• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Jun 27, 2006; 103(26): 9891–9896.
Published online Jun 20, 2006. doi:  10.1073/pnas.0603779103
PMCID: PMC1502549
Cell Biology

H2AX phosphorylation within the G1 phase after UV irradiation depends on nucleotide excision repair and not DNA double-strand breaks

Abstract

The variant histone H2AX is phosphorylated in response to UV irradiation of primary human fibroblasts in a complex fashion that is radically different from that commonly reported after DNA double-strand breaks. H2AX phosphorylation after exposure to ionizing radiation produces foci, which are detectable by immunofluorescence microscopy and have been adopted as clear and consistent quantitative markers for DNA double-strand breaks. Here we show that in contrast to ionizing radiation, UV irradiation mainly induces H2AX phosphorylation as a diffuse, even, pan-nuclear staining. UV induced pan-nuclear phosphorylation of H2AX is present in all phases of the cell cycle and is highest in S phase. H2AX phosphorylation in G1 cells depends on nucleotide excision repair factors that may expose the S-139 site to kinase activity, is not due to DNA double-strand breaks, and plays a larger role in UV-induced signal transduction than previously realized.

Keywords: ultraviolet light, xeroderma pigmentosum, chromatin structure, Wortmannin, human fibroblasts

H2AX phosphorylation plays a major role in nuclear events during meiosis and DNA double-strand breaks (DSBs). DSBs cause phosphorylation of histone H2AX on large areas of chromatin flanking the breaks (1). Phosphorylation, coupled with the acetylation-dependent condensation of chromatin, permits microscopic visualization of discrete nuclear foci (2). Phosphorylated H2AX (γH2AX) foci formation is a powerful tool used to study DSB formation and repair after genomic damage, chromosome dynamics, and signaling mechanisms (37). Analysis of knockout mice and cell lines implicate H2AX phosphorylation in genomic stability, tumor suppression, and spermatogenesis (8, 9).

Although H2AX phosphorylation in response to ionizing radiation and other radiomimetic agents garners a great deal of attention, its role in the UV-induced DNA damage response is not well characterized. Reports have limited γH2AX formation to S phase after exposure to UV-B (10, 11). We have reported previously coincidence of γH2AX, MRE11, and PCNA in foci that represent replication fork breakage in the S phase of SV40-transformed cells (12, 13). In contrast, a recent report finds that γH2AX is present in HeLa and HL60 cells at 1 h after UV-B irradiation in all phases of the cell cycle (14). These studies speculate that H2AX phosphorylation after UV irradiation is induced only by processing of UV-induced lesions into DNA DSBs, but we report a more complex response.

Historically, direct DNA breakage from UV-C has been undetectable in mammalian cells (15). Calculation of the maximum frequency of DNA breaks during excision repair after UV light, from these experiments, yields an estimate of 3,000–4,000 single strand breaks per diploid cell associated with nucleotide excision repair (NER) at a maximum rate within 1 h of 10–20 J/m2 with a separation of >109 Da (4 × 106 nt) between excision sites (1618). The number of photoproducts per cell under these conditions is ≈2 × 106, and repair operates on only ≈0.1% of the initial photoproducts at any one time. DSBs associated with direct UV damage or overlapping NER sites therefore are extremely rare.

We show that after UV-C (UV) irradiation (254 nm), in primary human fibroblasts, H2AX phosphorylation occurs in all phases of the cell cycle and is primarily displayed as a diffuse, even staining of the whole nucleus, which we term “pan-nuclear” staining. The intensity of pan-nuclear staining is cell cycle-dependent with two classes of intense γH2AX staining in S phase cells, whereas G1 cells contain the least intensely pan-nuclear-stained cells. Within the G1 phase of the cell cycle, H2AX is phosphorylated in the absence of DNA DSBs and is decreased in cells lacking core nucleotide excision repair factors.

Results

H2AX Is Phosphorylated After UV Irradiation.

Antibodies specific for the phosphorylation of Ser-139 at the C-terminal region of H2AX allow for detection of γH2AX by immunofluorescence microscopy and flow cytometry (19). In mock-treated fibroblasts, most cells do not contain levels of γH2AX that are detectable by immunofluorescence microscopy (see Fig. 3a). Four hours after exposure to an acute dose of 20 J/m2 UV, primary fibroblasts show increased levels of-γH2AX by flow cytometry (Fig. 1a). Gating cells based on nuclear content, measured by propidium iodide (PI) staining, reveals distinct patterns of G1 and S phase H2AX phosphorylation 4 h after UV irradiation (Fig. 1a). An acute dose of UV caused a 2- to 3-fold increase in γH2AX within G1 cells compared with mock-treated controls.

Fig. 1.
UV irradiation induces H2AX phosphorylation in all phases of the cell cycle in cycling and contact-inhibited cells. (a) Overlay of unirradiated (filled histogram) and UV-irradiated (open histogram) cells represents flow cytometry data that demonstrate ...
Fig. 3.
UV irradiation induces predominantly pan-nuclear H2AX phosphorylation in all stages of the cell cycle. (a) Immunofluorescence of γH2AX and CPDs in normal human fibroblasts after exposure to UV, ionizing radiation or mock treatment. Nuclear DNA ...

The S phase population, after irradiation, is more heavily phosphorylated and less homogeneous than the G1 population (Fig. 1b). UV-irradiated S phase cells form two classes based on their level of γH2AX intensity relative to controls (Fig. 1b). Cells in the first S phase class (class I) contain 5- to 6-fold more γH2AX and are present primarily during early and late S phase. The second S phase class (class II) contains 15- to 20-fold more γH2AX than mock-treated controls and are spread throughout S phase with the greatest density of cells in the mid-S phase region (Fig. 7, which is published as supporting information on the PNAS web site).

Contact Inhibited Cells Showed a Reduced Proportion of Cells in S Phase.

After UV irradiation, the G1 population of contact-inhibited cells still showed a 2- to 3-fold increase in γH2AX (Fig. 1a). Contact inhibition before UV irradiation altered the H2AX phosphorylation pattern within the remaining S phase cells. In contact-inhibited cells, only one S phase class was detectable that had similar high γH2AX intensity as class II of cycling cells (Fig. 1a). The class I phosphorylation in S phase requires ongoing DNA replication, and we have found that it is ATR-dependent (E.H. and J.E.C., unpublished data).

Kinetics of UV-Induced H2AX Phosphorylation.

H2AX phosphorylation follows a complex temporal pattern after UV irradiation. This pattern is in contrast to ionizing radiation exposure where γH2AX is detectable minutes after irradiation, reaches maximum levels within 30 min, and rapidly declines to low levels by 2 h (1, 13). After UV irradiation, H2AX phosphorylation appears within 1 h in all phases of the cell cycle. Flow cytometry of γH2AX content after UV exposure shows that maximum G1 γH2AX content is achieved after 2 h and is maintained out to 4 h (Fig. 2a). Between 4 and 8 h, G1 γH2AX phosphorylation begins to recede and is completely absent by 24 h. PI staining revealed that between 4 and 8 h, after UV irradiation, the previously arrested G1 cells begin to enter S phase. This movement of G1 cells into S phase coincides with a second wave of S phase-specific H2AX phosphorylation (Figs. (Figs.22 and 7).

Fig. 2.
Time course of γH2AX fluorescence after UV irradiation. (a) Histograms represent γH2AX fluorescence intensity at different time points after UV irradiation overlayed on the control sample. Determination of G1 and S phase population based ...

Presentation of γH2AX Foci Within the Nucleus.

Although flow cytometry offers detailed information regarding the total level of cellular H2AX phosphorylation, it does not reveal information regarding the distribution or appearance of γH2AX within the nucleus. Immunofluorescence microscopy of untreated fibroblasts shows that ≈25% of cells have a small number of dim nuclear γH2AX foci (ref. 20; Fig. 3a). Nearly all of these untreated γH2AX foci-positive cells are also BrdU positive, which indicates that they are S phase cells (Fig. 8, which is published as supporting information on the PNAS web site). Four hours after exposure to 20 J/m2 UV, human fibroblasts show varying nuclear H2AX phosphorylation patterns (Fig. 3b). Approximately 9% of the cells show bright γH2AX foci staining similar to the γH2AX foci seen after irradiation with ionizing radiation (see Fig. 5a). The decrease in γH2AX foci-positive cells from 25% with dim foci after mock treatment to 9% with bright foci after UV irradiation could be explained by the fact that in a fraction of the UV-irradiated cells, the γH2AX foci are masked by the far-stronger γH2AX pan-nuclear staining. Disruption of normal replication fork initiation and progression by UV also can reduce the number of S phase foci (21, 22).

Fig. 5.
H2AX phosphorylation after UV irradiation occurs in the absence of DNA DSBs. (a) Filled histogram represents flow cytometry of γH2AX fluorescence in untreated control cells, whereas open solid line and dashed line histograms represent cells treated ...

UV-Induced Pan-Nuclear γH2AX Staining.

In mock-treated human fibroblasts, only 1% of the cells show pan-nuclear γH2AX staining (data not shown). Four hours after 20 J/m2 UV irradiation, 22% of cells show bright pan-nuclear γH2AX staining (class I), whereas 24% of the cells show even brighter γH2AX staining (class II), which we termed “super-bright” pan-nuclear γH2AX staining (Fig. 3b). Colocalization with BrdU after UV irradiation reveals that S phase cells have either bright or super-bright pan-nuclear γH2AX staining (Fig. 3c). Therefore, the BrdU colocalization and the flow cytometry both show that the two classes (I + II) with bright and super-bright γH2AX staining after UV irradiation are S phase cells (Figs. 1b and and33c).

Four hours after 20 J/m2 UV irradiation, immunofluorescence microscopy reveals that 45% of the cells show even but dim H2AX phosphorylation over the whole nucleus that is clearly stronger than the H2AX phosphorylation detectable in mock-treated control cells (Fig. 3a). Flow cytometry and BrdU/γH2AX colocalization reveal that the cells with weak staining after UV irradiation are predominantly G1 cells (Fig. 1b and and33b). Thus, in the absence of replication, UV irradiation induces a low-intensity pan-nuclear H2AX phosphorylation.

To elucidate the mechanism of pan-nuclear γH2AX phosphorylation, we used spatially restricted UV irradiation: Cells were irradiated with 100 J/m2 UV through 3-μm isopore filters and probed for the presence of cyclobutane pyrimidine dimers (CPDs) and γH2AX by immunofluorescence microscopy. Four hours after spatial restricted UV irradiation, normal cells show circular regions in the nuclei, which are positive for CPD staining (Fig. 4). These CPD-positive regions colocalize almost completely with γH2AX staining. The irradiated regions clearly show uniform γH2AX staining and do not contain foci. Thus, spatially restricted UV irradiation does not elicit foci or nuclearwide H2AX phosphorylation.

Fig. 4.
UV irradiation restricted to subnuclear regions triggers phosphorylation of H2AX exclusively at sites of UV exposure and not throughout the nucleus. Normal human fibroblasts were irradiated through 3.0-μm isopore filters with 100 J/m2 UV. Overlay ...

H2AX Phosphorylation After UV Irradiation Is Not due to DNA DSB Formation.

There is strong historical evidence, discussed in the Introduction, that UV irradiation does not directly induce DNA DSB formation. To confirm that DNA DSBs are not present in the G1 cells after UV irradiation, we calibrated γH2AX intensity by flow cytometry within the G1 population of UV-irradiated cells. The measured γH2AX intensity of the G1 population of UV-irradiated cells equals the intensity profile of G1 cells that were irradiated with 5 Gy of ionizing radiation (Fig. 5a). Ionizing radiation (5 Gy) induces ≈175 DSBs per G1 cell (1), which are visible by immunofluorescence microscopy. After UV irradiation, γH2AX foci were seldom detected in G1 cells and, when foci were present, they never approached the number seen in cells irradiated with 5 Gy of ionizing radiation (Fig. 5a).

53BP1 and the phosphorylated form of NBS1(ser343) form foci in response ionizing radiation or replication-induced DNA breaks (Fig. 5 b and c; refs. 23 and 24). In G1 cells after exposure to UV irradiation, no phospho NBS1(ser343) fluorescence is evident; 53BP1 forms panstaining but no foci (Fig. 5 b and c). This response is consistent with a lack of DNA DSBs either induced by UV irradiation or formed by repairing UV-induced DNA lesions.

H2AX phosphorylation after exposure to ionizing radiation can be inhibited by the kinase inhibitor wortmannin (Fig. 5d). Wortmannin, however, does not affect H2AX phosphorylation after UV irradiation in G1 cells. Therefore, UV-induced H2AX phosphorylation is not performed by the same kinase that phosphorylates H2AX at the sites of DSBs after the exposure to ionizing radiation. This observation, together with the results acquired by immunofluorescence and flow cytometry, indicates that H2AX phosphorylation in G1 cells after UV irradiation is not due to DNA DSB formation.

UV-Induced H2AX Phosphorylation Requires Nucleotide Excision Repair Factors.

UV-induced DNA damage is removed primarily by NER. Mutants defective for either the xeroderma pigmentosum group A (XPA) protein or the xeroderma pigmentosum group C (XPC), which are early damage-recognition proteins, fail to carry out NER (25). Previous research has established that H2AX phosphorylation is not induced as a direct result of UV damage (26). We then tested whether XPA and XPC are required for UV-induced H2AX phosphorylation.

Four hours after 20 J/m2 UV, normal G1 cells show a 2.3-fold increase in γH2AX compared with mock-treated control cells, whereas the UV-induced γH2AX levels in XPC and XPA are only 1.2-fold higher than in mock-treated controls (Fig. 6a). The S phase population in both mutant cell lines was homogenous in its γH2AX phosphorylation pattern, showing only “bright” pan-nuclear staining, whereas the super-bright population seen in irradiated wild-type cells is absent (Fig. 6). Therefore, γH2AX phosphorylation after UV exposure within the G1 phase of the cell cycle largely depends on the nucleotide excision repair factors XPA and XPC. In addition, within S phase, absence of either XPA or XPC reduces the extent of H2AX phosphorylation.

Fig. 6.
Reduced levels of γH2AX after UV irradiation in NER-deficient cell lines. (a) Flow cytometry of γH2AX levels in normal, XP-A, and XP-C cell lines. Overlay of γH2AX levels of unirradiated cells (filled histogram) compared with cells ...

Discussion

Our investigation of H2AX phosphorylation by flow cytometry, in response to UV irradiation, showed that, in human fibroblasts, phosphorylation occurs in all phases of the cell cycle. This observation is in agreement with a report showing H2AX phosphorylation after exposure to UVB by Halicka et al. (ref. 14; Fig. 1).

Our immunofluorescence analysis revealed heterogeneous H2AX staining 4 h after UV exposure (Fig. 3 c and d) that was primarily diffuse, even, and nuclearwide. The lack of foci in most cells after UV irradiation is strikingly different from the bright distinct foci seen after exposure to ionizing radiation. In contrast to ionizing radiation, UV exposure leads to dramatically different levels of H2AX phosphorylation that depends on the phase of the cell cycle. Cells in the G1 phase of the cell cycle show low levels of H2AX phosphorylation in response to UV irradiation. UV exposure of S phase cells yields two distinct levels of γH2AX fluorescence intensity, bright and super-bright cells, which can be distinguished either by flow cytometry (Fig. 1c) or by immunofluorescence microscopy (Fig. 3). The bright cells are found in early and late S phase, whereas the super-bright cells occur throughout S phase but are concentrated in mid-S phase. Halicka et al. (14) also reported the presence of a HL-60 subpopulation 1 h after UVB irradiation with very bright H2AX phosphorylation and identified those as apoptotic cells. In our study, the super-bright UV-irradiated fibroblasts have the same γH2AX intensity as cells treated with the apoptosis inducer staurosporine (Fig. 9a, which is published as supporting information on the PNAS web site). But, 4 h after UV irradiation, these cells did not detach from the surface as staurosporine-treated cells did, and they were not positive for the early apoptosis marker Annexin V (Fig. 9b), nor was any apoptotic laddering present (unpublished data). Our previous results also showed that UV does not induce apoptosis in primary fibroblasts during the time periods we investigated (27). Therefore, it is unlikely that the UV-induced super-bright γH2AX-positive cells are apoptotic, unless γH2AX is a preapoptotic signal associated with early changes in chromatin structure. The precise function of nuclear-wide pan staining during the S phase remains to be determined but may represent nuclearwide conformational changes that expose the histone substrate to kinase activity during replication arrest.

One of our main findings is that UV-induced H2AX phosphorylation in G1 cells cannot be attributed to DNA DSBs. (Fig. 5). Flow cytometry revealed that the total G1 cellular phosphorylation 4 h after 20 J UV irradiation is approximately equal to the total G1 cellular phosphorylation 0.5 h after exposure to 5 Gy ionizing radiation. If the H2AX phosphorylation within G1 cells after UV irradiation was triggered by DNA DSBs, then UV-irradiated cells should display the same number of γH2AX foci as induced by exposure to ionizing radiation. Instead, H2AX phosphorylation in non-S phase cells, as determined by absence of BrdU incorporation, is present throughout the nucleus, and there are few γH2AX foci.

The finding that UV-induced H2AX phosphorylation in not due to DNA DSBs is supported by immunofluorescence microscopy to detect 53BP1 and phospho NBS1(ser343) foci. Our data clearly shows that 53BP1 and phospho NBS1(ser343) form foci that colocalize with γH2AX foci after exposure to ionizing radiation. After UV irradiation, neither 53BP1 nor phospho NBS1(ser343) form a significant number of foci in G1 cells. After exposure to UV radiation, 53BP1 is evenly distributed throughout the nucleus, whereas phospho NBS1 is not evident.

Finally, H2AX phosphorylation after exposure to ionizing radiation can be inhibited by wortmannin, which inhibits ATM and DNA-PKcs during repair of DNA DSB. We show that H2AX phosphorylation in G1 cells after UV irradiation is not affected by wortmannin (28, 29). This insensitivity indicates that H2AX phosphorylation, at least in G1 cells after UV irradiation, is not due to DNA DSBs and requires a distinct kinase.

An unsolved question is the molecular mechanism and function of H2AX phosphorylation after UV irradiation. We have shown that NER mutants are deficient in H2AX phosphorylation after UV irradiation in the G1 phase of the cell cycle. Thus, the process of repairing UV-induced DNA damage is necessary for H2AX phosphorylation in G1 cells and may expose the Ser-139 site to kinase activity.

Given that the temporal pattern of UV-induced H2AX phosphorylation in G1 cells and the repair kinetics of 6-4 photoproducts are very similar, it is possible that they are directly linked. Excision of 6-4 photoproducts from DNA in the spatially restricted regions (Fig. 4) would signal γH2AX formation coincident with persistent CPDs that are more slowly excised. Immediately after UV irradiation, recognition of 6-4 photoproducts triggers cell cycle arrest (30). Recent findings in yeast show that H2AX dephosphorylation is required for checkpoint recovery (31). Therefore, H2AX phosphorylation in G1 cells, which follows the kinetics of 6-4 photoproduct removal after UV irradiation, may play a role in maintaining an active checkpoint during the period of UV lesion repair.

Checkpoint maintenance is necessary to preserve genomic integrity (32), and genomic instability is reported for H2AX-deficient mouse cells under normal growth conditions (9). Thus, the absence of H2AX phosphorylation as a signal to maintain the G1 checkpoint could lead to an increase in premature entry into S phase and genomic instability. In summary, we suggest that a possible function for H2AX phosphorylation after UV irradiation could be to maintain cell cycle checkpoints, which protect the cell from genomic instability.

Materials and Methods

Cell Lines and Culture Conditions.

The following cell lines were used in this study: WT (GM05659+T), XP-A (XP200SF+T), and XP-C (GM02993+T). All cell lines are primary human fibroblasts immortalized by transfection with the human telomerase gene. Cells were cultured in Eagle's minimal essential media with Earle's balanced salt solution, supplemented with 10% fetal bovine serum/10 μg/ml streptomycin/10 units/ml penicillin/0.292 g/liter glutamine/25 μg/ml blasticidin. Cells were maintained between 50% and 90% confluency, passaged 24–48 h before use in experiments, and grown in a 37°C incubator at 5% CO2. For contact inhibition, cells were grown to 100% confluency, and normal media then was replaced with media containing 0.1% fetal bovine serum. The cells were grown for an additional 72 h under the low-serum conditions.

Irradiation and Drug Treatment.

Cells were exposed either to ionizing radiation from a Cs137 source or to UV light (254 nm at a fluency of 2.65 W·m−2). Where indicated, cells were labeled immediately after UV irradiation with 10 μM BrdU for 4 h. For micropore UV irradiation, a 3-μm isopore polycarbonate filter (Millipore) was placed on top of the cell monolayer before irradiation with 100 J/m2 UV light. For experiments involving wortmannin, cells were treated with 100 μM wortmannin 30 min before irradiation and during subsequent incubations.

Flow Cytometry.

Detection of cellular γH2AX was carried out by using the H2AX phosphorylation assay kit for flow cytometry (Upstate Biotechnology, Lake Placid, NY). The assay was carried out according to the manufacturer's instructions with one major modification: the FITC-labeled antibody was incubated overnight at 4°C instead of 20 min on ice. Cells were suspended in flow buffer (Ca2+- and Mg2+-free 1× PBS with 1% BSA containing 10 μg/ml PI and 100 μg/ml RNaseA) and analyzed by using a FACS Caliber Flow Cytometer (Becton Dickinson) equipped with cell quest. Data were analyzed by using both cell quest and flojo software (Tree Star, San Carlos, CA).

Immunofluorescence Microscopy.

Cells were grown on dual-chambered slides (Nalge Nunc), fixed with 4% paraformaldehyde for 10 min, and permeabilized with 0.2% Triton X-100 for 5 min except for slides used for 53BP1 immunofluorescence, which were fixed with 100% methanol for 20 min. Slides were either directly processed or air-dried for 5 min and stored at −80°C. After storage/methanol fixation, slides were repermeabilized with −20°C 50:50 acetone:methanol for 20 min, blocked in PBS/10% FBS/1% BSA for 1 h at 37°C, and incubated overnight at 4°C with the following primary antibodies: mouse monoclonal γH2AX (Ser-139), 1:1,000 (Upstate Biotechnology); rabbit polyclonal γH2AX (Ser-139), 1:1,000 (Novus Biologicals); rabbit polyclonal 53BP1, 1:300 (Cell Signaling Technology, Beverly, MA); rabbit polyclonal NBS1 (p-ser343) (Cell Signaling Technology); mouse monoclonal cyclobutane pyrimidine dimers, 1:5,000 (Medical & Biological Laboratories, Naka-ku Nagoya, Japan); and mouse monoclonal bromodeoxyuridine, 1:500 (BD Biosciences). Secondary antibodies labeled with Alexa 488 or Alexa 555 (Molecular Probes) were added at 1:1,000, and slides were incubated at 37°C for 1 h. Slides were mounted with ProLong Gold Antifade reagent containing DAPI (Molecular Probes). Images were acquired at room temperature with a Zeiss Axioplan 2 microscope equipped with a ×100 ZEISS Plan-NEOFLUAR 1.3 Oil objective lens and a Zeiss Axiocam color camera under the control of axiovision 4.2 software. Image processing with photoshop 6.0 (Adobe Systems, San Jose, CA) was applied to whole images only. Images used for comparison between different treatments and/or cell lines were acquired with the same instrument settings and exposure time and were processed equally.

Supplementary Material

Supporting Figures:

Acknowledgments

We are grateful to the XP Society (Poughkeepsie, NY) and The Luke O'Brien Foundation for their continued support and encouragement for the University of California, San Francisco, research program (J.E.C.). This work was supported by National Institute on Environmental Health Sciences Grant 1 R01 ES 8061 (to J.E.C.), Cancer Center Support Grant P30 CA82103 (principal investigator F. McCormick), and program project Grant P01 AR050440-01 (principal investigator E. H. Epstein). Postdoctoral support was provided by a subcontract from City College San Francisco funded by National Science Foundation Advanced Technology Education Grant 01-52 (to E.H.; principal investigator Dr. V. Natale) and University of California, San Francisco, Dermatology Training Grant T32 AR007175-27. Postdoctoral support was provided by the Swiss National Science Foundation by a Fellowship for Prospective Researchers PBBEA-102308 (to T.M.M.).

Abbreviations

CPD
cyclobutane pyrimidine dimer
DSB
double-strand break
NER
nucleotide excision repair
PI
propidium iodide.

Footnotes

Conflict of interest statement: No conflicts declared.

References

1. Rogakou E. P., Pilch D. R., Orr A. H., Ivanova V. S., Bonner W. M. J. Biol. Chem. 1998;273:5858–5868. [PubMed]
2. Park E. J., Chan D. W., Park J. H., Oettinger M. A., Kwon J. Nucleic Acids Res. 2003;31:6819–6827. [PMC free article] [PubMed]
3. Drouet J., Delteil C., Lefrancois J., Concannon P., Salles B., Calsou P. J. Biol. Chem. 2005;280:7060–7069. [PubMed]
4. Paull T. T., Rogakou E. P., Yamazaki V., Kirchgessner C. U., Gellert M., Bonner W. M. Curr. Biol. 2000;10:886–895. [PubMed]
5. Rogakou E. P., Boon C., Redon C., Bonner W. M. J. Cell Biol. 1999;146:905–916. [PMC free article] [PubMed]
6. Wang H., Wang M., Wang H., Bocker W., Iliakis G. J. Cell Physiol. 2005;202:492–502. [PubMed]
7. Ward I. M., Reina-San-Martin B., Olaru A., Minn K., Tamada K., Lau J. S., Cascalho M., Chen L., Nussenzweig A., Livak F., et al. J. Cell Biol. 2004;165:459–464. [PMC free article] [PubMed]
8. Celeste A., Difilippantonio S., Difilippantonio M. J., Fernandez-Capetillo O., Pilch D. R., Sedelnikova O. A., Eckhaus M., Ried T., Bonner W. M., Nussenzweig A. Cell. 2003;114:371–383. [PubMed]
9. Celeste A., Petersen S., Romanienko P. J., Fernandez-Capetillo O., Chen H. T., Sedelnikova O. A., Reina-San-Martin B., Coppola V., Meffre E., Difilippantonio M. J., et al. Science. 2002;296:922–927. [PubMed]
10. Furuta T., Takemura H., Liao Z. Y., Aune G. J., Redon C., Sedelnikova O. A., Pilch D. R., Rogakou E. P., Celeste A., Chen H. T., et al. J. Biol. Chem. 2003;278:20303–20312. [PubMed]
11. Ward I. M., Chen J. J. Biol. Chem. 2001;276:47759–47762. [PubMed]
12. Limoli C. L., Giedzinski E., Bonner W. M., Cleaver J. E. Proc. Natl. Acad. Sci. USA. 2002;99:233–238. [PMC free article] [PubMed]
13. Limoli C. L., Laposa R., Cleaver J. E. Mutat. Res. 2002;510:121–129. [PubMed]
14. Halicka H. D., Huang X., Traganos F., King M. A., Dai W., Darzynkiewicz Z. Cell Cycle. 2005;4:339–345. [PubMed]
15. Fornace A. J., Jr., Kohn K. W., Kann H. E., Jr. Proc. Natl. Acad. Sci. USA. 1976;73:39–43. [PMC free article] [PubMed]
16. Erixon K., Ahnstrom G. Mutat. Res. 1979;59:257–271. [PubMed]
17. Cleaver J. E., Bodell W. J., Morgan W. F., Zelle B. J. Biol. Chem. 1983;258:9059–9068. [PubMed]
18. Zorn C., Cremer C., Cremer T., Zimmer J. Exp. Cell Res. 1979;124:111–119. [PubMed]
19. Rogakou E. P., Nieves-Neira W., Boon C., Pommier Y., Bonner W. M. J. Biol. Chem. 2000;275:9390–9395. [PubMed]
20. Mirzoeva O. K., Petrini J. H. Mol. Cancer Res. 2003;1:207–218. [PubMed]
21. Kaufmann W. K., Cleaver J. E., Painter R. B. Biochim. Biophys. Acta. 1980;608:191–195. [PubMed]
22. Kaufmann W. K., Cleaver J. E. J. Mol. Biol. 1981;149:171–187. [PubMed]
23. Anderson L., Henderson C., Adachi Y. Mol. Cell. Biol. 2001;21:1719–1729. [PMC free article] [PubMed]
24. Schultz L. B., Chehab N. H., Malikzay A., Halazonetis T. D. J. Cell Biol. 2000;151:1381–1390. [PMC free article] [PubMed]
25. Costa R. M., Chigancas V., Galhardo Rda S., Carvalho H., Menck C. F. Biochimie. 2003;85:1083–1099. [PubMed]
26. Ward I. M., Minn K., Chen J. J. Biol. Chem. 2004;279:9677–9680. [PubMed]
27. Cleaver J. E., Bartholomew J., Char D., Crowley E., Feeney L., Limoli C. L. DNA Repair (Amst.) 2002;1:41–57. [PubMed]
28. Burma S., Chen B. P., Murphy M., Kurimasa A., Chen D. J. J. Biol. Chem. 2001;276:42462–42467. [PubMed]
29. Baumann P., West S. C. Proc. Natl. Acad. Sci. USA. 1998;95:14066–14070. [PMC free article] [PubMed]
30. Giannattasio M., Lazzaro F., Longhese M. P., Plevani P., Muzi-Falconi M. EMBO J. 2004;23:429–438. [PMC free article] [PubMed]
31. Keogh M. C., Kim J. A., Downey M., Fillingham J., Chowdhury D., Harrison J. C., Onishi M., Datta N., Galicia S., Emili A., et al. Nature. 2006;439:406–407.
32. Latif C., Harvey S. H., O'Connell M. J. ScientificWorldJournal. 2001;1:684–702. [PubMed]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...