Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2012 Nov 6.
Published in final edited form as: Nature. 2012 Mar 14;483(7390):434–438. doi: 10.1038/nature10895

Intrinsic coupling of lagging-strand synthesis to chromatin assembly

Duncan J Smith 1, Iestyn Whitehouse 1
PMCID: PMC3490407  NIHMSID: NIHMS411192  PMID: 22419157

Abstract

Fifty per cent of the genome is discontinuously replicated on the lagging strand as Okazaki fragments. Eukaryotic Okazaki fragments remain poorly characterized and, because nucleosomes are rapidly deposited on nascent DNA, Okazaki fragment processing and nucleosome assembly potentially affect one another. Here we show that ligation-competent Okazaki fragments in Saccharomyces cerevisiae are sized according to the nucleosome repeat. Using deep sequencing, we demonstrate that ligation junctions preferentially occur near nucleosome midpoints rather than in internucleosomal linker regions. Disrupting chromatin assembly or lagging-strand polymerase processivity affects both the size and the distribution of Okazaki fragments, suggesting a role for nascent chromatin, assembled immediately after the passage of the replication fork, in the termination of Okazaki fragment synthesis. Our studies represent the first high-resolution analysis—to our knowledge— of eukaryotic Okazaki fragments in vivo, and reveal the interconnection between lagging-strand synthesis and chromatin assembly.

During eukaryotic chromosome replication, both genetic and epigenetic information must be accurately duplicated. For chromatin architecture and modifications to be truly epigenetic—that is, heritable despite not being genetically encoded—complete disruption and dissociation of nucleosomes from the replication fork must be prevented to allow the rapid re-deposition of precisely located and appropriately modified histones on the nascent DNA. Histone chaperone complexes govern both nucleosome disassembly and assembly at the replication fork1.

DNA replication is inherently asymmetric. Okazaki fragment synthesis on the lagging strand necessitates the repeated production of single-stranded DNA and polymerization in the opposite direction to fork progression. Given the delay in lagging-strand synthesis and the rapidity of histone deposition behind the replication fork2, these two processes may be interlinked. A coordinated series of events occurs each time an Okazaki fragment is synthesized3. Each fragment is initiated via an RNA primer4 and up to 30 nucleotides of DNA, both synthesized by DNA polymerase α (Pol α)-primase5; subsequently, the clamp loading factor RFC mediates the replacement of Pol α by the sliding clamp PCNA and the processive polymerase Pol δ (ref. 6). Pol δ extends the nascent DNA chain through the 5′ end of the preceding Okazaki fragment7,8; this strand-displacement synthesis generates a 5′ RNA or DNA flap that is cleaved by nucleases such as the flap endonuclease Fen1 (ref. 9). Repeated cycles of extension and DNA cleavage followed by Pol δ idling10 produce a nick that migrates away from the replication fork and can be sealed by DNA ligase I11. Little to no DNA synthesized by the error-prone Pol α remains in the genome after replication12, suggesting that strand displacement generally replaces at least 30 nucleotides of DNA, but it remains unclear how the replication machinery coordinates the transition from Pol δ to DNA ligase. Indeed, despite their fundamental importance in replication, comparatively little is known about the in vivo properties of eukaryotic Okazaki fragments: although they are widely accepted to be shorter than the 1–2 kilobases (kb) observed in prokaryotes, a clear consensus has not emerged11,13–15. Moreover, we have little information about how nucleosome assembly may interact with lagging-strand synthesis. A plausible explanation for the difference in size between eukaryotic and prokaryotic Okazaki fragments is that eukaryotic DNA replication occurs within the context of chromatin.

To investigate the relationship between lagging-strand processing and chromatin assembly, we have developed strategies to analyse Okazaki fragments purified from S. cerevisiae. We find that Okazaki fragments are heterogeneously sized with a repeat length corresponding to that of nucleosomes. In addition, we find that ligation junctions occur preferentially around known nucleosome midpoints, and that these relationships can be altered by interfering with chromatin assembly or Pol δ processivity. Our data suggest an integrated model whereby nascent chromatin structure facilitates Okazaki fragment processing, which may provide a means to monitor nucleosome assembly behind the replication fork.

Nucleosome-sized Okazaki fragments

To enrich for ligation-competent Okazaki fragments, we constructed strains of S. cerevisiae in which DNA ligase I11 (CDC9) expression is driven from a doxycycline-repressible promoter and the encoded protein tagged with a temperature-sensitive degron16. The inability to ligate Okazaki fragments should result in the accumulation of nicked DNA that, after purification, can be radiolabelled using DNA polymerase and α-32P dCTP. Ligase inactivation in an asynchronous culture resulted in the accumulation of short heterogeneous DNA species whose abundance increased with duration of ligase inhibition (Fig. 1a). Notably, these fragments displayed an underlying periodicity reminiscent of the nucleosome repeat. Indeed, comparison of the nicked DNA to a nucleosome ladder indicated that a significant fraction of fragments seem to be sized according to the 165-bp nucleosomal repeat length in S. cerevisiae (Fig. 1b).

Figure 1. DNA ligase I depletion in S. cerevisiae leads to the accumulation of Okazaki fragments sized similarly to the nucleosome repeat.

Figure 1

Open in a new tab

a, Transcriptional repression of DNA ligase I (CDC9) results in the accumulation of nicked DNA. Cells carrying a doxycycline-repressible allele of the CDC9 gene were treated with doxycycline (Dox) for the indicated time. Purified genomic DNA was labelled using exonuclease-deficient Klenow fragment and α-32P dCTP and separated in a denaturing agarose gel. nt, nucleotides. b, The size of labelled Okazaki fragments mirrors the nucleosome repeat. Okazaki fragments (lane 2) were labelled as in Fig. 1a; nucleosomes were prepared from wild-type (lane 3) or repressible CDC9 strains (lanes 4 and 5) by in vivo Micrococcal Nuclease (MNase) digestion. The chromatin digestion patterns in lanes 4 and 5 indicate that CDC9 repression does not alter global chromatin structure. c, Okazaki fragments accumulate during S phase. Cells were arrested in G1 using α-factor, during which time CDC9 transcription was inhibited by the addition of doxycycline, and degradation of the protein stimulated by activation of the degron system using galactose and 37 °C. Okazaki fragments appear upon release of the culture into S phase (lanes 3 and 4). d, Okazaki fragments are bordered by ligatable nicks. Purified DNA was treated (lanes 2 and 4) or mock-treated (lanes 1 and 3) with the indicated ligase, and then labelled as in Fig. 1a. The inability to label fragments after ligase treatment confirms that labelled Okazaki fragments are flanked by ligatable nicks.

To clarify that we were observing Okazaki fragments, we confirmed that they did not appear in a G1-arrested culture until S-phase release (Fig. 1c). Additionally, we treated purified DNA with recombinant DNA ligase; this repaired the nicks and severely diminished our ability to label fragments (Fig. 1d), showing that our assay predominantly detects nicked DNA rather than species containing single-stranded gaps or flaps resulting from incomplete Okazaki fragment processing. Unrelated control experiments indicated that a 125-nucleotide species previously reported to be Okazaki fragments14,17 is 5S ribosomal RNA (Supplementary Fig. 2). Thus, we demonstrate that the mono-, di- and trinucleosome-sized fragments observed on ligase inhibition are generated in S phase and bordered by ligatable nicks—the essential properties of ligation-competent Okazaki fragments.

Global distribution of Okazaki fragments

To explore further the relationship between Okazaki fragments and nucleosomes, we developed a deep sequencing approach to map the strand, position and abundance of fragments across the S. cerevisiae genome. To allow yeast to complete S phase in the presence of nicked DNA we inactivated the DNA damage checkpoint by deleting the RAD9 gene. Additionally, to deplete ligase activity further we deleted the second DNA ligase (DNL4). DNL4 or RAD9 deletion does not affect Okazaki fragment size (Supplementary Fig. 3). Okazaki fragments were harvested from an asynchronous culture after a ~2.5 h ligase inactivation. Small single-stranded fragments were purified by anion exchange chromatography in alkaline conditions and sequencing primers were ligated directly to each end18 (Supplementary Fig. 4). Importantly, this method preserves strand identity.

After paired-end deep sequencing, we aligned the data to the S. cerevisiae genome and found that use of a single asynchronous culture was sufficient to attain complete coverage (a representative chromosome is shown in Fig. 2a, and a comparison of replicates in Supplementary Fig. 5). Sequencing reads aligning to the Watson or Crick strands showed a complementary distribution with strand bias being particularly pronounced close to replication origins19; such asymmetry is expected, given that replication forks proceed bidirectionally from origins (Fig. 2b). The strong strand bias observed around experimentally validated replication origins mapped at high resolution (Fig. 2c)20 demonstrates preferential sequencing of nascent lagging strands, unequivocally confirming that the DNA species enriched after ligase repression are Okazaki fragments. Detailed analysis of global Okazaki fragment distributions can provide mechanistic insight into replication-fork dynamics (S. McGuffee, D.J.S. and I.W., manuscript in preparation).

Figure 2. Sequenced Okazaki fragments show a pronounced bias towards the lagging strand.

Figure 2

Open in a new tab

a, Distribution of paired-end sequencing hits mapping to either the Watson (blue, above the axis) or Crick (orange, below) strands across S. cerevisiae chromosome 10. The locations of replication origins19 are indicated as grey dashed lines. Data are unsmoothed. b, The anticipated distribution of Okazaki fragments surrounding an efficient origin. c, Sequenced Okazaki fragments are strongly enriched in regions predicted to be replicated as the lagging strand. Log2 ratio of Watson strand: Crick strand fragments across a 50-kb window around ARS consensus sequences (ACS) confirmed previously20 to correspond to active origins. Data are smoothed to 200 bp.

Okazaki fragments terminate within nucleosomes

The periodic size of Okazaki fragments observed in Fig. 1 led us to investigate how the ends of Okazaki fragments relate to nucleosome positions found within the yeast genome. We aligned Okazaki fragment termini to a reference list of all consensus nucleosome midpoints (dyads) in S. cerevisiae21. Inherent bias towards smaller fragments during purification, library amplification and sequencing result in size distributions that differ substantially between starting material and sequenced fragments (Supplementary Fig. 6); therefore, we randomly selected fragments from our sequencing data to approximate the mononucleosome-sized fragments we detected by electrophoresis (Supplementary Fig. 6). Aligning both the 5′ and 3′ ends of mature (Fig. 3a) mononuclesome-sized fragments against the dyads of high-confidence nucleosomes indicated that the highest density of ends occurs at the dyad (Fig. 3b). Alignment of unfiltered data and dinucleosome-sized fragments is comparable to the mononucleosome size-selected subset (Supplementary Fig. 6). Therefore, rather than being overrepresented in the accessible internucleosomal linker DNA, we find that Okazaki fragment termination is most likely to occur at locations corresponding to nucleosome dyads.

Figure 3. Okazaki fragment termini are preferentially located at nucleosome dyads.

Figure 3

Open in a new tab

a, Schematic indicating the mature Okazaki fragment termini sequenced in this study. b, Okazaki fragment termini are enriched around nucleosome dyads. The distribution of termini from mononucleosome-sized Okazaki fragments around the top 50% consensus S. cerevisiae nucleosome dyad locations21 is shown. Unless otherwise indicated, all analyses use unsmoothed data normalized to the maximum signal in the analysed range, and area aligned such that Okazaki fragment (OF) synthesis by Pol δ proceeds from left to right. c, The distribution of Okazaki fragment termini correlates with nucleosome occupancy28 around TSSs. Data are aligned to TSSs such that the direction of transcription is from left to right: only Okazaki fragments synthesized in the same direction as transcription are analysed here; equivalent analyses for fragments synthesized in the opposite direction can be found in Supplementary Fig. 7. Data are smoothed to 5 bp. d, Okazaki fragments at nucleosome-depleted regions are disproportionately likely to be long. Equal numbers of reads were selected from the top (long), and the bottom three (short) length quartiles of sequenced fragments. The log2 ratio of long to short fragments is plotted. Data are smoothed to 5 bp.

DNA-bound proteins dissociate Pol δ

To confirm that the observed correspondence between nucleosome occupancy and Okazaki fragment end density was not biased by a small subset of nucleosomes, we analysed transcription start sites (TSSs), at which nucleosome positioning follows a well-defined pattern22. Alignment of Okazaki fragment termini around ~4,500 TSSs indicated a marked correlation between nucleosome occupancy and the density of Okazaki fragment ends (Fig. 3c and Supplementary Fig. 7) across a broad region surrounding TSSs. Precise nucleosome positioning around TSSs is determined by chromatin remodelling22,23, allowing us to infer that the correlation refers to remodelled, rather than intrinsically preferred, nucleosome locations.

If the ends of Okazaki fragments are dictated by nucleosomes then nicks should always be in register with positioned nucleosomes, regardless of internucleosomal spacing. Thus, nucleosomes separated by long linker DNA should be associated with longer Okazaki fragments. We took advantage of the differential nucleosome spacing surrounding the nucleosome-depleted region (NDR) at TSSs, comparing the distribution of equal numbers of randomly selected long and short fragments (defined, respectively, as the top quartile of sequenced fragments by length, and the bottom three quartiles). When the ratio of long to short fragments is plotted across this region (Fig. 3d and Supplementary Fig. 7), we find that fragments whose ends map within nucleosomes bordering the NDR are disproportionately long. Thus, Okazaki fragment length reflects nucleosome spacing.

As well as correlating with nucleosome occupancy, Okazaki fragment termini were enriched around known transcription factor binding sites (Fig. 3c)24,25. This suggested that, in addition to nucleosomes, sequence-specific DNA binding factors might directly influence the ends of Okazaki fragments. Such correlations would be expected if, following replication-fork passage, some transcription factors rapidly bound to replicated DNA and impeded Pol δ during strand-displacement synthesis, causing the polymerase to dissociate and leave a nick close to the site of collision. Furthermore, this model would also provide an explanation for the observed link between nucleosome occupancy and Okazaki fragment termination: nucleosomes contain numerous weak protein–DNA interactions, whose cumulative strength peaks around the dyad26. Polymerases can invade nucleosomes, but experience increasing resistance as they approach the dyad27,28. Therefore, if nucleosomes are rapidly re-established on the lagging strand behind the replication fork, and Pol δ invades them during strand-displacement synthesis, the likelihood of polymerase stalling and/or dissociation will increase with proximity to the dyad (see model in Supplementary Fig. 1), producing Okazaki fragments sized according to the nucleosome repeat. Other models that invoke biased Okazaki fragment initiation due to replisome pausing on encountering nucleosomes would produce distinct correlations around NDRs (Supplementary Fig. 8).

If polymerization by Pol δ is inhibited by sequence-specific DNA-bound transcription factors, Okazaki fragment ends should accumulate on the side of the factor that is first encountered by Pol δ. Aligning all transcription factor binding sites25 indicated a significant enrichment of precisely juxtaposed Okazaki fragment 5′ and 3′ termini around the replication-fork-proximal side of the binding site (Fig. 4a). However, when the Abf1, Reb1 and Rap1 binding sites were considered separately we found that, among transcription factors, these proteins most strongly biased the positioning of Okazaki fragment ends (Fig. 4b, c and Supplementary Fig. 9). Abf1, Reb1 and Rap1 have essential roles in chromatin organization24,29, and may therefore bind more rapidly and tightly to newly replicated DNA than the majority of transcription factors. We observed a strand-dependent enrichment of Okazaki fragment termini around the 3′ ends of tRNA genes, as well as transcription factor TFIIIB binding sites (Supplementary Fig. 10). Thus, it seems that diverse DNA binding proteins can influence the processing of Okazaki fragments and probably facilitate the dissociation of Pol δ, with tightly bound protein complexes stimulating termination more precisely than those, such as nascent histones, that have more diffuse binding regions.

Figure 4. Transcription factors with roles in nucleosome positioning stimulate dissociation of Pol δ.

Figure 4

Open in a new tab

a, Okazaki fragment termini are enriched on the replication-fork-proximal side of known transcription factor (TF) binding sites25. Mid, midpoint of TF binding site. b, c, The enrichment of Okazaki fragment termini around transcription factor binding sites can be attributed almost entirely to three factors—Abf1, Reb1 and Rap1—known to have roles in nucleosome positioning.

Chromatin assembly dictates nick location

Two testable predictions arise from the model that Pol δ dissociates via interaction with nascent nucleosomes: (1) disrupting nucleosome assembly will impair nucleosome-mediated Okazaki fragment termination and should, therefore, alter the size distribution of the fragments; and (2) reducing Pol δ processivity will result in a shift of Okazaki fragment termination sites away from the dyad, towards the replication-fork-proximal side of nucleosomes.

The deposition of (H3H4)2 tetramers on nascent DNA—the first stage in nucleosome assembly—is mediated in part by CAF-1, a multisubunit histone chaperone complex associated with the replisome via PCNA30. We deleted each subunit of CAF-1 individually, and compared the length of Okazaki fragments in the deletion strains to those with wild-type CAF-1 (Fig. 5a). Loss of any CAF-1 subunit abrogated the periodic sizing of Okazaki fragments, significantly increasing their average length. These data are consistent with a global delay in (H3H4)2 deposition following replication, with the resulting paucity of tetrasomes/nucleosomes on the lagging strand leading to increased strand displacement by Pol δ and thus to longer Okazaki fragments.

Figure 5. Impaired chromatin assembly and Pol δ processivity affect the size of Okazaki fragments and the location of their termini, respectively.

Figure 5

Open in a new tab

a, Deletion of any component of the CAF-1 complex (lanes 2–4) results in Okazaki fragments that are no longer sized according to nucleosomes. WT, wild type. b, Deletion of the pol32 subunit of Pol δ (lane 2) does not abrogate the chromatin-like size distribution of Okazaki fragments. c, In the absence of Pol32, the termini of mononucleosome-sized Okazaki fragments are shifted towards the replication-fork-proximal edge of the nucleosome: peak density occurs at a location consistent with the predicted edges of (H3H4)2 tetramers. See also Supplementary Fig. 11.

In S. cerevisiae, Pol δ is a heterotrimer whose Pol32 subunit increases processivity and PCNA binding affinity31; Pol δ lacking Pol32 shows decreased strand-displacement synthesis in vitro and probably in vivo32. Labelled Okazaki fragments from a pol32Δ strain were more heterogeneous but were still generally sized according to the nucleosome repeat (Fig. 5b). When the positions of the 5′ and 3′ termini of mononucleosome-sized fragments (165 ± 15 nucleotides, as per Fig. 3b) from the pol32Δ strain were aligned against nucleosome midpoints, a clear shift was observed towards the first edge of the nucleosome encountered by the polymerase (Fig. 5c and Supplementary Fig. 11). Therefore, when Pol δ processivity is perturbed, the polymerase dissociates more rapidly due to an inability to invade nascent nucleosomes assembled on Okazaki fragments. The new maximum end density occurs ~35–40 nucleotides from the dyad, consistent with the extent of DNA protected by (H3H4)2 tetramers33: nucleosome assembly occurs step-wise, with assembly of (H3H4)2 tetramers preceding recruitment of H2A/H2B34; it is therefore possible that the histone–DNA species encountered by Pol δ is a tetrasome rather than a nucleosome, although our data do not allow us to distinguish between these possibilities. Importantly, the distribution of Okazaki fragment termini around transcription factor binding sites was identical between wild-type and pol32Δ strains (Supplementary Fig. 9), allowing us to conclude that the shift towards the edge of nucleosomes does not simply represent a constant decrease of ~35–40 bp in the extent of strand displacement.

Discussion

Our data suggest a mechanism by which Pol δ may reliably remove DNA synthesized by Pol α while avoiding excessive strand-displacement synthesis. Repeated cycles of extension and cleavage can occur during the synthesis of a single Okazaki fragment. After removal of the RNA primer, all DNA flap structures generated by Pol δ are biochemically indistinguishable from one another, and an external mechanism is therefore required to measure the extent of strand displacement already carried out by Pol δ on each fragment. The removal of Pol δ by newly deposited histones represents a simple way to constrain Pol δ extension, and might allow nucleosome assembly to stimulate replication-fork progression directly. Although our data imply that nucleosomes strongly impede Pol δ, we note that both Fen1 and DNA ligase I can act efficiently on nucleosomal substrates35,36. We speculate that carrying out Okazaki fragment processing in the context of nucleosomes rather than DNA allows the ligation reaction to outcompete further strand displacement by Pol δ.

CAF-1 is not essential in S. cerevisiae, suggesting that other histone chaperones such as Asf1, Rtt106 and HIR37 allow sufficient replication-coupled or post-replicative nucleosome assembly for viability. Nevertheless, our results are consistent with a temporal delay in nucleosome assembly in CAF-1 mutants, which may give rise to previously reported silencing defects38. The importance of replication-coupled chromatin assembly in metazoa is illustrated by the severe phenotypes of CAF-1 or ASF1 disruption in human cell lines: ASF1 depletion leads to replication-fork stalling39, and CAF-1 depletion also precludes progression through S phase40.

Our studies demonstrate that the location of Okazaki fragment termini can be determined by interactions between the lagging-strand polymerase and nascent nucleosomes. This observation provides the first direct mechanistic evidence for the coupling of DNA replication to chromatin assembly on the newly replicated daughter genomes. Such coupling is fundamentally important given the part played by chromatin in the regulation of gene expression, as well as the potential for the epigenetic inheritance of precisely located modified nucleosomes. In addition, transcription factors known to have roles in the establishment of chromatin structure are apparently able to re-bind to DNA immediately after replication-fork passage. Unlike histones, which are present in sufficient number to be distributed to both daughter genomes, DNA-bound transcription factors are present at only one local copy per two daughters. Regulation of transcription factor re-binding could thus have a key role in asymmetric epigenetic inheritance.

METHODS

DNA purification

Yeast strains carrying degron-tagged, doxycycline-repressible alleles of CDC9 and a galactose-inducible UBR1 allele (see Supplementary Table 1 for a list of strains) were grown at 30 °C in YEP supplemented with 2% raffinose. At optical density (O.D.) 0.4, galactose and doxycycline were added to final concentrations of 2% and 40 mg l−1, respectively, and the culture shaken at 37 °C for 2.5 h. Fifty-millilitre cultures were used for labelling experiments, and 250-ml cultures for purification and library generation.

Genomic DNA was prepared from spheroblasts as described for medium resolution DSB mapping41. Following ligase repression, cells were collected by centrifugation, washed in SCE buffer (1 M sorbitol, 100 mM sodium citrate, 60 mM EDTA, pH 7.0) and spheroblasted for 3 min with 5 mg zymolyase 20T (USB) per 50-ml culture. Spheroblasts were washed with SCE, and resuspended in 480 μl lysis buffer (50 mM Tris-HCl, pH 8.0, 50 mM EDTA, 100 mM NaCl, 1.5% sarkosyl) containing 150 μg proteinase K (Fisher). Digestion was carried out for 2–16 h at 37 °C. After digestion, residual proteins and peptides were precipitated by adding 200 μl 5 M KOAc and spinning at 16,000g for 30 min at 4 °C. Nucleic acids were precipitated from the supernatant by addition of 500 μl isopropanol and centrifugation at 16,000g for 10 min. Pellets were washed twice with 500 μl 70% ethanol, resuspended in 200 μl STE buffer (10 mM Tris-HCl, pH 8.0, 1 mM EDTA, 100 mM NaCl) and digested with 25 μg RNase A (Sigma) and/or 10 U RIboshredder RNase blend (Epicentre) at 37 °C for 30 min. Genomic DNA was precipitated by addition of 20 μl NaOAc, pH 5.5 and 800 μl ethanol followed by centrifugation at 5,000g for 10 min at 25°C. Pellets were washed with 70% ethanol and resuspended in 1 μl TE (10mM Tris:Cl pH 7.5, 0.1mM EDTA) per ml original culture volume. DNA was stored at 4 °C and never frozen.

DNA labelling

Two microlitres of DNA (corresponding to the genomic DNA content of 2 ml cultured cells) was used in 20 μl labelling reactions containing 5 U Klenow (exo-)polymerase (NEB) and α-dCTP (Perkin Elmer) at a final concentration of 33 nM. Free label was removed using Illustra microspin G-50 columns (GE healthcare). Labelled DNA was separated in 1.3% denaturing agarose gels (50 mM NaOH, 1 mM EDTA). After electrophoresis, the gel was neutralized and DNA transferred to an uncharged nitrocellulose membrane (Hybond-N; GE healthcare) via capillary transfer. Membranes were exposed to phosphor screens or film.

Okazaki fragment purification

Genomic DNA purified as described earlier was denatured by heating to 95 °C for 5 min, rapidly cooled on ice and brought to 300 mM NaCl, pH 12. Purification was carried out in batch using 400 μl Source 15Q (GE healthcare), binding at 300 mM NaCl, pH 12 and eluting in 50 mM steps to 1,100 mM NaCl, pH 12. As determined by purification of fragments pre-labelled as above (see also Supplementary Fig. 4), fractions from 800–900 mM NaCl contained the majority of fragments of interest. DNA was ethanol precipitated and treated with 10 U Riboshredder RNase blend for 30 min at 37 °C to remove residual, undigested RNA: digestion products were removed using Illustra microspin G-50 columns to leave essentially pure Okazaki fragments.

Sequencing library generation

Adaptor primer pairs with single-stranded overhangs (shown schematically in Supplementary Fig. 4) were annealed by cooling from 95 °C and purified from 12% native polyacrylamide gels via standard methods. Sequences of the adaptor pairs are as follows. 5′ top, ACACTCTTTCCCTACACGACGCTCTTCCGATCT; 5′ bottom, NNNNNNAGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGT; 3′ top, /Phos/AGATCGGAAGAGCGGTTCAGCAGGAATGCCGAG; 3′ bottom, CTCGGCATTCCTGCTGAACCGCTCTTCCGATCTNNNNNN.

Up to 200 ng denatured purified Okazaki fragments were incubated at 16 °C overnight in a ligation reaction containing 1 μg of each primer pair and 1,000 U T4 DNA ligase (NEB). Unligated adaptors were removed using Illustra microspin S-300 columns (GE healthcare) and a second strand-synthesis reaction carried out at 72 °C using Taq polymerase (NEB). Products from ~200–1,000 bp were purified from 2.5% agarose gels run in TBE using Qiaquick kits (Qiagen). Purified libraries were amplified (16 cycles) using Illumina Truseq primers according to Illumina protocols, except that KOD hot start polymerase (Novagen) was used. Amplified libraries were purified from two sequential 2.5% agarose gels. Subsequent steps in the sequencing workflow were carried out according to standard procedures.

Nucleosome dyad and occupancy data

The top 50% of nucleosome dyads (by confidence score), taken from a meta-analysis21, were used in our analysis; nucleosome occupancy data (Fig. 3b) were from the YPD data set detailed previously42.

Supplementary Material

Supplementary Info

Acknowledgments

We thank S. McGuffee for assistance with data processing; S. Keeney, K. Marians, D. Remus, T. Tsukiyama, members of the Molecular Biology Program and Whitehouse laboratory for discussions and comments on the manuscript. This work was supported by a Louis V. Gerstner Jr Young Investigator Award and an Alfred Bressler Scholars Endowment Award to I.W. D.J.S. is an HHMI fellow of the Damon Runyon Cancer Research Foundation (DRG-#2046-10).

Footnotes

Supplementary Information is linked to the online version of the paper at www.nature.com/nature.

Author Contributions D.J.S. and I.W. designed experiments; D.J.S. performed experiments and analysed data; D.J.S. and I.W. interpreted results; the manuscript was drafted by D.J.S. and edited by D.J.S. and I.W.

Author Information Raw sequencing data and processed data are available at the Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE33786) under accession number 33786. Reprints and permissions information is available at www.nature.com/reprints. The authors declare no competing financial interests. Readers are welcome to comment on the online version of this article at www.nature.com/nature.

References

  • 1.Corpet A, Almouzni G. Making copies of chromatin: the challenge of nucleosomal organization and epigenetic information. Trends Cell Biol. 2009;19:29–41. doi: 10.1016/j.tcb.2008.10.002. [DOI] [PubMed] [Google Scholar]
  • 2.Sogo JM, Stahl H, Koller T, Knippers R. Structure of replicating simian virus 40 minichromosomes. The replication fork, core histone segregation and terminal structures. J Mol Biol. 1986;189:189–204. doi: 10.1016/0022-2836(86)90390-6. [DOI] [PubMed] [Google Scholar]
  • 3.Burgers PM. Polymerase dynamics at the eukaryotic DNA replication fork. J Biol Chem. 2009;284:4041–4045. doi: 10.1074/jbc.R800062200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Kaufmann G, Falk HH. An oligoribonucleotide polymerase from SV40-infected cells with properties of a primase. Nucleic Acids Res. 1982;10:2309–2321. doi: 10.1093/nar/10.7.2309. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Nethanel T, Kaufmann G. Two DNA polymerases may be required for synthesis of the lagging DNA strand of simian virus 40. J Virol. 1990;64:5912–5918. doi: 10.1128/jvi.64.12.5912-5918.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Waga S, Stillman B. Anatomy of a DNA replication fork revealed by reconstitution of SV40 DNA replication in vitro. Nature. 1994;369:207–212. doi: 10.1038/369207a0. [DOI] [PubMed] [Google Scholar]
  • 7.Ayyagari R, Gomes XV, Gordenin DA, Burgers PM. Okazaki fragment maturation in yeast. I. Distribution of functions between FEN1 AND DNA2. J Biol Chem. 2003;278:1618–1625. doi: 10.1074/jbc.M209801200. [DOI] [PubMed] [Google Scholar]
  • 8.Bae SH, Bae KH, Kim JA, Seo YS. RPA governs endonuclease switching during processing of Okazaki fragments in eukaryotes. Nature. 2001;412:456–461. doi: 10.1038/35086609. [DOI] [PubMed] [Google Scholar]
  • 9.Kao HI, Veeraraghavan J, Polaczek P, Campbell JL, Bambara RA. On the roles of Saccharomyces cerevisiae Dna2p and Flap endonuclease 1 in Okazaki fragment processing. J Biol Chem. 2004;279:15014–15024. doi: 10.1074/jbc.M313216200. [DOI] [PubMed] [Google Scholar]
  • 10.Garg P, Stith CM, Sabouri N, Johansson E, Burgers PM. Idling by DNA polymerase delta maintains a ligatable nick during lagging-strand DNA replication. Genes Dev. 2004;18:2764–2773. doi: 10.1101/gad.1252304. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Johnston LH, Nasmyth KA. Saccharomyces cerevisiae cell cycle mutant cdc9 is defective in DNA ligase. Nature. 1978;274:891–893. doi: 10.1038/274891a0. [DOI] [PubMed] [Google Scholar]
  • 12.Pavlov YI, et al. Evidence that errors made by DNA polymerase alpha are corrected by DNA polymerase delta. Curr Biol. 2006;16:202–207. doi: 10.1016/j.cub.2005.12.002. [DOI] [PubMed] [Google Scholar]
  • 13.Anderson S, DePamphilis ML. Metabolism of Okazaki fragments during simian virus 40 DNA replication. J Biol Chem. 1979;254:11495–11504. [PubMed] [Google Scholar]
  • 14.Bielinsky AK, Gerbi SA. Discrete start sites for DNA synthesis in the yeast ARS1 origin. Science. 1998;279:95–98. doi: 10.1126/science.279.5347.95. [DOI] [PubMed] [Google Scholar]
  • 15.Blumenthal AB, Clark EJ. Discrete sizes of replication intermediates in Drosophila cells. Cell. 1977;12:183–189. doi: 10.1016/0092-8674(77)90196-9. [DOI] [PubMed] [Google Scholar]
  • 16.Dohmen RJ, Varshavsky A. Heat-inducible degron and the making of conditional mutants. Methods Enzymol. 2005;399:799–822. doi: 10.1016/S0076-6879(05)99052-6. [DOI] [PubMed] [Google Scholar]
  • 17.Bielinsky AK, Gerbi SA. Chromosomal ARS1 has a single leading strand start site. Mol Cell. 1999;3:477–486. doi: 10.1016/s1097-2765(00)80475-x. [DOI] [PubMed] [Google Scholar]
  • 18.Ng P, et al. Gene identification signature (GIS) analysis for transcriptome characterization and genome annotation. Nat Methods. 2005;2:105–111. doi: 10.1038/nmeth733. [DOI] [PubMed] [Google Scholar]
  • 19.Nieduszynski CA, Hiraga S, Ak P, Benham CJ, Donaldson AD. OriDB: a DNA replication origin database. Nucleic Acids Res. 2007;35:D40–6. doi: 10.1093/nar/gkl758. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Eaton ML, Galani K, Kang S, Bell SP, MacAlpine DM. Conserved nucleosome positioning defines replication origins. Genes Dev. 2010;24:748–753. doi: 10.1101/gad.1913210. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Jiang C, Pugh BF. A compiled and systematic reference map of nucleosome positions across the Saccharomyces cerevisiae genome. Genome Biol. 2009;10:R109. doi: 10.1186/gb-2009-10-10-r109. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Whitehouse I, Rando OJ, Delrow J, Tsukiyama T. Chromatin remodelling at promoters suppresses antisense transcription. Nature. 2007;450:1031–1035. doi: 10.1038/nature06391. [DOI] [PubMed] [Google Scholar]
  • 23.Hartley PD, Madhani HD. Mechanisms that specify promoter nucleosome location and identity. Cell. 2009;137:445–458. doi: 10.1016/j.cell.2009.02.043. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Badis G, et al. A library of yeast transcription factor motifs reveals a widespread function for Rsc3 in targeting nucleosome exclusion at promoters. Mol Cell. 2008;32:878–887. doi: 10.1016/j.molcel.2008.11.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.MacIsaac KD, et al. An improved map of conserved regulatory sites for Saccharomyces cerevisiae. BMC Bioinformatics. 2006;7:113. doi: 10.1186/1471-2105-7-113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Hall MA, et al. High-resolution dynamic mapping of histone-DNA interactions in a nucleosome. Nat Struct Mol Biol. 2009;16:124–129. doi: 10.1038/nsmb.1526. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Bondarenko VA, et al. Nucleosomes can form a polar barrier to transcript elongation by RNA polymerase II. Mol Cell. 2006;24:469–479. doi: 10.1016/j.molcel.2006.09.009. [DOI] [PubMed] [Google Scholar]
  • 28.Churchman LS, Weissman JS. Nascent transcript sequencing visualizes transcription at nucleotide resolution. Nature. 2011;469:368–373. doi: 10.1038/nature09652. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Bai L, Ondracka A, Cross FR. Multiple sequence-specific factors generate the nucleosome-depleted region on CLN2 promoter. Mol Cell. 2011;42:465–476. doi: 10.1016/j.molcel.2011.03.028. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.Shibahara K, Stillman B. Replication-dependent marking of DNA by PCNA facilitates CAF-1-coupled inheritance of chromatin. Cell. 1999;96:575–585. doi: 10.1016/s0092-8674(00)80661-3. [DOI] [PubMed] [Google Scholar]
  • 31.Johansson E, Garg P, Burgers PM. The Pol32 subunit of DNA polymerase delta contains separable domains for processive replication and proliferating cell nuclear antigen (PCNA) binding. J Biol Chem. 2004;279:1907–1915. doi: 10.1074/jbc.M310362200. [DOI] [PubMed] [Google Scholar]
  • 32.Stith CM, Sterling J, Resnick MA, Gordenin DA, Burgers PM. Flexibility of eukaryotic Okazaki fragment maturation through regulated strand displacement synthesis. J Biol Chem. 2008;283:34129–34140. doi: 10.1074/jbc.M806668200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Dong F, van Holde KE. Nucleosome positioning is determined by the (H3-H4)2 tetramer. Proc Natl Acad Sci U S A. 1991;88:10596–10600. doi: 10.1073/pnas.88.23.10596. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Smith S, Stillman B. Stepwise assembly of chromatin during DNA replication in vitro. EMBO J. 1991;10:971–980. doi: 10.1002/j.1460-2075.1991.tb08031.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Chafin DR, Vitolo JM, Henricksen LA, Bambara RA, Hayes JJ. Human DNA ligase I efficiently seals nicks in nucleosomes. EMBO J. 2000;19:5492–5501. doi: 10.1093/emboj/19.20.5492. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36.Huggins CF, et al. Flap endonuclease 1 efficiently cleaves base excision repair and DNA replication intermediates assembled into nucleosomes. Mol Cell. 2002;10:1201–1211. doi: 10.1016/s1097-2765(02)00736-0. [DOI] [PubMed] [Google Scholar]
  • 37.Ray-Gallet D, et al. Dynamics of histone h3 deposition in vivo reveal a nucleosome gap-filling mechanism for h3.3 to maintain chromatin integrity. Mol Cell. 2011;44:928–941. doi: 10.1016/j.molcel.2011.12.006. [DOI] [PubMed] [Google Scholar]
  • 38.Zhang Z, Shibahara K, Stillman B. PCNA connects DNA replication to epigenetic inheritance in yeast. Nature. 2000;408:221–225. doi: 10.1038/35041601. [DOI] [PubMed] [Google Scholar]
  • 39.Groth A, et al. Regulation of replication fork progression through histone supply and demand. Science. 2007;318:1928–1931. doi: 10.1126/science.1148992. [DOI] [PubMed] [Google Scholar]
  • 40.Hoek M, Stillman B. Chromatin assembly factor 1 is essential and couples chromatin assembly to DNA replication in vivo. Proc Natl Acad Sci U S A. 2003;100:12183–12188. doi: 10.1073/pnas.1635158100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 41.Murakami H, Borde V, Nicolas A, Keeney S. Gel electrophoresis assays for analyzing DNA double-strand breaks in Saccharomyces cerevisiae at various spatial resolutions. Methods Mol Biol. 2009;557:117–142. doi: 10.1007/978-1-59745-527-5_9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 42.Kaplan N, et al. The DNA-encoded nucleosome organization of a eukaryotic genome. Nature. 2009;458:362–366. doi: 10.1038/nature07667. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Info
NIHMS411192-supplement-Supplementary_Info.pdf (11.4MB, pdf)

RESOURCES