Eukaryotic cells respond to DNA damage within the S phase by activating an intra-S checkpoint: a response that includes reducing the rate of DNA synthesis. In yeast cells this can occur via checkpoint-dependent inhibition of origin firing and stabilization of ongoing forks, together with a checkpoint-independent slowing of fork movement. In higher eukaryotes, however, the mechanism by which DNA synthesis is reduced is less clear. We have developed strategies based on DNA fiber labeling that allow the quantitative assessment of rates of replication fork movement, origin firing, and fork stalling throughout the genome by examining large numbers of individually labeled replication forks. We show that exposing S phase cells to ionizing radiation induces a transient block to origin firing but does not affect fork rate or fork stalling. Alkylation damage by methyl methane sulfonate causes a slowing of fork movement and a high rate of fork stalling, in addition to inducing a block to new origin firing. Nucleotide depletion by hydroxyurea also reduces replication fork rate and increases stalling; moreover, in contrast to a recent report, we show that hydroxyurea induces a strong block to new origin firing. The DNA fiber labeling strategy provides a powerful new approach to analyze the dynamics of DNA replication in a perturbed S phase. Eukaryotic cells respond to DNA damage within the S phase by activating an intra-S checkpoint: a response that includes reducing the rate of DNA synthesis. In yeast cells this can occur via checkpoint-dependent inhibition of origin firing and stabilization of ongoing forks, together with a checkpoint-independent slowing of fork movement. In higher eukaryotes, however, the mechanism by which DNA synthesis is reduced is less clear. We have developed strategies based on DNA fiber labeling that allow the quantitative assessment of rates of replication fork movement, origin firing, and fork stalling throughout the genome by examining large numbers of individually labeled replication forks. We show that exposing S phase cells to ionizing radiation induces a transient block to origin firing but does not affect fork rate or fork stalling. Alkylation damage by methyl methane sulfonate causes a slowing of fork movement and a high rate of fork stalling, in addition to inducing a block to new origin firing. Nucleotide depletion by hydroxyurea also reduces replication fork rate and increases stalling; moreover, in contrast to a recent report, we show that hydroxyurea induces a strong block to new origin firing. The DNA fiber labeling strategy provides a powerful new approach to analyze the dynamics of DNA replication in a perturbed S phase. Many types of DNA damage can cause mutations in the genome of a cell, not only by direct mutagenesis but also by generating lesions that are processed into mutations when DNA is replicated during S phase. Mechanisms that guard against this include multiple DNA repair systems and also cell cycle checkpoints that coordinate cell cycle progression with the DNA damage response (1Zhou B.B. Elledge S.J. Nature. 2000; 408: 433-439Crossref PubMed Scopus (2628) Google Scholar). One such checkpoint acts within the S phase to reduce the rate of DNA synthesis, presumably minimizing the risk of damage being fixed into potentially dangerous mutations before it can be repaired. The reduction in rates of DNA synthesis in the intra-S checkpoint may be due to any of a combination of parameters: the overall number of active origins, the temporal program of origin firing, the rates of movement of all active forks, and the occurrence of “fork stalling” events. Any or all of these parameters may be affected by DNA damage, either as a direct physical result of DNA lesions or via the action of checkpoint proteins. This issue has been addressed in some detail in the budding yeast Saccharomyces cerevisiae, in which replication from specific origins has been examined after treatment with methyl methane sulfonate (MMS) 1The abbreviations used are: MMS, methyl methane sulfonate; HU, hydroxyurea; IR, ionizing radiation; PIPES, 1,4-piperazinediethanesulfonic acid; PBS, phosphate-buffered saline; Gy, gray(s). 1The abbreviations used are: MMS, methyl methane sulfonate; HU, hydroxyurea; IR, ionizing radiation; PIPES, 1,4-piperazinediethanesulfonic acid; PBS, phosphate-buffered saline; Gy, gray(s). and hydroxyurea (HU) using a combination of Southern blot, two-dimensional gel, and density transfer analyses of replication intermediates. These techniques can separate effects on origin firing from effects on fork rate, at least on a population level, and they have shown that origin firing is blocked in response to MMS or HU (2Tercero J.A. Diffley J.F.X. Nature. 2001; 412: 553-557Crossref PubMed Scopus (557) Google Scholar, 3Shirahige K. Hori Y. Shiraishi K. Yamashita M. Takahashi K. Obuse C. Tsurimoto T. Yoshikawa H. Nature. 1998; 395: 618-621Crossref PubMed Scopus (359) Google Scholar) and that rates of fork movement are also reduced after MMS damage (2Tercero J.A. Diffley J.F.X. Nature. 2001; 412: 553-557Crossref PubMed Scopus (557) Google Scholar). The block to origin firing in yeast depends on the checkpoint kinases Mec1 and Rad53, whereas the reduction in fork rate appears to be independent of these kinases. Mec1 and Rad53 (homologues of human ATM/ATR and Chk2, respectively) are also central to several other aspects of the S phase checkpoint: the induction of a transcriptional program of damage response genes (4Allen J.B. Zhou Z. Siede W. Friedberg E.C. Elledge S.J. Genes Dev. 1994; 8: 2401-2415Crossref PubMed Scopus (343) Google Scholar, 5Aboussekhra A. Vialard J.E. Morrison D.E. de la Torre-Ruiz M.A. Cernakova L. Fabre F. Lowndes N.F. EMBO J. 1996; 15: 3912-3922Crossref PubMed Scopus (109) Google Scholar), the prevention of irreversible fork stalling after MMS damage (2Tercero J.A. Diffley J.F.X. Nature. 2001; 412: 553-557Crossref PubMed Scopus (557) Google Scholar, 6Lopes M. Cotta-Ramusino C. Pellicioli A. Liberi G. Plevani P. Muzi-Falconi M. Newlon C.S. Foiani M. Nature. 2001; 412: 557-561Crossref PubMed Scopus (618) Google Scholar, 7Sogo J.M. Lopes M. Foiani M. Science. 2002; 297: 599-602Crossref PubMed Scopus (675) Google Scholar), and the increase of dNTP levels in the cell after damage (8Chabes A. Georgieva B. Domkin V. Zhao X. Rothstein R. Thelander L. Cell. 2003; 112: 391-401Abstract Full Text Full Text PDF PubMed Scopus (350) Google Scholar). It is not clear, however, whether these additional checkpoint responses actually affect the rate of DNA synthesis. S phase responses to DNA damage have also been examined extensively in human cells. However, by contrast to the techniques described above, the standard assay for an S phase checkpoint response in mammalian cells, the radioresistant DNA synthesis assay, simply measures rates of overall DNA synthesis by pulse labeling a population of cells with tritiated thymidine after DNA damage. Because it only measures bulk synthesis, this assay cannot distinguish effects on origin firing from those on either fork movement or fork stalling. Moreover, it is affected not only by intra-S phase changes to DNA synthesis but also by inhibition of the G1-to-S transition. In addition, to correlate the incorporation of tritiated thymidine with DNA synthesis, it is necessary to assume that the specific activity of the endogenous dNTP pools remains constant. These pools may, however, be affected by changes in the rates of de novo nucleotide synthesis and/or nucleotide salvage after damage. Therefore, in the absence of a good range of efficient, sequence-defined early and late origins in mammalian genomes (which might facilitate the use of the same techniques employed to study S. cerevisiae), various alternative assays have been used to further investigate specific aspects of the mammalian S phase checkpoint. Size separation of 3H-labeled DNA on an alkaline sucrose gradient after treating cells with ionizing radiation (IR) led to the inference that origin firing is blocked because the proportion of small DNA fragments, assumed to represent recently fired origins, is reduced after IR damage (9Painter R.B. Young B.R. Proc. Natl. Acad. Sci. U. S. A. 1980; 77: 7315-7317Crossref PubMed Scopus (720) Google Scholar, 10Painter R.B. Kroc. Found. Ser. 1985; 19: 89-100PubMed Google Scholar). Longer fragments of labeled DNA, assumed to represent ongoing forks, were also found to be reduced but only after much higher doses of IR. A similar block to origin firing was observed after MMS and UV damage, with fork movement again being affected to a lesser extent and only after longer time periods (11Painter R.B. Mutat. Res. 1977; 42: 299-303Crossref PubMed Scopus (44) Google Scholar, 12Painter R.B. Mutat. Res. 1985; 145: 63-69PubMed Google Scholar). The response to both IR and UV was found to be deficient in ATM cells. It is important to note, however, that alternative interpretations of much of these data could be made because it is not possible to tell how the large and small DNA fragments actually originated, and like the radioresistant DNA synthesis method, this assay could be skewed by changes to dNTP levels as well as cell cycle effects outside the S phase. Indeed, a subsequent investigation of 3H labeling of DNA in asynchronous versus synchronized cell populations showed that at least 50% of the reduction in 3H labeling that follows exposure to IR in an asynchronous population was due to the complete prevention of S phase entry via a G1/S checkpoint, as opposed to any intra-S phase change in replication dynamics (13Lee H. Larner J.M. Hamlin J.L. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 526-531Crossref PubMed Scopus (40) Google Scholar). Nevertheless, the existence of a block to origin firing that is genuinely intra-S phase and ATM-dependent has been corroborated by a second method: two-dimensional gel analysis of replication in rDNA (one of the few areas in the mammalian genome showing sequence-defined “early” and “late” replication). This showed, at least qualitatively, that unfired origins could be blocked following IR damage within S phase, whereas fork movement appeared to be minimally affected, at least after moderate IR doses (14Larner J.M. Lee H. Little R.D. Dijkwel P.A. Schildkraut C.L. Hamlin J.L. Nucleic Acids Res. 1999; 27: 803-809Crossref PubMed Scopus (51) Google Scholar). Replication dynamics and their dependence on checkpoint proteins were not tested by the two-dimensional gel method after other forms of DNA damage such as alkylation by MMS. However, an alternative approach has been used to examine origin firing after aphidicolin or HU treatment, drugs that stall replication. This technique, involving the fluorescent labeling of characteristic patterns of “early S” and “late S” foci in Chinese hamster ovary cells, revealed an ATR/Chk1-dependent block to the appearance of late replication patterns when the cells are treated with aphidicolin (15Zachos G. Rainey M.D. Gillespie D.A. EMBO J. 2003; 22: 713-723Crossref PubMed Scopus (222) Google Scholar, 16Dimitrova D.S. Gilbert D.M. Nat. Cell Biol. 2000; 2: 686-694Crossref PubMed Scopus (135) Google Scholar). This was interpreted as a checkpoint-dependent block to origin firing; however, the method does not yield quantitative data on the numbers or proportions of affected origins in the labeled foci, nor can it address other parameters such as fork rate or fork collapse. Finally, all of the techniques described above, including those used to examine replication in yeast, rely on examining replication intermediates in populations of cells. Thus, all such approaches may miss important information that can be obtained by examining individual replication forks. To integrate all these different pieces of information using a single experimental system, a DNA fiber-labeling strategy has been developed in which all the various parameters determining DNA synthesis during the S phase can be assessed individually, on the level of single replication forks as opposed to whole cell populations. This method measures DNA synthesis across the entire genome, independently of sequence or structure; it is quantitative, and the results can be subjected to statistical analysis. The technique has been used in a systematic investigation of both the immediate and longer term changes to replication dynamics, which occur after a variety of DNA-damaging and replication-stalling stimuli. Cell Culture and Synchronization—HeLa cells were grown as monolayers in Dulbecco's modified Eagle's medium +10% fetal calf serum. Synchronization was carried out by adding 0.17 μm nocodazole (from stock solution 3.4 mm in Me2SO). After 4–5 h, rounded mitotic cells were shaken off into prewarmed PHEM buffer (60 mm PIPES, 25 mm HEPES, 10 mm EDTA, 2 mm MgCl2, pH 6.9), collected with minimal centrifugation (∼130 × g for 5 min) and replated in fresh medium. DNA damaging treatments were applied 15–16 h after replating, when the majority of cells were in the early S phase. The experiment in Fig. 4e was carried out in unsynchronized IMR90 cells, also grown in Dulbecco's modified Eagle's medium + 10% fetal calf serum. Flow Cytometry—Cell samples were prepared by trypsinizing, washing in cold PBS, and fixing for at least 2 h in 70% ethanol at 4 °C. The cells were then washed in complete PBS and incubated for 30 min in 0.5 ml of complete PBS containing 40 μg/ml propidium iodide and 0.5 mg/ml RNase A. Flow cytometry was carried out using a BD Biosciences FACScan. DNA Damaging Treatments—MMS (100% solution; Sigma) was added directly to the culture medium at final concentrations of 0.005–0.03% (0.59–3.54 mm). After 20-min treatments the MMS was removed, and cells were washed twice with MMS-free medium before incubating in further fresh medium. IR exposures were carried out at between 1 and 10 Gy (∼25–250 s; control cells were removed from the incubator for the same time period). HU (Sigma) was dissolved in water and added to the culture medium at final concentrations of 20 μm to 2 mm. Replication Labeling and DNA Fiber Spreads—The cells were single-labeled with 50 μm IdU for 10–60 min, or, for double-labeling, 10 μm or 20 μm IdU for 10 min and then 100 μm CldU for 20 min. In the experiments in Fig. 5, the cells were pulsed with 20 μm IdU for 10 min directly before DNA damage, then incubated with 50 μm thymidine for 15 min to wash out the IdU, and then kept in fresh medium before double-labeling 1.5–4.5 h later. DNA spreads were made as described by Jackson and Pombo (18Jackson D.A. Pombo A. J. Cell Biol. 1998; 140: 1285-1295Crossref PubMed Scopus (625) Google Scholar), with certain modifications. Briefly, the cells were trypsinized and resuspended in ice-cold PBS at 2.5 × 105 cells/ml. The labeled cells were diluted 1:8 in unlabeled cells, and 2.5 μl of cells were mixed with 7.5 μl of spreading buffer (0.5% SDS in 200 mm Tris-HCl, pH 7.4, 50 mm EDTA) on a glass slide. After ∼8 min the slides were tilted at ∼15°, and the resulting DNA spreads were air-dried, fixed in 3:1 methanol/acetic acid, and refrigerated overnight. Immunolabeling—The slides were treated with 2.5 m HCl for 1 h, washed several times in PBS, and blocked in 1% bovine serum albumin, 0.1% Tween 20. The slides were then incubated at room temperature with the following antibodies, rinsed three times in PBS, and then washed three times for 20 min in blocking buffer between each incubation: 1) overnight in 1:2000 rat anti-bromodeoxyuridine (detects CldU) (OBT0030F Immunologicals Direct); 2) 2 h in 1:1000 Alexafluor 633-conjugated anti-rat (A-21094 Molecular Probes); 3) 2 h in 1:500 mouse anti-bromodeoxyuridine (detects IdU) (MD5100 Caltag); and 4) 2 h in 1:1000 Cy3-conjugated anti-mouse (C-2181 Sigma). The slides were then counterstained for 20 min with 1:20 000 YOYO-1 in PBS (Molecular Probes) before rinsing three times in PBS and mounting in PBS/glycerol. Microscopy was carried out using a Zeiss LSM Meta 510 confocal microscope. S Phase Progression is Slowed by IR, MMS, and HU—Many techniques used to synchronize cells in S phase, such as aphidicolin, mimosine, or double thymidine blocks, interfere with replication forks and are likely to activate DNA damage responses. Therefore, in this study, HeLa cells were synchronized by nocodazole arrest, mitotic shake-off, and release for 16 h, at which point most cells are in the early S phase. Initially, we used such synchronized cells to examine the effects of various treatments on overall S phase progression. First, the cells were treated with 20-min pulses of 0.001–0.03% MMS, the MMS was removed, and S phase progression was followed by flow cytometry over the next 12 h. Fig. 1a shows that S phase was slowed in a dose-dependent manner, ranging from a mild effect after 0.005% MMS to nearly complete arrest over 12 h after the 0.03% treatment. Second, S phase progression was followed after exposure to 1 or 5 Gy of IR (exposures that should cause ∼36 and 180 double-stranded breaks/cell, respectively (17Rothkamm K. Lobrich M. Proc. Natl. Acad. Sci. U. S. A. 2003; 100: 5057-5062Crossref PubMed Scopus (1315) Google Scholar)). 1 Gy did not cause a detectable slowing of S phase, but 5 Gy resulted in a moderate slowing of S phase progression (Fig. 1b). Third, 5–100 μM HU was added to the cells in the early S phase. Again, a dose-dependent slowing of the S phase was observed (Fig. 1c); 5 μm HU had little effect, 20 μm caused a significant slowing of S phase, and 100 μm lead to arrest with a nearly 2C DNA content. Fork Movement Is Reduced by MMS and HU but Not by IR Damage—The DNA fiber labeling (DIRVISH) technique (18Jackson D.A. Pombo A. J. Cell Biol. 1998; 140: 1285-1295Crossref PubMed Scopus (625) Google Scholar) has been adapted in this study such that two distinguishable modified nucleotides, IdU and CldU (19Aten J.A. Bakker P.J. Stap J. Boschman G.A. Veenhof C.H. Histochem. J. 1992; 24: 251-259Crossref PubMed Scopus (113) Google Scholar), could be used to label replication within a single S phase. In this technique (itself adapted from the classical DNA fiber autoradiography technique (20Huberman J.A. Riggs A.D. J. Mol. Biol. 1968; 32: 327-341Crossref PubMed Scopus (636) Google Scholar) in which newly replicated DNA is labeled with tritiated thymidine), the cells are pulse-labeled with halogenated nucleotides, then collected, and lysed on a glass slide. By tipping the slide, DNA from the cells is spread out in the form of single fibers. This DNA is subsequently fixed, denatured, and immunolabeled to detect the halogenated nucleotides. In these experiments, all of the DNA was then counterstained in a third color with YOYO-1 DNA dye, allowing the exclusion of any broken or tangled fibers. Consecutive pulse labeling of the S phase cells with IdU and then CldU yields double-fluorescently labeled tracks on the DNA that can be interpreted unambiguously as either ongoing forks, newly fired origins, terminations, or fork stalling events (Fig. 2). The length of any track after a given labeling period is proportional to its fork rate, whereas counting the relative numbers of different track forms can determine changes in the rates of origin firing or fork stalling after DNA damage. DNA fiber assays were carried out after each of the three treatments examined in Fig. 1 to establish which parameter(s) of DNA synthesis contributed to the overall slowing of S phase seen by flow cytometry. To quantify any change in fork rates after DNA damage, the cells were exposed to 20-min pulses of MMS (0.005–0.03%) and then, after removal of the MMS, immediately labeled with IdU for 10–60 min before preparing DNA fiber spreads. The mean length of at least 50 IdU-labeled tracks was calculated for each time period. Fig. 3a shows that fork rates were reduced for at least 60 min after more severe MMS treatments. The severity of slowing was correlated with the MMS dose, but slowing was only observed above ∼0.01% MMS. Fig. 3b shows that reduction of cellular dNTP pools by treatment with hydroxyurea also slows replication forks in a dose-dependent manner. When cells were treated with sufficiently high levels of HU (above ∼100 μm), the forks were essentially stalled, and very little progression occurred over several hours (data not shown). By contrast to MMS and HU, IR did not cause detectable fork-slowing, even at doses that do reduce overall S phase progression. Fig. 3c shows no significant change in the mean lengths of tracks labeled after IR exposures of up to 5 Gy. Origin Firing Is Rapidly Inhibited after IR, MMS, or HU— DNA fiber labeling can be used to distinguish newly fired origins from ongoing forks using the experimental protocol outlined in Fig. 4a. Active replication forks prior to damage were labeled with IdU, and the cells were then treated with damage and the IdU was replaced by CldU. During the subsequent 20 min, any newly fired origins will generate tracks labeled along their entire length with CldU, and they can be counted against the number of double-labeled (ongoing) forks that were tagged with IdU prior to damage. Fig. 4b shows that origin firing was inhibited in response to MMS and that the severity of inhibition was dose-dependent over the range tested (20-min pulses of MMS at 0.005–0.02%). Exposure to IR also inhibited origin firing, but unlike the response to MMS, this may show a threshold between 1 and 2.5 Gy (Fig. 4c). No further decrease in origin firing was then seen after IR exposures up to 10 Gy (data not shown). This damage-insensitive subset of initiation events, seen after the maximum doses of both IR and MMS damage, may represent the proportion of the total origins, which are already committed to fire within this 20-min labeling period at the time of damage. The response of cells to HU was also tested in this origin blocking assay, because nucleotide depletion has been shown to inhibit origin firing via the S phase checkpoint in S. cerevisiae (2Tercero J.A. Diffley J.F.X. Nature. 2001; 412: 553-557Crossref PubMed Scopus (557) Google Scholar, 3Shirahige K. Hori Y. Shiraishi K. Yamashita M. Takahashi K. Obuse C. Tsurimoto T. Yoshikawa H. Nature. 1998; 395: 618-621Crossref PubMed Scopus (359) Google Scholar, 21Santocanale C. Diffley J.F.X. Nature. 1998; 395: 615-618Crossref PubMed Scopus (531) Google Scholar). In higher eukaryotes the S phase checkpoint response to HU has not been tested, but aphidicolin, which stalls replication by inhibiting DNA polymerases, does inhibit the appearance of late S phase foci in Chinese hamster ovary cells (16Dimitrova D.S. Gilbert D.M. Nat. Cell Biol. 2000; 2: 686-694Crossref PubMed Scopus (135) Google Scholar). Replication forks were prelabeled for 10 min with IdU as before, and then the IdU was replaced with CldU together with 250 μm HU. The accumulation of new (CldU-labeled) origins was then counted against the IdU-tagged ongoing tracks over the subsequent 2–6 h. Fig. 4d shows that origin firing is greatly reduced, such that it takes 6 h to accumulate the same number of origin firing events that occur in control cells in less than 1 h. It is unlikely that many new origins did fire but were simply not labeled because of nucleotide depletion, because most existing forks were able to progress, incorporating CldU, for a further 1–2 μm over the 6 h of HU arrest. To confirm this, the experiment was repeated using only 50 μm HU, a concentration that allows existing forks to elongate more extensively, growing by 3–4 μm over 3 h. As before, new origin firing was severely inhibited (Fig. 4d). Because of a recent report indicating an increase in origin firing after treatment of a modified hamster fibroblast cell line with HU (22Anglana M. Apiou F. Bensimon A. Debatisse M. Cell. 2003; 114: 385-394Abstract Full Text Full Text PDF PubMed Scopus (273) Google Scholar), this experiment was repeated using primary human fibroblasts instead of HeLa cells, and a similar inhibition of new origin firing was observed (Fig. 4e). Origin Firing Recovers at Different Rates after IR, MMS, and HU—A modified version of the origin-firing assay described above was used to assess recovery in the rate of firing over longer periods after DNA damage (Fig. 5a). As in Fig. 4, active replication forks were tagged with a pulse of IdU prior to DNA damage (Fig. 5a, tracks a), and then the IdU was washed out before MMS or IR were applied. This generates exclusively IdU-labeled (red) tracks representing the number of active replication forks before DNA damage. At time points from 1.5 to 4.5 h later, the cells were then double-labeled with consecutive pulses of IdU (red) and CldU (green). This protocol distinguishes any new origins actually firing at each time point (exclusively green or green at both ends: labeled c in Fig. 5a) from ongoing replication forks (red then green: b in Fig. 5a). These new origins were counted against the exclusively red tracks that form an internal control because they had been tagged identically in all the cells before any DNA damage. Fig. 5b shows that a 20-min pulse of 0.01% MMS (gray bars) elicited a sustained block to origin firing when compared with the levels occurring in undamaged cells (white bars); origin firing recovered to only a very limited extent during at least 4.5 h after the MMS treatment. In comparison, 5 Gy IR (Fig. 5c) caused a much more transient block to origin firing with significant recovery after only 1.5 h. By 3 h post-IR exposure, origin firing had returned to normal levels. The efficiency of origin firing recovery was also assessed after release from an HU arrest. As before, replication forks were prelabeled with IdU and then completely arrested by adding a high level of HU for 1–4 h. Upon release from HU, the IdU was replaced with CldU, and new origins fired within 1 h were counted against the prelabeled tracks. By comparison with either IR or MMS damage, origin firing recovered relatively well after a brief (1 h) HU arrest, but recovery became progressively less efficient after longer periods (2–4 h) (Fig. 5e). This is unlikely to be an artifact because of under-detection of CldU-labeled tracks after HU release, because the nucleotide balance within the cells recovered sufficiently fast to allow the origins that did fire to elongate by ∼3 μm within 30 min and 6 μm within 60 min (data not shown). Replication Forks Stall at an Elevated Rate after MMS and HU but Not after IR—The slowing of replication forks after MMS damage, which was documented in Fig. 3, could result from at least two distinct modes of altered fork progression. DNA damage may provoke a pan-nuclear change to a slower mode of replication, for example, by modification of all replication forks or a change to a different polymerase. Alternatively, there could simply be a series of transient stalling events at each fork in isolation as it encounters successive DNA lesions. If such fork stalling does occur within the time frame of a double-labeling experiment (Fig. 4a), it should be detectable in the form of IdU-labeled tracks, which fail to incorporate the subsequent 20-min pulse of CldU because they are currently stalled. These events will therefore appear as an elevated number of red-only tracks (Fig. 2). When the percentage of these red-only tracks was counted, a significant level of fork stalling was indeed found after higher MMS treatments (Fig. 6a), supporting the hypothesis that fork slowing occurs via stochastic stalling events. By contrast, IR did not cause significant fork stalling, consistent with the lack of overall fork slowing after IR damage (Fig. 6b). In the case of HU treatment, all forks are essentially stalled by sufficiently high levels of HU. In lower levels of HU, however, replication does proceed at reduced speed (Fig. 3b), and in this situation there is elevated fork stalling, detectable in as little as 5 μm HU and increasing in a dose-dependent fashion to very high levels when S phase cells are subjected to 20 or 50 μm HU (Fig. 6c). This work comprises the first systematic investigation of all the various parameters that determine the rate of DNA synthesis in mammalian cells during S phase and the ways in which these parameters are affected by DNA damage. The fiber labeling technique developed here is an improvement on other methods that have been used to investigate S phase checkpoint responses because it unambiguously separates changes in the rate of origin firing from changes in the rates of fork movement and fork stalling. Using this technique, each of these parameters can be examined quantitatively and under comparable conditions, using the same experimental method throughout. (Labeling cells with short pulses of modified nucleotides does not in itself perturb the S phase (23Hamlin J.L. Exp. Cell Res. 1978; 112: 225-232Crossref PubMed Scopus (21) Google Scholar) or activate the S phase checkpoint in yeast (24Vernis L. Piskur J. Diffley J.F.X. Nucleic Acids Res. 2003; 31: e120Crossref PubMed Scopus (46) Google Scholar), so the technique should measure only changes in DNA synthesis that are induced by IR, MMS, or HU.) Fiber labeling also offers the advantage of revealing replication dynamics on the level of individual forks rather than as an average of an entire cell population. It does not allow any analysis of replicon clustering in relation to higher order chromatin or nuclear structure, but it does allow subtle yet potentially important effects on a minority of individual forks to be detected and quantified. Effects of IR, MMS, and HU on Replication Dynamics: Mammalian Cells Compared with S. cerevisiae—This study shows that different forms of DNA damage affect replication in different ways. Moderate levels of ionizing radiation, alkylation by MMS, or nucleotide depletion by HU can all slow down the overall progression of S phase. In the case of IR, this slowing appears to be entirely due to a rapid but fairly transient block to origin
The DNA damage response (DDR) checkpoint is activated when DNA is damaged or when DNA replication forks stall. The DDR checkpoint plays a critical role in preserving the integrity of stalled replication forks; this is essential for subsequent fork resumption, faithful and complete genome replication, and cell survival. The mechanisms by which the DDR checkpoint preserves stalled replication forks are still incompletely understood. Many substrates of the DDR checkpoint kinases have been identified over the years, but in many cases the functional consequences of phosphorylation are still unclear. Emerging as a complementary approach, recent advances in biochemical reconstitution of DNA replication have made it possible to characterise specific mechanisms of DNA replication regulation by the DDR checkpoint. In this review, we discuss the role of DNA replication in the activation of the DDR checkpoint and how this checkpoint regulates different aspects of DNA replication. We then distinguish between checkpoint action locally at the site of replication stalling and more globally, and we discuss how these functions contribute to coordinating complete replication of the genome in the face of replication stress.
The yeast ARS binding factor 1 (ABF1)—where ARS is an autonomously replicating sequence—and repressor/activator protein 1 (RAP1) have been implicated in DNA replication, transcriptional activation, and transcriptional silencing. The ABF1 gene was cloned and sequenced and shown to be essential for viability. The predicted amino acid sequence contains a novel sequence motif related to the zinc finger, and the ABF1 protein requires zinc and unmodified cysteine residues for sequence-specific DNA binding. Interestingly, ABF1 is extensively related to its counterpart, RAP1, and both proteins share a region of similarity with SAN1, a suppressor of certain SIR4 mutations, suggesting that this region may be involved in mediating SIR function at the silent mating type loci.
Getting loaded—make mine a double! Chromosomal DNA replication initiates bidirectionally by loading two ring-shaped helicases onto DNA in opposite orientations. How this symmetry is achieved has been puzzling because replication initiation sites contain only one essential binding site for the initiator, the origin recognition complex (ORC). Coster and Diffley now show that both helicases are loaded by a similar mechanism. Efficient loading requires binding of two ORC complexes to two ORC binding sites in opposite orientations. Natural origins were found to be partially symmetrical, containing functionally relevant secondary ORC sites. Sites can be flexibly spaced, but introducing an intervening “roadblock” prevented loading, suggesting that individual helicases translocate toward each other on DNA to form a stable double ring. Science , this issue p. 314
The coronavirus disease 2019 (COVID-19) pandemic, which is caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), is a global public health challenge. While the efficacy of vaccines against emerging and future virus variants remains unclear, there is a need for therapeutics. Repurposing existing drugs represents a promising and potentially rapid opportunity to find novel antivirals against SARS-CoV-2. The virus encodes at least nine enzymatic activities that are potential drug targets. Here, we have expressed, purified and developed enzymatic assays for SARS-CoV-2 nsp13 helicase, a viral replication protein that is essential for the coronavirus life cycle. We screened a custom chemical library of over 5000 previously characterized pharmaceuticals for nsp13 inhibitors using a fluorescence resonance energy transfer-based high-throughput screening approach. From this, we have identified FPA-124 and several suramin-related compounds as novel inhibitors of nsp13 helicase activity in vitro. We describe the efficacy of these drugs using assays we developed to monitor SARS-CoV-2 growth in Vero E6 cells.
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