Article |
Address correspondence to David M. Gilbert, Department of Biochemistry and Molecular Biology, State University of New York Upstate Medical University, 750 East Adams St., Syracuse, NY 13210. Tel.: (315) 464-8723. Fax: (315) 464-8750. E-mail: gilbertd{at}mail.upstate.edu
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Abstract |
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Key Words: ß-globin; DNA replication; cell cycle; heterochromatin; nuclear organization
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Introduction |
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To investigate the relationship between replication timing and subnuclear position, we have exploited the ability of Xenopus egg extracts to initiate replication within mammalian nuclei isolated from cells staged at any time during G1 phase. With nuclei isolated 1 h after mitosis, heterochromatic and euchromatic domains are replicated in this in vitro system in no particular order. However, with nuclei isolated 2 h after mitosis, the overall temporal order for replication of these domains is preserved in vitro (Dimitrova and Gilbert, 1999b). This time period (12 h after mitosis), designated as the timing decision point (TDP),* is coincident with the spatial repositioning of chromosomal domains within the nucleus, providing a provocative temporal coincidence between replication timing and subnuclear position. These previous experiments monitored the general positions of whole populations of chromosomal domains but did not examine individual genes or their relationship to transcription. Here, we compare the developmentally regulated ß-globin locus, which is transcriptionally silent and late-replicating in CHO cells (Taljanidisz et al., 1989), to the active and early-replicating dihydrofolate reductase (DHFR) gene locus. The ß-globin locus is an excellent candidate for a locus silenced by a developmentally regulated replication timing switch. At both the human and the mouse ß-globin locus, over 200 kb of DNA is early-replicating and DNaseI sensitive in erythroid cells, but late-replicating and DNaseI resistant in nonerythroid fibroblasts (Dhar et al., 1988; Epner et al., 1988). In mousehuman hybrids, general deacetylation and transcriptional silencing of the human ß-globin locus is accompanied by its localization adjacent to murine centromeric heterochromatin (Schubeler et al., 2000).
An important question is whether the ß-globin locus acquires the replication timing program of the heterochromatin domain that it juxtaposes, and whether this juxtaposition is required to delay its replication timing program. In this report, we demonstrate that the CHO ß-globin locus is localized close to the periphery of the nucleus and replicated in the middle of S phase, coincident with the replication of peripheral heterochromatin. By contrast, the DHFR locus is more internally located and is replicated within the first 30 min of S phase. We further demonstrate that the differential replication timing program of these two loci is established 12 h after mitosis and that, during this same period of time, the ß-globin locus is localized to the periphery of the nucleus. These results are consistent with a model (Gilbert, 2001; Heun et al., 2001) in which replication timing of at least some loci is determined by association with a heterochromatic subnuclear compartment.
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Results |
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An example of FISH results with differentially labeled DHFR and ß-globin probes is shown in Fig. 1
A. This example shows a nucleus with replicated doublets for both DHFR alleles and unreplicated singlets for both ß-globin alleles. To verify that these probes could effectively discern a difference in replication timing between the early-replicating DHFR and late-replicating ß-globin loci, the number of nuclei showing doublets for one or both of each allele was scored and the data were expressed in three ways (Fig. 1, BD). First, the total percentage of doublet and singlet alleles revealed more than twice as many doublets for the DHFR locus (Fig. 1B). One advantage of the FISH method is that each cell can be examined individually. In principle, doublets for a late-replicating locus should only be observed within nuclei that already contain doublets for an early-replicating locus. When the percentage of nuclei showing more doublet alleles for one locus versus the other (1:0, 2:0, and 2:1) were scored (Fig. 1 C), a 10-fold preference for nuclei with a majority of doublet DHFR loci was observed. This preference for DHFR doublets increased to 100-fold when the analysis was restricted to the
30% of S phase nuclei that displayed doublets for both alleles of one locus and singlets for both alleles of the other locus (Fig. 1 D). Although we cannot rule out the possibility that the ß-globin locus replicated before the DHFR locus in a few cells, the rare exceptions likely reflect the frequency with which DHFR doublets are obscured from vision. These results agree with previous studies of replication timing in CHO cells (Taljanidisz et al., 1989) and verify the utility of FISH methodology as an assay for replication timing at these loci.
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To more directly assess whether ß-globin doublets appear during replication of the peripheral heterochromatin, the percentage of doublets for DHFR versus ß-globin was scored as a function of the replication pattern during S phase in nuclei that were simultaneously stained for BrdU and subjected to FISH. To accomplish this, we had to be confident that the distorted replication patterns shown in Fig. 2 A (FISH + BrdU) could be correctly categorized into early, middle, and late patterns. We defined early patterns as those displaying diffuse BrdU staining throughout the interior of the nucleus but noticeably absent BrdU staining at the nuclear periphery. Middle-replicating patterns were defined as nuclei with BrdU staining at the periphery and internal areas with significantly less BrdU staining. Late replication patterns displayed several large foci. Using these criteria, it was found that the percentage of nuclei displaying each pattern at various times during S phase was very similar to that shown in Fig. 2 B. Next, the percentage of either DHFR (Fig. 2 D) or ß-globin (Fig. 2 E) alleles present as doublets within nuclei of each pattern was scored. As expected, DHFR doublets appeared in nuclei with early replication patterns within 30 min after the release from the aphidicolin block. Middle and late replication patterns were first observed at 4.5 and 6.5 h into S phase, respectively, and a high percentage of these nuclei harbored DHFR doublets at all time points after their appearance. By contrast, ß-globin doublets were not observed above background levels in nuclei with early replication patterns at any time during S phase. Nuclei with middle-replicating patterns had an increasing percentage of ß-globin doublet versus singlet alleles as cells proceeded through S phase, whereas nuclei with late-replicating patterns harbored a high percentage of doublet alleles from the onset of their appearance. Moreover, 73.3% of the ß-globin doublets observed during midS phase were within 1 µm of the periphery, whereas only 22.2% of DHFR doublets were found within this zone (not shown). These data clearly demonstrate that ß-globin doublets appear at the periphery during the middle replication pattern.
As discussed above, theoretically, the appearance of doublets is not necessarily coincident with replication. DHFR alleles were largely separated into distinct FISH signals within 30 min after the onset of S phase and so must have physically separated in less than 30 min. However, we can conclude that the ß-globin loci are replicated during the middle S phase replication pattern only if we assume that the ß-globin loci also separate shortly after replication. Moreover, with both the correlative method (Fig. 2 C) and the direct method (Fig. 2 D), there was an increase in the percentage of DHFR alleles displaying doublets as cells proceeded from early to middle S phase, coincident with the appearance of ß-globin doublets. The simplest explanation for this increase is that chromosomal domains adjacent to the DHFR locus may replicate in midS phase, resulting in further separation of daughter DHFR strands and enhanced doublet resolution. However, it was formally possible that there is a general increase in the separation of daughter strands in midS phase, and that ß-globin alleles are actually replicated earlier but take a longer time to separate than do DHFR alleles. To address this concern, we evaluated the colocalization of ß-globin and DHFR doublets with BrdU label. Colocalization of FISH and BrdU signals is complicated by the highly variable signal-to-noise ratios obtained with FISH and the variable intensity of BrdU signal at different sites within the nucleus. However, computer-assisted colocalization analysis using a confocal microscope allowed us to identify alleles showing significant colocalization of the FISH signal with BrdU signal (Fig. 3) . As the analysis of individual nuclei by this method was laborious, we concentrated on doublets within type III or IV nuclei. Within experimental error, yellow doublets represent alleles that have both replicated and separated within the BrdU pulse labeling time. As shown in Fig. 3 C, ß-globin doublets colocalized with BrdU at a much higher frequency than DHFR doublets. In many cases, the ß-globin doublets appeared to be very close to peripheral heterochromatin (Fig. 3 B). Very few DHFR doublets were found within the peripheral zone (defined here as <1 µm from the perimeter of DAPI staining), and none of these colocalized with BrdU. These data strongly suggest that daughter ß-globin loci are separated shortly after their replication, providing direct evidence for the replication of ß-globin alleles in midS phase, coincident with the replication of peripheral heterochromatin.
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The ability to identify specific loci by FISH provided us with the opportunity to determine whether peripheral localization of the ß-globin locus is established coincident with the establishment of its replication timing. At various time points after mitosis, the closest distance of each ß-globin and DHFR allele to the periphery of the nucleus was measured and normalized to the nuclear radius. To control for the possibility of invaginations in the nuclear envelope, which are not readily identified with DNA stains, the nuclear periphery was highlighted with an antibody against the nuclear lamina (Fig. 5 C), as has been done in similar studies (Dernburg et al., 1996; Parreira et al., 1997). Results (Fig. 5 D) revealed that, at 1 h after mitosis, the ß-globin locus was no closer to the periphery than the DHFR locus, and the distribution of these two genes within the nucleus was similar. However, at 2 and 3 h after mitosis, the median distance of the ß-globin locus to the periphery was significantly smaller than the DHFR locus. In fact, at 2 and 3 h after mitosis, the majority of nuclei had at least one ß-globin allele with a distance/radius ratio of 0.2, whereas very few nuclei had DHFR alleles that close to the periphery. We conclude that the ß-globin locus is positioned adjacent to the periphery of the CHO nucleus between 1 and 2 h after mitosis, coincident with the establishment of a replication timing program.
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Discussion |
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We were surprised to find that the CHO ß-globin locus associates with the periphery rather than centromeres. Several reports, including studies of the ß-globin locus, have found transcriptional silencing in both mouse and human cells to be correlated with localization close to centromeric heterochromatin (Brown et al., 1999; Francastel et al., 1999; Schubeler et al., 2000). In addition, we found the ß-globin locus adjacent to centromeres in mouse C127 fibroblasts (unpublished data). On the other hand, there is extensive evidence in S. cerevisiae that genes can be silenced by their localization to the periphery (Andrulis et al., 1998; Cockell and Gasser, 1999; Heun et al., 2001). Our results suggest that the nuclear periphery can also serve as a silencing compartment in mammalian cells. Since both mouse and human cells appear to have late-replicating hypoacetylated chromatin at the nuclear periphery (Croft et al., 1999; Taddei et al., 1999), it is likely that some silenced genes will be found localized to the periphery in these species as well.
The fact that the peripherally localized and transcriptionally silent ß-globin locus is replicated in midS phase, distinctly before the latest replicating sequences, raises two important questions. First, are silenced genes ever replicated as late as constitutive heterochromatin, or is the middle of S phase reserved for replication of facultative heterochromatin? Second, do silenced genes generally take on the replication timing of the compartment with which they associate, as the ß-globin genes appear to have done in CHO cells? Obviously, this report provides only a beginning, concentrating on a single locus that has been shown to switch its replication timing program during development. There is clearly a need for additional studies relating transcriptional silencing, association with heterochromatin, and replication timing in order to address these questions.
Our results provide additional evidence that nuclear position is established at a distinct point after nuclear formation (Csink and Henikoff, 1998; Dimitrova and Gilbert, 1999b; Bridger et al., 2000). However, they provide only a series of snapshots and cannot evaluate the mobility of these sequences during the cell cycle. In fact, the stability of these positions is still unresolved and may vary for different sequences and under different metabolic conditions. Photobleaching studies of fluorescently labeled chromatin in living cells have detected very little movement during the cell cycle (Abney et al., 1997; Marshall et al., 1997; Zink et al., 1998; Manders et al., 1999). However, other studies, also in living cells, have revealed rather substantial movements of certain centromeres (Shelby et al., 1996) and transfected sequences (Li et al., 1998; Tumbar and Belmont, 2001). Furthermore, Bridger et al. (2000) reported that the peripheral localization of the largely late-replicating human chromosome 18 was lost upon entry of cells into quiescence. When stimulated to reenter the cell cycle, chromosome 18 remained late-replicating in the first S phase but did not return to the periphery until the second cell cycle. This experiment indicates that subnuclear localization is not necessary to maintain the replication timing program that was established at the TDP. This conclusion was recently supported by direct visualization of origin sequences in living yeast cells (Heun et al., 2001). These studies suggest that late-replicating sequences in budding yeast associate with the periphery during early G1 phase and acquire a chromatin modification that delays their replication time and persists even though the sequences may wander from the periphery later in the cell cycle. The lack of a perfect association of the ß-globin locus with the periphery in the experiments reported herein is also consistent with some degree of dynamic movement of this locus within the nucleus. Hence, early G1 phase may represent a unique window of time during which association with heterochromatin can influence chromosome architecture and replication timing for the remainder of the cell cycle, regardless of the positional fate of those domains thereafter.
How might this occur? We have previously proposed a model in which proteins regulating chromosome architecture are removed from chromatin during mitosis and reassociate at the TDP (Dimitrova and Gilbert, 1999b; Gilbert, 2001). As sequences are repositioned after mitosis, the clustering of chromosomal segments that associate with similar proteins (e.g., constitutive heterochromatin and/or repetitive DNA segments) would seed a locally high concentration of these proteins, analagous to the concentration of Sir proteins at telomere clusters in S. cerevisiae (Cockell and Gasser, 1999; Gasser, 2001; Heun et al., 2001). Association with these nuclear compartments could influence the structure of facultative heterochromatin. Once established, this chromatin architecture may be stable for the remainder of the cell cycle, whether or not subnuclear position is maintained. Particular chromatin configurations could establish thresholds for the accessibility of replication origins to initiation factors, thereby influencing when during S phase replication origins can fire. The results presented here support this model by demonstrating that a specific developmentally regulated locus is positioned adjacent to peripheral heterochromatin early in G1 phase, coincident with a modification of this locus that delays the timing of its replication during S phase.
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Materials and methods |
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Probes and FISH
Cosmid cSc26, encompassing 40 kb of DNA downstream of the DHFR gene, was a gift from J. Hamlin (University of Virginia, Charlottesville, VA). To clone the Chinese hamster (Cricetulus griseus) ß-globin locus, we hybridized a Chinese hamster genomic lambda FIX II library (catalogue no. 946910; Stratagene) with a partial cDNA probe corresponding to exons 1 and 2, and part of exon 3 from the human ß-globin gene (gift of M. Aladjem, National Cancer Institute, Bethesda, MD) and isolated lambda phage
MCB. A 600-bp SacI cross-hybridizing fragment derived from this phage was cloned (pSac600) and sequenced (EMBL/GenBank/DDBJ under accession nos. AF314660 and AF314661), revealing 88% identity to the previously cloned (Lee et al., 1992) golden hamster ß-major globin chain cDNA. In addition, oligonucleotide primers corresponding to the golden hamster embryonic z gene (Li et al., 1992) were used to amplify a fragment from Chinese hamster genomic DNA that was then cloned (pHEB) and used to identify four overlapping phage (
JHC2,
JHC4,
JHC6, and
JHC8). A 400-bp PvuII/SacI PCR fragment from
JHC2 was then sequenced (EMBL/GenBank/DDBJ under accession nos. AF314658 and AF314659), revealing 96% homology to the golden hamster ßlike y-globin gene cDNA. Phage DNA was purified with a Wizard lambda DNA purification kit (Promega) for making FISH probes.
FISH analysis was performed as described (Bickmore and Carothers, 1995; Boggs and Chinault, 1997) with some modifications. Cultured cells were pulse labeled with BrdU 90 min, detached with trypsin, swollen in 0.075 M KCl at 37°C for 17 min, and fixed at 0°C for 10 min with fresh 3:1 methanol/acetic acid (fixative). Cells were centrifuged and resuspended in fixative at 0°C for 20 min, centrifuged, and resuspended again in fixative, and then dropped onto clean slides and air dried. Slides were stored at -20°C until use. For in vitro reactions, intact G1 phase nuclei were prepared by digitonin treatment and introduced into Xenopus egg extracts at 10,000 nuclei/µl extract, as described (Wu et al., 1997). Reactions were stopped by diluting nuclei 1:10 in cold nuclear isolation buffer (NIB; 10 mM NaCl, 3 mM MgCl2, 10 mM Tris-HCl, 0.5% NP40). 100 µl of nuclei were overlaid on 300 µl 1:2 glycerol/extraction buffer (3x EB; 100 mM KCl, 5 mM MgCl2, 20 mM Hepes, pH 7.5, 2 mM 2-ME), cytocentrifuged onto clean glass slides, and air dried. Fixative was dropped onto the slides, which were then air dried and stored as above.
Probes cSc26 and either JHC2 or
MCB were labeled with either digoxigenin-dUTP or biotin-dUTP using a commercial nick translation kit (Roche). For each slide, labeled DHFR and ß-globin probes (50100 ng each) were coprecipitated with 5 µg sheared herring sperm DNA (Sigma) and 1 µg sonicated hamster genomic DNA. Probes were denatured 5 min at 84°C and preannealed at 37°C for 15 min before hybridization. For hybridization, slides were treated with RNase A (100 µg/ml) in 2x SSC at 37°C for 1 h, rinsed in 2x SSC, dehydrated in 70, 90, and 100% ethanol for 5 min each, and air dried. Slides were then incubated in 70% formamide/2x SSC at 84°C (in our hands, 72°C was not sufficient) for 3 min, and then transferred quickly to ice-cold 70, 90, and 100% ethanol for 2 min each and air-dried. Denatured probes were applied to each slide under a coverslip, sealed with rubber cement, and incubated at 37°C overnight. Slides were then washed 4 times for 3 min each in 50% formamide/2x SSC at 45°C, 2x SSC at 45°C, and then 0.1x SSC at 60°C. Probe detection and BrdU detection (with an AMCA-labeled secondary antibody) were performed exactly as described (Bickmore and Carothers, 1995), and only BrdU-positive nuclei were scored. For the characterization of replication patterns after FISH (Fig. 2), hybridization was performed for only a single probe (detected with Texas red), and BrdU-labeled DNA was detected by FITC-conjugated antimouse IgG (Jackson Laboratory) at a final concentration of 14 µg/ml. Nuclei were observed with a Nikon Labophot-2 equipped with a 100x Planapo lens (NA = 1.4), and images were collected with a 35-mm camera and P1600 slide film. Slides were scanned with a Nikon 2000 slide scanner and composed in Adobe® PhotoshopTM software using only standard brightness and contrast adjustments.
To examine colocalization between FISH and BrdU, images were collected by confocal microscopy (MRC 1024ES; Bio-Rad Laboratories). The red and green images in the same focal plane were scanned sequentially to prevent bleed-through and were then merged. Background was subtracted, and colocalization analysis was performed with LaserSharp software (Bio-Rad Laboratories).
Labeling of nascent DNA in vivo and in Xenopus egg extracts
Xenopus egg extracts were prepared and handled as described (Wu et al., 1997; Dimitrova and Gilbert, 1998). The labeling of nascent DNA with 30 µg/ml BrdU in cultured cells, 50 µM biotin-11-dUTP in Xenopus egg extracts, and the detection of BrdU- and biotin-substituted DNA were performed as described (Dimitrova and Gilbert, 1999b).
Determination of gene position
Simultaneous detection of DHFR or ß-globin and the nuclear lamina was performed as described (Itoh and Shimizu, 1998), except that probes were denatured and preannealed (as described above) before adding to slides. One digoxigenin-labeled probe was used at a time. After secondary antibody incubation, the nuclear lamina was detected using mouse monoclonal anti-lamin A&C IgM (Covance) and Texas redconjugated goat antimouse IgM (Vector Laboratories). Images were collected by confocal microscopy (MRC 1024ES; Bio-Rad Laboratories) with 0.5 µm Z-series. The shortest distance from the brightest FISH pixel(s) to the center of the lamina signal, and the longest nuclear diameter within the medial plane were measured with LaserSharp-Processing software.
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Footnotes |
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Acknowledgments |
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This work was supported by National Institutes of Health grant GM57233-01 and National Science Foundation grant 0077507 to D.M. Gilbert.
Submitted: 9 April 2001
Revised: 13 June 2001
Accepted: 15 June 2001
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References |
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