Visualizing the dynamics of chromosome structure

Chromosoma DOI 10.1007/s00412-007-0109-5

RESEARCH ARTICLE

Visualizing the dynamics of chromosome structure formation coupled with DNA replication Eisuke Gotoh

Received: 2 February 2007 / Revised: 2 April 2007 / Accepted: 18 April 2007 # Springer-Verlag 2007

Abstract A basic question of cell biology is how DNA folds to chromosome. Numbers of examples have suggested the involvement of DNA replication in chromosome structure formation. To visualize and identify the dynamics of chromosome structure formation and to elucidate the involvement of DNA replication in chromosome construction, Cy3-2′-deoxyuridine-5′-triphosphate direct-labeled active replicating DNA was observed in prematurely condensed chromosomes (PCCs) under a confocal scanning microscope utilized with drug-induced premature chromosome condensation (PCC) technique that facilitates the visualization of interphase chromatin as condensed chromosome form. S-phase PCCs revealed clearly the drastic dynamics of chromosome formation that transits during Sphase from a ‘cloudy nebula’ to numerous numbers of ‘beads on a string’ and finally to ‘striped arrays of banding structured chromosome’ along with the progress of DNA replication. The number, distribution, and shape of replication foci were also measured in individual subphases of Sphase more precisely than reported previously; maximally, ∼1,400 foci of 0.35 μm average radius size were scored at the beginning of the S-phase, and the number reduced to ∼100 at the end of the S-phase. Drug-induced PCC clearly provided Communicated by J. Diffley E. Gotoh (*) Division of Genetic Resources, National Institute of Infectious Diseases, 1-23-1, Toyama, Shinjuku-ku, Tokyo 162-8640, Japan e-mail: [email protected]

E. Gotoh Department of Radiology, Jikei University School of Medicine, 3-25-8, Nishi-Shimbashi, Minato-ku, Tokyo 116, Japan

the new insight that eukaryote DNA replication is tightly coupled with the chromosome condensation/compaction for the construction of the higher-ordered structure of the eukaryote chromosome.

Introduction Chromosome condensation/compaction is a basic and profound principle of cell biology. Much recently accumulated evidences have implied that eukaryote DNA replication/ transcription is involved in the compaction of chromosomes (Zink et al. 1998; Manders et al. 1999; Samaniego et al. 2002); however, its involvement remains unclear. Eukaryote DNA replication commences at thousands of multiple replication origins at the beginning of the S-phase and then proceeds spatially and temporally synchronizing in an ordered manner through the S-phase. Cytogenetical observation of the replicating DNA in the nucleus is certainly a most direct approach to elucidate the dynamics of chromosome formation and DNA replication. In earlier studies, the fiber autoradiography of DNA labeled with [3H]thymidine revealed the synchronized synthesizing replicon of size ranging from 50 to 300 kbp (Fakan and Hancock 1974; Edenberg and Huberman 1975; Hand 1978). Then, the thymidine analogue bromodeoxy uridine (BrdU) has been introduced to localize and measure the replication foci (Nakamura et al. 1986; Mills et al. 1989; Nakayasu and Berezney 1989). More recently, the replication regions and chromosome formation in living cells were visualized using Cy5-2′-deoxyuridine-5′-triphosphate (dUTP) directly labeled fluorescent DNA (Manders et al. 1999). The recent development of a laser scanning microscopy (LSM) and new fluorochromes such as Cy3 and Cy5 have much improved the resolution of the observed DNA replication regions.

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The accumulated findings of the dynamic organization of the replication factory in the nuclei have raised interest in replication DNA and how it is folded within the interphase nuclei during S-phase and how the folding changes lead to a higher-ordered whole chromosome. Numbers of supporting examples have actually showed the relationship of DNA replication and chromosome conformation (Zink et al. 1998; Manders et al. 1999; Samaniego et al. 2002). However, the results shown in these reports were derived from the observation in interphase nuclei; thus, the resolution and the resulting information were limited because chromosomes are only visible in the mitotic stage of the cell cycle and are invisible as decondensed structure in interphase (Manders et al. 1999; Gotoh and Durante 2006). Premature chromosome condensation (PCC) is a useful and unique technique that allows the interphase nuclei to be visualized as a condensed form of mitotic chromosome (Johnson and Rao 1970; Johnson et al. 1970; Sperling and Rao 1974b; Gotoh and Durante 2006); thus, a lot of studies including DNA replication and DNA repair have been achieved using the PCC method (Hittelman and Rao 1976; Rao et al. 1977; Hanks and Rao 1980; Mullinger and Johnson 1980; Lau and Arrighi 1981; Mullinger and Johnson 1983; Hittelman and Pollard 1984; Hittelman 1986). Conventional PCC has been carried out by cell fusion using either fusogenic viruses (Johnson and Rao 1970) or polyethylene glycol (Pantelias and Maillie 1983; cell fusion-mediated PCC), but this use is usually problematic, and resulting chromosomes are a mixture of those of the inducer and recipient cells (Gotoh and Durante 2006). These drawbacks of the technique have been recently overcome with a much easier and more rapid technique using calyculin A or okadaic acid, specific inhibitors of protein phosphatases (drug-induced PCC technique; Gotoh et al. 1995; Gotoh and Asakawa 1996; Asakawa and Gotoh 1997; Durante et al. 1998; Gotoh and Durante 2006). Druginduced PCC is now becoming much more popular and has been used in a wide range of cytogenetic applications (Gotoh and Asakawa 1996; Asakawa and Gotoh 1997; Gotoh et al. 1999; Terzoudi et al. 2003; El Achkar et al. 2005; Gotoh and Tanno 2005; Srebniak et al. 2005; Terzoudi et al. 2005). The morphology of prematurely condensed chromosomes (PCCs) of the interphase nucleus depends on the cell-cycle position at the time of PCC induction either by cell fusion or drugs (Johnson and Rao 1970; Gotoh et al. 1995; Gotoh and Durante 2006); G1-, S-, or G2-PCCs show the characteristic appearance: (1) G1-phase nuclei produced univalent (i.e., unreplicated) chromosomes (G1-PCCs), (2) Sphase nuclei converted to a ‘pulverized’ appearance that consisted of univalent and bivalent (i.e., replicated) chromosomes (S-PCCs), and (3) G2-phase nuclei showed bivalent chromosomes, similar to mitotic chromosomes, although less

condensed (G2-PCCs) (Gotoh and Durante 2006). This cellcycle-dependent morphological transition is typically seen through the S-phase: (1) In early S, thin filamentous fibers spread widely, (2) in middle S, the ‘pulverized’ appearance of chromosomes consist of thin and thick chromatin fiber, and (3) in late S, most of the chromosomes condensed to show bivalent chromosomes like mitotic chromosomes (Johnson and Rao 1970; Rao 1977; Rao et al. 1977). These different characteristic appearances seen in different subphases of S-PCCs might be useful for identifying the involvement of chromosome formation and DNA replication in individual substages of the S-phase; thus, PCC is a possible way to address the above question. In the present study, we utilized the drug-induced PCC technique; Cy3-dUTP direct-labeled fluorescent DNA of individual substages of S-phase nuclei was condensed prematurely, and the dynamics of chromosome condensation/ compaction and the distribution of replication foci on chromosomes was analyzed. The results strongly suggested that DNA replication and chromosome condensation/compaction are tightly coupled in constructing the higher-ordered structure of the eukaryote chromosome. A hypothetical model for the integration of DNA replication and chromosome conformation will be also discussed.

Materials and methods Cell, chemicals and antibodies Human normal karyotype fibroblast cell line GM05389 was obtained from Coriell Cell Repositories (New Jersey). Calyculin A was purchased from Wako Chemicals (Osaka, Japan) and dissolved in 100% ethanol; 100 μM of stock solution was stored at −20°C. Cy3-dUTP was purchased from Amersham (Upsala, Sweden). Sizes of 425–600-μmdiameter acid-washed glass beads were from Sigma (Missouri). SC56 (anti-proliferating cell nuclear antigen [PCNA] mouse monoclonal IgG) was purchased from Santa Cruze Biochemicals (California), and fluorescein isothiocyanate (FITC)-conjugated antimouse-IgG goat-IgG was purchased from Jackson ImmunoResearch Lab (Pennsylvania). Cy3-dUTP labeling by a bead-loading method Cy3-dUTP was loaded in the cells using a bead-loading procedure as described previously (McNeil and Warder 1987; Manders et al. 1999; Ito et al. 2002). The procedure facilitates incorporation of the analogues in the cell nucleus in a very short time after transient permeabilization of the cell membranes (Manders et al. 1999). Human fibroblast cells GM05389 were seeded on a 35-mm glass-base culture dish (Iwaki, Japan) designed for observation under inverted

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microscopes and were maintained exponential and asynchronously in minimum essential medium (MEM) supplemented with 15% fetal calf serum at 37°C in 5% CO2 atmosphere with 95% humidification. The culture medium was replaced with 37°C-prewarmed 50 μl of 10 μM Cy3-dUTP in MEM medium; 425–600-μm-diameter glass beads were sprinkled onto the cells, tapping and rocking the dish several times, and then the beads were rinsed off with prewarmed phosphatebuffered saline (PBS; 37°C). It usually took a couple of minutes to complete the loading procedure. Prewarmed (37°C) MEM was added to the culture and incubated for 10 min after starting of Cy3-dUTP loading. Then, cells were subjected to PCC using calyculin A or were immunostained.

Laser scanning microscopy Blue, green, or red confocal images were collected using a Confocal Laser Scanning microscopy system Zeiss LSM510 (Jena, Germany) equipped on an inverted epifluorescence microscope Zeiss Axiovert100 M (Jena, Germany). A water immersion objective lens Zeiss C-Apochromat 63×/1.2 w corr. (Jena, Germany) was used. UV, Ar, or He–Ne laser was tuned at 366, 488, 568 nm to excite DAPI, FITC, and Cy3, respectively. The fluorescence signals from each fluorochromes were recorded separately in multiple scans to minimize optical cross-talk. Digital images were manipulated using Zeiss LSM510 software (Jena, Germany), then processed using Adobe Photoshop (California) for the figures.

Immunostaining After Cy3-dUTP loading, immunostaining for PCNA detection was done as follows: Cells were rinsed three times with ice-cold PBS, then fixed with increasing series of −20°C cold ethanol (50, 70, 95, and 99.5%). Cells were then rehydrated with decreasing series with −20°C cold ethanol (95, 70, and 50%) and washed three times with ice-cold PBS. Cells were incubated with anti-PCNA mouse monoclonal IgG, SC56, and diluted 100 times in 0.2% NP40 in PBS, for 30 min at room temperature. Cells were washed twice with 0.2% NP40 in PBS, then incubated with FITC-conjugated antimouse-IgG goat IgG, and diluted 2,000 times in 0.2% NP40 in PBS, for 30 min at room temperature. Finally, cells were washed three times with 0.2% NP40 in PBS, then observed under a confocal microscope. Premature chromosome condensation Immediately after the Cy3-dUTP loading procedure, chromosomes were condensed prematurely using 50 nM of calyculin A. PCC using calyculin A was carried out as described elsewhere (Gotoh et al. 1995; Asakawa and Gotoh 1997; Johnson et al. 1999; Ito et al. 2002) except that the incubation time was kept as short as possible (10 min) to obtain a high spatial and temporal resolution, otherwise, replication foci can be merged together resulting in a less spatial/temporal resolution. For cytogenetic scoring, 30 min incubation was used to obtain sufficient number of chromosomes (Gotoh et al. 1999; Gotoh and Tanno 2005), but chromosomes begin to condense within the first 5 min of incubation (Gotoh et al. 1999), and a substantial number of chromosomes condensed after 10 min incubation. After incubation with calyculin A, cells were harvested, swollen in 75 mM KCl for 10 min at 37°C, fixed three times with methanol/acetic acid (3:1), dropped on a glass slide, and air dried. Finally, DNA was counterstained with 200 ng/ml of DAPI, mounted with PBS, and covered with a coverslip.

Results Observation of replication foci in S-phase interphase nuclei Cy3-dUTP was loaded into the exponentially growing cells as described previously (McNeil and Warder 1987; Manders et al. 1999; Ito et al. 2002). This method allows the replicating DNA to be Cy3-fluorescently imaged within a very short lapse of time. Figure 1 shows the distribution of Cy3fluorescent DNA in interphase nuclei observed under a laser scanning confocal microscope. Cy3-dUTP was actually incorporated during the 10 min of the bead-loading procedure. PCNA was detected simultaneously to confirm that the Cy3-dUTP-incorporated regions are the replication foci (Essers et al. 2005). The Cy3-labeled nuclei showed the diagnostic distribution for different subphases of the Sphase as described in the previous reports using BrdUlabeled DNA (Nakamura et al. 1986; Nakayasu and Berezney 1989) or Cy5-dUTP-labeling DNA (Manders et al. 1999): (1) In early S, hundreds of small and discrete foci were evenly distributed over the nuclei (with the exception of nucleoli regions; Fig. 1c), (2) in middle S, the foci fused together resulted in the size increased and the number decreased (Fig. 1g), (3) in late S, the size of foci further increased, the number of foci decreased, and the shape became more irregular (Fig. 1k), and (4) in very late Sphase, the size of foci increased much, the number of foci was markedly decreased to about ten, some of them showed a characteristic appearance of ring- or horseshoe-like structures (Nakamura et al. 1986; Manders et al. 1999) (Fig. 1o, indicated by arrows), and most of foci localized at nuclear periphery regions. PCNA distribution was well consisted with Cy3-dUTP foci particularly in the early Sphase (Fig. 1b), but after the middle S-phase, some discrepancy was seen between PCNA and Cy3 localizations (Fig. 1f,j,n). This discrepancy was similar as previously reported (Bravo and Macdonald-Bravo 1987), and the

Chromosoma Fig. 1 Localization of DNA replication foci and PCNA in interphase nuclei of individual subphase of S-phase. From left to right column, a, b, c, d early S-phase, e, f, g, h middle Sphase, i, j, k, l late S-phase, and m, n, o, p very late S-phase. a, e, i, m Nuclear DNA was counterstained with 200 nM of DAPI. b, f, j, n PCNA was detected using SC56 (antiPCNA mouse-IgG) and FITC-conjugated antimouse-IgG goat-IgG. c, g, k, o Replicating DNA was directly labeled using Cy3-dUTP by the bead-loading method (McNeil and Warder 1987; Manders et al. 1999). Note that there was absolutely no background Cy3 signal in the Cy3-dUTP unincorporated nucleus (e, f, g, h, indicated by arrow). d, h, l, p Merged image of DAPI, FITC, and Cy3. Scale bar, 5 μm

reason is still unclear, but the function of PCNA might be different in the early- and after the middle-S-phase (Bravo and Macdonald-Bravo 1987). Observation of replication foci on prematurely condensed S-phase chromosomes The replication foci distribution was fairly well defined in interphase nuclei as shown in Fig. 1; however, little was known about how replicating DNA is folded to higherorder chromosomes because chromosomes are invisible in interphase as they decondense. To visualize the chromosome compaction dynamics and replication regions more precisely in S-phase chromatin, the advantage of the drug-induced PCC method was taken (Gotoh et al. 1995; Asakawa and Gotoh 1997; Johnson et al. 1999; Ito et al. 2002). The cells were not synchronized using a DNA synthesis inhibitor such as thymidine because cell synchronization can give some bias in DNA replication, and consequently, all phases of replication can be observed. The individual substages of the S-phase can be easily identified by typical diagnostic appearances seen in different phases of S-PCCs (Mullinger and Johnson 1983; Gollin et al. 1984; Hameister and Sperling 1984; Savage et al. 1984;

Gotoh et al. 1995; Gotoh and Durante 2006). A drastic conformational change of chromosome structure formation coupled with the replication foci distribution was, as shown in Fig. 2, clearly revealed in PCCs after Cy3-dUTP loading. The Cy3-dUTP loading procedure was finished within 10 min followed by 10 min of PCC induction and fixation. Accordingly, only DNA replicated in this short lapse fluoresces. Thus, the observed S-PCCs in the present study reflected the replication stages at most 20 min (10 min for Cy3-dUTP labeling and 10 min for PCC induction; see “Materials and methods”) before the cell fixation. The different diagnostic appearance of different S-PCCs facilitates the DNA replication stages to be identified precisely. The size and distributions of Cy3-dUTP seen on PCCs seemed to be well comparable to those seen in interphase nuclei (see to compare, Figs. 1 and 2), but more clear and precise spatial and temporal distribution of replication foci on chromosomes were clearly seen on PCCs studies: (1) In early S-phase, PCCs showed a cloudy spreading mass of thin fibers like a ‘nebula.’ Numerous fine granular foci were distributed homogeneously on overall the fibers (Fig. 2c), showing a ‘beads on a string’ appearance that are similar to the same named or ‘particles on a string’ structures observed under an electron microscope (Olins and Olins 1974;

Chromosoma Fig. 2 DNA replication regions on prematurely condensed chromosomes (PCCs) of different substages of S-phase. Ten minutes after Cy3-dUTP loading, cells were condensed prematurely using 50 nM of calyculin A (Gotoh et al. 1995). From left to right column, a, b, c early S-phase PCCs, d, e, f middle S-PCCs, g, h, i late S-PCCs, and j, k, l very late S-PCCs. a, d, g, j DAPI-counterstained DNA, b, e, h, k Cy3dUTP-labeled DNA replication region, and c, f, i, l merged image of DAPI and Cy3. l Centromeric region (arrow) or telomeric region (arrowhead) replicates in very-late S-phase are indicated. i, l Late S- and very late S-PCCs already condensed as mitotic chromosomes, but these PCCs were actually Sphase chromosomes because they incorporated Cy3-dUTP. G2/M chromosomes are easily distinguished from late or verylate S chromosomes as G2/M chromosomes do not incorporate Cy3-dUTP (data not shown). Inset in c is a higher magnification of the boxed portion. Scale bar, 10 μm. DNA replication regions seen on prominent fiber of PCCs. m early-S-phase (918 foci scored) and n middle Sphase (707 foci scored). Replication foci are clearly seen as ‘beads on a string’ structure, some of these are indicated by an arrowhead. Scale bar, 10 μm

Thoma et al. 1979). (2) In middle S-phase, typical ‘pulverized’ PCCs were recognized, and the size of foci increased with the number of foci decreased and distributed unevenly on chromosomes. The foci became much brighter (Fig. 2f). (3) In late S-phase, chromosomes showed mostly condensed like mitotic chromosomes. Cy3-dUTP incorporated regions were recognized as band arrays inserted in the condensed chromosome (Fig. 2i, indicated by arrows). The similar appearance of replication foci along longitudinally on chromosomes were previously reported on metaphase of kangaroo-rat kidney PtK1 cells (Ma et al. 1998). The size of foci increased, and the number decreased to the level that was easy to score. (4) In very late S-phase, the number of foci further reduced and predominantly localized at centromeric or telomeric regions (Fig. 2l, indicated by arrows). These regions are actually known as satellite heterochromatic DNA regions where DNA replicates at very late S

(O’Keefe et al. 1992). Although PCC was induced in the present study using phosphatase inhibitor calyculin A, the different foci distributions should not be an artifact by the chemical because Cy3-dUTP was already incorporated in the replicating DNA at the end of the loading procedure before the start of PCC by calyculin A (see the results obtained in interphase nuclei, Fig. 1). The number and the size of replication foci seen in individual different subphase of S-PCCs correlated well with those seen on corresponding interphase nuclei, which facilitated identification of the stage of S-PCCs and that of corresponding interphase nuclei. It is also notable that the size of the replication foci in each stage seems well correlate to the degree of condensation/compaction of chromosomes, as proposed by the earlier study (Buongiorno-Nardelli et al. 1982): Very fine granular replication foci were distributed on thin chromatin fiber that formed ‘beads on a string’ in the early

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S-phase (Fig. 2c), whereas in the late S-phase, larger and fused foci were flanked with thickly condensed chromatin and showed ‘band-arrayed chromosomes’ as G- or R-banding chromosomes (Fig. 2f). In the present study, PCC was conducted using phosphatase inhibitor calyculin A; therefore, there still remains some possibility that the chemical may boost or accelerate compaction of chromosomes in particular in the late S-phase. Nevertheless, the different degree of condensation seen in different subphases of SPCCs should reflect the different degree of chromosome conformation of individual subphases because all cells were subjected to PCC by exactly the same time (10 min) of calyculin A exposure. Sarkar et al. (2002) also tried to induce artificial chromosome condensation in interphase using microinjection of trivalent cations (cobalt hexamine trichloride), although the chemical was different from calyculin A. They showed that the part of nucleus exposed to trivalent cations transiently condensed then decondensed as the cations diffused out of the nucleus. The experiment was done in interphase nuclei of living cells; therefore, images were not clear because of strong artificial halos, but the result seems to suggest that chromosome conformational changes are already introduced in interphase nuclei as the progress of DNA replication. Therefore, it might be more likely that DNA replication and chromosome formation couple together, rather than independently proceed each other. PCNA detection was also tried on PCCs, but the signals were too faint to observe (data not shown). Presumably, PCNA loosely bound to the replication factory at this stage and might detach during hypotonic treatment (75 mM KCl) done for chromosome preparation.

Table 1. In previous reports, 100–350 sites were scored in the early S-phase (Nakayasu and Berezney 1989; Manders et al. 1996; Manders et al. 1999). Other studies, using a LSM coupled to multidimensional image analysis, scored ∼1,100 replication foci (Jackson and Pombo 1998; Ma et al. 1999). However, the scoring in these studies was done on interphase nuclei. In the present study, the number of foci scored a maximum of ∼1,400 in the early S-phase that was much precise than ever reported, then decreased to ∼100 in the very late S-phase. The size changing of foci during S-phase was also shown clearly. While the maximum size of foci changed from early to late S-phase, the minimum size changed less through the S-phase. Therefore, the 0.08–0.2-μm size of replication foci could be a fundamental unit of replication. The spacing between replication foci clusters was ranged from 85.3 (0.64 μm) to 536 kbp (4.02 μm; average, 208 kbp, 1.56 μm) based on the assumption that the 0.75-μm length of a 30-nm chromatin fiber is equivalent to 100 kbp (linear packing ratio of ∼40:1; Berezney et al. 2000). Fiber autoradiography of DNA labeled with [3H]thymidine revealed the synchronized synthesizing replicon of spacing distance ranging from 50 to 300 kbp (Fakan and Hancock 1974; Edenberg and Huberman 1975; Hand 1978). Therefore, the spacing of replication foci seems to vary more than previously reported values. The measure of the spacing of foci were also reported on stretched DNA fiber with detergent, and the average distance of foci was calculated as 144 kbp (Jackson and Pombo 1998), which is in fairly good agreement with the present and the previous studies, although the intact structure of foci was no more retained in the stretched fiber after lysis treatment.

The number, size and spacing of replication foci in different subphase of S

Discussion

The ‘beads on a string’ structure was more evidently seen on the prominent fiber shown in Fig. 2m,n, and the distance between foci was easily measured than those seen in interphase nuclei. The number, size, and the spacing of the foci of different S-phase stages were summarized in

In the present study, the dynamics of DNA compaction and DNA replication foci distribution were analyzed precisely in interphase nuclei by Cy3-dUTP direct-labeling method. In particular, we took advantage of the drug-induced PCC method (Gotoh and Durante 2006); the chromosomal conformational change and DNA replication foci distribu-

Table 1 The number, size, and spacing of replication foci of different subphase of S Subphase of S

Early

Middle

Late

Very late

Number of foci (max–min)a Size of foci (max–min; μm radius)b,c Spacing between foci (max–min; μm)b,c

1,046±187 (1,396–758) 0.35±0.01 (0.08d–0.62) 1.56±0.68 (0.64–4.02)

678±83 (887–601) 0.52±0.06 (0.1d–0.62) 1.01±0.11 (0.33–1.86)

450±132 (607–237) 0.98±0.23 (0.24–1.44) N.D.e

153±42 (245–98) 1.1±0.44 (0.32–4.1) N.D.e

a

For each subphase, at least 12 spreads were scored, except for very late S (ten spreads to be scored were available). Measured using the Zeiss LSM510 software c Randomly selected 20 points were measured, and the average and the error were calculated. d These data were beyond the resolution of optical microscope and measured on digitized images. e The measure was not done because PCCs at these stages mostly condense as mitotic chromosomes (see Fig. 2), thus the measure of the spacing seems less meaningful. b

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some condensation (Loupart et al. 2000; Marheineke and Hyrien 2001). However, other replication protein such as PCNA and Rfc4 did not influence the chormosome condensation (Krause et al. 2001; Prasanth et al. 2004). Therefore, little is still known about how the chromatin is folded within the interphase or how the folded chromatins change as chromosomes form (Cook 1995). The morphological change of chromosome structure coupled with the transition of replication foci distribution, as shown in Fig. 2, seems to support the tight coupling of DNA replication and chromosome structure formation. Previously reported evidence and the results of this study show that: (1) The different appearance of condensation in different phase of S-PCCs is thought to be depend on the different degrees of chromosome conformation at the time of induction of PCC (Johnson and Rao 1970; Rao 1977; Rao et al. 1977). In late or very late S-phase, in particular, chromosome conformation already changes like mitotic chromosomes (Fig. 2f in the present study). (2) Chromosomes condense asynchronous and the different degree of condensation is depend on the time of chromatin replication (Kuroiwa 1971). (3) Chromosomes are not fully diffused but are compartmentalized in interphase nuclei (Cremer et al. 1993; Ferreira et al. 1997; Berezney et al. 2000), and each chromosome occupy the ‘territory’ and do not intermingle (Hadlaczky et al. 1986; Cremer et al. 1993; Swedlow and Hirano 2003). (4) Late replication foci were prealigned during interphase. They moved only subtly to

tion during the S-phase was clearly seen on PCCs of different sub-S-phase. Therefore, drug-induced PCC would be a useful tool that provides new insights of the dynamics of chromosome formation and DNA replication. More accumulated evidence suggested the role of DNA replication in chromosome condensation/compaction (Pflumm 2002). In earlier studies, the relationship of DNA replication and chromosome condensation was proposed by several studies, particularly by a fusionmediated PCC study (Sperling and Rao 1974a; Mullinger and Johnson 1983). However, the limited available methodologies at that time did not allow the precise mechanism to be cleared. More recently, molecular genetic studies have provided supporting evidence for the idea that mutation in genes (XCDT1, cdt1, Orc2, Orc3, Orc5, MCM10) required for DNA replication showed an abnormal phenotype in chromosome condensation (Maiorano et al. 2000; Nishitani et al. 2000; Pflumm and Botchan 2001; Christensen and Tye 2003; McHugh and Heck 2003; Prasanth et al. 2004) or aberrant replication timing resulting in abnormal chromoa

early S-phase

b

middle S-phase

'beads on a string'

R 'pulverized chromosome' c

late S-phase

'band arrayed structured chromosome' d

G2 to prophase

e

mitosis

replication factory (active;red, inactive;blue) unreplicated DNA (univalent chromosome) old replicated DNA (bivalent chromosome, Cy3-dUTP not incorporated) newly replicated DNA (bivalent chromosome, Cy3-dUTP incorporated) generated force pulling replication foci

Fig. 3 A hypothetical two-dimensional model for chromosome conformational change involving DNA replication based on the models proposed by Cook (1995) or Pflumm (2002). a Early S-phase. DNA replication starts at multiple origins and proceeds bidirectionally. Early S-PCCs are seen as ‘beads on a string’ appearance. b Middle S-phase. As DNA replication proceeds, replicated DNA passes through the replication factory, and some tension is generated. The generated tension may pull back the replication factories close together so as to release the tension. Replication factories may in turn fuse together, and chromosomes compact. Middle S-PCCs are seen as well-known ‘pulverized chromosomes’ appearance. c Late S-phase. Most of DNA finished replication, and conformation was changed. Late S-PCCs are seen as ‘tandem band arrayed structured chromosomes’ as mitotic chromosomes. d G2 to prophase. After finishing of DNA replication, chromosome conformation changed as mitotic chromosomes but still so elastic that they are packed in nucleus. Before fixation, each chromosome occupies individual chromosome territory (CT) in the interphase nucleus, thus observed as compartment regions (colored). e Mitosis. After prophase, chromosomes further shorten in the longitudinal axis of chromosomes, consequently a straight rod-shaped recognizable chromosome formed as usually seen by cytologists. For simplicity, the model is shown in two dimensions, and the scaling is arbitrary. The model intends not to depict actual events of chromosome conformation change but to help imagine how DNA replication is involved in chromosomal conformation. As the real chromosomes condense three-dimensionally, other elements such as coiling and helical winding should be considered together to construct a stereoscopic hierarchical structure of eukaryote chromosomes (Woodcock and Dimitrov 2001; Swedlow and Hirano 2003)

Chromosoma

generate recognizable chromosomes presumably because of the shortening of the longitudinal chromosome axis (Manders et al. 1999). (5) The gross structure of an interphase chromosome territories is directly related to that of the prophase chromosomes (Manders et al. 1999). (6) The structure of mitotic chromosomes and nuclear chromosome territories are closely related (Manders et al. 1999), and the different bands of mitotic chromosomes are presented as distinct domains regarded as subchromosomal foci within chromosome territories (Zink et al. 1999). (7) The global chromosome territories are conserved during cell cycling; although conflicts still remain, some studies reported that chromosome territories transmitted through mitosis (Manders et al. 1999; Gerlich et al. 2003; Gerlich and Ellenberg 2003), whereas others reported that positional relations of chromosome territories are lost either at mitosis (Walter et al. 2003) or early G1 (Essers et al. 2005). (8) The spatio-temporal organization of DNA replication is determined by the specific nuclear order of these stable chromosomal units (Sadoni et al. 2004). (9) Chromatin domains with the dimension of replication foci may be fundamental units of chromosomal architecture (Berezney et al. 2000). (10) DNA replication occurs at fixed sites, and replicated DNA move through the replication center (Berezney and Coffey 1975; Pardoll et al. 1980; Hozak et al. 1993). (11) DNA replication contributes to a longitudinal contraction of the chromosome axis (Hearst et al. 1998). (12) Functional replication origins are critical requirement for longitudinal condensation of the chromosome axis (Pflumm and Botchan 2001). The results presented in this study and the above previous findings strongly suggesting that DNA replication, nuclear organization, and chromosome condensation are mutually integrated to construct a higher-order structure of eukaryote chromosomes. Several models for eukaryotic chromosome architecture have been proposed (Marsden and Laemmli 1979; Woodcock et al. 1984; Woodcock and Dimitrov 2001; Swedlow and Hirano 2003; Kireeva et al. 2004), but they are still controversial, and many of things remain unclear. In addition, these models do not take account the role of DNA replication/transcription. DNA/RNA polymerase are known to be tightly immobilized to the replication/ transcription factories (Cook 1999). DNA polymerase is thought to ‘reel in template DNA and extrude replicated DNA,’ rather than the enzyme tracking along the DNA template (Hozak et al. 1996; Cook 1999). In the context of this model, some kind of mechanical tension force should be generated in the template DNA while DNA replication goes on, because the factory does not freely suspended in the nucleus; consequently, this force may pull and aggregate the replication foci of both sides, releasing the tension in the DNA strand, resulting in the formation of chromosomes as seen in mitosis. Based on the above mechanism

and the observed findings listed above, Fig. 3 shows a hypothetical model for the relationship of DNA replication and chromosome conformational changes, which shows how interphase chromatin is constructed to chromosomes observed in mitosis. During the S-phase, the chromosomal conformation changes as DNA replicates, and the chromosome formation would be mostly completed at the end of DNA replication (Fig. 3a,b,c). From G2 to prophase, chromosomes are still more elastic, less condensed, folded several times, and prealigned in interphase nuclei (Manders et al. 1999). Chromosomes, at these phases, would be observed as chromosome territories (Cremer et al. 1993; Berezney et al. 2000; Fig. 3d). Entering in mitosis, chromosomes would condense even more, as they shorten the longitudinal axis to form a solid and rod-shaped appearance of recognizable mitotic chromosomes (Manders et al. 1999) (Fig. 3e). In this model, conformational change is simply illustrated two-dimensionally, but the real architecture of chromosomes occurs three-dimensionally. It is mostly unclear how DNA replication/transcription conduct to make up a three-dimensional hierarchical structure of chromosomes coupled with twisting/folding/winding or other motions, and must be a most principle challenge in cell science.

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