type and in XPC deficient cells. Our results indicate that the bulk of DDB2 interacts with lesions independently of XPC and that there is little interaction between the two recognition proteins on damaged DNA.

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Results

Generation of cell-lines expressing YFP-tagged DDB2 and GFP-tagged CUL4A To study the cellular distribution and dynamics of DDB2 we tagged the protein with enhanced yellow fluorescent protein (EYFP), which was fused to the C-terminus of murine DDB2 (Figure 1B) resulting in a DDB2-EYFP fusion protein that was stably expressed in human fibroblasts (MRC5-SV) (Moser et al., 2005). EYFP was also fused to the N-terminus of DDB2 and stably expressed in the same cells. Both fusion proteins were found predominantly in the nucleus (Fig. 1A). Western blot analysis using anti- DDB2 antibodies showed that DDB2-EYFP migrates in SDS-PAGE with a mobility corresponding to the expected size of the full-length fusion protein (~75 kDa; Fig 1C; Dualan et al., 1995). The fluorescently tagged DDB2 protein binds to UV-damaged DNA (Moser et al., 2005) and is proteolytically degraded after UV irradiation as shown by western blot analysis of whole cell extracts (Fig. 1C). This indicates that the fusion protein exhibits wild-type behavior. To quantify the expression level of DDB2-EYFP, we compared the fluorescence of a solution of EYFP protein with a known concentration with the fluorescence of cells expressing DDB2-EYFP. The average concentration of DDB2-EYFP in living cells was 0.7 ± 0.1 µM (data not shown), which corresponds to ~1,2.105 molecules in a 300 µm3 nucleus. The number of endogenous DDB2 molecules per cell is ~1.105, indicating that DDB2-EYFP is expressed at levels comparable to endogenous DDB2 (Keeney et al., 1993). Moreover, western blot analysis using a polyclonal antibody against DDB2 (Wang et al., 2005b) demonstrated that DDB2-EYFP is expressed at about the same level as endogenous DDB2 in the same cells (Fig 1C). DDB2-EYFP was also expressed in XPC-deficient and XPA- deficient human fibroblasts (XP20MA-SV and XP12RO-SV, respectively) and displayed a similar cellular distribution as observed in wild-type cells (data not shown). In addition to DDB2, CUL4A was fused to EGFP and expressed in HeLa cells, where it was found predominantly in the nucleus (Fig 1A, B). The cellular distribution of endogenous CUL4A and DDB2 was mainly nuclear (Fig 1A), showing that the localization of the fluorescent fusion proteins reflects that of the endogenous proteins.

DDB2 and CUL4A have the same mobility in undamaged cells Analysis of purified proteins from insect cells and whole cell lysates suggested that DDB2 exists in at least four different assembly states in vitro: monomeric DDB2, heterodimeric DDB1-DDB2 and as a protein complex containing either ROC1/CUL4A or ROC1/CUL4A with COP9 (Kulaksiz et al., 2005). However, DDB2 isolated from cells is co-purified with DDB1, CUL4A and all subunits of COP9 (Groisman et al., 2003). To investigate whether DDB2 is part of different protein complexes in vivo we employed fluorescence recovery after photobleaching (FRAP) (Houtsmuller and Vermeulen, 2001; Rabut and Ellenberg, 2005), in which a small strip (2 µm width) spanning the nucleus was bleached and the fluorescence recovery in the strip was

66 DDB2 dynamics

Figure 1. Expression of DDB2- EYFP and EGFP-CUL4A. (A) Localization of DDB2 and CUL4A. The upper panel in (A) shows the distribution of DDB2- EYFP in an MRC5 cell, the middle image shows the distribution of EGFP-CUL4A in a HeLa cell and the lower image shows the localization of endogenous DDB2 and CUL4A detected with specific antibodies in primary human cells (VH10). (B) Repre- sentation of the DNA constructs encoding fluorescently tagged DDB2 and CUL4A. (C) Western blot analysis of MRC5-SV40 cells expressing DDB2-EYFP. Whole cell extracts (10 µg) of non UV-irradiated cells and UV- irradiated cells (2 and 4 hours after irradiation with 20 J.m-2), were probed with antibodies against DDB2, DDB1 and CUL4A.

Figure 2. FRAP analysis of DDB2 mobility in non-damaged cells (NoDa). (A) Example of a DDB2-EYFP cell in which a strip (2 µm) spanning the nucleus is bleached. (B) Recovery plots of DDB2-EYFP (n = 14) and XPG- EGFP (n = 23), normalized between 0 and post-bleach intensity after full equilibration. (C) Normalized recovery plots of DDB2-EYFP at 37°C and 27°C (n = 12), DDB2(R273H)-EYFP (n = 13) at 37°C and EGFP-CUL4A (n = 11) at 37°C. (D) Recovery of DDB2-EYFP normalized to pre-bleach intensity (red line). Monte Carlo simulation assuming a diffusion constant of 2.4 µm2/s (green line) and the residuals (blue lines at the bottom) that are a measure for the quality of the simulation. (E) Recovery of XPG-EGFP normalized to pre-beach intensity (red line), Monte Carlo simulation assuming a diffusion constant of 4.7 µm2/s (green line) and the residuals (blue lines).

67 Chapter 4 monitored (Fig. 2A). Most DDB2-EYFP was mobile in non UV-irradiated cells monitored at 37°C (Fig. 2B) and similar mobility was measured at 27°C (Figure 2C). Curves generated by Monte Carlo simulations (Farla et al., 2004) were fitted to experimental FRAP curves, yielding an effective diffusion constant (Deff) for DDB2 in undamaged cells of 2.4 ± 0.4 µm2/s (Fig. 2D). To compare the mobility of DDB2 to that of other NER proteins we performed FRAP on cells expressing NER endonuclease XPG-EGFP (Zotter et al., 2006). The rationale to investigate XPG is that the molecular weight of this protein (180 kDa; O'Donovan et al., 1994) is about the same as the molecular weight of heterodimeric DDB1-DDB2 (~175 kDa). The 2 recovery of XPG-EGFP (Fig. 2B,E, Deff = 4.7 ± 1.0 µm /s) was significantly faster than the recovery of DDB2-EYFP. In addition to DDB2, we have tagged the CUL4A protein with EGFP (Fig. 1B) and measured its mobility in non-UV irradiated cells. The FRAP curves of EGFP-CUL4A and DDB2-EYFP at 37°C were similar, showing that both proteins move through the nucleus with the same mobility (Fig. 2C). To estimate the size of the protein complexes in which DDB2 resides, we compared the mobility of DDB2 and CUL4A with that of other GFP-tagged NER factors (TFIIH, ERCC1, XPG and XPA) and free EGFP (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006). For simplicity, we assumed that the shape of the different NER factors is the same and that the mobility of these proteins is determined predominantly by their molecular weight. Based on these assumptions, we estimate that the majority of DDB2 and CUL4A have a diffusion rate consistent with high molecular weight complexes of 500 to 700 kDa (Figure S1). CUL4A was recently reported to be part of several protein complexes, varying in size between 300 and 700 kDa (He et al., 2006), which is in agreement with our estimate. Immuno- precipitation studies have demonstrated that the naturally occurring DDB2 protein with an R273H substitution in the WD40 motif is unable to interact with the E3 ubiquitin ligase core proteins DDB1 and CUL4A (Chen et al., 2001; Rapic-Otrin et al., 2003; Shiyanov et al., 1999a). In contrast to these studies, purified R273H mutant DDB2 forms a complex with DDB1 in vitro (Wittschieben et al., 2005). We generated a YFP- tagged DDB2 (R273H) protein, which was expressed in human HeLa cells. The mobility of DDB2 (R273H) and wild-type DDB2 was similar (Fig 2C), suggesting that mutant DDB2 is part of the DDB-CUL4A-ROC complex in vivo. Although DDB2 can exist in different assembly states in vitro (Kulaksiz et al., 2005), our results indicate that the majority of DDB2 in vivo resides in high molecular weight complexes of at least 500 kDa. Such a molecular weight is consistent with the size of the DDB-CUL4A- ROC (280 kDa) complex with the COP9 signalosome (450 kDa).

DDB2 recruitment to lesions is independent of XPC To investigate how DDB2 associates with damaged DNA in vivo, we analyzed its binding to UV lesions in human fibroblasts that expressed DDB2-EYFP and that were subjected to local UV irradiation (100 J.m-2 through 5 µm pores) with UV-C light (Fig.

68 DDB2 dynamics

3A) (Moné et al., 2004). DDB2-EYFP accumulated at 37°C with a half-time (t1/2) of 40 seconds (Fig. 3D), which is significantly faster than other GFP-tagged NER proteins tested so far. Similar result were obtained with DDB2 tagged with the red fluorescent protein mCherry (Shaner et al., 2004; data not shown). To verify that the fluorescent tag does not affects the binding kinetics of DDB2 we measured accumulation of EYFP-DDB2 (i.e. N-terminally tagged DDB2; Fig. 1B), which displayed similar binding behavior as C-terminally tagged DDB2 (Fig. 3D). To monitor the binding of DDB2 and another NER protein simultaneously, we expressed DDB2- mCherry and XPG-EGFP in XPG deficient rodent cells and measured the binding kinetics of both proteins upon local UV-C irradiation (Zotter et al., 2006). Results show that DDB2 binds considerably faster than XPG (t1/2 of 40s and 200s, respectively) and that the binding kinetics of DDB2 in rodent and human cells is similar (data not shown). To investigate the kinetics of CUL4A recruitment to sites of DNA damage we transiently expressed EGFP-CUL4A in human HeLa cells (Fig. 3B) and exposed cells to local UV-irradiation. Clear accumulation of GFP-CUL4A at UV-irradiated areas was detected. Since CUL4A alone does not bind to damaged DNA, this indicates that the GFP-CUL4A interacts with endogenous DDB2 (Li et al., 2006). The binding kinetics of EGFP-CUL4A (t1/2 of 47seconds) and DDB2-EYFP (t1/2 of 40 seconds) were similar (Fig. 3D), suggesting that they reflect the assembly of the DDB-CUL4A- ROC1 protein complex onto UV-damaged DNA. In agreement, DDB1-mCherry also binds to UV-damaged DNA with similar kinetics (S. Alekseev et al., manuscript in preparation). To investigate whether the association of DDB2 with UV-damaged DNA involves a temperature-dependent step, for instance an enzymatic reaction, we analyzed its binding kinetics at 27°C. The initial slope of the assembly curves at 37°C and 27°C are similar (Fig. 3E), indicating that the association of DDB2 with damaged DNA is not temperature sensitive. Previous NER assembly approaches showed that exogenously expressed DDB2 binds to UV-induced DNA lesions in the absence of functional XPC (Fitch et al., 2003; Li et al., 2006; Wakasugi et al., 2002). In this study, we show that endogenous DDB2 accumulates at sites of localized UV damage in cells lacking XPC, XPA, XPG and XPF (Fig. S2), indicating that the binding of DDB2 does not require any of these NER proteins. Although binding of DDB2 can be detected in several XP cell-lines, it is possible that the kinetics of binding are different in NER deficient cells. To investigate this, we expressed DDB2-EYFP in XP-C cells (XP20MA-SV40; see Rademakers et al., 2003). Similar binding kinetics were measured in XP-C cells and wild-type cells (Fig. 3C, 3D). These findings show that DDB2 rapidly binds to damaged DNA independently of XPC.

Most DDB2 is bound to chromatin after global DNA damage induction The distribution of DDB2-EYFP in non-UV irradiated living fibroblasts is homogeneous although the protein is partially excluded in the nucleoli, possibly due to their very high macromolecular concentration (Fig. 4A-C). To investigate if UV-

69 Chapter 4

Figure 3. Binding kinetics of DDB2 after local UV irradiation. (A) Example of a DDB2-EYFP cell that shows local accumulation after local UV-C irradiation with 100 J/m2 through 5 µm pores at 37°C. (B) Example of a HeLa cells expressing EGFP-CUL4A and (C) an XP-C cell expressing DDB2-EYFP. Both fusion proteins accumulate after local UV-C irradiation through 5 µm pores at 37° (B-C). (D) Quantification of accumulation kinetics of DDB2-EYFP in MRC5 cells (brown line, n = 9), EYFP-DDB2 in MRC5 cells (green line, n = 8), DDB2-EYFP in XP-C cells (red line, n = 3) and EGFP-CUL4A in HeLa cells (blue line, n = 3). Curves were normalized to the plateau value. Time point t=0 is defined as the start of UV irradiation. (E) Quantification of assembly kinetics of DDB2-EYFP at 37°C (n = 9) and 27°C (n = 8). The local accumulation of DDB2-EYFP was measured and plotted as a percentage of the total EYFP fluorescence in the cell nucleus. Error bars depict the SD.

Figure 4. Nuclear distribution of DDB2 after global UV exposure. (A-B) Representative confocal images of DDB2-EYFP cells (C-E) Example of a DDB2-EYFP cell (C) that was transfected with cerulean-histone H2A (D). (E) Merge of the two signals. (K) Line scan of the arrow in E. (F-G) Representative confocal images of DDB2-EYFP cells that were exposed to global UV-C irradiation (16 J/m2). (H-J) Globally UV- irradiated DDB2-EYFP expressing cell (H) that was transfected with cerulean-histone H2A (I). (J) Merge of the two signals. (L) Line scan of the arrow in J. (M) Example of a cell expressing DDB2-EYFP, that was globally UV-C irradiated (16 J/m2) in metaphase and monitored until cell division was completed.

70 DDB2 dynamics irradiation results in nuclear redistribution of DDB2 we globally irradiated cells (16 J.m-2), inducing a uniform distribution of damage throughout the nucleus. Confocal microscopy showed that the distribution of DDB2 changed upon UV irradiation, which is not observed for other NER factors except TFIIH (Volker et al., 2001). Irradiated cells show regions of dense and less dense DDB2 fluorescence (Fig. 4F-H), similar to the spatial distribution of chromatin (Kimura and Cook, 2001). In agreement, we observed co-localization between Cerulean-tagged histone H2A and DDB2-EYFP after global UV irradiation (Fig. 4H-J and line-scan Fig. 4L), which was not observed in non UV-irradiated cells (Fig. 4C-E and line-scan Fig. 4K). This indicates that DDB2 binds to UV-damaged chromatin in interphase nuclei (Rapic-Otrin et al., 1997). Previous studies have shown that XPC readily associates with interphase- and mitotic chromatin in the absence of DNA damage (Hoogstraten, 2003; van der Spek et al., 1996). In contrast to XPC, the distribution of DDB2 in metaphase cells was homogenous and no association with mitotic chromatin could be observed (Fig 4M). Global irradiation (16 J.m-2) of cells in metaphase resulted in rapid association of DDB2 with the condensed mitotic chromosomes after UV exposure (Fig. 4M). After mitosis, daughter cells displayed the characteristic chromatin localization as observed in irradiated interphase nuclei (Fig. 4M). Binding of GFP-tagged XPG to mitotic chromatin was not observed after global UV-C irradiation (data not shown). It is interesting to note that DDB2 is the only NER protein studied so far that displays this distribution pattern in interphase nuclei upon UV irradiation. Our findings show that DDB2 has the ability to bind to UV-damaged interphase and metaphase chromatin.

The amount of DDB2 that is proteolytically degraded is UV dose-dependent To investigate whether YFP-tagged DDB2 is degraded upon UV irradiation we analyzed whole cell extracts by western blotting using antibodies against DDB2 (Figure 1C). Our analysis shows that DDB2-EYFP is degraded within 4 hours (Fig. 1C) and that expression of DDB2 is restored after 20 hours (data not shown). The amount of DDB1 and CUL4A remains unchanged after UV irradiation (Fig 1C). Next, we used live-cell imaging to measure the degradation kinetics of DDB2-EYFP in living UV- irradiated cells up to 6 hours. We observed that locally UV irradiated cells (100 J.m-2) showed a significant decrease in total nuclear fluorescence (within 6 hours) in contrast to cells that were not hit by local UV irradiation (Fig. 5A). Cells that were globally UV-irradiated (16 J.m-2) showed a complete loss of nuclear fluorescence with a t1/2 of ~2 hours (Fig. 5B, 5D). To unambiguously show that the decrease in nuclear fluorescence is due to proteolytic degradation of DDB2-EYFP we treated cells with proteasome inhibitor MG-132 and observed that DDB2-EYFP levels remained unchanged up to 6 hours after 16 J.m-2 (Fig. 5C). Irradiation with UV doses of 8 and 4 J.m-2 resulted in degradation of ~75% and ~50% of the DDB2 pool, respectively (Fig. 5D). Non UV-irradiated DDB2-EYFP cells (Fig. 5D) and globally irradiated HeLa cells transfected with EYFP-NLS (8 J.m-2) showed no change in nuclear

71 Chapter 4 fluorescence between 0 and 6 hours. Previous experiments showed that DDB2 is degraded in several NER mutant cell-lines, including XP-C (Rapic-Otrin et al., 2002). To determine if the rate of degradation is affected by the ongoing NER process, we measured degradation of DDB2-EYFP in XP-C and XP-A cells. The initial breakdown of DDB2 was faster in both XP lines than in wild-type cells irradiated with the same UV-dose (Fig. 5D). The kinetics of degradation in repair proficient cells resembles the repair kinetics of 6-4 PPs in the UV dose range employed (van Hoffen et al., 1995). We suspect that breakdown of DDB2-EYFP in cells irradiated with UV doses of 4 and 8 J.m-2 is not complete because a significant fraction of 6-4 PPs is removed before all DDB2 is degraded. For example, NER proficient cells remove ~50% of their 6-4 PPs in the first hour after a dose of 8 J.m-2 (Moser et al., 2005). It is possible that the breakdown of DDB2 is faster in NER deficient cells due to the persistence of DNA damage in the genome of these cells. It is unclear why DDB2 is not completely degraded in XP-C and XP-A cells (Fig. 5D). Although a single high dose of UV-C can result in the almost complete break-down of DDB2, such a situation, and thus complete break-down of DDB2, is not likely to occur in cells of an organism that is exposed to solar UV irradiation. Together, our data indicate that the amount of DDB2 that is proteolytically degraded is UV-dose dependent and that the rate of DDB2 breakdown depends on ongoing NER.

All DDB2 can bind to lesions in UV-irradiated cells To examine the properties of bound and unbound DDB2 upon UV-irradiation, we applied strip-FRAP on globally irradiated cells (Fig. 6A). The FRAP curves showed incomplete recovery (1 hour after 16 J.m-2 at 37°C) indicative of a significant immobile fraction (Fig. 6C). We observed fast but incomplete recovery (within 2 seconds) of DDB2 in irradiated cells. This initial very fast recovery (t1/2 = 0.1 s) is probably due to fluorescence blinking, given that the average lifetime of GFP in a dark non-emissive state is around 2 seconds (Garcia-Parajo et al., 2000). Blinking of GFP is the result of switching between an emissive (anionic) on-state and a non-emissive (neutral) off-state (Dickson et al., 1997). The off-time is independent of excitation intensity whereas the on-time is shortened at high excitation intensities (such as the FRAP bleach pulse), which makes blinking a light-induced process (Garcia-Parajo et al., 2000). Comparable fast but incomplete recovery was measured with EYFP and EGFP tagged histone H2A (Fig. 6C and data not shown), which are immobile chromatin proteins on this time-scale (Phair and Misteli, 2000). In contrast to wild-type DDB2, the R273H mutant DDB2 protein was not immobilized after UV-C irradiation (Fig 6C). This confirms the inability of DDB2 R273H to bind to damaged DNA (Hwang et al., 1998) and demonstrates that the R273 residue is essential for binding to damaged DNA. Monte Carlo simulations of the experimental FRAP curves, which were corrected for blinking, (Farla et al., 2004) yielded an immobile fraction of 85 ± 15% for DDB2, measured 1 hour after UV irradiation at a dose of 8 or 16 J.m-2 (Fig.

72 DDB2 dynamics

Figure 5. UV-induced degradation of DDB2. (A) Two DDB2-YFP cells are locally UV- damaged (nuclei marked by the green dotted contour line), while a third cell is not hit by UV light (indicated by the red dotted contour line). The same three cells are shown shortly (40 seconds) after UV irradiation and six hours after UV irradiation. (B) Example of a DDB2-EYFP cell that has been globally UV- irradiated (16 J/m2) and monitored for five hours. (C) Example of a globally UV- irradiated (16 J/m2) DDB2-EYFP cell that has been treated with 50 µM MG-132. (D) Quantification of the total nuclear fluorescence in the absence of UV-C irradiation (green line, NoDa, n = 8) and after global UV irradiation (GloDa) in MRC5 human cells with 4 (light-purple line, n = 10), 8 (red line, n = 12), and 16 J/m2 (light-blue line, n = 10) and in XP-C (dark-blue line, n = 15) and XP-A cells with 16 J/m2 (dark-purple line, n = 7). All values were corrected for background fluorescence and the initial total nuclear fluorescence intensity was set to 100%. Error bars depict the standard error of the mean.

Figure 6. FRAP analysis of DDB2 mobility in UV-irradiated cells. (A) Example of a globally UV-irradiated DDB2-EYFP cell in which a narrow strip (2 µm) across the nucleus is bleached. (B) Recovery kinetics of DDB2- EYFP in undamaged cells (red line, NoDa, n = 14), in locally damaged cells (green line, outside LoDa, n = 14) and in globally UV-irradiated cells (4 J/m2; blue line, GloDa, n = 15). The recovery plots are normalized to 0 and to the pre-bleach intensity. (C) Shows the recovery plots of DDB2-EYFP in globally damaged cells 1 hour after 16 J.m2 (purple line, GloDa, n = 16) and in globally damaged cells 4 hours after 16 J.m2 (blue line, GloDa, n = 15). The recovery plots of mutant R273H DDB2 1 hour after 16 J.m2 are shown in red (n = 12). The recovery of EYFP-histone H2A is also shown (green line, n = 15). The recovery plots are normalized to 0 and to the pre-bleach intensity. (D) Normalized recovery of DDB2-EYFP in cells irradiated with 4 J/m2 (blue line, GloDa). The recovery plot is normalized to post-bleach intensity and compared to DDB2-EYFP in undamaged cells (red line, NoDa). (E) UV-immobilized fraction of DDB2-EYFP in MRC5 cells 1 hour after global irradiation with 4, 8 and 16 J/m2 and 4 hours after 16 J/m2. The immobilized fractions depicted in (E) were determined by Monte Carlo simulations. The bound fractions obtained from the ten best simulations that fitted the experimental data were averaged. The error bars represent the standard deviation.

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6E). Such a UV-C dose produces approximately 3.105 6-4 PP and 1.106 CPD (Perdiz et al., 2000), indicating that there are sufficient damages to bind all DDB2. Even 4 hours after UV irradiation (16 J.m-2) we still observed a significant immobilization of DDB2 (65 ± 5%, Fig. 6C, 6E). It is likely that this immobilization reflects binding to CPDs since the majority of 6-4PPs is removed within 4-6 hours (Moser et al., 2005; van Hoffen et al., 1995). For comparison, the fraction of immobilized XPC-EGFP decreases to background levels ~2 hours after global irradiation at a UV dose of 8 J.m-2 (Hoogstraten, 2003), indicating that NER complexes assembled on slowly repaired CPDs can not be detected. There is significant degradation of DDB2 between 1 (~15% degradation; Fig. 5D) and 4 hours after UV (~85% degradation; Fig. 5D). Considering this degradation, we estimate that the number of DDB2 molecules bound to damaged DNA between 1 and 4 hours after UV decreases with a factor of ~8. A lower UV dose (4 J.m-2) induced a less pronounced immobilization of DDB2 (40 ± 9%; Fig. 6B, 6E). Unbound DDB2 in these UV-irradiated cells has the same mobility as DDB2 in unchallenged cells (Fig. 6D). Accordingly, the mobility of DDB2 outside local UV-induced DNA damages (100 J.m-2 through 5 µm pores) was similar to the mobility in undamaged cells (Fig. 6B), indicating that DDB2 molecules that are not bound to a lesion have the same mobility and thus a similar complex composition as DDB2 in non-damaged cells. These findings indicate that DDB2 binds to lesions in a UV-dose dependent manner up to complete binding of all DDB2 molecules. Moreover, DDB2 that dissociates from lesions has a similar diffusion rate as DDB2 in undamaged cells, suggesting that the composition of the DDB2 complex that dissociates from lesions is similar to that in undamaged cells.

Release of DDB2 from lesions is XPC independent To determine the dissociation kinetics of DDB2 from UV-damaged DNA, we applied fluorescence loss in photobleaching (FLIP). A region distant from the local UV-induced DNA damage was continuously bleached (Figure 7A) and the decrease of fluorescence in the locally damaged area was measured (Figure 7B). The half-time (t1/2) of the FLIP curve is indicative for the koff of DDB2 at sites of DNA damage. Directly after bleaching we observed a decrease of fluorescence intensity, which corresponds to the non-bound DDB2 pool. The overall decrease in the locally UV-irradiated area was much slower due to the binding of DDB2 to UV-induced damage. The t1/2 of release of DDB2 at sites of UV damage (at 37°C) was ~110 seconds (Fig. 7B). Surprisingly, the dissociation kinetics of DDB2 from UV-lesions was similar in cells lacking XPC (Fig. 7B), indicating that XPC has no influence on the release of DDB2. The residence time of DDB2 was significantly longer at 27°C (t1/2 ~ 220 seconds; Fig. 7B) compared to 37°C (t1/2 ~ 110 seconds), indicating that the dissociation of DDB2 from UV- damaged DNA is temperature sensitive.

To compare the release kinetics of DDB2 with that of other NER proteins, we have

74 DDB2 dynamics applied FLIP on cell-lines expressing XPC-EGFP and XPG-EGFP (Hoogstraten, 2003; Zotter et al., 2006). Residence times of 28 seconds and 45 seconds were measured for XPC-EGFP and XPG-EGFP, respectively (t1/2 of the FLIP curves shown in Figure 7B). Comparison of the residence time of DDB2 with the residence times of XPC- EGFP and XPG-EGFP shows that DDB2 resides significantly longer on UV-damaged DNA than XPC (about 4 times longer since t1/2 values are 110 seconds and 28 seconds, respectively) and about 2 times longer than the NER endonuclease XPG.

Discussion Mammalian nucleotide excision repair is initiated by the XPC complex, which has high affinity for UV-damaged DNA (Sugasawa et al., 1998; Volker et al., 2001). Besides XPC, mammalian cells express a second UV-damage recognition protein with high affinity for UV lesions: DDB2, which is part of an E3 ubiquitin (ub) ligase containing CUL4A, ROC1 and DDB1 (Groisman et al., 2003). Currently, it is not clear how these two protein complexes cooperate in the DNA damage recognition step of mammalian NER in the living cell. To investigate the interplay between DDB2 and XPC in vivo we used quantitative live cell imaging by employing fluorescently-tagged proteins. Figure 7. Dissociation kinetics of DDB2. (A) Example of a DDB2-EYFP cell after local UV- irradiation with 100 J/m2. A strip of about one third of the cell nucleus opposite to the local damage was continuously bleached and the fluorescence decrease in the local damage was monitored. (B) Quantification of the decrease in fluorescence signal in the local damaged nuclear area. The half-time (t1/2) of a FLIP curve corresponds to the residence time of a protein molecule in the locally damaged area. FLIP was performed on DDB2-EYFP in MRC5 human fibroblasts at 37°C (purple line, n = 10) and at 27°C (orange line, n = 10), DDB2-EYFP in XP20MA human XP-C fibroblasts (blue line, n = 11), XPF-EGFP in UV135 Chinese hamster ovary XP-G cells (green line, n = 8) and XPC- EGFP in XP4PA human XP-C fibroblasts (red line, n = 9). The error bars represent the standard error of the mean.

DDB2 resides in a high molecular weight protein complex We measured the mobility of EYFP-tagged DDB2 and of EGFP-tagged CUL4A in nuclei of living human fibroblasts (Fig. 2) and found that the mobility of both fusion proteins is similar and remarkably low. The apparent diffusion constants of DDB2 and

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CUL4A (~2.5 µm2/s) correspond to that of globular protein complexes with a molecular weight between 500 and 700 kDa (Fig. S1). This suggests that the majority of DDB2 resides in the DDB-CUL4A-ROC1 (280 kDa) E3 ub ligase complex, possibly associated with COP9 (450 kDa). The naturally occurring R273H mutant DDB2 protein displays a similar slow behavior as wild-type DDB2 (Fig 2), suggesting that this protein is assembled into the DDB-CUL4A-ROC1 complex. In agreement with these results, DDB2 R273H was recently demonstrated to form a complex with DDB1 in vitro (Wittschieben et al., 2005). Although CUL4A and DDB2 have the same mobility in living cells, it should be noted that CUL4A is also part of a number of other protein complexes not containing DDB2 and therefore its mobility is not necessarily directly related to that of DDB2 (He et al., 2006). About 15 proteins containing WD40 repeats were shown to associate with DDB1-CUL4A (He et al., 2006). All of these DDB1- interacting proteins have a MW of ~50 kDa, comparable to DDB2. This might explain why the mobility of CUL4A is similar to the mobility of DDB2. Our results show that transiently transfected EGFP-CUL4A, which can be part of all of these DDB1-CUL4A complexes, accumulates at UV-damaged sites in significant quantities. CUL4A does not accumulate at UV-damaged sites unless associated with DDB2 (Li et al., 2006). Therefore, the accumulation of EGFP-CUL4A indicates that a significant fraction of CUL4A complexes contains DDB2. Recently, DDB2 was shown to bind to UV- damaged DNA after knock- down of DDB1 and CUL4A (El-Mahdy et al., 2006; Li et Figure S1. Relation between the apparent diffusion constant and al., 2006). In the absence of the molecular weight of a protein. The mobility of EGFP alone (30 kDa) and of the EGFP-tagged NER proteins XPA (70 kDa), DDB1, the proteins CUL4A XPG (210 kDa), ERCC1/XPF (215 kDa), and TFIIH (500 kDa) and DDB2 do not interact, was determined by FRAP and subsequent Monte Carlo simulati- (He et al., 2006) showing that ons, yielding apparent diffusion constants for all fusion proteins and EGFP alone. The data points were fitted with the exponential DDB2 alone binds to UV- equation y = 246x-0,73, where x is the molecular weight in kDa damaged DNA (Li et al., and y the apparent diffusion constant in µm2/s. Using this 2006). Our experiments equation, diffusion constants were derived for monomeric DDB2 (78 kDa), dimeric UV-DDB (205 kDa), multi-protein complex suggest that the majority of DDB-CUL4A-ROC1 (310 kDa) and DDB-CUL4A-ROC1 DDB2 in living DDB1 associated with COP9 (760 kDa). The calculated diffusion containing cells is part of the constants are plotted in the same graph as the measured diffusion constant for DDB2-EYFP that fits well to the expected diffusion DDB-CUL4A-ROC1 constant of DDB-CUL4A-ROC1 associated with COP9. All complex, rather than free values include an additional 30 kDa due to the presence of DDB2. UV-induced DNA yellow/green fluorescent protein.

76 DDB2 dynamics damage triggers the binding of DDB2 and CUL4A to UV-damaged chromatin (Fig. 3, 4). Mutant DDB2 with a R273H substation does not become immobilized after UV-C irradiation (Fig 6), demonstrating that the R273 residue is essential for binding to damaged DNA. DDB2 molecules that dissociate from lesions have the same mobility and thus complex composition as DDB2 in undamaged cells (Fig. 6). This suggests that DDB2 that is released from damaged DNA is bound to COP9, which is the negative regulator of the DDB2 E3 complex. Our findings are consistent with a model in which activation of the DDB2 E3 ub ligase complex (by dissociation of COP9) only occurs on damaged chromatin and in which the bulk of DDB2 that dissociates from lesions is inactivated by binding to COP9. In this way the ub ligase activity is specifically directed to chromatin at DNA damaged sites.

DDB2 binds to DNA lesions independently of XPC To investigate if the dynamic behavior of DDB2 is dependent on functional NER, we measured the accumulation of endogenous and YFP-tagged DDB2 in NER deficient cells. Binding of endogenous DDB2 to UV-damaged DNA can be detected in the absence of functional XPC, XPA, XPG or XPF protein (Fig. S2). Moreover, the association kinetics of DDB2-EYFP with UV-damaged DNA is not influenced by XPC (Fig. 3). Up to 85% of the cellular DDB2 pool becomes immobilized after UV (Fig. 6), which is considerably higher than the percentage of the cellular XPC pool (~25%) that is immobilized after a similar UV dose (W. Vermeulen, unpublished data). The number of endogenous DDB2 molecules per cell is ~1.105 (Keeney et al., 1993), whereas there are ~3.104 XPC molecules per cell (Araujo et al., 2001). Given that the expression level and the damage-bound fraction of XPC is lower than that of DDB2 (Araujo et al., 2001; Keeney et al., 1993), we estimate that shortly after UV the number of damage- bound DDB2 molecules is ~10 times higher Figure S2. Accumulation of endogenous DDB2 and than the number of damage-bound XPC TFIIH (p89 subunit) after local UV-damage -2 molecules (85% of 1.105 vs. 25% of 3.104 (30 J.m ) in wild-type human cells (VH10) and several NER mutant cell-lines (XP-C, XP-A, XP-G for DDB2 and XPC, respectively). This and XP-F). The left column show the immuno- comparison is possible since the fluores- fluorescence labeling with anti-XPB (green) and the right column shows labeling with anti-DDB2 (red) cently-tagged proteins are present in the antibody.

77 Chapter 4 same amounts as their endogenous counterpart (see materials and methods and results section). Immunoprecipitation experiments have indicated that XPC and DDB2 directly interact even in the absence of UV-lesions (Sugasawa et al., 2005). It has been suggested that DDB2 recruits XPC to UV-induced lesions resulting in DDB2 displacement (Sugasawa et al., 2005). If each DDB2 protein would recruit an XPC molecule, resulting in the dissociation of DDB2 due to inactivation by ubiquitination (Sugasawa et al., 2005), the kinetics of DDB2 dissociation should be XPC-dependent. In vivo measurements reveal that DDB2 resides about four times longer on UV- damaged DNA than XPC and that the release of DDB2 from UV-lesions is not affected by XPC (Fig. 7). These findings, in combination with the difference in the number of damage-bound XPC and DDB2 molecules, suggest that DDB2 does not physically interact with XPC on UV-damaged DNA. It is possible that there is an interaction between the two recognition factors on some (possibly poorly recognized) lesions resulting in the DDB2-mediated ubiquitination of XPC (Sugasawa et al., 2005), but our results indicate that the bulk of DDB2 binds to lesions independently of XPC.

DDB2 primes chromatin around lesions to be targeted by NER Based on our findings we propose the following model for DDB2 function in the living cell: DDB2 is part of an inactive E3 ub ligase complex that contains DDB1, CUL4A and the COP9 signalosome (Groisman et al., 2003). Essentially all DDB2 binds to UV- damaged DNA at high UV doses independently of XPC. It is likely that the E3 ub ligase binds to damages as a holocomplex, since DDB2 and CUL4A have the same rate of binding to lesions. A DDB2 molecule that binds to DNA damage is not inactivated immediately, but can dissociate and rebind several times to UV-damaged DNA, since the residence time of DDB2 is ~2 minutes, whereas the half-time of breakdown is two orders of magnitude larger (~2 hours). This indicates that a DDB2 molecule can bind about 75 times to UV-damaged DNA before it is degraded. The binding-deficient DDB2 mutant K244E is resistant to degradation. It is has been suggested that binding to damaged DNA is the trigger for DDB2 breakdown (Nichols et al., 1996; Rapic- Otrin et al., 2002). Our findings show that binding of DDB2 to UV-damaged DNA is not the primary trigger for its proteolytic degradation. Other factors are likely to be involved in the regulation of DDB2 breakdown, such as the checkpoint protein Claspin (Praetorius-Ibba et al., 2007). However, the exact trigger for DDB2 breakdown remains to be elucidated.

Our findings show that DDB2 interact with UV-damaged DNA independently of XPC. It is attractive to speculate that DDB2 marks lesions to be targeted by NER. Several studies have shown that DDB2 interacts with chromatin remodeling complexes, such as p300 histone acetyltransferase (Datta et al., 2001; Rapic-Otrin et al., 2002). The DDB2 E3 ub ligase was recently shown to ubiquitinate histone H2A, H3 and H4 (Kapetanaki et al., 2006; Wang et al., 2006), linking DDB2 to chromatin remodeling.

78 DDB2 dynamics

We propose a scenario in which the binding of DDB2 to a lesion and subsequent remodeling creates a local chromatin environment that facilitates the assembly of a NER complex. The ubiquitination of histone H3 and H4 results in a weakened interaction with DNA (Wang et al., 2006). Thus, DDB2-dependent histone ubiquitination could result in the dissociation of histones from the DNA. Evidence for this is that histone H3 was recently shown to be incorporated into newly repaired chromatin in a CAF-1-dependent manner (Polo et al., 2006). Alternatively, the DDB2- dependent ubiquitination of histones might facilitate binding of XPC through the ubiquitin-associated (UBA) domains of RAD23B (Bertolaet et al., 2001; Kapetanaki et al., 2006). After the initial massive binding of DDB2 to UV-lesions, thereby marking of these sites for NER, the effective DDB2 concentration is gradually lowered due to its proteolytic degradation. Breakdown may be important because otherwise potential binding sites for XPC are occupied by DDB2. Evidence for this comes from in vitro studies using purified components, showing that NER is inhibited by the addition of UV-DDB in the absence of ubiquitination factors (e.g. E1, E2 and ub itself). Inhibition is relieved when ubiquitination factors are added (Sugasawa, 2006; Sugasawa et al., 2005). The importance of DDB2 degradation in vivo is illustrated by the compromised removal of CPD upon interference with DDB2 proteolysis (El-Mahdy et al., 2006; Groisman et al., 2003; Wang et al., 2005a).

Together, our results give insight into the early steps of DNA damage detection. DDB2 diffuses in the undamaged nucleus as part of an E3 ubiquitin ligase. In UV-damaged cells most DDB2 binds to DNA lesions. We provide evidence that DDB2 functions independently of damage recognition protein XPC. Although DDB2 is not part of the pre-incision complex, it has a central role in NER, probably by preparing chromatin around DNA lesions for assembly of the pre-incision complex.

Materials and Methods

Cell lines. Cell lines used in this study were human fibroblasts MRC5-SV expressing DDB2-EYFP and EYFP-DDB2, XPC deficient XP20MA-SV expressing DDB2-EYFP, XPA deficient XP12RO-SV, XPC deficient XP4PA-SV expressing XPC-EGFP (Hoogstraten, 2003), XPA deficient XP2OS-SV expressing EGFP-XPA (Rademakers et al., 2003), XPG deficient XPCS1RO-SV expressing XPG-EGFP (Zotter et al., 2006), XPB deficient XPCS2BA-SV expressing XPB-EGFP (Hoogstraten et al., 2002), HeLa cells, Chinese hamster ovary cells: UV135 expressing XPG-EGFP (Zotter et al., 2006), and 43-3B expressing ERCC1-GFP (Houtsmuller et al., 1999). The expression level of all GFP-tagged repair proteins is comparable to the level of endogenous proteins as shown by western blot analysis (Hoogstraten, 2003; Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006). MRC5-SV cells were cultured in Ham’s F12 supplemented with 400 µg/ml G418, XP20MA-SV cells were cultured in DMEM supplemented with 600 µg/ml G418 and HeLa cells were cultured in DMEM. The cell lines expressing GFP-tagged NER proteins were cultured in a 1:1 mixture of DMEM/Ham’s F10 medium. All media contained glutamine (Gibco, Breda, The Netherlands) supplemented with antibiotics and 10% FCS and all cells were cultured at 37°C in an atmosphere of 5% CO2.

DNA constructs. Murine DDB2 cDNA was ligated in-frame into pEYFP-N1 and pEYFP-C1 (Moser et al., 2005) (Clontech Laboratories, California, USA) and stably expressed in MRC5-SV cells and XP-C cells

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(XP20MA-SV). DDB2 was also ligated with EGFP and mCherry. CUL4A cDNA was ligated in frame with EGFP and transiently transfected in HeLa cells using lipofectamine 2000 (Invitrogen, Breda, the Nether- lands) according to the manufacturer's instructions. DDB2-EYFP was transiently transfected in XP12RO- SV cells and monitored 5 days after transfection. PCR-based overlap extension (Ho et al., 1989) was used to generate a DDB2 mutant protein with an R273H substitution. Briefly, two products were amplified from the DDB2 gene using two different primer sets each containing the same nucleotide substitution (details available on request). The first PCR product covers the first 837 bp of the DDB2 gene, whereas the second PCR product encompasses the last 512 bp of the DDB2 gene. Both products contain the region with the G818A mutation that results in an R273H amino acid substitution. In a subsequent “fusion” reaction, both overlapping products were added together and amplified using PCR. The generated product was subsequently inserted into pEYFP-N1 with EcoRI and BamHI and transiently transfected into HeLa cells.

Quantification of DDB2-EYFP expression level. To quantify the expression level of DDB2-EYFP, the cellular fluorescence of living cells was compared to the fluorescence of purified EYFP protein from E coli. The concentration of purified EYFP protein was determined by the extinction coefficient of 72,000 M-1. cm-1 (Kremers et al., 2006). Confocal slices of EYFP solutions (1 and 4 µm) and living cells (n = 44) were quantified using Metamorph software (Molecular Devices Corporation, Sunnyvale, California, USA).

Immunoblot analysis. Western blot analysis was performed as described previously (Fousteri et al., 2006). DDB2 was detected with mouse polyclonal anti-DDB2 antibodies (1:2000, a gift from Dr. A.A. Wani), DDB1 was detected with goat polyclonal anti-DDB1 antibodies (1:600, Abcam, Cambridge, USA) and CUL4A was detected with rabbit polyclonal CUL4A antibodies (1:300, Santa Cruz, California, USA). Pro- tein bands were visualized via chemiluminescence (ECL-Plus, Amersham Biosciences, Uppsala, Sweden) using Horseradish Peroxidase (HP)-conjugated secondary antibodies and exposure to ECL-Hyperfilms (Amersham Biosciences, Uppsala, Sweden).

Immunofluorescence. Immunolabeling was carried out as described before (Moser et al., 2005). Primary antibodies used were: mouse monoclonal antibody against XPB (a gift from J.M. Egly, IGMC, Illkirch, France), rabbit polyclonal antibody against DDB2 (Acris, Hiddenhausen, Germany), rabbit polyclonal against CUL4A (1:100, Santa Cruz, California, USA) and goat polyclonal against DDB1 (1:300, Abcam, Cambridge, USA). Secondary antibody used were: alexa 488-conjugated goat anti-mouse antiserum (Mo- lecular probes/Invitrogen, California, USA) and Cy3-conjugated goat anti-rabbit antiserum (Jackson Im- munoResearch Laboratories, West Grove, Pennsylvania). All antibodies were diluted in PBS containing 0.15% glycine and 0.5% bovine serum albumin.

Microscopic analysis. Assembly and degradation kinetics were measured on a Zeiss Axiovert 200M wide field fluorescence microscope, equipped with a 100x Plan-Apochromat (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Germany) and a Cairn Xenon Arc lamp with monochromator (Cairn research, Kent, U.K.). Images were recorded with a cooled CCD camera (Coolsnap HQ, Roper Scientific, USA). FRAP and FLIP experiments were performed on Zeiss LSM 510 confocal microscope, equipped with a 63x Plan-A (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Germany) and a 60 mW Argon laser (488 and 514 nm). Both microscopes were equipped with an objective heater and cells were examined in microscopy medium (137 mM NaCl, 5.4 mM KCl, 1,8 mM CaCl2, 0.8 mM MgSO4, 20 mM D-glucose and 20 mM HEPES) at 37°C or 27°C.

UV irradiation. For all experiments, cells were irradiated with a UV source containing four UV lamps (Phi- lips TUV 9W PL-S) above the microscope stage. The UV dose rate was measured to be 3 W.m-2 at 254 nm. For induction of global UV-damage, cells were rinsed with microscopy medium and irradiated for 1.5 seconds (4 J.m-2), 3 seconds (8 J.m-2), 6 seconds (16 J.m-2), or 12 seconds (32 J.m-2). For induction of local UV-damage, cells were UV irradiated through a polycarbonate mask (Millipore Billerica, Massachusetts, USA) with pores of 5 µm (Moné et al., 2001) and subsequently irradiated for 39 seconds (100 J.m-2).

Fluorescence Recovery after Photobleaching (FRAP). FRAP analysis was used to measure the mobility of DDB2-EYFP and EGFP-CUL4A as described by Houtsmuller and co-workers (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006). The data was normalized to pre- bleach intensity or to post-bleach intensity after full equilibration using the second data point for normali-

80 DDB2 dynamics

zation to 0. Normalization to post-bleach intensity was used to compare the shape of different FRAP curves.

Fluorescence Loss in Photobleaching (FLIP). FLIP analysis was performed by continuously bleaching a third of a locally UV-irradiated nucleus as previously described (Essers et al., 2002; Hoogstraten et al., 2002). Fluorescence in the locally damaged area was monitored with low laser intensity. All values were background corrected.

Monte Carlo simulations. For analysis of FRAP data, experimental FRAP curves were normalized to pre- bleach values and corrected for monitor bleaching. FRAP curves were generated by Monte Carlo simulations in which three parameters were varied: diffusion constant (ranging from 0.04 to 36 µm2/s), immobile fraction (0, 10, 20, 30, 40, 50, 60, 70, 80, 90, 99.9 %) and time spent in an immobile state (0.25, 0.5, 1, 2, 4, 8, 16, 32, 64, 128, 256, 512, ∞ s) (Farla et al., 2004; Zotter et al., 2006). Fluoresce blinking was also simulated based on FRAP measurements on cells expressing EGFP- H2A and EYFP-H2A (Rabut and Ellenberg, 2005). Simulated curves that fitted the experimental curves were selected based on the method of least squares. The reported error is the standard deviation calculated from ten independent simulations. Monte Carlo simulations were based on a model of random diffusion in an ellipsoid volume and simple binding kinetics representing binding to immobile elements. Diffusion was simulated by deriving novel positions M(x+ dx, y+ dy, z+ dz) for all mobile molecules M(x, y, z) after each time step, where dx = G(r1), dy = G(r2) and dz= G(r3), ri is a random number (0 ≤ ri ≤ 1) chosen from a uniform distribution, and G(ri) is an inversed cumulative Gaussian distribution with µ = 0 and σ2 = 6Dt, where D is the diffusion coefficient and t is time measured in unit time steps. Immobilization was based on simple binding kinetics described by: kon/koff = Fimm / (1 - Fimm), where Fimm is the relative number of immobile molecules. The chance for each particle to become immobilized was defined as Pimmobilise = kon = koff .Fimm / (1 - Fimm), where koff = 1/ timm, and timm is the average time a molecule is immobile, measured in unit time steps. The chance for each particle to be released from an immobile state was defined as: Pmobilise = koff = 1/ timm. The FRAP procedure was simulated on the basis of an experimentally derived three-dimensional laser intensity profile providing a chance for each molecule to get bleached during simulation of the bleach pulse based on the 3D position of the molecule.

Assembly kinetics. Cells were grown in glass bottom dishes (MatTek, Ashland, Massachusetts, USA) and locally UV-irradiated as described by van Driel and co-workers (Moné et al., 2004; Zotter et al., 2006). Accumulation after local irradiation was quantified with Object-Image software. Time courses were normalized with respect to the plateau level. Start of the UV irradiation was defined as t=0.

Degradation kinetics. To measure the degradation of DDB2-EYFP, cells were globally irradiated with 4, 8, 16 and 32 J.m-2. For the treatment of cells with proteasome inhibitors, MG-132 (Sigma, Zwijndrecht, the Netherlands) was dissolved in DMSO at 10 mM. Three hours prior to UV irradiation medium was replaced with microscopy medium containing 50 µM MG132 (Pajonk et al., 2005). Cells were monitored for 6 hours and the fluorescence intensities at every time point were quantified.

Acknowledgement

This work was supported by ZonMW, the Netherlands (grant 912-03-012). A polyclonal antibody directed against DDB2 was a kind gift of Drs. A.A. Wani and Q.E. Wang (The Ohio State University, Ohio, U.S.A.). CUL4A cDNA was kindly provided by Drs. T. Chiba and K. Tanaka (Tokyo Metropolitan Institute of Medical Science, Japan), the Cerulean-H2A construct by Dr. S. Diekmann (Department of Molecular Biology, Jena, Germany) and the mCherry cDNA by Dr. R.Y. Tsien (University of California, San Diego, U.S.A.). The authors thank are grateful to Drs. A. Politi, C. Di- nant and P.O. Mari for stimulating discussion and Drs. R.Th. Dame, F. Tessadori and M.J. Moné for critical reading of the manuscript.

81 82 Chapter 5 Spatial Organization of Nucleotide Excision Repair Proteins After UV-induced DNA Damage in the Human Cell Nucleus

Liliana Solimando, Martijn S. Luijsterburg, Lorella Vecchio, Wim Vermeulen, Roel van Driel, and Stanislav Fakan

Submitted

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Abstract Nucleotide excision repair (NER) is an evolutionary conserved DNA repair system that is essential for the removal of UV-induced DNA damage. In this study we have investigated how NER is compartmentalized in the interphase nucleus of human cells at the ultrastructural level by using electron microscopy in combination with immuno- gold labeling. We analyze the role of two nuclear compartments, i.e. condensed chromatin domains and the perichromatin region. The latter contains partly decondensed and transcriptionally active chromatin at the surface of condensed chromatin domains. We have studied the distribution of damage-recognition protein XPC and of XPA, which is a central component of the chromatin-associated NER complex. Both XPC and XPArapidly accumulate in the perichromatin region after UV irradiation, whereas only XPC is moderately enriched in condensed chromatin domains. This indicates that DNA damage is detected by XPC throughout condensed chromatin domains, whereas DNA repair preferentially occurs in the perichromatin region. Our results suggest that UV-damaged DNA inside condensed chromatin domains is relocated to the perichromatin region. In addition, we provide evidence that UV-damaged chromatin undergoes large-scale decompaction, possibly related to the relocation process. Our results offer novel insight into the dynamic spatial organization of DNA repair in the human nucleus and reveal that the dynamic spatial organization of DNA repair is more complex than previously anticipated.

Introduction Nucleotide excision repair (NER) is a highly conserved DNA repair system that removes a variety of damages from the genome (Hoeijmakers, 2001). Placental mammals fully rely on NER for the removal of UV-induced lesions, such as cyclobutane pyrimidine dimers (CPDs) and 6-4 photoproducts (6-4PPs). Both types of helix distorting lesions are potent inhibitors of transcription by RNA polymerase II (RNAP II) and replication by DNA polymerases δ and ε (Lehmann et al., 2007). Two closely related NER pathways exist that are referred to as global genome NER (GGR) and transcription-coupled NER (TCR). TCR removes lesions exclusively from transcribed genes, whereas GGR repairs lesions elsewhere in the genome (Hanawalt, 2002). Damage recognition in GGR is carried out by the XPC protein complex, which subsequently recruits the repair machinery to process the lesions (Araki et al., 2001; Buterin et al., 2005; Maillard et al., 2007; Sugasawa et al., 1998; Volker et al., 2001). DNA damage in actively transcribed genes is detected by a stalled RNAP II in concert with the CSA and CSB proteins, which are essential for TCR (Brueckner and Cramer, 2007; Fousteri et al., 2006; Hanawalt, 2000; Laine and Egly, 2006). Although CPDs are more abundant than 6-4 PPs following UV irradiation, the latter photoproduct is removed approximately five times faster than CPDs (Mitchell and Nairn, 1989). CPD removal by GGR is inefficient, probably because of the lower affinity of XPC for this type of lesion. For the recognition of CPDs in GGR, XPC is assisted by UV-DDB,

84 Spatial organization of NER which is essential for efficient repair of CPDs (Groisman et al., 2003; Hwang et al., 1998; Tang et al., 2000). TCR is particularly important for the removal of lesions such as CPDs that are inefficiently cleared away by GGR (Balajee and Bohr, 2000; Hanawalt, 2000; Mullenders and Berneburg, 2001). After the damage recognition step in GGR and TCR, both pathways converge to a common molecular mechanism that involves approximately 30 proteins in mammals (Araujo et al., 2000; de Laat et al., 1999; Lindahl and Wood, 1999). The ten-subunit protein complex TFIIH is the first to be recruited to the damaged DNA after the recognition step by XPC or CSB (Fousteri et al., 2006; Giglia-Mari et al., 2004; Volker et al., 2001; Yokoi et al., 2000). TFIIH utilizes its XPD helicase subunit to unwind the DNA around a lesion (Coin et al., 2007). The XPA protein is a central organizer of the NER complex and is involved in the proper orientation of the incision proteins XPG and ERCC1/XPF together with RPA (de Laat et al., 1998; Missura et al., 2001). Following dual incision by the two NER endonucleases, a stretch of ~30 nucleotides is released after which DNA polymerase δ is loaded by PCNA and fills the remaining gap, which is subsequently sealed by XRCC1-ligase III (de Laat et al., 1999; Hoeijmakers, 2001; Lindahl and Wood, 1999; Moser et al., 2007). After repair synthesis and ligation, CAF-1 is recruited to the repair site to deposit new histones and re-establish the chromatin state (Green and Almouzni, 2003; Polo et al., 2006).

Although NER has been studied extensively in vitro and in vivo, we are beginning to understand the kinetics (i.e. temporal organization) of NER complex assembly on a chromatin template in nuclei of living cells. However, little is known about the spatial organization of DNA repair in the nucleus. It has been known for many years that the cell nucleus is a highly compartmentalized structure that contains distinct structural domains. For example, the nucleus contains chromatin-dense subchromosomal domains of about 100 – 500 nm that are surrounded by interchromatin space that contains little or no chromatin (Cremer et al., 2004). Each dense chromatin domain contains several Mbs of DNA and a chromosome consists of a number of these dense chromatin domains. The perichromatin region, constituting a shell of partly decondensed chromatin at the surface of condensed chromatin domains, is a nuclear domain where replication and transcription mainly take place (Cmarko et al., 1999; Fakan, 1994; Fakan and van Driel, 2007; Jaunin and Fakan, 2002). Newly replicated DNA in the perichromatin region is rapidly internalized into the condensed chromatin area (Jaunin et al., 2000). Polycomb (PcG) proteins are also concentrated in this compartment, suggesting that PcG-mediated silencing also occurs mainly at the periphery of condensed chromatin (Cmarko et al., 2003).

In this study we investigate where nucleotide excision repair occurs in the nucleus of human cells. Our results indicate that the NER proteins XPC and XPA become significantly enriched in the perichromatin region after global UV-C irradiation. This

85 Chapter 5 suggests that NER predominantly takes place in the same chromatin compartment where transcription and replication occur. In addition, we provide evidence for considerable chromatin decondensation in response to UV-induced DNA damage, which might facilitate relocation of DNA lesions to the perichromatin region. Together, our results provide novel insight into the dynamic spatial organization of DNA repair in the human cell nucleus

Results

Visualisation of DNA damage and NER proteins by fluorescence microscopy To study whether DNA repair by NER is compartmentalized in the human nucleus or if it occurs throughout the genome, we have analyzed the nuclear distribution of the NER proteins XPC and XPA before and after UV-C irradiation at the ultrastructural level by immuno-gold labeling on ultrathin sections. To this aim, we generated cell lines stably expressing XPC-EGFP and EGFP-XPA (Politi et al., 2005; Rademakers et al., 2003). The fusion proteins were expressed in human repair-deficient XP4PA-SV and XP2OS-SV cells, lacking functional XPC and XPA, respectively. Expression of the fusion proteins re-established UV-resistance of the cell lines, showing that the EGFP- tagged NER proteins are functional (Hoogstraten et al., 2003; Rademakers et al., 2003). Moreover, the nuclear concentration of the fusion proteins was similar to that of the endogenous proteins in wild type cells (Politi et al., 2005; Rademakers et al., 2003). The subnuclear distribution of EGFP-tagged NER proteins has been analysed rather than that of the endogenous proteins, because antibodies against the latter did not recognize their antigens after embedding and sectioning. In contrast, anti-GFP antibodies did work well under these conditions. Fig. 1D and H and Fig. 1F and H show that in formaldehyde fixed cells that are analyzed by light microscopy, anti-EGFP antibody labeling show precisely the same intranuclear distribution as the intrinsic fluorescence of XPC and XPA in cells that express XPC-EGFP and EGFP-XPA, respectively. This validates the use of anti-EGFP antibodies in our studies.

Cells expressing XPC-EGFP and EGFP-XPAwere UV-C irradiated (60 J.m-2) and subsequently immuno-labeled for CPDs, using an anti-CPD antibody (Fig 1A, D) (Mori et al., 1991). While hardly any signal was detected in control cells, irradiated cells showed a nuclear signal in every cell (Fig 1A, D), confirming that significant numbers of CPDs are produced under these conditions. Confocal microscopy showed that within the resolution of light microscopy, the distribution of EGFP-XPA is homogeneous throughout the nucleus in UV irradiated and control cells (Fig 1E). In contrast, XPC- EGFP displayed a typical chromatin-like distribution in irradiated and control cells (Fig 1B). This is probably due to the affinity of this protein for DNA in general (Shivji et al., 1994). Global UV-C irradiation did not result in a visible redistribution of these two repair proteins at the light microscopic level (Fig 1B, E). To show that XPC and

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XPA are engaged in NER after UV-C irradiation, we performed strip-FRAP (Hout- smuller and Vermeulen, 2001) on globally irradiated cells and control cells. Significant immobilization of XPC-EGFP (Fig1G) and of EGFP-XPA (Fig 1H) was detected after UV-C irradiation indicating that a significant fraction of both protein pools participates in DNA repair under our experimental conditions (see also (Rademakers et al., 2003).

Figure 1.Immunofluorescence and live cell microscopy. (A). Example of XPC-EGFP cells immunofluorescently labeled against CPDs in non-irradiated control cells (left panel) or 10 minutes after global UV-C irradiation at 60 J.m-2 (right panel). The EGFP signal is shown in green (upper panel) and the anti-CPD staining of the same cells in red (lower panel). (B). Example of EGFP-XPA cells immuno fluorescently labeled against CPDs in non- irradiated control cells (left panel) or 10 minutes after global UV-C irradiation at 60 J.m-2 (right panel). The EGFP signal is shown in green (upper panel) and the anti- CPD staining of the same cells in red (lower panel). (C) Confocal slice of living XPC-EGFP cells either non-irradiated (upper panel) or globally irradiated at 60 J.m-2 (lower panel). (D). Confocal slice of XPC-EGFP cells immunofluorescently labeled against EGFP in cells 10 minutes after global UV-C irradiation at 60 J.m-2. The intrinsic EGFP signal is shown in green (upper panel) and the anti-EGFP staining of the same cell in red (lower panel). (E) Confocal slice of living EGFP- XPA cells either non-irradiated (upper panel) or globally irradiated at 60 J.m-2 (lower panel). (F). Confocal slice of EGFP- XPA cells immunofluorescently labeled against EGFP in cells 10 minutes after global UV-C irradiation at 60 J.m-2. The intrinsic EGFP signal is shown in green (upper panel) and the anti-EGFP staining of the same cell in red (lower panel). (G) Line scan of the arrows depicted in the upper and lower panel of D, reflecting intrinsic XPC-GFP fluorescence (green line) and anti-EGFP signal (red line) in the same cell. (H) Line scan of the arrows depicted in the upper and lower panel of F, reflecting intrinsic EGFP-XPA (green line) fluorescence and anti-EGFP signal (red line) in the same cell. (I) FRAP analysis of XPC-EGFP in non-irradiated cells (blue line, n = 9) or globally irradiated cells at 60 J.m-2 (red line, n = 9). (J) FRAP analysis of EGFP-XPAin non-irradiated cells (blue line, n = 9) or globally irradiated cells at 60 J.m-2 (red line, n = 9).

Ultrastructural distribution of UV-induced DNA lesion. To analyze the spatial distribution of UV-induced DNA lesions upon global UV-C irradiation, we used immuno-gold labeling of ultrathin sections with antibodies that specifically bind to CPDs (Mori et al., 1991). We discriminate between two nuclear

87 Chapter 5 compartments, condensed chromatin domains and the perichromatin region. The former are defined as nuclear areas that are relatively weakly contrasted by the EDTA regressive staining method. The perichromatin region is operationally defined as an about 80 nm wide shell at the surface of condensed chromatin domains. From the approximate size of the EGFP molecule and the primary and secondary antibody it can be estimated that the gold particle is located somewhere in a sphere of 40 nm around the antigen. Therefore, this definition includes antigens up to about 80 nm at both sides of the surface of condensed chromatin domains, including those in dispersed chromatin and RNP-containing structures, such as perichromatin fibrils and granules, near the surface.

Figure 2.. Ultrastructural localization of CPDs. Example of a human XP2OS fibroblast immuno-labeled using a specific antibody against CPDs that has been A) mock-treated and B) fixed 10 min after global UV-C irradiation at 60 J.m-2. Samples were contrasted with the EDTA regressive technique in order to reduce the contrast of chromatin. Arrowheads indicate condensed chromatin, arrows designate perichromatin region. The scale bar is 0.5 µm.

Global UV irradiation at 60 J.m-2 resulted in a significant increase in CPD labeling density in the perichromatin region and condensed chromatin domains compared to non-irradiated control cells, showing that CPDs are present throughout these two nucleoplasmic domains (Fig 2). Quantitative analysis of the CPD distribution in condensed chromatin domains and the perichromatin region (defined as an about 80 nm wide shell at the surface of condensed chromatin domains) shows that the perichromatin region is two- fold less labeled than condensed chromatin domains (Fig 5A). This is most likely due to the fact that chromatin in the perichromatin region is more unfolded compared to condensed chromatin domains (Bouchet-Marquis et al., 2006) (for a review see (Fakan, 2004; Fakan and van Driel, 2007)). These results show that the perichromatin region contains an about two-fold lower concentration of DNA lesions compared to dense chromatin domains. Assuming that the probability of UV-

88 Spatial organization of NER induced DNA damage is the same in both compartments, this indicates an about two- fold lower concentration of chromatin in the perichromatin region compared to condensed chromatin domains.

Ultrastructural distribution of protein factors involved in NER To establish whether DNA repair by NER occurs in specific nuclear regions we have analyzed the distribution of EGFP-XPA and XPC-EGFP using anti-EGFP antibodies in non-irradiated cells or after 10 min and 60 min following global UV-C irradiation

Figure 3. Ultrastructural localization of EGFP-XPA and XPC-EGFP using anti-EGFP antibodies in non- irradiated and UV-irradiated cells. A) shows an example of an XP2OS EGFP-XPA fibroblast that has been mock treated. B) shows a nucleus globally irradiated with UV-C at 60 J.m-2. Similarly, C) shows a XP4PAXPC-EGFP cell that was mock treated and D) UV-C irradiated cell nucleus. The shown examples were fixed 10 min following UV irradiation. Cell monolayers were fixed in paraformaldehyde and thin sections were subsequently immuno-labeled with antibodies specific for EGFP. Samples were contrasted with the EDTA regressive technique in order to reduce the contrast of chromatin. The arrows designate the perichromatin region and the arrowheads condensed chromatin domains. N indicates the nucleolus. Scale bar is 0.5 µm.

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(60 J.m-2). After UV irradiation, the labeling intensity of XPA in condensed chromatin domains slightly decreased (Fig 3A,B and Fig 5C), whereas XPC accumulated about 1.5-fold compared to non-irradiated cells (Fig 3C,D and Fig 5C). The UV-induced accumulation of both proteins in the perichromatin region was between 3 and 6-fold, which is considerably higher than the accumulation in condensed chromatin domains (Fig 5B). Since the average chromatin density in the perichromatin region is about 2- fold lower than in condensed chromatin domains, as can be inferred from the difference in CPD labeling (see above), this shows that these two NER proteins accumulate (per unit chromatin) about 10-fold more in the perichromatin region than in condensed chromatin in response to UV damage. This indicates that about one order of magnitude more pre-incision complexes are assembled in the perichromatin region than in compact chromatin domains after UV irradiation. There is no systematic difference between the labeling at 10 min and at 60 min after UV irradiation (Fig 5B,C), indicating that the preferential accumulation in the perichromatin region is not due to, for instance, slow diffusion of proteins into the condensed chromatin domain. Our results indicate that repair of UV-induced lesions by mammalian NER mainly takes place in the perichromatin region.

Figure 4. Increase of dense chromatin areas upon global UV-C irradiation. XP2OS EGFP-XPA cells were A) mock-treated or B) fixed 10 min after global UV irradiation at 60 J.m-2 and subsequently stained with osmium ammine to specifically visualize DNA (dark colored). The analyzed sections were collected at the middle nuclear plane of the cells. IC indicates interchromatin compartment. The scale bar is 1.25 µm.

90 Spatial organization of NER

Expansion of condensed chromatin domains after DNA damage To establish whether UV irradiation results in detectable changes in large-scale chromatin structure, we analyzed ultrathin sections of UV irradiated cells using osmium ammine, a Feulgen-type stain specifically visualizing DNA (Fig 4A,B). Quantitative evaluation of the surface area of condensed chromatin domains showed that their surface area increases about 2-fold after UV irradiation compared to non- irradiated cells, indicating that the increase in volume of the chromatin domains is about 3-fold (Fig 5D). This is particularly visible in cells fixed 10 min after irradiation, while after 60 min the increase is somewhat less, probably due to partial repair of the damaged DNA sites (Fig 5D). An increase in the volume of condensed chromatin domains is indicative of considerable chromatin decondensation. These results indicate that UV irradiation triggers a transient decondensation of chromatin.

Figure 5. Quantitative evaluations of ultrastructural data. A) shows a quantitative evaluation of CPD labeling density in irradiated cells either 10 min (n= 9) or 1 h (n=10) following irradiation in i) condensed chromatin domains (orange bars) and ii) perichromatin region (blue bars). Values were corrected for residual labeling in non-irradiated cells (n=20). The values represent the average gold grain number per µm2 nuclear surface area ± SEM. B and C) show a quantitative evaluation of EGFP-XPA (blue bars) and XPC-EGFP (orange bars) labeling density (gold particles per µm2 nuclear area) in the perichromatin region (PR) shown in B) or in condensed chromatin domains shown in C). The labeling density was determined in non-irradiated cells (n=20 for XPC and XPA), and in nuclei 10 min (n=15 for XPC and XPA) and 1 hr (n=15 for XPC and XPA) following irradiation. Values represent the average ± SEM. D) shows a quantitative evaluation of the fraction of the nucleus occupied by condensed chromatin domains (%) before (n=29) and after global UV irradiation at 60 J.m-2 determined 10 min (n=15) or 1 hr (n=15) following UV exposure. The values represent the mean ± SEM. NoDa: no UV irradiation. GloDa: after global UV irradiation

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Discussion The results presented in this study indicate that nucleotide excision repair in human cells takes place predominantly in the perichromatin region, an about 100 nm wide area on the border of compact chromatin domains in the interphase nucleus (Fakan and van Driel, 2007). Transcription and replication have also been shown to be concentrated in this nuclear region (Cmarko et al., 1999; Fakan and van Driel, 2007; Jaunin and Fakan, 2002; Jaunin et al., 2000). We have used immuno-electron microscopy to identify the spatial distribution of UV-induced lesions (CPDs) and two core NER proteins, i.e. the NER damage-recognition protein XPC and the NER protein XPA, which play a crucial role in damage verification and regulation of dual incision during NER (Missura et al., 2001; Sugasawa et al., 1998; Volker et al., 2001). By using cell lines that stably express fluorescent protein tagged XPC and XPA (XPC-EGFP and EGFP-XPA), we have carried out light microscopic and electron microscopic studies in parallel. The light microscopic observations support the electron microscopic data, as far as the resolution of the former allows. FRAP experiments show that after global UV-C irradiation a substantial fraction of the XPC and XPAmolecules is bound to UV-damaged chromatin (Fig 1). High-resolution electron microscopic analysis revealed that accumulation of repair factors occurs preferentially in the perichromatin region, suggesting that this is the main site of assembly of the NER protein complex (Fig 3 and Fig 5B).

The distribution of XPC at the light microscopic level is similar to that of chromatin in non-irradiated cells (Fig 1B), reflecting its affinity for undamaged DNA (Shivji et al., 1994; Sugasawa et al., 1998). In contrast, XPA is diffusely distributed throughout the nucleoplasm (Fig 1E; Rademakers et al., 2003). As expected, immuno-gold labeling shows that after UV irradiation CPDs are present throughout the condensed chromatin domains and in the perichromatin region (Fig 2 and Fig 5A) (Gazave et al., 2005). The concentration of damage-recognition protein XPC in the condensed chromatin domains increases about 2-fold in the first 10 min after UV exposure, most probably reflecting its binding to 6-4 photoproducts (Fig 5B). In agreement with this notion, live cell experiments have shown that under these experimental conditions a maximal amount of the XPC protein pool (~25%) is bound to damaged DNA (Vermeulen, unpublished). Given the fact that in these cells roughly 20% of the nucleus is occupied by condensed chromatin domains (Fig 4 and Fig 5D), we can estimate that a 2 to 3-fold increase in XPC concentration in condensed chromatin domains after global UV irradiation can be expected, which is also what we measure (see Material and Methods). No significant increase in XPAbinding in condensed chromatin domains is observed, despite the fact that under these conditions about 30% of the XPA molecules are bound to damaged DNA sites (Rademakers et al., 2003). In contrast, both XPC and XPAaccumulate between 3 and 6-fold in the perichromatin region (Fig. 5B). This is an underestimate of the real accumulation per unit chromatin, since

92 Spatial organization of NER chromatin in the perichromatin region is more decondensed, resulting in a lower local chromatin concentration. Assuming that the CPD concentration is a crude measure for the chromatin concentration, we find that the chromatin concentration in condensed chromatin domains is about 2- fold higher compared to the perichromatin region (Fig 5A). This indicates a 10-fold increase in XPC and XPA binding per unit chromatin in the perichromatin region. Therefore, our results reveal that at least one order of magnitude more pre-incision NER complexes are assembled in this subchromosomal region in response to UV damage than in condensed chromatin domains. XPC accumulation in condensed chromatin is considerably lower than in the perichromatin region (~2-fold enrichment compared to non-irradiated cells), whereas no enrichment of XPA is detected here. Since XPC is exclusively involved in global genomic NER (GGR) (van Hoffen et al., 1995), our results suggest that GGR predominantly takes place in the perichromatin region. As most transcription takes place in the peri chromatin region it might be expected that transcription-coupled NER also occurs in this region. During the first sixty minutes after UV irradiation the amount and distribution of bound XPC and XPA hardly changes (Fig 5B,C) and the concentration of damaged sites hardly decreases (Fig 5A). Since under our experimental conditions the number of UV-induced DNA lesions is high compared to the number of available NER proteins, one does indeed expect only a moderate decrease in the number of CPDs (Moser et al., 2005; van Hoffen et al., 1995).

Our results suggest that UV-induced DNA lesions are detected by XPC throughout the condensed chromatin domains, but that the pre-incision NER complex is assembled predominantly in the perichromatin region, i.e. at or near the surface of the condensed chromatin domains. This implies that damaged sites are translocated to the surface of condensed chromatin domains. Although the mechanism of this process is not known, it may be the same as that used for relocating replicating DNA transiently to the perichromatin region and newly synthesized DNA back into the condensed chromatin domain (Jaunin et al., 2000). The need for translocation to the perichromatin region may explain in part why TCR is much faster than GGR (Bohr et al., 1985; Mellon et al., 1987), since active genes are predominantly located in the perichromatin area and do not need translocation. We observe an about three-fold increase in volume of condensed chromatin domains after UV irradiation, reflecting a considerable decondensation of the chromatin fiber. It is tempting to speculate that this decondensation is required for the relocation of damaged chromatin. Several studies suggested considerable decondensation of chromatin upon induction of UV-induced DNA damage or formation of double-strand breaks (Carrier et al., 1999; Kruhlak et al., 2006; Murga et al., 2007; Rubbi and Milner, 2003; Wang et al., 2006; Ziv et al., 2006), and chromatin relaxation thus might be a more general response to DNA damage. Chromatin decondensation in response to UV irradiation has been suggested to depend on p53 (Rubbi and Milner, 2003), the small acidic GADD45 protein (Carrier et al.,

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1999) and members of the ING tumor suppressor family (Kuo et al., 2007; Wang et al., 2006). In this study, we demonstrate a substantial enlargement of condensed chromatin domains in UV-irradiated human cells reflecting considerable chromatin deconden sation. It cannot be excluded that in addition local chromatin condensation might take place possibly caused by inhibition of transcription due to the presence of UV lesions (Monéet al., 2001). Together, our study provides novel insight into the spatial organization of the DNA repair process by NER. Major changes in chromatin structure seem to occur after the detection of UV-damaged sites, followed by translocating damaged DNA sites to the perichromatin region. The molecular mechanism of this process is unclear. It likely involves binding of XPC, which shows a significant accumulation inside condensed chromatin domains after UV irradiation. Alternatively, the UV-DDB protein complex may be involved, which is required for CPD repair and binds rapidly to UV-induced DNA lesions (Dualan et al., 1995; Luijsterburg et al., 2007; Takao et al., 1993). It is conceivable that UV-DDB is required to facilitate translocation of CPDs to the perichromatin region.

This study, together with previous work, establishes that major chromatin-associated processes, such as transcription, replication, Polycomb-mediated gene silencing and DNA repair, preferentially occur at the interface between condensed chromatin domains and the interchromatin space, i.e. the perichromatin region. They raise the fundamental question what mechanisms are involved in translocating DNA lesions to the perichromatin region and probably back again into condensed chromatin domains after repair. It is likely that it is similar to the mechanism responsible for the movement of replicating DNA. Together, our results add a spatial dimension to our understanding of NER in the human cell nucleus, revealing that the spatiotemporal organization of an essential chromatin-association process is more complex than anticipated.

Materials and Methods

Cell lines and culture conditions. Cell lines used in this study were human XPC deficient fibroblasts (XP4PA-SV) expressing XPC-EGFP (Politi et al., 2005) and human XPAdeficient fibroblasts (XP2OS-SV) expressing EGFP-XPA (Rademakers et al., 2003). The cell lines were cultured in a 1:1 mixture of DMEM/Ham’s F10 medium. All media contained glutamine (Gibco, Breda, The Netherlands), were supplemented with antibiotics and 10% FCS, and all cells were cultured at 37°C in an atmosphere of 5% CO2.

Global UV-C irradiation. For all experiments, cells were irradiated with a UV-C source containing four UV lamps (TUV 9W PL-S; Philips, Eindhoven, The Netherlands). The UV dose rate was 1.48 W.m-2 at 254 nm as measured with an SHD 240/W detector connected to an IL 1700 radiometer (International Light Techno- logies, Peabody, MA, USA). For induction of global UV-C damage, cells were rinsed with phosphate buffered saline (PBS) and irradiated for 40 seconds resulting in a UV dose of 60 J.m-2. Control cells were mock-treated.

Immunolabeling for fluorescence microscopy. Cells were fixed with 4% formaldehyde in PBS for 15 min at 4°C, permeabilized in 0.5% Triton X-100 (Serva, Heidelberg, Germany) in PBS for 5 min, and incubated with 100 mM glycine in PBS for 10 min to block unreacted aldehyde groups. Cells were rinsed with PB (130 mM KCl, 10 mM Na2HPO4 and 2.5 mM MgCl2 pH 7.4) and equilibrated in WB (PB containing 0.5% BSA,

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0.2% gelatin and 0.05% Tween 20; Sigma-Aldrich, St. Louis, MO. USA). Antibody steps and washes were in WB. EGFP labeling was carried out using a rabbit polyclonal Ab against EGFP (1:5000; Eusera, Edmon- ton, Alberta, Canada), and detection was by donkey anti-rabbit Ig coupled to Cy3 (1:500; Jackson Immu- noResearch Laboratories, West Grove, Pennsylvania). Immunolabeling of CPDs was performed using mouse monoclonal Ab TDM-2 (Mori et al., 1991). For this, the above steps were repeated, but prior to labeling DNA was denatured with 2 M HCl for 30 min at 37°C and blocked in 10% BSA in PB for 15 min. Detection was done using donkey anti-mouse Ig coupled to Cy3 (1:500 ; Jackson ImmunoResearch Laboratories). Samples were mounted in Vectashield (Vector Laboratories, Burlingame, CA, USA).

Fluorescence microscopy. Fluorescence microscopy was performed on a Zeiss Axiovert 200M widefield fluorescence microscope, equipped with a 100x Plan-Apochromat (1.4 NA) oil immersion lens (Zeiss, Ober- kochen, Germany) and a Cairn Xenon Arc lamp with monochromator (Cairn research, Kent, U.K.) Images were recorded with a cooled CCD camera (Coolsnap HQ, Roper Scientific, USA). A 375-490 excitation filter, 490 dichroic mirror and 525-40 band-pass emission filter was used for EGFP imaging (monochromator: 470 nm ± 20 nm) and a 375-580 excitation filter, 585 dichroic mirror and 620-60 band-pass emission filter was used for Cy3 imaging (monochromator: 550 nm ± 20 nm). Confocal fluorescence microscopy was performed on a Zeiss LSM 510 confocal microscope, equipped with a 63x Plan-A (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Germany) and a 60 mW Argon laser (488 and 514 nm).

Fluorescence recovery after photobleaching (FRAP). FRAP analysis was used to measure the mobility of XPC-EGFP and EGFP-XPA as described by Houtsmuller and co-workers (Hoogstraten et al., 2002; Hout- smuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006). Briefly, a strip of 512x50 pixels was imaged with 38 ms per frame at zoom 8 (1 pixel is 0.04 µm by 0.04 µm). After 5-8 images, a strip of 512x40 pixels was bleached by applying 5 scans at maximal 488 nm and 514 nm laser intensity (AOTF 100%, total time 160 ms) and the fluorescence recovery was acquired by scanning at least 100 images. The data was corrected for background fluorescence and normalized to pre-bleach intensity. FRAP experiments were performed on a Zeiss LSM 510 confocal microscope, equipped with a 63x Plan-A (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Germany) and a 60 mW Argon laser (488 and 514 nm). The LSM510 was equipped with an objective heater and cells were examined in microscopy medium (137 mM NaCl, 5.4 mM KCl, 1,8 mM CaCl2, 0.8 mM MgSO4, 20 mM D-glucose and 20 mM HEPES) at 37°C.

Sample preparation for electron microscopy. Cell monolayers on round coverslips (diam. 12 mm) were quickly washed in DMEM without FCS and fixed in situ with 4% paraformaldehyde in 0.1 M Sörensen phosphate buffer (pH 7.4) for 1 h at 4 °C. After washing at 4 °C in the same buffer to remove unbound fixa- tive, the specimens were dehydrated in increasing concentrations of ethanol and subsequently infiltrated with London Resin White (LR White, London resin company limited, Berkshire, England). Coverslips were attached to LR White resin-filled capsules and allowed to polymerize for 48 h at 60 °C. The embedded cells were separated from the coverslips by a short treatment with liquid nitrogen and cut parallel to the coverslip surface with a diamond knife using a Leica Ultracut UCT ultramicrotome. Ultrathin sections were mounted on Formvar-carbon-coated nickel grids (EMS, Warrington, PA, USA) and processed for immunogold labeling or placed on uncoated gold grids for osmium ammine staining. Immunogold labeling for electron microscopy. Ultrathin section were preincubated on a drop of 1% normal goat serum (NGS, Nordic Immunology Laboratories, Tilburg, The Netherlands) in PBS for 3 min at room temperature. Incubation with anti-EGFP polyclonal primary antibodies diluted (1:100) in PBS containing 0.1% bovine serum albumin (BSA, Fluka, Buchs, Switzerland) and 0.05% Tween 20 (Sigma-Aldrich, St. Louis, MO, USA) was carried out for 17h at 4° C in a humid chamber. After rinsing with PBS/Tween and incubating with PBS for 15 min, grids were treated again with NGS as above and labeled for 30 min at room temperature with colloidal gold particle-conjugated secondary antibodies diluted in PBS. Detection of the polyclonal anti-EGFPAb was done by incubating the grids with a goat anti-rabbit (Aurion, Wageningen, The Netherlands) coupled with 15 nm colloidal gold particles (1:3 in PBS; Aurion, Wageningen, The Nether- lands). Grids were finally rinsed with PBS and ultrapure water and subsequently air-dried. For CPD detection, ultrathin sections were treated with 3 M HCl for 20 min at room temperature to denature dsDNA. After washing with water, sections were immunolabelled with a mouse monoclonal anti-CPD antibody (Kamiya Biomedical Company, Seattle, WA, USA) diluted 1/800, using the protocol described above. Detection was carried out using a goat anti-mouse IgG antibody conjugated with 15 nm colloidal gold particles (Aurion) diluted 1/3 in PBS. As controls of labeling specificity, some grids were treated as above except that the

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primary antibody was omitted.

Most preparations were stained with the EDTA regressive technique differential for nuclear nucleoprotein constituents (Bernhard, 1969) in a way which kept chromatin weakly contrasted. They were first stained with 5% uranyl acetate for 1 min, then treated for 3 min with 0.02 M EDTA, and finally with lead citrate for 1 min. To better evaluate if there were differences in the chromatin area between the irradiated and non-irradiated cells, some thin sections were stained using a Feulgen type reaction specific for DNA (Cogliati and Gautier, 1973). They were placed on uncoated gold grids, hydrolyzed with 5N HCl for 20 min at room temperature, rinsed with distilled water and stained with 0.2% osmium ammine solution (saturated with SO2) for 1 h at room temperature. Electron micrographs were acquired with a Philips CM 12 or CM 10 electron microscopes operating at 80 kV, using a 30- to 40-µm objective aperture.

Quantification of CPD and of NER protein distribution. To determine the nuclear distribution of CPDs and repair proteins, we have quantified the signal obtained with anti-CPD antibodies and anti-EGFP antibodies as described above. Electron micrographs were acquired at a final magnification of x31000, using a CCD Morada camera (Olympus Soft Imaging System, Lakewood, CO, USA). The signal in two nuclear regions was quantified: i) condensed chromatin domains (CC) and ii) the perichromatin region (PR). The PR was arbitrarily defined as a layer of 100 nm width, 50 nm on each side of the border of condensed chromatin domains. The labeling intensity of CPDs and repair proteins was expressed as the number of gold particles found in each subnuclear compartment per surface area of the cell nucleus (µm2). This allows averaging of data obtained for different nuclei. The surface area of the nucleus was determined morphometrically using NIH ImageJ 1.34 software (Office of Research Services, Bethesda, MD, USA). The difference in surface area of condensed chromatin domains upon UV irradiation was quantified morphometrically also using NIH ImageJ software. The surface area of condensed chromatin domains was expressed as fraction of the surface area of the whole nucleus.

Estimating the expected accumulation of XPC in dense chromatin domains. The expression level of XPC-EGFP is similar to that of endogenous XPC in wild-type cells and is roughly 25,000 molecules (Araujo et al., 2001). About 20% of the nucleus is occupied by dense chromatin domains (Fig 4), meaning that if homogenously distributed there will be ~5,000 molecules in dense chromatin domains in non-irradiated cells. Global UV irradiation caused about 25% of the XPC-EGFP pool to be immobilized (Vermeulen, unpublished results) corresponding to 6,250 molecules that would accumulate in dense chromatin domains on top of the 5,000 already present. Thus, the expected increase of XPC in dense chromatin domains is 6,250 + 5,000 = 11,250/5,000 = 2.25-fold increase. The measured increase in dense chromatin domains is about 2-fold, which corresponds well with the expected value.

Acknowledgement The authors thank Mrs. J. Fakan and Mrs. F. Voinesco (Centre of Electron Microscopy, University of Lausanne) for their excellent technical assistance, Mr. W. Blanchard for his photographic work, Mrs. L. Hautle for helping in editing the manuscript, Dr. M.J. Moné for initial experiments and Dr. F. Tessadori for critical reading of the manuscript. The authors also acknowledge Dr. J. Goedhart (Swammerdam Inst. for Life Sciences, University of Amsterdam, Amsterdam, The Netherlands) for technical assistance and Drs. E.M.M. Manders and T.W.J. Gadella for support. This work was supported by the Swiss National Science Foundation (grant 3100AO-109333) and by the Netherlands Organisation for Health Research and Development (ZonMW, grant 912-03-012).

96 Chapter 6 Choreography of Nucleotide Excision Repair Unraveled by Live-Cell Imaging and Kinetic Modeling

Martijn S. Luijsterburg, Gesa von Bornstaedt, Antonio Z. Politi, Audrey M. Gourdin, Martijn J. Moné, Daniël O. Warmerdam, Joachim Goedhart, Wim Vermeulen, Thomas Höfer, and Roel van Driel

In preparation

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Abstract Each process that controls the functioning of our genome is carried out by tens of proteins that cooperate in time and space. Examples are transcription initiation, replication machines and DNA repair systems. Remarkably little in known about how such complex systems are assembled and function in the living cell. Therefore, we have carried out a detailed analysis of the kinetics of the nucleotide excision repair (NER) system in living cells, which is responsible for the removal of UV-induced DNA lesions. Results have been integrated in a comprehensive, quantitative and predictive model, which describes the kinetics of DNA repair in terms of the joint action of seven repair factors and involves six enzymatic steps. The model gives novel and detailed insight into DNA repair complex assembly and functioning. Among others, we show that all proteins in chromatin-associated complexes rapidly exchange with soluble pools in the nucleus, making the formation of a functional multi-protein complex a time-consuming process. Kinetic proofreading seems essential to drive the repair process to completion. Since NER is considered a paradigm for other chromatin- associated processes, results can be extended to other systems in the nucleus.

Introduction Nucleotide excision repair (NER) is a versatile DNA repair pathway that removes a variety of helix-distorting lesion from the genome. Placental mammals fully rely on NER for the removal of UV-induced DNA damage such as 6-4 photoproducts (6-4PP) and cyclobutane pyrimidine dimers (CPD) (de Laat et al., 1999; Friedberg, 2001; Hoeijmakers, 2001). NER requires the coordinated action of ~25 polypeptides that assemble and disassemble onto the chromatin fiber and mediate removal of UV- induced DNA lesions (Aboussekhra et al., 1995; Park and Choi, 2006). The general mechanism of NER involves damage recognition, DNA unwinding, excision of a ~30 nucleotides long single-stranded DNA fragment containing the damage and resynthesis of the generated gap by the replication machinery (de Laat et al., 1999). Lesions in actively transcribed genes are removed by a specialized NER pathway known as transcription-coupled NER (TCR), in which stalled RNA polymerase II is the damage recognition signal (Bohr et al., 1985; Mellon et al., 1987). The TCR-specific proteins CSA and CSB play an important role in the initiation of this NER sub-pathway (Fousteri et al., 2006). The majority of lesions, however, are removed by global genome NER (GGR) that removes damages throughout the genome. Damage recognition in GGR is carried out by XPC-HR23B in concert with the UV-DDB complex (Sugasawa et al., 1998; Tang et al., 2000). Binding of XPC to damaged DNA triggers the recruitment of the ten-subunit TFIIH complex that utilizes its helicase activity to locally unwind the DNA surrounding the lesion (Coin et al., 2007). The locally unwound DNA is further stabilized by RPA, XPA and the 3’ endonuclease XPG (Evans et al., 1997b). In the partially unwound state, XPA associates with the DNA lesion, RPA binds to the DNA strand opposite to the damage and XPG stabilizes the emerging repair complex

98 Choreography of NER factors and carries out the 3’ incision (Camenisch et al., 2007; de Laat et al., 1998b; Evans et al., 1997b). Then endonuclease ERCC1/XPF is recruited to the pre-incision complex and carries out the 5’ incision resulting in excision of the DNA-lesion (Park and Choi, 2006; Wakasugi and Sancar, 1999). DNA pol δ is subsequently loaded by PCNA to fill in the generated gap, which is sealed by LigIII-XRCCI to complete repair (Essers et al., 2005; Hoeijmakers, 2001; Moser et al., 2007). The requirements for individual NER factors to assemble onto lesions are well established based on the analysis of mutant cell lines. For example, XPC binding is strictly required for the recruitment of all subsequent repair factors in GGR and open complex formation is needed for the efficient recruitment of XPG and ERCC1/XPF (Moné et al., 2004; Volker et al., 2001; Zotter et al., 2006). Recruitment of repair synthesis factor PCNA and chromatin restoration factor CAF1 depends on dual incision (Essers et al., 2005; Green and Almouzni, 2003). Several cartoon models for NER have been proposed in which repair factors bind seemingly irreversibly until repair is completed followed by the simultaneous release of all NER proteins (Riedl et al., 2003; Volker et al., 2001). Studies on other chromatin-associated processes have indicated that transcription factors and other chromatin-associated proteins exchange rapidly between the bound and free diffusing state. Previous studies on individual NER proteins have shown that repair factors transiently interact with chromatin in vivo (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006), but understanding of the precise choreography of NER factors during repair in living cells is currently lacking. Moreover, little is know about how binding properties of repair proteins change during the course of repair, the rate of the enzymatic reactions during NER and the efficiency of repair complex assembly. Current cartoon models for NER do not explain the dynamic behaviour of this versatile repair system in living cells. In this study, we have systematically analyzed several kinetic parameters of GFP-tagged core NER factors during the repair process in living cells. We show that different NER proteins associate with the DNA lesion with remarkably different binding kinetics. NER proteins continuously and rapidly exchange between the bound and freely diffusing state. To monitor progression of the repair process in time, we have measured the levels of bound NER proteins for several hours following UV irradiation. Based on these and previous quantitative in vivo data, we have developed a comprehensive kinetic model for NER, which quantitatively reproduces many aspects of the repair process in vivo. The model indicates that the NER process progresses through a sequential set of intermediates separated by in practice irreversible enzymatic steps, including unwinding, incision and DNA synthesis. Repair proteins assemble in a stochastic fashion on each intermediate until a catalytically active complex is formed that allows progression to the next repair intermediate. Using this model, we explore the dynamic behaviour and properties of the NER system to gain detailed insight into how it operates in vivo. Among others, the model reveals that the assembly of the repair complex is remarkably slow, since more than 99% of the repair time is spent on

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NER complex formation. Our approach provides detailed insight into the kinetic properties of NER in vivo and unravels the complex choreography of this repair process, which is a paradigm for other chromatin-associated processes.

Results

Approach: quantitative in vivo measurements and kinetic modelling To understand the molecular mechanism of NER in living cells we have systematically measured kinetic parameters of several fluorescent protein (FP)-tagged repair proteins, in living human fibroblasts or CHO cells, such as assembly rates of the NER complex immediately after induction of UV damage, dissociation kinetics and levels of bound NER factors as repair progresses. Based on these and previous quantitative measurements (Hoogstraten et al., 2002; Houtsmuller et al., 1999; Politi et al., 2005; Rademakers et al., 2003; Zotter et al., 2006), we have developed a comprehensive kinetic model for NER that is based on (i) irreversible enzymatic reactions that result in different repair intermediates (e.g. unwound DNA and incised DNA) and (ii) reversible binding of NER proteins to different repair intermediates. Six different repair intermediates are considered based on previous experiments (Evans et al., 1997a; Mu et al., 1997; Riedl et al., 2003; Shivji et al., 1995) (Fig. 4): damaged DNA (Ddam), partially unwound DNA (D*), fully unwound DNA (D+), incised DNA (Din), re- synthesized DNA (Dsynth) and chromatinized DNA (Dchrom). The stepwise formation of these 6 repair intermediates by irreversible enzymatic reactions is a concept that is useful to interpret the in vivo measurements. Moreover, this concept is the basis for a comprehensive kinetic model that quantitatively reproduces many aspects of NER in vivo. To verify and improve the model we have measured kinetics of repair proteins in various NER-deficient cell lines, as well in the presence of repair synthesis inhibitors. The model is interrogated to obtain detailed insight into the in vivo functioning of this chromatin-associated process.

Estimating the concentration of DNA damage sites following local UV irradiation Measurements of UV-induced DNA lesions produced in naked DNA and in E. coli genomic DNA have suggested that the amount of lesions increases linearly with the UV dose (Chandrasekhar and Van Houten, 2000; Kuluncsics et al., 1999). However, similar measurements on chromatinized DNA in mammalian cells have yielded conflicting results (Perdiz et al., 2000; van Hoffen et al., 1995). Because the concentration of DNA lesions is an important parameter, we have carried out experiments to estimate the concentration of DNA lesions produced under our experimental conditions. Since the bound fraction of repair proteins depends on the concentration of DNA lesions in the cell, we have studied the behaviour of GFP-tagged repair proteins at different UV fluencies and by irradiating areas of different size of cell nuclei. The amount of XPG- GFP (Zotter et al., 2006) in CHO cells that were locally irradiated through 5 µm pores

100 Choreography of NER factors did not increase significantly between 100 and 200 J.m-2 and was never more than ~10% of the XPG pool (Fig. 1A). Similar results were obtained with several other repair proteins, showing that NER appears to saturate under these conditions (data not shown). Two hypotheses could explain these results: (i) the amount of damages increases with the UV dose and one or more of the involved proteins become limiting for repair (Politi et al., 2005), or (ii) the amount of induced damage saturates at UV doses over 100 J.m-2. To test the two hypotheses, cells were irradiated at 100 J.m-2 using 3, 5 and 8 µm pores (corresponding to irradiated areas of 7, 20, and 50 µm2, respectively; Fig. 1C). In the presence of a limiting factor, we expect no increase in the bound fraction for different irradiated areas. However, we found that the amount of XPG involved in repair increased linearly with the irradiated surface and exceeded the 10% of the XPG pool (4.7%, 9.1% and 21.8%; Fig. 1B). These results are in agreement with the second hypothesis: none of the NER proteins becomes limiting under our experimental conditions and the amount of DNA lesions saturates at higher UV dose. We estimate (based on data of (Perdiz et al., 2000)) that local UV irradiation at 100 J.m-2 through 5 µm pores (our standard condition) introduces approximately 5.104 6-4 PPs (Fig 1D), corresponding to a nuclear concentration of about 0.3 µM.

The kinetics of NER proteins recruitment to damaged DNA We have previously established a novel method to measure assembly of the NER complex in living cells immediately following local UV-C irradiation (Moné et al., 2004). The initial slope of the measured binding curves is indicative of the rate of assembly of repair complexes. Employing this method, we measured the assembly

Fig 1. Estimation of the amount of 6-4 PPs under experimental conditions. A) UV-C dose (in J.m-2) response curve (measured in % of protein bound in the locally irradiated area) of UV135 cells expressing XPG-EGFP locally irradiated through 5 µm irradiated at UV-C fluencies ranging from 50 – 200 J.m-2. B) Quantification of percentage of XPG-EGFP bound following local UV-C irradiation at 100 J.m-2 using pores of 3, 5 or 8 µm. C) Representative images of XPG-EGFP cells locally irradiated at 100 J.m-2 using 3, 5 or 8 µm. D) Estimate of the amount of 6-4 PPs produced at different UV fluencies (including experimental conditions employed in this study) based on extrapolation of data by Perdiz and co-workers (Perdiz et al., 2000).

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kinetics for ERCC1-GFP (t1/2 ~ 65) and TFIIH-EGFP (t1/2 ~ 113s) and showed that the incorporation of both proteins in repair complexes occurs with similar rates (~20- 30 molecules/s) on UV-damaged chromatin (Moné et al., 2004). To extend this analysis, we have measured the assembly kinetics of XPC-EGFP, XPG-EGFP and EGFP-XPA in human cells (Fig. 2A). All three GFP-tagged NER proteins are stably expressed in NER-deficient cells at levels similar to wild-type levels (Hoogstraten et al., 2003; Politi et al., 2005; Rademakers et al., 2003; Zotter et al., 2006). Binding of XPC-EGFP and XPG-EGFP started immediately after local UV-C irradiation at 100 J.m-2 through 5 µm pores and reached a maximum within 10 minutes with a t1/2 of 100 and 200 seconds, respectively (Fig 2B). Surprisingly, binding of XPAwas considerably slower and reached a maximum only after 1 hour with a t1/2 of 600 seconds (Fig 2B). Given that the concentrations of XPC (3.104 molecules per cell), XPG (8.104 molecules per cell) and XPA (2.105 molecules per cell) are different (Araujo et al., 2001), which was confirmed by flow cytometry (data not shown) and western blot analysis (Hoogstraten et al., 2003; Rademakers et al., 2003; Zotter et al., 2006), we estimate that the assembly rate of all three proteins is ~15 molecules/s, which is in the same range as previously reported for TFIIH and ERCC1 (Moné et al., 2004).

Our experiments revealed that the maximal level of bound XPA and its t1/2 of association are considerably higher than those of XPC and XPG (Fig 2B). This prompted us to investigate how the amount of bound NER factors changes during the time period in which the majority of 6-4 PPs are removed (van Hoffen et al., 1995). To this aim, we locally irradiated cells at 100 J.m-2 through 5 µm pores and monitored the level of bound protein during 5 hours after UV irradiation. After reaching a maximum, the bound XPC-EGFP and XPG-EGFP levels gradually decreased with a t1/2 of ~1 hour (Fig 2B). A similar decrease in bound ERCC1-GFP (t1/2 ~ 1 hr) has been described elsewhere (Politi et al., 2005). In contrast, the bound level of EGFP-XPA stayed high for a considerable longer time (t1/2 ~ 2.5 hr) and decreased to background levels only after about five hours (Fig. 2B). It should be noted that the levels of bound repair proteins never fully returns to zero on the time-scale of a few hours, possibly reflecting repair complexes that are assembled on slowly repaired CPDs. In addition to several pre-incision NER proteins, we also monitored the bound levels of repair synthesis protein EGFP-PCNA (Essers et al., 2005) and EGFP-RPA (Gourdin and Vermeulen, unpublished) during 5 hours. The levels of bound EGFP-PCNA and EGFP-RPA continued to increase for several hours and reached a maximum with a t1/2 of over 1 hr (Fig. 2C and not shown). PCNA and RPAbound levels did not decrease significantly within 9 hrs following UV irradiation (Fig. 2B).

These results show that, although NER proteins incorporate in nascent NER complexes with similar initial rates, the overall kinetics of pre-incision protein XPC, XPG, and ERCC1 is remarkably different from that of XPA, RPA and PCNA. The maximal

102 Choreography of NER factors amount of bound XPC, XPG and ERCC1 is reached ~10 min after irradiation, whereas the maximal amount of bound PCNA and RPA is reached much later (after 4 hrs).

Fig 2. Levels of bound NER proteins during several hours after UV irradiation. A) Representative examples of cells expressing XPG-EGFP, EGFP-XPA and EGFP-PCNA at various time-points after local UV irradiation at 100 J.m-2 using 5 µm pores. B) Quantification of levels of bound XPC-EGFP, XPG-EGFP and EGFP-XPA during 5 hours following local irradiation. The curves are normalized to absolute number of proteins given by the bound fraction (%) times the nuclear amount (in molecules) of these repair factors as measured previously (see table 3 and Araujo et al., 2001). C) Quantification of levels of bound of EGFP-PCNA during 5 hours following local irradiation, normalized to 1.

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XPA displays intermediate behaviour and the maximal amount of bound XPA is reached after ~ 1 hr. Furthermore, the bound level of XPC, XPG, and ERCC1 returned to background level within 1hr after damage induction, XPA required more than two times longer, whereas PCNA and RPA were still bound to DNA after 9 hr. As we will show later, this remarkable behaviour of NER proteins can be fully resolved within the framework of the kinetic model.

Dissociation kinetics of NER proteins The core NER factors are engaged in repair for several hours following UV irradiation. This is a highly dynamic process in which NER proteins continuously exchange between the bound and free diffusing state. To gain insight into the rate at which NER proteins exchange, we determined dissociation kinetics of NER proteins from locally damaged sites using FLIP (Fig 3A; Luijsterburg et al., 2007). Dissociation of EGFP- tagged pre-incision proteins XPC, TFIIH, XPG, XPA, RPA and ERCC1/XPF from repair complexes was complete on a minute time-scale and we determined dissociation half-times (t1/2) of 20s (RPA), 25s (XPC), 50s (TFIIH, XPG, ERCC1/XPF) and 80 s (XPA) (Fig 3B). These results reveal that all pre-incision factors exchange rapidly between the bound and freely diffusing state and are incorporated only shortly in repair complexes. Curve fitting showed that dissociation of RPA, XPG and ERCC1 is mono-exponential and that dissociation of XPC (t1/2 of 9s and 50s) and XPA (t1/2 of 52s and 190s) is biphasic, suggesting that at least two distinct dissociation rates contribute to the measured dissociation kinetics. Dissociation kinetics of repair synthesis factor PCNA displayed strong biphasic behaviour with a t1/2 of 10s and 225s for each component, which is in agreement with previous measurements (Fig. 3C) (Essers et al., 2005). These results show that various NER factors dissociate with different kinetics from repair complexes: XPC and RPA dissociation is two to three- fold-fold faster than that of TFIIH, XPG and ERCC1, whereas dissociation of XPA is almost two-fold slower. PCNA dissociation is 5 to 10-fold slower than that of pre- incision proteins. These differences in dissociation kinetics are likely to reflect the affinity of NER proteins for different repair intermediates or emerging repair complexes. To understand for which repair intermediates NER proteins have affinity, we analyzed dissociation kinetics of NER factors in various mutant cell lines and in the presence of drugs that inhibit DNA repair synthesis. Both approaches block the formation of specific repair intermediates, which allows us to test whether NER factors have affinity for these intermediates.

XPC affinity is lowered upon full pre-incision complex formation Several studies have suggested that the affinity of XPC for repair complexes is lowered by the association of a specific NER factor (for example XPG) or by DNA unwinding (open complex formation) favouring dissociation of XPC (Riedl et al., 2003; Tapias et al., 2004). To investigate if the dissociation rate of XPC depends on open complex

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Fig 3. Dissociation kinetics of NER proteins. A) Represen- tative examples of cells expressing EGFP-XPA and EGFP-PCNA after local irradiation, which are continuously bleached opposite to the local damage until all fluorescent molecules are bleached. B) Quantifica- tion of FLIP experiments on locally irradiated cells expressing XPC-EGFP, XPG-EGFP, EGFP- XPA, RPA-EGFP and ERCC1-GFP, which is a measure for the dissociation kinetics of the investigated proteins. Quantification of FLIP experiments in the absence or presence of HU and AraC on locally irradiated cells expressing C) EGFP-PCNA and D) EGFP-XPAor ERCC1- GFP. E) Quantification of FLIP experiments on XPC-mVenus expressed in various locally irradiated NER-deficient CHO and human cell lines (see material and methods for details on the cells). formation or specific NER factors such as XPG, we have tagged XPC with the YFP variant mVenus (Kremers et al., 2006) and transiently expressed the fusion protein in various NER-deficient human and CHO cell lines and subsequently tested if the dissociation kinetics of XPC is changed. We specifically compared cells that are deficient and proficient in open complex formation (such as XP-B and XP-F cells, respectively) and cells lacking functional XPG. Dissociation kinetics of XPC from locally UV-irradiated sites when XPC binding had reached its maximum was ~4-fold slower (t1/2 ~ 100s) in cells with mutations in XPB, XPA and XPG compared to wild type cells (t1/2 ~ 25s; Fig 3E). Biochemical studies have shown that these proteins are essential for open complex formation in contrast to ERCC1/XPF, which is not required (Evans et al., 1997b). Similar slow XPC dissociation kinetics was measured in cells with compromised ERCC1 or XPF proteins (Fig 3E), which are proficient in open complex formation (Evans et al., 1997b). These results suggest that the affinity of XPC is not lowered following open complex formation or by the association of a specific NER protein. Our results are consistent with a scenario in which the affinity of XPC is lowered after dual incision, or at least until a full pre-incision complex is assembled. Consequently, we conclude that in addition to damaged DNA, XPC binds reversibly to the complexes associated to unwound and incised DNA (Fig 4).

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XPA has affinity for repair-synthesis intermediates The high levels of bound XPA several hours after UV irradiation after the levels of bound XPC, XPG and ERCC1 have returned close to background levels, prompted us to investigate whether XPA is bound to repair intermediates to which XPC, TFIIH, XPG and ERCC1/XPF are not bound. Given that XPA strongly binds RPA, which is involved in DNA repair synthesis (Li et al., 1995a; Shivji et al., 1995), we determined whether XPAhas an affinity for repair intermediates in the DNA repair synthesis stage, i.e. late in the repair process. To this end, we measured dissociation kinetics of several pre-incision factors and of PCNA in the presence of DNA synthesis inhibitors hydroxyurea (HU) and cytosine-β-arabinofuranoside (AraC). Both drugs strongly inhibit repair- patch synthesis and subsequent ligation following UV irradiation (Mullenders et al., 1987; Smith and Okumoto, 1984). Cells were treated with drugs for 30 min about 1 hr after local UV irradiation, after which dissociation kinetics were measured by FLIP (Luijsterburg et al., 2007). EGFP-PCNA dissociation was significantly slowed down in the presence of synthesis inhibitors (Fig 3C), consistent with its participation in repair synthesis (Shivji et al., 1995) and binding to incised DNA. Treatment of cells expressing XPC-EGFP (data not shown) or ERCC1-EGFP (Fig 3D) with HU and AraC did not result in a detectable change in dissociation kinetics of these proteins. In contrast, EGFP-XPAdissociation became significantly faster when DNA synthesis was inhibited (Fig 3D). Curve fitting showed that the biphasic dissociation kinetics of XPA (t1/2 of 52s and 190s) in untreated cells became mono-exponential (t1/2 of 40s) upon treatment with HU and AraC, suggesting that the slower dissociation reaction occurs after repair synthesis and the faster one before. These results suggest that, in contrast to the other pre-incision proteins, XPAhas an affinity for repair synthesis intermediates, possibly through the interaction between XPA and RPA (Li et al., 1995a; Moser et al., 2007). It also shows that XPA has a different affinity for repair intermediates before repair synthesis compared to after repair synthesis. In agreement with these findings, chromatin immuno-precipitation experiments using antibodies against the late NER protein XRCCI , which forms a complex with Lig III, pulled down RPA and XPA but not XPC or TFIIH, supporting the idea that XPA binds to repair intermediates after repair DNA synthesis (Moser et al., 2007). Consequently, we conclude that RPA, PCNA and XPA bind reversibly to the intermediate characterized by re-synthesized DNA, but that the other pre-incision proteins have no affinity for re-synthesized DNA (Fig 4).

Description of the kinetic model Based on the above and other quantitative in vivo measurements (Moné et al., 2004), we have developed a mechanistic kinetic model for mammalian NER. The model only considers GGR and not TCR, since the contribution of the latter pathway to the total NER activity in cells after UV irradiation is less than 1% (Moné et al., 2004). Below we first outline the overall structure of the model. Its framework is formed of six repair

106 Choreography of NER factors intermediates, characterized by six different states of the DNA during the repair process (indicated by the green boxes in Fig. 4; table 1). The transition between repair intermediates is mediated by enzymatic steps (red arrows in Fig. 4; table 1). Because these steps are essentially irreversible, they in principle drive the repair process to completion. Each enzymatic step can occur only after the correct complement of proteins is assembled on the DNA (the proteins indicated in the box corresponding to each intermediate).

Box I: formation of the DNA-XPC-TFIIH complex NER is initiated by the binding of XPC to lesions and subsequent recruitment of TFIIH (Box I; (Volker et al., 2001; Yokoi et al., 2000)). Although RPA and XPA have been suggested to initiate NER (Wakasugi and Sancar, 1999), this is not supported by in vivo experiments (Rademakers et al., 2003; Volker et al., 2001). The time-dependent changes in the concentration of free damage ([Ddam]), XPC bound to damage ([DdamC]) and XPC and TFIIH bound to damage ([DdamCT]) can be described by (notations are

Fig 4. Framework of the kinetic model for NER. The framework for NER consists of six repair intermediates (represented by green boxes) and irreversible enzymatic steps (indicated by red arrows). The notations used in the model, such as abbreviation of proteins and symbols for enzymatic reactions are given in table 1. For details, see the main text.

107 Chapter 6 given in table 1 and 2):

(1)

(2)

(3)

The Ddam-XPC-TFIIH complex is able to partially unwind the DNA (reaction α) resulting in D*-XPC-TFIIH (Evans et al., 1997b). Unwinding of DNA is reversible and if unwound DNA is devoid of any proteins it will re-anneal back to damaged DNA via reactions ε1 and ε2.

Box II: formation of a partially unwound DNA Box II represent the partially unwound repair intermediate (Fig 4), to which repair proteins XPG, XPA and RPA, as well as XPC and TFIIH, bind reversibly. All proteins bind and dissociate independently, Proteins except for ERCC1/XPF, which C XPC can only bind to repair complexes T TFIIH that contain XPA, consistent with G XPG A XPA data from intact cells (Volker et F ERCC1-XPF al., 2001). In contrast to the R RPA sequential binding of XPC and P PCNA Enzymatic recations TFIIH to damaged DNA (Ddam), α initial unwinding we assume random complex α' full inwinding assembly on unwound DNA. In β dual incision γ repair synthesis agreement, pre-melted lesions are δ re-chromatinization repaired without XPC, showing ε1 re-annealing partially unwound DNA that XPC is not strictly required ε2 re-annealing fully unwound DNA Repair intermediate for the binding of subsequent Box I: Ddam damaged DNA NER proteins (Mu and Sancar, Box II: D* partially unwound DNA (~8 nt) Box III: D+ fully unwound DNA (~30 nt) 1997). However, XPC is strictly Box IV: Din incized DNA required to produce the partially synth Box V: D re-synthesized DNA unwound repair intermediate in Box VI: Dchrom re-chromatinzed DNA the first place, to which sub Table 1. Notations for proteins, intermediates and enzymatic sequent NER factors can bind. reactions. This table lists abbreviations for repair proteins and This explains why binding of all repair intermediates and symbols for enzymatic reactions used in this study. NER proteins depends on XPC in

108 Choreography of NER factors

Specific notations (1-3) [Ddam] Concentration of damaged DNA (µM) c -1 -1 kon On-rate constant of XPC (µM .s ) [C] Free concentration of XPC c -1 Koff Off-rate constant of XPC (s ) [DdamC] Concentration of damaged DNA bound by XPC (µM) t -1 koff Off-rate constant of TFIIH (s ) [DdamCT] Concentration of damaged DNA bound by XPC and TFIIH (µM) t -1 -1 kon On-rate constant of TFIIH (µM .s ) [T] Free concentration of TFIIH (µM) α Catalytic rate constant of partial unwinding of the DNA (s-1) General notations [X] Free concentration of protein X x -1 -1 kon On-rate constant of protein X (µM .s ) x -1 koff Off-rate constant of protein X (s ) ζ Catalytic rate constant of enzymatic reaction ζ (s-1) Specific notations (4-5) τ Characteristic time to fill a repair intermediate box (s) y0 Lesion concentration at time-point t (µM) t Time (s)

Table 2. Description for terms in equations. This table defines the terms in differential equations (1) through (5). vivo (Rademakers et al., 2003; Volker et al., 2001). Box II represents 48 different states, i.e. 47 different protein assembly states and naked partially unwound DNA. The rationale is that 6 protein can theoretically be assembled on D* (26 states), however, this number is reduced (with 24) to 48 states (26 - 24) since ERCC1-XPF only binds complexes containing XPA. Unwinding of DNA is in principle reversible and if the DNA is devoid of any protein (such as D* in box II and D+ in box III) it will re-anneal dam * back to D (Box I). We consider two possible re-annealing reactions: ε1 (from D ) and + ε2 (from D ). Differential equations similar to the ones shown in equations (1) through (3) are obtained for the time-dependent changes in concentration of all possible DNA states.

Box III: fully unwound DNA We assume a two-step unwinding model as suggested by Evans et al. (Evans et al., 1997b). Full unwinding was shown to require XPC, RPA, XPA, XPG, but not ERCC1/XPF. Likewise, in the model full unwinding (reaction α’) occurs when XPC- TFIIH-XPG-XPA-RPAare present simultaneously. In the fully unwound state (D+; box III), we again assume reversible and mutually independent association-dissociation of the pre-incision factors XPC, TFIIH, XPG, XPA, RPA, whereas ERCC1/XPF again only binds to complexes containing XPA. Hence, box III represents 47 protein

109 Chapter 6 assembly states on D+ and naked fully unwound DNA.

Box IV: Dual incision Dual incision of fully unwound DNA (Box III) requires the endonucleases XPG and ERCC1-XPF and dual incision is stimulated by TFIIH, XPA, RPA and possibly XPC (Evans et al., 1997b; Winkler et al., 2001). In the model, incision is carried out when D+-TFIIH-RPA-XPG-XPA-ERCC1/XPF or D+-XPC-TFIIH-RPA-XPG-XPA-ERCC1 /XPF is formed. Incised DNA (Din; Box IV) is a substrate for reversible binding of all pre-incision proteins. In addition, PCNA can bind reversible to Din since binding of PCNA requires a free 3’-OH group, which is generated by the ERCC1-XPF incision. In the model, 95 different protein complexes can be assembled in Box IV.

Box V: Repair synthesis Repair synthesis (Dsynth; Box V) is carried out after Dsynth-XPA-RPA-PCNA is assembled (reaction γ1). We assume that XPAis bound during repair synthesis or binds to repair intermediates after repair synthesis, since the dissociation kinetics of XPA is severely affected by inhibition of repair synthesis (i.e. step γ in the model). In contrast, this treatment has no effect on the other pre-incision proteins (Fig 3). This assumption is also supported by ChIP experiments with antibodies against the late NER factor XRCC1-Lig III, which pulled down RPA and XPA, but not XPC and TFIIH (Moser et al., 2007). Requirement of RPAand PCNA to perform repair synthesis has been shown previously (Shivji et al., 1995). Repair factors RPA, XPA and PCNA bind reversibly and independently to re-synthesized DNA (Dsynth; box V). Box V contains 8 assembly states: up to 7 different protein complexes and Dsynth devoid of proteins.

Box VI: completion of the repair process Progression to a chromatinized repair intermediate (Dchrom) via enzymatic reaction δ occurs when Dsynth-RPA-PCNA is formed. In the model, RPA and PCNA bind reversibly to the Dchrom, while all other NER proteins have no affinity for this repair intermediate. In agreement, levels of bound EGFP-PCNA, EGFP-RPAand EGFP-RFC (Gourdin and Vermeulen, unpublished) are high up to at least 4 hrs after UV irradiation. At that time other repair proteins are no longer bound.

Kinetic model The complete model consists of 214 coupled differential equations, which describe the concentration of the different assembly states in time (207 equations) as well as the time-dependent changes in the concentration of repair factors (7 equations). By lack of experimental evidence, we do not assume cooperative binding. Repair proteins have a single kon and koff for each repair intermediate. However, we assume that XPC, TFIIH, * XPG, XPA and ERCC1/XFF have the same kon and koff for partially (D ) and fully + unwound DNA (D ). Likewise, we assume the same kon and koff values for RPA

110 Choreography of NER factors binding to fully unwound (D+) and incised DNA (Din) since both these repair inter mediates contain single-stranded DNA. The initial lesions concentration is 60.000 6-4 PPs based on previously described estimates (Fig 1D).

Quantitative parameters of the model We fitted the model (Fig 4) to the measured in vivo binding kinetics and dissociation kinetics of repair proteins XPC, XPG, XPA, RPA, PCNA and previously measured kinetics on TFIIH and ERCC1-XPF (Hoogstraten et al., 2002; Moné et al., 2004). Concurrently, the model was fitted to dissociation kinetics of XPC in NER-deficient cells, as well as to dissociation kinetics of PCNA, RPA and XPA in the presence of repair-synthesis inhibitors. Protein concentrations (all in the range of 0.3-1.4 µM) were taken from the literature (Araujo et al., 2001) or estimated (table 3). Changes in free protein concentration in time due to transient binding to repair intermediates were included in the model. It should be noted that the measured in vivo binding and dissociation kinetics of individual NER factors is the result of an ensemble of different association rate constants ( kon) and dissociation rate constants ( koff), since repair proteins not necessarily bind different repair intermediates with the same affinity. For example, the affinity of RPA is higher for fully unwound DNA compared to partially unwound DNA (Blackwell and Borowiec, 1994). Using the mathematical model is it possible to untangle the complex interactions between the different reactions and derive kon and koff values for individual NER factors for each repair intermediate, and estimate the kcat of the enzymatic reactions that occurs throughout the enzymatic process (Table 3). Hence, modelling of the in vivo measurements on NER protein dynamics allows one to derive key biochemical parameters that cannot be measured directly in vivo.

Simulation of the NER process and comparison to experimental data A critical test of the model is whether it can reproduce the in vivo kinetics of NER. The model faithfully reproduces the binding kinetics of NER factors over a period of 5 hrs in addition to the relatively fast dissociation kinetics of all repair factors (Fig 5A, B, C). Moreover, it predicts the long-term binding behaviour of TFIIH and ERCC1-XPF during the repair course (Fig 5A). The in vivo data could not be fitted with a scheme in which NER factors assemble in a strict sequentially fashion and dissociate simultaneously, as suggested previously (Politi et al., 2005). Although the measured binding rates could be reproduced partially with such scheme, the predicted dissociation kinetics of repair factors was quite different from the measured values. If binding would be strict sequential, XPC was predicted to dissociate with a t1/2 > 10 min while the ERCC1-XPF dissociation t1/2 would be < 1s (data not shown). The random and reversible assembly of repair factors on various repair intermediates explains the measured in vivo kinetics of the NER factors much better.

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Enzymatic rates during NER Fitting the model to the recruitment and dissociation kinetics of all repair proteins yielded binding rate constants (kon) and dissociation rate constants (koff) for all NER factors. It also allowed estimation of catalytic rate constants ( kcat) for the enzymatic reactions (table 3). The fits for the catalytic rates are in the expected range for such enzymatic reactions and slower enzymatic rates could not reproduce the experimental data. Characteristic times for the enzymatic steps (calculated as 1/ kcat) show that the

Fig 5. Modelling the kinetics of NER proteins based on experimental data. A) Binding kinetics of various GFP-tagged NER proteins (indicated by diamonds) and fitted curves by the mathematical model (lines) to the data. The curves are plotted in proteins bound in the locally damaged area in µM, based on the concentrations of NER proteins given in table 3. B) Dissociation kinetics of various GFP-tagged NER proteins (diamonds) together with fitted curves by the model (lines). C) Data (diamond) and fitted curves (lines) of FLIP experiments in the absence or presence of HU and AracC, and of XPC-mVenus in wild-type and XPF-deficient cells. partial (α) and full unwinding (α’) of DNA each take 9 s and 2 s, respectively, whereas dual incision (β) is faster (0.6 s, see table 3 for kcat values). In agreement, a viral helicase has previously been shown to unwind short DNA duplexes with a kcat similar to our fitted value (Porter et al., 1998). Re-annealing of partially (ε1) or fully unwound (ε2) DNA is fast and takes 0.1 s and 0.3 s, respectively. The re-synthesis of the DNA (γ) takes about 30 s (Fig 4) and the final enzymatic step in the model representing

112 Choreography of NER factors chromatin restoration (δ) takes ~80 s. Thus, the enzymatic reactions during NER (given by the sum of 1/kcat values), including unwinding, dual incision, repair synthesis and chromatin restoration take about 2 min to complete.

Affinity of NER proteins for repair intermediates In addition to catalytic rates, kon and koff values for binding and dissociation of NER proteins (given in table 3) allowed us to calculate the affinity (calculated as Kd = koff/kon) of repair proteins for each repair intermediate. The Kd values (nM to µM range; see Fig 6B and table 3) are all in the excepted range (ref) for chromatin- associated proteins. From these calculations, it follows that XPG, TFIIH and ERCC1/XPF bind with high affinity to unwound DNA, but have at least one order of magnitude lower affinity for incised DNA. Thus, these proteins loose their affinity after dual incision. In contrast, the affinity of RPA to incised DNA is increased and the affinity of XPA is only moderately decreased, which explains why the bound levels of these proteins remain high for extended periods of time.

Time-dependent changes in concentration of repair intermediates The change in concentration of the different repair intermediates as a function of time can be calculated from the model (Fig 6A). It is given by the sum of the concentrations of all protein assembly states on a given repair intermediate. For example, the amount of Ddam at any time-point is given by [Ddam] + [ Ddam C] + [ Ddam CT]. Under the given experimental conditions of concentration of lesions and of NER proteins, half of the damaged sites (Ddam) are converted to unwound DNA within ~30 min as predicted by the model (Fig. 6A). The rate of lesion removal is predicted by the model based on primary biochemical parameters derived from the association and dissociation kinetics of repair factors measured in vivo. Direct in vivo measurements of the rate at which 6-4 PPs are removed gave very similar values (Moser et al., 2005; van Hoffen et al., 1995). This independent experimental result supports our model and indicates that the obtained parameters are realistic. The model predicts that partially unwound DNA and especially fully unwound DNA never reach high levels, since the enzymatic reactions to unwind and incise (i.e. kcat of α, α’ and β) are relatively fast. It should be noted that kcat values provide information on how fast enzymatic reactions are carried out once a catalytically active protein complex is assembled. The kcat values do not reflect the time it takes to assemble such an active complex. A maximal amount of incised DNA (Din) accumulates around 1 hour after UV irradiation. After that, the levels of Din gradually decrease over a period of 5 hrs (Fig 6A).

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A quantitative measure for the rate at which a given repair intermediate is produced is the characteristic time (τ), which indicates when half of the initial lesion concentration is converted to a given repair intermediate. For example, the characteristic time (τ) for Din is given by (see also table 2 for notations):

(4)

The standard deviation (σ) for τ is given by:

(5)

From equation (4) and (5) it follows that the characteristic time (τ) for incised DNA is 40 ± 37 min. To determine τ, the enzymatic reaction to the next repair intermediate is blocked. For example, when both γ reactions are set to zero, half of the original amount of damages (Ddam) is converted to Din in 40 ± 37 min. This indicates that there is a high variation from lesion to lesion in the time it takes to be incised. Some lesions are incised within minutes after UV irradiation while others take up to 1 hour to be incised. It should be noted that τ reflects the time it takes to assemble a catalytically active complex and the time to carry out the enzymatic reactions, which is discussed in more detail in the next section. At present, it is not possible to obtain reliable τ values for D* and D+, because the re-annealing (ε) reactions of unwound DNA back to Ddam do not allow this calculation. It follows from equation (4) and (5), in which [Din] is substituted for [Dsynth] and [Dchrom], that τ for Dsynth and Dchrom is 2.5 ± 2.1 and 3.0 ± 2.0 hours, respectively. This shows that it takes a considerable time to re-chromatinize DNA that has been repaired as predicted by the kinetic model. In agreement with these values, DNA repair synthesis (reflecting Dsynth) still occurs 5 to 6 hours after UV irradiation (Rubbi and Milner, 2001; Vincent et al., 2001). Moreover, incorporation of histone H3 at sites of repair (reflecting Dchrom) and binding of CAF1 occurs up to 6 hours after DNA damage induction and possibly longer (Polo et al., 2006). Additionally, results indicate that there is a high degree of variation between lesions in the time it takes to be converted to the Dsynth and Dchrom repair intermediate. The τ values reported here for Din, Dsynth and Dchrom are independent of the concentration of lesions. Hence, these values reflect an intrinsic property of the NER system.

Repair complex assembly on repair intermediates is slow Deriving key parameters from the experimental data, such as catalytic rates and changes in repair intermediate concentration in time, allows us to gain detailed insight into the molecular mechanism of NER in vivo. For example, it follows from the model

114 Choreography of NER factors that the enzymatic steps to produce incised DNA (Din, given by the sum of α, α’ and β) take only ~12 s. On the other hand, the characteristic time (τ) to produce Din is roughly 40 min, which means that half of the damages are incised in the first 40 min following irradiation. Therefore, to produce Din, less than 1% of the time is spent on the enzymatic reactions and more than 99% of the time is spent on the assembly of a catalytically active protein complex. All pre-incision repair factors continuously bind to and dissociated from to the complex characterized by fully unwound DNA. Therefore, a complex that contains all proteins and is competent to perform dual incision is formed only at low probability. The majority of complexes that are assembled are not competent to carry out dual incision and require additional

Fig 6. Concentration of repair intermediates and affinity of NER proteins derived from the model solutions. A) Model prediction of the concentration of repair intermediates (expressed in µM) in time derived from fitting the model to the data shown in Fig 5. B) Affinity of repair factors for different repair intermediates (measured as Kd in µM), derived from fitted on-rates constants and off-rate constants, calculated as Kd=koff/kon.

115 Chapter 6

Table 3. Kinetic parameter and calculations derived from model solutions. The nuclear amounts of NER factors XPC, XPA and XPG is based on published data (Araujo et al., 2001), while RPA and PCNA amounts were estimated at 250,000 molecules per cell, and TFIIH and ERCC1-XPF were estimated at 65,000 and 50,000 molecules per cell, respectively, based on previous estimates (Houtsmuller et al., 1999; Moné et al., 2004). The nuclear concentrations are calculated assuming a nuclear volume of 0.3 pL. Using the model, the association and dissociation rate constants for NER factors for each repair intermediate are estimated by fitting to the experimental data. Subsequently, Kd values are calculated by Kd = koff/kon. Catalytic rate constants (Kcat) are derived from fitting the model to the data, and characteristic times for enzymatic reactions are given by 1/kcat. The time to produce repair intermediates (τ), as calculated using equations (4) and (5), is also listed. For a description of abbreviations see table 1. association/dissociation events. This indicates that the fast exchange of NER factors between the bound and freely diffusing state does not necessarily reflect involvement in DNA repair, but rather reflects dissociation and re-binding events that occasionally result in the assembly of a catalytically active complex. Similarly, the enzymatic steps to produce Dsynth take less than 1 min, while τ for this repair intermediate is roughly 2.5 hours. Hence, less than 1% of the time is spent on enzymatic reactions, whereas by far most time (>99%) is spent on assembly of complexes that are capable of performing repair synthesis. The overall repair process is kinetically driven by six in practice irreversible reactions. Such mechanism, known as kinetic proofreading (Hopfield,

116 Choreography of NER factors

1974; Reardon and Sancar, 2004), confers high specificity to the repair process in which substrate specificity is not present at the level of the individual repair factors, which have only limited substrate specificity. Consequently, the seemingly slow repair complex assembly is an efficient quality control tool (Kanaar et al., 2008).

Discussion

In this study, we have systematically determined kinetic parameters of seven NER factors in the nuclei of living cells. Results have been integrated in a comprehensive, quantitative and predictive model, which describes the kinetics of DNA repair in ters of the cooperative action of seven repair proteins and involves six enzymatic steps. We currently do not include the activity of UV-DDB or chromatin remodelling prior to NER in the model since it is not understood how these processes influence the kinetics of NER. Clearly, NER is more complex than the mechanism that we propose in this study. Additional steps include, amongst others, recruitment of RFC, DNA polymerase δ or ε, XRCC1-LigIII and CAF1 to restore the chromatin structure, and chromatin remodelling such as NER-specific H2A ubiquitylation (Bergink et al., 2006; Green and Almouzni, 2003; Moser et al., 2007). Nonetheless, the combined approach of live-cell imaging and kinetic modelling allows us to unravel general characteristics of the NER process overall. This approach provides novel and mechanistic insight into DNA repair complex assembly and functioning, which will be disused in more detail below.

No stepwise directional assembly of repair complexes It is the often-held view that assembly of the repair complex is sequential. In other words, there is a strict order in which repair factors assembly into repair-competent complexes at the site of DNA damage from freely diffusion constituents (Riedl et al., 2003; Volker et al., 2001). Mathematical modelling of NER reveals that directional sequential assembly would result in pre-incision factor dissociation kinetics that are very different than observed experimentally. For instance, modelling shows that XPC dissociation kinetics would be much slower (>800s) while ERCC1-XPF would dissociate rapidly (<1s) if protein assembly is strict sequential (assuming reversible binding) (data not shown). The rationale is that XPC will be the first protein to bind to a lesion and it will remain bound, while subsequent proteins are incorporated into the nascent repair complex. Since this involves more than six repair factors it may take a considerable time to assembly a fully functional repair complex, containing all proteins at the same time. Hence, the dissociation kinetics of XPC would be much longer than that of a later protein (e.g. ERCC1-XPF) that binds to the nascent NER complex. However, in vivo measurements indicate that XPC and ERCC1-XPF bind for 25s and 50s, respectively, arguing against stepwise sequential assembly until a repair complex is fully assembled. Accordingly, in vivo measurement of GFP-RPA show that this

117 Chapter 6

protein exchanges rapidly (t1/2 of 20s) at sites of damage with a similar rate as the damage-recognition protein XPC. However, RPA is a pre-incision as well as a DNA- synthesis factor. If RPAwould associate with pre-incision complexes and would remain associated during the DNA synthesis step in NER, it would be bound much longer that observed experimentally. Evidently, numerous RPA dissociation/association events occur while repair is in progress, arguing against stepwise directional assembly of repair complexes. A direct consequence of strict sequential assembly and simultaneous release is that the amount of a repair factor that is recruited earlier in the chain is higher than a factor that is recruited later in the chain (Politi et al., 2005). In vivo measurements indicate that the level of bound XPG, XPA and RPA is considerably higher than the amount of XPC that is bound during repair. This behaviour is faithfully reproduced by our model, which is based on random assembly of repair factors. Moreover, we show that XPA exchanges with repair sites when XPC is not longer bound. This would not be possible in a strict sequential assembly mechanism in which XPAbinding is only possible after XPC binding. However, the binding of two proteins to repair complexes is sequential. Binding of TFIIH only occurs after binding of XPC to damaged DNA. Subsequently, TFIIH binds other repair proteins independent of XPC. Similarly, ERCC1-XPF only associates with complexes containing XPA. These sequential steps have been included in the model. These data strongly indicate that NER complex assembly is not strictly sequential.

Stochastic assembly of NER complexes and irreversible enzymatic reactions drive the repair process to completion Rather than sequential assembly, many aspects of the kinetics of NER can be faithfully reproduced if repair complex formation is assumed to be stochastic, in other words if repair factors assemble randomly on repair intermediates. Such an assembly mechanism results in the formation of many possible protein complexes on repair intermediates. For instance, 47 different assembly states are possible, each containing one or more of the following components: XPC, TFIIH, XPG, XPA, RPAand ERCC1- XPF on partially and fully unwound DNA (see Fig 4). Modelling reveals that repair factors only occasionally form a catalytically active complex capable of performing a specific enzymatic reaction, such as unwinding or incising the DNA. Progress made by assembly of the correct, i.e. functional, protein complex on a repair intermediate is fixed by irreversible enzymatic reactions. These irreversible reactions ensure that repair is directional and drive the whole repair process to completion. Thus, there is a directional formation of repair intermediates that are formed in a strict order, but the assembly of protein complexes on these different intermediates is stochastic. Stochastic assembly has been suggested previously for NER complexes (Kesseler et al., 2007; Reardon and Sancar, 2003) in agreement with our model. In previous models, based on in vitro measurements on CPD repair, RPA, XPA and XPC were suggested to bind randomly to DNA lesions. It is likely that such a mechanism is employed for the

118 Choreography of NER factors recognition of CPDs (Reardon and Sancar, 2003), which are poorly recognized by XPC and require UV-DDB for efficient removal (Tang et al., 2000). The in vivo measurements in the present study mainly reflect assembly of repair complexes on 6-4 PPs. We therefore do not consider CPD repair in our model. Recognition of 6-4 PPs during GGR in vivo is carried out strictly by XPC (Sugasawa et al., 1998; Volker et al., 2001). In agreement, both XPA and RPA do not bind to sites of repair in cells deficient for XPC, showing that their binding is strictly dependent on XPC (Rademakers et al., 2003; Volker et al., 2001). Clearly, RPA and XPA bind independently of XPC during transcription-coupled repair (TCR), but since the latter repair pathway contributes less than 1% to the total NER activity in cells this can only be detected using ChIP (Fousteri et al., 2006; Jiang and Sancar, 2006). Once XPC binds to a 6-4PP and recruits TFIIH, this results in partial unwinding of the DNA. In our model, all pre-incision factors then assembly at random on this partially unwound repair intermediate. The exception is ERCC1-XPF, which only binds complexes already containing XPA (Volker et al., 2001).

Assembly and functioning of multi-protein complexes is inherently slow and needs kinetic proofreading to proceed Essentially all processes that control the functioning of our genome are each carried out by tens of proteins that have to come together to exert their function. Examples are transcription initiation complexes, replication machines and DNA repair mechanisms. Mathematical modelling of NER suggests that protein complex assembly on repair intermediates is a time consuming process, which is an inherent property of multi- protein complexes. Our results reveal that about 99% of the repair time is spent on the assembly of catalytically active repair complexes while ~1% of the repair time is used to perform enzymatic reactions. For instance, roughly half of the DNA lesions are incised within 40 min after UV irradiation, while the unwinding (α and α’) and dual incision (β) reactions that take place to produce incised DNA take about 12 s (Table 3). Similarly, complex assembly on other repair intermediates is also highly time- consuming. Such slow complex assembly suggest a kinetic proofreading mechanism (Hopfield, 1974; Reardon and Sancar, 2004) for NER in vivo, which confers high specificity to the repair process even though this specificity is not present at the level of the individual repair factors. Specific and non-specific binding events often do not display too different binding kinetics and specificity of proteins is often determined by differences in koff (i.e. slower dissociation of specifically bound proteins), while kon is often the same for specific and non-specific association (Hopfield, 1974). Due to this non-specific binding, it is useful to split a complex process up in different steps, which are often separated by irreversible (enzymatic) reactions. Such a mechanism, known as kinetic proofreading, increases the specificity with which biochemical reactions take place above the level of available free energy difference between “correct” and “incorrect” substrates (Hopfield, 1974). NER naturally fits this concept since the

119 Chapter 6 affinity of lesion-recognition protein XPC is rather low, while the NER system as a whole removes lesions with high substrate specificity. The reduction of errors is achieved by the ability to “reject” false substrates and it allows a system to cope with the binding of non-specific proteins. Hence, the continuous building up and tearing down of repair complexes that only occasionally result in enzymatic activities can then be envisaged as quality control (Kanaar et al., 2008). Previous in vivo measurements on components of the RNA pol I and RNA pol II transcription machinery in mammalian cells revealed that also the onset of transcription is an unlikely event (Darzacq et al., 2007; Dundr et al., 2002). Mathematical modelling of mRNA and rRNA transcription suggested that about 1% of the RNA pol I and II binding events resulted in elongation and the formation of RNA. Consequently, more than 99% of the binding events on promoters are non-productive (Darzacq et al., 2007; Dundr et al., 2002). Together with our results, this indicates that complex assembly of the major chromatin-associated processes is highly time-consuming. We propose that this reflects quality control by kinetic proofreading, which confers high specificity to the entire process even though this degree of specificity is absent at the level of the individual components. The overall repair reaction is then kinetically driven by in practice irreversible enzymatic steps, which are essential to complete repair. Without such irreversible steps repair would be exceedeingly slow, since assembly of the large multi-protein complex that is able to carry out all enzymatic reactions during NER would have a low probability.

Experimental approaches to test model prediction Our combined approach of in vivo measurements at the single cell level and mathematical modelling provides detailed insight into the complex choreography of repair proteins in the nuclei of living cells. Critical predicted parameters identified by the model are, amongst others, the long time that is spent on complex assembly and the time-dependent changes in repair intermediate concentration. These parameters can be tested experimentally by lowering the rates of enzymatic reactions, perhaps by using specific drugs. The model predicts that lower rates for relatively fast enzymatic reactions, such as for example dual incision, should have little effect on the repair efficiency. The rationale behind this is that assembly of a complex capable to perform dual incision is much more time-consuming than the time it takes to perform dual incision itself. Model and experimental results imply that XPA has affinity for re- synthesized DNA. It may be that XPAhas an additional function after repair gap filling besides its function in lesion verification before dual incision. Consequently, its affinity to repair intermediates beyond repair synthesis is mediated by protein-protein interactions, rather than binding to DNA itself. Interestingly, the model predicts that RPAand PCNA bind to a repair intermediate for which the other NER factors examined in this study have no affinity. The concentration of this repair intermediate reaches a plateau about 4 hrs after UV exposure and levels of bound RPA and PCNA stay high

120 Choreography of NER factors for at least 8 hrs following UV irradiation. It is at present unclear what the precise nature of the repair intermediate is to which RPA and PCNA bind at these late time- points. Experiments triggered by the model are to perform ChIP experiments around 4-5 hours following UV irradiation using antibodies against RPA and PCNA to study the nature of the substrate to which these proteins bind. In addition, the model may be extended by studying chromatin-remodelling steps prior to UV as well as study in detail the kinetics of XRCC1-Lig IIII and DNA pol δ to gain further insight into DNA synthesis during NER as well as kinetics of CAF1 to unravel the kinetics of chromatin restoration following NER. Results of these model-driven experiments will help to obtain comprehensive understanding of a complex chromatin-associated process in full.

Materials and Methods

DNA constructs. The XPC cDNA was ligated in-frame with mVenus resulting in XPC-mVenus, which was transiently transfected in several NER mutant cell lines using lipofectamine 2000 (Invitrogen, Breda, the Ne- therlands) according to the manufacturer's instructions. RPA70 cDNA (Henricksen et al., 1994) was cloned in frame with EGFP in pEGFP-N1 (Clontech Laboratories, California, USA) and stably expressed in SV40 transformed MRC5 human fibroblasts.

Cell lines. Cell lines used in this study were human fibroblasts XPC deficient XP4PA-SV expressing XPC- EGFP (Politi et al., 2005), XPA deficient XP2OS-SV expressing EGFP-XPA (Rademakers et al., 2003), MRC5-SV expressing RPA70-EGFP (Gourdin and Vermeulen. Unpublished), Chinese hamster ovary cells: UV135 expressing XPG-EGFP (Zotter et al., 2006), 43-3B expressing ERCC1-GFP (Houtsmuller et al., 1999), CHO9 expressing EGFP-PCNA (Essers et al., 2005) and CHO K1. The expression level of all GFP- tagged repair proteins is comparable to the level of endogenous proteins as shown by western blot analysis (Essers et al., 2005; Hoogstraten et al., 2002; Hoogstraten et al., 2003; Houtsmuller et al., 1999; Rademakers et al., 2003; Zotter et al., 2006). Immunoblot analysis of MRC5 RPA70-EGFP cells with anti-RPA70 antibodies (B-6:sc-28304, mouse monoclonal 1:1000; Santa Cruz, California, USA) showed that endogenous RPA70 and RPA70-EGFP were expressed at similar levels. Hybridization with anti-GFP antibodies revealed that there was no breakdown product detectable, indicating that all the measured fluorescence in the cells is derived from the full-length protein (Gourdin and Vermeulen. Unpublished results). The following NER mutant cell lines were used: human XP-B (XPCS2BA-SV; Vermeulen et al., 1994), XP-A (XP12RO-SV; Satokata et al., 1992), XP-F (XP2YO-SV; Yagi et al., 1991) and CHO XP-B/ERCC3 (27.1; Hall et al., 2005), XP-G/ERCC5 (UV135 ;MacInnes et al., 1993) and ERCC1 (43-3B; van Duin et al., 1986). All cell lines expressing GFP-tagged NER proteins were cultured in a 1:1 mixture of DMEM/Ham’s F10 medium. All media contained glutamine (Gibco, Breda, The Netherlands) supplemented with antibiotics and 10% FCS and all cells were cultured at 37°C in an atmosphere of 5% CO2.

Microscopic analysis. Binding kinetics were measured on a Zeiss Axiovert 200M wide field fluorescence microscope, equipped with a 100x Plan-Apochromat (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Ger- many) and a Cairn Xenon Arc lamp with monochromator (Cairn research, Kent, U.K.). Images were recorded with a cooled CCD camera (Coolsnap HQ, Roper Scientific, USA). FLIP experiments were performed on Zeiss LSM 510 confocal microscope, equipped with a 63x Plan-A (1.4 NA) oil immersion lens (Zeiss, Ober- kochen, Germany) and a 60 mW Argon laser (488 and 514 nm). Both microscopes were equipped with an objective heater and cells were examined in microscopy medium (137 mM NaCl, 5.4 mM KCl, 1,8 mM CaCl2, 0.8 mM MgSO4, 20 mM D-glucose and 20 mM HEPES) at 37°C or 27°C.

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lerica, Massachusetts, USA) with pores of 5 µm and subsequently irradiated for 39 seconds (100 J.m-2) (Luijsterburg et al., 2007; Moné et al., 2004).

Binding kinetics. Cells were grown in glass bottom dishes (MatTek, Ashland, Massachusetts, USA) and locally UV-irradiated as described by van Driel and co-workers (Luijsterburg et al., 2007; Moné et al., 2004). Individual cells were subsequently monitored for up to 6 hours. Accumulation of GFP-tagged repair proteins after local irradiation was quantified with Object-Image software. Time courses were normalized with respect to the plateau level. Start of the UV irradiation was defined as t=0. The bound fraction of GFP-tagged NER proteins in the local damage was calculated by: Bound (%) = (Ispot-Ioutspot)* pixelsspot / (Inucleus-Ibackground)* pixels nucleus, where Ispot and Ioutspot are the average pixel intensities inside the damaged spot and outside the spot, respectively. Inucleus is the average pixel intensity of the nucleus, including the spot and Ibackground is the average pixel intensity outside the cell

Fluorescence Loss in Photobleaching (FLIP). FLIP analysis was performed by continuously bleaching a third of a locally UV-irradiated nucleus as previously described (Hoogstraten et al., 2002; Luijsterburg et al., 2007). Fluorescence in the locally damaged area was monitored with low laser intensity. All values were background corrected. To determine the exchange rate in the presence of repair synthesis inhibitors, cells were first locally irradiated. One hour following irradiation, cells were treated with 100 mM hydroxyurea (HU) and 10 µM cytosine-β-arabinofuranoside (AraC) for 30 minutes. FLIP was subsequently performed in the presence of HU and AraC.

Acknowledgement This work was supported by the Netherlands Organisation for Health Research and Development (ZonMW, grant 912-03-012).

122 Chapter 7 Heterochromatin Protein 1 is Involved in the DNA Damage Response

Martijn S. Luijsterburg, Christoffel Dinant, Elżbieta Wiernasz, Hannes Lans, Jan Stap, Maria I. Fousteri, Daniël O. Warmerdam, Maartje C. Brink, Saskia Lagerwerf, Jurek W. Dobrucki, Gert Jansen, Wim Vermeulen, Leon H.F. Mullenders, Adriaan B. Houtsmuller, Pernette J. Verschure, and Roel van Driel

Submitted

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Abstract The mammalian genome is protected against genotoxic stress by DNA damage response (DDR) mechanisms that include DNA repair. Heterochromatin protein 1 (HP1) family members are chromatin-associated proteins involved in transcription and replication. In this study we show that the HP1 isoforms HP1α, HP1β and HP1γ are recruited to DNA lesions in living human cells. This response to DNA damage strictly requires the chromo shadow domain of the HP1 protein and occurs at sites of double strand breaks and at sites of helix-distorting lesions. Evidence is provided that HP1 is not involved DNA repair itself. We show that several HP1-interacting proteins are also recruited to DNA damage, including chromatin remodelling factors and DNA and histone methyltransferases. Loss of HP1 results in high sensitivity to UV-induced damage in the nematode C. elegans, providing evidence for involvement of HP1 in a DDR mechanism. These results suggest a novel function of HP1 proteins and reveal a link between maintenance of epigenetic information and DNA damage responses.

Introduction The genetic component of chromatin (i.e. the DNA) can be damaged by various sources including ionizing and ultraviolet (UV) radiation. Cells respond to genotoxic stress by activating interwoven DNA damage response (DDR) pathways including DNA repair, cell cycle arrest, repression of transcription, senescence and apoptosis (Bartek and Lukas, 2007; Huen and Chen, 2008; Shimada et al., 2008). To restore the genetic information after it has been compromised, several sophisticated DNA repair pathways have evolved, each dealing with specific lesions (Hoeijmakers, 2001). Nucleotide excision repair (NER) removes bulky adducts and UV-induced pyrimidine-pyrimidone (6-4) photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs) from the genome (de Laat et al., 1999). NER involves lesion detection by stalled RNA Polymerase II (for transcription-coupled NER or TC-NER) or XPC and DDB2 complexes (for global genome NER or GG-NER) and subsequent unwinding of DNA (TFIIH, RPA, XPA), incision of the damaged strand (XPG and ERCC1-XPF) and DNA resynthesis, which is performed by the DNA replication machinery (de Laat et al., 1999; Fousteri and Mullenders, 2008; Friedberg, 2001; Hoeijmakers, 2001). DNA double strand breaks (DSB) induced by, amongst others, ionizing radiation, are removed by homologous recombination (HR) or non-homologous end-joining (NHEJ). Repair by HR requires a homologous template, which is available during S and G2 phases. Therefore HR is active during these stages of the cell-cycle, whereas NHEJ repairs DSBs mainly in G1(Kanaar et al., 2008).

Chromatin is the substrate for all genome-associated processes in eukaryotes such as replication, transcription and DNA repair. Besides the genetic information, chroma tin encodes inherited epigenetic information by means of DNA methylation, histone

124 HP1 is involved in the DNA damage response variants and post-translational modifications of histones (Henikoff et al., 2004; Jenuwein and Allis, 2001; Turner, 2002). The maintenance of epigenetic information is crucial to regulate gene expression profiles and maintain cellular identity. How epigenetic information is maintained during DNA repair is currently not understood. A major obstacle for NER in eukaryotic cells is the packaging of damaged DNA into chromatin. In general, chromatin is considered inhibitory to processes associated with DNA and the rate of repair in a chromatin context is decreased to about 10% of that on naked DNA (Hara et al., 2000; Ura et al., 2001; Wang et al., 1991). To overcome this obstacle, cells employ a number of chromatin modifying activities to create a more permissive environment for repair locally. It has been demonstrated that NER in mono- and di-nucleosomes is considerably enhanced by ATP-dependent remodelling factors like SWI/SNF and ACF1 (containing ISWI) (Hara and Sancar, 2002; Ura et al., 2001), although it is unclear whether these factors also stimulate NER in vivo. Several other mechanisms, including histone ubiquitylation (Kapetanaki et al., 2006; Wang et al., 2006a), histone acetylation (Brand et al., 2001; Fousteri et al., 2006; Martinez et al., 2001) and small acidic proteins (e.g. GADD45 and ING) that increase the accessibility of nucleosomal DNA, have been suggested to enhance NER in a chromatin environment (Carrier et al., 1999; Cheung et al., 2001; Kuo et al., 2007; Smith et al., 2000; Smith et al., 1996; Wang et al., 2006b). Following removal of DNA injuries, cells need to restore the original chromatin structure to maintain the epigenetic information, which requires specialized factors (Groth et al., 2007). For instance, CAF1 is recruited to NER sites followed by incorporation of histone H3.1, reflecting a chromatin reassembly step after repair (Green and Almouzni, 2003; Polo et al., 2006). Such a model in which chromatin rearrangements precede and follow DNA repair, has been put forward as the access-repair-restore model that has proven very useful for understanding how NER operates on a chromatin template (Green and Almouzni, 2002; Smerdon, 1991). This model is conceptually appealing and predicts that epigenetic regulators are involved in chromatin rearrangements before, during or after DNA repair.

The heterochromatin protein 1 (HP1) isoforms, HP1α, HP1β and HP1γ are versatile epigenetic regulators with functions in chromatin organization, transcription regulation and DNA replication (Lomberk et al., 2006; Maison and Almouzni, 2004). All three HP1 isoforms share a common dimerization motif allowing them to form homo and heterodimers (Cowieson et al., 2000). The versatility of HP1 proteins in various chromatin-interacting processes raises the question whether these proteins are involved in DNA damage response mechanisms. HP1 proteins, amongst others localize to pericentromeric heterochromatin (Mateos-Langerak et al., 2007; Schmiedeberg et al., 2004). Expression of genes is considerably decreased when HP1 is targeted to them (Ayyanathan et al., 2003; van der Vlag et al., 2000). We previously demonstrated that targeted binding of HP1 triggers formation of heterochromatin (Verschure et al., 2005).

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However, depletion of chromatin-bound HP1 did not result in loss of heterochromatin, suggesting that HP1 is not involved in heterochromatin maintenance (Mateos-Langerak et al., 2007). Although previous studies have led to the idea that HP1 proteins are mainly involved in heterochromatin formation it is becoming increasingly clear that this view needs revision. For instance, analysis of HP1 binding sites along the Drosophila genome revealed that HP1 also associates with actively transcribed genes (de Wit et al., 2007). Moreover, the HP1γ isoform interacts with RNAPII and is mainly found in euchromatin (Minc et al., 2000; Vakoc et al., 2005). HP1 proteins directly bind to methylated H3 at Lys9 via their N-terminal chromodomain (Jacobs and Khorasanizadeh, 2002). In addition, HP1 dimers interact with a plethora of nuclear proteins via their C-terminal chromoshadow domains, including Suv(3-9), Suv(4-20), Dnmt1, CAF1 as well as chromatin remodelling proteins ACF1 and BRG1 (Eskeland et al., 2007; Fuks et al., 2003; Murzina et al., 1999; Nielsen et al., 2002; Thiru et al., 2004). This makes HP1 a candidate to serve as a binding platform for the recruitment of epigenetic regulators in response to DNA damage.

In this study, we present evidence for a novel function of the HP1 proteins in the response to DNA damage. We demonstrate that the HP1 proteins are recruited to several types of DNA injuries such as UV-lesions and double strand breaks (DSBs) in living human cells. A comprehensive in vivo analysis shows that recruitment of HP1 to UV-induced lesions critically depends on the CSD and is independent of the binding and activity of pre-incision NER proteins. Moreover, we demonstrate that loss of HP1 renders the nematode C. elegans highly sensitive to UV irradiation, providing evidence for involvement of HP1 in the DNA damage response.

Results

HP1 proteins are recruited to UV-lesions by the chromoshadow domain To study if HP1 proteins are recruited to sites of UV-induced DNA damage, we used UV-C light that induces the formation of CPDs and 6-4 PPs. Cell nuclei were locally irradiated with either a UV-C laser (266 nm; Dinant et al., 2007) or with a UV-C lamp (254 nm) through a polycarbonate mask with pores of 5 µm, resulting in localized DNA damage (Luijsterburg et al., 2007; Moné et al., 2004). Both methods trigger recruitment of NER proteins but not of factors involved in other repair pathways, demonstrating that these methods produce bona fide NER-specific lesions. Human HeLa cells and mouse NIH/3T3 were transfected with fluorescent protein-tagged HP1 and mCherry-tagged DDB2 and subsequently irradiated. At irradiated sites that are marked by the accumulation of fluorescent protein-tagged DDB2, we observed recruitment of all three HP1 isoforms (mRFP-HP1α, SCFP3a-HP1β and EGFP-HP1γ) in both cell types (Figure 1A-C). In agreement, endogenous HP1β also accumulated after local UV irradiation in primary human fibroblasts (data not shown). To confirm

126 HP1 is involved in the DNA damage response these results, we carried out photobleaching experiments on mouse cells stably expressing EGFP-HP1β that were globally UV-C irradiated at 25 J.m-2. Bleaching one half of a cell nucleus (n = 5 cells) and monitoring the equilibration of bleached and non-bleached molecules confirmed that a small fraction of EGFP-HP1β was immobilized in UV-irradiated but not in control cells on a time-scale of several minutes (Figure 1D,E).

HP1 proteins contain three distinct domains: the N-terminal chromo-domain (CD), the C-terminal chromoshadow-domain (CSD) and the hinge region that separates the CD from the CSD. This prompted us to investigate whether a specific HP1 domain is responsible for the recruitment to DNA damage. Different deletion mutants of HP1β, lacking CD, CDS or hinge (Figure 2A), were tagged with EGFP or mCherry and tested for recruitment to UV-irradiated regions. Interestingly, recruitment of mCherry-HP1β (∆CD) and EGFP-HP1β (∆hinge) following UV irradiation was observed, but EGFP- HP1β (∆CSD) failed to accumulate (Figure 2B-D). To test whether the CSD (amino acids 98 – 185 of HP1β) is sufficient for recruitment, we fused this domain to EYFP and show that CSD-EYFP is indeed recruited to sites of localized UV irradiation (Figure 2E). To verify that the CD is not required for HP1 binding, we examined recruitment of HP1 in MEFs deficient for Suv39h1 and Suv39h2 (Peters et al., 2001; Schotta et al., 2004). These proteins are methyl-transferases responsible for tri-methy lation of H3K9, which is a binding site for HP1 via its CD (Cheutin et al., 2003). Following UV irradiation, SCFP3a-HP1β clearly accumulated at sites of damage, confirming that binding of HP1 does not require the CD (data not shown). These results demonstrate that all three HP1 isoforms are recruited to sites of UV-induced DNA damage and that this recruitment depends on the CSD.

HP1 recruitment to UV lesions is independent of NER Cells from placental mammals are fully dependent on NER for the removal of UV- induced DNA injuries. Several chromatin-related events are triggered by UV-lesions, such as recruitment of CAF-1 followed by incorporation of histone H3.1 and ubiquitylation of H2A at lysine 119. These events strictly depend on the binding and activity of early binding NER proteins, involved in recognition and stabilising of lesions (Bergink et al., 2006; Green and Almouzni, 2003; Polo et al., 2006). This favours a scenario in which such chromatin rearrangements occur only after repair by NER. To investigate whether HP1 recruitment is a late event following repair we tested accumulation of HP1β in repair-deficient XP-A cells that have compromised GG- and TC-NER. Unexpectedly, SCFP3a-HP1β accumulated in two XPA mutant cell lines upon UV irradiation, showing that its recruitment does not require dual incision (Figure 3a). This shows that HP1β binding is not a late step that occurs after DNA repair is finished. We then considered the possibility that HP1 binding is an early step following damage detection and tested accumulation in DDB2-deficient and XPC-deficient

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Figure 1. Recruitment of HP1s to UV- damage. (A) Nuclear localization ofmRFP- HP1α, (B) SCFP3a- HP1β and (C) EGFP-HP1γ in living HeLa cells locally irradiated at 100 J.m-2 though 5 µm pores or irradiated using a UV- C laser. The site of local DNA damage is indicated by accumulation of DDB2-mVenus or DDB2-mCherry (left column in A,B and C). (D) immunolocalisation of endogenous HP1β in locally UV-irradiated conflu- ent normal human fibroblasts through 3 µm pores. UV- damaged sites are visualized by local accumulation of XPA. Combined FLIP-FRAP analysis on NIH-3T3 cells expressing EGFP-HP1β. Cells were either mock treated (E) or globally irradiated at 25 J.m-2 (F). Half of a cell nucleus was bleached and the loss of fluoresence (FLIP) was measured in the non-bleached half (blue line) while the recovery of fluoresence (FRAP) was measured in the bleached half (red line).

Figure 2. HP1β deletion mutants. (A) Re- presentation of DNA constructs encoding fluorescently tagged HP1β deletion mutants. The CD is indicated in red, the CSD in blue and the hinge in yellow. Indicated numbers represent amino acids. (B) Nuclear localization of mCherry-HP1β (∆CD), (C) EGFP- HP1β (∆hinge), (D) EGFP-HP1β (∆CSD) or (E) EYFP-CSD in living HeLa cells locally irradiated at 100 J.m-2 through 5 µm pores or irradiated using a UV-C laser. The site of local DNA damage is indicated by accumu lation of DDB2-mVe- nus or DDB2-mCherry

128 HP1 is involved in the DNA damage response human cells, as well as XPC-inactivated mouse embryonic fibroblasts (MEF), which express very low levels of DDB2. Accumulation of SCFP3a-HP1β was observed in all cell lines, showing that HP1 binding is independent of the activity of GG-NER proteins (Figure 3B-D). Damage detection during TC-NER requires stalled RNAPII and subsequent recruitment of NER factors depends on the coupling-factor CSB (Fousteri et al., 2006). However, recruitment of HP1 was also observed in CSB-deficient cells derived from CS-B patients, indicating that it did not require TC-NER (Figure 3E).

One explanation for recruitment of HP1 to DNA damage in NER deficient cells is that the cells that we monitored were in S-phase. Lesions induced by UV light can be converted to other types of lesions (e.g. DSBs) during replication, which are removed by other repair pathways than NER. To exclude this possibility, we determined the cell-cycle stage by expressing mCherry-PCNA together with SCFP3a-HP1β and DDB2-mVenus in repair-deficient XP-A cells and wild-type MRC5 cells. Recruitment of HP1β was observed in wild-type and NER-deficient in S-phase as well as non S- phase cells, as shown by the distribution of PCNA (Supplemental figure S1A and -B). To verify these results, we examined the distribution of endogenous HP1β in human cells that do not proliferate (i.e. G0 cells), using Ki67 antigen, as a marker for cell proliferation. Clear accumulation of endogenous HP1β was observed in G0 cells (Ki67 negative cells) at sites of UV irradiation (Supplemental figure S1C), showing that HP1 accumulation is not due to stalled replication. Together, these results show that HP1 binding following UV irradiation occurs in both cycling and quiescent cells and is independent of the activity of pre-incision NER proteins and CSB.

We then investigated whether HP1 accumulates longer on damaged DNA in cells in which UV lesions are not repaired. To test this, accumulation of SCFP3a-HP1β following local UV damage was measured in XPA-deficient cells (XP12RO) for several hours. Accumulation of the fusion protein was still observed ~4 hrs after irradiation (Figure 4A). Conversely, in XPA-deficient cells that were transiently transfected with mVenus-XPA (to restore the repair capacity), bound HP1β levels gradually decreased in time and HP1 accumulation was almost lost 4 hrs following UV irradiation (Figure 4B). Accordingly, accumulation of EYFP-tagged CSD of HP1β in HeLa cells was also lost ~4 hrs after local irradiation (data not shown). These results show that binding of HP1 is mainly triggered by the presence of 6-4PPs because this type of lesion is removed within ~4 hours following repair whereas CPDs are still abundant at this time. To our knowledge, this is the first example of a protein that is recruited to sites of UV-induced DNA damage independent of the known NER factors.

HP1 recruitment to UV lesions is independent of ATR In addition to DNA repair, cells respond to damaged DNA by activating ATR/ATM

129 Chapter 7 kinase signaling pathways resulting in activation of cell cycle checkpoint (e.g. Chk1 and Chk2) and phosphorylation of a plethora of proteins involved in the DDR (Bartek and Lukas, 2007). UV-induced DNA damage activates ATR, which is required for ubiquitylation of H2A following UV damage (Bergink et al., 2006). It is currently unclear if ATR activation requires processing of DNA lesion by NER or whether ATR binds directly to damaged DNA (Choi et al., 2007; O'Driscoll et al., 2003). Since HP1 recruitment to sites of DNA damage does not require NER, we examined accumulation of HP1α, β and γ in Seckel cells, which have severely reduced ATR expression (O'Driscoll et al., 2003). Accumulation of the HP1 isoforms was not affected in ATR mutant cells following local UV irradiation (Figure 3F). This demonstrates that HP1 recruitment to sites of DNA damage does not require DNA damage-induced signalling mediated by the ATR kinase.

HP1 is recruited to double-stranded breaks To investigate whether the recruitment of HP1 is restricted to UV-induced DNA damage or if it is a more general response to different types of lesions, we tested HP1 recruitment upon producing double strand DNA breaks (DSBs). Cells were irradiated with α-particles from a radioactive Americium (Am-241) source thereby producing linear tracks of DSBs in cell nuclei (Aten et al., 2004; Stap et al., 2008; Williams et al., 2007). Human U2OS and mouse NIH/3T3 cells were irradiated with α-particles and clear linear tracks of γH2A.X were observed by immunolabelling. Accumulation of EGFP-HP1β in mouse cells and endogenous HP1β in human U2OS cells co-localized with the linear γH2AX pattern. (Figure 5A,B). In addition, accumulation of GFP-HP1α and GFP-HP1γ was observed in MRC5 cells upon α-particle irradiation, showing that all HP1 isoforms are recruited to DSBs (Figure 5; data not shown). Strikingly, complete co-localization of HP1β and γH2AX was observed near DSBs (Figure 5), which is different from the observed localization of many DSB repair proteins (e.g. Rad51 or DNA-PK) that are usually found only in a small area within larger γH2AX domains (Aten et al., 2004; Bekker-Jensen et al., 2006). This suggests that HP1 associates with a chromatin area larger than the damaged DNA region. Mammalian cells utilize homologous recombination (HR; which is only operational in S and G2 ) or non- homologous end-joining (NHEJ) to remove DSBs from the genome (Wyman and Kanaar, 2006). The latter pathway is initiation by the KU70/80 dimer (Mari et al., 2006), which was shown to interact with HP1α (Song et al., 2001). To test whether HP1 accumulation at DSBs depends on NHEJ or KU70 in particular, we irradiated wild-type and KU80-deficient CHO cells with α-particles. Clear recruitment of endogenous HP1β was observed in both cell types at all γH2A.X tracks, showing that HP1 association is independent of NHEJ (Figure 5C). Co-localization of HP1 with γH2A.X tracks was detected in all cells, indicating that HP1 binding is independent of the cell cycle stage. This argues against recruitment by homologous recombination (HR). Thus, similar to the NER-independent recruitment to UV-lesions, we provide evidence that

130 HP1 is involved in the DNA damage response

HP1 associates with DSBs independent of NHEJ and possibly independent of HR.

Figure 4. Long-term accumulation of HP1β in repair-proficient and repair-deficient cells. (A) XP-A cells were transected with mVenus-XPA (to complement the repair-deficient phenotype), DDB2- mCherry and SCFP3a-HP1β. Cells were irradiated at 100 J.m-2 and accumulation of HP1β was monitored for 4 hrs follwing UV irradiation. (B) XP-A cells were transfected with DDB2-mVenus and SCFP3a-HP1β. Cells were irradiated at 100 J.m-2 and accumulation of HP1β was monitored for 4 hrs follwing UV irradiation. The accumulation of DDB2-mVenus or DDB2-mCherry indicates the site of local damage. Accumulation of mVenus-XPA is observed at the site of DNA damage and in nucleoli.

Figure 3. Recruitment of HP1β in NER-deficient cells. (A) Nuclear localization of SCFP3a-HP1β or EGFP- HP1β in human fibroblasts deficient for XPA, (B) XPC, (C) DDB2, (D) MEFs deficient for XPC, (E) human fibroblasts deficient for CSB and (F) human seckel cells, which have severly reduced expression of ATR kinase. All cells were locally irradiated at 100 J.m-2 though 5 µm pores or irradiated using a UV-C laser. The site of local DNA damage is indicated by accumulation of DDB2-mVenus, DDB2-mCherry or XPC-mVenus or by a square (in D and F).

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Figure 5. Recruitment of HP1β to DSBs. (A) Wild- type U2OS cells were irradiated with α-particles and subsequently labeled for endogenous HP1β (green) and γH2A.X (red). (B) Mouse cells expressing EGFP-HP1b (green) were irradiated with α-particles and subsequently labeled for γH2A.X (red). (C) Ham- ster cells deficient in Ku80 were irradiated with α-particles and subsequently labeled for endogenous HP1β (green) and γH2A.X (red). A merged image of HP1 and γH2A.X is shown.

Figure 6. Recruitment of HP1-binding proteins to UV-damage. (A) Accumulation of Dnmt1- EGFP and (B) EYFP-BRG1 in living HeLa cells or MRC5- Sv cells locally irradiated at 100 J.m-2 though 5 µm pores or irradiated using a UV-C laser. (C) Accumulation of Dnmt1-EGFP, and (D) EYFP-BRG1 in living XP-A or XP- C cells locally irradiated at 100 J.m-2 though 5 µm pores or irradiated using a UV-C laser. (E) Accumulation of SCFP3a-HP1β in Dnmt1 knock-out cells and (F) Accumu- lation of EGFP-Suv4-20h1 in locally irradiated MRC5-SV cells.

Several HP1-interacting proteins are recruited to UV-lesions Having established that HP1 proteins are recruited to UV-induced DNA damage through the CSD, we next investigated whether proteins that interact with this domain also accumulate at sites of DNA damage. DNA methyltransferase 1 (Dnmt1) directly interacts with the CSD of HP1 and has previously been shown to accumulate at damaged DNA sites (Mortusewicz et al., 2005). Indeed, EGFP-Dnmt1 accumulated upon UV irradiation (Figure 6A). If Dnmt1 methylates DNA at the locally irradiated site, this may

132 HP1 is involved in the DNA damage response trigger the binding of MeCP2, which is a protein that binds methylated DNA (Ho et al., 2008). Also, EGFP-tagged MeCP2 did not accumulate after localized UV irradiation (data not shown). We also tested accumulation of EYFP-tagged H3K9 methyl transferase Suv39H1 (Krouwels et al., 2005) at sites of UV-lesions. Local UV irradiation did not result in recruitment of EYFP-Suv39H1 (data not shown). This demonstrates that H3K9Me3 is not involved in HP1 recruitment to DNA damage, consistent with the findings that HP1 is recruited in cells deficient for Suv39h and that the CD is not required for HP1 binding (data not shown, Figure 2). Interestingly, the H4K20 methyl-tranferase EGFP-Suv4-20h1 (Schotta et al., 2004; Yang et al., 2008) did accumulate following UV irradiation (Figure 6F). Previous studies showed that H4K20me3 depends on Suv3-9 and direct interactions between HP1 and Suv4-20 (Schotta et al., 2004). It appears that after UV damage HP1 recruits Suv4-20 independently of Suv3-9 at damaged sites.

Previous in vitro experiments have suggested a role for ATP-dependent chromatin remodelling factors like SWI/SNF and ACF1 during DNA repair (Hara and Sancar, 2002; Ura et al., 2001). Interestingly, ACF1 and the SNF2-like BRG1 protein directly interact with the CSD of HP1 and we tested whether these factors are recruited to sites of damage in vivo (Eskeland et al., 2007; Nielsen et al., 2002). Upon UV irradiation of HeLa cells, we observed accumulation of EYFP-BRG1 and ACF1-EGFP, which can form a heterodimer with ISWI, showing that ATP-dependent remodelling factors are recruited to sites of UV-induced DNA damage (Figure 6B, data not shown). Similar to HP1, accumulation of EGFP-Dnmt1, EYFP-BRG1 and ACF1-EGFP was also detected in NER-deficient cells (Figure 6C, D, data not shown), suggesting that HP1, Dnmt1, ACF1 and BRG1 may be involved in a DNA damage response pathway other than DNA repair. To investigate if the accumulation of HP1 depends on Dnmt1, we expressed SCFP3a-HP1β in Dnmt1 knock-out cells (Rhee et al., 2000). Results show that HP1β accumulates upon UV irradiation in the absence of Dnmt1 (Figure 6E), suggesting that HP1 is not recruited to DNA lesions by this protein.

Loss of HP1 renders C.elegans highly sensitive to UV irradiation Because knockout of HP1 in mammalian cells is lethal (Filesi et al., 2002; Schotta etal., 2004), we used the nematode C. elegans to test whether HP1 is functionally required for the DNA damage response. Two HP1 homologues (HPL-1 and HPL-2) are present in C. elegans and loss of both HP1 proteins is also lethal in worms (Coustham et al., 2006; Schott et al., 2006). To circumvent this problem, we employed animals lacking HPL-1 and carrying a temperature-senstive allele of HPL-2, which is expressed at 20°C but not at 25°C (Coustham et al., 2006). To test whether loss of HPL-1 and HPL-2 results in sensitivity to UV, we exposed eggs of single and double mutant animals (grown at 25°C) carrying null alleles of hpl-1 and hpl-2 to UV-B radiation (80 J.m-2). Wild-type and NER-deficient xpa-1 null eggs (Stergiou et al., 2007) were also

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Figure 7. Survival of C. elegans HP1 knock-out worms upon UV irradiation. (A) Hatching of wild-type, hpl- 1, hpl-2 and hpl-2/hpl-1 mutant eggs 8 hours following collection. (B) Hatching of wild-type, hpl-1, hpl-2 and hpl-2/hpl-1 mutant eggs 8 hours following collection and subsequetly irradiated with UV-B at 80 J.m-2. (C) Quantification of hatching and non-hatching eggs following UV-irradiation releative to non- irradiated eggs. In addition to wild-type (red bars), hpl-1 (light-yellow bars), hpl-2 (dark-yellow bars) and hpl-2/hpl-1 (blue bars) mutant eggs, the survival of xpa mutant eggs was also quantified (purple bars).

Supplemental figure S1. HP1β accumulation is independent of cell cycle. Representative images of human XP-A cells expressing mCherry-PCNA (red), SCFP3a-HP1β (cyan) and DDB2-mVenus (yellow) following local irradiation at 100 J.m-2 through 5 µm pores. (A) Accumulation of HP1β is observed outside S- phase when PCNA is homogenously distributed in the cell nucleus and (B) in S-phase cells (indicated by the typical S-phase pattern of PCNA. (C) Immunolocalisation of endogenous HP1β in locally UV-irradiated (3 µm pores, 100 J.m-2) in quiescent human fibroblasts as visualised by Ki67 negative staining and in cycling cells as visualised by Ki67 positive staining.

134 HP1 is involved in the DNA damage response irradiated at 80 J.m-2. The survival of irradiated eggs was subsequently determined compared to non-irradiated eggs (Figure 7). Wild-type and hpl-1 or hpl-2 single mutant worms did not exhibit increased sensitivity to UV-B. Strikingly, UV-B irradiation caused an immediate growth arrest in hpl-2/hpl-1 double mutant worms comparable to NER-deficient xpa-1 mutant worms. Similar results were obtained when juvenile hpl- 2/hpl-1 worms were irradiated instead of eggs (data not shown). This indicates that either HPL-1 or HPL-2 is sufficient for a normal response to DNA damage, but that loss of both HP1 proteins renders C. elegans highly sensitive to UV irradiation. These results reveal an essential role for the HP1 proteins in the DNA damage response in C. elegans. The association of HP1 proteins with several types of DNA lesions in mammalian cells suggests that this DNA damage response mechanism in conserved between C. elegans and humans.

Discussion In this study, we provide evidence for a novel function of the HP1 proteins in the DNA damage response. We show that all three isoforms of human HP1 (α, β and γ) are recruited to sites of UV-induced DNA damage (Figure 1) and to areas of DSBs (Figure 5; data not shown). Moreover, HP1 is also recruited to DNA lesions repaired by base excision repair (Zarebski and Dobrucki, unpublished results). This indicates that HP1 is involved in a variety of DNA repair systems in mammalian cells. Our finding that C. elegans without HP1 is highly UV sensitive, suggests that HP1 is an essential component of the DNA damage response. Unexpectedly, recruitment of HP1 to DNA damage is independent of any of the known damage recognition proteins, such as XPC and DDB2 in NER and KU70/80 in DSB repair. Moreover, HP1 binding is independent of DNA-damage induced signalling by ATR kinase (Figure 3). We show that HP1 binding does not require active NER. However, loss of DNA damage-induced HP1 binding sites occurs only in cells that are proficient in NER. HP1 remains bound under conditions that the NER system is inactive (Fig. 4). This shows that the NER system and the HP1 system communicate with each other. In conjunction, these results indicate that the HP1 proteins are involved in NER via a novel mechanism that acts in parallel to the well-studied NER repair pathway. HP1 is likely involved in a process that modifies and/or re-establishes the correct chromatin structure at the sites of DNA damage and that serves various repair systems.

Recently, Ayoub et al. (Ayoub et al., 2008) presented evidence that suggested that HP1β is mobilized from DNA breaks, seemingly in contrast with results presented here. This may be due to the fact these authors use an exceedingly harsh method to inflict DNA damage in living cells: 200 iterations of 100% power from a 405 nm diode laser under conditions that the DNA is photo-sensitized with Hoechst and BrdU. Very likely, under these conditions severe and diverse types of photodamage are induced in the DNA (Dinant et al., 2007; Williams et al., 2007). Therefore, the data of Ayoub et al. (Ayoub

135 Chapter 7 et al., 2008) cannot easily be compared to our conditions, which have been chosen so that either the NER or the DSB repair system is activated.

Recruitment of HP1 to UV-lesions is strictly dependent on the C-terminal chromo shadow domain (CSD) and does not require the N-terminal chromodomain (CD) or hinge region (Figure 2). Therefore is unlikely that interaction with methylated H3K9 plays a major role. The CSD of HP1 is known to interact with many nuclear proteins, including chromatin remodelling factors ACF1 and BRG1, maintenance DNA methyltransferase Dnmt1 and histone chaperone CAF-1 (Eskeland et al., 2007; Fuks et al., 2003; Murzina et al., 1999; Nielsen et al., 2002). We have shown that, similar to HP1 recruitment, Dnmt1 is recruited to UV-damaged areas in the nucleus independently of the activity of pre-incision NER proteins (Figure 6). However, recruitment of HP1 did not depend on Dnmt1, showing that HP1 does not bind to DNA lesions via this protein. Alternatively, HP1 may serve as a loading platform for various epigenetic regulators that are involved in DNA damage response mechanisms and are uncoupled from NER. In this scenario, HP1 would recruit proteins such as Dnmt1, Suv4-20, BRG1 and ACF1 to damage. Previous studies have demonstrated that, in contrast to HP1, recruitment of CAF-1 depends on dual incision (Green and Almouzni, 2003) and it is not clear if HP1 and CAF-1 interact at sites of DNA damage.

What is the molecular mechanism and the function of HP1 in DNA damage response? Evidently, binding of HP1 at sites of DNA damage after UV damage does neither require any of the known damage response proteins (XPC, DDB2 or Ku80), nor DNA repair activity. Consistent with these results, HP1 is not required for NER on naked DNA in vitro (Aboussekhra et al., 1995). Binding of HP1 fully depends on its chromoshadow domain, excluding a role of H3K9 methylation. Accordingly, Suv39H1 does not accumulate on damaged DNA and HP1β still accumulates in cells deficient for Suv39H1. Moreover, HP1 binding does not depend on the presence of the DNA methyltransferase Dnmt1, which interacts with the chromoshadow domain. In contrast, the H4K20 methyl-tranferase Suv4-20h1 did accumulate, possibly cooperating with HP1 in modifying chromatin structure. Our studies unveil an unexpected, intriguing and apparently ubiquitous link between DNA repair systems and chromatin modifiers. It is tempting to speculate that HP1 is involved in re-establishing the correct chromatin state after DNA repair. HP1β was recently shown to associate with inactive RNA pol II resulting in gene repression, while HP1γ binds to elongating RNA pol II, linking the HP1 proteins directly to regulation of transcription (Mateescu et al., 2008; Smallwood et al., 2008). It is attractive to speculate that HP1 proteins play a role in regulation of transcription in chromatin containing UV lesions or DNA breaks, since DNA damage inhibits transcription independently of DNA repair (Moné et al., 2001; Solovjeva et al., 2007). This process involves a yet unknown damage detection system in addition to the known detection pathways.

136 HP1 is involved in the DNA damage response

Materials and Methods

Cell lines. Cell lines used in this study were HeLa, U2OS, CHOK1, NIH/3T3, NIH/3T3 EGFP-HP1β (Ma- teos-Langerak et al., 2007), VH10 hTERT immortalised normal human fibroblasts, ATR-deficient GM18366- hTERT Seckel cells (Bergink et al., 2006), Dnmt1-deficient HCT116 cells (Rhee et al., 2000), Suv3-9h double knockout MEFs (Peters et al., 2001; Schotta et al., 2004) and KU80-deficient XR-V15B CHO cells (Mari et al., 2006). The NER-deficient SV40-immortalized cell lines were XP4PA (XP-C), XP20S (XP-A), XP12RO (XP-A), XP23PV (XP-E), MEFs XPC-/- and CS1AN (CS-B). All cell lines were cultured in a 1:1 mixture of DMEM/Ham’s F10 medium. All media contained glutamine (Gibco, Breda, the Netherlands) supplemented with antibiotics and 10% FCS and all cells were cultured at 37°C in an atmosphere of 5% CO2. For immunolocalisation experiments of endogenous HP1β hTERT immortalised human fibroblasts were grown to confluency for approximately 10 days. Subsequently, cells were synchronised in G0 phase by keeping them for a minimum of 5 days in medium supplemented with 0.2% FCS (serum starved cells).

DNA constructs. HP1α and HP1β cDNA were ligated in frame with mRFP and super cyan fluorescent protein 3a (SCFP3a) resulting in mRFP-HP1α and SCFP3a-HP1β, respectively. EGFP-HP1β and EGFP-HP1γ were a gift from Dr. P. Hemmerich (Schmiedeberg et al., 2004). All constructs were transiently transfected in various cell lines cells using Lipofectamine 2000. EGFP-HP1β was stably expressed in mouse NIH-3T3 cells as described previously (Mateos-Langerak et al., 2007). HP1β (∆CD) was tagged with mCherry as described elsewhere (Mateos-Langerak et al., 2007) and EGFP- HP1β (∆CSD) and EGFP-HP1β (∆hinge) were kindly provided by Dr. T. Misteli (Cheutin et al., 2003). EYFP-tagged CSD was provided by Dr. Y. Hi- rako (Hayakawa et al., 2003). The genes encoding NER proteins XPC, DDB2 and XPA were fused to the yellow fluorescent protein variant, monomeric Venus (mVenus), resulting in XPC-mVenus, DDB2-mVenus and mVenus-XPA. In addition, DDB2 was fused to mCherry (Luijsterburg et al., 2007). MeCP2 cDNA (kind gift from M.C. Cardoso) was fused to EGFP and EGFP-Dnmt1 was provided by Dr. H. Leonardt (Mortuse- wicz et al., 2005). EYFP-BRG1 was provided by Dr. T. Misteli (Phair et al., 2004) and ACF1-EGFP by Dr. P.D. Varga-Weisz (Collins et al., 2002). EYFP-Suv3-9H1 was provided by Dr. R.W. Dirks and EGFP-Suv4- 20H1 and H2 by Dr. T. Jenuwein. The cDNAs for SCFP3a and mVenus were provided by Dr. J. Goedhart (Kremers et al., 2006)and mCherry and mRFP cDNA by Dr. R.Y. Tsien (Campbell et al., 2002; Shaner et al., 2004).

UV-C irradiation. Lamp-induced UV damage was inflicted using a UV source containing four UV lamps (Philips TUV 9W PL-S) above the microscope stage. The UV dose rate was measured to be 3 W.m-2 at 254 nm. For induction of global UV-damage, cells were rinsed with medium and irradiated for 9 seconds (25 J.m- 2). For induction of local UV-damage, cells were UV irradiated through a polycarbonate mask (Millipore Billerica, Massachusetts, USA) with pores of 3 or 5 µm for 39 or 11.7 seconds (100 or 30 J.m-2) (Luijsterburg et al., 2007). Laser-induced UV damage was inflicted by using a 2 mW pulsed (7.8 kHz) diode pumped solid state laser emitting at 266 nm (Rapp OptoElectronic, Hamburg GmbH). The laser was connected to a Zeiss LSM 510 confocal microscope with an Axiovert 200 M housing adapted for UV by all-quartz optics. A special adaptor (ZSI-A200, Rapp OptoElectronic) to fit in the aperture slider position of an Axiovert 200 microscope was developed by Rapp OptoElectronic to focus the laser on a sample. For local UV-C irradiation experiments, cells were grown on 25 mm diameter quartz coverslips (010191T-AB, SPI supplies) (Dinant et al., 2007).

α-particle irradiation. Cells were plated in custom-made culture dishes containing an ultra-thin Mylar bottom. (Aten et al., 2004; Stap et al., 2008). The Mylar membrane was coated with carbon to improve attach- ment of cells. Cells were incubated for 24 hours at 37 ºC and subsequently irradiated using an americium (Am-241) source with an activity of 140 kBq. The americium source was placed underneath the dish, just touching the Mylar membrane. The source was placed at an angle of 30° with the horizontal plane to obtain long linear arrays of DSBs. Cells were irradiated at room temperature for 0.5 min and subsequently fixed using paraformaldehyde (final concentration 2%).

Immunofluorescence. Fixed cells were immunostained as described previously (Aten et al., 2004; Stap et al., 2008). The following primary antibodies were used: mouse monoclonal antibody against γH2A.X (clone JBW301, Upstate Biotechnology, Waltham, MA, USA), mouse monoclonal antibody against XPA (clone

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12F5, Abcam) and rat monoclonal antibody against HP1β (1:250; a kind gift from Dr. Prim Singh (Wreggett et al., 1994). Secondary antibodies: goat anti-mouse Cy3 and goat anti-rat FITC (both from Jackson Immu- noResearch Laboratories, West Grove, Pennsylvania, USA). All antibodies were diluted in PBS containing 0.1% Triton-X 100 and 1% Foetal Calf Serum. Fluorescence microscopy images were acquired using a Leica DM RA HC microscope (Leica Microsystems, Wetzlar, Germany) equipped with a PLAN APO 100x / 1.40 oil objective and a cooled CCD camera (KX1400, Apogee Instruments, CA, USA).

Microscopic analysis. Cells were imaged on a Zeiss LSM 510 confocal microscope, equipped with a 63x Ultrafluar (1.2 NA) glycerol immersion lens (Zeiss, Oberkochen, Germany) and a 30 mW Argon laser (488 and 514 nm), or on a Zeiss Axiovert 200M wide field fluorescence microscope, equipped with a 100x Plan- Apochromat (1.4 NA) oil immersion lens (Zeiss, Oberkochen, Germany) and a Cairn Xenon Arc lamp with monochromator (Cairn research, Kent, U.K.). Images were recorded with a cooled CCD camera (Coolsnap HQ, Roper Scientific, USA). Both microscopes were equipped with an objective heater and cells were examined in microscopy medium (137 mM NaCl, 5.4 mM KCl, 1,8 mM CaCl2, 0.8 mM MgSO4, 20 mM D- glucose and 20 mM HEPES) at 37°C.

Combined FLIP and FRAP. FRAP analysis was used to measure the immobilization of EGFP-HP1β after global UV irradiation as described by Houtsmuller and co-workers (Hoogstraten et al., 2002; Zotter et al., 2006). Briefly, ~50% of the nuclei of NIH/3T3 cells expressing EGFP-HP1β were bleached using 100% power of a 488 and 514 laser line. The loss of fluorescence from the non-bleached half (FLIP) and the gain of fluorescence in the bleached half (FRAP) were determined. The data was corrected for background values and normalized to pre-bleach intensity.

C. elegans UV-B survival assay. C. elegans strains used were Bristol N2 (wild type), hpl-1(tm1624), hpl- 2(tm1489), hpl-2(tm1489); hpl-1(tm1624) (constructed using PCR to confirm deletions) and xpa-1(ok698). The hpl-1(tm1624) and xpa-1(ok698) are likely null alleles (Schott et al., 2006; Stergiou et al., 2007), whereas hpl-2(tm1489) is a null allele that causes severe growth delay and sterility only at 25°C but not at 20°C (Coustham et al., 2006). Thus, it was possible to test UV-sensitivity by growing animals at 20°C and trans- ferring them to 25°C directly following UV-irradiation. To test UV-sensitivity, eggs were collected from gravid adult animals by ClNaO/NaOH treatment and placed on culture plates seeded with HT115 bacteria at 20°C. Eight hours following egg collection, eggs were irradiated using two FS20 erythemal UV-B lamps (Phillips, Eindhoven, the Netherlands), after which survival rate was determined by counting the amount of hatching eggs (surviving animals) and non-hatching, dead eggs. Additional details are available upon request.

Acknowledgement This work was supported by ZonMW, the Netherlands (grant 912-03-012). The authors thank Drs. P. Hemmerich, T. Misteli, R.W. Dirks, R.Y.Tsien, J. Goedhart, H. Leonardt, M.C. Cardoso, P.D. Varga-Weisz, K.E. Schuebel and T. Jenuwein for kindly providing constructs and cells and Dr. P. Singh for providing HP1 antibodies. The authors are grateful to the Caenorhabditis Genetics Center and the Japanese National Bioresource Project for hpl-1, hpl-2 and xpa-1 strains. The authors thank Drs. E.M.M. Manders and T.W.J. Gadella (Center for advanced microscopy (CAM)/UvA) for support.

138 Chapter 8 Perspective: Assembly of Multi-Protein Complexes that Control Genome Function

Christoffel Dinant, Martijn S. Luijsterburg, Wim Vermeulen, Adriaan B. Houtsmuller and Roel van Driel

In preparation

139 Chapter 8

Introduction

The nucleus is the information repository of the cell. In this ~300 µm3 container, mammalian cells store the genetic material. The 3 billion base-pairs of the human genome associate with histone proteins to form a chain of basic units of chromatin: the nucleosomes. Functioning of the genome is controlled by a variety of processes that each involve tens of proteins that cooperate in space and time. Examples are transcription initiation complexes, replication machineries and DNA repair mechanisms. In vivo studies have revealed that many proteins that control genome function rapidly diffuse inside the mammalian nucleus (Phair and Misteli, 2000) with apparent diffusion rates ranging between ~1 and 10 µm2/s depending on the shape and the size of the molecule. If such a diffusing protein encounters a site for which it has a finite affinity, it binds for a certain time depending on its dissociation rate. Many nuclear proteins exchange fast between the bound and free state at the scale of seconds (e.g. transcription factors) or minutes (e.g. repair proteins; Luijsterburg et al., 2007; Phair et al., 2004). Due to the short binding time of individual chromatin-binding proteins, the formation of a multiprotein complex consisting of tens of protein molecules is a low probability event, unless special processes are implemented. Although the binding kinetics of many individual proteins have been measured, little is known about how proteins assemble into the functional multiprotein complexes that are involved in controlling genome function in living cells. In this perspective, we focus on the general mechanism and kinetics of the assembly of multiprotein complexes involved in genome functioning, studied using live-cell imaging. We discuss how proteins find their target site on the genome, a process that has mainly been studied using bacterial proteins binding to naked DNA. Recent in vivo studies suggest that similar mechanisms are used by chromatin-binding proteins in eukaryotes. Additionally, we give an overview of the binding kinetics of proteins involved in genome controlling processes, such as transcription of rRNA and mRNA genes, replication of the genome and DNA repair. Finally, we discuss how live-cell studies aided by mathematical modelling have unveiled characteristic properties of assembly of multi-protein complexes on the chromatin fibre, which appears to be similar for different genome controlling processes. We argue that complex data sets from kinetic studies cannot be interpreted intuitively and require mathematical modelling to gain comprehensive insight into the dynamic organization and functioning of multi-protein complexes that control genome function.

How do site-specific proteins find target site on chromatin? Essentially all processes that control genome function are carried out by complexes that contain tens of proteins that assemble on specific genomic sites to exert their function. Assembly of such multi-protein complexes at specific sites is often initiated by a recognition protein that is able to, for example, discriminate between damaged and

140 Perspective: Dynamic organization of chromatin-associated processes non-damaged DNA (initiating DNA repair) or recognize a specific sequence in a promoter region (initiating transcription). Several mechanisms have evolved to ensure that site-specific proteins bind their target sites with sufficient rates. Proteins often bind to specific and non-specific sites with similar on-rates. Differences in affinity are often determined by a slower rate of dissociation (Hopfield, 1974). Site-specific proteins diffuse through the nucleus until they transiently interact with chromatin by chance. Since non-specific (i.e. low affinity) sites are usually present in large excess over specific sites, it is more likely that a protein binds a non-specific site (Misteli, 2008). Thus, once a protein interacts with a low affinity site on chromatin it will be transiently bound and then dissociate (since it has a higher dissociation rate from low affinity sites) until it encounters a specific site (i.e. high affinity) from which it dissociates more slowly. Multiple association/dissociation events on non-specific sites will take place before a protein binds to a specific site more stably (Gorski et al., 2006; Misteli, 2007). Some proteins associate with their target sites several orders of magnitude faster than diffusion-limited association (~ 108 M-1s-1) to DNA in vitro (Berg et al., 1981; Riggs et al., 1970). This rapid association can be accounted for by movement of a protein from its initial non-specific site to its target site by one- dimensional (1D) diffusion along the DNA by a sliding and/or hopping mechanism, which involves electrostatic DNA-protein interactions (Berg et al., 1981; Elf et al., 2007; Halford and Marko, 2004; Shimamoto, 1999). It is not likely that many proteins bind to DNA with association rates above the diffusion limit in vivo due to the high concentration of salt (equivalent to >150 mM NaCl) in cells, which reduces electrostatic DNA-protein interactions and thus one-dimensional diffusion (Halford and Marko, 2004). Sliding implies diffusion along the DNA helix without losing contact, whereas hopping involves dissociation and re-binding near to the site where the protein last dissociated. Hopping may occasionally occur to a more distant site on linear DNA, since the genome is present in a highly folded state (Halford and Marko, 2004). However, experiments show that hopping proteins tend to associate close to the site where they last dissociated. It may be that this is due to long-range electrostatic DNA-protein interactions that “pull” the protein back to the DNA (Halford and Marko, 2004).

Many DNA-binding proteins are able to move along a DNA strand without dissociating from it, including restriction enzymes, transcription factors and DNA repair proteins (Blainey et al., 2006; Coppey et al., 2004; Gowers and Halford, 2003; Gowers et al., 2005; Graneli et al., 2006; Harada et al., 1999; Shimamoto, 1999). However, at physiological ionic strength, proteins only diffuse along the DNA over distances of about 50 bp, suggesting that 1D diffusion is not the main mode of translocation of site- specific proteins (Gowers et al., 2005). Several examples indicate that site-specific proteins in eukaryotes also have affinity for non-specific DNA sites. For instance, repair factor XPC continuously associates with and dissociates from chromatin and

141 Chapter 8 occasionally encounters a lesion that it binds to with higher affinity. More precisely, at each moment 55% of the XPC molecules are bound to DNA for on average 300 ms in undamaged cells (Hoogstraten and Vermeulen, in press). These 300 ms likely represent the time that XPC diffuses along the DNA until it dissociates or encounters a DNA lesion. Other damage recognition proteins, Rad51 and hOgg1, were shown by single molecule techniques to diffuse along double stranded DNA and bind with higher affinity upon encountering a lesion (Blainey et al., 2006; Graneli et al., 2006). Similarly, proteins involved in transcription initiation were found to move along DNA by 1D diffusion and bind more tightly once encountering promoter regions or other regulatory elements (Elf et al., 2007; Harada et al., 1999). Probably, proteins alternate between 1D sliding over short distances and short hops near the dissociation site (1D hop) interrupted by periods of free diffusion. In eukaryotes, sliding is likely restricted to the linker regions that have no histones bound (Hannon et al., 1986). In agreement, promoter localization by RNA polymerase is equally efficient for promoters in the linker DNA of chromatin in living cells as for promoters in naked DNA (Hannon et al., 1986). Moreover, the C-terminus of p53 was shown to mediate 1D-diffision along DNA and a p53 mutant lacking this domain was unable to bind specific promoters in vivo (McKinney et al., 2004), suggesting that 1D-diffusion is also important for locating target sites in a chromatin context. Since sliding is mainly due to electrostatic interactions, the sliding properties will be determined by the distribution of (mainly positively) charged residues on the protein surface that interact with DNA. A protein that displays 1D diffusion is often envisioned to track the major groove of DNA, thus spiralling around the helix as it diffuses along the DNA. Another possibility is that proteins diffuse freely on the DNA surface (termed 2D-diffusion). Experiments revealed that such a mechanism is indeed employed by bacterial proteins and modelling showed that 2D diffusion would allow a protein to bypass obstacles such as nucleosomes (Kampmann, 2004). Therefore, 2D diffusion of proteins might be more relevant in a chromatin context. Mere 3D diffusion is not sufficient to explain the high rates at which proteins associate with target sites (Halford and Marko, 2004). Mechanism that increase the rate of binding are diffusion along the DNA (either tracking the major groove or by diffusing freely on the cylindrical DNA surface) for short stretches combined with “hopping” and “jumping”. Clearly, the contribution of these processes to finding the target site depends on the biophysical properties of the protein and the number and nature of the binding sites.

Assembly of multi-protein complexes that control genome function Finding of a target site by a recognition-protein is only the starting point for genome controlling processes. Upon recognition of an origin of replication, DNA lesion or promoter by a recognition protein, multi-protein complexes often containing tens of proteins are assembled that subsequently carry out replication, DNA repair or transcription. Here, we give an overview of live-cell imaging studies on proteins that

142 Perspective: Dynamic organization of chromatin-associated processes form such multi-protein complexes during DNA repair, transcription and replication. First, we discuss the mechanism of protein complex assembly during DNA repair by the nucleotide excision repair system. This process has been studied extensively and may therefore serve as a paradigm for other chromatin-associated processes.

Assembly of DNA repair complexes. To protect the integrity of the genome, multiple DNA repair mechanisms have evolved to deal with specific DNA injuries (Essers et al., 2006; Hoeijmakers, 2001). For example, nucleotide excision repair (NER) removes helix-distorting injuries that affect one of the DNA strands, whereas homologous recombination (HR) and non- homologous end-joining (NHEJ) repair double strand breaks (DSBs). Particularly the kinetics of proteins involved in NER have been well studied. NER involves the assembly of repair complexes that contain up to 30 polypeptides, which cooperate in space and time. Damage-recognition protein XPC was recently shown to continuously associate and dissociate with chromatin for on average 300 ms per binding event (Hoogstraten and Vermeulen, in press) At any moment, about half of the XPC molecules is bound to chromatin and half diffuses freely. It is likely that the chromatin-bound fraction of XPC diffuses along the DNA during this short period, similar to 1D diffusion reported for some restriction enzymes and the lac repressor (Elf et al., 2007). In the absence of damage, other NER proteins XPA, XPG, and ERCC1/XPF diffuse rapidly through the nucleus with rates that correspond to their molecular size (Houtsmuller et al., 1999; Luijsterburg et al., 2007; Rademakers et al., 2003; Zotter et al., 2006). Upon DNA damage induction by UV-C light, XPC occasionally encounters a helix-distorting lesion to which it binds more stably (t1/2 = 25s) (Hoogstraten et al., 2008). Lesion detection by XPC subsequently triggers assembly of NER complexes on the site of damage from freely diffusing constituents. NER proteins XPG, TFIIH and ERCC1/XPF rapidly exchange (t1/2 ~1 min) with emerging repair complexes, while XPA exchanges somewhat slower (~2 min) and, as mentioned above, XPC exchanges is somewhat faster (t1/2 = 25s) (Houtsmuller et al., 1999; Luijsterburg et al., 2007; Rademakers et al., 2003; Zotter et al., 2006). Fast exchange of proteins between the diffusing and bound state may ensure a high free concentration of these factors allowing a dynamic competition for binding sites, which may facilitate cross-talk between different processes that control genome function. However, this fast exchange also makes the formation of functional multi-protein repair complexes a time-consuming process, since binding of all proteins to the same site at the same time has a low probability. Rather, there will be an ensemble of mostly in complete assembled repair complexes. Only a few complexes will have all the repair proteins bound at the same time. This does not appear specific for DNA repair complexes, since protein complex assembly was also shown to be slow during other genome-controlling processes (see below and (Darzacq et al., 2007; Dundr et al., 2002)). Indeed, mathematical modeling suggests that by far most of the repair time is

143 Chapter 8 spent on assembly of functional complexes, which is an inherent property of large multi-protein complexes (Luijsterburg and van Driel, unpublished). NER involves some of the same enzymatic reactions (e.g. helix unwinding and DNA synthesis) that are carried out during transcription and replication, which involves proteins that are shared between these different processes, such as TFIIH, RPA and PCNA. The nucleoplasmic pool of TFIIH is in a continuous equilibrium between RNA pol I, RNA pol II transcription and NER (if DNA damage is present). The binding time of TFIIH at transcription sites is in the order of seconds, while TFIIH is bound much longer to NER intermediates during repair (t1/2 ~50 s) (Hoogstraten et al., 2002). Similarly, PCNA and RPA are shared factors between replication and several repair pathways including NER (Essers et al., 2005; Solomon et al., 2004). PCNA is bound for about 3 minutes during replication, while its exchange rate at sites of DNA damage is almost twice as long (t1/2 ~300s). RPA is also more stably associated with repair sites than with replication (Solomon et al., 2004). Thus, it appears that TFIIH, RPA and PCNA have higher affinities for DNA repair intermediates than for sites of transcription and replication, respectively.

Repair of DSBs by HR involves assembly of a protein complex that includes the formation of a Rad 51 filament (in which the actual recombination events takes place) and binding of chromatin remodelling factors (Rad54) and additional repair proteins such as Rad52 and RPA. Live cell imaging revealed that Rad51 filaments are highly stable and the residence time of Rad51 proteins is in the order of hours. Exchange rates of Rad52 (~1min) and Rad54 (~10 s) are much faster (Essers et al., 2002). Thus, it appears that Rad51 is a structural component during HR, which serves as a binding platform for several other repair proteins that exchange rapidly between the bound and free diffusing state. Such a mechanism is reminiscent of replication in which PCNA is bound relatively stably. Besides HR, DSBs can be repaired by NHEJ if a sister chromatid is not present (i.e. such as in G1), which involves, among others, the ring-shaped Ku70/80 dimer and the kinase DNA-PKcs. Current models suggest that multiple Ku70/80 dimers encircle the broken DNA ends and become trapped there. After ligation of the broken ends, it has been suggested that Ku70/80 could be permanently trapped (Mari et al., 2006). However, live-cell imaging challenged this model and revealed that binding of Ku70/80 to broken DNA ends is reversible and that exchange between bound and soluble pools occurs within 40s. Moreover, Ku70 was shown to recruit LigIV via XRCC4 to broken DNA ends, which in turn joins the broken ends together (Mari et al., 2006). Similar to Ku70, DNA-PKcs was shown to exchange between soluble and DNA bound pools within 1 min. Nevertheless, when DNA-PKcs cannot be phosphorylated or perform its kinase activity, a much larger fraction of the DNA-PKcs pool is bound for longer times (Mari et al., 2006; Uematsu et al., 2007). These studies highlight the importance of post-translational modification of repair proteins and show that, in case of DNA-PKcs, phosporylation is needed to

144 Perspective: Dynamic organization of chromatin-associated processes lower its affinity for repair intermediates.

Collectively these studies show that, with the exception of Rad51 filaments, repair machineries assemble at sites of DNA damage from freely diffusing components and form short-lived complexes on damaged chromatin. Affinity differences between specific and non-specific sites are small. Likely, the affinity of repair proteins is precisely tuned such that transient binding is sufficient to assemble complexes at specific (in this case damaged) sites with an acceptable rate, while at the same the low affinity ensures that complex assembly at non-specific sites is not very likely. Indeed, a recent study showed that tethering of DSB repair proteins Mre11, Rad50 or Nbs1 to chromatin elicits a DNA damage response including activation of Chk1/Chk2 and cell cycle arrest (Soutoglou and Misteli, 2008). This confirms that binding of a single repair protein with a low koff (high affinity binding) is sufficient to trigger a cellular response.

Assembly of transcription initiation and elongation complexes. Transcription involves assembly of a multi-protein transcription initiation complex on the chromatin fibre. Various live-cell imaging studies in combination with kinetic modelling have unveiled that, like DNA repair proteins, transcription factors, co- activators and RNA polymerases (RNA pol) bind rapidly and reversibly to target-sites (promoters) in a stochastic fashion (Becker et al., 2002; Hager et al., 2006). Occasionally, these factors assemble in a way that leads to onset of transcription and the production of RNA (Darzacq et al., 2007). Several transcription factors and co- activators (e.g. GR, GRIP-1, p53, TFIIB and TFIIH) diffuse rapidly through the nucleus and at any given time about 15% - 25% of these proteins is bound for 3-5 s to chromatin (Chen et al., 2002; Hoogstraten et al., 2002; Mueller et al., 2008). Although short residence times (i.e. several seconds) are common for transcription factors, some have binding times in the order of 1 min (e.g. AR and TBP) (Chen et al., 2002; Farla et al., 2004). The majority (~85%) of RNA pol II in living cells rapidly exchanges within on average 6 s, reflecting reversible binding to promoters or to non-specific sites with low affinity (Darzacq et al., 2007). The remaining 15% of RNA pol II molecules binds for on average about 1 min, reflecting transcription initiation attempts at the promoter. About half of these 1 min binding events leads to elongation with ~70 nt/s during which RNA pol II exchanges slowly from transcribed genes (~10 min) (Darzacq et al., 2007; Kimura et al., 2002). These live-cell studies combined with mathematical modelling reveal that only 1% of the RNA pol II binding events at promoters lead to a complete RNA molecule, showing that the majority of RNA pol II-promoter interactions are not productive (Darzacq et al., 2007). This at first sight inefficient onset of transcription may indicate that there is only a low probability of forming an active transcription initiation complex at a promoter. Intriguingly, a small fraction (~5%) of elongating polymerases pauses for several minutes without dissociating.

145 Chapter 8

Thus, at any time, at least one-fourth of transcribed genes have a paused RNA pol II bound resulting in discontinuous production of mRNAs (Darzacq et al., 2007), providing an explanation for transcriptional bursts. Recently, the steady-state level of a bound transcription factor to a natural promoter was found to oscillate in vivo on a time-scale of 20-40 min after transcription initiation by stimulation with a hormone, while the exchange rate of this transcription factor with chromatin was in the order of 1 min. These slow oscillations in steady-state levels of bound transcription components have also been detected by ChIP (Metivier et al., 2006) and it has been suggested that these slow cycles reflect stable interactions of activators with promoters leading to mRNA production, while fast cycles reflect transient interactions of activators that do not result in mRNA production (Metivier et al., 2006).

Live-cell imaging suggested that rapid exchange (within 1 min) of transcription factors reflects active transcription, whereas slow cycles reflect gradual changes in accessibility of binding sites at promoters (resulting in 20-40 min oscillations) (Karpova et al., 2008). In agreement, the occupancy of histones at promoters was also found to oscillate and to inversely mirror the slow cycle of a transcriptional activator. However, it is currently not clear, why the number of accessible binding sites of transcription factors gradually changes in time and whether this is functionally important for transcription regulation. In addition to RNA pol II, transcription of rRNA genes by the RNA pol I system is also highly dynamic (Dundr et al., 2002). The majority of pre-initiation factors (UBF1 and 2), transcription factors (TAFI48) as well as individual subunits of RNA pol I rapidly exchanges within 5 s at rRNA genes, while TFIIH exchange in nucleoli is considerably slower (~25s) (Hoogstraten et al., 2002). Similar to RNA pol II transcription, only 1-3% of the RNA pol I binding events result in elongation, which is inefficient in terms of association/dissociation steps needed to initiate transcription. However, because there are several hundred transcription factors binding events per second, such an 'inefficient' mechanism still results in rapid assembly of functional complexes in time. Interestingly, RNA pol I subunits exchange on rRNA promoters about 4 times slower in S-phase, during which the rRNA transcriptional output is much higher than in G1, suggesting that longer binding time of individual RNA pol I subunits to promoters results in more efficient assembly of functional RNA pol I complexes able to produce an rRNA (Gorski et al., 2008). Thus, modulation of RNA pol I assembly kinetics is a mechanism to control the transcriptional output of rRNA genes in living cells. In conclusion, these studies show that the assembly of active transcription initiation complexes is slow, similar to the formation of repair complexes. It is not likely that tens of proteins bind to the same site at the same time and the onset of transcription therefore takes multiple association/ dissociation events. Additionally, the 'inefficiency' of complex assemble during transcription serves as a regulatory mechanism that reduces the incorporation of wrong (i.e. non-specific binding) proteins in the complex, a process named kinetic

146 Perspective: Dynamic organization of chromatin-associated processes proofreading (see below).

Assembly of replication complexes. Faithful duplication of the genome is essential to maintain genome integrity and cellular identity. Besides DNA polymerases δ and ε, many proteins are involved in replication, including clamp-loader PCNA, endonuclease Fen1, ssDNA-binding protein RPAand DNA ligase 1 (Lig1) (Gorski et al., 2008; Liu et al., 2004; McCulloch and Kunkel, 2008). The leading strand is synthesized continuously during replication, while lagging strand synthesis requires the discontinuous synthesis and joining of Okazaki fragments. Fen1, RPA and Lig1 (and probably DNA polymerase δ), are specifically required for synthesis of the lagging strand (McCulloch and Kunkel, 2008). Initiation of replication involves origin recognition complex (ORC) proteins, which bind chromatin and exchange rapidly with soluble pools within 2 min (McNairn et al., 2005). This transient binding of ORC proteins to chromatin indicates that the mechanism to assemble these replication initiation proteins likely involves sliding and hopping, similar to assembly of repair and transcription initiation complexes. Replication of DNA occurs in discrete areas in the nucleus termed replication foci or factories, which appear at the onset of S phase and disappear as soon as replication is finished (Leonhardt et al., 2000). These microscopically visible replication foci represent clusters of replication forks and associated replication machineries and are often several MDa in size. Although replication foci appear to be static structures, the proteins that make up these structures exchange rapidly between the bound and freely diffusing pool (Kitamura et al., 2006; Misteli, 2007).

Live-cell imaging has shown that there is a remarkable difference in exchange kinetics between different replication proteins to replication foci. For example, exchange of clamp-loader PCNA at replication foci takes 3 min on average (Essers et al., 2005; Solovjeva et al., 2005; Sporbert et al., 2002), while proteins involved in lagging strand synthesis exchange much faster compared to PCNA. For instance, Fen 1 and DNA lig 1 exchange at replication foci takes about 1s (Solovjeva et al., 2005; Sporbert et al., 2005). Similarly, RPA exchanges in seconds (redistribution in 1 min) (Sporbert et al., 2002). The exchange rates of these proteins are in the same range as the time it takes to synthesize one Okazaki fragment (estimated at 6 s), which is roughly 180 bp in size (Jackson and Pombo, 1998; Solovjeva et al., 2005; Sporbert et al., 2005), suggesting that these proteins bind during the synthesis of one Okazaki fragment and dissociates upon completion of such a fragment. Fen1 and DNA lig1 are indeed required for Okazaki fragment maturation, performing removal of the RNA-DNA primer and sealing of the remaining nick, respectively (Liu et al., 2004). This favours a scenario in which proteins such as RPA, Fen1 and DNA lig1 exchange after each Okazaki fragment has been synthesized, whereas PCNA remains bound for the synthesis of multiple Okazaki fragments (Sporbert et al., 2005). Thus, it seems that PCNA serves

147 Chapter 8 as a binding platform for many proteins that are transiently involved in replication, including proteins involved in epigenetic maintenance such as Dnmt1, which methylates the newly synthesized DNA during replication (Schermelleh et al., 2007). The use of a binding platform for transiently interacting proteins is conceptually similar to transcription elongation, during which RNA pol II exchanges very slowly, while other proteins, such as TFIIB or TFIIH, can transiently interact with the active transcription machinery (Darzacq et al., 2007). Whether DNA polymerases act similarly as RNA pol II with respect to chromatin association kinetics has not been studied so far.

Understanding assembly and functioning of genome- controlling complexes. The study of GFP-tagged proteins involved in chromatin processes generates large and complex sets of data that generally cannot be interpreted intuitively (Phair and Misteli, 2001). Kinetic modelling of quantitative in vivo data is a powerful tool in obtaining mechanistic insight into genome-controlling processes. It allows estimation of biophysical properties of proteins such as diffusion, association and dissociation rate constants that cannot be determined directly in vivo. Moreover, modelling of the kinetic properties of chromatin-associated systems provides detailed insight into their behaviour (Darzacq et al., 2007; Dundr et al., 2002; Politi et al., 2005). Luijsterburg and van Driel, unpublised results). Although it is the often-held view that multi-protein complexes are assembled in a stepwise fashion, this is not supported by several in vivo studies. These studies combined with mathematical modelling revealed that proteins only occasionally form the complete protein complex on chromatin. For example, components of the RNA pol I machinery frequently associate with ribosomal genes with a rate of several hundred molecules per second, but only occasionally (1-10% of the binding events) form the correct complex that is capable of transcription (Dundr et al., 2002). Likewise, in vivo studies and modelling revealed that only 1% of the RNA pol II-promoter interactions results in completion of an mRNA (Darzacq et al., 2007), suggesting the formation of the complete protein complex is a low probability event. Recent modelling of the kinetics of NER in terms of the collective action of eight repair proteins and six enzymatic steps, revealed that protein complex assembly during NER is remarkably slow compared to the enzymatic processes. More than 99% of the repair time is spent on the assembly of catalytically active complexes, whereas enzymatic reactions during repair (such as unwinding of DNA and dual incision) take only a small fraction of the total repair time (Luijsterburg and van Driel, unpublished). Although the term inefficient has been used for these processes, it should be noted that if several hundred molecules bind per second and even only 1% of the binding events results in assembly of a functional complex, this still results in complex formation on specific sites at sufficient high rates. In conclusion, a low probability to assemble the correct protein complex appears a shared characteristic of different genome controlling processes.

148 Perspective: Dynamic organization of chromatin-associated processes

Figure 1. Model for binding of a site-specific protein to a target site and subsequent assembly of a multi- protein complex on that site. A site-specific protein (orange oval) diffuses rapidly inside the nucleus and binds non-specifically to chromatin (represented by the light-green line), dissociates and re-binds close to the site it dissociated from (so-called “hopping”). Finally, it moves along chromatin by 1-dimensional diffusion and encounters a specific site (dark-green) to which the protein binds more stably. The orange protein mediates the assembly of a complex consisting of two additional proteins (light-purple and dark-purple). Binding of all these proteins is stochastic and 8 different assembly states can be formed on the specific site consisting of one or a combination of the 3 proteins or the site can be devoid of any protein. Once the “correct” complex containing all 3 proteins is formed, the specific site is modified (for example acetylated shown in red) resulting in dissociation of the orange and dark-purple protein while the light-purple protein remains bound (since it has affinity for the altered state while the other proteins do not). The red arrow reflects an enzymatic step (in this case acetylation, which is in principal irreversible). This altered state is the substrate for a new set of proteins (the green and yellow protein and the light-purple protein from the last box) to bind to. Com- plex assembly is again stochastic and 8 different assembly states can be formed. Assembly of the “correct” complex containing all three proteins in the second box results in an enzymatic step that produces mRNA, resulting in dissociation of the yellow and light-purple protein. The probability of the overall reaction (i.e. binding of 5 different proteins to the same site) is increased by splitting the reactions in assembly of complex 1 and complex 2 separated by an irreversible enzymatic reaction. The irreversible step drives the reaction forward. Completion of processes involving more proteins can be kinetically driven by multiple irreversible reactions.

In this scenario, proteins bind stochastically (i.e. not in a fixed order of binding) to specific sites on chromatin to form an ensemble of protein complexes with different subunit composition. The correct protein complex, capable of performing a specific function (repair, transcription, replication), is formed with a low probability. This “chaotic” view on complex assembly is radically different from the often-held view that complex assembly on chromatin occurs through an ordered and stepwise mechanism in which each protein is incorporated in a long-lived chromatin-bound complex. This stochastic mechanism of complex assembly is schematically depicted in figure 1. Completion of a process in which assembly of large protein complexes is a low probability event can be kinetically driven by irreversible (often-enzymatic) reactions, which split up the process in smaller sub-processes that are executed sequentially. Such a mechanism, known as kinetic proofreading, increases the specificity of molecular processes above the level of difference in free energy between correct and incorrect

149 Chapter 8 substrates (Hopfield, 1974). This creates a system that rejects 'wrong' proteins, despite the similar kon rates of specific and non-specific interactions. Many genome- controlling processes involve enzymatic reactions including ATP-hydrolysis, DNA unwinding, cleavage of DNA, DNA synthesis and ligation, which due to their often irreversible nature can drive genome controlling processes to completion (see fig 1). Moreover, these irreversible reactions allow the specificity of DNA-binding proteins for specific sites and the specificity at which multi-protein complex are formed to be increased at a level that is not present at the level of individual proteins. Thus, it appears that cells have evolved robust mechanisms to assemble multi-protein complexes at specific sites on chromatin with high accuracy. These mechanisms can only be unravelled using live-cell analyses combined with mathematical modelling and this combined approach is expected to provide insight into behaviour of systems that control the genome, in the years to come.

Acknowledgement This work was supported by ZonMW, the Netherlands (grant 912-03-012).

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174 175 176 Summary

Summary

The nucleus is the control centre of the mammalian cell. It is in this container that the genetic material DNA is stored as chromatin. Despite the fact that DNA is a rather stable molecule, it is not inert and can be compromised in many ways during the lifetime of an organism. Integrity of the genome is continuously challenged by intracellular metabolism (such as the production of free radicals in mitochondria) as well as external sources such as ionizing and ultra-violet radiation. Chapter 1 gives an introduction into the field of DNA repair and summarizes evolutionary conserved mechanism to remove mutagenic and cytotoxic lesions from the genome. These mechanisms include base excision repair (BER), mismatch repair (MMR), double-strand break repair by homologous recombination (HR) and non-homologous end-joining (NHEJ) and Fanconi anemia repair. Nucleotide Excision Repair (NER) is a versatile DNA repair mechanism that removes a variety of structurally unrelated lesions from the genome. Placental mammals fully depend on NER for the removal of the UV-induced photoproducts. This repair pathway is the major topic of this thesis and is discussed in detail in chapter 2. In addition, we discuss chromatin remodelling events that are required to facilitate NER on a chromatin template.

The ten-subunit TFIIH complex has a dual role in transcription and in NER. Defects in TFIIH subunits can result in a combined disorder of Xeroderma Pigmentosum (XP) and Cockayne Syndrome (CS). Defects in the 3’ endonuclease XPG can also result in combined XP/CS and TFIIH and XPG were suggested to be part of the same protein complex (Ito et al., 2007). To determine if XPG and TFIIH are assembled into nascent NER complexes as a single factor, we studied the kinetics of GFP-tagged XPG in living cells in chapter 3. Using photobleaching techniques, immuno-labelling experiments and by measurements on the binding kinetics of repair factors to nascent repair complexes directly after DNA damage induction, we demonstrate that XPG and TFIIH are separately incorporated into NER complexes (Zotter et al., 2006). Moreover, by employing several cell lines defective in one of the TFIIH subunits, we show that recruitment of XPG to UV-induced DNA damage depends on functional TFIIH (Zotter et al., 2006). In support of our data, it was recently demonstrated that TFIIH accumulation at sites of DNA-damage is not hampered by the absence of functional XPG (Dunand-Sauthier et al., 2005; Thorel et al., 2004).

Mammalian nucleotide excision repair is initiated by the XPC complex, which has high affinity for UV-damaged DNA (Sugasawa et al., 1998; Volker et al., 2001). Besides XPC, mammalian cells express a second UV-damage recognition protein with high affinity for UV lesions: DDB2, which is part of an E3 ubiquitin (ub) ligase containing CUL4A, ROC1 and DDB1 (Groisman et al., 2003). This raises the question how these damage sensors co-operate to detect lesions. In chapter 4, the binding and

177 Summary dissociation kinetics of DDB2 to UV-damaged chromatin was studied. Moreover, DDB2 was expressed in cells lacking functional XPC (Luijsterburg et al., 2007). Various photobleaching and immunolabeling experiments reveal that DDB2 bind to damaged-DNA independently of XPC (Luijsterburg et al., 2007), in contrast to other NER factors (Volker et al., 2001). The other subunits of the E3 ligase complex (DDB1 and CUL4A) bind to UV-lesions with similar kinetic as DDB2, demonstrating that the entire E3 complex binds to UV-induced DNA damage. We demonstrate that there is little physical interaction between the two damage sensors on UV-lesions and propose a scenario in which DDB2 prepares UV-damaged chromatin for assembly of the NER complex, possibly by histone ubiquitylation.

The nucleus is highly compartmentalized and electron microscopic studies have revealed that chromosomes are made up of several dense chromatin domains. At the border of these domains is a region called perichromatin, which contains dispersed chromatin fibers (Fakan, 2004a; Fakan, 2004b; Jaunin and Fakan, 2002; Jaunin et al., 2000). Transcription and replication mainly takes place in perichromatin (Cmarko et al., 1999; Jaunin et al., 2000), showing that this is an important functional compartment in the nucleus. In chapter 5, we studied where NER actually takes place in the nucleus using electron microscopy. NER proteins are significantly enriched in perichromatin upon UV irradiation. These results demonstrate that NER takes place in the same functional domain as transcription and replication. In addition, we provide evidence for considerable chromatin decondensation in response to UV-induced DNA damage, which might facilitate relocation of DNA lesions to the perichromatin region. Together, our results provide novel insight into the spatial organization of DNA repair in the human cell nucleus.

In chapter 6, we have systematically analyzed kinetic parameters of all pre-incision NER factors, in addition to repair synthesis factors RPA and PCNA during the repair process in living cells. Different NER proteins associate with the DNA lesion with remarkably different binding kinetics. NER proteins continuously and rapidly exchange between the bound and freely diffusing state, suggesting that repair proteins dissociate and associate while repair is in progress. Based on these quantitative in vivo data, we have developed a comprehensive kinetic model for NER, which quantitatively reproduces many aspects of the repair process in vivo. The model indicates that the NER process progresses through a sequential set of intermediates separated by in practice irreversible enzymatic steps, including unwinding, incision and DNA synthesis. Repair proteins assemble in a stochastic fashion on each intermediate until a catalytically active complex is formed that allows progression to the next repair intermediate. Using this model, we explore the dynamic behaviour and properties of the NER system to gain detailed insight into how it operates in vivo. Among others, the model reveals that the assembly of the repair complex is remarkably slow, since

178 Summary more than 99% of the repair time is spent on NER complex formation. Our approach provides detailed insight into the kinetic properties of NER in vivo and unravels the complex choreography of this repair process, which is a paradigm for other chromatin-associated processes.

Chromatin contains inherited information, which is stored in the genetic material DNA. An additional layer of epigenetic information is stored in the histone proteins that package DNA. These proteins can be modified by acetylation, methylation, phosphorylation and ubiquitylation and these modifications constitute a “code” that is “read” by a variety of epigenetic factors, which in turn regulate genome function. One of such epigenetic regulators is the HP1 family and HP1 proteins play roles in maintenance of heterochromatin, replication and transcription. In chapter 7, we demonstrate that that the HP1 isoforms HP1α, HP1β and HP1γ are recruited to DNA lesions in living human cells. This response to DNA damage strictly requires the chromoshadow domain of the HP1 protein and occurs at sites of double strand breaks and at sites of helix-distorting lesions. Evidence is provided that HP1 is not involved DNA repair itself. We show that several HP1-interacting proteins are also recruited to DNA damage, including chromatin remodelling factors and DNA and histone methyltransferases. Loss of HP1 results in high sensitivity to UV-induced damage in the nematode C. elegans, providing evidence for involvement of HP1 in a DNA damage response mechanism. These results suggest a novel function of HP1 proteins and reveal a link between maintenance of epigenetic information and DNA damage responses.

In the final chapter 8, we give an overview of how site-specific proteins locate target site in the genome. The kinetics of proteins involved in essential genome-controlling functions such as transcription, replication and DNA repair is discussed. Moreover, we discuss mechanisms to increase the probability that a multi-protein complex is assembled on a specific site and argue that full understanding of the dynamics of genome controlling processes requires live-cell imaging combined with mathematical modelling.

179 Samenvatting

Samenvatting voor de leek

Ieder mens bestaat uit miljarden cellen waaruit de verschillende organen en weefsel zijn opgebouwd. Zo zijn er groepen cellen die samen de hersenen, de lever of andere organen in het lichaam vormen. De instructies oftwel het bouwplan om een menselijk lichaam te maken liggen opgeslagen in het DNA wat er uitziet als een wenteltrap. Bijna alle cellen bevatten een 2 meter lange DNA draad met daarop alle instructies die de cel nodig heeft om te weten wat voor een soort cel hij is. Deze lange draad is netjes opgerold en past in de kern van een cel die een volume heeft van minder dan een miljoenste van een miljoenste van een liter. Dus stel je een pak melk voor en neem hier één miljoenste deel van. Dit is een druppel die je nog net met het blote oog kunt zien. Neem van deze druppel weer één miljoenste deel en je hebt de inhoud van een kern van een cel. Het is dus zeer opmerkelijk dat in deze onzichtbare druppel ruim 2 meter DNA past! Met het blote oog zijn kernen van cellen niet te zien en er is in dit proefschrift dan ook gebruik gemaakt van microscopen die objecten velen malen kunnen vergroten zodat cellen en hun kernen zichtbaar gemaakt kunnen worden. Terug naar het DNA in de kern. Het is van groot belang dat de instructies opgeslagen in het DNA niet verloren gaan doordat bijvoorbeeld het DNA beschadigd raakt of breekt. Het DNA kan bijvoorbeeld kapot gaan door straling afkomstig van de zon. Dit is een van de grootste problemen voor austronauten in de ruimte, die niet meer beschermd worden door de dampkring. Zij lopen veel schade aan hun DNA op door straling van de zon. Het gevolg van schade aan het DNA kan zijn dat cellen direct dood gaan omdat de cel niet meer kan functioneren. Anderzijds kunnen er fouten in het DNA ontstaan waardoor de instructies onleesbaar worden. Zo kan bijvoorbeeld de boodschap verloren gaan dat iedere cel zich maar maximaal 60 keer mag vermenigvuldigen. Cellen kunnen zich dan ongeremd gaan delen en er ontstaat kanker.

Hoe voorkomen cellen nu dat de boodschap onleesbaar wordt door schade aan het DNA? De cel bevat werkpaarden in de vorm van eiwitten. Dit zijn kleine moleculen waarvan soms wel één miljoen exemplaren in de cel aanwezig zijn die verschillende taken uitvoeren, zoals het kopiëren en het aflezen van het DNA. Zo zijn er ook eiwitten die beschadigd DNA repareren. Verschillende van dit soort eiwitten werken samen (soms wel met 30 eiwitten tegelijk) en vormen een DNA reparatiesysteem. In hoofdstuk 1 staan de verschillende reparatiesystemen in de cellen van mensen beschreven. In dit proefschrift is één van deze systemen (die schade door ultra-violet licht van de zon opruimt) in detail onderzocht. Dit reparatiesystem staat beschreven in hoofdstuk 2. Al kunnen we dus cellen en celkernen met de microscoop zien, reparatie-eiwitten zijn te klein om te kunnen waarnemen. In dit proefschrift is onder de microscoop onderzocht hoe reparatie-eiwitten samen werken om schade aan het DNA te repareren in levende cellen. De reparatie-eiwitten zijn hierbij zichtbaar gemaakt door er een lampje aan te hangen wat licht uitzendt. Dit lampje is een

180 Samenvatting fluorescerend eiwit (vaak groen, maar soms ook geel en rood) wat afkomstig is uit lichtgevende kwallen of koraal. Door dit lichtgevende eiwit aan een reparatie-eiwit te koppelen kunnen we het licht van het lampje opvangen met de microscoop en metingen verrichten aan hoe snel de verschillende eiwitten hun werk doen in levende cellen. In hoofdstuk 3, 4, 6 en 7 hebben we gebruikt gemaakt van deze truc om te meten hoe snel reparatie-eiwitten de plekken waar het DNA kapot is kunnen vinden in de cel en daarna samen werken om de schade op te ruimen. Dit klinkt niet al te ingewikkeld, maar stel je maar eens voor dat je met een blinddoek om door de Kalverstraat in Amsterdam rent, samen met 19 andere mensen die ook een blinkdoek om hebben. Je bent op zoek naar één bepaalde winkel in deze lange straat en jullie moeten alle 20 tegelijkertijd in deze winkel staan in een zo kort mogelijke tijd. Al klinkt dit bijna onmogelijk, dit is precies wat reparatie-eiwitten moeten doen om plekken te vinden waar het DNA kapot is om het daarna samen te repareren! In hoofdstuk 6 rekenen we met behulp van een wiskundig model uit hoe snel reparatie-eiwitten tegelijk op plekken waar het DNA beschadigd is terecht komen. De berekeningen maken duidelijk dat de reparatie- eiwitten héél veel pogingen nodig hebben om elkaar te vinden en samen de schade op te ruimen. Ondanks dat het dus heel vaak niet lukt om elkaar op tijd te vinden in de kern (net zoals je waarschijnlijk met je blinkdoek om in de Kalverstraat de 19 anderen personen ook niet kunt vinden), lukt het toch vaak genoeg wel. De eiwitten ruimen samen veel schades op in het DNA (soms wel meer dan 50.000) in een tijd van een aantal uur, wat een erg uitzonderlijke prestatie is. Dit verklaart waarom we buiten in de zon kunnen lopen (waarbij iedere cel in de huid ongeveer 30.000 schades per uur aan het DNA oploopt) zonder dat we hier direct nadelige effecten van ondervinden. Ons reparatiesysteem kan de schade aan het DNA behoorlijk effectief repareren.

In hoofdstuk 5 bekijken we cellen onder een speciale microscoop die objecten zo vaak kan vergroten dat we het opgevouwen DNA in de kern kunnen zien. Nu blijkt dat de reparatie-eiwitten niet overal de plekken waar het DNA beschadigd is opzoeken en repareren, maar dit alleen op specifieke plaatsen in de kern doen. Het lijkt er dus op dat DNA wat beschadigd is naar een bepaalde plek in de kern wordt gebracht om daar gerepareerd te worden. Dit is natuurlijk erg opmerkelijk want hoe breng je de beschadigde stukken van een 2 meter lange DNA draad naar elkaar toe zonder dat de hele draad enorm in de knoop komt te zitten? In het voorbeeld van de Kalverstraat zou dit betekenen dat de winkels in de straat zich zo rangschikken dat alle winkels waar jij en de andere 19 mensen naartoe moeten bij elkaar komen te liggen. Daarna is de kans natuurlijk groter dat alle 19 mensen, nog steeds met blinkdoek om, tegelijkertijd in de juiste winkel terecht komen. In hoofdstuk 8 vergelijken we DNA reparatie-systemen met andere systemen in de kern die ook gebaseerd zijn op het bij elkaar komen van tientallen eiwitten op dezelfde plek om samen iets voor elkaar te krijgen. Het is waarschijnlijk dat andere systemen op een vergelijkbare manier werken/functioneren als DNA reparatie. Kortom, het werk dat in dit proefschrift beschreven staat, heeft ons

181 Samenvatting duidelijker gemaakt hoe reparatie-eiwitten samen werken om schade aan het DNA te vinden en te herstellen. Het opruimen van schade aan het DNA is essentieel voor het gezond houden van het DNA en voor het leesbaar houden van de daarin opgeslagen boodschap. Het onderzoeken en volledig begrijpen van hoe schade aan het DNA wordt gerepareerd is zeer belangrijk omdat het onstaan van kanker, maar ook het ouder worden van cellen (en dus van mensen), direct te maken heeft met het niet goed opruimen van schades aan het DNA.

182 Curriculum Vitae

Curriculum Vitae

De auteur van dit proefschrift werd geboren op 19 maart 1980 te Utrecht. Na in 1997 zijn HAVO diploma aan het St. Gregorius college in Utrecht behaald te hebben begon hij in datzelfde jaar aan de studie moleculaire biologie aan de hogeschool van Utrecht. Als onderdeel van deze studie begon hij in september 2000 aan een 9 maanden durend afstudeervak bij de vakgroep immunologie aan de Universiteit van Utrecht onder begeleiding van Dr. C.P. Broeren. Tijdens deze periode heeft hij een methode ontwikkeld om zoogdier cellen MHC klasse II moleculen tot expressie te laten brengen waaraan gedefinieerde peptiden gebonden zijn. Nadat hij zijn studie in 2001 afrondde begon hij in datzelfde jaar aan de studie medische biologie aan de Vrije Universiteit Amsterdam. Als afsluiting van deze studie begon hij in februari 2003 aan een 7 maanden durend afstudeervak bij de vakgroep Fysica van complexe systemen aan de Vrije Universiteit Amsterdam onder begeleiding van Dr. R.Th. Dame. Hier werd onderzoek gedaan naar de rol van architecturele eiwitten in de organisatie van bacterieel chromatine met behulp van scanning force microscopy. Na het afronden van de studie medische biologie bleef de auteur nog een half jaar werkzaam als onderzoeker bij dezelfde groep. In januari 2004 begon de auteur als promovendus bij de vakgroep kernorganisatie van de Universiteit van Amsterdam onder de begeleiding van Prof. dr. R. van Driel. Het hier uitgevoerde onderzoek aan de dynamiek van DNA reparatie eiwitten in levende cellen staat beschreven in dit proefschrift. Vanaf 1 mei 2008 is de auteur werkzaam als postdoc bij de groep van Dr. N.P. Dantuma verbonden aan het Karolinska Instituut in Stockholm, Zweden. Hier zal onderzoek worden verricht aan de rol van histon ubiquitylatie in gen expressie.

List of publications

Dame, R.T., Luijsterburg, M.S., Krin, E., Bertin, P.N., Wagner, R. and Wuite, G.J. (2005) DNA bridging: a property shared among H-NS-like proteins. J Bacteriol, 187, 1845-1848.

Dame, R.T.*, van Mameren, J.*, Luijsterburg, M.S., Mysiak, M.E., Janicijevic, A., Pazdzior, G., van der Vliet, P.C., Wyman, C. and Wuite, G.J. (2005) Analysis of scanning force microscopy images of protein-induced DNA bending using simulations. Nucleic Acids Res, 33, e68.

Luijsterburg, M.S.*, Noom, M.C.*, Wuite, G.J. and Dame, R.T. (2006) The architectural role of nucleoid-associated proteins in the organization of bacterial chromatin: a molecular perspective. J Struct Biol, 156, 262-272.

183 Curriculum Vitae

Zotter, A.*, Luijsterburg, M.S.*, Warmerdam, D.O., Ibrahim, S., Nigg, A., van Cappellen, W.A., Hoeijmakers, J.H., van Driel, R., Vermeulen, W. and Houtsmuller, A.B. (2006) Recruitment of the nucleotide excision repair endonuclease XPG to sites of UV-induced DNA damage depends on functional TFIIH. Mol Cell Biol, 26, 8868-8879.

Mateos-Langerak, J., Brink, M.C., Luijsterburg, M.S., van der Kraan, I., van Driel, R. and Verschure, P.J. (2007) Pericentromeric heterochromatin domains are maintained without accumulation of HP1. Mol Biol Cell, 18, 1464-1471.

Williams, E.S., Stap, J., Essers, J., Ponnaiya, B., Luijsterburg, M.S., Krawczyk, P.M., Ullrich, R.L., Aten, J.A. and Bailey, S.M. (2007) DNA double-strand breaks are not sufficient to initiate recruitment of TRF2. Nat Genet, 39, 696-698

Luijsterburg, M.S.*, Goedhart, J.*, Moser, J.*, Kool, H., Geverts, B., Houtsmuller, A.B., Mullenders, L.H.F, Vermeulen, W. and van Driel, R. (2007) Dynamic in vivo interaction of DDB2 E3 ubiquitin ligase with UV- damaged DNA is independent of damage-recognition protein XPC. J Cell Sci, 120, 2706-2716.

Hoogstraten, D., Bergink, S., Verbiest, V.H.M., Luijsterburg, M.S., Geverts, B., Raams, A., Dinant, C., Hoeijmakers, J.H.J., Vermeulen, W., and Houtsmuller, A.B. (2008) Versatile DNA damage probing by the global genome nucleotide excision repair protein XPC. (J Cell Sci, In press)

Solimando, L*, Luijsterburg, M.S*., Vecchio, L*., van Driel, R., Fakan, S. (2008) Spatial Organization of Nucleotide Excision Repair Proteins After UV-induced DNA Damage in the Human Cell Nucleus (submitted)

Luijsterburg, M.S.*, Dinant, C.*, Wiernasz, E., Stap, J., Warmerdam, D.O., Lans, H., Brink, M.C., Dobrucki, J.W., Vermeulen, W., Mullenders, L.H.F., Houtsmuller, A.B., Verschure, P.J., van Driel, R. (2008) Heterochromatin Protein 1 is Involved in the DNA Damage Response (submitted)

Luijsterburg, M.S., White, M., van Driel, R., and Dame R.T. (2008) The Major Architects of Chromatin: Architectural Proteins in Bacteria, Archaea and Eukaryotes (submitted)

Alekseev, S. Luijsterburg, M.S., Pines, A., Geverts, B., Mari, P.O., Giglia-Mari, G. Lans, H., Houtsmuller, A.B., Mullenders, L.H.F., Hoeijmakers, J.H.J., and Vermeulen, W. (2008) Cellular concentrations of DDB2 regulate dynamic binding of DDB1 at UV-induced DNA damage (submitted)

184 Curriculum Vitae

Brink, M.C., Piebes, D., Luijsterburg, M.S., Kooistra, S., van Driel, R., and Verschure, P.J. (2008) MeCP2 targeting induces decondensation of higher order chromatin structure in vivo (In preparation)

Luijsterburg, M.S., von Bornstaedt,G., Politi, A.Z., Gourdin, A.M., Moné, M.J., Warmerdam, D.O., Goedhart, J, Vermeulen, W., Höfer, T., and van Driel, R. (2008) Choreography of Nucleotide Excision Repair Proteins Unravelled by Live-Cell Imaging and Kinetic Modelling (In preparation)

Dinant, C.*, Luijsterburg, M.S.*, Vermeulen, W., Houtsmuller, A.B., and van Driel, R. (2008) Assembly of Multi-Protein Complexes that Control Genome Function (In preparation)

* Equal author contributions

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