UV-DDB-dependent regulation of nucleotide excision repair kinetics in living cells

DNA Repair 8 (2009) 767–776

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UV-DDB-dependent regulation of nucleotide excision repair kinetics in living cells Ryotaro Nishi a,b,c , Sergey Alekseev d , Christoffel Dinant d,e , Deborah Hoogstraten d , Adriaan B. Houtsmuller e , Jan. H.J. Hoeijmakers d , Wim Vermeulen d , Fumio Hanaoka b,f,1 , Kaoru Sugasawa a,b,c,∗ a

Biosignal Research Center, Organization of Advanced Science and Technology, Kobe University, Kobe, Hyogo 657-8501, Japan SORST, Japan Science and Technology Agency, Kobe, Hyogo 657-8501, Japan Genome Damage Response Research Unit, RIKEN Discovery Research Institute, Wako, Saitama 351-0198, Japan d MGC-CBG, Department of Cell Biology and Genetics, Erasmus Medical Center, 3000 CA Rotterdam, The Netherlands e Department of Pathology, Erasmus Medical Center, 3000 CA Rotterdam, The Netherlands f Graduate School of Frontier Biosciences, Osaka University, Suita, Osaka 565-0871, Japan b c

a r t i c l e

i n f o

Article history: Received 16 July 2008 Received in revised form 30 December 2008 Accepted 17 February 2009 Available online 21 March 2009 Keywords: Nucleotide excision repair Dynamics XPC UV-DDB

a b s t r a c t Although the basic principle of nucleotide excision repair (NER), which can eliminate various DNA lesions, have been dissected at the genetic, biochemical and cellular levels, the important in vivo regulation of the critical damage recognition step is poorly understood. Here we analyze the in vivo dynamics of the essential NER damage recognition factor XPC fused to the green fluorescence protein (GFP). Fluorescence recovery after photobleaching analysis revealed that the UV-induced transient immobilization of XPC, reflecting its actual engagement in NER, is regulated in a biphasic manner depending on the number of (6-4) photoproducts and titrated by the number of functional UV-DDB molecules. A similar biphasic UV-induced immobilization of TFIIH was observed using XPB-GFP. Surprisingly, subsequent integration of XPA into the NER complex appears to follow only the low UV dose immobilization of XPC. Our results indicate that when only a small number of (6-4) photoproducts are generated, the UV-DDB-dependent damage recognition pathway predominates over direct recognition by XPC, and they also suggest the presence of rate-limiting regulatory steps in NER prior to the assembly of XPA. © 2009 Elsevier B.V. All rights reserved.

1. Introduction DNA is highly susceptible to damage caused by exposure to agents from both exogenous and endogenous sources. Unrepaired DNA lesions may induce mutations as well as chromosomal aberrations, thereby leading to cellular malfunctioning including cancer, and they may also cause cellular senescence or cell death implicated in damage-induced ageing. Multiple genome maintenance processes that counteract the deleterious effects of DNA lesions have evolved. The heart of this defense system is formed by several DNA repair mechanisms [1]. One of the most versatile DNA repair pathway is nucleotide excision repair (NER), which is able to eliminate a wide variety of lesions that destabilize the DNA double helix, such as ultraviolet light (UV)-induced (6-4) photoproducts (6-4PPs) and cyclobutane pyrimidine dimers (CPDs) [2]. The severe clinical

∗ Corresponding author at: Biosignal Research Center, Organization of Advanced Science and Technology, Kobe University, 1-1 Rokkodai, Nada-ku, Kobe, Hyogo 6578501, Japan. Tel.: +81 78 803 5960; fax: +81 78 803 5970. E-mail address: [email protected] (K. Sugasawa). 1 Present address: Faculty of Science, Gakushuin University, 1-5-1 Mejiro, Toshima-ku, Tokyo 171-8588, Japan. 1568-7864/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.dnarep.2009.02.004

consequence associated with three human rare autosomal recessive diseases, xeroderma pigmentosum (XP), Cockayne syndrome (CS) and trichothiodystrophy (TTD) [3], which are based on inherited NER defects, illustrates the biological relevance of this process. So far, seven NER-deficient genetic complementation groups for XP (XP-A through XP-G), two for CS (CS-A and CS-B) and one for TTD (TTD-A) have been identified, and in all cases the responsible genes have been cloned [4,5]. On the other hand, one XP group, called a variant form of XP (XP-V), is exceptional since mutations confer defects in translesion DNA synthesis but not in NER [6,7]. NER consists of two subpathways: transcription-coupled NER (TC-NER), which removes DNA damage specifically from the transcribed strand of active genes, and global genome NER (GG-NER), which surveys the entire genome for damage. A major difference in the molecular mechanism of these two modes of NER is evident in the damage recognition step. While RNA polymerase II stalling at a damaged site is probably employed as a damage sensor in TC-NER [8], a complex containing the XPC protein, an XP-related gene product, plays an essential role in damage recognition in GG-NER [9–11]. Subsequent steps after damage recognition are thought to be shared by both subpathways: first, transcription factor IIH (TFIIH), which is composed of ten subunits containing two helicase proteins (XPB and XPD), unwinds the DNA duplex around the lesion in the

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presence of XPA, replication protein A and XPG [12–19]. An oligonucleotide of approximately 30 nucleotides including the lesion is then excised by two structure-specific endonucleases, ERCC1-XPF and XPG, which make incisions at sites 5 and 3 to the lesion, respectively [20–23]. The resulting single-stranded gap is filled by DNA polymerase (␦ or ␧) in conjunction with proliferating cell nuclear antigen (PCNA) and replication factor C, and the DNA strands are finally rejoined by DNA ligase I or DNA ligase III/XRCC1 [24–26]. XPC is part of a stable heterotrimeric complex with one of the two mammalian homologs of Saccharomyces cerevisiae Rad23 (RAD23A or RAD23B), which stabilizes and stimulates XPC, and centrin 2, which is also known as a centrosomal protein [9,27–33]. XPC is an essential protein for the initiation of GG-NER and binds various DNA lesions in vitro including UV-induced 6-4PPs [9,34]. Biochemical analyses revealed that XPC binds a certain structure of DNA, which is generated by the presence of lesions, probably enabling the recognition of structurally unrelated DNA lesions [35,36]. On the other hand, UV-damaged DNA binding protein (UV-DDB) also participates in damage recognition in GG-NER. UV-DDB was initially identified as a heterodimer consisting of the DDB1 and DDB2 subunits, the latter of which is implicated in XP group E [37–41]. Purified UV-DDB exhibits a much higher binding affinity and specificity than XPC for both major UV-induced photolesions CPDs and 6-4PPs [9]. Although XP-E cells show obvious defects in the GG-NER of CPDs, the same cells can remove 6-4PPs efficiently from genomic DNA, indicating that UV-DDB in vivo would be important particularly for the recognition of CPDs [42,43]. UV-DDB can be recruited to UV-damaged site even in the absence of XPC and it promotes recruitment of XPC [44–48]. Although UV-DDB is dispensable for cell-free NER reaction [24,49], in vitro reconstituted NER reaction was stimulated by the addition of UV-DDB under certain conditions [47,50,51]. Furthermore, UV-DDB physically interacts with XPC and the associated E3 ligase containing cullin 4A and Roc1 ubiquitylates XPC according with UV irradiation [52]. These findings suggest that UV-DDB is important for damage sensing in vivo. Thus far, in vitro NER reactions have been successfully reconstituted with defined damaged DNA substrates and a set of highly purified proteins [24,49,50]. These studies have provided detailed insights into the reaction mechanism of GG-NER. However, it remains to be determined how the initial step of GG-NER, damage sensing, is regulated in time and space within a living mammalian cell nucleus. Since the initial steps of complex biological processes would be important for its regulation, it is expected that GG-NER may be regulated by initiating factors such as XPC and UV-DDB. To understand the molecular mechanism regulating GG-NER in vivo, we previously assessed the mobility and reaction kinetics of several GFP-tagged NER proteins in living cells by the fluorescence recovery after photobleaching (FRAP) technique, both in the presence and absence of UV irradiation [53–58]. However, how UVDDB-dependent damage recognition of XPC is coordinated within GG-NER and how UV-DDB participates in the further assembly of other NER factors into functional NER complexes remain elusive. To address these key issues, in the present study, we analyzed in further detail the in vivo dynamics of the GFP-tagged XPC protein. Our results not only provide novel insights into the concerted actions of XPC and UV-DDB in damage recognition, but also shed light on some previously uncovered aspects of regulation, including those governing the later step of NER in living cells.

XPCS2BASV (XPB-deficient), and XP4PASV (XPC-deficient), as well as stable transfectants were cultured at 37 ◦ C in an atmosphere of 5% CO2 with Dulbecco’s modified Eagle’s medium containing 10% fetal bovine serum (FBS). 2.2. Establishment of cell lines Two stably transformed cell lines, XP2OSSV cells expressing GFP-XPA and XPCS2BASV cells expressing XPB-GFP, were established as previously described [57,59]. The cDNA encoding GFP-XPC was cloned into the vector pIREShyg (Clontech), from which the GFP-XPC and hygromycin B resistance genes were transcribed as a single mRNA. Twenty micrograms of the construct was linearized and electroporated into a 100-mm dish of XP4PASV cells using Gene Pulser II (Bio-Rad). Stable transformants were selected initially with 200 ␮g/ml hygromycin B (Invitrogen), which gave rise only to clones overexpressing GFP-XPC. The concentration of hygromycin B in the culture medium was then gradually reduced to lower the selective pressure, which resulted in a natural drop of protein levels, thereby allowing isolation of a clone expressing GFP-XPC at nearly physiological levels. 2.3. Measurement of the repair rates of UV photolesions in vivo To avoid dilution of lesions by DNA replication, cells (in 100mm dishes) were treated for 2 h with 6 mM thymidine before each experiment. The cells were then irradiated with 10 or 40 J/m2 of UVC (under a germicidal lamp with a peak at 254 nm) and further cultured for various time periods in the presence of 6 mM thymidine. Genomic DNA was purified with the QIAamp Blood mini kit (Qiagen), and the levels of remaining 6-4PPs and CPDs were measured by an enzyme-linked immunosorbent assay (ELISA) using the lesion-specific monoclonal antibodies 64M-2 and TDM-2, respectively [60]. 2.4. Immunohistochemistry

2. Materials and methods

Prior to immunostaining, 4 × 105 WI38 VA13 cells were cultured overnight in the presence of 0.01% (w/v) polystyrene microsphere beads (Polysciences) in a 25-cm2 flask. After unincorporated beads were washed out with culture medium, the cells were mixed with an equal number of XP4PASV transformant cells expressing GFPXPC and seeded in 35-mm glass-bottom dishes (MatTech) at a density of 1.5 × 105 cells/dish for immunohistochemistry. The cells were fixed with 1.6% (v/v) formaldehyde (Wako Pure Chemicals) for 15 min at 4 ◦ C. The dishes were washed twice with ice-cold phosphate-buffered saline (PBS) and subsequently permeabilized with 0.5% (v/v) Triton X-100 in PBS for 10 min on ice. The cells were again washed twice with ice-cold PBS and then incubated with 3% (v/v) FBS in PBS to block non-specific antibody adsorption. During the following procedures, the dishes were washed three times with PBS after each incubation. The cells were incubated at room temperature for 1 h with an anti-XPC (FL) antibody and then for 1 h with an anti-rabbit IgG antibody conjugated with Alexa Fluor 594 (Molecular Probes, 1:500 dilution). Both antibodies were diluted with PBS containing 0.05% (v/v) Tween 20 and 0.5% (v/v) FBS. Fluorescence microscopy was performed using an Olympus IX71 instrument and Metamorph software (Mitani). For the analysis of localization of GFP-XPC, living cells were treated at 37 ◦ C for 10 min with 10 ␮g/ml of Hoechst 33342 and analyzed with the same microscope system.

2.1. Cell culture

2.5. Preparation of cell lysates

Simian virus 40-transformed human fibroblasts from a normal individual (WI38 VA13) or XP patients, XP2OSSV (XPA-deficient),

For immunoblot analysis of the expression level of XPC or DDB2, cells in 60-mm dishes were washed twice with ice-cold PBS and

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lysed on ice for 1 h with 500 ␮l CSK buffer (10 mM Pipes [pH 6.8], 3 mM MgCl2 , 1 mM EGTA, 0.1% [v/v] Triton X-100, 10% [v/v] glycerol, 0.25 mM phenylmethylsulfonyl fluoride, 1 mM dithiothreitol, 1× protease inhibitor cocktail [Complete, Roche Diagnostics]) containing 0.3 M NaCl. After the cell lysates were scraped into Eppendorf tubes, the dishes were washed with 500 ␮l of the same buffer, which was then combined with the recovered lysates. Soluble protein fractions were obtained by centrifugation for 10 min at 20,000 × g, while the resulting precipitates were washed twice with the same buffer and resuspended in 500 ␮l of the same buffer with the aid of sonication. 2.6. Confocal microscopy Two days prior to each experiment, cells were seeded onto 24mm coverslips. Imaging and FRAP analysis were performed on an LSM 510 META confocal laser scanning microscope (Carl Zeiss). A strip of area spanning the nucleus was photobleached for 100 ms at 100% of laser intensity (laser current set at 6.1 A). Recovery of the fluorescence intensity in the bleached area was monitored every 100 ms over 29 sec at 0.5% laser intensity. At least 20 cells were analyzed in each experiment. Relative fluorescence intensity (defined as the ratio of fluorescence before and after bleaching) was plotted as a function of time. To examine the kinetics of the XPC accumulation into the locally UVC-damaged area, an LSM 510 system equipped with UVC laser emitting at 266 nm was used (Rapp OptoElectronic, Hamburg GmbH) [61]. After local UVC irradiation, live cell images were monitored every 10 s over 390 s. For local UVC irradiation experiments, cells were grown on 25-mm quartz coverslips (010191T-AB, SPI Supplies). 2.7. Transient expression of mCherry-mDDB2 Cells were transiently transfected with a construct for expression of mouse DDB2 fused to monomeric Cherry (mCherry-mDDB2) using Lipofectamine 2000 (Invitrogen). At 12 h post-transfection, cells expressing mCherry-mDDB2 were identified by red fluorescence and subjected to FRAP analysis. 2.8. Knockdown of DDB2 with siRNA Stably transformed XP4PASV cells expressing GFP-XPC were seeded on 25-mm coverslips and transfected with 40 nM siRNA against DDB2 using Lipofectamine 2000. The siRNA, which was labeled with Alexa Fluor 555 at the 3 end of the sense strand, was purchased from Qiagen (Hs DDB2-1: target sequence 5 CCCAGAUCCUAAUUUCAAA-3 ). The corresponding control RNA (5 -CCCAGAUCCUACCUUCAAA-3 ) was also purchased from Qiagen. The cells were cultured for 3 days and then subjected to further experiments. 2.9. Antibodies and other methods Anti-XPC (FL) and anti-RAD23B antibodies were obtained as previously described [32]. The anti-serum raised against purified mouse XPA was kindly provided by Andre P.M. Eker (Erasmus Medical Center, Rotterdam). The anti-DDB2 monoclonal antibody was established by the MAB Institute Co. Ltd. (Yokosuka, Japan). For immunoblotting, proteins were separated by SDS-PAGE and transferred onto polyvinylidene difluoride membranes (Immobilon-P, Millipore). Except for DDB2, detection was performed by chemiluminescence using an anti-rabbit IgG antibody conjugated to alkaline phosphatase (Sigma) and CDP-Star as a substrate (Roche Diagnostics). For detection of DDB2, the horseradish peroxidase-conjugated anti-mouse IgG antibody (R&D Systems)

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and Immobilon Western (Millipore) were employed as a secondary antibody and a substrate, respectively. Quantification was carried out with a luminescent imaging analyzer (LAS-1000 plus, Fujifilm) and accompanying software (ImageGauge v3.45), and blots were also exposed to X-ray films (RX-U, Fujifilm). Concentrations of purified proteins were determined by the Bradford method [62] using the Bradford-ULTRA reagent (Novexin) and bovine serum albumin as a standard. 3. Results 3.1. Generation and characterization of cells expressing GFP-tagged XPC To visualize the DNA damage recognition step of GG-NER in living cells, we generated a cell line that stably expresses N-terminally FLAG- and GFP-tagged XPC (GFP-XPC). Previously, we generated a cell line that expresses a C-terminally tagged XPC [63]. Although this protein is functional in NER, its expression was variable and its stability appeared to be affected by the position of the tag (data not shown). For this reason we generated the alternative N-terminally tagged version. The fusion protein was expressed in the human SV40-immortalized cell line, XP4PASV, which is defective in endogenous XPC. Expression levels of the GFP-XPC protein were examined by immunoblot analysis with an anti-XPC antibody, indicating that GFP-XPC was expressed at a slightly higher level compared to endogenous XPC in a normal human fibroblast cell line WI38 VA13 (Fig. 1A). Immunoprecipitation using an antiFLAG antibody revealed that complex formation of GFP-XPC with both RAD23B and centrin 2 was not affected by the presence of GFP tag (data not shown). To assess GG-NER activity, these cell lines were irradiated with 10 J/m2 UVC and the amounts of 6-4PPs remaining in genomic DNA were measured at various time points by ELISA using a lesion-specific antibody. As shown in Fig. 1B, the transformed cells removed 6-4PPs with kinetics similar to that of the control WI38 VA13 cells. Furthermore, we observed that GFPXPC localized exclusively within the nucleus, like endogenous XPC (Fig. 1C) [64] and C-terminally tagged XPC [63]. Taking these results together, we conclude that the exogenously expressed GFP-XPC is fully functional in NER. Although we showed that the tagged protein is expressed at near physiological levels when measured over the entire population of transformed cells, this analysis does not reveal intercellular variation. Since differences in expression levels may affect the overall mobility of GFP-tagged NER proteins in response to UV irradiation [55,57], we carefully examined protein expression levels of GFP-XPC in individual cells in a large population by immunohistochemistry using the anti-XPC antibody (Fig. 1D and E) and compared this distribution with the expression profile of endogenously produced XPC in normal human fibroblasts. To allow direct comparison, we performed comparative immunofluorescence [65], in which the transformed cells were co-cultured with WI38 VA13 cells labeled with polystyrene beads (Fig. 1D). Within the population of WI38 VA13 cells a rather homogeneous expression level of endogenous XPC was observed, whereas the expression profile of GFP-XPC in transformants appeared much broader, with a considerable proportion of the population expressing protein levels much higher than endogenous levels, as frequently observed for exogenously expressed proteins (Fig. 1E). Although in Fig. 1D the nuclei of cells expressing GFP-XPC look larger than those of control cells, nuclear staining of transformed cells with propidium iodide suggested that the cell line was not polyploid (data not shown). In addition, we also determined the distribution profile of the GFP signal in living cells, which showed an intercellular distribution pattern similar to that determined by immunostaining (data not shown). Hereafter, only cells expressing physiological levels of GFP-XPC (corresponding to

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Fig. 1. Characterization of transformed XP4PASV cells stably expressing GFP-XPC. (A) XPC-deficient cells (XP4PASV) were stably transformed with an GFP-XPC expression construct. Cell lysates were prepared from one of the transformed clones as well as from the parental XP4PASV cells and repair-proficient WI38 VA13 control cells. Six micrograms of soluble proteins (S) and the corresponding pellet fractions (P) were subjected to immunoblotting with the anti-XPC (FL) antibody. RAD23B was detected on the same membrane as a loading control. (B) The indicated cells were irradiated with 10 J/m2 of UVC and allowed to repair induced lesions for various time periods. The 6-4PPs remaining in genomic DNA were quantified and plotted as a function of time. The mean values and standard errors were calculated from at least two independent experiments. (C) Fluorescence microscopic images of transformed XP4PASV cells expressing GFP-XPC, which were stained with Hoechst 33342. The image was taken with 40 times magnification. (D) Co-cultures of WI38 VA13 cells and transformed XP4PASV cells expressing GFP-XPC were subjected to immunohistochemistry using an anti-XPC (FL) antibody. Arrowheads indicate WI38 VA13 cells that had incorporated polystyrene beads. Arrows present GFP-positive stable transformants of XP4PASV cells. These images were taken with 40 times magnification. (E) Expression profiles of endogenous XPC in WI38 VA13 cells (solid bars) and of GFP-XPC in transformed XP4PASV cells (open bars) were obtained based on the intensity of anti-XPC fluorescence signals of the co-cultures as shown in (D). For each expression profile, fluorescence signals of more than 100 cells were measured. Classification of relative expression levels was determined arbitrarily.

arbitrary units 2 and 3 in Fig. 1E) were subjected to an analysis of protein mobility. 3.2. UV-induced alteration of XPC mobility in vivo To further understand the regulation of XPC and to quantitatively determine its reaction kinetics in living cells, we systematically examined the in vivo mobility of GFP-XPC after treatment with various doses of UVC. A significantly slower recovery of GFPXPC fluorescence was observed immediately after treatment with 10 J/m2 UVC (Fig. 2A). As reported before, this UV-dependent transient immobilization of part of the GFP-XPC molecules is found specifically for NER factors [55,57,59], thus likely reflects the binding of XPC to UV-damaged sites and its engagement in repair reactions. In line with this idea, UV-dependent immobilization of XPC was not observed with mutant XPC, which cannot bind to DNA [63]. Notably, the mobility of GFP-XPC returned to the level of that seen for undamaged cells by 5 h after UVC irradiation (Fig. 2A). Within this timeframe the removal of 6-4PPs was almost completed (Fig. 1B), while CPDs were only poorly repaired in the SV40-transformed cell lines, regardless of the presence or absence of functional XPC (Supplementary Fig. S1), in accordance with the previous report [42]. These correlative findings suggest that ongoing GG-NER of 6-4PPs rather than that of CPDs mainly causes the observed decrease in XPC mobility after UVC irradiation. Next, we examined the dynamics of GFP-XPC in more detail using varying doses of UVC (Fig. 2B). Although immobilization of GFP-XPC was apparently more pronounced with increasing UVC, we did not observe a simple proportional relationship between XPC mobility and UVC dose, i.e., the reduction in XPC mobility saturated at relatively low UVC doses, between 5 and 10 J/m2 . Surprisingly,

with higher doses of UVC, further retardation of the fluorescence recovery was observed, which was then saturated again at extremely high doses, around 80–100 J/m2 (Fig. 2B). Although the behavior of GFP-XPC with high dose UVC, such as 100 J/m2 , may not be physiologically relevant, apparent further retardation of the mobility was also observed with 20 J/m2 UVC. Interestingly, even when high dose UVC was applied (40 J/m2 ), reduction in the XPC mobility was partially recovered within 5 h after UVC irradiation (Supplementary Fig. S2A). Within this time frame, about 40% of 6-4PPs were repaired (Supplementary Fig. S2B), whereas CPDs were hardly removed (data not shown) as observed after 10 J/m2 UVC irradiation (Supplementary Fig. 1). These data suggest that the reduction in the XPC mobility caused by high dose UVC irradiation is also attributable mainly to its engagement in GGNER of 6-4PPs rather than that of CPDs. We reproducibly observed this biphasic dose-dependent transient immobilization of GFP-XPC upon UVC treatment of cells. This remarkable biphasic character of XPC entrapment on damaged DNA suggests that the presence of an intricate regulatory mechanism of GG-NER that depends on the number of lesions generated. Even though it is also known that UVC irradiation induces crosslinking between protein and DNA, exposure times used in this study generate negligible crosslinking [66]. Therefore, it is suggested that we observed transient immobilization of GFP-XPC, not covalent binding with DNA. 3.3. UV-DDB regulates the mobility of XPC in UV-irradiated cells To elucidate the molecular basis underlying the biphasic regulation of XPC mobility, we examined the involvement of UV-DDB, which has been implicated in damage recognition in GG-NER [9,46,47,52]. First, FRAP analysis was performed with stably trans-

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Fig. 2. Alteration of XPC mobility in response to UV irradiation. (A) FRAP analysis of GFP-XPC in transformed XP4PASV cells was carried out without UV irradiation (ND), or immediately or 5 h after irradiation with 10 J/m2 of UVC. (B) FRAP analysis of GFP-XPC was performed after treatment of transformed XP4PASV cells with various doses of UVC irradiation.

formed cells expressing GFP-XPC and transiently transfected with mouse DDB2 fused to mCherry at the N-terminus, a derivative of the auto-fluorescent protein DsRed (mCherry-mDDB2) (Fig. 3) [67]. Since it is reported that the DDB1 level exceeds that of DDB2 in vivo [68,69], overexpression of mCherry-mDDB2 in addition to endogenous DDB2 was expected to increase the total number of functional UV-DDB complexes in GFP-XPC cells. Cells expressing mCherry-mDDB2 could be easily distinguished by the presence of red fluorescence (Fig. 3A). As shown in Fig. 3B, expression of mCherry-mDDB2 hardly affected the mobility/immobile fraction of GFP-XPC after UVC irradiation at 5 J/m2 . In striking contrast, however, a dramatic further retardation of the fluorescence recovery was observed in cells expressing mCherry-mDDB2 after 10 J/m2 UVC. These data suggest that the DNA damage-induced transient immobilization of GFP-XPC is in part dependent on the amount of DDB2, and the secondary transient immobilization might be explained by direct binding of XPC to lesions, independent of UV-

DDB, which only becomes significant after a certain threshold of lesion density. To further analyze the molecular mechanism regulating damage recognition by UV-DDB and XPC, we performed the reciprocal experiment, in which expression of endogenous DDB2 was reduced by siRNA. Transfection with specific siRNAs against human DDB2 efficiently suppressed expression of endogenous DDB2 (Fig. 4A). Since the siRNAs were directly labeled with a fluorescent dye (Alexa Fluor 555), only transfected cells with red fluorescence could be identified and chosen for the following FRAP analysis (Fig. 4B). Knockdown of DDB2 per se had little effect on the mobility of GFPXPC in the absence of UV damage (Fig. 4C). Reducing the DDB2 concentration by siRNA concomitantly reduced the immobilization of XPC after 5 or 10 J/m2 UVC (Fig. 4D). Taken together, these results suggest that the reduction in in vivo mobility of XPC, particularly the first step of immobilization caused by relatively low UVC doses, is titrated by the number of functional UV-DDB complexes. There are at least two possible explanations for the observed effects of UV-DDB on the mobility of XPC in UV-irradiated cells: UV-DDB (i) promotes recruitment of XPC to UV-induced lesions, or (ii) prevents (or delays) its release from sites where GG-NER occurs. To examine these possibilities, we first determined the assembly kinetics of GFP-XPC in the presence and absence of DDB2 at pre-steady states (Kon > Koff ) by directly measuring the rate of accumulation on locally UVC-damaged sites, using a confocal microscope equipped with a UVC laser (Fig. 4E) [61]. A short pulse of UVC that generates both 6-4PPs and CPDs [61] in a defined part of the nucleus caused a rapid accumulation of GFP-XPC in the irradiated area, which reached a maximum of approximately 2.5 times the level before irradiation within 400 s (Fig. 4E). Notably, when DDB2 was knocked down, a significantly less pronounced accumulation of GFP-XPC was observed, as compared to control conditions. Although we cannot completely exclude the possibility that the release of XPC is affected by the knockdown of DDB2 in this assay, the initial increase in the accumulating GFP fluorescence was reduced in DDB2-knockdown cells. These results strongly support the idea that UV-DDB facilitates the recruitment of XPC to UV-damaged sites in vivo at relatively low UV exposure.

3.4. Differential regulation of the in vivo dynamics of TFIIH and XPA in UV-irradiated cells Fig. 3. Effect of ectopic expression of mCherry-mDDB2 on XPC UV-dependent immobilization of XPC. (A) Confocal microscopic images of transformed XP4PASV cells stably expressing GFP-XPC, which were transiently transfected with a mCherrymDDB2 expression construct. Arrowhead and arrow indicate transfected and untransfected cells, respectively. (B) FRAP analysis of GFP-XPC in transformed XP4PASV cells with (+DDB2) or without ectopic expression of mCherry-mDDB2. The cells were pretreated with 5 or 10 J/m2 of UVC as indicated.

The above data suggest that UV-DDB is involved in the repair of not only CPDs but also 6-4PPs. Consistent with this notion, recent evidence indicated that DDB2 also stimulates 6-4PP removal, but only at relatively low doses of UVC [46]. To determine whether overexpression of DDB2 can further stimulate GG-NER or NER complex assembly, we analyzed the live cell dynamic properties of two other

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Fig. 4. Reduced expression of endogenous DDB2 affects XPC mobility in UV-irradiated cells. (A) Transformed XP4PASV cells expressing GFP-XPC were mock-treated or transfected with either siRNA against DDB2 or the corresponding control RNA which carries two base substitutions. Cell lysates were prepared 3 days after transfection, and 10 ␮g of soluble proteins were subjected to immunoblot analysis using an anti-DDB2 antibody. RAD23B was detected as a loading control. (B) Confocal microscopic image of transformed XP4PASV cells expressing GFP-XPC (green), which were transfected with Alexa Fluor 555-labeled siRNA against DDB2 (red signal). The image was taken 3 days after transfection. (C and D) Transformed XP4PASV cells expressing GFP-XPC were treated either with an siRNA against DDB2 (DDB2 KD) or the control RNA. FRAP analysis of these cells was carried out without UVC irradiation (C), or after UVC irradiation at 5 or 10 J/m2 (D). (E) A pulse of the UVC laser was applied to a point within the nucleus of cells expressing GFP-XPC, which had been treated with either the DDB2 siRNA (DDB2 KD) or the corresponding negative control RNA. The relative fluorescence intensity in the irradiated area (the value before irradiation was set as 1) was monitored and plotted as a function of time.

NER factors, TFIIH and XPA, both of which are involved in steps following initial damage recognition. To examine the mobility of TFIIH, we used XP-B fibroblasts (derived from patient XPCS2BA) that stably express physiological levels of biologically active XPB C-terminally tagged with GFP (XPB-GFP) [59]. FRAP analysis of XPB-GFP after increasing doses of UVC showed that it exhibited a dose-dependent behavior similar to that of GFP-XPC: XPB-GFP was rapidly immobilized after UV irradiation with an initial saturation between 5 and 10 J/m2 , and there was a secondary dose-dependent

immobilization at higher doses of UVC (Fig. 5A). In the presence of ectopically expressed mCherry-mDDB2, the mobility of XPB-GFP appeared to be slightly reduced even in unirradiated cells, suggesting that DDB2 might be involved in other cellular functions including transcription. Nevertheless, immobilization of XPB-GFP was much more pronounced in cells irradiated with 10 J/m2 UVC with overexpressed mCherry-mDDB2 (Fig. 5B). Finally, the in vivo mobility of XPA was analyzed by using an XPA-deficient cell line (XP2OSSV) expressing GFP-XPA [57]. Intrigu-

Fig. 5. Effect of UVC irradiation and ectopic DDB2 expression on the mobility of XPB. (A) Transformed XPCS2BASV cells expressing XPB-GFP were treated with various doses of UVC as indicated and subjected to FRAP analysis. (B) Cells expressing XPB-GFP were transiently transfected with a mCherry-mDDB2 expression construct (+DDB2) or mock-transfected. The cells were subjected to FRAP analysis before and after UVC irradiation at 10 J/m2 .

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Fig. 6. Effects of UVC irradiation and ectopic DDB2 expression on the mobility of XPA. (A) Transformed XP2OSSV cells expressing GFP-XPA were treated with various doses of UVC as indicated and subjected to FRAP analysis. (B) Cells expressing GFP-XPA were transiently transfected with a mCherry-mDDB2 expression construct (+DDB2) or mock-transfected. The cells were subjected to FRAP analysis before and after UVC irradiation at 10 J/m2 .

ingly, FRAP analysis of these cells irradiated with various doses of UVC revealed that GFP-XPA exhibited a kinetic behavior completely different from that of XPC and XPB: the mobility of GFP-XPA was reduced by 10 J/m2 UVC, like for the other NER factors, but no further immobilization was observed with higher doses, up to at least 100 J/m2 (Fig. 6A). Furthermore, the ectopic expression of mCherrymDDB2 did not affect the mobility of GFP-XPA either in unirradiated or UV-irradiated cells (Fig. 6B). These results suggest that only a limited amount of XPA may be engaged in NER even though much more lesions were generated and more damage recognition complexes were assembled. 4. Discussion 4.1. UV-DDB-dependent damage recognition is predominant when low doses of UV are applied In the present study, our FRAP analyses revealed that the in vivo mobility of XPC is reduced in UVC-irradiated cells, as previously reported for other NER factors [55,57–59]. This observation most likely means that XPC is trapped at lesion sites and engaged in NER. However, XPC also exhibited unique behaviors in that its UV-induced immobilization seems to be saturated at relatively low UVC doses (5 or 10 J/m2 ). This reduction in XPC mobility appears to depend on the presence of 6-4PPs, because it could not be observed at 5 h after UVC irradiation (Fig. 2A), by which time point the GG-NER of 6-4PPs was mostly completed. Since repair of CPDs appeared quite inefficient in the SV40-transformed cell lines used (Supplementary Fig. 1) [42], the extent of ongoing CPD repair, if any, was probably too small to be detected by measuring the overall mobility of XPC. Based on several data, this first-step immobilization of XPC can be attributed to the UV-DDB-mediated pathway of GG-NER damage recognition. First, siRNA knockdown of DDB2 largely restored the mobility of XPC up to the level observed for unirradiated cells (Fig. 4). With UVC doses at 40 J/m2 or higher, a dose-dependent reduction in XPC mobility was still observed in DDB2-knockdown cells (data not shown), strongly suggesting that the second-step immobilization is due to direct damage binding of XPC itself. Second, ectopic expression of DDB2 substantially potentiated the XPC immobilization caused by 10 J/m2 of UVC irradiation, while it hardly affected XPC mobility after 5 J/m2 of UVC (Fig. 3). We deduce that this difference observed between the two UVC doses could be explained by the relative numbers of UV-DDB molecules and generated lesions, particularly 6-4PPs. It has been calculated that 1.5 × 105 and 3 × 105 6-4PPs are generated in a human diploid cell after UVC irradiation at 5 and 10 J/m2 , respectively [70]. By quantitative immunoblot analysis using the purified UV-DDB complex as a

standard, we estimated the number of endogenous DDB2 molecules in our GFP-XPC-expressing cell line as approximately 1 × 105 per cell (data not shown). Given that UV-DDB can recognize and bind to 6-4PPs in a nearly quantitative manner in vivo, endogenous UVDDB would be sufficient to cover most of the 6-4PPs generated by 5 J/m2 UVC. This would imply that one UV-DDB complex would only be able to bind to one 6-4PP, and then cannot be reused for a subsequent lesion, consistent with the reported degradation of UV-DDB after damage detection [41,71,72]. On the other hand, UVDDB may be limiting when the UVC dose is increased up to 10 J/m2 , so that the expression of additional DDB2 would help in trapping more XPC. Direct damage recognition by XPC may become more pronounced when higher doses of UV are applied. Notably, in DDB2 knockdown cells, the immobilization of XPC does not appear pronounced with such low UVC doses (Fig. 4). The presence of residual DDB2 in the knockdown cells (Fig. 4A) suggests that the UV-DDBdependent pathway is predominant in the GG-NER of 6-4PPs, when the number of lesions is low. In striking contrast to what is known for CPDs, the stimulatory role of UV-DDB in the GG-NER of 6-4PP has been under debate since human XP-E cells and Chinese hamster cells, which both lack functional DDB2, can apparently remove 6-4PPs efficiently from the global genome and display considerable levels of residual UV-induced unscheduled DNA synthesis [3]. It was recently reported that UV-DDB can promote NER, particularly when the number of lesions generated is relatively small [46]. Our results are consistent with this notion, and the apparent contribution of the UV-DDB-dependent pathway to 6-4PP repair may vary depending not only on UV doses but also on the cell types examined, since the expression level of DDB2 may be substantially affected by multiple factors, such as status of the p53 tumor suppressor [42]. The kinetic analysis of XPC accumulation in response to local UVC damage indicates that UV-DDB facilitates the recruitment of XPC to lesion sites, since the initial increase in fluorescence intensity was greatly reduced by knockdown of DDB2 (Fig. 4E). However, the absolute levels of accumulating GFP signals also seem different, and it is possible that UV-DDB may also affect the release of XPC from damaged sites. It was recently shown that DDB2 and XPC are ubiquitylated in UVC-irradiated cells [52], which we propose may be important for the efficient transfer of lesions between the two damage recognition factors. The presence of these extra steps may take some time, thereby retaining XPC at specific lesions. In addition, since ubiquitylated XPC seems to exhibit moderately higher affinity for 6-4PP compared to unmodified XPC, modification of XPC itself might affect its retention time. On the other hand, it has been recently shown that in vivo dynamics of YFP-tagged DDB2 appeared largely independent of XPC [56]. This is not inconsistent with our data, since it was shown that UV-DDB can bind to UV-damaged sites in the absence of XPC and that the number of DDB2 molecules

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Fig. 7. Schematic representation of damage recognition involving UV-DDB and XPC complex under various conditions. (A, B and C) Recognition of 6-4PP with 5, 10 J/m2 or relatively high dose of UV, respectively. (D) Recognition of 6-4PP with ectopically overexpressed DDB2. N6-4PP and NUV-DDB mean the numbers of generated 6-4PP and UV-DDB molecules, respectively. See text for detail.

in a cell exceeds that of XPC [12,69] (it should be noted that the YFP-tagged DDB2 was additionally expressed in the presence of endogenous DDB2). Based on above data, we deduce in vivo damage recognition in GG-NER (Fig. 7). When the amount of DNA damage was lower than molecular number of UV-DDB, damage recognition would predominantly be accomplished by UV-DDB and XPC complex (Fig. 7A). A certain fraction of damage may also be recognized directly by XPC complex if the damage amount greatly exceeds the number of UVDDB (Fig. 7B and C), while ectopically expressed UV-DDB helps the recruitment of XPC to damaged sites (Fig. 7D).

4.2. The regulatory mechanism of NER in living cells To obtain further insights into the overall regulatory mechanism of NER, we also analyzed the dynamics of NER factors involved in later stages of NER. Notably, we found that the in vivo mobility of XPB-GFP, a subunit of TFIIH, seems to be regulated by UVC irradiation as well as by UV-DDB in a fashion similar to that observed with XPC (Fig. 5). These results suggest that the overall mobility of TFIIH in UVC-irradiated cells may be regulated directly by XPC, probably through their reported physical interaction [12,19,73]. Furthermore, ectopically expressed DDB2 also enhanced the immobilization of TFIIH, especially upon UVC irradiation, suggesting that the recruitment of TFIIH is dependent on the formation of the damage recognition complex. Although TFIIH is involved not only in GG-NER but also in transcription and TC-NER, our observation is not surprising because TFIIH appears to have higher affinity for NER sites than for transcription sites [59], and the number of ongoing TC-NER events must be lower than the number of GG-NER events. Compared to XPC and XPB, GFP-XPA showed different behavior (Fig. 6). FRAP analysis indicates that the percentage of XPA molecules involved in NER does not increase if UVC dose is 10 J/m2

or higher, suggesting that this cell line has a similar saturating UVdamage level for repair in the range of 10 J/m2 as for the other cell lines. This is not because XPA is present in a large excess over XPC and TFIIH, judging from the estimated numbers of XPC, TFIIH, and XPA molecules per cell [12,64,74]. Thus, although both XPC and TFIIH are present at a lesion site, participation of XPA appears to be restricted, especially with high dose UVC. A possible explanation is that there could be a rate-limiting factor, such as XPG, which may be involved in an earlier step of NER than XPA as determined in vivo [11]. It has been reported that XPG interacts strongly with TFIIH and enhances the unwinding of duplex DNA around lesions [12,14,15,75]. However, in vitro experiments concerning the assembly order of NER factors have suggested that XPG may function after XPA [76]. Another possibility is that XPA might be present in two or more different states (possibly determined by a post-translational modification), and only a particular portion may be available for NER, as previously suggested [77]. In this regard, it is notable that XPA has been reported to be related to the ATR-mediated checkpoint pathway [78]. When cells are exposed to UVC above a certain dose, checkpoint signaling may be triggered and prevent XPA from participating in NER. Alternatively, under high UV doses, which cells cannot handle, there would be abortive NER that is triggered by XPC followed by TFIIH but then abrogated as the cells are on the way to undergo cell death in the presence of very high levels of damage. It may be beneficial to trigger cell death instead of trying to repair as survival of the heavily damaged cell might be dangerous for leading to cancer. It is also possible that instead of XPA, some other factor may interact with XPC and/or TFIIH at lesion sites, which may further modulate damage signaling (e.g., by regulating checkpoint controls and/or apoptosis). To understand the overall regulatory mechanism of NER and signal transduction related to NER, further studies could use the chromatin immunoprecipitation technique to identify the protein complexes assembled at lesion sites in vivo.

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Conflict of interest None. Acknowledgments We thank all lab members in the Biosignal Research Center, Kobe University and at RIKEN for beneficial discussions and encouragement. We also thank all the members in the Erasmus Medical Center, especially S. Bergink, for helpful discussion. This work was supported by Grants-in-Aid from the Ministry of Education, Culture, Sports, Science and Technology of Japan, and by the Solution Oriented Research for Science and Technology (SORST) from the Japan Science and Technology Agency. This work was also financially supported by the Bioarchitect Research Project of RIKEN. R.N. was supported by the RIKEN Special Postdoctoral Researchers Program. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.dnarep.2009.02.004. References [1] J.H.J. Hoeijmakers, Genome maintenance mechanisms for preventing cancer, Nature 411 (2001) 366–374. [2] E.C. Friedberg, G.C. Walker, W. Siede, R.D. Wood, R.A. Schultz, T. Ellenberger, DNA Repair and Mutagenesis, 2nd edn., ASM Press, Washington, D.C., 2006. [3] D. Bootsma, K.H. Kraemer, J.E. Cleaver, J.H.J. Hoeijmakers, Nucleotide excision repair syndromes: xeroderma pigmentosum, Cockayne syndrome, and trichothiodystrophy, in: C.R. Scriver, A. Beaudet, W.S. Sly, D. Valle (Eds.), The Metabolic and Molecular Basis of Inherited Disease, McGraw-Hill Book Co, New York, NY, 2001, pp. 677–703. [4] E.C. Friedberg, How nucleotide excision repair protects against cancer, Nat. Rev. Cancer 1 (2001) 22–33. [5] G. Giglia-Mari, F. Coin, J.A. Ranish, D. Hoogstraten, A. Theil, N. Wijgers, N.G. Jaspers, A. Raams, M. Argentini, P.J. van der Spek, E. Botta, M. Stefanini, J.-M. Egly, R. Aebersold, J.H.J. Hoeijmakers, W. Vermeulen, A new, tenth subunit of TFIIH is responsible for the DNA repair syndrome trichothiodystrophy group A, Nat. Genet. 36 (2004) 714–719. [6] R.E. Johnson, C.M. Kondratick, S. Prakash, L. Prakash, hRAD30 mutations in the variant form of xeroderma pigmentosum, Science 285 (1999) 263–265. [7] C. Masutani, R. Kusumoto, A. Yamada, N. Dohmae, M. Yokoi, M. Yuasa, M. Araki, S. Iwai, K. Takio, F. Hanaoka, The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta, Nature 399 (1999) 700–704. [8] V. van den Boom, E. Citterio, D. Hoogstraten, A. Zotter, J.-M. Egly, W.A. van Cappellen, J.H.J. Hoeijmakers, A.B. Houtsmuller, W. Vermeulen, DNA damage stabilizes interaction of CSB with the transcription elongation machinery, J. Cell Biol. 166 (2004) 27–36. [9] D. Batty, V. Rapic’-Otrin, A.S. Levine, R.D. Wood, Stable binding of human XPC complex to irradiated DNA confers strong discrimination for damaged sites, J. Mol. Biol. 300 (2000) 275–290. [10] K. Sugasawa, J.M.Y. Ng, C. Masutani, S. Iwai, P.J. van der Spek, A.P.M. Eker, F. Hanaoka, D. Bootsma, J.H.J. Hoeijmakers, Xeroderma pigmentosum group C protein complex is the initiator of global genome nucleotide excision repair, Mol. Cell 2 (1998) 223–232. [11] M. Volker, M.J. Mone, P. Karmakar, A. van Hoffen, W. Schul, W. Vermeulen, J.H.J. Hoeijmakers, R. van Driel, A.A. van Zeeland, L.H.F. Mullenders, Sequential assembly of the nucleotide excision repair factors in vivo, Mol. Cell 8 (2001) 213–224. [12] S.J. Araujo, E.A. Nigg, R.D. Wood, Strong functional interactions of TFIIH with XPC and XPG in human DNA nucleotide excision repair, without a preassembled repairosome, Mol. Cell. Biol. 21 (2001) 2281–2291. [13] E. Evans, J. Fellows, A. Coffer, R.D. Wood, Open complex formation around a lesion during nucleotide excision repair provides a structure for cleavage by human XPG protein, EMBO J. 16 (1997) 625–638. [14] E. Evans, J.G. Moggs, J.R. Hwang, J.-M. Egly, R.D. Wood, Mechanism of open complex and dual incision formation by human nucleotide excision repair factors, EMBO J. 16 (1997) 6559–6573. [15] D. Mu, M. Wakasugi, D.S. Hsu, A. Sancar, Characterization of reaction intermediates of human excision repair nuclease, J. Biol. Chem. 272 (1997) 28971–28979. [16] L. Schaeffer, V. Moncollin, R. Roy, A. Staub, M. Mezzina, A. Sarasin, G. Weeda, J.H.J. Hoeijmakers, J.-M. Egly, The ERCC2/DNA repair protein is associated with the class II BTF2/TFIIH transcription factor, EMBO J. 13 (1994) 2388–2392. [17] L. Schaeffer, R. Roy, S. Humbert, V. Moncollin, W. Vermeulen, J.H.J. Hoeijmakers, P. Chambon, J.-M. Egly, DNA repair helicase: a component of BTF2 (TFIIH) basic transcription factor, Science 260 (1993) 58–63.

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