| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | ADVANCED SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 Department of Pathology, Nagoya University Graduate School of Medicine, 65 Tsurumai-cho, Showa-ku, Nagoya 466-8550, Japan
2 Division of Molecular Pathology, Center for Neurological Disease and Cancer, Nagoya University Graduate School of Medicine, 65 Tsurumai-cho, Showa-ku, Nagoya 466-8550, Japan
 |
Abstract |
|---|
 Top Abstract IntroductionResultsDiscussionExperimental proceduresReferences |
|---|
, Pol 
, Pol 
, and REV1) has been shown to carry out TLS by itself or in combination with another enzyme in vitro. Recently, the C-terminal region of mammalian REV1 (the total 1251 residues in human) was found to interact with Pol , Pol , and Pol , as well as with the REV7 subunit of another TLS enzyme, Pol 
. Thus, it is proposed that REV1 plays a pivotal role in TLS in vivo. We here describe our study on the localization of human REV1 protein (hREV1) in nondamaged and ultraviolet (UV)-irradiated cells. Ectopically expressed hREV1 in mammalian cells was localized to the nucleus and exhibited dozens of tiny foci in approximately 3% of nondamaged cells. The percentage of focus-forming cells markedly increased after UV irradiation in a time- and dose-dependent manner. The focus formation was associated with UV-induced DNA damage. Interestingly, although the hREV1 foci in S-phase cells colocalized with PCNA foci, suggesting the association of hREV1 with the replication machinery, hREV1 focus formation was observed not only in the S phase but also outside S phase. Furthermore, it was found that the hREV1 focus formation after UV irradiation required a region near the C-terminal (826–1178). |
Introduction |
|---|
|
TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
|---|
and Pol 
. To circumvent this situation, cells have low-fidelity DNA polymerases specialized for replicating past DNA lesions. Such low-fidelity polymerases are able to carry out translesion DNA synthesis (TLS), in which a nucleotide is inserted opposite a lesion and extended further before the major replicative DNA polymerase resumes DNA synthesis. However, the low fidelity of TLS enzymes affects the accuracy of the synthesized DNA, resulting in the introduction of mutations (Kunz et al. 2000; Prakash & Prakash 2002; Friedberg et al. 2004).
In the budding yeast, Saccharomyces cerevisiae, most of the UV-induced mutations are generated during TLS, involving Rev1 and Pol (Lawrence & Maher 2001). Rev1 has a catalytic domain as deoxycytidyl transferase in the central region, which contains multiple motif sequences conserved among the Y-family DNA polymerases (Nelson et al. 1996b; Ohmori et al. 2001; Haracska et al. 2002). Rev1 incorporates dCMPs opposite an abasic site, an undamaged guanine, and some of modified guanines, but not opposite a CPD or a (6-4) photoproduct in vitro. Pol is a complex of Rev3 and Rev7, the former being a catalytic protein with a polymerase domain similar to Pol and the latter being an accessory protein of unknown function (Nelson et al. 1996a). The major function of Pol in TLS is assumed to be extending primer ends paired at various DNA lesions (Lawrence 2002). UV-induced mutagenesis by Pol is defective in the rev1-1 mutant carrying an amino acid substitution in the BRCT domain near the N-terminal, indicating that Rev1 plays a second, noncatalytic function that is necessary for Pol to carry out TLS in vivo (Nelson et al. 2000; Rajpal et al. 2000). Human REV1, REV3, and REV7 homologs were recently identified, and their biologic properties are considered to be very similar to those of their yeast counterparts (Gibbs et al. 1998, 2000; Xiao et al. 1998; Lin et al. 1999; Murakumo et al. 2000). Human cells expressing an hREV1 antisense RNA or a ribozyme against hREV1 mRNA had a lower UV-induced mutation frequency when compared to control cells (Gibbs et al. 2000; Clark et al. 2003). hREV3 and hREV7 together form human Pol , and hREV3 anti-sense RNA expression in human cells also reduced UV-induced mutation frequency (Gibbs et al. 1998; Murakumo et al. 2000). hREV7 was found to form a complex with hREV1 as well as with hREV3, but not with both hREV1 and hREV3 at the same time (Murakumo et al. 2001; Masuda et al. 2003). Furthermore, significant similarity of hREV7 to MAD2 implies that it has some roles in cell cycle checkpoint mechanism (Chen & Fang 2001; Pfleger et al. 2001; Murakumo 2002); however, the biologic function of hREV7 remains unclear. Biochemical studies on human Pol have been hampered because of the difficulty in producing recombinant hREV3 protein.
Human cells have four Y-family enzymes (Pol , Pol , Pol , and REV1), each of which has a different specificity for DNA lesion to be bypassed. Pol , a human homolog of S. cerevisiae Rad30, can bypass a thymine–thymine (T-T) CPD efficiently and accurately, inserting A-A opposite the lesion, and its loss of function increases the risk of a cancer-prone syndrome, the variant form of xeroderma pigmentosum (Johnson et al. 1999; Masutani et al. 1999). Pol , which is the second human homolog of the S. cerevisiae Rad30, is an extraordinarily error-prone polymerase on undamaged templates, inserting G more frequently than A opposite T; however, it can insert an A opposite the 3'-T of a T-T (6-4) photoproduct while not extending further (Tissier et al. 2000a, 2000b). Pol is able to bypass a benzo[a]pyrene-adducted guanine, whereas it cannot bypass either a T-T CPD or a (6-4) photoproduct (Rechkoblit et al. 2002; Suzuki et al. 2002; Zhang et al. 2002). In some cases, a lesion may be bypassed by the combined action of two polymerases. For example, an abasic site can be bypassed by yeast Rev1 and Pol , with the former inserting a dCMP opposite the lesion and the latter extending the replication a short distance beyond the inserted nucleotide (Nelson et al. 1996b). A (6-4) T-T photoproduct, a highly mutagenic lesion resulting from its large structural distortion, may be bypassed by the sequential action of Pol and Pol (Johnson et al. 2000). To accomplish such TLS actions efficiently, those enzymes might interact with each other in vivo. The direct interaction between Pol and Pol has been noted (Kannouche et al. 2002). Furthermore, recent publications demonstrate that hREV1 interacts with Pol , Pol , and Pol at the C-terminal region (1130–1251) as it does with hREV7 (Murakumo et al. 2001; Guo et al. 2003; Ohashi et al. 2004). These results suggest that hREV1 plays an important role for TLS events in vivo.
Here we report the analyses of hREV1 localization in mammalian cells. Green fluorescent protein (GST)-fused or FLAG-tagged hREV1 was localized to the nucleus showing dozens of tiny, sharp foci in about 3% of hREV1-expressing cells. The population of the hREV1 focus-forming cells dramatically increased after UV irradiation. Analysis by localized UV irradiation through a micropore membrane filter revealed that hREV1 accumulated at regions of UV-induced DNA damage. About half of the hREV1 focus-forming cells after UV irradiation were in S phase, and the hREV1 foci in S-phase cells co-localized with proliferating cell nuclear antigen (PCNA) foci, suggesting the association of hREV1 with the replication machinery. Interestingly, in cell cycle analysis, it was revealed that most of GFP-hREV1 focus-forming cells after UV irradiation were distributed in G1 and early S phase. Furthermore, the hREV1 focus formation required a region (826–1178) near the C-terminal. These results indicate that hREV1 re-localizes in response to UV-induced DNA damage in a cell cycle-dependent manner and that the C-terminal region of hREV1, which is not conserved between yeast and human, may play an important role in hREV1 function in vivo.
|
Results |
|---|
|
TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
|---|
To investigate biologic properties of hREV1 in cells, we visualized the hREV1 molecules by tagging hREV1 protein with GFP. The expression vector pEGFP/hREV1 was constructed by inserting the full-length hREV1 cDNA into the pEGFP-C3 for producing the GFP-fusion hREV1 protein (GFP-hREV1) in cells. COS-7 or MRC-5 cells were transfected with the pEGFP/hREV1, and the cells were fixed 48 h after transfection. Localization of the GFP-hREV1 protein was analyzed by fluorescence microscopy. In most cells, GFP-hREV1 protein was diffusely distributed in the nucleus (Fig. 1Aa), and in some cells, GFP-hREV1 formed dozens of small and sharp foci, which were evenly distributed in the nucleus (Fig. 1Ad). This phenomenon was also observed in living cells (Fig. 1Ac,f). As a control, localization of GFP protein when the empty vector pEGFP-C3 was transfected was analyzed. Diffuse distribution of GFP protein in both the nucleus and the cytoplasm was observed; none of the cells formed foci (data not shown). In order to further confirm that hREV1 formed foci, we expressed hREV1 protein with an N-terminal FLAG tag (FLAG-hREV1) in COS-7 cells by transfecting the expression vector pcDNA/FLAG-hREV1. We were able to confirm by immunofluorescence microscopy that FLAG-hREV1 also formed foci in a manner similar to GFP-hREV1 (Fig. 1Ab,e). This phenomenon was observed not only in cells with a high level of the hREV1 expression but also in cells with a comparatively low level of the hREV1 expression evaluated grossly by fluorescence microscope (Fig. 1B). Therefore, it seems unlikely that the focus formation of hREV1 is an artifact by protein over-expression in cells.
|
Because S. cerevisiae REV1 is known to be involved in UV-induced mutagenesis, we examined whether or not hREV1 localization changes in response to UV irradiation. COS-7 or MRC-5 cells transfected with pEGFP/hREV1 were UV-irradiated at 8 J/m2 48 h after transfection, fixed 8 h after UV irradiation and analyzed for the localization of GFP-hREV1 protein. The GFP-hREV1 protein was localized to the nucleus, as seen in the non-UV-irradiated cells, and interestingly, the population of the GFP-hREV1 focus-forming cells significantly increased compared to non-UV-irradiated cells. In most focus-forming cells, hundreds of sharp and tiny foci evenly distributed in the nucleus were observed (Fig. 1Ag). The same results were obtained when the FLAG-hREV1 protein transiently expressed in cells was immunocytochemically detected after UV irradiation, or when the GFP-hREV1 protein was observed in living cells after UV irradiation (Fig. 1Ah,i). These results indicated that many hREV1 foci were newly formed in response to UV irradiation. It should be noted that some cells had large or unevenly distributed spots. In addition, some cells contained only a few foci. In light of this, we defined ‘nuclear foci’ as sharp and tiny speckles that were evenly distributed in the nucleus, as shown in Fig. 1. We next calculated the focus formation rates in GFP-hREV1-expressing cells. About 3% of the GFP-hREV1-expressing cells showed nuclear focus formation without UV irradiation, whereas about 25% of the cells with GFP-hREV1 expression showed nuclear focus formation after UV irradiation. To analyze the dynamics of the nuclear focus formation, COS-7 or MRC-5 cells expressing GFP-hREV1 were UV-irradiated at 8 J/m2 and fixed at various time points postirradiation for a time course analysis. As shown in Fig. 2A (COS-7) and 2B (MRC-5), the number of focus-positive cells increased in a time-dependent manner with a peak percentage of about 25% at 8 h after UV irradiation. The percentages of GFP-hREV1 focus-forming cells were also analyzed to evaluate UV-dose dependency. COS-7 or MRC-5 cells transiently expressing GFP-hREV1 were UV irradiated at various UV doses and fixed 8 h after UV irradiation. The percentage of GFP-hREV1 focus-forming cells increased in a dose-dependent manner with a peak of about 25% at 12 J/m2 in COS-7 cells (Fig. 2C) and about 30% at 16 J/m2 in MRC-5 cells (Fig. 2D). To eliminate the possibility that focus formation was accelerated as a result of up-regulation of GFP-hREV1 expression induced by UV irradiation, 20 µg/mL of cycloheximide, an inhibitor of protein synthesis, was added in the medium after UV irradiation and analyzed for GFP-hREV1 focus formation. As shown in Fig. 2E, no significant difference was observed in the focus formation between the cells cultured in the medium with and without cycloheximide. These results indicate that GFP-hREV1 focus formation is induced by UV irradiation in a time- and dose-dependent manner via relocalization of the protein.
|
T-T CPD is one of the major DNA lesions generated by UV irradiation. To evaluate the relationship between hREV1 foci and UV-induced DNA damage, we examined the recruitment of GFP-hREV1 to UV-induced DNA lesions represented by T-T CPDs. COS-7 cells expressing GFP-hREV1 were subjected to localized UV irradiation at 80 J/m2 through a micropore membrane filter. Cells were immunocytochemically stained by using TDM-2 monoclonal antibody, which recognizes the T-T CPD DNA lesion. In this experiment, the UV-irradiated areas were demonstrated as the TDM-2-positive areas as shown in Fig. 3B,E, and a large portion of the GFP-hREV1 foci were localized to these areas (Fig. 3A–C,G). As a control, cells expressing GFP alone were also analyzed. No accumulation of GFP protein was observed (Fig. 3D–F). These results demonstrate that hREV1 foci accumulate around regions of UV-induced DNA damage.
|
To investigate the biologic significance of hREV1 foci in cells, we analyzed the association of hREV1 with the replication machinery. First, we checked whether GFP-hREV1 focus-forming cells were in S phase or not. If the hREV1 foci represent the sites of TLS where the replicative DNA polymerases stall, we can expect that hREV1 foci should be observed only in cells actively synthesizing DNA. COS-7 cells expressing GFP-hREV1 were incubated in the medium with 10 µM 5-bromo-2'-deoxy-uridine (BrdU) for 15 min before fixation, and then the cells were immunofluorescently stained with anti-BrdU antibody. In contrast to the previous anticipation, our results showed that only a minor fraction of the GFP-hREV1 focus-forming cells showed BrdU uptake. Such as cells presented in Fig. 4A, some of the GFP-hREV1 focus-forming cells indeed showed BrdU uptake; however, others did not show any BrdU uptake at all, whereas the nearby cell did show an active BrdU uptake. When a number of cells were analyzed, we found that approximately two-thirds of the cells with GFP-hREV1 foci were negative for BrdU uptake without UV irradiation (Fig. 4B). Furthermore, this tendency was observed in cells after UV irradiation. About two-fifths of the GFP-hREV1 focus-forming cells showed BrdU uptake 8 h after UV irradiation at 8 J/m2 and the rest did not (Fig. 4B). We confirmed that the GFP-hREV1 foci in BrdU-negative cells were resistant to Triton-X 100 treatment prior to fixation (data not shown).
|
hREV1 foci are formed in G1 and S phase
Because it was revealed that more than half of the cells with GFP-hREV1 foci were not in S phase, we tried to assess the cell cycle of the GFP-hREV1 focus-forming cells by using the laser scanning cytometer, LSC2 system (Olympus). This equipment allows us to analyze the cell cycle status of a cell population fixed on a coverslip by means of evaluating DNA content of each cell, and hence enables us to assess the cell cycle of individual cells. MRC-5 cells transiently transfected with the pEGFP/hREV1 were fixed 8 h after UV irradiation at a dose of 8 J/m2 or without UV irradiation. Cells were then stained with propidium iodide and subjected to cell cycle analysis by using the LSC2 system. The histograms of nontreated and UV-irradiated cell populations showed an increase of percentage of cells in G1 phase after UV irradiation, suggesting cell cycle delay at G1 phase after UV irradiation (Fig. 5A). Then, the GFP-hREV1 focus-forming cells in the UV-irradiated cell population were identified, and their DNA indexes were analyzed. It was revealed that the majority of the GFP-hREV1 focus-forming cells showed their DNA indexes below 1.5 with a peak index of 1.0 (Fig. 5B), indicating that hREV1 foci were formed in G1 and early S phase. We obtained a similar result by using COS-7 cells, confirming the cell cycle-dependent focus formation of GFP-hREV1 (data not shown).
|
Because S. cerevisiae Rev1 and Rev7 belong to the same mutagenic pathway and mammalian REV1 interacts REV7 via its C-terminal region (Murakumo et al. 2001), we checked whether hREV7 expression level affects hREV1 focus formation or not. HEK293 cells with reduced expression of hREV7 protein were generated using the RNA interference (RNAi) technology. HEK293 cells were transfected with the pcPURU6ß/shREV7, and several stable cell lines were obtained after puromycin selection. The results by Western blot analysis showed that the hREV7 protein expression in the shREV7-treated cells was reduced to 10–20% of the control cells stably transfected with empty vector (Fig. 6A). Then, the hREV7-knockdown cells and the control cells were transfected with the pEGFP/hREV1, and the cells were analyzed for GFP-hREV1 focus formation with and without UV irradiation. The percentages of the GFP-hREV1 focus-forming cells were about 3–6% both in the hREV7-knockdown cells and in the control cells without UV irradiation (Fig. 6B), and the percentages increased in the hREV7-knockdown cells as well as in the control cells 8 h after 4 or 8 J/m2 UV irradiation (Fig. 6B). These findings indicated that hREV7 protein level did not affect the hREV1 focus formation, suggesting that hREV7 is not required for hREV1 focus formation.
|
In order to investigate the domain structure of hREV1 protein, we analyzed the regions of hREV1 necessary for its nuclear localization and focus formation. hREV1 cDNA fragments corresponding to hREV1 amino acid residues 1–1178, 1–1036, 1–825, 1–152, 153–1251, 387–1251, 826–1251, 1037–1251, and 387–825 were individually placed into the vector pEGFP-C3 in order to construct a series of vectors for producing GFP-fused truncated hREV1 proteins in cells (Fig. 7A). The 1–152 and 387–825 regions contain the BRCT and the catalytic domain, respectively, and the 1037–1251 contains a domain required for the interaction with other Y-family polymerases. COS-7 cells were transfected with each expression vector, and the intracellular localization of the expressed GFP-fusion protein was analyzed. Western blot analysis using anti-GFP antibody revealed that the fusion proteins were of the correct molecular masses (Fig. 7B). The GFP-fusion protein carrying hREV1 amino acid residues 1–1251, 1–1178, 1–1036, 1–825, 153–1251, or 826–1251 was localized mainly to the nucleus, whereas that carrying 387–1251 or 387–825 was mainly to the cytoplasm, and that carrying 1–152 or 1036–1251 was to both the nucleus and the cytoplasm, as GFP alone (Fig. 7C). These results suggest that a major signal for the nuclear localization of hREV1, which leads the GFP-fusion proteins to the nucleus, might reside within the 152–387 amino acid region of hREV1, and a minor signal might be present within the 826–1036 region, which leads the GFP-fusion protein with hREV1 826–1251 to the nucleus but not the protein with hREV1 387–1251. A very small fraction of the GFP-fusion protein with hREV1 153–1251, which contains the major nuclear localization signal region, was retained in the cytoplasm (Fig. 7C, panel 6). There is a possibility that this cytoplasmic retention might be due to the result of the degradation protein product of the GFP-fusion protein (Fig. 7B, lane 6). Next, we examined the focus-forming ability of each truncated hREV1 protein fused with GFP. COS-7 cells transiently expressing each GFP-fusion protein were UV-irradiated at a dose of 8 J/m2, fixed 8 h after irradiation, and analyzed for focus formation. As shown in Fig. 7C, GFP-fusion protein carrying hREV1 amino acid 1–1251, 1–1178, 153–1251, or 826–1251 showed the nuclear focus formation, whereas that carrying hREV1 1–1036, 1–825, 1–152, or 1037–1251 did not. GFP-fusion protein carrying hREV1 387–1251 or 387–825 was localized to the cytoplasm so that it did not show the nuclear focus formation. These results suggest that the domain for the nuclear focus formation very probably resides in the region of amino acid 826–1178 of hREV1 (the region F in Fig. 7A). Similar results were also obtained when the intracellular localization and the focus-forming abilities of the GFP-fusion hREV1 proteins were analyzed in MRC-5 cells.
|
|
Discussion |
|---|
|
TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
|---|
has been suggested because in an in vivo study, a mutation in the BRCT domain near the N-terminal of scRev1 affected the bypass of UV-induced DNA damages by scPol (Nelson et al. 2000). By contrast, Guo et al. (2004) recently reported that scRev1 protein enhanced scPol -catalyzed nucleotide insertion opposite the acetylaminofluorene-dG DNA adduct and strongly stimulated scPol -catalyzed extension from opposite the lesion, and that in the in vitro study, this stimulatory activity of scRev1 required the C-terminal 205 amino acids. REV1 proteins of higher eukaryotes have extended C-terminal regions, the sequences of which are very strictly conserved in mammals, chicken, and frog (Xenopus laevis). The C-terminal extended regions (the I region in Fig. 7A) of the human and mouse REV1 proteins were found to interact with the other TLS polymerases, Pol , Pol , and Pol , and with the REV7 subunit of Pol , although REV1 interactions scarcely affected the enzymatic activities of Pol and Pol in vitro (Guo et al. 2003; Ohashi et al. 2004).
In the present study, we analyzed the hREV1 localization in cells with and without UV-induced DNA damage. One purpose of this study was to assess the involvement of hREV1 protein in the DNA damage response in vivo. Our results revealed that although hREV1 formed nuclear foci in a small fraction of nonirradiated cells, the percentage of hREV1 focus-forming cells increased dramatically after UV irradiation. hREV1 nuclear foci in S-phase cells colocalized with PCNA foci, the landmarks of the replication factories. Furthermore, in the localized UV irradiation experiments, hREV1 foci accumulated in regions with DNA damage. These findings strongly suggest that hREV1 re-localizes to the replication machinery in S-phase cells in response to UV irradiation, probably at the sites of DNA lesions. The major lesions induced by UV irradiation are CPDs and (6-4) photoproducts. However, TLS through these lesions does not require the deoxycytidyl transferase activity of hREV1. Thus, the hREV1 focus formation around the UV-induced DNA lesions may indicate that hREV1 plays a noncatalytic role at these sites. This role might be associated with the functions of other TLS polymerases, because the colocalization of hREV1 with Pol , Pol and Pol was detected (Ohashi et al. 2004; Tissier et al. 2004). Damage-induced focus formation of hREV1 was also observed when cells were exposed to benzo[a]pyrene, which generates DNA adducts mainly at guanine (Mukhopadhyay et al. 2004), and this lesion is bypassed by Pol accurately and efficiently (Rechkoblit et al. 2002; Suzuki et al. 2002; Zhang et al. 2002). Thus, all members of the Y-family polymerases form nuclear foci in response to DNA damage and may be closely associated in the foci (Kannouche et al. 2001; Bergoglio et al. 2002; Ohashi et al. 2004).
We analyzed the cell cycle status of the hREV1 focus-forming cells in detail, which revealed that hREV1 foci are mainly formed in G1 and early S phase. In addition, our results using localized UV irradiation showed that hREV1 clustered foci generated around the DNA lesion were observed not only in S phase but also outside S phase. These results suggest a possibility that hREV1 might be associated with damaged DNA not only in S phase but also in G1 phase. Because it is generally believed that TLS is carried out in S phase at the site of replication fork arrest, the hREV1 focus formation in G1 phase are not accompanied with the TLS events. Two possibilities concerning hREV1 focus formation in G1 phase are (i) hREV1 is involved in excision repair and (ii) hREV1 is introduced at DNA damaged sites in the G1 phase to recruit DNA repair proteins to the lesions before replication. More recently, it was reported that scRev1 participates in the formation of replication-independent frame-shift mutation induced by UV irradiation in cell cycle-arrested, stationary-phase yeast cells (Heidenreich et al. 2006). At present, it is not known whether these foci in G1 phase have any biologic meanings. However, taken together with their results and the present findings, there is a possibility that hREV1 has unknown functions outside the S phase, especially in G1 phase, which are not associated with active TLS events.
Tissier et al. (2004) recently reported similar results concerning hREV1 focus formation, in which 10–15% of the cells expressing GFP-hREV1 formed foci before UV irradiation and approximately 70% after UV irradiation. There is a discrepancy between their results and ours in the percentages of the hREV1 focus-forming cells. A similar discrepancy also was found in studies of Pol (Bergoglio et al. 2002; Ogi et al. 2005). Such discrepancies might be caused by the difference in the definition of "nuclear foci" in each study. As described in the Results section, we defined "nuclear foci" as the clearly sharp and tiny speckles evenly distributed in the nucleus in a manner similar to PCNA foci. We excluded cells with large, unevenly distributed spots or few spots from cells with "nuclear foci." Therefore, the population of focus-forming cells in our study was lower than those in their study. However, even with our stringent criteria, a small fraction of cells with GFP-hREV1 expression showed focus formation without UV irradiation. The GFP-hREV1 foci in a non-UV-irradiated cell are significantly less numerous than in a UV-irradiated cell (Fig. 1A); therefore, spontaneous DNA damage may be the cause of the hREV1 focus formation in non-UV-irradiated cells.
We determined the region of hREV1 required for focus formation to be within the region of amino acid residues 826–1178, which is the region adjacent to the domain required for the interaction with hREV7, Pol , Pol , and Pol (Murakumo et al. 2001; Guo et al. 2003; Ohashi et al. 2004). Our finding is distinctly different from the result reported by Tissier et al. (2004), which indicated that the N-terminal half fragment as well as the C-terminal half fragment of hREV1 showed focus formation. The possible reason why the difference between the two studies occurred must be the difference of fluorescent constructs used in each study. The GFP-fusion protein carrying hREV1 amino acid residues 1–1036 or 1–825, which did not form nuclear foci in our study, included the conserved region of 419–825, whereas the N-terminal half construct used in their study representing amino acid 1–730 did not include the full of the conserved region. The GFP-fusion protein carrying hREV1 387–825 was retained in the cytoplasm without nuclear focus formation (Fig. 7C, panel 10). In view of this, there is a possibility that the conserved region of hREV1 has an inhibiting effect on the nuclear focus formation when the GFP-fused hREV1 1–1036 or 1–825 is expressed in cells, although the protein is localized to the nucleus. Following UV irradiation, the GFP-fusion protein harboring hREV1 amino acid 1–1178, which lacked a portion of the interaction domain with hREV7, Pol , Pol , and Pol , showed clear focus formation in cells, and the percentage of focus-forming cells was almost the same as that of cells with GFP-hREV1 (data not shown), indicating that the focus formation of hREV1 is independent of its interaction with other TLS polymerases. This was supported by the present finding that hREV7 knockdown did not affect the focus-forming ability of hREV1, and also by the finding that GFP-hREV1 focus formation was observed in XP-variant cells (Tissier et al. 2004).
How does hREV1 form foci after DNA damage? A potential mechanism of hREV1 focus formation in G1-phase cells is through binding of hREV1 to proteins involved in excision repair or direct binding to damaged DNA, although there is not sufficient supportive data at present. One possible mechanism of the hREV1 focus formation in S-phase cells is through direct binding of hREV1 to PCNA, as suggested for the focus formation of other TLS polymerases (Vidal et al. 2004; Ogi et al. 2005). In the case of Pol , the PCNA binding site was identified to be the 420-KKGLIDYY-427 sequence with a similarity to the consensus sequence of PCNA-binding motif, Q-x-x-(I, L, M)-x-x-FF (Vidal et al. 2004; Haracska et al. 2005), in which the YY-to-AA substitution drastically diminished the focus formation after UV-irradiation (Vidal et al. 2004). For focus formation of Pol , the C-terminal 97 amino acid residues are sufficient for localization into nuclear foci, which include a C2HC zinc finger motif, bipartite nuclear localization signal and putative PCNA binding site, and the FF-to-AA substitution in the putative PCNA binding site abolished the focus formation upon DNA damaging (Ogi et al. 2005). Also, in the case of Pol , the C-terminal 119 amino acids are sufficient for correct localization into nuclei and nuclear foci (Kannouche et al. 2001), which contains a C2H2 zinc finger motif, a nuclear localization signal, and a PCNA binding site, very similarly to the case of Pol , and all of the three motifs are required for correct localization (unpublished results by P. Kannouche and A. R. Lehmann, cited in the reference of Ogi et al. 2005). Recently, the RAD18 function was found to be necessary for the focus formation of Pol , and the RAD18 protein was shown to directly bind to a C-terminal region (158 amino acids) of Pol (Watanabe et al. 2004). We find no sequence with strong similarity to the consensus sequence of PCNA binding site in the 826–1178 region of hREV1; however, more recently, Ross et al. (2005) showed by using the mammalian two-hybrid assay that a region between amino acid residues 923 and 1047 of hREV1 mediates an interaction with PCNA, and that these C-terminal domains of hREV1 are necessary for effective DNA damage tolerance in the DT40 cells. Our present results showing the domain of hREV1 required for the focus formation are exactly compatible with their findings of the PCNA binding domain of hREV1. Furthermore, Ross et al. (2005) showed that binding of RAD18 to hREV1 was not detected by the same assay. Further experiments are definitely required in order to clarify the mechanisms of the hREV1 nuclear focus formation and its roles in DNA damage response and TLS.
|
Experimental procedures |
|---|
|
TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
|---|
cDNA fragments of full-length hREV1 or truncated hREV1, encoding amino acids 1–152, 1–825, 1–1036, 1–1178, 153–1251, 387–825, 387–1251, 826–1251, or 1037–1251, were placed into the vector pEGFP-C3 (Clontech) in order to produce GFP-fusion proteins in cells (pEGFP/hREV1s). Full-length hREV1 cDNA was inserted into the vector pcDNA3.1(+) (Invitrogen) with a FLAG sequence at the 5'-terminal of its open reading frame in order to produce FLAG-tagged hREV1 protein in cells (pcDNA/FLAG-hREV1). The cDNA fragment cloned into each vector was produced by polymerase chain reaction with the Pfu DNA polymerase (Stratagene). The plasmid DNAs were amplified in Escherichia coli DH5
and purified with the Qiagen plasmid kit (Qiagen).
Cell culture and reagents
COS-7 (monkey kidney fibroblast), MRC-5 (human fetal lung fibroblast) and HEK293 (human fetal kidney epithelial cell) cells were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 8% fetal bovine serum (FBS). For transient transfection experiments, cells were grown on appropriate dishes and transfected with expression vectors using the Lipofectamine 2000 transfection reagent (Invitrogen) according to the manufacturer's protocol. The cells were used for analyses 48 h after transfection.
Generation of hREV7 knockdown cell lines
The shRNA (short hairpin RNA) expression vector for silencing hREV7 expression was constructed using the pcPURU6ß icassette plasmid vector (iGENE Therapeutics), which contains a human U6 promoter. The target sequence of hREV7 is GAC AAG ACC TCA ACT TTG G. HEK293 cells were transfected with the hREV7 shRNA expression vector (pcPURU6ß/shREV7) or a control empty vector by using the Lipofectamine 2000 transfection reagent (Invitrogen), and 0.5 µg/mL puromycin was added to the medium 72 h after transfection. Several stable transfectants, each of which was derived from a single colony, were obtained after puromycin selection, and the hREV7 expression levels were assessed with Western blotting using anti-hREV7 antibody.
UV irradiation
Cells grown on a coverslip in a 35-mm culture dish were rinsed twice with PBS, and were then UV-irradiated at 254 nm with a UV254 lamp at a fluence rate of 1.0 J/m2/s without PBS. Cells were incubated at 37 °C in fresh growth medium in a CO2 incubator until analyzed. For localized UV irradiation, cells grown on a coverslip were rinsed twice with PBS and UV-irradiated through a 3.0-µm IsoporeTM membrane filter (Millipore) at 80 J/m2 without PBS. They were then incubated at 37 °C in fresh growth medium in a CO2 incubator until analyzed.
Antibodies
Mouse monoclonal anti-FLAG M2 antibody was purchased from Sigma. Mouse monoclonal anti-PCNA antibody was purchased from Dako Cytomation. Mouse monoclonal anti-BrdU antibody was purchased from Roche. TDM-2 mouse monoclonal antibody for the detection of T-T CPDs was kindly provided by Dr T. Matsunaga of Kanazawa University. Mouse monoclonal anti-GFP antibody was purchased from Nacalai Tesque. Rabbit polyclonal anti-hREV7 antibody was produced by immunization with keyhole limpet hemacyanin-conjugated peptide containing the C-terminal 19 amino acids of hREV7 and affinity purified as previously described (Murakumo et al. 2000). Goat anti-mouse IgG secondary antibody conjugated with Alexa 488 or Alexa 594 was purchased from Molecular Probes.
Western blot analysis
Cells were harvested in a microcentrifuge tube using a cell scraper and disrupted in cell lysis buffer (20 mM HEPES, pH 7.6, 350 mM NaCl, 1 mM ethylenediaminetatraacetic acid, 10% glycerol, 1 mM DTT, 1 mM phenylmethylsulfonyl fluoride (PMSF), 10 µg/mL leupeptin, 5 µg/mL pepstatin A) with three freeze and thaw cycles. The cell lysates were clarified by centrifugation (20 000x g) for 10 min and boiled for 2 min with the same amount of 2x sample buffer (62.5 mM Tris-HCl, pH 6.8, 2% sodium dodecyl sulfate (SDS), 25% glycerol, 5% 2-mercaptoethanol, 0.01% bromophenol blue). Proteins were subjected to SDS-polyacrylamide gel electrophoresis (PAGE) and transferred on to a polyvinylidene difluoride membrane (Millipore). After blocking with 5% bovine serum albumin (BSA) in Tris- buffered saline with Tween 20 buffer (20 mM Tris-HCl, pH 7.6, 137 mM NaCl, 0.1% Tween 20) for 1 h, the membrane was probed with anti-GFP antibody (2.2 mg/mL) at 1 : 1000 dilution or anti-hREV7 antibody (2 mg/mL) at 1 : 1000 dilution followed by an incubation with the secondary antibody conjugated to horseradish peroxidase (Dako Cytomation). After intensive washing, the antigen–antibody complexes were detected with the ECL Western blotting detection reagent (Amersham Pharmacia Biotech).
Fluorescent immunocytochemistry
For visualization of GFP-fusion proteins, cells transiently expressing GFP-fusion proteins grown on a coverslip were rinsed twice with PBS and fixed by soaking with 4% paraformaldehyde at 4 °C for 20 min. After washing, the cells were mounted on a slide glass and GFP-fusion proteins were visualized under a fluorescence microscope. The fluorescent images were captured with the FLUOVIEW FV500 confocal laser scanning microscope (Olympus) or the DP70 CCD (charge-coupled device) camera (Olympus). For fluorescent immunocytochemical staining with anti-FLAG antibody, cells grown on a coverslip were fixed by soaking in 100% cold methanol for 20 min at –20 °C and permeabilized with 0.5% Triton X-100 for 5 min at room temperature (RT). After blocking with 1% BSA in PBS for 30 min at 37 °C, cells were incubated with anti-FLAG antibody (4.4 mg/mL) at 1 : 500 dilution with 1% BSA in PBS for 30 min at 37 °C. This was followed by an incubation with the secondary antibody conjugated to Alexa 488 (2 mg/mL) at 1 : 1000 dilution with 1% BSA in PBS for 30 min at 37 °C. After intensive washing, cells were mounted and FLAG-tagged proteins were visualized as described previously. For PCNA staining, cells grown on a coverslip were fixed by soaking with 100% cold methanol for 20 min at –20 °C, and then incubated for 30 s with cold acetone in order to extract the soluble part of PCNA. After blocking with 1% BSA in PBS for 30 min at 37 °C, cells were incubated with anti-PCNA antibody (0.57 mg/mL) at 1 : 200 dilution with 1% BSA in PBS for 30 min at 37 °C, followed by an incubation with the Alexa 594-conjugated secondary antibody for 30 min at 37 °C. For BrdU staining, cells grown on a coverslip were incubated in growth medium with 10 µM BrdU (Roche) for 15 min and fixed by soaking with 99% ethanol +1% acetic acid for 20 min at –20 °C. They were incubated with anti-BrdU antibody for 30 min at 37 °C, followed by an incubation with the Alexa 594-conjugated secondary antibody for 30 min at 37 °C. For the detection of DNA lesions represented by T-T CPD, cells treated with localized UV irradiation were fixed by soaking with 4% formalin at RT for 10 min. After permeabilization with 0.5% Triton X-100 for 5 min at 4 °C, cells were exposed to 2 M HCl for 10 min. They were blocked with 20% FBS in PBS for 30 min at 37 °C, then incubated with TDM-2 antibody at 1 : 1000 dilution with 5% FBS in PBS for 30 min at 37 °C, followed by an incubation with the Alexa 594-conjugated secondary antibody for 30 min at 37 °C.
Cell cycle analysis
COS-7 or MRC-5 cells with GFP-hREV1 expression grown on a coverslip were fixed with 100% ethanol overnight at –20 °C. Cells were rinsed with PBS and incubated in PBS with 50 µg/mL propidium iodide and 100 µg/mL RNase A for 30 min at 37 °C. After mounting, cells were subjected for cell cycle analyses by using the laser scanning cytometer, LSC2 system (Olympus).
|
Acknowledgements |
|---|
|
Footnotes |
|---|
* Correspondence: E-mail: murakumo{at}med.nagoya-u.ac.jp
|
References |
|---|
|
TopAbstractIntroductionResultsDiscussionExperimental proceduresReferences |
|---|
to replication foci. J. Cell Sci.
115, 4413–4418.Chen, J. & Fang, G. (2001) MAD2B is an inhibitor of the anaphase-promoting complex. Genes Dev.
15, 1765–1770.
Clark, D.R., Zacharias, W., Panaitescu, L. & McGregor, W.G. (2003) Ribozyme-mediated REV1 inhibition reduces the frequency of UV-induced mutations in the human HPRT gene. Nucleic Acids Res.
31, 4981–4988.
Friedberg, E.C., McDaniel, L.D. & Schultz, R.A. (2004) The role of endogenous and exogenous DNA damage and mutagenesis. Curr. Opin. Genet. Dev. 14, 5–10.[CrossRef][Medline]
Friedberg, E.C., Walker, G.C. & Siede, W. (1995) DNA Repair and Mutagenesis. Washington, DC: American Society for Microbiology.
Gibbs, P.E.M., McGregor, W.G., Maher, V.M., Nisson, P. & Lawrence, C.W. (1998) A human homolog of the Saccharomyces cerevisiae REV3 gene, which encodes the catalytic subunit of DNA polymerase . Proc. Natl. Acad. Sci. USA
95, 6876–6880.
Gibbs, P.E.M., Wang, X.-D., Li, Z., et al. (2000) The function of the human homolog of Saccharomyces cerevisiae REV1 is required for mutagenesis induced by UV light. Proc. Natl. Acad. Sci. USA
97, 4186–4191.
Guo, C., Fischhaber, P.L., Luk-Paszyc, M.J., et al. (2003) Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J. 22, 6621–6630.[CrossRef][Medline]
Guo, D., Xie, Z., Shen, H., Zhao, B. & Wang, Z. (2004) Translesion synthesis of acetylaminofluorene-dG adducts by DNA polymerase is stimulated by yeast Rev1 protein. Nucleic Acids Res.
32, 1122–1130.
Haracska, L., Acharya, N., Unk, I., et al. (2005) A single domain in human DNA polymerase iota mediates interaction with PCNA: implications for translesion DNA synthesis. Mol. Cell. Biol.
25, 1183–1190.
Haracska, L., Prakash, S. & Prakash, L. (2002) Yeast Rev1 protein is a G template-specific DNA polymerase. J. Biol. Chem.
277, 15546–15551.
Heidenreich, E., Eisler, H. & Steinboeck, F. (2006) Epistatic participation of REV1 and REV3 in the formation of UV-induced frameshift mutations in cell cycle-arrested yeast cells. Mutat. Res. in press.
Johnson, R.E., Kondratick, C.M., Prakash, S. & Prakash, L. (1999) hRAD30 mutations in the variant form of xeroderma pigmentosum. Science
285, 263–265.
Johnson, R.E., Washington, M.T., Haracska, L., Prakash, S. & Prakash, L. (2000) Eukaryotic polymerases and act sequentially to bypass DNA lesions. Nature
406, 1015–1019.[CrossRef][Medline]
Kannouche, P., Broughton, B.C., Volker, M., Hanaoka, F., Mullenders, L.H.F. & Lehmann, A.R. (2001) Domain structure, localization, and function of DNA polymerase , defective in xeroderma pigmentosum variant cells. Genes Dev.
15, 158–172.
Kannouche, P., Fernandez de Henestrosa, A.R., Coull, B., et al. (2002) Localization of DNA polymerases and to the replication machinery is tightly co-ordinated in human cells. EMBO J.
21, 6246–6256.[Medline]
Kunz, B.A., Straffon, A.F.L. & Vonarx, E.J. (2000) DNA damage-induced mutation: tolerance via translesion synthesis. Mutat. Res. 451, 169–185.[Medline]
Lawrence, C.W. (2002) Cellular roles of DNA polymerase and Rev1 protein. DNA Repair
1, 425–435.[CrossRef][Medline]
Lawrence, C.W. & Maher, V.M. (2001) Eukaryotic mutagenesis and translesion replication dependent on DNA polymerase and Rev1 protein. Biochem. Soc. Trans.
29, 187–191.[CrossRef][Medline]
Lin, W., Xin, H., Zhang, Y., et al. (1999) The human REV1 gene codes for a DNA template-dependent dCMP transferase. Nucleic Acids Res.
27, 4468–4475.
Masuda, Y., Ohmae, M., Masuda, K. & Kamiya, K. (2003) Structure and enzymatic properties of a stable complex of the human REV1 and REV7 proteins. J. Biol. Chem.
278, 12356–12360.
Masutani, C., Kusumoto, R., Yamada, A., et al. (1999) The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase . Nature
399, 700–704.[CrossRef][Medline]
Mukhopadhyay, S., Clark, D.R., Watson, N.B., Zacharias, W. & McGregor, W.G. (2004) REV1 accumulates in DNA damage-induced nuclear foci in human cells and is implicated in mutagenesis by benzo[a]pyrenediolepoxide. Nucleic Acids Res.
32, 5820–5826.
Murakumo, Y. (2002) The property of DNA polymerase zeta: REV7 is a putative protein involved in translesion DNA synthesis and cell cycle control. Mutat. Res. 510, 37–44.[Medline]
Murakumo, Y., Ogura, Y., Ishii, H., et al. (2001) Interactions in the error-prone postreplication repair proteins hREV1, hREV3, and hREV7. J. Biol. Chem.
276, 35644–35651.
Murakumo, Y., Roth, T., Ishii, H., et al. (2000) A human REV7 homolog that interacts with the polymerase catalytic subunit hREV3 and the spindle assembly checkpoint protein hMAD2. J. Biol. Chem.
275, 4391–4397.
Nelson, J.R., Gibbs, P.E.M., Nowicka, A.M., Hinkle, D.C. & Lawrence, C.W. (2000) Evidence for a second function for Saccharomyces cerevisiae Rev1p. Mol. Microbiol. 37, 549–554.[CrossRef][Medline]
Nelson, J.R., Lawrence, C.W. & Hinkle, D.C. (1996a) Thymine-thymine dimer bypass by yeast DNA polymerase . Science
272, 1646–1649.[Abstract]
Nelson, J.R., Lawrence, C.W. & Hinkle, D.C. (1996b) Deoxycytidyl transferase activity of yeast REV1 protein. Nature 382, 729–731.[CrossRef][Medline]
Ogi, T., Kannouche, P. & Lehmann, A.R. (2005) Localisation of human Y-family DNA polymerase : relationship to PCNA foci. J. Cell Sci.
118, 129–136.
Ohashi, E., Murakumo, Y., Kanjo, N., et al. (2004) Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells
9, 523–531.
Ohmori, H., Friedberg, E.C., Fuchs, R.P.P., et al. (2001) The Y-family of DNA polymerases. Mol. Cell 8, 7–8.[CrossRef][Medline]
Pfleger, C.M., Salic, A., Lee, E. & Kirschner, M.W. (2001) Inhibition of Cdh1-APC by the MAD2-related protein MAD2L2: a novel mechanism for regulating Cdh1. Genes Dev.
15, 1759–1764.
Prakash, S. & Prakash, L. (2002) Translesion DNA synthesis in eukaryotes: a one- or two-polymerase affair. Genes Dev.
16, 1872–1883.
Rajpal, D.K., Wu, X. & Wang, Z. (2000) Alteration of ultraviolet-induced mutagenesis in yeast through molecular modulation of the REV3 and REV7 gene expression. Mutat. Res. 461, 133–143.[Medline]
Rechkoblit, O., Zhang, Y., Guo, D., et al. (2002) trans-Lesion synthesis past bulky benzo[a]pyrene diol epoxide N2-dG and N6-dA lesions catalyzed by DNA bypass polymerases. J. Biol. Chem.
277, 30488–30494.
Ross, A.L., Simpson, L.J. & Sale, J.E. (2005) Vertebrate DNA damage tolerance requires the C-terminus but not BRCT or transferase domains of REV1. Nucleic Acids Res.
33, 1280–1289.
Suzuki, N., Ohashi, E., Kolbanovskiy, A., et al. (2002) Translesion synthesis by human DNA polymerase on a DNA template containing a single stereoisomer of dG-(+)- or dG-(-)-anti-N2-BPDE (7,8-dihydroxy-anti-9,10-epoxy-7,8,9,10-tetrahydrobenzo[a]pyrene). Biochemistry
41, 6100–6106.[CrossRef][Medline]
Tissier, A., Frank, E.G., McDonald, J.P., Iwai, S., Hanaoka, F. & Woodgate, R. (2000b) Misinsertion and bypass of thymine-thymine dimers by human DNA polymerase . EMBO J.
19, 5259–5266.[CrossRef][Medline]
Tissier, A., Kannouche, P., Reck, M.-P., Lehmann, A.R., Fuchs, R.P.P. & Cordonnier, A. (2004) Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol and REVl protein. DNA Repair
3, 1503–1514.[CrossRef][Medline]
Tissier, A., McDonald, J.P., Frank, E.G. & Woodgate, R. (2000a) Pol, a remarkably error-prone human DNA polymerase. Genes Dev.
14, 1642–1650.
Vidal, A.E., Kannouche, P., Podust, V.N., Yang, W., Lehmann, A.R. & Woodgate, R. (2004) Proliferating cell nuclear antigen-dependent coordination of the biological functions of human DNA polymerase . J. Biol. Chem.
279, 48360–48368.
Watanabe, K., Tateishi, S., Kawasuji, M., Tsurimoto, T., Inoue, H. & Yamaizumi, M. (2004) Rad18 guides pol to replication stalling sites through physical interaction and PCNA monoubiquitination. EMBO J.
23, 3886–3896.[CrossRef][Medline]
Xiao, W., Lechler, T., Chow, B.L., et al. (1998) Identification, chromosomal mapping and tissue-specific expression of hREV3 encoding a putative human DNA polymerase . Carcinogenesis
19, 945–949.
Zhang, Y., Wu, X., Guo, D., Rechkoblit, O. & Wang, Z. (2002) Activities of human DNA polymerase in response to the major benzo[a]pyrene DNA adduct: error-free lesion bypass and extension synthesis from opposite the lesion. DNA Repair
1, 559–569.[CrossRef][Medline]
Received: 11 October 2005
Accepted: 28 November 2005
This article has been cited by other articles:
|
|
 |
|
  L. S. Waters, B. K. Minesinger, M. E. Wiltrout, S. D'Souza, R. V. Woodruff, and G. C. Walker Eukaryotic Translesion Polymerases and Their Roles and Regulation in DNA Damage Tolerance Microbiol. Mol. Biol. Rev., March 1, 2009; 73(1): 134 - 154. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Zhang, S.-H. Yang, and A. D. Sharrocks Rev7/MAD2B Links c-Jun N-Terminal Protein Kinase Pathway Signaling to Activation of the Transcription Factor Elk-1 Mol. Cell. Biol., April 15, 2007; 27(8): 2861 - 2869. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Guo, T.-S. Tang, M. Bienko, J. L. Parker, A. B. Bielen, E. Sonoda, S. Takeda, H. D. Ulrich, I. Dikic, and E. C. Friedberg Ubiquitin-Binding Motifs in REV1 Protein Are Required for Its Role in the Tolerance of DNA Damage Mol. Cell. Biol., December 1, 2006; 26(23): 8892 - 8900. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. Nakajima, L. Lan, S.-i. Kanno, N. Usami, K. Kobayashi, M. Mori, T. Shiomi, and A. Yasui Replication-dependent and -independent Responses of RAD18 to DNA Damage in Human Cells J. Biol. Chem., November 10, 2006; 281(45): 34687 - 34695. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. S. Waters and G. C. Walker The critical mutagenic translesion DNA polymerase Rev1 is highly expressed during G2/M phase rather than S phase PNAS, June 13, 2006; 103(24): 8971 - 8976. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Shen, S. Jun, L. E. O'Neal, E. Sonoda, M. Bemark, J. E. Sale, and L. Li REV3 and REV1 Play Major Roles in Recombination-independent Repair of DNA Interstrand Cross-links Mediated by Monoubiquitinated Proliferating Cell Nuclear Antigen (PCNA) J. Biol. Chem., May 19, 2006; 281(20): 13869 - 13872. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | ADVANCED SEARCH | TABLE OF CONTENTS |
