The spatio-temporal organization of DNA replication sites is identical in primary, immortalized and transformed mammalian cells

Daniela S. Dimitrova and Ronald Berezney*

Department of Biological Sciences, State University of New York at Buffalo, Buffalo, NY 14260, USA

* Author for correspondence (e-mail: berezney{at}acsu.buffalo.edu)

Accepted 1 August 2002


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We investigated the organization of DNA replication sites in primary (young or presenescent), immortalized and transformed mammalian cells. Four different methods were used to visualize replication sites: in vivo pulse-labeling with 5-bromo-2'-deoxyuridine (BrdU), followed by either acid depurination, or incubation in nuclease cocktail to expose single-stranded BrdU-substituted DNA regions for immunolabeling; biotin-dUTP labeling of nascent DNA by run-on replication within intact nuclei and staining with fluorescent streptavidin; and, finally, immunolabeling of the replication fork proteins PCNA and RPA. All methods produced identical results, demonstrating no fundamental differences in the spatio-temporal organization of replication patterns between primary, immortal or transformed mammalian cells. In addition, we did not detect a spatial coincidence between the early firing replicons and nuclear lamin proteins, the retinoblastoma protein or the nucleolus in primary human and rodent cells. The retinoblastoma protein does not colocalize in vivo with members of the Mcm family of proteins (Mcm2, 3 and 7) at any point of the cell cycle and neither in the chromatin-bound nor in the soluble nucleoplasmic fraction. These results argue against a direct role for the retinoblastoma or nuclear lamin proteins in mammalian DNA synthesis under normal physiological conditions.

Key words: DNA replication patterns, Primary and transformed mammalian cells, Senescence, Retinoblastoma protein, Minichromosome maintenance proteins, Nuclear lamin proteins


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The nucleus is the control center of the eukaryotic cell, where the major genomic processes of DNA replication, repair, transcription, RNA processing and transport occur and are precisely coordinated in space and time. There is a growing realization that the coordination and regulation of the different nuclear activities is mediated, at least in part, by the spatial compartmentalization and higher order arrangement of chromosomal domains, structural components and enzymatic and regulatory activities (Berezney, 2002Go; Lamond and Earnshaw, 1998Go). It has been demonstrated that DNA replication (Nakamura et al., 1986Go; Nakayasu and Berezney, 1989Go; O'Keefe et al., 1992Go) and concomitant chromatin assembly and maturation (Krude, 1995Go; Murzina et al., 1999Go; Shibahara and Stillman, 1999Go; Taddei et al., 1999Go), DNA repair (Jackson et al., 1994Go), transcription and RNA processing (Wei et al., 1999Go) take place at discrete sites within the nucleus. The dynamic changes in the spatial distribution of such sites and associated activities during development and differentiation (Blumenthal et al., 1974Go; Ferreira and Carmo-Fonseca, 1997Go; Leibovici et al., 1992Go; Micheli et al., 1982Go), during cell cycle progression (Bravo and Macdonald-Bravo, 1987Go; Dimitrova et al., 1999Go; Hozak et al., 1994Go; Leonhardt et al., 1992Go; Ma et al., 1998Go; Manders et al., 1992Go; O'Keefe et al., 1992Go; Rossi et al., 1999Go), or relative to the metabolic activity of the cell (Dimitrova and Gilbert, 2000bGo; Spector, 1996Go; Zeng et al., 1997Go) are a subject of active investigation.

The integrity of the nuclear organization is compromised in a variety of diseases. Major structural components and functional domains of the nucleus undergo dramatic changes during viral infection (Maul, 1998Go; Monier et al., 2000Go; Wilcock and Lane, 1991Go), malignant transformation, or in organisms with genetic syndromes. Examples of altered nuclear components in transformed cells include the nuclear matrix (Davido and Getzenberg, 2000Go; Pienta et al., 1989Go), chromatin (Chadee et al., 1999Go; Chadee et al., 1995Go; de Campos Vidal et al., 1998Go; Herrera et al., 1996Go; Vassilev et al., 1995Go), nuclear bodies and speckles (Gordon et al., 2000Go; Manuelidis, 1984Go; Spector et al., 1992Go) and replication proteins (Bechtel et al., 1998Go). Regulatory activities play a key role in establishing nuclear architecture that favors tumor suppression (Chuang et al., 1997Go; Linares-Cruz et al., 1998Go). Genetic disorders have been traced to mutations in genes encoding for components of chromatin [Rett syndrome (Amir et al., 1999Go)], of the DNA replication, recombination and repair machinery [e.g. ICF syndrome (Hansen et al., 1999Go)], and nuclear structural components, such as the nuclear lamins (Wilson et al., 2001Go).

We set out to investigate whether differentiation or malignant transformation of mammalian cells is accompanied by significant changes in nuclear architecture and the number and distribution of DNA replication, transcription and RNA processing sites. In this study, we have compared the spatio-temporal patterns of DNA replication sites within normal, immortalized and transformed mammalian cells of various origin. Unlike a recent study which reports a fundamental difference in the organization of DNA replication sites in primary versus immortalized cell lines (Kennedy et al., 2000Go), we find that the number and distribution of these sites throughout the S-phase is strikingly similar in all cell lines examined and is independent of the technique used to visualize the replication sites. We further report that neither the nuclear lamin proteins nor the retinoblastoma protein (pRb) are significantly associated at any stage in S-phase with replication sites and that pRb does not colocalize with minichromosome maintenance (Mcm) protein family members.


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Cell culture
CHO AA8 (immortal Chinese hamster ovary fibroblasts, ATCC), CHOC 400 [CHO cell derivative containing an amplified DHFR gene locus (Milbrandt et al., 1981Go)], JH1 [SV40-transformed CHOC 400 cell line derivative (Wu et al., 1998Go)], TE671 (human medulloblastoma), HT1080 (human fibrosarcoma, ATCC), MG63 (human osteosarcoma, ATCC), HeLa (human cervix adenocarcinoma, ATCC) and MCF7 (human breast adenocarcinoma, ATCC) were propagated in Dulbecco's modified Eagle's medium (DMEM, Life Technologies), supplemented with 5% fetal bovine serum (FBS, Life Technologies). BALB/c 3T3 cells (immortal mouse fibroblasts, ATCC) were grown in the presence of 10% calf serum (Life Technologies). The human colon carcinoma cell lines HCT8 and HCT116 were grown in RPMI 1640 or McCoy 5A medium (Life Technologies), respectively, supplemented with 5% FBS. The primary cell lines WI38 (normal diploid human female lung fibroblasts, ATCC), NHF1 (normal diploid human foreskin fibroblasts) and NRK (normal rat kidney epithelial cells) were grown in DMEM in the presence of 10% FBS. Cells were synchronized by aphidicolin arrest at the G1/S border after either mitotic selection (release from nocodazole block), or serum starvation, as described previously (Dimitrova et al., 1999Go; Fox et al., 1991Go; Ma et al., 1998Go).

Visualization of DNA replication sites
The immunofluorescent labeling of DNA replication sites was performed in 4 different ways. (1) 5-bromo-2'-deoxyuridine (BrdU)-substituted DNA was immunolabeled by the acid-depurination protocol as described (Dimitrova et al., 1999Go), using a monoclonal anti-BrdU antibody (Becton Dickinson) and FITC- or Texas Red-conjugated donkey anti-mouse secondary antibodies (Jackson ImmunoResearch Laboratories). (2) Detection of BrdU using the nuclease cocktail (BrdU labeling and detection kit, cat. no. 1296736, Roche) was performed exactly as recommended by the manufacturer. (3) Sites of DNA synthesis within digitonin-permeabilized cells were labeled by in situ run-on reaction in a replication cocktail (30 mM HEPES, pH 7.6, 7 mM MgCl2, 1 mM dithiothreitol, 100 µM each dATP, dGTP and dCTP, 25 µM biotin-11-dUTP, 400 µM each GTP, CTP and UTP, 4 mM ATP, 40 mM creatine phosphate, 20 µg/ml creatine phosphokinase). Biotin-dUTP was detected with Texas Red-conjugated streptavidin (Amersham). The digitonin permeabilization of cells under conditions that preserve intact nuclear membranes has been described (Dimitrova and Gilbert, 1998Go). (4) PCNA and RPA were immunolabeled as described (Dimitrova et al., 1999Go). Nuclear lamin proteins were detected using rabbit polyclonal antibodies (gift of Dr G. Blobel, Rockefeller University).

In the triple labeling experiments in Figs 7,8,9, nascent DNA was labeled with 5-chloro-2'-deoxyuridine (CldU) and visualized using a rat anti-BrdU antibody (Harlan/SeraLab) as described (Dimitrova et al., 1999Go). Nucleolin was detected with a mouse monoclonal anti-nucleolin antibody (Santa Cruz), the retinoblastoma protein — with either a rabbit polyclonal (NeoMarkers; a gift from Dr D. Goodrich, Roswell Park Cancer Institute) or a mouse monoclonal antibody (a gift from Dr T. Melendy, SUNY Buffalo), and Mcm7 — with a mouse monoclonal antibody (Santa Cruz).



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Fig. 7. The numerous pRb granules are not preferentially associated with perinucleolar regions and do not colocalize with CldU-labeled replication sites during any stage of S-phase. An asynchronous culture of WI38 cells was pulse-labeled for 5 minutes with 10 µM CldU, fixed with formaldehyde and stained with anti-pRb rabbit polyclonal antibodies and anti-nucleolin mouse monoclonal antibodies. The cells were briefly fixed again with formaldehyde before mild HCl hydrolysis and labeling of CldU-substituted DNA with rat monoclonal anti-BrdU antibodies. DNA was stained with DAPI. Images corresponding to double- and triple-labeling protocols were pseudocolored as indicated within each panel. Sites that co-localize appear yellow for red/green, light blue or blue-green for blue/green, and magenta for blue/red merged images, respectively. Based on the appearance of the CldU replication patterns, the cells were classified into early-S (A), mid-S (B) and late-S (C) categories. Besides the strong nucleolar signal, nucleolin exhibits a weaker diffuse nucleoplasmic staining within formaldehyde-fixed cells. This soluble nucleoplasmic form of nucleolin can be removed completely by mild extraction with Triton X100 prior to fixation (not shown). Note the nearly complete lack of colocalization (yellow color) between the pRb (red) and CldU (green) signals in the double-labeled images.

 


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Fig. 8. pRb does not colocalize with Mcm proteins within licensed chromatin in permeabilized cells. An aliquot of the same CldU-labeled W138 asynchronous cell culture used for the results shown in Fig. 7 was extracted for 3 minutes on ice with 0.5% Triton X100 in cytoskeleton (CSK) buffer. The cells were then fixed with formaldehyde and stained with anti-pRb rabbit polyclonal antibodies, anti-Mcm7 mouse monoclonal antibodies and anti-BrdU rat monoclonal antibodies as described in Fig. 7. The cells were classified into G1, early-S, middle-S and late-S according to the presence of detergent-resistant Mcm7 chromatin association, the presence or absence of CldU staining and the type of CldU replication pattern. DNA was stained with DAPI. Images corresponding to double- and triple-labeling protocols were pseudocolored as indicated within each panel. Sites that co-localize appear yellow for red/green, light blue or blue-green for blue/green, and magenta for blue/red merged images, respectively. Identical results were obtained with anti-Mcm2 and anti-Mcm3 antibodies. Compared to Fig. 7, longer exposure times were used to capture the pRb images due to the weaker fluorescent signal as a result of the release of a fraction of nuclear pRb by the nonionic detergent extraction procedure.

 


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Fig. 9. pRb and Mcm proteins are not present at the same nuclear sites within intact cells. An aliquot of the same CldU-labeled WI38 asynchronous cell culture was stained with anti-pRb, anti-Mcm7 and anti-BrdU antibodies as described in Fig. 7 without a prior Triton X100 extraction. Cells were categorized according to the type of CldU replication pattern. Images corresponding to double- and triple-labeling protocols were pseudocolored as indicated within each panel. Sites that co-localize appear yellow for red/green, light blue or blue-green for blue/green, and magenta for blue/red merged images, respectively. Note the presence of Mcm proteins throughout the nucleus at all cell cycle stages.

 

The cells were imaged on a Zeiss epifluorescent microscope equipped with a RTE CCD camera (Princeton Instruments), controlled by IP Lab software (Scanalytics), or on an Olympus BX51 epifluorescent microscope equipped with a Sensicam QE CCD camera (The Cooke Corporation), controlled by Image-Pro Plus software (Media Cybernetics). The images were assembled in an Apple G3 Powerbook using Adobe Photoshop 5.0.2 and Claris Draw 1.0v4 software.


    Results and Discussion
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Eukaryotic cells respond in distinct ways to physiological or environmental signals. Primary cells in culture divide a limited number of times and then go into a growth arrest state, known as replicative senescence (Hayflick and Moorhead, 1961Go). Senescence ensues naturally as a result of prolonged passaging of the primary cell cultures, or in response to oxidative damage or the activation of certain oncogenes (Lundberg et al., 2000Go). However, eukaryotic cells can also acquire the potential to grow indefinitely in culture (McLean, 1993Go). Immortalized cells either preserve a normal karyotype and the differentiated phenotype characteristic of primary cells, or undergo transformation and develop features characteristic of tumor cells (e.g. abnormal response to growth factors and hormones, loss of differentiated phenotype, loss of contact inhibition, polyploidy and/or chromosomal aberrations, etc.).

Whereas a significant amount of information has accumulated regarding the changes in cell morphology, chromosome structure, enzymatic activities and cell cycle regulatory activities that distinguish normal proliferating cells, senescent cells or immortal/transformed cells, very little is known about the relationship between senescence or immortalization and the spatio-temporal regulation of DNA replication. Scattered reports have been published on the appearance and sequence of DNA replication patterns within normal or tumor cells of various derivation (Fox et al., 1991Go; Humbert et al., 1992Go; Humbert and Usson, 1992Go; Kill et al., 1991Go; O'Keefe et al., 1992Go; van Dierendonck et al., 1989Go), but few attempts have been made to conduct a comprehensive study of these patterns within aging cells at different passage levels or within immortalized cells at different stages of transformation. Therefore, we set out to investigate whether there are differences in the replication patterns between transformed, immortal and young or aging primary mammalian cells.

We collected a number of rodent and human cell lines that belong to these groups. Cells were synchronized at the beginning of S-phase and then released for various times to prepare cell populations at different stages of S-phase (Fig. 1). To test whether the cell synchronization procedures might influence the distribution of the replication sites, replication patterns were also analyzed in exponentially growing cells (Fig. 2). Furthermore, to ensure that the method employed to visualize the replication patterns does not produce artifactual results and lead to inaccurate conclusions, we used four different techniques to reveal sites of DNA replication.



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Fig. 1. The distribution of nuclear DNA replication sites in primary, immortalized and transformed mammalian cells follows identical spatio-temporal patterns during S-phase. Cell cultures synchronized at different stages of S-phase after release from aphidicolin block [5 min (G1/S), 1 hour, 2 hours and 4 hours (early-S), 6 hours and 8 hours (mid-S), 10 hours and 12 hours (late-S)] were pulse-labeled for 5 minutes with 30 µg/ml BrdU, fixed and sites of BrdU incorporation were detected with a nuclease/anti-BrdU cocktail. Based on the number, size, shape and distribution of fluorescent foci, the replication sites were classified into five major types of patterns, which were similar in all cell lines.

 


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Fig. 2. HCl hydrolysis (gray columns) and nuclease digestion (black columns) techniques for visualization of BrdU-labeled nascent DNA reveal identical frequency of the five major replication patterns within S-phase nuclei of exponentially growing primary and immortalized mammalian cells. Shown are results from a single experiment. Identical results were obtained in three independent experiments.

 

Visualization of BrdU-labeled DNA replication sites through the use of nuclease/anti-BrdU antibody cocktail
In the first series of experiments, the cell cultures were synchronized at the G1/S border by aphidicolin arrest. Aliquots of each cell culture were released from the block for different time periods (5 minutes, 1 hour, 2 hours, 4 hours, 6 hours, 8 hours, 10 hours or 12 hours) and pulse-labeled for 5 minutes with BrdU just prior to fixation and immunolabeling of the sites of BrdU incorporation with fluorescent anti-BrdU antibodies. Since the immunodetection of BrdU is possible only within single-stranded DNA regions, two methods were used to create such regions. First, a nuclease cocktail was used to generate short single-stranded tracts accessible for binding of the anti-BrdU antibodies (Dolbeare and Gray, 1988Go; Fox et al., 1991Go). Although this technique is not frequently used for labeling of replication foci, it is considered to be the one that best preserves the nuclear architecture (Dolbeare and Gray, 1988Go; Kennedy et al., 2000Go). Fig. 1 shows a gallery of immunofluorescent images representative of different stages of S-phase within a subset of the cell lines analyzed, including primary (WI38, NHF1 and NRK), immortal (CHOC 400 and 3T3) and transformed (JH1, HeLa, MCF7, TE671 and HT1080) cells. We and others have previously reported three major types of replication patterns in several mammalian cell lines, corresponding to early (type I), middle (type II) and late (type III) stages of S phase (Ma et al., 1998Go; Nakayasu and Berezney, 1989Go). A finer discrimination of the replication patterns, however, led to their classification into five major types (Dimitrova and Gilbert, 1999Go; Humbert and Usson, 1992Go; O'Keefe et al., 1992Go). Based on the number, size, shape and distribution of the fluorescent foci, the replication patterns in the cell lines analyzed in this study were similarly classified into five types.

The type IA pattern is detectable for a brief period of time (~30 minutes) at the onset of S-phase. It consists of a relatively low number (several dozens to few hundred) of small discrete foci scattered throughout the nuclear interior, but excluded from the peripheral, nucleolar or heterochromatic regions. Importantly, this pattern was observed in all cell lines (Fig. 1), indicating that DNA replication initiates at the beginning of S-phase within nuclear sites that are similarly distributed in the different cell lines.

It has been suggested that the arrest of cells at the G1/S border leads to the artificial activation of additional replication origin sites that are not normally utilized in unperturbed cell cycles (Li et al., 2000Go; Taylor, 1977Go). The results presented in Fig. 1, as well as previously published reports (Dimitrova and Gilbert, 1999Go; Dimitrova and Gilbert, 2000bGo; Fox et al., 1991Go; Jackson, 1995Go), demonstrate that this does not apply to a tight aphidicolin block administered for limited time intervals. Our observation of type IA patterns within exponentially growing cell cultures (Fig. 2) at a frequency ~5-10% of the BrdU-positive nuclei is consistent with the duration of this pattern observed in our synchronized cell studies (~30 minutes within a typical ~10-hour S-phase). We conclude that limited exposure to aphidicolin does not change significantly the number of early-S-phase replication sites or induce artifacts in the overall replication patterns observed from early to late S-phase.

The number and fluorescent intensity of foci rapidly increased as the cells were allowed to progress further into S-phase (1-5 hours), until the nuclear interior appears virtually filled with foci, including the regions close to the nuclear periphery. This pattern, consisting of small and numerous discrete granules [several hundreds to one- to two-thousand (Ma et al., 1998Go)], is here classified as type IB (Fig. 1). It is the major replication pattern observed until the middle of S-phase. Electron microscopy and biochemical data accumulated by different labs have demonstrated that the euchromatic regions of the genome are involved in the DNA replication process during this time (Nakayasu and Berezney, 1989Go; O'Keefe et al., 1992Go; van Dierendonck et al., 1989Go; Williams and Ockey, 1970Go).

A striking change in the distribution of the replication foci marks the transition from early to middle S-phase. Although the size and intensity of the individual foci did not change as compared to early-S, the number of the DNA replication sites dropped drastically and their localization switched from the internal euchromatic to predominantly perinuclear, perinucleolar and/or intranucleolar regions of the genome, known to accommodate mostly heterochromatin. This replication pattern, classified here as type II, persisted for 2-3 hours during mid-S (5-8 hours after release from aphidicolin arrest) in all cell lines (Fig. 1).

During late S-phase, the size of the replication sites increased and their shape became irregular. Within type IIIA (~1-2 hours), the number of replication sites (several hundred) was intermediate between types IB and II. The fluorescent foci were both at the nuclear periphery and scattered throughout the interior and, consistent with previous observations (Nakayasu and Berezney, 1989Go; O'Keefe et al., 1992Go; van Dierendonck et al., 1989Go), they often had a ring-, chain- or horseshoe-like appearance (Fig. 1). The number of replication foci decreased again at the very end of S-phase and their morphology changed to more compact granules (this is especially pronounced in the rodent cell lines — see the NRK, CHOC 400, JH1 panels in Fig. 1). This pattern, classified as type IIIB (~1 hour), consisted of a small number of heterogeneously sized (often extremely large) sites over peripheral or interior nuclear regions. They stained intensely with DAPI, which is characteristic of the constitutive heterochromatin.

In contrast to the early- and mid-S-phase patterns, not all cell lines showed identical morphology, number and distribution of replication sites within mid/late-S types II, IIIA and IIIB. For example, the late-S sites in the TE671, HCT116 and HT1080 human tumor cell lines remained relatively small in size and, in TE671 and HCT116, they occupied predominantly perinuclear regions with few sites in the nuclear interior (Fig. 1). In addition, whereas the rat and Chinese hamster cell lines showed a very small number (often just one) of large, homogeneously stained and often multi-lobed type IIIB sites, the human cell lines contained a higher number (a dozen or more) of late-S sites with a more heterogeneous morphology. The observed differences in the late-S replication patterns were not specific for the primary vs. the transformed cells, but rather reflected features characteristic of a certain cell type. For example, the replication patterns were indistinguishable between the primary NRK, the immortal CHO AA8 (not shown) and CHOC 400 cells, and the SV40-transformed JH1 cells. Likewise, the replication patterns were similar between the primary human cell lines (WI38 and NHF1) and the transformed HeLa cells.

When the replication patterns were analyzed within young (starting at passage 18), ageing (passage 35-45) or presenescent (passage 50) primary human fibroblasts (WI38), no major differences were found, with the exception of a marked decrease in the percentage of cells incorporating BrdU during the late passages, as expected for a senescent cell culture (data not shown). We conclude that there are no fundamental differences in the distribution of DNA replication sites and the spatio-temporal sequence of replication patterns between primary, immortal or transformed mammalian cells.

Visualization of BrdU-labeled DNA replication sites after mild HCl hydrolysis
Our observations illustrated in Fig. 1 are in sharp contrast with the conclusions reached in a recent study on the distribution of replication sites in primary mammalian fibroblasts (Kennedy et al., 2000Go). These investigators reported that primary cell lines exhibited early-S replication pattern that lasted for approximately 3 hours into S-phase and consisted of 5 to 20 perinucleolar foci. This unusual early-S-phase pattern was observed in asynchronous cell cultures, as well as in synchronized cell populations irrespective of the synchronization method (i.e., contact inhibition with or without an aphidicolin or hydroxyurea block). In contrast, immortalized cells exhibited numerous early-S replication sites. Kennedy and colleagues (Kennedy et al., 2000Go), further suggested that previous observations of numerous replication sites in early-S phase in primary cells are incorrect due to the use of immunolabeling protocols (namely HCl hydrolysis) that destroy nuclear structure. It was suggested that the use of DNase I to expose single-stranded DNA regions for binding by the anti-BrdU antibodies better preserves nuclear organization. Since we agree that HCl treatment (used in the majority of the previous studies) is destructive for many nuclear antigens, and, when used inappropriately, could produce artifacts, we applied in this present study four different techniques for immunolabeling of DNA replication sites to aliquots of the same cell cultures. This enabled a direct comparison among the results obtained with each technique.

To optimize the comparison, cells from the same synchronized cell populations employed in the nuclease labeling method (Fig. 1) were used for the HCl method. The same five S-stage-dependent replication patterns were observed as found following nuclease treatment (compare Figs 1 and 3). In addition, a comprehensive analysis of the percent of each replication pattern in exponentially growing cells is shown in Fig. 2. Fourteen different cell lines ranging from primary to immortalized to transformed cells were compared following application of the nuclease or HCl labeling protocols. No significant differences were observed in the distribution of the five replication patterns as a function of the labeling method.



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Fig. 3. Spatial organization of DNA replication sites in primary, immortalized and transformed mammalian cells detected by limited HCl depurination of BrdU-substituted DNA. Aliquots of cells from the same synchronized cell populations as in Fig. 1 were used.

 

While we cannot definitively explain the discrepancies between our results and those of Kennedy and colleagues (Kennedy et al., 2000Go), one possibility is that the nuclease labeling procedure used in their study failed to efficiently immunolabel the BrdU sites within the primary cell lines. In this regard, Dolbeare and Gray demonstrated that the sole use of an endonuclease may not be sufficient to expose enough single-stranded DNA for significant immunodetection. Addition of exonuclease III to the nuclease/antibody cocktail, however, resulted in the generation of more extensive single-stranded DNA regions, which were very efficiently recognized by the anti-BrdU antibodies (Dolbeare and Gray, 1988Go). It is, therefore, conceivable that, under the conditions used by Kennedy and colleagues (Kennedy et al., 2000Go), DNase I (an endonuclease) generated mostly nicks in the chromosomal DNA of the primary fibroblasts (as testified by the TUNEL staining performed in that study), but failed to create sufficient number of single-stranded DNA regions for good immunolabeling. It is interesting to consider that the late-S-phase primary cells and the immortalized and transformed cells in general might have been more sensitive to the DNase I digestion and, therefore, were more efficiently stained with the anti-BrdU antibodies.

This possibility is corroborated by the published observation that chromatin of pRb-deficient primary mouse fibroblasts exhibits higher sensitivity to micrococcal nuclease digestion than chromatin of pRb-positive primary mouse or human fibroblasts (Herrera et al., 1996Go). Intriguingly, this higher sensitivity is especially pronounced during the progression of the cells from late-G1-phase into early-S-phase. Higher nuclease sensitivity is associated with relaxed chromatin structure, which potentially results from the elevated levels of histone H1 and H3 phosphorylation (Hohmann, 1983Go; Lu et al., 1994Go) within pRb-deficient cells at this time of the cell cycle due to deregulated cyclin-dependent kinase 2 (Cdk2) activity (Chadee et al., 1999Go; Herrera et al., 1996Go). Similar to the pRb-deficient primary fibroblasts, many transformed cell lines either lack pRb, or express mutant inactive pRb (Nevins, 2001Go; Niculescu et al., 1998Go). Thus, pRb-negative primary and transformed cells, as well as tumor cells expressing various oncogenes may be more susceptible to BrdU labeling via the nuclease cocktail technique due to the higher accessibility of chromatin to nuclease attack (Chadee et al., 1995Go; Herrera et al., 1996Go; Taylor et al., 1995Go). The possibility of different chromatin/chromosome organization (and different nuclease sensitivity, respectively) between primary and transformed cells (de Campos Vidal et al., 1998Go; Takaha et al., 2002Go) and of chromatin rearrangements, taking place during the progress of S-phase, is an exciting direction to pursue.

We emphasize that, unlike Kennedy et al. (Kennedy et al., 2000Go) who used post-microscopy deconvolution image processing, we have not applied any computer processing to our raw microscopic images. Deconvolution algorithms, when applied inappropriately, can lead to a significant loss of relevant immunofluorescent signal. Weakly fluorescing replication foci, typical of nuclei at the onset of S-phase, would be especially susceptible to elimination by harsh deconvolution processing after inefficient immunolabeling, thus leading to in silico generation of artifactual results.

Visualization of DNA replication sites without generation of single-stranded DNA regions
To further test our conclusions, we also applied two protocols that do not require the generation of single-stranded DNA regions to visualize the location of replication sites. First, biotinylated dUTP, incorporated into DNA, can be directly immunolabeled with fluorescent streptavidin without the need of DNA denaturation. The cells have to be permeabilized to allow access of the biotin-dUTP to the nucleus. Following permeabilization, DNA synthesis continues at the same sites that were active in vivo (Krude, 1995Go; Mills et al., 2000Go). Unfortunately, most previous studies were performed with cells permeabilized with the non-ionic detergents Triton X-100 or Nonidet P40. These detergents severely compromise both the overall structure and the replication capacity of mammalian nuclei (Dimitrova and Gilbert, 1998Go), which raises legitimate concerns about the degree of preservation of nuclear organization (Kennedy et al., 2000Go). To optimize comparisons, we used aliquots of the same cell cultures from Fig. 1 to prepare permeabilized cells with intact nuclei (Fig. 4A). We previously demonstrated that controlled permeabilization with digitonin produces nuclei with uncompromised nuclear structure, fully preserved replication capacity and unaltered distribution of replication proteins and origin sites (Dimitrova and Gilbert, 1998Go; Dimitrova and Gilbert, 1999Go; Dimitrova et al., 1999Go). As illustrated in Fig. 4B,C, the labeling of nascent DNA in situ by run-on replication in the presence of biotin-11-dUTP generates the same replication patterns as the in vivo labeling with BrdU. This is in agreement with previous reports on biotin-dUTP labeling of mammalian nuclei (Kill et al., 1991Go; Mills et al., 2000Go; Nakayasu and Berezney, 1989Go).



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Fig. 4. Visualization of replication sites within permeabilized primary or transformed human cells after nuclear run-on replication. (A) Controlled permeabilization with digitonin produces cells with intact nuclei (impermeable to large molecules that lack nuclear localization signal (e.g. Texas Red IgG, right panel) and well preserved nuclear structure. Intact nuclei prepared from WI38 (B) or HeLa (C) cells were introduced into in vitro replication cocktail and nascent DNA at replication forks generated in vivo was labeled with biotin-dUTP. Nuclei were fixed with 70% ethanol and biotin-dUTP was detected with Texas Red-conjugated streptavidin. DNA was stained with 0.1 µg/ml 4',6-diamidino-2-phenylindole (DAPI).

 

Finally, in order to visualize replication sites by a method that does not rely on the detection of nascent DNA, we stained aliquots of the same cell cultures as in Fig. 1 with antibodies specific for protein components of mamalian replication forks. This was done either independently, or in combination with BrdU staining. As expected, the immunolabeling of PCNA and RPA, two of the best characterized replication proteins, produced replication patterns identical to those generated by the immunolabeling of nascent DNA (not shown). The two proteins colocalized at discrete nuclear sites (Fig. 5I), whose distribution dynamically changed during S-phase.



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Fig. 5. The replication fork proteins RPA and PCNA, but not nuclear lamins, colocalize with BrdU-labeled nascent DNA within replication foci. HeLa and WI38 cells were pulse-labeled with BrdU, extracted with Triton X100 to remove soluble proteins and fixed with freshly prepared formaldehyde. (A-C) One aliquot of the cells was treated with HCl and then stained simultaneously with lamin B- (green) and BrdU-specific (red) antibodies. A second aliquot was first stained for lamin B [green (D-G)] or lamin A/C [red (J-Q)], the cells were fixed again with formaldehyde, then treated with HCl and stained for BrdU (red in D-F and green in J-Q). (H) The same protocol was used to stain for PCNA (red) and BrdU (green). (I) A fourth aliquot of cells was stained directly with anti-PCNA (red) and anti-RPA (green) antibodies. Results obtained with HeLa cells are shown in panels A-I. Identical results were obtained with WI38 cells.

 

To verify that these are active DNA replication sites, we performed double-labeling experiments to demonstrate that protein components of replication forks colocalize with BrdU-labeled sites. We emphasize that this double-labeling must be performed in a specific order to avoid artifactual results. As previously discussed, the mild HCl hydrolysis required for immunolabeling of BrdU sites is destructive for many nuclear proteins (Humbert et al., 1992Go). To illustrate this point, we immunolabeled the lamin B proteins, whose established localization at the nuclear boundary can be used as a reliable reference point. The immunolabeling of the lamin B proteins was done either before, or following HCl treatment.

As evident from Fig. 5A-C, prior HCl hydrolysis destroyed the integrity of the nuclear lamina and resulted in a dispersed distribution of the lamin B epitopes, in agreement with the results reported by Kennedy et al. (Kennedy et al., 2000Go). However, when the lamin B staining was performed first, and the fluorescent anti-lamin antibodies were covalently fixed to their specific sites before the cells were treated with HCl (Fig. 5D-F), the exclusive localization of the lamin B antibodies to the nuclear envelope was preserved [most of the few internal lamin B foci could be traced to invaginations of the nuclear envelope (results not shown)]. Identical distribution is obtained when lamin B proteins are stained without any HCl treatment. We did not detect colocalization of internal lamin B foci and BrdU sites during any stage of S-phase (examples of early-S and mid-S nuclei are shown in Fig. 5A-F and G, respectively), and neither before (Fig. 5D-G), nor after (Fig. 5A-C) HCl hydrolysis. Identical results were obtained with lamin A/C-specific antibodies (Fig. 5J-Q). In contrast, PCNA (Fig. 5H) or RPA sites (not shown) colocalized with the BrdU sites, as expected for replication fork proteins.

These results are in agreement with many previous studies, but differ strikingly from the observations on the cell cycle distribution of these proteins reported by Kennedy and colleagues (Kennedy et al., 2000Go). They report that abundant DNA replication-related proteins, such as PCNA and CAF-1 localize to only a few discrete perinucleolar foci during G1- and early S-phase. Such observations are inconsistent with previous studies of these proteins in both primary and immortalized mammalian cells. For example, both RPA and PCNA are present in a soluble form during the entire cell cycle, thus generating strong uniform nuclear staining. A chromatin-bound fraction of these proteins, which localizes to the DNA replication sites, is detected only during S-phase, after a prior extraction of the soluble fraction (Bravo and Macdonald-Bravo, 1987Go; Dimitrova and Gilbert, 2000aGo; Dimitrova et al., 1999Go). Therefore, the very low number (5 to 20) of discrete foci of total nuclear PCNA and CAF-1 during G1-phase and the first 2-3 hours of S-phase in formaldehyde-fixed cells described by Kennedy and colleagues (Kennedy et al., 2000Go) is bizarre and is not consistent with the known behavior of these proteins (Bravo and Macdonald-Bravo, 1987Go; Dimitrova et al., 1999Go; Humbert et al., 1992Go; Krude, 1995Go; Marheineke and Krude, 1998Go; Murzina et al., 1999Go; Shibahara and Stillman, 1999Go).

The retinoblastoma protein does not colocalize with DNA replication sites or mammalian Mcm proteins
pRb is a key regulator of cell cycle progression (Zheng and Lee, 2001Go) and is present in a functionally active, hypophosphorylated form during late mitosis/G1-phase, or in a functionally inactive, unphosphorylated or hyperphosphorylated forms during G0- and throughout late-G1/S/G2-phases, respectively (Ezhevsky et al., 2001Go; Ho and Dowdy, 2002Go; Moberg et al., 1996Go). The estimated number of pRb molecules per human cell is of the order of 1 million (Goodrich et al., 1991Go) and the levels of this protein do not vary excessively during the cell cycle (Buchkovich et al., 1989Go; Mihara et al., 1989Go; Muller et al., 1997Go). It is difficult to imagine that a million molecules of an important cell cycle and differentiation regulator known to have numerous genomic targets (Dyson, 1998Go; Wells et al., 2000Go; Zheng and Lee, 2001Go) would coalesce into only 5-20 perinucleolar foci within mammalian nuclei, as observed by Kennedy et al. (Kennedy et al., 2000Go). Moreover, this immunofluorescent pattern differs from the findings on the subnuclear localization of pRb family members in both primary, immortalized and transformed cells reported by several other groups (Bartek et al., 1992Go; Cinti et al., 2000Go; Fortunato and Spector, 1998Go; Mittnacht and Weinberg, 1991Go; Szekely et al., 1991Go; Zini et al., 2001Go). Most of these studies found numerous pRb-positive foci and/or speckles scattered throughout the nucleus. However, differences have also been reported, mostly concerning the size, number and intensity of the pRb granules (Szekely et al., 1991Go). Whereas some of these differences could be explained by the different functional status of pRb in primary vs. transformed cells (i.e. normal vs. mutant pRb forms) (Cinti et al., 2000Go; Szekely et al., 1991Go), others clearly derive from the use of different fixation protocols (Szekely et al., 1991Go) or from the utilization of unrelated anti-pRb antibodies (Bartek et al., 1992Go; Mittnacht and Weinberg, 1991Go; Szekely et al., 1991Go), which recognize different epitopes and, potentially, different nuclear subpopulations of pRb.

Since, to our knowledge, Kennedy et al. (Kennedy et al., 2000Go) are the only investigators who have directly examined the relative distribution of pRb and nuclear DNA replication sites and in view of the apparent discrepancies, we decided to reinvestigate this important issue using our collection of anti-pRb antibodies. From the eight anti-pRb antibodies, which we tested, only two (see Materials and Methods) gave positive immunofluorescent signal with primary human WI38 and NHF1 fibroblasts. In agreement with the observations reported by Szekely et al. (Szekely et al., 1991Go), we found that formaldehyde fixation of the cells resulted in a more uniformly granular staining pattern (e.g. see Fig. 6), whereas methanol/acetone fixation resulted in a smaller number of heterogeneously sized pRb granules (not shown). Since methanol treatment is known to both extract and fix proteins through precipitation, it is likely that these differences result from the loss and/or aggregation of pRb molecules in cells fixed with organic solvents. We, therefore, used formaldehyde fixation in the remaining experiments.



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Fig. 6. pRb is present at hundreds of sites scattered throughout mammalian nuclei. An asynchronous culture of WI38 cells was pulse-labeled for 5 minutes with 30 µg/ml BrdU, fixed with freshly prepared formaldehyde and stained with pRb-specific rabbit polyclonal antibodies (A,D,G) and a nuclease/anti-BrdU labeling cocktail (B,E). DNA was stained with DAPI. (H) pRb foci (red) density is higher within nuclear regions with lower density of DNA (blue).

 

Under our experimental conditions, pRb exhibited exclusively nuclear localization and all interphase nuclei within asynchronously growing primary fibroblast cell cultures were positive. In agreement with previous reports in the literature (Bartek et al., 1992Go; Mittnacht and Weinberg, 1991Go), we observed some variability in the intensity of the pRb immunofluorescent signal (Fig. 6). The weaker-staining nuclei were generally, albeit not exclusively, BrdU-negative (e.g. Fig. 6A-C) and, thus, it is likely that most of them are G1-phase nuclei, since the levels of pRb are known to increase ~twofold in the course of the cell cycle (Buchkovich et al., 1989Go; Mihara et al., 1989Go; Szekely et al., 1991Go). In all nuclei the pRb labeling appeared as hundreds of fluorescent foci scattered throughout the nucleus and often showed a bias towards nuclear regions with lower DNA density [Fig. 6G-I (Szekely et al., 1991Go)].

Simultaneous immunolabeling of pRb and the major nucleolar protein nucleolin (Ginisty et al., 1999Go) demonstrated that the nucleolar regions contain numerous pRb foci (Fig. 7), consistent with a role for the Rb family of proteins in the regulation of rDNA transcription (Ciarmatori et al., 2001Go; Hannan et al., 2000Go). We, however, did not find any cells in the entire exponentially growing population (Figs 6, 7), which exhibited the limited perinucleolar pRb staining pattern described by Kennedy et al. (Kennedy et al., 2000Go). Additionally, merged images of pRb and DNA replication sites following triple labeling experiments revealed that pRb was present at hundreds of extranucleolar sites, which showed little, if any, colocalization with CldU-substituted nascent DNA during all stages of S-phase (Figs 7,8,9).

The absence of pRb within nuclear DNA replication sites argues against a direct role for pRb in this process. This view is supported by the notion that DNA replicates during S-phase when pRb is normally rendered inactive by hyperphosphorylation (Ezhevsky et al., 2001Go; Lundberg and Weinberg, 1998Go). Furthermore, our immunocytochemical results are in agreement with the observations reported by Goodrich et al. (Goodrich et al., 1991Go) on the in vivo physiological effect of active pRb on DNA replication. In these experiments, the microinjection of purified functional pRb into pRb-negative cultured mammalian cells prevented G1 cells from entry into S-phase, but did not have any major effect on those cells, which were already in S-phase. We conclude that pRb does not have a direct role in mammalian DNA synthesis during S-phase.

Our results, however, do not rule out the possibility that pRb might have an indirect role in certain aspects of S-phase regulation in normal cells or under conditions of stress (Knudsen et al., 2000Go; Sever-Chroneos et al., 2001Go). For example, it has been shown that pRb, independent of its phosphorylation status, associates with (Pradhan and Kim, 2002Go; Robertson et al., 2000Go) and inhibits the activity of the maintenance DNA methyltransferase (Dnmt1) (Pradhan and Kim, 2002Go). Since active Dnmt1, but not pRb, is present at DNA replication sites (Leonhardt et al., 1992Go; Rountree et al., 2000Go), it is possible that pRb and Dnmt1 associate in the nucleosolic compartment. Thus, pRb might modulate Dnmt1 activity by controlling the levels of chromatin-bound functional Dnmt1. The importance of properly regulated Dnmt1 is underscored by the observation that deregulated methyltransferase activity accompanies neoplastic transformation of mammalian cells (El-Deiry et al., 1991Go; Kautiainen and Jones, 1986Go).

An alternative role for pRb in the regulation of genome replication is suggested by the finding that human pRb interacts in vitro with a member of the Mcm family of proteins, Mcm7 (Sterner et al., 1998Go). Mammalian Mcm proteins begin to load onto chromatin in late telophase (Dimitrova et al., 2002Go; Dimitrova et al., 1999Go) in a process known as replication licensing (Blow, 2001Go). It is generally believed that Mcm-s, together with ORC, Cdc6 and Cdt1, assemble into pre-replication complexes (pre-RCs) at chromosomal replication origins. The loading of Mcm-s during late telophase completes the assembly of fully functional pre-RCs and the licensing capacity of mammalian nuclei remains unchanged throughout G1-phase (Dimitrova et al., 2002Go). By analogy with the role of Rb family members in transcriptional repression during G1-phase, which at least in part is mediated by association with promoter-bound E2F protein family members (Dyson, 1998Go; Wells et al., 2000Go; Zheng and Lee, 2001Go), association of pRb with origin-bound Mcm family members could represent a potential mechanism for keeping the origins dormant until a protein kinase nuclear environment favoring activation of the origins has been established at the G1/S-phase transition. To date, evidence for the existence in vivo of pRb-Mcm7 complexes, however, has not been presented. Hence, we decided to investigate whether pRb and Mcm proteins are found in close proximity within nuclei of primary and transformed mammalian cells.

Mcm proteins bind tightly to licensed chromatin and this interaction is resistant to mild detergent extraction (Dimitrova et al., 2002Go; Dimitrova et al., 1999Go). Therefore, in order to look at the active, possibly origin-bound form of Mcm-s, we subjected exponentially growing cultures of WI38 and HeLa cells to a Triton X100 extraction procedure prior to fixation, which removes all soluble or loosely bound cytoplasmic and nuclear proteins. To mark DNA replication sites, the cells were pulse-labeled with CldU immediately before extraction. After fixation with formaldehyde, the cells were triple labeled with antibodies specific for pRb, CldU and one Mcm family member (Mcm2, 3 or 7). Representative results of WI38 cells with anti-Mcm7 antibodies are shown in Fig. 8. Identical results were obtained with HeLa cells and with Mcm2- and Mcm3-specific antibodies.

Even though we used asynchronous cell cultures, it is possible to classify precisely the cells according to the cell cycle stage. G1 nuclei are CldU-negative and Mcm-positive, since the Mcm proteins are tightly bound to chromatin and, therefore, resistant to Triton extraction during G1-phase. S-phase nuclei are CldU-positive and can further be categorized into early-, mid- or late-S subgroups based on the characteristic replication patterns and on the decreasing amount of chromatin-bound Mcm-s. Finally, G2-phase nuclei (not shown) are both CldU- and Mcm-negative. The mild extraction procedure, which we applied, removed also a significant fraction of pRb. Nevertheless, most nuclei, including those in S-phase, remained positive for pRb, even though the fluorescence intensity varied between individual nuclei and was generally low in CldU-positive nuclei. pRb is known to associate tightly with the nuclear matrix only during G1-phase when it is hypophosphorylated and active (Mancini et al., 1994Go). Unlike nuclear matrix results obtained through the application of high salt extraction procedures, our experiments show that a small fraction of pRb resists low salt/detergent extraction also during S-phase. This observation seems to contradict previously published observations on the low-salt extractability of pRb (Mittnacht and Weinberg, 1991Go). We do not know the exact reason for this discrepancy, but we believe that differences in buffer composition and extraction procedures may provide a potential answer. We note that hypotonic treatment used in previous studies (Mittnacht and Weinberg, 1991Go) is known to disrupt nuclear architecture (Zatsepina et al., 1997Go), whereas the cytoskeleton buffer used by us may preserve molecular interactions better. Even though unexpected, our observations that small amounts of pRb resist low-salt extraction during S-phase are corroborated by a recently published report (Wells et al., 2000Go), which found that, surprisingly, Rb family proteins are bound in vivo to a number of gene promoters at this cell cycle time. The implications of these discoveries remain to be unveiled.

Consistent with previous reports on the lack of colocalization between Mcm proteins and DNA replication sites (Dimitrova et al., 1999Go; Krude et al., 1996Go; Todorov et al., 1995Go), we did not observe spatial coincidence between chromatin bound Mcm-s and CldU-labeled nascent DNA during any stage of S-phase (Fig. 8). Importantly, pRb and Mcm7 also did not significantly overlap (Fig. 8). Identical results were obtained with Mcm2- and Mcm3-specific antibodies (data not shown). To address the possibility that Mcm-s and pRb might interact within the soluble nucleoplasmic fraction, we also conducted experiments where the cells were fixed directly after CldU pulse-labeling without prior detergent extraction (Fig. 9). Again, no significant overlap was detected between the pRb- and Mcm-specific immunofluorescent signals. Thus, the lack of spatial proximity between pRb and Mcm proteins makes it unlikely that extensive interactions between these proteins take place in vivo within mammalian nuclei. This notion is further strengthened by the observation that microinjection of a functional pRb mutant with deleted N-terminal part (which mediates the interaction with Mcm7) is sufficient to block cultured human cells in G1-phase and to prevent their entry into S-phase (Goodrich et al., 1991Go). Furthermore, expression of a constitutively active form of pRb resistant to Cdk phosphorylation has no effect on the establishment or maintenance of mammalian pre-RCs (Sever-Chroneos et al., 2001Go). The significance of the biochemically detected interactions between pRb and Mcm7 remains unclear at present (Sterner et al., 1998Go).

In conclusion, the results presented in this report are in good agreement with a plethora of previous reports in the literature, in which the replication patterns during S-phase were studied individually in primary, immortal or transformed mammalian cells. We conducted a comprehensive study of the distribution of DNA replication sites in over a dozen mammalian cell lines with different proliferation capacities. Through the use of four independent approaches for visualization of these sites, we have provided an explanation for some puzzling discrepancies in the literature and have convincingly demonstrated that there are no fundamental differences in the distribution of DNA replication sites and the spatio-temporal sequence of replication patterns between normal, immortalized or transformed mammalian cells. These studies further demonstrate that pRb and nuclear lamin proteins A, B and C are not significantly associated with replication sites in either early-S or any other stage of S-phase. Moreover, no evidence was found for limited perinucleolar foci containing labeled replicating DNA, nuclear lamins and/or pRb as reported by Kennedy et al. (Kennedy et al., 2000Go). Simultaneous examination of Mcm proteins and pRb enabled us to extend these studies to determine whether pRb, together with Mcm-s, while not associating with active replication sites, might associate with replication origins and thus play a potential role in origin regulation or in the transformation of pre-RCs into active replication complexes. Our findings show for the first time the lack of association of pRb with Mcm-labeled sites in G1-phase, as well as throughout S-phase.

The validity of the major replication patterns described in this and many past studies is further substantiated by recent in vivo studies, whereby the dynamics of DNA replication sites are visualized directly in live cells through the microinjection of fluorescent nucleotides (Manders et al., 1999Go; Pepperkok and Ansorge, 1995Go) or through the stable transfection of cells with GFP-tagged variants of protein components of the replication machinery (Leonhardt et al., 2000Go; Somanathan et al., 2001Go). Taken together with the findings of similar distributions of replication sites in other eukaryotes, including plant cells (Fuchs et al., 1998Go; Fujishige and Taniguchi, 1998Go; Lafontaine and Lord, 1974Go; Sparvoli et al., 1976Go), we conclude that replication patterns are generally conserved in at least the higher eukaryotic organisms. Observed differences occur during the mid/late stages of S-phase and possibly reflect differences in DNA sequence content and unique chromatin organization in different cell types, rather than the stage of transformation or the replicative capacity of the cells. Deciphering the mechanistic basis for this highly conserved spatio-temporal programming for the replication of the genome and its associated replication factors within the context of a global nuclear architecture is an exciting challenge for future investigation.


    Acknowledgments
 
We are indebted to M. Stachowiak, S. Matsui, D. Kaufman, T. Beerman and B. Nicholson for the gift of cell lines, to G. Blobel for the anti-lamin antibodies and to T. Melendy and D. Goodrich for anti-pRb antibodies. This work was supported by NIH grant GM23922 to R.B.


    References
 Top
 Summary
 Introduction
 Materials and Methods
 Results and Discussion
 References
 

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