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and 
during S phase
1 Biochemistry Group, Leibniz Institute for Age Research–Fritz-Lipmann-Institute e. V., Beutenbergstraße 11, D-07745 Jena, Germany
2 GSF, Institute of Immunology, Marchioninistr. 25, D-81377 München, Germany
3 National University of Ireland Galway, Department of Biochemistry, Galway, Ireland
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and 
, with Psf2, which is a component of the GINS complex as well as with Mcm5 and 7, subunits of the putative replicative DNA helicase complex. Hence, Cdc45 may play an important role in elongation of DNA replication by bridging the processive DNA polymerases and with the replicative helicase in the elongating machinery. |
Introduction |
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-primase (pol ) (Nasheuer et al. 2002). Mcm2–7 is a hetero-hexameric complex that belongs to the AAA ATPase helicase family. Most of the Mcm subunits in a cell are associated to the Mcm2–7 hetero-hexamer with a 1 : 1 : 1 : 1 : 1 : 1 stoichiometry, although there are likely to be single Mcms and Mcm sub-complexes (for review see Forsburg 2004 and references therein). Mcm4, 6 and 7 subunits bind tightly together to form a trimeric complex. Human Mcm2 is loosely associated to the Mcm4, 6, 7 sub-complex (Musahl et al. 1995). Mcm3 and Mcm5 as well as Mcm3 and Mcm7 together form dimers. The Mcm3/Mcm5 dimer binds weakly to the other MCMs, probably through Mcm7 (Paul et al. 1996).
Cdc45 has a critical function in the initiation and elongation of DNA replication. Chromatin association of Cdc45 requires the converging action of Cdks and DDK (Mimura & Takisawa 1998; Zou & Stillman 2000) leading to the concept that this protein represents the key regulator for the initiation of DNA replication (see, e.g., Dolan et al. 2004). However, further studies in yeast and Xenopus showed that Cdc45, in addition to its role in initiation (Hopwood & Dalton 1996; Mimura et al. 2000; Tercero et al. 2000), is also required for the elongation step of DNA replication (Tercero et al. 2000). Origin association of Cdc45 corresponds well to origin activation, whereas Mcm proteins associate with origins prior to their final activation (Aparicio et al. 1999; Masuda et al. 2003). Yeast Cdc45 physically interacts with Mcm proteins (Hopwood & Dalton 1996; Sheu & Stillman 2006), and an interaction between human Cdc45 and Mcm7 has been identified in vitro (Kukimoto et al. 1999). Moreover, it has been suggested that Cdc45 has a role as a processivity factor for the Mcm helicase (Masuda et al. 2003; Pacek & Walter 2004). As Cdc45 is significantly less abundant than the Mcm proteins this implies that Cdc45 may be a limiting factor in the initiation of DNA replication (Edwards et al. 2002).
Eukaryotic DNA replication is carried out by the three essential DNA polymerases: pol , pol and pol . Pol is the only enzyme that can start DNA replication de novo. Pol consists of four subunits designated as p180, p68, p58 and p48 (Nasheuer et al. 2002). By a process called DNA polymerase switch, pol and pol elongate the pol -initiated DNA strands. Pol (p261, p59, p17 and p12) and pol (p125, p66, p50 and p12) also comprise four subunits each that are conserved from yeast to man. Pol and pol are essential in yeast and depletion of either of them reduces the synthesis rate during Xenopus DNA replication (Fukui et al. 2004 and references therein). Several groups have shown that pol and interact with Cdc45. Besides determining pol as a physical binding partner of Cdc45 (Kukimoto et al. 1999), fission yeast studies revealed that a deletion of the smallest subunit of pol , dpb4, was synthetically lethal with cdc45 (Spiga & D’Urso 2004). Genome-wide analyses of budding yeast also showed that the deletion of either pol subunits dpb3 or dpb4 was synthetically lethal with a cdc45 mutation (Tong et al. 2004). The genetic interaction between Cdc45 and pol supports the earlier finding that the loading of Xenopus pol requires the presence of Cdc45 (Mimura et al. 2000).
The hetero-tetrameric protein complex GINS consists of Sld5, Psf1, Psf2 and Psf3 (Kubota et al. 2003; Takayama et al. 2003). This complex is well conserved from yeast to humans (Takayama et al. 2003) and it has an important role in cell viability and DNA replication (for review see Aparicio et al. 2006). Several groups have reported that S. cerevisiae and Xenopus Cdc45, Mcm2–7 and GINS form a large complex after initiation of DNA replication (Gambus et al. 2006; Kanemaki & Labib 2006; Moyer et al. 2006). In the absence of Psf2, Mcm2–7 and Cdc45 cannot associate during S phase (Gambus et al. 2006). Recent data from budding yeast and X. laevis indicate that GINS and Cdc45 also interact with pol . However, the GINS complex, but not Cdc45 stimulates the DNA polymerase activity of pol
in vitro (Shikata et al. 2006). Inhibition of DNA replication with aphidicolin causes physical uncoupling of the Cdc45/MCM2–7/GINS complex from the DNA polymerases and the sites of DNA synthesis in Xenopus egg extracts (Pacek et al. 2006).
Despite these genetic and biochemical evidences, the function of Cdc45 is still illusive. Particularly the functions of the human protein are only partially understood. In order to analyze functions of human Cdc45 during the cell cycle, we established a new monoclonal antibody against human Cdc45 based on a cDNA coding for human Cdc45 L (Saha et al. 1998). This antibody was used to analyze the intracellular localization and physical interactions of human Cdc45 with other replication factors.
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In order to analyze the function of human Cdc45, hybridoma cell lines secreting antibodies against this protein were established. The clone C45-3G10 produced an antibody that can be used in Western blotting, immunoprecipitation and immunofluorescence. In Western blots this antibody recognized the purified recombinant His-tagged Cdc45 (Fig. 1A, right part) as well as a single band with the expected molecular mass in whole cell extracts (Fig. 1A, left part). On the same Western blot, the purified recombinant His-tagged protein ran slightly slower than the endogenous Cdc45. Immunoprecipitations of either the antibody alone or the antibody with whole cell extract are shown in Fig. 1B. The C45-3G10 antibody precipitated a single Cdc45 protein band. In contrast, the meanwhile available polyclonal antibody sc-20 686 from Santa Cruz detected multiple bands in Western blotting and immunoprecipitated human Cdc45 (data not shown).
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To improve our understanding of the intracellular localization of human Cdc45 and its interaction partners, its chromatin binding was analyzed by Western blotting and immunofluorescence microscopy. We first examined to find whether it is a component of the pre-RC. To this end, HeLaS3 cells were treated with mimosine and arrested at the G1/S border (Fig. 2A, Mimo 0 h). In addition, cells were synchronized with thymidine in S phase for analyzing proteins and their chromatin association when pre-RCs were converted into ICs (Fig. 2A, TdR 3 h). To investigate cells in G2, a thymidine/nocodacole treatment (TN) was performed as described in Experimental procedures (Fig. 2A, TN 0 h and TN 2 h). This led to cell populations that were enriched in late S/G2 and G2/M stages, respectively. Synchronized cells were homogenized and the extracts were fractionated into a detergent-soluble, a DNase I-soluble, and a high salt-soluble fraction. The detergent-soluble fraction 1 contained cyto- and nucleoplasmic proteins and maybe replication factors that were only loosely chromatin-bound. Fraction 2 represented proteins that became soluble after DNase I treatment, that is, proteins that were less easily extracted from DNA, such as clamp forming proteins and associated polypeptides. The remaining high salt fraction 3 consists of solubilized proteins that bound tightly to chromatin and/or to the nuclear matrix.
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Association of Cdc45 with the elongating DNA polymerases and in S phase
These findings suggested an involvement of Cdc45 at the replication fork and so we started to investigate potential interactions between Cdc45 and the DNA polymerases , and in human cells (for detailed analysis of the DNA polymerase subunits in Western blots and immunofluorescence see Supplementary Figs S2 and S3). To investigate possible binding partners of human Cdc45, co-localization studies of different subunits of the replicative DNA polymerases and Cdc45 were performed. Cdc45 (red) and the DNA polymerase subunits (green) displayed a spot-like appearance in S phase (Fig. 3A). Using the co-localization analysis tool of Zeiss (software LSM 510 META 3.2) to extract only those dots that had green and red portions, we could generate a new sub-figure indicated as "mask." In this derived figure the black dots represent the co-localized spots of two proteins in the indicated immunofluorescent data set. The overlapping signals (mask) indicated that Cdc45 co-localized with the pol subunits p180 and p68 as well as with the pol subunit p261 (Fig. 3A). Unfortunately, it was not possible to investigate co-localization with pol because the antibodies against Cdc45 and pol were both elicited in the same species. However, the results indicate a close proximity of Cdc45 with the replicative DNA polymerases.
To further study possible interactions of human Cdc45 protein and the DNA polymerase subunits, immunoprecipitations with Cdc45 antibodies and the same fractionated extracts as in Fig. 2 were performed (Fig. 3B, C). Cdc45 was extracted from G1, S, S/G2 and G2/M cells. Chromatin-associated Cdc45 appeared in the high salt-soluble fraction (fraction 3) of S, S/G2 and G2/M cells. Only in S phase, Cdc45 was able to pull down the catalytic subunits of pol and pol (Fig. 3B). Interestingly, in contrast to pol , Cdc45 was also associated with pol in late S/G2 (Fig. 3C). Moreover, both Cdc45–DNA polymerase complexes behaved differently. The interaction of the pol –Cdc45 complex with chromatin was weaker than that of the Cdc45–pol complex, since the co-precipitation of Cdc45 and pol was already observed in the detergent-solubilized fraction (fraction 1), whereas the binding between Cdc45 and pol only became apparent in the high salt-soluble fraction (fraction 3). In G2/M extracts no interactions between Cdc45 and pol or pol were visible (Fig. 3C). Since the smaller subunits of pol and pol have similar molecular masses as the antibody heavy chains, we were unable to get conclusive results for these subunits. Despite of a co-localization of Cdc45 with PCNA (Supplementary Fig. S1b) and an interaction of Cdc45 with pol we were not able to co-immunoprecipitate Cdc45 and PCNA (data not shown), which is believed to be a processivity factor for pol . Also, it was not possible to obtain a positive result for the immunoprecipitation of Cdc45 with any of the pol subunits. This outcome may have been due to the transient nature of the interaction of Cdc45 with pol as suggested previously (Zou & Stillman 2000).
Complex formation of Cdc45 with pol was verified by immunoprecipitation with an antibody against pol . In extracts from S phase cells pol was immunoprecipitated from the detergent-soluble, the DNase I-soluble and the high salt-soluble fraction (Fig. 4). However, an interaction of the catalytically active p261 subunit with both Mcm7 and Cdc45 was only detected in the high salt-soluble fraction. Taken together, these results suggest that a complex containing pol , Cdc45 and Mcm7 exists in human cells. Furthermore, Cdc45 interacted with pol , although this complex was more easily extractable than the pol –Cdc45 complex.
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Our investigations pointed to the direction that human Cdc45 not only interacts with DNA polymerases (Fig. 3) but also with the Mcm subunits (Fig. 4). To test whether Cdc45 interacts with the Mcm2–7 complex in human cells, immunoprecipitations and immunofluorescence analyses were performed. Cells were synchronized in S phase using a thymidine double block followed by a 2 h release into fresh medium before cells were harvested (for details see Experimental procedures). Initial results using the extraction procedure as in the previous experiments revealed that Cdc45 co-immunoprecipitated Mcm7, but Mcm2 was not associated with Cdc45 (data not shown). However, this failure to detect an interaction of Cdc45 and Mcm2 could be due to relatively high non-ionic detergent concentrations in the extraction condition. Therefore, extracts were prepared using a slightly modified method to that described previously (Mendez & Stillman 2000). We now started with the elution of free protein (FP) from the cytoplasm and the easily disrupted nuclei of HeLaS3 cells by hypotonic extraction conditions. Then, the loosely-bound chromatin proteins were first eluted and this fraction was named chromatin fraction 1 (C1). The subsequently obtained DS fraction (DNaseI/ salt extraction) represented proteins that became soluble after DNase I treatment, for instance clamp forming proteins and associated polypeptides, and proteins eluted by low salt. The remaining tightly-bound proteins from chromatin and/or from the nuclear matrix were consecutively extracted by high-salt treatment (fractions 1 M and 2 M, respectively).
Interestingly a substantial amount of Cdc45 was found to be chromatin-associated with this more gentle extraction method (compare Figs 2B and 5A). Furthermore, under this condition it was possible to extract significant amounts of Cdc45 from chromatin by DNase I/250 mM salt treatment (Fig. 5A; DS fraction). Strikingly, there was still a considerable amount of Cdc45 on the chromatin that was extracted by high salt (fractions 1 M and 2 M). In parallel, the behavior of Mcm proteins was investigated. The Mcm2 protein mainly appeared in the first two fractions as free and easily extractable protein. Only minor amounts of Mcm2 remained in the DS, 1 M and 2 M fractions (Fig. 5A). Mcm4 showed similar properties as Mcm2 by appearing only in the first two fractions. It was not possible to obtain signals for Mcm4 in any of the last three fractions. This was maybe due to weak antigen detection by the Mcm4 antibody but in fractions FP and C1 the Mcm4-recognizing antibody produced a similar or more intense signal than those for Mcm5, Mcm6 and Mcm7, which could be found in all five fractions. In contrast to Mcm4, small amounts of Mcm6 and Mcm7 were detectable in fractions 1 M and 2 M (Fig. 5A). These findings indicate that most of the Mcm proteins were already extracted after DNase I/250 mM salt treatment.
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The treatment with TritonX-100 and SDS was a critical preparation step as otherwise it was not possible to investigate chromatin-bound Mcm proteins, in accordance with previous findings (Krude et al. 1996). In S-phase cells the Mcm signals (green) and the Cdc45 signals (red) were punctated, which also is depicted in the fluorescence intensity diagrams drawn along the white arrows (software LSM 510 META 3.2). The signals from Cdc45 and Mcm7 co-localized well, whereas only a small portion of the Mcm2 and Cdc45 signals were in close proximity as can be seen in both the mask picture and the fluorescence diagrams of Fig. 5B. These results show that during S phase human Cdc45 was in close proximity to chromatin-bound Mcm7 and, to a lesser extent, to Mcm2. The different behavior of Mcm2 and Mcm7 in these experiments motivated us to investigate the two Mcm proteins and their co-localization pattern. Thymidine-synchronized HeLaS3 cells were treated and analyzed as described before. Mcm2 (green) and Mcm7 (red) displayed speckled patterns in S phase (Fig. 5C) and co-localized to a higher extent than Cdc45 and Mcm7.
To determine if there are physical interactions between Cdc45 and Mcm proteins during S phase, co-immunoprecipitations were performed. Immunoprecipitated Cdc45 was detected in all fractions. While Cdc45 did not co-immunoprecipitate Mcm2 in any case, Cdc45 was associated with Mcm5 and Mcm7 (Fig. 5D). These findings do not exclude that Cdc45 may also form a complex containing Mcm2 or physically bind to Mcm2, but under the investigated conditions such an interaction was not detectable in human cell extracts. We were not able to analyze Mcm6 as a possible binding partner of Cdc45 because the Mcm6 signal appeared in the same region as incomplete reduced antibody bands of the rat monoclonal antibody against Cdc45 (data not shown). Moreover, Cdc45 and Mcm5 were also co-immunoprecipitated with the Mcm7 antibody (Fig. 5D).
Association of Cdc45 and Psf2, a subunit of the GINS complex
In Xenopus, the GINS complex is associated with the Mcm2–7 complex, Cdc45 and pol during S phase (Shikata et al. 2006). We wanted to verify this association in human cells and therefore tested if it was possible to co-precipitate Psf2 with Cdc45, as we had previously observed an interaction of human Cdc45 with the DNA polymerases and , and with subunits of the Mcm2–7 complex. Immunoprecipitations were performed using the same extracts as described for Fig. 5 and Cdc45 was precipitated from all investigated fractions (Fig. 6). A faint but reproducible signal for Psf2 appeared as co-precipitated protein in the C1 fraction (Fig. 6), suggesting that human Cdc45 interacts with the GINS complex. Thus, the human Cdc45 is also part of a large replisomal protein complex (see Fig. 7) comparable to that of the yeast and the Xenopus system.
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Differential extraction procedures revealed that the extraction of human Cdc45 became detergent- and DNase I-resistant in S phase, where salt extractions were necessary to fully dissociate Cdc45 from chromatin (Fig. 2) consistent with an earlier study (Saha et al. 1998). In accordance with the discussed function of Cdc45 as triggering factor for the pre-RC to IC conversion (Mimura & Takisawa 1998; Mimura et al. 2000), human Cdc45 was not tightly bound to chromatin, even in late G1. However, it became firmly associated with chromatin during S and G2 phases of the cell cycle, as determined by Western blotting (Fig. 2) and immunoprecipitation (Fig. 3). In contrast to Cdc45, Mcm7, a known component of the pre-RC, was tightly associated with chromatin in G1. This led us to conclude that the human Cdc45 protein is not a component of the pre-RC consistent with findings from yeast and Xenopus (Aparicio et al. 1997; Mimura et al. 2000). Our data support the hypothesis that Cdc45 may act as a key regulator of origin firing at the conversion step of the pre-RC to the IC (Mimura & Takisawa 1998; Mimura et al. 2000). Moreover, Cdc45 stayed in the chromatin-bound fractions in G2. This extended chromatin association of Cdc45, which was also observed in Xenopus (Pacek & Walter 2004), indicates a slow disassembly of Cdc45-containing protein complexes at the end of S phase and in G2. Alternatively, Cdc45 may help in surveying the genome for incompletely replicated regions, and/or may assist DNA polymerases in finishing duplication of cellular chromosomal DNA. The latter view was supported by findings that Cdc45 was still associated with pol in late S/G2.
In contrast to yeast, Xenopus and Drosophila (Loebel et al. 2000; Mimura et al. 2000; Zou & Stillman 2000; Moyer et al. 2006; Sheu & Stillman 2006) protein–protein interactions of human Cdc45 beyond that of recombinant proteins, such as Mcm7 and the p68 subunit of pol (Kukimoto et al. 1999), have not been reported yet. In immunofluorescence microscopy experiments we observed a close proximity between Cdc45 and the two largest subunits of pol , that is, p180 and p68 (Fig. 3). Despite this, we were not able to co-precipitate p180 and/or p68 with Cdc45, and vice versa (data not shown). This may be due to the weak and transient interaction of Cdc45 with pol as discussed previously (Zou & Stillman 2000). However, the published data of an interaction of pol and Cdc45 are controversial (Mimura & Takisawa 1998; Mimura et al. 2000; Zou & Stillman 2000; Uchiyama et al. 2001) and recent findings revealed that Mcm10 rather than Cdc45 facilitates the loading of pol into replication origins, at least in yeast (Ricke & Bielinsky 2004).
In contrast to the interaction studies with pol , Cdc45 associated with the elongating DNA polymerases and in human cells. Binding of pol to Cdc45, which was detected in both S phase and late-S-to-early-G2 phase cells, that is, in cells that had completed or were in the process of completing DNA replication (Fig. 3C), has not yet been described. Despite a detected unspecific adsorption of pol to the immunoprecipitation matrix the signal for pol in the co-immunoprecipitation reaction (fraction 1) was stronger and suggests that pol is still bound to Cdc45 in late S/early G2 phase. The timing of this association suggests a function of the Cdc45–pol complex late in DNA replication, such as the processing of Okazaki fragments or signaling of late replication events. Interestingly, both Cdc45–DNA polymerase complexes behaved differently, since Cdc45–pol was more readily extracted and appeared in fraction 1, whereas the Cdc45–pol complex appeared in fraction 3 after treatment with high-salt. This elution behavior of pol is consistent with results from other groups showing that a significant amount of pol can only be eluted with very stringent conditions, which require high salt or high concentrations of detergent (Rytkönen et al. 2006 and H. Pospiech, personal communication). These results indicate that leading and lagging strand replicases are differentially extracted from cells. Nevertheless these results need to be treated with caution since interactions of pol and Cdc45 determined in extracts eluted with high salt might not reflect physiological conditions. However, Cdc45–pol interactions have already been observed genetically in the yeasts S. pombe and S. cerevisiae (Zou & Stillman 2000; Spiga & D’Urso 2004), and recent publications reported a physical interaction of pol , GINS and Cdc45 in Xenopus (Shikata et al. 2006). We show here that in human cells the binding of pol and Cdc45 only appeared in S phase implying a modulation of both proteins in DNA replication (Fig. 3). Furthermore, human Cdc45 interacts with Psf2, a component of the GINS complex, during S phase (Fig. 6) which is consistent with observations in S. cerevisiae, D. melanogaster and Xenopus, where interactions of Cdc45, GINS and the Mcm2–7 complex were observed (Gambus et al. 2006; Moyer et al. 2006; Pacek et al. 2006; Sheu & Stillman 2006).
The Mcm2–7 complex comprises the putative eukaryotic replicative DNA helicase. Several sub-complexes have been reported: Mcm4, 6, and 7 with DNA helicase activity in vitro, Mcm3/5 and Mcm2 that binds loosely to the Mcm4/6/7 sub-complex (for review see Forsburg 2004 and references therein). By using immunofluorescence microscopy we observed a partial co-localization of Cdc45 and Mcm2 and a more obvious overlap of Cdc45 signals with Mcm7. The extraction of unbound replication protein during fixation (Krude et al. 1996) turned out to be indispensable for Mcm investigations by immunofluorescence. As described earlier it is possible to visualize Mcm proteins at the replication fork, and Mcm7 was found in replication foci from Drosophila (Claycomb et al. 2002). Despite earlier difficulties to detect the Mcm subunits at sites of ongoing replication in immunofluorescence the Mcm2–7 complex was associated with stalled replication forks in Xenopus (Pacek et al. 2006) and it was concluded that the Mcm2–7 complex travels with the replication fork. Binding of human subunits of the Mcm2–7 complex to Cdc45 was investigated by immunoprecipitation using extracts of S phase cells. We observed interactions of human Mcm5 and Mcm7 with Cdc45 (Fig. 5). Interestingly, in accordance with results from S. pombe (Dolan et al. 2004) and Mus musculus (Kneissl et al. 2003), we did not observe a stable complex formation of Mcm2 and Cdc45 in human cells. This is consistent with earlier findings of human Mcm2–7, where only a loose binding of Mcm2 to the other Mcm subunits was observed (Musahl et al. 1995). Our Western blotting results (Fig. 5A) revealed that human Mcm2 was mainly extracted as "free protein" (FP fraction) and with an EDTA containing buffer (fraction C1; chromatin-bound and easily extractable proteins), but hardly any Mcm2 could be detected in the following fractions after these gentle treatments. In contrast to our findings, Sheu and Stillman recently detected all subunits of the Mcm2–7 complex to be associated with Cdc45 which could be due to a higher stability of the S. cerevisiae MCM complex than the human protein complex or due to their less stringent immunoprecipitation conditions (Sheu & Stillman 2006).
In summary, we showed that the human Cdc45 protein interacts with Mcm5, a component of the stable extractable Mcm3/Mcm5 sub-complex of the Mcm complex as well as with Mcm7, a subunit of the trimeric sub-complex Mcm4, 6 and 7. Furthermore, we observed interactions of human Cdc45 with Psf2, a component of the GINS complex, and with the replicative DNA polymerases and . The binding of Cdc45 to the Mcm complex, Psf2 and to the elongating DNA polymerases may facilitate an increased processive movement of the replication fork (summarized in Fig. 7). We conclude that human Cdc45 is part of the elongation complex, as a protein that forms a hinge between the replicative DNA helicases and the elongating DNA polymerases.
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Experimental procedures |
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The indicated proteins were analyzed using the following antibodies: anti-Mcm4–H-300, anti-Mcm7–141.2, anti-Cdc45–sc-20 685, anti-PCNA–pc10 (all Santa Cruz); anti-Mcm2, 4, 5 and 6 rabbit antisera (kindly provided by R. Knippers); anti-Psf2, anti-CENP-F–ab5 (both abcam); anti-Mcm2–BM28, anti-pol p261–pol catalytic subunit clone 34 (both BD Biosciences); anti-pol p125–PDG-5G1, anti-pol p50–PDK-7B4, anti-pol p180–2CT25 (Zlotkin et al. 1996), anti-pol p68–anti-mp68, anti-BrdU–CBL187, goat/donkey anti-rat/rabbit/mouse conjugated with Cy2/Cy3/Cy5, goat anti-rat conjugated with HRP (all Dianova), goat anti-mouse/rabbit conjugated with HRP (Promega) and rabbit anti-chicken HRP (Sigma). To analyze human Cdc45 a new monoclonal rat antibody (C45-3G10) was generated. Recombinant human Cdc45 protein was expressed in E. coli BL21 as a His6 tag fusion protein from pRSET-C that contained the cDNA of Cdc45 cloned between BamHI and XhoI (Saha et al. 1998, kindly provided by A. Dutta). The purified protein was injected into rats. Hybridoma cell lines producing monoclonal antibodies against Cdc45 were established according to standard procedures.
Cell culture and synchronization
HeLaS3 cells (ATCC CCL 2.2) were grown in DMEM with 10% fetal calf serum (FCS) in a 10% CO2 incubator at 37 °C. Synchronization in G1 phase (Mimo 0 h) was performed for 16 h using L-mimosine (Sigma) at a final concentration of 0.3 mM (adapted from Krude 1999). Cell cycle arrest at the G1/S transition was achieved by two subsequent thymidine (TdR, Sigma) blocks (5 mM thymidine) for 16 h, separated by a 10 h growth period without TdR. To investigate active replication forks cells were treated with thymidine as described before and then released into fresh medium for 2 or 3 h (TdR 2 h or TdR 3 h as indicated). To synchronize HeLaS3 cells in late S/G2 and in G2/M, cells were pre-synchronized by a single TdR block. Six hours after TdR release, nocodazole (Sigma) was added to a final concentration of 40 ng/mL for 3 h. Cells were directly harvested (TN 0 h) or released into fresh medium and allowed to proceed into G2/M by incubation for 2 h (TN 2 h).
The cell cycle status of the population was analyzed with an EPICS XL MCL flow cytometer (Beckmann Coulter) by measuring the fluorescence of cells stained with propidium iodide (Sigma). The synchronized cells were routinely tested for signs of apoptotic processes. Neither PARP cleavage nor a sub-G1 peak could be detected (data not shown).
Cell extract preparation and Western blotting
Logarithmically growing HeLaS3 cells (control cells) were lysed using 1xTBS pH 7 with 1% NP40. This yielded whole cell extracts.
Synchronized HeLaS3 cells were fractionated with a previously described method (Riva et al. 2004). Briefly, cells were solubilized in a detergent-containing buffer (10 mM Tris–HCl, pH 7.4, 2.5 mM MgCl2, 0.5% NP-40, 1 mM DTT, 1 mM PMSF, 0.5 nM okadaic acid and protease inhibitors). After centrifugation, the supernatant was named as fraction 1 that represented cytoplasmic, nucleoplasmic and loosely chromatin-bound proteins. The insoluble material was then digested with 150 units DNase I (Sigma) per 107 cells for 30 min at 25 °C in digestion buffer (10 mM Tris–HCl, pH 7.4, 10 mM NaCl, 5 mM MgCl2, 0.2 mM PMSF, 0.5 nM okadaic acid and protease inhibitors). The obtained fraction 2 was called as DNase I-soluble fraction and contained proteins that were harder to be released from DNA, including those that were associated with these proteins, and possibly also proteins that were bound to short stretches of DNA. The remaining insoluble material was finally extracted with high salt buffer (10 mM Tris–HCl, pH 8.0, 2 M NaCl and protease inhibitors) yielding fraction 3. The high salt soluble material included proteins that were not extractable after DNase I treatment, that were tightly bound to DNA, or that belonged to protein complexes whose protein–protein or protein–DNA interactions were only destroyed by high salt conditions.
Mcm interaction studies were carried out with extracts using a slightly modified, previously described fractionation method (Mendez & Stillman 2000). Cells were synchronized in S phase by a double thymidine block followed by a release of the cells in fresh medium, harvested and resuspended in buffer A (10 mM HEPES, pH 7.5, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM DTT, 1x protease inhibitor cocktail (Sigma)) at 5 x 107 cells/mL and incubated for 10 min on ice. These HeLaS3 cell extracts contain free proteins (FPs) from the cytoplasm and the nucleoplasm. After a low speed centrifugation (1300 g, 5 min, 4 °C) the supernatant (FP fraction) was obtained. The FP fraction was further clarified by high speed centrifugation (14 000 g, 15 min, 4 °C). Nuclei were washed twice in buffer A, resuspended in 8 mM EDTA, 1 mM DTT, 1x protease inhibitor cocktail (Sigma) and incubated for 30 min at 4 °C. This first chromatin extraction procedure yielded chromatin fraction 1 (C1). Insoluble chromatin-bound proteins were obtained by centrifugation (1500 g, 5 min, 4 °C), and the supernatant (chromatin fraction 1–C1) was further clarified by high speed centrifugation (14 000 g, 15 min, 4 °C). The remaining chromatin was washed twice, resuspended in buffer A plus 2000 U/mL DNase I (Roche) and incubated for 30 min at RT and a further 30 min at 4 °C with 1 volume 0.5 M NaCl. Solubilized chromatin-bound proteins were obtained by high speed centrifugation (14 000 g, 5 min, 4 °C). The remaining pellet was successively incubated on ice for 30 min with 1 M and 2 M salt buffer. Fraction 1 M and 2 M were obtained by high speed centrifugation (14 000 g, 5 min, 4 °C), respectively. To analyze these, the protein equivalent of about 7.5 x 104 cells was loaded per lane of an SDS gel. For Western blotting standard procedures were used.
Immunoprecipitation
Immunoprecipitations were carried out in IP buffer (50 mM HEPES, pH 7.5, 150 mM NaCl and 0.5% NP-40) using protein G SepharoseTM 4 Fast Flow (Amersham Biosciences, part of GE Healthcare) or protein A Sepharose (Sigma). Fast Flow beads alone (negative control) or beads pre-coupled to desired antibodies were, after blocking with "Perfect Block" (MoBiTec), mixed with extracts of 106 cells in 500 µL IP buffer and rotated overnight at 4 °C. Mixing of cell extracts in 500 µL IP buffer decreased the salt concentrations in the extracts so that the immunoprecipitation was carried out at physiological conditions. To prevent a co-immunoprecipitation (Co-IP) mediated by DNA, ethidium bromide was added at a final concentration of 20 ng/µL. Unbound material was removed by washing 4 times with IP buffer. The immunoprecipitated proteins and possible co-IP partners were analyzed by Western blotting.
Immunofluorescence microscopy
Samples for immunofluorescence microscopy were prepared as follows: Cells were grown in quadriPERM dishes (VivaScience) on cover slides (Roth). BrdU, 5-bromo-1-(2-deoxy-ß-D-ribofuranosyl) uracil (Sigma) was used to mark S-phase cells. For pulse labeling cells were incubated 15 min with BrdU at a final concentration of 32 µM or as mentioned otherwise. The cover slides were then washed twice with PBS and fixed with 4% formaldehyde (Sigma) in PBS for 10 min at 4 °C. Afterwards, cells were permeabilized with 0.25% TritonX-100 in PBS for 10 min. To study the Mcm proteins, cells were permeabilized with 0.1% TritonX-100 and 0.02% SDS in PBS for 30 min at room temperature (modified according to Krude et al. 1996). For BrdU staining, DNA was denatured with 1 M HCl for 30 min at room temperature. To block unspecific protein binding the treated cells were incubated with 5% BSA in PBS overnight at 4 °C. The next day cover slides were washed 3 times with PBS and then simultaneously incubated with the respective primary antibodies (diluted in 1% BSA and 0.5% TritonX-100 in PBS) for 1 h at room temperature in a humidified chamber. After washing 3 times with PBS the appropriate secondary antibodies were incubated for 1 h at room temperature in a dark humidified chamber. After antibody incubation, cells were washed 2 times with PBT (PBS with 2% TritonX-100) before DNA staining with 2 µM To-Pro3 (Molecular Probes) in PBT for 45 min. After washing 3 times with PBS, the cover slides were mounted with mounting medium (20 mM Tris–HCl, pH 8.0, 90% glycerol and 200 µM 1,4-diaza-bicyclo(2.2.2)octane (DABCO)), and sealed with nail polish. Images were taken using a LSM 510 META microscope with standard software LSM 510 META 3.2 (Carl Zeiss GmbH, Jena, Germany).
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Footnotes |
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a Present address: Radiation Oncology and Biology, Radiobiology Research Institute, Churchill Hospital Headington, Oxford OX3 7LJ, UK.
* Correspondence: E-mail: fgrosse{at}fli-leibniz.de or h.nasheuer{at}nuigalway.ie |
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Accepted: 17 March 2007
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