Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Curr Biol. Author manuscript; available in PMC 2011 December 7.
Published in final edited form as:
PMCID: PMC3108027

Developmental Control of Late Replication and S phase Length



Fast, early embryonic cell cycles have correspondingly fast S phases. In early Drosophila embryos, forks starting from closely spaced origins replicate the whole genome in 3.4 min, ten times faster than in embryonic cycle 14 and a hundred times faster than in a wing disc. It is not known how S phase duration is regulated. We examined prolongation of embryonic S phases, its coupling to development and relationship to the appearance of heterochromatin.


Imaging of fluorescent nucleotide incorporation and GFP-PCNA gave exquisite time resolution of S phase events. In the early S phases, satellite sequences replicated rapidly despite a compact chromatin structure. In S phases 11–13, a delay in satellite replication emerged in sync with modest and progressive prolongation of S phase. In S phase 14, major and distinct delays ordered the replication of satellites into a sequence that occupied much of S phase. This onset of late replication required transcription. Satellites only accumulated abundant heterochromatin protein 1 (HP1) after replicating in S phase 14. By cycle 15, satellites clustered in a compact HP1 positive mass, but replication occurred at decondensed foci at the surface of this mass.


The slowing of S phase is an active process not a titration of maternal replication machinery. Most sequences continue to replicate rapidly in successive cycles, but increasing delays in the replication of satellite sequences extend S phase. Although called constitutively heterochromatic, satellites acquire the distinctive features of heterochromatin, compaction, late-replication, HP1 binding and aggregation at the chromocenter, in successive steps coordinated with developmental progress.

Keywords: Drosophila, embryo, cell cycle, S phase, MBT, heterochromatin, satellite sequences


In a typical eukaryotic cell cycle, different regions of the genome replicate at different times during S phase[1, 2]. We understand little of the mechanisms underlying this phenomenon, but the temporal hierarchy of replication is linked to the organization of DNA into chromatin [3]. In phylogenetically diverse organisms, DNA that is packaged into heterochromatin is late replicating. Recent advances have identified proteins and protein modifications that distinguish euchromatic and heterochromatic sequences, but we still know little of how these molecular features relate to the distinguishing behaviors of heterochromatin, its high compaction, transcriptional quiescence and late replication. Even though the behaviors of heterochromatin are unchanged between distantly related organisms, they change during development of an individual.

During development there is a trend of euchromatin converting to heterochromatin as cells become increasingly specialized. This formation of “facultative” heterochromatin is thought to reflect and mediate developmental control of gene activity, which becomes increasingly canalized as differentiation progresses. In one well-studied example of developmental change, one of the two mammalian X chromosomes in female embryos is inactivated near the time of gastrulation [2, 4]. The inactive X develops all the hallmarks of heterochromatin, compaction, transcriptional quiescence and late replication.

Other regions of the genome, particularly large blocks of simple repeat sequences called satellite sequences, are more consistently heterochromatic and late replicating. Though referred to as constitutively heterochromatic, the behavior of even these sequences comes under developmental control during the transformative events of early embryogenesis.

The embryos of many organisms exhibit global changes as they transit from early rapid cycles, which serve to increase cell number, to the initial events of morphogenesis at a stage called the Mid Blastula Transition, or MBT [59]. Changes in DNA replication accompany this embryonic progression [10, 8, 7]. The preblastoderm mitotic cycles in Drosophila embryos (cycles 2 through 9) are about 8.6 min long. In these preblastoderm cycles, nuclei are large, chromatin is substantially dispersed, and numerous origins initiate replication in near synchrony to duplicate the genome within a 3.4 min S phase [10]. The mitoses occur in synchrony without cytokinesis and the nuclei undergo precisely choreographed movements in the syncytial cytoplasm. The rapid cycles gradually begin to slow as the nuclei reach the cortex to form the blastoderm in late cycle 9. After slight changes during four gradually slowing cycles, cycle 14 changes abruptly in character and length (Figure 1A). S phase lasts 50 min, more than ten-fold longer than the early S phases. Coincident with this S phase prolongation, the embryo activates zygotic transcription, destroys many maternal mRNA’s, invests the previously syncytial nuclei in cellularization membranes, and initiates morphogenesis. The cell cycle, which previously was driven by maternally provided gene products, first comes to rely on zygotic gene expression. The loss of maternally provided mitotic activator Cdc25 prevents immediate progress to mitosis and so creates a G2 phase [11, 12]. Triggering mitosis from this G2 quiescence is spatially controlled by the time of transcription of Cdc25string [13, 11, 14]. The abrupt onset of these events in cycle 14 marks the MBT of the Drosophila embryo.

Figure 1
Contributions to S phase Length

From embryo to adult, two modifications of replication prolong S phase about 200 fold (Figure 1B) [10]; first, origin frequency is reduced thereby increasing the length of DNA replicated by each fork; and second, different regions of the genome start to replicate at different times within S phase spreading the task out over a longer time. While stalling or slowing of replication forks could also contribute to S phase duration, no systematic change in these parameters has been observed between early embryos and somatic cycles [10]. Although the relative contributions of replication fork spacing and late replication to changes in embryonic S phase duration are unknown, focal incorporation of BrdU late in S phase 14 implicated late replication in S phase prolongation [14].

The different satellite sequences of Drosophila constitute about 30% of the genome [15] ( We have focused our analyses on predominant classes of satellite sequence. The satellite sequences take on attributes of heterochromatin during development. Even prior to cycle 10, we find that satellite sequences are localized in foci of selectively compacted chromatin. At blastoderm formation, the sequences throughout the genome replicate toward the end of S phase implying synchronous completion of replication of all regions of the genome. The last replicating sequences in the latter blastoderm cycles are confined to restricted spots that include satellite sequences. The MBT and cycle 14 are associated with a dramatic prolongation of S phase and the emergence of sequential replication of a series of satellite sequences in a “late replication program”. Each satellite sequence has a distinct schedule of late replication within S phase. Onset of late replication in cycle 14 is prevented by introduction of the RNA polymerase inhibitor α-amanitin late in cycle 13, suggesting that the transition requires an active process and cannot be attributed to simple titration of maternal replication factors. Dramatic recruitment of HP1 to foci of satellite sequences does not occur until after replication in cycle 14. We conclude that emergence of late replication is responsible for the early embryonic slowing of S phase and that the different properties of heterochromatin are introduced at a series of distinct transitions during development.


Emerging features of heterochromatin in syncytial blastoderm nuclei

We developed new approaches to analyze DNA replication during the short syncytial cycles. To examine the chromosomal regions replicated in the closing moments of the short syncytial S phases (Schematic in Figure 2A), we injected fluorescently tagged deoxynucleotide triphosphate (Alexa546-dUTP) into blastoderm embryos and then fixed. Since the labeling period was shorter than mitosis, embryos that were at the end of S phase when label was injected progressed only part way through mitosis by the time of fixation. Indeed, all prometaphase, most metaphase, some anaphase, and no telophase nuclei were labeled. Thus, the label in anaphase nuclei represents the latest incorporation in the previous S phase, and the position of the label along the anaphase chromosomes reveals the location of these last replicating sequences. The last replicating sequences changed with successive blastoderm cell cycles (Figure 2A). In cycle 11, the label was widely distributed in a speckled pattern. In cycle 13, the label was localized to the leading (pericentromeric) regions of the separating anaphase chromosomes. Cycle 12 showed an intermediate pattern. We conclude that replication of the pericentromeric regions of the chromosomes finishes later than bulk replication in cycle 13 and that this pattern is established progressively during the blastoderm cycles.

Figure 2
DNA Incorporation in Syncytial Cycles and in Cycle 14

We examined S phase dynamics in real time using injected GFP-PCNA as a marker of replication forks [16]. Interphases 11, 12 and 13 begin with rapidly emerging fine-grained speckles of fluorescence throughout the nucleus that declined about five min later. In addition, foci of fluorescence were seen near the apex of the nuclei where the pericentric regions lie. In each successive cycle, these brighter foci intensified, appeared later, and persisted longer (Figure 3 and movies 13). We conclude that the extension of S phase in cycles 11–13 is substantially attributable to these late replicating regions.

Figure 3
Injection of PCNA-GFP Demonstrates the Changing Character of the Syncytial Divisions

Satellite sequences, which are largely pericentric, comprise the bulk of Drosophila constitutive heterochromatin. We visualized several satellites by in situ hybridization (Figure S1), and present data from two of them; an 11 Mb X-chromosomal repeat of 359 bases, and a 3.4 Mb Y-chromosomal repeat of the simple sequence AATAC (Figure S1). These two satellites have distinct schedules of replication representing a range of behaviors seen among all the satellites. In situ probes detect a coherent focus of signal for each locus (see Figure 5A and S1).

Figure 5
Satellite Sequences Shift to Late Replication According to Individual Schedules

To examine the timing of satellite replication in S phase 13, we labeled with Alexa-dUTP, and tested for coincidence of incorporated label and hybridization with 359 and AATAC probes. We categorized embryos as early, mid or late S phase based on nuclear size and pattern of incorporation and collected data from multiple nuclei. Embryos in early S phase showed diffuse nuclear incorporation with lacunae such that a few nuclei showed no signal overlying either 359 or AATAC foci, and many nuclei showed a weak signal overlying 359, but no signal overlying AATAC (Figure 5A). Embryos in late S phase showed localized incorporation that often included signal overlying AATAC but not over 359. Mid S phase embryos had nuclei with general fluorescence that were positive for both 359 and AATAC (Figure 5A). We conclude from this that replication of 359, AATAC and bulk DNA largely overlap in cycle 13, but that satellite sequence replication is slightly displaced to a later time, and that the amount of this displacement is greater for AATAC replication than it is for 359 replication. Notably, the replication of satellites was not simply slow; initiation of replication was delayed and this delay was satellite specific.

Satellite sequences are generally heterochromatic and their chromatin structure is thought to underlie the replication behavior. Since chromatin structure changes during the embryonic cycles, the process of heterochromatin formation might impact on the late replication process. For this reason, we examined two features of heterochromatin, compaction and binding of the heterochromatin protein 1, HP1, as markers for heterochromatinization of the satellites.

Compacted heterochromatic sequences are visible as bright regions in nuclei stained with DNA specific fluorescent dyes (cycles 9 and 10 in Figure S2 and Figure 4A, respectively). Uniformly condensed anaphase chromosomes had uniform staining, showing that selective dye binding is not responsible for focal satellite staining (data not shown). In situ hybridization with the 359 probe revealed that the biggest of the foci of compacted DNA corresponded to this large satellite (Figure 4A and S2). Careful staging showed focally concentrated DNA throughout interphase, and in all cycles examined (Figure S2 and data not shown). To assess compaction independently of dye binding, we measured the volume of the 359-in situ signal using 3D microscopy (Figure S3). The 359 sequences were compact with a volume that was virtually unchanged between mitosis and interphase. During mitosis (anaphase measurements), the volume of the 359 focus relative to total chromosomal volume approximated its relative DNA content (Figure S3), while its interphase volume was roughly an order of magnitude less than expected based on proportionality. Thus, while other features of heterochromatin are not yet apparent (see below), compacted chromatin is present during the preblastoderm cycles and the 359 satellite lies within compacted chromatin during these early cycles. Since replication occupies much of the early interphase, the satellite sequences are not fully decondensed throughout replication, but the data are compatible with transient and incomplete decompaction during replication (see discussion).

Figure 4
Chromatin Compaction and HP1 Accumulation in the Blastoderm Embryo

To probe heterochromatin formation further, we injected recombinant GFP-HP1 into embryos. Its localization parallels previous descriptions [17]. Notably, GFP-HP1 is nuclear during interphase with much of the fluorescence homogenously distributed during cycles 11–13, but foci that appear in mid cycle 11 intensify in each subsequent cycle (movie 4). Relative to later HP1 accumulation (see below) these early foci are faint and are small; they are notably smaller than the foci of late replication or domains of satellite sequences (compare Figure 4B and and33).

In summary, satellite sequences are selectively compacted throughout early embryonic cycles, and their replication, which is initially coincident with bulk DNA replication, is delayed in the later blastoderm cycles. The replication delay increases in cycles 11, 12 and 13, and different satellites show different extents of delay. However, even in the last syncytial blastoderm cycle the delays are small and there is extensive overlap in the replication of different sequences. Although we observe small foci of HP1 during syncytial blastoderm cycles, these foci do not correspond to the major satellites, which lie in larger foci and recruit HP1 only later (see below and discussion).

The MBT and prolongation of S phase in cycle 14

Proliferation slows in cycle 14 and S phase reaches about 50 min (Figure 1A). To examine the distribution of recently incorporated nucleotide, we injected Alexa-dUTP into embryos and fixed them within two min. Fixed embryos were staged and the distribution of fluorescence analyzed. Early injections gave widespread incorporation, while later injections gave increasingly restricted incorporation. The nuclei of pulse labeled embryos fixed at 5 min into S phase 14 showed relatively uniform incorporation. In contrast, the nuclei of embryos 15 min into S phase showed focal labeling (Figure 1C). Real-time imaging gave a more detailed view of this transition (movie 5); the accumulating GFP-PCNA suggested that widespread replication began by 2.5 min and a progressive shift to a grainy distribution beginning at about 5 min suggests that many of the early initiating replicons completed replication three to nine min later. We conclude that the bulk of the genome is replicated rapidly toward the beginning of cycle 14, and that replication of localized domains occupies the remainder of the long S phase.

To examine the regions of the genome replicating at different times during this S phase, we injected Alexa-dUTP at different times and examined label distribution along the chromosomes at the next anaphase (Figure 2B). Incorporation should mark all sequences replicated after injection. The distribution of label on the chromosomes became increasingly restricted with later injection times. Faint telomeric labeling was oserved mid S phase, but the predominant and latest labeling regions were pericentromeric, the position of the bulk of the heterochromatin in Drosophila [15] (Figure 2B).

To determine whether satellite sequences are among the late replicating sequences, we localized the satellites by FISH and defined the timing of nucleotide incorporation at these loci (Figure 5B and C). Early in S phase 14, pulses of Alexa-dUTP failed to label either 359 or AATAC foci. Incorporation in the region of 359 began about 2 min after S phase initiation and was completed by 18 min. Incorporation in the AATAC region occurred between about 18 min and 28 min. Other satellites replicated late with different schedules (data not shown). We conclude that several satellite sequences become dramatically late replicating in cycle 14, and that replication of each sequence block takes about 10–15 minutes.

We used GFP-PCNA to view S phase 14 in real time (Figure 6A and movie 5). PCNA is widely distributed only for the first 10 min of S phase 14. For the remainder of S phase, PCNA is localized to a group of bright foci that diminish first in number and then in intensity, finally fading into background at about 50 min. This suggests that bulk DNA replication is followed by a prolonged period of sequential replication of localized blocks of late replicating DNA.

Figure 6
Real-time Data for Replication and Heterochromatin Protein Binding

Real-time imaging of GFP-HP1 (movie 4) and RFP-HP1 (Figure 6A and movie 5) reveal progressive emergence of heterochromatin during cycle 14. Interphase 14 begins with faint foci of HP1 similar to and only slightly brighter than the foci in earlier cycles. While it is difficult to track individual foci with assurance, there is a clear trend in which the number and intensity of HP1 foci increase predominantly from 10 min to 25 min of S phase (e.g. Figure 6A frames 10:19 to 27:27). At high temporal resolution (movie 4), it appears that the early faint foci undergo a partial breakup (e.g. 42:50 frame) and re-establishment, perhaps with duplication (e.g. 52:12 frame) between about 5 min and 15 min of S phase. We conclude that a major accumulation of HP1 occurs during replication, but this analysis does not reveal whether any, some or all of the intense foci of HP1 evident later in cycle 14 represent de novo accumulation versus intensification of pre-existing foci.

To test for coupling of late replication and association of HP1, we simultaneously imaged both GFP-PCNA and RFP-HP1 (Figure 6A and movie 5). PCNA appears to first associate with the faint HP1 foci when they undergo the breakup described above (movie 5 and Figure 6A, 10:19 frame) suggesting that the breakup may correspond to replication of these foci. Later, when more obvious bright foci of PCNA accumulate (e.g. Figure 6A, frame 18:52), the PCNA foci do not overlap the HP1 foci. This surprising lack of overlap suggests that the late replicating sequences are not associated with abundant HP1 early in cycle 14.

To assess HP1 association with specific sequences, we probed for colocalization of HP1 and satellite sequences. At the beginning of S phase 14, no GFP-HP1 was detected at sites of 359 or AATAC hybridization (Figure S5 and data not shown). Indeed, HP1 did not appear to concentrate at the 359 site until after completion of its replication, becoming obvious 35 to 40 min into cycle 14 (Figure S5). We did not detect HP1 accumulation over AATAC in cycle 14 (not shown). We conclude that satellites can become late replicating prior to association with HP1.

In summary, the replication of satellite sequences are greatly delayed in cycle 14, and a program of sequential replication of distinct blocks of the genome extends S phase. Notably, different blocks of satellite sequences exhibit distinct replication profiles. Though recruitment of HP1 to chromatin is complex, its abundant association to late replicating regions follows rather than anticipates their replication.

Restructuring of heterochromatin in cycle 15

S phase 14 is transitional, beginning before many MBT events and introducing a prolonged interphase during which the MBT occurs. We examined satellite replication in S phase 15, the first post-MBT S phase. New experimental challenges, which resulted from cellularization and the switch from synchronous to patterned divisions, were circumvented by labeling with the cell permeable precursor, BrdU, staging embryos by mitotic domains [13, 11] and assessing progression of S phase by the gradual restriction of incorporation. Labeling of 359 becomes evident when the nuclei have numerous localized foci of BrdU incorporation, after the decline of widespread incorporation, and it is completed before the more restricted late pattern of replication (Figure 6D). AATAC sequences are replicated later than 359 (data not shown). Thus, in contrast to the other satellites we examined, which shift to late replicating in cycle 14, replication of the 359 satellite overlaps bulk DNA replication in cycle 14 and shifts in 15.

The postmitotic distribution of HP1 in cycle 15 exhibits new features. Rather than faint individual foci, a coherent mass of localized HP1 marks the leading edge of the late telophase chromosomes as the cycle 15-nucleus forms (Figure 6B and movie 6). We take this marked clustering of HP1 as an indicator of a step in the development of the chromocenter.

Cycle 15 still lacks a G1 phase, and dispersed bright speckles of PCNA marked immediate onset of replication except in the region of the chromocenter. At about 15 min into S phase, a PCNA halo appeared around the HP1 and intensified. The HP1domain then fragmented into HP1-bright foci on a background of somewhat less bright fluorescence, the HP1-dim regions (e.g. Figure 6B, 28:48 frame). PCNA strongly localized to the HP1-dim regions and was substantially excluded from the HP1-bright regions. Dynamic associations of HP1-bright foci with adjacent HP1-dim/PCNA-bright were detected (Figure 6C). As replication of a region completed and localized PCNA declined, the HP1-dim region shrunk and disappeared in concert. This behavior suggests that the compacted HP1 positive chromatin is transiently unfolded into a less compacted structure and that replication occurs in this less compacted region.

In summary, S phase 15 differs from 14 in at least two ways. First, replication of 359, which was largely coincident with bulk replication in cycle 14, is shifted to a distinctly later time in S phase 15. Second, heterochromatin is marked by HP1 from the beginning of interphase and it forms a more coherent chromocenter. The replication of this chromocenter is delayed, and its replication is accompanied by transient decompaction.

A switch in late replication at the MBT

An appealing model suggests that exponential amplification of nuclei titrates a maternal supply of replication machinery to prolong S phase. This is a passive model in that it does not involve new gene expression. We tested the effect of inhibition of embryonic transcription on subsequent cycle(s).

Injection of α–amanitin in cycle 13 caused a premature and synchronous mitosis 14 (movie 7). Mitosis 14 occurred so soon after mitosis 13 (21 min in movie 7) that S phase 14 (normally 50 min) must have been shortened. Since the premature mitosis 14 was successful (i.e. no bridging), S phase was completed in a reduced amount of time rather than being cut short. To achieve this effect, the α–amanitin had to be injected prior to cycle 14. Thus, RNA polymerase activity is required prior to cycle 14 to achieve the normal prolongation of S phase 14.

We also looked directly at the effect of α–amanitin on replication. We injected α–amanitin into one pole of cycle 13 embryos, incubated until aged to the time of late S phase 14, and then injected Alexa-dUTP. The α–amanitin caused an extra rapid cycle and higher nuclear density near the site of its injection, while distant nuclei were still in cycle 14 at the time of labeling (Figure 7). The nuclei that had advanced to cycle 15 showed widespread incorporation characteristic of earlier S phases. In contrast, the areas still in cycle 14 showed a restricted replication pattern typical of late S phase 14. Hybridization with 359 and AATAC probes reveal numerous nuclei with simultaneous labeling of both satellites, a feature seen in early syncytial S phases, in the accelerated part of the embryo, while only AATAC was labeled, characteristic of late replication, in the cycle 14 area (Figure 7 and S8). We conclude that onset of the late replication program is α–amanitin sensitive.

Figure 7
Transcription is required for the onset of late replication at the MBT

These findings show that α–amanitin blocks both the increased duration of S phase 14 and onset of the distinctive late replication program of the satellite sequences. We conclude that the egg carries sufficient replication machinery to support rapid S phases beyond the time of the normal MBT, and that if titration of maternal factors triggers onset of late replication, it does so by engaging an RNA polymerase dependent process.


Our work shows that heterochromatin, long recognized as a key factor in the developmental programming of gene expression, also plays an integral role in the timing of the early embryonic cell cycles. Satellite sequences successively acquire features of heterochromatin, becoming late replicating by cycle 14, which prolongs S phase. This prolongation of S phase slows the early cell cycles and allows the progression to MBT [16].

Our description of the successive introduction of the features of heterochromatic reveals a lack of interdependency of these features. For example, since it occurs earlier, the compaction of the satellite sequences is independent of late replication and of HP1 binding. We also have been able to visualize the events of late replication with unprecedented spatial and temporal resolution that gives insights into the replication of compacted HP1-bound chromatin.

Onset of late replication of satellites prolongs embryonic S phases

Origin spacing could contribute to S phase length (Figure 1). However, EM studies show that origin spacing changes only slightly from 7.9 kb to 10.6 kb from preblastoderm embryos to cycle 14 [10, 18]. Since forks are thought to converge at a rate of 3 kb/min, the additional separation would extend S phase by about 1 min [10], a minor contribution to the change from a 3.4 to a 50 min S phase.

S phase duration would also increase if all of the replicons did not replicate at the same time [1, 18, 10]. However, asynchrony in replicon firing can happen in two ways, organized and unorganized. By unorganized asynchrony, we mean that origins fire at different times without regard to their position in the genome. In this case early and late firing origins can be juxtaposed. When replication from an early firing origin reaches an adjacent later firing origin just before it fires, one, rather than two, forks replicates the inter-origin distance, doubling replication time. Greater unorganized asynchrony will result in passive replication of later origins and reduce the number of origins. Thus, given the known origin spacing, unorganized asynchrony is unlikely to make a very major contribution to the more than ten-fold increase in S phase length between pre-blastoderm cycles and cycle 14.

If replication asynchrony is organized so that large regions of the genome (replication units) have many similarly behaving replicons, early initiated forks invading a late region from the outside will not have time to replicate a significant portion of the large domain. The insulation resulting from distance can greatly magnify the impact of asynchrony on S phase duration. Organization of genomes into large replication units is widespread but poorly understood. We show that the satellite sequences are replication units, and that embryonic changes in S phase duration result from change in the schedules of their replication. In preblastoderm cycles, satellite sequences replicate early, finishing in synch with general replication (Figure 8A). Subsequently, satellite replication is increasingly delayed in parallel to S phase prolongation. Importantly, when satellite replication is late, it is deferred, not slow. For example, AATAC sequences begin to replicate 18 min into S phase 14 (Figure 6E). Each type of satellite sequence exhibits distinct replication delays. The 359 sequence has almost no delay in S phase 14, while AATAT and AATAACATAG (data not shown) finish replicating after 359 but before AATAC. The stereotyped schedules suggest that each replication unit has a characteristic “lateness” parameter. This lateness parameter appears to be continuously variable in that there are many replication units each with its own schedule of replication.

Figure 8
The Changing Character of Replication in Embryonic S phases

Though its replication is delayed, a unit such as AATAC replicated quickly once initiated (10 min). Although the 359 satellite is more slowly replicating (~15 min), we suggest that it may be composed of separately and asynchronously replicating subdomains that we sometimes resolve (e.g. Figure 5B 2–4). We conclude that the dynamics of replication within a replication unit changes only modestly during the early cycles (from 3.4 to roughly 10 min).

In summary, our results argue that by S phase 14, the genome is replicated as a series of units each of which replicates relatively quickly, but that a temporal program of sequential replication of these units creates a long S phase (concept embodied in the schema shown in Figure 8). This replication program resembles a consensus view of replication in slowly replicating cells of mammals and plants. We conclude that progression from coincident replication of all of the replication units in a rapid S phase to sequential replication in a prolonged S phase 14 underlies prolongation of early embryonic S phases in Drosophila.

Developmental regulation of heterochromatin and its role in late replication

In widely divergent species and biological settings, heterochromatin has common characteristics including, compaction, transcriptional quiescence, late replication, “repressive” histone modifications, and association of specific heterochromatin proteins. This intimate association suggests mechanistic coupling of these features. If this were so, the various heterochromatin characteristics would emerge coordinately at the same moment during development. Instead, our observations show temporal uncoupling during early Drosophila embryogenesis.

Although it was suggested that heterochromatin appears at cycle 14 [19], both the cytological and biochemical manifestations of heterochromatin develop progressively [2022] (Victoria Foe, p.c.). Foci of compacted chromatin that align with satellite sequences appeared in pre-blastoderm embryos prior to, and hence independently of, late replication and HP1 recruitment (Figures 4A, S2, S3). Furthermore, HP1 binding to satellite sequences occurred late in cycle 14, after the onset of late replication. Since HP1 would have to decorate the satellite sequences at the onset of cycle fourteen if it were required to suppress early replication and promote late replication, we conclude that the late replication of satellite sequences is specified independently of the HP1 binding. Finally, satellite sequences reorganize; the previously independent foci of satellites aggregate into a large coherent HP1 positive region at the very beginning of interphase 15. This intimate association of satellites, which makes the chromocenter more coherent, is downstream of cycle 14 events and the MBT.

Together our findings show that satellite sequences acquire the features of heterochromatin progressively. Compaction is present early, late replication is introduced subsequently, and recruitment of HP1 and then chromocenter maturation follow. Onset of position effect variegation suggests that heterochromatic suppression of transcription begins in G2 of cycle 14 and mounts subsequently [23]. Thus, heterochromatin does not form in a single step, and it acquires increasing influence during critical developmental events surrounding the MBT and gastrulation.

Replicating compacted sequences

If compaction of chromatin prevents replication, decompaction might accompany or provoke replication. Our real-time observations of PCNA and HP1 in cycle 15 show replication adjacent to, but not overlapping HP1-bright foci of compacted chromatin. A more diffuse HP1 region appears adjacent to bright HP1 foci; PCNA overlies these fainter partner foci. Each partner focus appears and disappears as the PCNA signal rises and declines. We conclude from this that replication does not occur in the compacted domain and that the sequences in the compacted HP1-bright focus unfurl during replication.

The persistence of in situ foci for 359 and AATAC shows that the satellites are not fully decondensed during replication. The size of partner HP1 foci also argues for limited decompaction. If an entire focus of compacted HP1-bright chromatin were to disperse, it would expand in volume, but the partner focus is about the same size as the brighter parent focus. Thus, we suggest that that a partner focus represents decompaction of a portion of the sequences harbored in the adjacent HP1-bright focus.

Following replication heterochromatic sequences rapidly recompact. After an initial expansion, the partner HP1 focus does not grow throughout replication, and it shrinks and disappears as replication declines. When pulsed with fluorescent nucleotides for less than the replication time of the satellite, fluorescence overlies the compacted satellite sequence. Thus, we suggest that DNA is “spooled” out of compacted foci, replicated and returned to compacted foci shortly after replication. We roughly estimate the duration of replication-associated decompaction in embryonic cycle 15 as one min. The dynamics, which are not easily consistent with decompaction of large topological domains, suggest that active replication forks drive local unfolding of chromatin structure, but we cannot exclude the possibility that transient decompaction might promote replication.

The developmental program

We are interested in mechanisms that couple the changing cell cycle behavior with development. Previous work suggested that the gradual prolongation of early cycles is secondary to gradual prolongation of S phase [16, 24]. A model in which the exponentially increasing amounts of DNA titrates replication components to prolong S phase is attractive, but not presently supported.

Our results show that if a titration mechanism governs S phase duration, it is indirect. Injection of α-amanitin in cycle 13 prevented onset of late replication, accelerated S phase 14, and caused an early synchronous mitosis. Thus, activity of at least one of the DNA dependent RNA polymerases is required to slow S phase, and the replication “hardware” needed for a rapid S phase are not limiting. Accordingly, if a titration mechanism were involved, the titrated component would regulate an upstream process. For example, transcription is restricted prior to cycle 14, and titration of a repressor might derepress transcription in late cycle 13, indirectly triggering onset of late replication.

Three findings suggest an abrupt switch to late replication at the beginning of cycle 14: the dramatic increase in S phase length, the accompanying switch of satellite sequences to delayed replication, and the requirement for transcription in cycle 13 for this transition. However, we see that the late replication program of cycle 14 is anticipated by slight delays in replication of satellite sequences in cycles 12 and 13. These early changes suggest a more progressive process. We propose that early slight changes in replication timing and transcription initiate a positive feedback process that precipitates an abrupt change at the MBT. Rapid cell cycles suppress transcription [7, 25] and limit the time available to modify newly replicated chromatin, but, once the cycle begins to slow, transcription and heterochromatin modifications would accelerate to create conditions permissive for late replication, which would further slow the cycle.

Experimental Procedures

Fly stocks, embryo manipulation and imaging

Drosophila melanogaster Sevelen (Sev) flies were used as wild-type. Embryos were collected on agar plates containing grape juice, aged appropriately, dechorionated for 2 min in 50% bleach, and washed in water. For injections, embryos were aligned on agar plates, transferred to coverslips, desiccated for approximately 8–10 min and overlaid with Halocarbon oil (Sigma). Flies expressing Histone H2AvD-GFP [26] or injected Sev were used for live analysis using a spinning disc confocal [16]Fixed image analysis used a DeltaVision (Applied Precision, Inc) RT microscope system (Olympus IX70 microscope).

Monitoring Replication

Cellularized (cycles 15 and 16) were labeled with BrdU as described [27]. Syncytial embryos were injected with the labeled nucleotide-triphosphate (50–100 μM Alexa546-dUTP), incubated for 2 to 15 minutes (no detectable incorporation occurred for 1.5 min), fixed (37% HCHO, 10 min) and hand devitellinized [28]

Probes and FISH

A 131 bp section of the 359 bp repeat was amplified (primers: 5′-CCCTCCTTACAAAAAATGCG and 5′-AAAATGGTCACATAGATG) from genomic DNA (digested with Sst I and Hinf I to constrain the rounds of PCR to single copies of the repeat) in the presence of DIG-dUTP (Roche cat# 11093088910). AATAC, AATAT, 1.686, and other short sequence repeats (30mers) were synthesized and end labeling reaction using terminal deoxynucleotide transferase (Roche cat# 0333356600) to incorporate biotin-, Cy5-, or Alexa488-labeled dCTP (from Roche and Invitrogen) [29]. Embryos (generally labeled as above) were prepared for FISH by denaturing DNA in 2.5N HCl/0.1% Triton X-100 for 5 min, rinsed 2 × for 5min in 0.1M Na BO2, then rinsed with PTx. Hybridization conditions: 5XSSC,40% HCONH2, 100 μg/ml E.coli tRNA, 50 μg/ml Heparin, 0.1% Triton X-100, 1–5 ng of probe/μl, at 30°C with rocking overnight. To combine HP1 detection and FISH, we first stained fixed embryos with anti-HP1 (monoclonal C1A9; Developmental Studies Hybridoma Bank) and fluorescent secondary, then washed (PTx) and post-fixed with 5% HCHO for 20 minutes, prior to the typical FISH protocol.

Preparation and Injection of Fluorescent Proteins

GFP fusion constructs of PCNA and HP1 have proven useful for tracking localization to chromatin in other systems where they appear to mark replication forks and heterochromatin respectively [3032]. To construct GFP and cherry (RFP) fusion proteins, GFP and cherry genes were PCR amplified from the Drosophila gateway collection and cloned into the NdeI and BamHI sites of Pet28a to create Pet28-GFP and Pet28-cherry. Oligos used for GFP and cherry amplification were 5′ – AAACATatggtgagcaagggcgagg - 3′ and 5′ - AAAGGATCCcttgtacagctcgtccatgcc – 3′. PCNA was PCR amplified using the oligos 5′ - aaagaattcATGTTCGAGGCACGCCTGGGTC - 3′ and 5′ - aaactcgagTTATGTCTCGTTGTCCTCGATC – 3′ and cloned into Pet28-GFP as an EcoRI and XhoI fragment. HP1 was PCR amplified using the oligos 5′ - aaaggatccATGGGCAAGAAAATCGACAAC – 3′ and 5′ – aaagcggccgcTTAATCTTCATTATCAGAGTA – 3′ and cloned into Pet28-GFP and Pet28-cherry as a BamHI and NotI fragment. 6XHis-GFP-PCNA, 6His-GFP-HP1, and 6His-RFP-HP1 were expressed in BL-21 DE3 pLysS bacteria (Stratagene) and purified on nickel agarose beads according to the manufactures instructions (Qiagen). Purified proteins were dialyzed into 40 mM HEPES, pH = 7.4, 150 mM KCl and concentrated using a Vivaspin Centrifugal device. GFP-PCNA, GFP-HP1 and RFP-HP1 were injected at a needle concentrations of 10 mg/ml.

Supplementary Material










Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.


1. LIMA-DE-FARIA A. Differential uptake of tritiated thymidine into hetero- and euchromatin in Melanoplus and Secale. J Biophys Biochem Cytol. 1959;6:457–466. [PMC free article] [PubMed]
2. LYON MF. Sex chromatin and gene action in the mammalian X-chromosome. Am J Hum Genet. 1962;14:135–148. [PubMed]
3. Brown SW. Heterochromatin. Science. 1966;151:417–425. [PubMed]
4. Lucchesi JC, Kelly WG, Panning B. Chromatin remodeling in dosage compensation. Annu Rev Genet. 2005;39:615–651. [PubMed]
5. Dalle Nogare DE, Pauerstein PT, Lane ME. G2 acquisition by transcription-independent mechanism at the zebrafish midblastula transition. Dev Biol. 2009;326:131–142. [PubMed]
6. Edgar BA, Kiehle CP, Schubiger G. Cell cycle control by the nucleo-cytoplasmic ratio in early Drosophila development. Cell. 1986;44:365–372. [PubMed]
7. Edgar BA, Schubiger G. Parameters controlling transcriptional activation during early Drosophila development. Cell. 1986;44:871–877. [PubMed]
8. Newport J, Kirschner M. A major developmental transition in early Xenopus embryos: I. characterization and timing of cellular changes at the midblastula stage. Cell. 1982;30:675–686. [PubMed]
9. Newport J, Kirschner M. A major developmental transition in early Xenopus embryos: II. Control of the onset of transcription. Cell. 1982;30:687–696. [PubMed]
10. Blumenthal AB, Kriegsein HJ, Hogness DS. The units of DNA replication in Drosophila melanogaster chromosomes. Cold Spring Harb Symp Quant Biol. 1974;28:205–223. [PubMed]
11. Edgar BA, O’Farrell PH. Genetic control of cell division patterns in the Drosophila embryo. Cell. 1989;57:177–187. [PMC free article] [PubMed]
12. O’Farrell PH, Edgar BA, Lakich D, Lehner CF. Directing cell division during development. Science. 1989;246:635–640. [PubMed]
13. Foe VE. Mitotic domains reveal early commitment of cells in Drosophila embryos. Development. 1989;107:1–22. [PubMed]
14. Edgar BA, O’Farrell PH. The three postblastoderm cell cycles of Drosophila embryogenesis are regulated in G2 by string. Cell. 1990;62:469–480. [PMC free article] [PubMed]
15. Lohe AR, Hilliker AJ, Roberts PA. Mapping simple repeated DNA sequences in heterochromatin of Drosophila melanogaster. Genetics. 1993;134:1149–1174. [PubMed]
16. McCleland ML, Shermoen AW, O’Farrell PH. DNA replication times the cell cycle and contributes to the mid-blastula transition in Drosophila embryos. J Cell Biol. 2009;187:7–14. [PMC free article] [PubMed]
17. Kellum R, Raff JW, Alberts BM. Heterochromatin protein 1 distribution during development and during the cell cycle in Drosophila embryos. J Cell Sci. 1995;108(Pt 4):1407–1418. [PubMed]
18. McKnight SL, Miller OL. Electron microscopic analysis of chromatin replication in the cellular blastoderm Drosophila melanogaster embryo. Cell. 1977;12:795–804. [PubMed]
19. Spofford J. Position-effect variegation in Drosophila. 1c 1976.
20. Vlassova IE, Graphodatsky AS, Belyaeva ES, Zhimulev IF. Constitutive heterochromatin in early embryogenesis of Drosophila melanogaster. Mol Gen Genet. 1991;229:316–318. [PubMed]
21. Bongiorni S, Prantera G. Imprinted facultative heterochromatization in mealybugs. Genetica. 2003;117:271–279. [PubMed]
22. Eissenberg JC, Reuter G. Cellular mechanism for targeting heterochromatin formation in Drosophila. Int Rev Cell Mol Biol. 2009;273:1–47. [PubMed]
23. Lu BY, Ma J, Eissenberg JC. Developmental regulation of heterochromatin-mediated gene silencing in Drosophila. Development. 1998;125:2223–2234. [PubMed]
24. Sibon OC, Stevenson VA, Theurkauf WE. DNA-replication checkpoint control at the Drosophila midblastula transition. Nature. 1997;388:93–97. [PubMed]
25. Shermoen AW, O’Farrell PH. Progression of the cell cycle through mitosis leads to abortion of nascent transcripts. Cell. 1991;67:303–310. [PMC free article] [PubMed]
26. Clarkson M, Saint R. A His2AvDGFP fusion gene complements a lethal His2AvD mutant allele and provides an in vivo marker for Drosophila chromosome behavior. DNA Cell Biol. 1999;18:457–462. [PubMed]
27. Shermoen AW. Drosophila Protocols. CSHL Press; 2000.
28. Minden J, Namba R, Mergliano J, Cambridge S. Photoactivated gene expression for cell fate mapping and cell manipulation. Sci STKE. 2000;2000:pl1. [PubMed]
29. Dernburg AF. Situ Hybridization to Somatic Chromosomes. In: Sullivan W, Ashburner M, Hawley RS, editors. Drosophila Protocols. 2000. pp. 25–55.
30. Yamaguchi K, Hidema S, Mizuno S. Chicken chromobox proteins: cDNA cloning of CHCB1, -2, -3 and their relation to W-heterochromatin. Exp Cell Res. 1998;242:303–314. [PubMed]
31. Leonhardt H, Rahn HP, Weinzierl P, Sprorbert A, Cremer T, Zink D, Cardoso MC. Dynamics of DNA replication factories in living cells. J Cell Biol. 2000;149:271–280. [PMC free article] [PubMed]
32. Meister P, Poidevin M, Francesconi S, Tratner I, Zarzov P, Baldacci G. Nuclear factories for signalling and repairing DNA double strand breaks in living fission yeast. Nucleic Acids Res. 2003;31:5064–5073. [PMC free article] [PubMed]