FACT–(H3-H4) complex stimulates Pol α activity to coordinate DNA synthesis with nucleosome assembly

Wenshuo Zhang; Shichen Geng; Jiawei Xu; Yuantao Tang; Guanzhong Jiao; Jiayi Yang; Guojun Shi; Jiale Wu; Fangnan Qin; Ning Gao; Jianxun Feng; Qing Li

doi:10.1126/sciadv.aeh7889

. 2026 Sep 25;12(39):eaeh7889. doi: 10.1126/sciadv.aeh7889

FACT–(H3-H4) complex stimulates Pol α activity to coordinate DNA synthesis with nucleosome assembly

Wenshuo Zhang ^1,^†, Shichen Geng ^1,^†, Jiawei Xu ^1,^†, Yuantao Tang ¹, Guanzhong Jiao ¹, Jiayi Yang ¹, Guojun Shi ¹, Jiale Wu ¹, Fangnan Qin ¹, Ning Gao ², Jianxun Feng ^1,^*, Qing Li ^1,^*

PMCID: PMC13614383 PMID: 42789711

Abstract

DNA replication is an exquisitely regulated process that requires tight coordination at the eukaryotic replication fork, operating within the compacted chromatin environment. Minimizing exposure of naked DNA is critical to prevent nuclease degradation and genome instability, a hallmark of cancer. Cells, therefore, require a surveillance mechanism that links nucleosome assembly status to DNA replication. Here, we show that histone H3-H4 delivered by the chaperone facilitates chromatin transaction (FACT) directly stimulates the DNA polymerase activity of the polymerase α (Pol α, primase-polymerase complex) at replication forks. Histone H3-H4, but not histone H2A-H2B, promotes assembly of a FACT–(H3-H4)–Pol1 complex in S phase, where FACT associates with the Pol1 catalytic subunit in an H3-H4–dependent manner. This H3-H4–bridged interaction enhances Pol α primer extension on both DNA- and RNA-primed templates, increasing synthesis rate without altering primer patterning. Mechanistically, the Pol1 N-terminal domain, particularly its α1 and α2 helices, mediates complex assembly; the Pol1-Δα1α2 mutation disrupts this interaction and impairs primer extension in vitro. In cells, disrupting this complex reduces Okazaki fragment synthesis and slows replication fork progression. Among replication-coupled histone chaperones, only FACT interacts with Pol α, and this association is attenuated when chromatin assembly factor 1 (CAF-1)– and regulator of Ty1 transposition protein 106 (Rtt106)–dependent nucleosome assembly is compromised. Together, our results support a model whereby the FACT–(H3-H4)–Pol α complex serves as a sensor of H3-H4 availability, coordinating DNA synthesis with replication-coupled nucleosome assembly to safeguard genome stability.

INTRODUCTION

DNA replication is one of the most sophisticated cellular processes, requiring precise coordination at the replication fork (1, 2). The replisome, a multiprotein complex, unzips the parental DNA helix and duplicates the separated strands during DNA replication (3, 4). Physical interactions between replisome components ensure a coordinated framework for efficient DNA synthesis (5). In eukaryotes, the replisome operates within a chromatin environment, adding a profound layer of complexity (6–9). In DNA replication, newly synthesized DNA must be rapidly assembled into chromatin to maintain genomic integrity—a process known as replication-coupled (RC) nucleosome assembly (10–13). Limiting the time that DNA remains naked is crucial to avoid nuclease attacks, which can lead to genome instability—a hallmark of cancer (6, 8, 14). To prevent the exposure of naked DNA, a surveillance mechanism should be in place to monitor the synchronization between nucleosome assembly and DNA replication speed, thereby avoiding severe issues caused by uncoupled progression.

Histone chaperones play essential roles in regulating RC nucleosome assembly. They mediate histone dynamics—including deposition, transfer, and assembly—at the replication fork, thereby ensuring efficient nucleosome formation on nascent strands (12, 13, 15–17). Among them, the FACT (facilitates chromatin transaction) complex plays a role in both disassembly and reassembly of histones, functioning in both recycling parental histones ahead of the replication machinery and depositing newly synthesized histones onto replication fork (18–22), and thereby is essential for RC nucleosome assembly. CAF-1 (chromatin assembly factor 1) is specifically recruited to replication forks via interaction with proliferating cell nuclear antigen (PCNA) and replication protein A (RPA), where it primarily delivers newly synthesized H3-H4 histones to nascent DNA, promoting the de novo assembly of nucleosomes (23–26). In addition, the yeast histone chaperone Rtt106 (regulator of Ty1 transposition protein 106) functions cooperatively with CAF-1 and FACT, binding both new and parental H3-H4 histones, contributing to the maintenance of epigenetic information during DNA replication (21, 22, 24, 25, 27–29). These histone chaperones collectively form a coordinated network, ensuring efficient and accurate chromatin reassembly coupled with DNA replication.

The absence of key RC histone chaperones such as CAF-1 or FACT frequently results in a reduced replication rate (30–38). Accumulating evidence indicates that several replisome components can directly interact with histones and/or histone chaperones (8, 13, 17). While these physical interactions between replisome components and chromatin factors likely provide a mechanistic basis for coordinating multiple processes at the replication fork within chromatin, the precise molecular mechanisms underlying these functional connections remain poorly understood.

Notably, multiple replisome components interact with the histone chaperone FACT, which often assembles into a module consisting of replisome component(s), histones, and FACT (20, 39, 40). Among the replisome components that interact with the FACT-histones complex, several are enzymes, including the replicative helicase Mcm2-7 and the primase-polymerase complex DNA polymerase α (Pol α) (18–20). FACT is essential for helicase activity on nucleosomal DNA templates but not on naked DNA in cell-free systems (41, 42), indicating that it facilitates helicase function by promoting the disassembly of parental nucleosomes and recycling of parental histones. Downstream of unwinding, Pol α synthesizes the initial RNA-DNA chimeric primer essential for DNA replication (43, 44). Eukaryotic Pol α typically comprises four subunits: the largest catalytic polymerase subunit Pol1 (POLA1 in humans), the regulatory B subunit Pol12 (POLA2 in humans), and two primase subunits, Pri1 and Pri2 (PRIM1 and PRIM2 in humans) (43, 44). After the primase first synthesizes an 8 to 10 nucleotides (nt) RNA segment, the primer is then transferred to Pol1 for limited extension, producing a composite primer of ∼20 to 35 nt (45–50). Previous studies have shown that FACT interacts directly with Pol1 (51, 52) and that Pol1 also exhibits histone-binding activity toward both H3-H4 and histone H2A-H2B (53–56). However, whether and how the interactions of Pol1 with FACT and/or histones modulate Pol α activities remain to be elucidated.

Here, we demonstrate that H3-H4, but not H2A-H2B, specifically promotes the interaction between FACT and Pol1 during S phase. This ternary interaction stimulates the primer extension activity of Pol1. Furthermore, we mapped this interaction to the Pol1 N-terminal (Pol1-N) domain, which is required for binding to both H3-H4 and FACT. Disruption of these interactions impairs Okazaki fragment synthesis and slows replication fork progression. In addition, the FACT–(H3-H4)–Pol1 interaction is compromised under conditions of deficient RC nucleosome assembly. Collectively, our findings support a model in which the FACT–(H3-H4)–Pol α complex functions as a replisome-embedded sensor that monitors the availability of H3-H4 histones destined for nucleosome assembly into nascent DNA strands, thereby ensuring tight coordination between DNA replication and nucleosome assembly.

RESULTS

Histone H3-H4, but not H2A-H2B, promotes the interaction between FACT and Pol α

To test whether histones are required for the interaction between FACT and Pol α, we first characterized the interaction dynamics among FACT, histones, and Pol α based on fluorescent assays in living cells (Fig. 1A and fig. S1A). Because FACT directly interacts with Pol1, the largest subunit of Pol α (51, 52), we focused on Pol1 in this study. Using a bimolecular fluorescence complementation (BiFC) assay, the fluorescence signal arises from the complementation of two nonfluorescent mVenus fragments (e.g., Vn173 and Vc155) brought together by their respective fusion partners. In yeast, FACT consists of two essential subunits, Spt16 and Pob3. We tagged the Spt16 subunit with the N terminus of mVenus (Spt16-Vn173), while the Pol1 with the C terminus of mVenus (Pol1-Vc155) (fig. S1A). The BiFC signals confirmed that Spt16 interacts with Pol1, and both Spt16 and Pol1 interact with histone H3-H4 (fig. S1B). The control experiments coexpressing the individual Vn173 fragment with Pol1-Vc155, Vc155 with H3-Vn173, and Vn173 with Spt16-Vc155 in cells yielded no noticeable background signals, ruling out artifacts from nonspecific binding (fig. S1, C to E).

We further combined BiFC with the fluorescence resonance energy transfer (FRET) (Fig. 1A). In this system, Spt16 and Pol1 were fused with Vn173 and Vc155, respectively, while H3 was fused with mCerulean. Upon excitation at 440 nm, the interaction between H3-mCerulean and the reconstituted Spt16-Pol1-mVenus BiFC complex triggers FRET, resulting in mVenus emission (Fig. 1, A to C). The corrected FRET signal was calculated to reflect the BiFC-FRET efficiency. Expressing mCerulean or mVenus alone (Fig. 1B, 1 and 2), or coexpressing them (Fig. 1B, 3; negative control), did not trigger corrected FRET signals, whereas an mCerulean-mVenus fusion yielded a strong signal [Fig. 1, B (4; positive control) and C]. Notably, robust corrected FRET signals were detected in cells coexpressing Spt16-Pol1-mVenus and H3-mCerulean during S phase [Fig. 1, B (5) and C], demonstrating that Spt16, H3-H4, and Pol1 interact with each other in the S phase. These signals were notably enriched in S-phase cells compared to non-S–phase cells [Fig. 1, B (compare 5 and 6) and C], indicating a spatiotemporal specificity of the Pol α–(H3-H4)–FACT ternary complex formation.

FACT is the histone chaperone for both H3-H4 and H2A-H2B (40, 57), and Pol1 also binds both H3-H4 and H2A-H2B (54, 56). To determine the specific type and role of histones in the FACT-Pol α interaction, we tested the interactions in the spt16-m mutant background (spt16-K692AR693A in the Spt16 middle domain), which exhibits a substantial defect in binding H3-H4, but not H2A-H2B (21). Notably, BiFC signals of Spt16-Pol1, Spt16-H3, and Pol1-H3 pairs were notably reduced in spt16-m mutant cells (Fig. 1D and fig. S2, A to D). However, we did not detect an apparent reduction of BiFC signals in the Pol1-H2B pair in spt16-m mutant cells (Fig. 1D and fig. S2D). These results indicate that histone H3-H4 is crucial for the formation of both the FACT-Pol1 and Pol1-(H3-H4) complexes.

To further test this idea, we performed in vitro pull-down assays using purified yeast H3-H4, H2A-H2B, FACT complex, and Pol α complex (Fig. 1E and fig. S3, A to C). Green fluorescent protein (GFP)–tagged FACT complex (Spt16-GFP) was first incubated with either H3-H4 or H2A-H2B, followed by the addition of the Pol α complex. Western blot analysis of the GFP-FACT–associated proteins revealed that while both H2A-H2B and H3-H4 could be pulled down by FACT (Fig. 1E), Pol1–calmodulin binding peptide (CBP) was exclusively copurified when FACT was preincubated with H3-H4 (Fig. 1E; compare lanes 12 to 15 with lanes 6 to 11). With equal amounts of FACT, higher Pol1-CBP signals were observed as the amounts of H3-H4 increased (Fig. 1E; compare lanes 12 and 13 with lanes 14 and 15). As a control, we purified the FACT complex with the Spt16-m mutation for the pull-down assay (fig. S3A). Notably, the Spt16-m mutation reduced H3-H4 and Pol1-CBP signals in the FACT-associated proteins (fig. S4), confirming that H3-H4 binding of FACT is crucial for the formation of the Pol1–(H3-H4)–FACT complex.

To validate these interactions in vivo, we performed Pol1–tandem affinity purification (TAP) immunoprecipitation (IP) in G₁- and S-phase cells and observed that both FACT (Spt16 and Pob3) and histone H3-H4 signals associated with Pol1 were notably increased in S phase (Fig. 1F and fig. S5A). Moreover, the FACT and histone H3-H4 signals associated with Pol1 were notably decreased in the spt16-m mutant cells in S phase (Fig. 1F). Notably, both H3K4me3, a parental chromatin mark, and H3K56ac, a newly synthesized histone mark, were detected in the Pol1-associated complex. H3K56ac signals were notably decreased in spt16-m mutant cells, whereas H3K4me3 levels remained unchanged. Collectively, these results demonstrate that the H3-H4 binding ability of Spt16 is essential for the formation of the FACT–(H3-H4)–Pol α ternary complex, and the supply of newly synthesized histones may be more critical for regulating the assembly of this complex.

The FACT–(H3-H4)–Pol α ternary complex enhances the DNA polymerase activity of Pol α

To evaluate whether FACT–(H3-H4)–Pol α interaction regulates Pol α’s enzymatic activity and because the N-terminal domain (NTD) of Pol1 directly binds histones, we conducted a primer extension assay to examine its DNA polymerase activity (Fig. 2A). We designed several primer-template (P-T) substrates and confirmed their efficient annealing via native polyacrylamide gel electrophoresis (PAGE; Fig. 2B and fig. S6).

Fig. 2. — (A) Schematic of the in vitro Pol α primer extension assay using a DNA-primed template substrate. (B) Equal moles of 5′-Cy3–labeled dA15 primer and dT70 template were annealed to form the P-T junction (P_DNA-T₇₀) substrate. Primer (P), template (T), and annealed P-T junction were detected on a native gel via SYBR Gold staining and in the Cy3 channel. M, DNA marker. (C) Coomassie Brilliant Blue (CBB) staining of purified Pol α, WT or Spt16-m mutant FACT complexes, and recombinant yeast H3-H4. (D) Primer extension assays of Pol α with FACT–(H3-H4) complex or Spt16-m mutant FACT–(H3-H4) complex using the P_DNA-T₇₀ as the substrate. Products at the indicated time points were separated on a denaturing PAGE and imaged in the Cy3 channel. (E) Quantification of the grayscale intensity of full-length products in (D). (F) Primer extension assays of Pol α with FACT–(H3-H4) complex using P_DNA-T₁₂₅ as the substrate. Products at the indicated time points were separated on a denaturing PAGE and imaged in the Cy3 channel. (G) Quantification of the relative grayscale intensity of full-length products in (F). (H) Quantification of the relative grayscale intensity of intermediate products between 35 and 60 nt in (F). (I) Primer extension assays of Pol α with FACT–(H3-H4) complex or Spt16-m mutant FACT–(H3-H4) complex using P_DNA-T₁₂₅ as the substrate. Products at the indicated time points were separated on a denaturing PAGE and imaged in the Cy3 channel. (J) Quantification of the relative grayscale intensity of full-length products in (I). All quantification data are presented as mean ± SE from three independent biological replicates (n = 3). Differences between groups in (E), (G), and (J) were analyzed by two-way analysis of variance (ANOVA); *P < 0.05, ***P < 0.001, ****P < 0.0001.

First, we used a 70-nt poly-dT DNA as a template, annealed with a Cy3-labeled 15-nt poly-dA primer, to form a P-T junction (P_DNA-T₇₀) as a substrate for DNA polymerase (Fig. 2B), as previously described (46, 58, 59), and performed the primer extension assay as previously shown in the demonstration of the RPA’s role in regulating Pol α activity (58, 60). Pol α and FACT complexes were purified from yeast (Fig. 2C), and recombinant yeast histone H3-H4 was purified from Escherichia coli cells. After adding the Pol α complex to the reaction tube, DNA products were resolved on a denaturing PAGE gel at the indicated time points (Fig. 2, A and D). As expected, DNA synthesis products ranging from ∼15 (primer length) to 70 nt (full-length products) were generated, with a sharp band at ∼70 nt increasing in intensity over time. Meanwhile, a slight enrichment of the intermediate smear around 40 nt was observed compared to other intermediate ladders, consistent with the idea that DNA Pol α naturally has a limited DNA extension ability (45–47, 50). Notably, adding the preformed FACT–(H3-H4) complex increased the full-length product at 70 nt (Fig. 2, D and E), indicating that the FACT–(H3-H4) complex pronouncedly stimulates Pol1’s activity.

Given that the full-length product exceeds the reported primer extension length in cells, we hypothesized that the enhanced activity arises from FACT–(H3-H4)–Pol1 interactions that facilitate substrate binding by Pol1, thereby helping the enzyme overcome elongation barriers. To test this, we designed a longer template of 125 nt (P_DNA-T₁₂₅) to better reveal the intrinsic extension limitations of Pol1 and to determine whether FACT–(H3-H4) binding could promote elongation beyond these limitations and enable the generation of full-length products. Using this substrate, we observed distinct bands corresponding to the full-length 125 nt, an intermediate around 70 nt, and enriched intermediate smears above 30 nt, with clear periodicity in the peak distribution (Fig. 2F). These results are consistent with a defining property of Pol α, its limited processivity, which restricts the length of synthesized DNA products. Notably, the addition of FACT–(H3-H4) enhanced the formation of full-length products on the 125 nt as well (Fig. 2, F and G). We further quantified the conversion efficiency of intermediate products into full-length products. By tracking the time-dependent accumulation dynamics, we analyzed intermediate products between 35 and 60 nt and found that, in the presence of FACT–(H3-H4), these products reached peak levels much earlier than in its absence (20 min versus 40 min). The abundance of these intermediates decreased more rapidly in the presence of FACT–(H3-H4), accompanied by a rapid increase in longer intermediate products (Fig. 2H). Together, these results suggest that FACT–(H3-H4) alleviates the intrinsic elongation limitations of Pol1, likely by promoting more favorable substrate binding. To rigorously confirm that this enhancement specifically results from FACT–(H3-H4) binding to Pol1, we included the FACT complex with the Spt16-m mutation as a control. We observed that the stimulation effect is notably reduced compared to wild-type (WT) in both P_DNA-T₇₀ and P_DNA-T₁₂₅ substrates (Fig. 2, D, E, I, and J). In addition, while adding H3-H4 alone can slightly increase Pol1’s activity, they are not as efficient, as the FACT–(H3-H4) complex (fig. S7) results indicate that proper formation of the FACT–(H3-H4)–Pol1 complex is required for its stimulation of primer extension activity.

FACT–(H3-H4) stimulates Pol1 primer extension on both RNA- and DNA-primed templates by accelerating DNA synthesis

In cells, Pol α extends from an RNA primer synthesized by its primase subunit, and Pol1 preferentially acts on substrates with an RNA P-T junction (46, 61, 62). To validate our findings in a more biologically relevant context, we designed several substrates with a 5′-triphosphate–labeled RNA primer. A 60-nt DNA sequence was used as the template, with the first 5-nt poly-dT overhang and the following 10 nt containing a mixed sequence annealed to a 10-nt 5′-triphosphate–labeled RNA primer (P_RNA-T₆₀). To label the products, we included a dG in the template to ensure that the first nucleotide incorporated during primer extension is a Cy3-labeled dC (Fig. 3A). The products generated on the P_RNA-T₆₀ showed a distinct band around 60 nt and intermediate products above 35 nt (Fig. 3B). Similarly, we incubated the reaction at 30°C. We observed a distinct band around 35 nt, and with increasing time, additional intermediate products formed, culminating in a full-length product at 60 nt (fig. S8A). To systematically evaluate whether the intrinsic low processivity of Pol α persists on extended templates even when using a physiological RNA primer, we also tested a substrate containing an RNA primer and a 110-nt poly-dT DNA template (P_RNA-T₁₂₅). Consistent with the results obtained using DNA-primer substrates, we observed clear bands at the full-length 125 nt, an intermediate around 70 nt, and enriched intermediate smears above 35 nt, with some periodicity in the enriched peaks (fig. S9A). We observed that full-length products were generated at a much faster rate than with the DNA-primer substrate in both RNA-primer substrates, consistent with the idea that RNA primers are more favorable for Pol1 binding. Moreover, using an RNA primer with a random-sequence template (P_RNA-T_mix) yielded a clear full-length product without prominent intermediate stalling (fig. S8B), confirming that Pol α efficiency is highly dependent on sequence context and that the random sequence template has a higher activity (43, 58, 59).

Fig. 3. — (A) The procedure and scheme of the in vitro primer extension assay using RNA-primed template substrates. (B) Effect of adding WT or Spt16-m mutant FACT (together with H3-H4) on the primer extension activity of Pol α on RNA-primed template substrates. An RNA primer with a 5′-triphosphate group was annealed to a template to form the RNA P-T junction (P_RNA-T₆₀). Cy3-labeled 2′-deoxycytidine 5′-triphosphate (dCTP) was incorporated during primer extension and detected to assess Pol α activity. (C) Quantification of the relative grayscale intensity of full-length products in (B). Data are presented as mean values ± SE from three independent biological replicates, and the difference between two groups was analyzed using two-way ANOVA (**P < 0.01 and ****P < 0.0001). (D) Reaction progress curve of Pol α–catalyzed DNA-primer (P_DNA-T₁₂₅) extension. The abundance of full-length products was determined on the basis of relative grayscale intensity at each time point. Data represent mean ± SE from three independent replicates. Solid and dashed lines denote fits to a logistic growth model, which was used to calculate the half-time (t_1/2) and rate constant (K). (E) Reaction progress curve of Pol α–catalyzed RNA-primer (P_RNA-T₆₀) extension. The abundance of full-length products was determined on the basis of relative grayscale intensity at each time point. Data represent mean ± SE from three independent replicates. Solid and dashed lines denote fits to a logistic growth model, which was used to calculate the half-time (t_1/2) and rate constant (K).

Next, we applied the conditions described above to determine whether FACT–(H3-H4) promotes Pol α primer extension. Compared with Pol α alone, addition of FACT–(H3-H4) increased the amount of full-length products on all RNA-primed substrates (P_RNA-T₆₀, P_RNA-T₁₂₅, and P_RNA-T_mix) (Fig. 3, B and C; and figs. S9, A and B, and S10), indicating that FACT–(H3-H4) enhances Pol α primer extension from RNA primers. Similarly, as observed with the DNA-primer substrates, the Spt16-m FACT complex showed a reduced stimulatory effect (Fig. 3, B and C, and fig. S9, C and D). Adding both H2A-H2B and H3-H4 with the FACT complex generates aggregates in the lanes and also fails to show a stimulation effect. We reasoned that this may be due to difficulties in generating proper FACT binding with both (H3-H4) and H2A-H2B in vitro (fig. S10). Alternatively, the FACT–(H3-H4)–Pol1 interaction may not involve H2A-H2B at this moment. Together, these results suggest that FACT–(H3-H4) enhances Pol1 primer extension on both DNA and RNA primers.

Last, to quantify the kinetics of this stimulation, we asked whether FACT–(H3-H4) would alter the final reaction equilibrium or primarily accelerate the reaction rate. We therefore performed primer extension assays in the presence or absence of FACT–(H3-H4) over an extended time course until product formation reached a stationary phase and then measured the amount of full-length product (Fig. 3, D and E). We observed that the time-dependent accumulation of full-length products followed a sigmoidal profile. Fitting the data with a logistic growth model yielded the best-fit parameters. The parameter t_1/2 represents the time required to reach half-maximal product formation and reflects the lag phase preceding steady-state synthesis. The parameter K corresponds to the apparent elongation rate constant during the exponential phase and reflects the efficiency of productive DNA chain extension after reaction start. Using P_DNA-T₁₂₅ as a substrate, addition of the FACT–(H3-H4) complex did not change the final yield of full-length products but increased the synthesis rate, leading to an earlier arrival at equilibrium, with t_1/2 decreasing from 167.0 to 122.4 min (Fig. 3D and fig. S11, A and B). Similarly, using P_RNA-T₆₀ as a substrate (fig. S11, C and D), the addition of FACT–(H3-H4) did not alter the final yield of full-length products but increased the synthesis rate, shifting t_1/2 from 103.3 to 73.9 min (Fig. 3E). These results support the idea that FACT–(H3-H4) accelerates the DNA synthesis rate of Pol α.

Consistently, the apparent elongation rate constant K increased from 0.051 to 0.070 min⁻¹ when P_RNA-T₆₀ was used as the substrate, whereas only a modest change was observed with P_DNA-T₁₂₅ as the substrate (0.053 to 0.056 min⁻¹) (Fig. 3, D and E). Because the RNA-primer substrate requires a handoff of the primer from primase to the polymerase core, these results suggest that FACT–(H3-H4) may facilitate this switching process. Alternatively, binding of FACT–(H3-H4) to Pol1 may exert an allosteric effect that promotes primer extension.

Together, these results indicate that FACT–(H3-H4) stimulates Pol1’s primer extension activity on both RNA- and DNA-primed templates by accelerating the rate of DNA synthesis and that this stimulation depends on proper formation of the FACT–(H3-H4)–Pol1 complex.

The N terminus of Pol1 contributes to the FACT–(H3-H4)–Pol1 interaction

The N terminus of Pol1 (Pol1-N) contains a characterized histone-binding motif (54, 56). AlphaFold-Multimer v2 prediction confirmed that the Pol1-N domain interacts with histone H3-H4 (fig. S12). Ctf4 [chromosome transmission fidelity protein 4, AND-1 (Acidic Nucleoplasmic DNA-binding Protein 1) in vertebrates] functions as a trimeric scaffold that anchors Pol α–primase to the CMG helicase at the DNA replication fork to support efficient priming and coordinated Okazaki fragment synthesis on the lagging strand, and this interaction critically depends on the N-terminal CIP motif of Pol1 (63, 64). To define the structural basis of this FACT–(H3-H4)–Pol1 interaction, we predicted the interfaces among Pol1-N (1 to 330 amino acids), the H3-H4 tetramer, FACT [Spt16 (1 to 959 amino acids) and Pob3 (1 to 477 amino acids)], and the Ctf4-C domain (461 to 927 amino acids), as it was shown that FACT interacts with H3-H4 and Ctf4 interacts with Pol1-N via its C terminus (Fig. 4, A and B, and fig. S13). The predicted complex indicates that Pol1-N contains four α helices (α1 to α4; in red). Specifically, α1 interacts with the middle domain of Spt16 (Spt16-MD; yellow), α2 binds one H3-H4 dimer (H3-H4; blue and green), and α3 and α4 interact with Ctf4 (light purple) (Fig. 4B and fig. S13C). Notably, the linker upstream of α2 forms an antiparallel β sheet with the H4 N-terminal tail, while a random coil domain, C6, interacts with the Spt16-NTD (light orange) (Fig. 4B and fig. S13C). Similar multimeric assemblies were independently predicted by AlphaFold 3 (figs. S15 and S16).

Fig. 4. — (A) Structural domains of *Saccharomyces cerevisiae* full-length Pol1, FACT subunits (Spt16 and Pob3), and AlphaFold-Multimer v2–predicted structural domains of Pol1-N. CTD, C-terminal domain. (B) AlphaFold-predicted structure of the (Pol1-N)–(Spt16-NTD-DD-MD/Pob3-DD-MD)–(H3-H4)₂–(Ctf4-C) complex. Colors: Pol1-N, gray [α helices α1 to α4 and coil 6 (C6); red]; Spt16-NTD, light orange; Spt16-DD/MD, deep/light yellow; Pob3-DD/MD, light gray; H3/H4, blue/green; Ctf4-C, light purple. (C) Front (left) and back (right) views of the predicted structure of *S. cerevisiae* (Pol1-N)–(Spt16-MD)–(H3-H4)₂. Only α1 and α2 of Pol1-N (red), Spt16-MD (yellow), H3 (blue), and H4 (green) are shown. (D) In vitro pull-down assay of H3-H4 with recombinant GST-Pol1-N229. Bound proteins were separated by SDS-PAGE and visualized by CBB staining. Asterisk (*) indicates a nonspecific band. (E) In vitro pull-down assay of H3-H4 by WT and truncated forms of GST-Pol1-N229. Proteins were visualized by CBB staining. (F) Quantification of the relative intensity of bound histones in (E). Data are presented as mean ± SD from three independent experiments. Statistical differences were analyzed by one-way ANOVA (n.s. P > 0.05 and ***P < 0.001). (G) Pol1-TAP IP assay to assess the contribution of α1, α2, and C6 of the Pol1-N domain to the interaction between Pol1 and FACT–(H3-H4). Proteins in WCE and Pol1-CBP precipitates were detected by Western blotting with the indicated antibodies. A representative gel from two independent experiments is shown. (H) Table summarizing the degree of Pol1-FACT–(H3-H4) interaction shown in (G) across different mutants of Pol1-N domain. +++, strong interaction; ++, middle interaction; +, weak interaction. (I) Pol1-TAP IP results for WT and *pol1-*Δα1α2 mutant cells in G₁ and S phases, analyzed by Western blotting. A representative gel from two independent experiments is shown.

To further refine this architecture, we generated a structural model of the Pol1-N (1 to 330 amino acids), the H3-H4 tetramer, and the Spt16-MD (675 to 959 amino acids). This model also positions Pol1-α1 at Spt16-MD and Pol1-α2 at H3-H4 (Fig. 4C and figs. S14 and S16). Sequence alignments from budding yeast to human revealed strong conservation of these structural elements, particularly α1 to α5 and the C6, which are evolutionarily conserved (fig. S17). Consistently, AlphaFold Multimer v2 predictions of the (Pol1-N)–(H3-H4)₂–(Spt16-MD) complex across species showed highly conserved architectures (fig. S18), suggesting an evolutionarily conserved mechanism for FACT–(H3-H4)–Pol α interaction.

To experimentally validate these interfaces, we purified glutathione S-transferase (GST)–tagged budding yeast Pol1-N (1 to 229 amino acids) and structure-guided truncation mutants (Δα1, Δα2, Δα3, and Δα4) from E. coli (fig. S19A) and performed GST pull-down assays with recombinant yeast H3-H4. Compared with the GST control, H3-H4 was readily detected in pulldowns with WT GST-Pol1-N229 (Fig. 4D). Notably, deletion of α2 markedly reduced H3-H4 binding, whereas other truncations had little effect (Fig. 4, E and F). In contrast, GST-Pol1-N229 failed to pull down yFACT in the absence of histones (fig. S19B). These results indicate that the Pol1-N 229 region directly binds H3-H4, with α2 playing a key role, while this fragment alone is insufficient for FACT binding.

To assess complex assembly in vivo, we purified Pol α complexes from yeast cells expressing TAP-tagged WT or truncated Pol1. In WT Pol1-TAP complexes, both FACT subunits (Spt16 and Pob3), H3 and H4, newly synthesized histones marked by H3K56ac, and parental histones marked by H3K4me3 were detected. Consistent with structural predictions, signals for FACT and histones were notably reduced in the α1 deletion mutant (Δα1) and moderately reduced in the α2 mutant (Δα2), whereas deletion of α3 or α4 had little effect (fig. S20). Δα3 and Δα4 partially impaired the association of Pol1 with Mcm2 and Ctf4, in agreement with our structural prediction that Pol1-α3 and -α4 mediate binding to Ctf4 and its indirect interaction with Mcm4 (fig. S20). These results indicate that the interaction of Pol1 with Ctf4 may not directly contribute to the assembly of the Pol1–(H3-H4)–FACT complex. In addition, deletion of the C6 region (ΔC6) reduced FACT association without affecting histone binding. Notably, simultaneous deletion of α1 and α2 (Δα1α2) caused a marked reduction in both FACT and histone signals (Fig. 4, G and H). Together, these results confirm that the α1 and α2 helices are critical for the FACT–(H3-H4)–Pol1 complex formation.

Last, we examined these interactions across the cell cycle using Pol1-TAP IP. Fluorescence-activated cell sorting (FACS) analysis confirmed successful synchronization of WT and pol1-Δα1α2 cells in G₁ and S phase (fig. S5B). As expected, the association of FACT and H3-H4 with Pol1 was notably higher in the S phase than in G₁. FACT and H3-H4 signals were notably reduced in S-phase pol1-Δα1α2 cells compared with WT (Fig. 4I). Together, these results demonstrate that the α1 and α2 helices are essential for the FACT–(H3-H4)–Pol1 complex formation during S phase.

The FACT–(H3-H4)–Pol α interaction is required to maintain replication speed and Okazaki fragment synthesis

To further investigate the functional consequence of FACT–(H3-H4)–Pol1 interaction in DNA replication, we purified the Pol α complex with the Pol1-Δα1α2 mutation and conducted primer extension assays. Using the P_DNA-T₁₂₅ substrates, the mutant Pol α complex showed reduced primer extension activity compared with the WT proteins, indicating that the α1α2 region by itself has some role in promoting Pol1’s activity. Alternatively, the mutant under this condition may generate an inhibitory effect on Pol1-core’s activity. Moreover, the addition of FACT-H3-H4 produced little stimulation (Fig. 5, A and B, and fig. S21). These results demonstrate that the α1α2 is required for FACT-H3-H4 stimulation of Pol1’s DNA polymerase activity.

Fig. 5. — (A) Primer extension activity of WT and Pol1-Δα1α2 forms of Pol α using P_DNA-T₁₂₅ as the substrate for this assay. Products at indicated time points were separated by denaturing PAGE and imaged in the Cy3 channel. (B) Quantification of the primer extension products shown in (A). Data are represented as mean ± SE from three independent biological replicates (n = 3). Statistical significance between groups was determined by two-way ANOVA (****P < 0.0001). (C) The scheme of the DNA combing assay and representative DNA tracks. Top: Schematic of the DNA combing assay with BrdU labeling. Each BrdU tract indicates two divergent replication forks that start from the same origin. Bottom: Representative images of BrdU tracks (green) and full-length DNA (red) in WT and Pol1-N mutant cells. (D) Dot plots of BrdU track lengths in WT, and Pol1-N mutant cells. The mean and SD from a representative of two independent biological replicates are shown in the figures, and the mean values are indicated at the bottom of the panel. Differences were analyzed using a nonparametric t test (n.s. P > 0.05, *P < 0.05, and ****P < 0.0001). (E) The scheme of Okazaki fragment purification (left) and the result of Okazaki fragment synthesis in WT, and Pol1-N mutant cells by autoradiography (right). A representative gel from two independent experiments is shown. Left: In Cdc9-degraded cells, unligated Okazaki fragments are purified, end labeled with α-³²P-dCTP, separated on an alkaline gel, and detected by autoradiography. Right: The purified Okazaki fragments from WT and Pol1-N mutant cells; the marker for DNA size is labeled on the left of the panel. (F) Quantification of the grayscale intensity of each lane in (E).

Next, we examined cell cycle progression and DNA replication in cells with disrupted FACT–(H3-H4)–Pol1 interactions. Cells lacking the α1 and α2 motifs in Pol1-N (pol1-Δα1α2) showed a 5- to 10-min delay in S-phase progression (fig. S22) and compromised cell growth (fig. S23). To measure replication fork progression and origin usage, we performed DNA combing assays. Briefly, cells were arrested in G₁ and released into S phase, and nascent DNA was labeled with incorporated 5-bromo-2′-deoxyuridine (BrdU). Each BrdU track represents two sister forks emerging from a single origin, and the average BrdU track length reflects the average fork speed (Fig. 5C). Compared with WT cells (∼19.28 kb), the average BrdU track length was significantly reduced in Δα1 (∼16.60 kb), Δα2 (∼14.96 kb), and Δα1α2 (∼14.37 kb) mutants but not in ΔC6 (∼18.24 kb) (Fig. 5D). To rule out the reduced replication fork progression due to altered origin firing, we measured origin density by analyzing the average distance between origin centers. No noticeable differences were observed between WT and Pol1 mutants, indicating that these mutations do not affect replication origin selection (fig. S24). These results indicate that the α1α2-mediated interaction between Pol1 and H3-H4 contributes to maintaining replication fork speed.

We further performed chromatin immunoprecipitation (ChIP) assays to assess chromatin association of replisome components in WT and pol1-Δα1α2 cells. At early-firing origins ARS305 and ARS607, Mcm4, a subunit of the Mcm2-7 complex, was enriched in G₁ and decreased as replication progressed. Mcm4 loading at fired replication origins was not notably altered in pol1-Δα1α2 mutant cells (fig. S25, A and B), and Cdc45 binding also remained unperturbed (fig. S25, C and D), indicating that the mutation does not affect replisome assembly or integrity.

Because Pol α initiates Okazaki fragment synthesis during lagging-strand replication, we next examined Okazaki fragment production. Okazaki fragments were purified from Pol1-N mutant cells with DNA ligase I (Cdc9) degradation (65). Unligated fragments were end labeled with α-³²P–2′-deoxycytidine 5′-triphosphate (dCTP), separated on denaturing agarose gels, and detected by autoradiography (Fig. 5E). Okazaki fragment signals were markedly reduced in Δα1, Δα2, and Δα1Δα2 mutant cells and moderately reduced in ΔC6 cells compared with WT (Fig. 5, E and F). Together, these results demonstrate that the FACT–(H3-H4)–Pol1 interaction is critical for maintaining replication fork progression and efficient Okazaki fragment synthesis in vivo.

The FACT–(H3-H4)–Pol α interaction senses the RC nucleosome assembly level to adjust the replication speed

To further assess the role of FACT–(H3-H4)–Pol α interaction in the regulation of replication, we assessed the cell cycle progression in spt16-m mutant cells. We found that spt16-m mutant cells exhibited a delay in S-phase progression (fig. S26A). We then used the DNA combing assay to monitor DNA replication fork progression and origin selection under unperturbed conditions (fig. S26B). Our results showed a substantial reduction in the average length of BrdU tracks in spt16-m mutant cells (∼13.77 kb) compared to WT cells (∼19.31 kb) (fig. S26C), indicating a decrease in replication fork progression in spt16-m relative to WT. No noticeable difference in the average distance of origins between WT and spt16-m cells (∼40 kb) was detected (fig. S26D), suggesting that spt16-m mutation does not affect the overall selection of replication origin. Moreover, the spt16-m mutant cells showed a substantial reduction in purified Okazaki fragments (fig. S27, A and B). Thus, the ability of FACT to bind H3-H4 is critical for the formation of the FACT–(H3-H4)–Pol α complex and CMG complex, which is necessary for Okazaki fragment synthesis to maintain an appropriate DNA replication fork progression rate.

In addition to FACT, the histone H3-H4 chaperones CAF-1 and Rtt106 also function in RC nucleosome assembly (24, 25, 27). To test whether these chaperones interact with Pol1, we purified TAP-tagged Cac2 (a CAF-1 subunit), Rtt106, and Spt16 from yeast cells expressing Flag-tagged Pol1 (Fig. 6A). Although H3K56ac signals were detected in all three chaperone complexes, Pol1-Flag was detected only in the Spt16-TAP complex and not in the Cac2-TAP or Rtt106-TAP complexes (Fig. 6A). Consistently, BiFC signals were observed only for the Pol1-Spt16 pair but not for Pol1-Cac1 or Pol1-Rtt106 (Fig. 6B). To determine whether these histone chaperone mutations affect the FACT–(H3-H4)–Pol1 interaction, we purified Pol1-TAP complex from cells lacking CAF-1 and/or Rtt106. Both FACT and histone signals associated with Pol1 were notably reduced in cac1Δ and cac1Δ rtt106Δ mutant cells and moderately reduced in rtt106Δ cells (Fig. 6C). Consistently, BiFC analysis showed that the Pol1-H3 interaction was impaired in cac1Δ and rtt106Δ mutants, with the strongest reduction in the double mutant (Fig. 6D). Together, these results suggest that the FACT–(H3-H4)–Pol α interaction may act as a regulatory mechanism that senses the status of RC histone chaperone–mediated nucleosome assembly during replication, thereby facilitating the maintenance of efficient replication fork progression.

Fig. 6. — (A) IP assays using Cac2-TAP, Rtt106-TAP, and Spt16-TAP IP to purify Pol α. Proteins in WCE and those associated with each histone chaperone were detected using specific antibodies. A representative figure from two independent experiments is shown. (B) BiFC results of Cac1-Pol1, Rtt106-Pol1, and Spt16-Pol1 interactions in yeast cells. Pol1 was fused with Vc155, while Cac1, Rtt106, and Spt16 were fused with Vn173. All fusion proteins were expressed under their endogenous promoters. Cells harboring only Cac1-Vn173, Rtt106-Vn173, or Spt16-Vn173 served as negative controls. The percentage of cells with BiFC fluorescence signals is shown at the bottom-left corner of each image. Scale bars, 3 μm. (C) Pol1-TAP IP assays to purify histone H3-H4 and FACT complex in WT, *cac1*Δ, *rtt106*Δ, and *cac1*Δ *rtt106*Δ strains. The proteins in WCE and those associated with Pol1-CBP were detected using specific antibodies. A representative figure from three independent experiments is shown. (D) BiFC visualization and corresponding quantification of Pol1-H3 interactions in WT, cac1Δ, *rtt106*Δ, and *cac1*Δ *rtt106*Δ strains. Representative images show the percentage of cells with positive BiFC fluorescence at the bottom-left corner (scale bars, 3 μm). Cellular fluorescence intensity was quantified using ImageJ (n = 120 cells per strain). Data are represented as mean ± SD from a representative of two independent biological replicates. Statistical significance was determined using a nonparametric t test (*P < 0.05; ****P < 0.0001).

DISCUSSION

Our research has uncovered a previously unknown mechanism for regulating DNA replication through the FACT–(H3-H4)–Pol α interaction, in which nucleosome assembly provides feedback regulation of DNA replication, facilitating the mutual adaptation of the two processes and thereby maintaining genomic stability. We found that the FACT–(H3-H4) complex stimulates the DNA polymerase activity of Pol α, thereby maintaining replication fork progression under normal conditions. Furthermore, we demonstrated that histone H3-H4, but not H2A-H2B, is required for the interaction between FACT and Pol α during the S phase. Disruption of this interaction results in reduced Okazaki fragment synthesis and slower replication progression. This interaction is also regulated by CAF-1– and Rtt106-mediated RC nucleosome assembly pathways. Together, these results indicate that the FACT–(H3-H4)–Pol α interaction serves as a general mechanism for orchestrating DNA synthesis and RC nucleosome assembly at the replication fork (Fig. 7).

Fig. 7. — Model of the FACT–(H3-H4)–Pol α–mediated CARE (chromatin assembly–regulated early-warning) system. In WT cells, histone H3-H4 promotes the interaction between FACT and Pol α, and the formation of the FACT–(H3-H4)–Pol α complex is critical for stimulating Pol α activity, thereby maintaining Okazaki fragment synthesis and DNA replication speed. RC nucleosome assembly mediated by CAF-1 and Rtt106 can also regulate the FACT–(H3-H4)–Pol α interaction and DNA replication. When histone supply is deficient, due to the defective histone binding of FACT (*spt16-m*), Pol1-N domain mutations (*pol1-*Δα1α2), or impaired RC nucleosome assembly defective (*cac1*Δ and *rtt106*Δ), the primer synthesis activity of Pol α decreases. This reduction in Pol α activity leads to reduced Okazaki fragment synthesis and slowed replication speed, which prevents a marked mismatch between DNA synthesis and nucleosome assembly. Collectively, we propose a chromatin assembly–regulated early-warning system, termed CARE, which senses H3-H4 availability to coordinate DNA synthesis with nucleosome assembly.

Nucleosome assembly is a stepwise process, in general, wherein the initial step involves histone H3-H4 tetramer binding with DNA, followed by the subsequent addition of two H2A-H2B dimers. Moreover, the H2A-H2B dimers that constitute nucleosomes exhibit high dynamicity, readily dissociating from already assembled nucleosomes and exchanging with other H2A-H2B dimers. FACT consists of multiple domains and can interact with both H3-H4 and H2A-H2B. We and others have demonstrated that the Spt16-MD is required for the interaction between FACT and histone H3-H4 (20, 21, 66, 67). Here, we demonstrated that histone H3-H4, but not H2A-H2B, promotes the interaction between FACT and Pol α complex, raising the possibility that histone sensing of H3-H4 is more critical in regulating the coupling between nucleosome assembly and DNA synthesis. Considering that H3-H4 assembly is the first step in nucleosome assembly, it is reasonable to regard H3-H4 as a marker for histone supply and nucleosome assembly status. On the basis of this, the involvement of histone H3-H4—rather than H2A-H2B—in promoting the FACT-Pol α interaction further strengthens the intrinsic link between regulating Pol α enzymatic activity and synchronizing it with histone availability. When a sufficient level of histone H3-H4 is present, FACT binds to H3-H4, making it more readily accessible, facilitating the formation of the Pol1–(H3-H4)–FACT complex, and thereby stimulates the primer synthesis activity of the Pol α complex. This dynamic interaction thus serves as a regulatory mechanism that coordinates Pol α function with histone supply levels.

It is well established that before entering the S phase, genomic DNA organizes into a mature chromatin structure with a complete epigenetic landscape. Consequently, the amount of parental histones during RC nucleosome assembly is usually sufficient in the presence of parental histone recycling pathways. Consistent with this idea, while Pol1 also contributes to parental histone transfer via its Pol1-N domain (53, 54) and FACT was observed at the fork front side with a parental histone hexamer and the Mcm2 N-terminal histone-binding domain, the interaction between FACT–(H3K4me3-H4) and Pol α is not affected much, even in the spt16-m mutant background. However, in the absence of CAF-1 and Rtt106, or in a spt16-m background, new histone supply is insufficient, and the FACT–(H3K56ac-H4) levels are reduced. This may notably impair the FACT–(H3K56ac-H4)–Pol α interaction, leading to diminished primer synthesis activity by Pol α and a subsequent deceleration of DNA replication. Therefore, we propose that the FACT–(H3-H4)–Pol1 complex may serve as a general mechanism for sensing histone supply, especially newly synthesized histone levels regulated by RC nucleosome assembly pathways, to fine-tune DNA replication speed.

Pol α plays a crucial role in DNA replication, with its priming and polymerase activities highly regulated in cells (45). The NTD of Pol1, including a flexible region (∼1 to 330 amino acids), and its core-bridging extended N-terminal region (∼331 to 500 amino acids) are vital for the eukaryotic Pol α–primase complex (46, 63, 68). Cryo–electron microscopy studies reveal that Pol1-NTD is extended and dynamic, shifting alongside the catalytic core during primer synthesis (47). This flexibility allows Pol1-NTD to make transient contact with Pri1. When the RNA primer reaches 8 to 10 nt, it was proposed that Pol1-NTD transiently interacts with Pri1-NTD, leading to the release of Pri1 from template DNA, facilitating the switch from RNA to DNA synthesis (47). In addition, studies on replisome structures revealed the extrinsic regulatory significance of Pol1-NTD within the complete replisome. As a key interaction platform, Pol1-NTD participates in multisite binding with CMG helicase and Ctf4 adaptor during Pol α recruitment to replication forks (63, 69). We found that the α1α2 region (1 to 90 amino acids) of Pol1-NTD contributes to Pol1’s interactions with both histones and FACT, and this interaction mode is conserved from yeast to humans. Disruption of Pol1-α1α2 does not affect its interaction with Ctf4, or with Mcm2-7, key components of the replisome, but does reduce FACT–(H3-H4) binding. Notably, pol1-Δα1α2 results in a diminished Okazaki fragment synthesis. Thus, the N terminus of Pol1 is not merely a passive binding interface but plays a crucial role in maintaining or regulating the intrinsic catalytic activity of Pol α.

Our in vitro assay showed that α1α2 deletion notably reduced Pol α activity. One possibility is that this deletion may structurally generate an inhibitory effect of Pol1-N on the catalytic activity of Pol1 within the Pol α complex under these conditions. Alternatively, the Pol1-α1α2 region can somewhat influence the catalytic core of Pol1 to promote its activity, and the interaction of FACT–(H3-H4) further reinforces this activity, thereby stimulating the primer extension activity of Pol α. Thus, the reduced Okazaki fragment signals observed in pol1-Δα1α2 cells may arise from the combined effects of two intertwined factors: (i) the intrinsic loss of the N terminus’s contribution to basal Pol1-α1α2 activity (FACT independent) and (ii) the concomitant loss of the crucial stimulatory regulation provided by FACT–(H3-H4) via this exact region (FACT dependent). Given the essential role of Pol1-NTD in interactions with Pri1-NTD and the RNA-primer switching process, it would be interesting to test whether the α1α2 region plays a role in mediating the interaction between Pol1-NTD and Pri1-NTD and whether the FACT–(H3-H4) binding promotes the switching process from the primase core to the polymerase core, thereby reinforcing the stimulatory effect of Pol1-NTD. Collectively, the unique intrinsically disordered structure of Pol1-NTD may serve as a critical interaction hub that coordinates the spatiotemporal specificity of Pol α at replication forks, a feature fundamental to maintaining normal eukaryotic DNA replication and genome stability.

Pol α–primase executes canonical primer synthesis via two sequential catalytic steps: First, the Pri1 and Pri2 subunits of Pol α synthesize a short RNA oligomer (typically 7 to 10 nt); second, the Pol1 and Pol12 subunits extend this RNA to produce ∼30-nt RNA-DNA chimeric primers under physiological conditions (45–50). Despite a well-defined canonical length in vivo, in vitro assays show highly variable primer lengths, often diverging from the 30-nt standard. This heterogeneity mainly results from extrinsic factors such as P-T architecture, enzyme-substrate ratios, and incubation time, which notably influence Pol α product distribution. Proteins such as RPA and RFC modulate Pol α priming and extension, dynamically governing DNA synthesis, as shown in early biochemical studies using poly(A) and dT templates (58, 59). These factors can promote or restrict elongation based on their concentrations, affecting product length and abundance. Recent structural studies reveal that RPA remodels Pol α when encountering secondary structures, aiding bypass of hindrance and continuation of synthesis (70). Our primer extension assays confirmed Pol α’s ability to produce long DNA strands, much beyond the usual 30 nt. Even under conditions favoring long synthesis, a notable peak at 30 to 40 nt suggests an inherent processivity barrier limiting primer length to about 30 nt. In vivo, this processivity is further regulated by systems such as PCNA and Ctf4, which restrict Pol α activity and promote polymerase switches (63, 71). These insights explain the discrepancy between in vitro variability and the fixed primer size in vivo, highlighting both intrinsic properties of Pol α and cellular constraints that enforce the typical 30-nt primer length during replication.

We found that using both DNA- and RNA-primer substrates, FACT–(H3-H4) enhances the Pol α complex’s primer extension activity. The enhancement did not alter the periodicity of limited DNA extension, but it accelerated the rate, yielding intermediate, longer, and even full-length products. When conducting the primer extension assay over an extended period to reach the stationary phase of product formation, we found that the time-dependent accumulation of full-length products exhibited a sigmoidal profile. Notably, when the data were fitted using nonlinear regression to a logistic growth model, we observed that the addition of the FACT–(H3-H4) complex did not change the final yield of full-length products but did increase the synthesis rate, leading to an earlier arrival at equilibrium in both DNA-primer and RNA-primer substrates. The earlier arrival at equilibrium at the DNA-primer substrates raises the possibility that FACT–(H3-H4) may behave similar to a biocatalyst, notably advancing the synthesis rate. In addition, the apparent elongation rate parameter K changed when using RNA-primer substrates by adding FACT–(H3-H4) but remains unaltered when using DNA-primer substrates. These results indicate that the stimulation effect of FACT–(H3-H4) may have a different mechanism in DNA-primer or RNA-primer conditions. Because the RNA-primer substrate requires a handoff of the primer from primase to the polymerase core and Pol1 prefers RNA-primed substrate to DNA-primed substrate (46, 61, 62), these results indicate that FACT–(H3-H4) may facilitate this switching process. Alternatively, binding of FACT–(H3-H4) to Pol1 may exert an allosteric effect that promotes primer extension. Structural studies in the future would help provide insights into the mechanism.

Pol α plays a crucial role in DNA replication during the S phase. Throughout the DNA replication process, the continuous primer synthesis on the lagging strand drives the formation and maturation of Okazaki fragments. Here, we showed that disruption of FACT–(H3-H4)–Pol α interaction resulted in slowed S phase progression, reduced Okazaki fragment synthesis, and decreased DNA replication speed globally. Notably, disrupting the Pol1-N-(H3-H4) interaction in the pol1-Δα1α2 background did not affect CMG integrity much under normal conditions based on Cdc45 and Mcm4 ChIP analysis, indicating that the reduced DNA replication speed mainly stems from the effect on Pol α activity. A previous study has shown that histone H3-H4 promotes the formation of Asf1–(H3-H4)–Mcm2-7 complex ahead of the replication fork and provides a mechanism to regulate helicase unwind activity by histone supply (30, 72). Therefore, histone supply may control the DNA replication rate through multiple mechanisms.

Together, our findings demonstrate that DNA replication can be monitored to fine-tune DNA synthesis in response to nucleosome assembly status. The DNA replication system evaluates histone supply and nucleosome assembly levels by sensing the abundance of FACT–(H3-H4), which interacts with Pol1. This interaction further modulates the primer synthesis activity of Pol α, thereby adjusting DNA replication rates and ensuring coordination between DNA replication and nucleosome assembly. This coordination is essential for maintaining genomic stability, as it protects newly synthesized DNA by packaging it into nucleosomes, which then fold into higher-order chromatin structures based on the epigenetic information carried by histones. This precautionary mechanism is critical for preventing severe DNA damage resulting from exposed DNA; therefore, we refer to it as a chromatin assembly–regulated early-warning system (CARE system) (Fig. 7).

Clinical studies have shown that a mutation in the POLA1-α1α2 region [Ile⁸⁵→Ser (I85S)] is linked to an X-linked syndrome involving intellectual disability, severe growth retardation, microcephaly, and hypogonadism. The POLA1-I85S mutant notably impairs DNA replication capacity under conditions of replication stress (73). When replication stress or DNA damage occurs, the checkpoint pathway is activated to temporarily halt replication, allowing for repair and preventing the accumulation of errors. It was established that the S-phase checkpoint closely monitors Pol α activity and is also regulated by Pol α activity. With the CARE system to avoid uncoupled status, which may result in DNA damage and the S-phase checkpoint to repair the damage once activated, the two systems may jointly safeguard genome stability.

MATERIALS AND METHODS

Yeast strains and culture conditions

All Saccharomyces cerevisiae yeast strains used in this study were derived from W303-1 (leu2-3,112 trp1-1 can1-100 ura3-1 ade2-1 his3-11,15) (74). All genotypes, plasmid constructions, and oligos were listed in the tables S1 to S4. Yeast extract, peptone, and dextrose (YPD) and SC (synthetic complete) media were used to culture all yeast strains. Culture media and the culture conditions used in each assay were described in Materials and Methods details.

Bimolecular fluorescence complementation

Vn173 and Vc155 fragments were fused with the C terminus of genes of interest, which were linked with GGGGSGGGGS after translation, respectively (75). For Spt16-H3 BiFC, Vn173 was integrated into the C terminus of Spt16 with its endogenous promoter on a plasmid, and Vc155 was integrated into the C terminus of H3 with P_GAL1 promoter on a plasmid. For Spt16-Pol1 BiFC, Spt16-Vn173 was constructed as described above, and Vc155 was integrated into the C terminus of Pol1 with P_GAL1 promoter on the plasmid. For Pol1-H3 BiFC, Vn173 was integrated into the C terminus of H3 at its genomic loci, and Pol1-Vc155 was constructed as described above. The BiFC assays in Fig. 6B were performed by fusing Vc155 to Pol1 and Vn173 to the Cac1 subunit of CAF-1, Rtt106, and the Spt16 subunit of FACT. Pol1-Vc155 expression was under the intrinsic promoter of P_Pol1; the expression of Cac1-Vn173, Rtt106-Vn173, and Spt16-Vn173 was under P_CAC1, P_RTT106 and P_SPT16. The yeast strain carrying POL1-VC155 was crossed with yeast strains carrying fusion Vn173, respectively. For constitutive expression, yeast cells were cultured in SC medium with 5% glucose and then collected for imaging. For induced expression, yeast cells were cultured in SC medium with 2% raffinose and 0.02% glucose. Cells were then transferred into SC medium with 2% galactose, with 3 hours of induction at 25°C, followed by imaging. All the data are acquired on UltraView VoX Spinning Disk Confocal Microscopy (PerkinElmer) and Dragonfly (Andor).

Intensity-based FRET measurement with three filter microscopies

Yeast cells containing the BiFC-FRET system were collected and plated onto 0.17-mm-thick dishes with a glass slide at the bottom. Then, images were taken with an Olympus IX81 inverted microscope equipped with a 100×, numerical aperture = 1.45 oil immersion objective lens and a cooled charged-coupled device. Excitation light was delivered by an X-Cite light source. For image acquisition, the MetaMorph software (Molecular Devices) was used. In all experiments, the excitation intensity was attenuated to 25% of the maximal power of the light source. Images were acquired using 1 × 1 binning mode and 400-ms integration times. For quantitative FRET measurements, the method of sensitized FRET was previously described in detail (76–78). Images were acquired sequentially through yellow fluorescent protein (YFP), FRET, and cyan fluorescent protein (CFP) filter channels. Here, the filter sets were YFP channel (excitation filter: 504/12 nm; dichroic mirror: 400/521/607/700 nm; emission filter: 529/39 nm; Semrock), FRET channel (excitation filter: 427/10 nm; dichroic mirror: 400/521/607/700 nm; emission filter: 529/39 nm; Semrock), and CFP channel (excitation filter: 427/10 nm; dichroic mirror: 400/521/607/700 nm; emission filter: 472/30 nm; Semrock). The background images were subtracted from the raw images before carrying out the FRET calculation. Corrected FRET (F^C) was calculated on a pixel-by-pixel basis for the entire image using the following equation: F^C = FRET – (a × YFP) – (b × CFP), where YFP, FRET, and CFP correspond to background-subtracted images of cells coexpressing YFP and CFP acquired through the YFP, FRET, and CFP channels, respectively. The “a” and “b” are the fractions of bleed-through of YFP and CFP fluorescence through the FRET filter channel, respectively. To quantify FRET efficiency, we used FR = [FRET – (b × CFP)]/(a × YFP), a relative value that varies with changes in energy transfer to quantify the FRET signal. The FRET ratio (FR) represents the fractional increase in YFP emission due to FRET. Thus, in the absence of energy transfer, FR has a predicted value of 1 (79).

Tandem affinity purification

TAP-tagged purifications were performed as described (25, 80). Briefly, exponentially growing yeast cells were harvested by centrifugation and washed with 10% glycerol. Yeast cells were resuspended in equal volumes of buffer A [25 mM tris-HCl (pH 8.0), 100 mM NaCl, 1 mM EDTA, 10 mM MgCl₂, and 0.01% NP-40] containing protease inhibitors [1 mM dithiothreitol (DTT), 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM benzamidine, 1 mM Pefabloc, and deoxyribonuclease I (DNase I; 15 kU/ml)] and frozen in liquid nitrogen. Frozen cells were ground in a freezer mill (SPEX SamplePrep^LLC 6870). Cell lysates were clarified by centrifugation (20,450g for 40 min) and then incubated with immunoglobulin G (IgG) sepharose beads (GE Healthcare) for 2 hours at 4°C. IgG-bound proteins were digested with TEV protease for 2 hours at 16°C after extensive washing. The resulting proteins were further incubated with calmodulin beads (Agilent Technologies), eluted with SDS sample buffer, resolved on SDS-PAGE gel, and detected by Western blotting.

Protein purification for in vitro assays

yFACT complex purification for in vitro pulldown: GFP tag was integrated into the C terminus of the SPT16 gene at its endogenous genomic locus. The GFP purification was performed as previously described with minor modifications (19). Briefly, yeast cells were resuspended in equal volumes of buffer A with 200 mM NaCl and treated with DNase I (15 kU/ml). Cell lysates were incubated with GFP nanobody beads for 2 hours at 4°C. After binding, the beads were washed three times with buffer A with 500 mM NaCl to remove nonspecifically associated proteins. The beads coupled with the FACT complex were stored at 4°C.

Yeast DNA Pol α complex purification for in vitro pulldown: TAP tag was integrated into the C terminus of the POL1 gene at its genomic locus. The TAP purification was performed as described above with minor modifications. Briefly, yeast cells were resuspended in equal volumes of buffer A with 100 mM NaCl and treated with DNase I (15 kU/ml). Cell lysates were incubated with IgG beads for 2 hours at 4°C, and the beads were washed three times with buffer A with 300 mM NaCl. The IgG bead–bound proteins were digested with TEV protease, and the eluted DNA Pol α complex solution was stored at 4°C.

yH3-H4 purification: BL21 (DE3) strain carrying yH3-H4 expression vector was grown to reach optical density at 600 nm (OD₆₀₀) to 0.6 to 0.8. Isopropyl-β-d-thiogalactopyranoside (IPTG) was added to a final concentration of 0.4 mM, with induction at 37°C for 1 hour. Cells were washed once with 1× phosphate-buffered saline (PBS), resuspended in buffer C [10 mM Hepes·KOH (pH 7.6), 1 mM EDTA, 10% glycerol, 1 mM DTT, 0.1 mM PMSF, 2 mM benzamide, and 0.1 M NaCl], and sonicated three times, 30 s each, to break cells. The solution was spun down at 10,000 rpm for 10 min at 4°C. The pellets were resuspended in 0.25 M HCl using a homogenizer and incubated at 30°C for 20 min. The solution was then spun down at 10,000 rpm for 10 min. The supernatant was collected and neutralized with about 0.125 volume of 2 M tris base. The solution was dialyzed in buffer C overnight at 4°C and then loaded onto a SOURCE 15S column. The peak was collected at about 1 M NaCl concentration. The purified proteins were stored at −80°C.

yH2A-H2B purification: BL21 (DE3) strain carrying yH2A expression vector and JM109 (DE3) carrying yH2B expression vector were grown in LB medium at 37°C to reach OD₆₀₀ of about 0.6 to 0.8. IPTG was added to a final concentration of 0.4 mM. H2A was induced at 37°C for 1 hour, and H2B was induced at 37°C for 16 hours. Cells were washed with cold 1× PBS and spun down at 30,000 rpm for 15 min at 4°C. The supernatant was collected and neutralized with a 2 M tris base. The solution was dialyzed overnight in buffer C at 4°C and then loaded onto a SOURCE 15S column. The peak was at about 1 M NaCl concentration, and the eluted proteins were stored in −80°C.

Purification of Pol α from yeast cells for primer extension assay: The protocol was followed as previously described (69). Briefly, cells were grown to an OD₆₀₀ of about 2.0, and expression of Pol α was induced at 30°C for 2 hours by the addition of 2% galactose. Cells were collected, washed, and resuspended in buffer D [25 mM tris-HCl (pH 7.2), 10% glycerol, 0.02% NP-40-S, and 1 mM DTT] with 400 mM NaCl and protease inhibitors [0.3 mM PMSF, 7.5 mM benzamidine, 0.5 mM AEBSF, 1 mM leupeptin, 1 mM pepstatin A, and aprotinin (1 mg/ml)]. The frozen cells were ground in a freezer mill, and the resulting powders were then thawed in buffer D with 400 mM NaCl and protease inhibitors. The mixture was spun down at 235,000g for 1 hour at 4°C. The concentration of NaCl was reduced to 300 mM by dilution in a buffer lacking NaCl. The solution was then incubated with 2 mM calcium chloride and calmodulin affinity resin at 4°C for 90 min. Then, the beads were washed with buffer D with 300 mM NaCl and 2 mM CaCl₂, and proteins were eluted with buffer D with 300 mM NaCl, 2 mM EDTA, and 2 mM EGTA. Eluted fractions were diluted in buffer D to 120 mM NaCl and were loaded onto a 5-ml Mono Q column. Bound proteins were eluted and dialyzed in buffer D with 150 mM NaCl.

In vitro pull-down assay

For GFP pulldown, the FACT complex coupled GFP nanobody beads were washed three times with buffer A200 [25 mM tris-HCl (pH 7.5), 200 mM NaCl, 1 mM EDTA, and 0.01% Triton X-100] and incubated with the indicated proteins in buffer A200 for 2 hours at 4°C. After extensive washing, the beads were incubated with another indicated protein in buffer A200 overnight at 4°C. Protein-bound beads were eluted with SDS sample buffer, resolved by SDS-PAGE gels, and detected by Western blotting.

For GST pulldown, purified GST-tagged proteins were immobilized on Glutathione Sepharose beads in binding buffer A150 [25 mM tris-HCl (pH 7.5), 150 mM NaCl, 1 mM EDTA, and 0.01% Triton X-100] and protease inhibitors. Immobilized proteins were incubated with purified prey proteins (recombinant yeast H3-H4 tetramers or FACT complex) at 4°C for 2 hours with gentle rotation. Protein-bound beads were eluted with SDS sample buffer and resolved by SDS-PAGE gels. Band intensities were quantified using ImageJ (v1.54f), and binding efficiencies were normalized to input levels or the WT control.

In vitro primer extension assay of Pol α

Primer extension assays were performed on the basis of previously described protocols (46) with some modifications. For P-T junction substrates preparation, briefly, to prepare a 10 μM stock of the P-T substrates, 10 μl of 100 μM commercially synthesized 5′-Cy3–labeled DNA primers (synthesized by Tsingke Biotechnology) or 5′-triphosphate–labeled RNA primers (synthesized by Azenta Life Sciences) was combined with 10 μl of 100 μM corresponding DNA templates (synthesized by Tsingke Biotechnology) in 80 μl of TE buffer [10 mM tris-HCl (pH 8.0) and 1 mM EDTA] (primer sequences and DNA template sequences are shown in table S4). Annealing was achieved by heating the mixture to 95°C for 5 min, followed by a controlled cooling ramp to 4°C. The assembly quality of the P-T substrates was checked using 20% native PAGE (Fig. 2B and fig. S6).

For primer extension assay, all reaction mixtures were assembled on ice (4°C). Purified FACT and yH3-H4 proteins (100 nM each) were preincubated in the reaction buffer for 1 hour, followed by the addition of 50 nM Pol α. Assays using the P_DNA-T₇₀ substrate were performed in a low-salt buffer [50 mM tris-HCl (pH 8.0), 5 mM MgCl₂, bovine serum albumin (BSA; 0.05 mg/ml), and 1 mM DTT]. Assays involving the P_DNA-T₁₂₅, P_RNA-T₆₀, P_RNA-T₁₂₅, and P_RNA-T_mix substrates were conducted in the buffer [50 mM tris-HCl (pH 8.0), 150 mM NaCl, 5 mM MgCl₂, BSA (0.05 mg/ml), and 1 mM DTT]. Reactions were initiated by adding 1 μM P-T substrate and indicated nucleotide mixtures and then incubated at 37°C for the reaction unless otherwise specified. For DNA-primed P-T substrates, reactions were supplemented with 0.1 mM 2′-deoxyadenosine 5′-triphosphate (dATP). For RNA-primed P-T substrate, the nucleotide mixture contained 0.1 mM dCTP-Cy3 and stoichiometric concentrations of deoxynucleotide triphosphates (dNTPs) calculated proportionally to the required extension length. Specifically, 4.5 mM dATP was added for the 45-nt extension of P_RNA-T₆₀; 11.0 mM dATP was added for the 110-nt extension of P_RNA-T₁₂₅; and 0.5 mM dATP, 2.3 mM 2′-deoxyguanosine 5′-triphosphate (dGTP), and 1.7 mM 2′-deoxythymidine 5′-triphosphate (dTTP) were added for the mixed sequence extension of P_RNA-T_mix.

At designated time points, reactions were quenched with 2.5 volumes of denaturing loading buffer [90% formamide, 10 mM EDTA, and bromophenol blue (5 mg/ml)]. Samples were denatured at 70°C for 10 min and immediately snap chilled on ice to maintain the single-stranded DNA (ssDNA) state. Extension products were resolved on 18% denaturing polyacrylamide gels containing 8 M urea. Gels were prerun at 55°C for 40 min before loading and electrophoresed at 55°C for 60 min. Fluorescent signals were detected at an excitation wavelength of 568 nm using an Amersham Typhoon scanner (GE Healthcare) or an Amersham Imager 600 RGB system (GE Healthcare), and gels were subsequently stained with SYBR Gold (Invitrogen) and using a ChemiDoc Imaging System (Bio-Rad). Gel densitometry was quantified using ImageJ (v1.54f) with inverted grayscale measurements.

Kinetic data from primer extension assays were analyzed to quantify the full-length products. The time-dependent accumulation of full-length products exhibited a sigmoidal profile. Data were fitted using nonlinear regression in GraphPad Prism (version 9.5.3, GraphPad Software) to a logistic growth model, and best-fit parameters were obtained using the built-in nonlinear curve fitting algorithm with least-squares minimization. Logistic fitting was used as an empirical model to facilitate quantitative comparison between conditions and was not intended to imply a specific mechanistic kinetic scheme (81). The model

Y = Y_{m} / (1 + e^{[- K * (t - t_{1 / 2})]})

In this model, t_1/2 represents the time required to reach half-maximal product formation and serves as a quantitative measure of the lag phase preceding steady-state synthesis. The parameter K corresponds to the apparent elongation rate constant during the exponential phase and reflects the efficiency of productive DNA chain extension following reaction onset.

DNA combing assay

The method was modified from previous studies (82, 83). Yeast cells were cultured in YPD medium at 30°C to OD₆₀₀ of about 0.5 and arrested at the G₁ phase by α-factor (5 mg/ml, 1000×) at 25°C. Cells were spun down at 2500 rpm for 5 min at 4°C, washed twice with cold ddH₂O, and then released into fresh YPD medium with BrdU (0.4 mg/ml) at 23°C for 40 min to label nascent DNA. Cells (about 50 OD₆₀₀) were collected, washed twice with cold ddH₂O, and washed once with cold buffer z [1.2 M sorbitol and 50 mM tris-HCl (pH 7.4)]. The pellets were resuspended with 4.35 ml of buffer Z (10 mM β-mercaptoethanol freshly added) and digested with 106.5 μl of zymolyase at 28°C for about 35 min to get spheroplasts. The efficiency of digestion was evaluated by measuring OD₆₀₀ in 1% SDS, with a decrease of more than 90%. The pellets were then resuspended with 500 μl of buffer z (prewarmed at 42°C) and mixed with 500 μl of 1% low melting point agarose gel (melted at 65°C and kept at 42°C). A total of 90 μl of cell plugs was generated in casting molds (90 μl) and solidified at 4°C for 30 min. The plugs were then digested in proteinase K buffer [10 mM tris-HCl (pH 7.5), 50 mM EDTA, 1% sarkosyl, and proteinase K (2 mg/ml)] twice for 2 days (2 ml of proteinase K buffer for five plugs) at 42°C. After the digestion, plugs were washed with TE₅₀ [10 mM tris-HCl (pH 7.0) and 50 mM EDTA (pH 8.0)] five times for 10 min and can be stored at 4°C in TE₅₀ for a week. One plug was picked up each time and washed once with 5 ml of 1× MES buffer, pH 6.2 (50 mM MES) for 5 min. The 1× MES buffer was diluted from 500 mM MES, pH 6.2, which was adjusted by adding 500 mM MES sodium to 500 mM MES hydrate. The plug was then transferred to 300 μl of fresh 1× MES buffer, pH 6.2, and melted at 65°C for 15 min. After cooling down, 3 μl of β-agarase I was added to the solution, and the digestion was carried out at 42°C overnight. The solution was spun down at 13,300 rpm on the following day. A total of 40 μl of DNA solution in 1× MES buffer, pH 6.2, was added onto a clean glass slide, and a silanized coverslip (COV-001, Genomic Vision) was covered in the solution and incubated for 5 min. Then, the DNA was stretched at a speed of 300 μm/s and cross-linked at 60°C for 2 hours. DNA was fixed in 70, 90, and 100% ethanol for 5 min, respectively, and then denatured in 1 M NaOH for 22 min and neutralized with 1× PBS. BrdU was detected with a rat monoclonal antibody, followed by a goat anti-rat coupled to Alexa Fluor 488, and ssDNA was detected using anti-ssDNA and goat anti-mouse coupled to Alexa Fluor 546. Images of DNA fibers were recorded with Dragonfly (Andor). The number of DNA fibers calculated per sample was more than 120; two biological replicates were scored and statistically analyzed, and a representative result from one replicate was shown.

Chromatin immunoprecipitation–quantitative polymerase chain reaction assay

Briefly, exponentially growing cells (MATa) were synchronized with α-factor (5 μg/ml) for 2 hours at 25°C and then released into YPD medium with or without 200 mM HU (Sigma-Aldrich). Samples were collected at the indicated time points and temperatures for the following ChIP assay. The procedure of the following ChIP was described previously (84) with minor modifications based on special requirements. Cells were incubated with 1% formaldehyde (Sigma-Aldrich) at 25°C for 20 min and quenched with 0.125 M glycine at 25°C for 5 min. After bead beating with glass beads added, the protoplasts were sonicated by a Bioruptor (Diagenode) to chromosomal DNA fragments about 200 to 500 bps in length and then immunoprecipitated with specific antibodies in demand. All ChIP DNA was enriched by applying the indicated antibodies, except that TAP-tagged ChIP requires IgG beads (GE Healthcare) without further recognition by Protein G Sepharose (GE Healthcare). After extensive washing, the DNA sample was boiled with 10% Chelex 100 (Bio-Rad) for reverse cross-linking. The final ChIP DNA products were analyzed by Real-Time PCR (CFX96, Bio-Rad) using primers that amplify replication origins. The percentage of ChIP DNA relative to total input DNA was calculated. Three biological replicates were performed and shown.

Budding yeast Okazaki fragment purification and in vitro labeling

Cells carrying Cdc9-AID* degron were cultured to an OD₆₀₀ of about 0.4. 3-indoleacetic acid was added to the final concentration of 1 mM and induced at 30°C for 3 hours. A total of 50 ml of cells was collected, centrifuged at 3000g for 5 min, and washed with 10 ml of SCE buffer [1 M sorbitol, 100 mM sodium citrate, and 60 mM EDTA (pH 7.0)]. A total of 5 mg of 20T zymolase, together with β-mercaptoethanol, was added to digest the cell wall at 28°C for 30 min. Then, the spheroblasts were washed with 10 ml of SCE buffer and digested with 480 μl of lysis buffer containing 150 μg of proteinase K (Thermo Fisher Scientific) at 37°C for 2 to 16 hours. The remaining proteins or peptides were precipitated by adding 200 μl of 5.0 M KOAc and spun down at 16,000g for 30 min. The supernatant was collected, and 500 μl of isopropanol was added to precipitate the DNA, which was then centrifuged at 16,000g for 10 min. The pellet DNA was washed twice with 500 μl of 70% ethanol and centrifuged at 16,000g for 1 min. Then, the pellet was resuspended using 200 μl of STE buffer, and 25 μg of ribonuclease A was added to digest RNA at 37°C for 1 hour. The DNA was precipitated by adding 20 μl of 3 M NaOAc (pH 5.5) and 800 μl of ethanol and then was centrifuged at 5000g for 10 min. The pellet was washed once with 1 ml of ethanol and lastly resuspended in 50 μl of TE buffer. The DNA sample should be stored at 4°C and cannot be frozen.

For end labeling of the Okazaki fragment by radioactive probes, the total reaction system is 20 μl, containing 5 U of Klenow (exo⁻) polymerase, 33 nM α-³²P-dCTP, and 2 μl of DNA sample. The reaction is carried out at 37°C for 30 min and then inactivated at 75°C for 20 min. Using Illustra MicroSpin G-50 columns (GE Healthcare), the free radioactive probes were removed, and the DNA sample was eluted. Alkaline loading buffer was added to the DNA sample, and a 10 μl aliquot was loaded onto a 2% alkaline agarose gel to separate the DNA at 150 V for 3 hours (using ice to keep the buffer cool). After electrophoresis, DNA was transferred to the positively charged nitrocellulose film (Hybond-N⁺, GE Healthcare) overnight. The film was exposed to the phosphor screen and scanned by the Typhoon FLA 9500 imager (GE Healthcare). Using ImageJ (v1.54f), the grayscale intensity of bands was quantified.

Statistics and reproducibility

At least two independent experiments were performed to generate each dataset. Statistical analysis was performed by GraphPad Prism 9.5.3. The statistical test used for each experiment is stated in the figure legends.

Acknowledgments

We thank all members of the Li laboratory for technical and intellectual support throughout the project. We thank the National Center for Protein Sciences (Beijing) and the Core Facilities of Life Sciences at Peking University for assistance.

Funding:

This work was supported by the National Natural Science Foundation of China (NSFC 32530020 and 31830048 to Q.L. and NSFC 32370623 to J.F.), and the Fundamental and Interdisciplinary Disciplines Breakthrough Plan of the Ministry of Education of China (JYB2025XDXM503). Q.L. is also supported by the Hong Kong Fong’s Family Foundation.

Author contributions:

Conceptualization: Q.L., J.F., and N.G. Formal analysis: W.Z., S.G., J.X., J.F., and Q.L. Funding acquisition: Q.L. and J.F. Investigation: W.Z., S.G., J.X., Y.T., G.S., G.J., F.Q., J.Y., J.F., and Q.L. Methodology: W.Z., S.G., J.X., J.F., and Q.L. Project administration: Q.L. Supervision: Q.L., J.F., and N.G. Visualization: W.Z., S.G., J.X., J.F., and Q.L. Writing—original draft: W.Z., S.G., J.X., J.F., and Q.L. Writing—review and editing: W.Z., S.G., J.X., N.G., J.W., J.F., and Q.L.

Competing interests:

The authors declare that they have no competing interests.

Data, code, and materials availability:

All data and code needed to evaluate and reproduce the results in the paper are present in the paper and/or the Supplementary Materials. Plasmids and yeast strains generated in this study are available from the corresponding author upon request without restriction (li.qing@pku.edu.cn). Details regarding the generated materials can be found in tables S1 and S2 in the Supplementary Materials.

Supplementary Materials

This PDF file includes:

Figs. S1 to S27

Tables S1 to S5

References

sciadv.aeh7889_sm.pdf^{(4.3MB, pdf)}

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Associated Data

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

Supplementary Materials

Figs. S1 to S27

Tables S1 to S5

References

sciadv.aeh7889_sm.pdf^{(4.3MB, pdf)}

Data Availability Statement

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PERMALINK

FACT–(H3-H4) complex stimulates Pol α activity to coordinate DNA synthesis with nucleosome assembly

Wenshuo Zhang

Shichen Geng

Jiawei Xu

Yuantao Tang

Guanzhong Jiao

Jiayi Yang

Guojun Shi

Jiale Wu

Fangnan Qin

Ning Gao

Jianxun Feng

Qing Li

Roles

Abstract

INTRODUCTION

RESULTS

Histone H3-H4, but not H2A-H2B, promotes the interaction between FACT and Pol α

Fig. 1. H3-H4, but not H2A-H2B, promotes the interaction of FACT with DNA Pol α.

The FACT–(H3-H4)–Pol α ternary complex enhances the DNA polymerase activity of Pol α

Fig. 2. FACT–(H3-H4) promotes primer extension activity of Pol α in vitro.

FACT–(H3-H4) stimulates Pol1 primer extension on both RNA- and DNA-primed templates by accelerating DNA synthesis

Fig. 3. FACT–(H3-H4) promotes Pol α activity on RNA-primed template and accelerates its synthesis rate.

The N terminus of Pol1 contributes to the FACT–(H3-H4)–Pol1 interaction

Fig. 4. Pol1-N domain is critical for the formation of the FACT–(H3-H4)–Pol α ternary complex.

The FACT–(H3-H4)–Pol α interaction is required to maintain replication speed and Okazaki fragment synthesis

Fig. 5. The ability of Pol1-N to bind FACT–(H3-H4)2 is essential for replication elongation and Okazaki fragment synthesis.

The FACT–(H3-H4)–Pol α interaction senses the RC nucleosome assembly level to adjust the replication speed

Fig. 6. CAF-1 and Rtt106 influence the assembly of the FACT–(H3-H4)–Pol α complex.

DISCUSSION

Fig. 7. Model of the chromatin assembly–regulated early-warning system mediated by the FACT–(H3-H4)–Pol α complex.

MATERIALS AND METHODS

Yeast strains and culture conditions

Bimolecular fluorescence complementation

Intensity-based FRET measurement with three filter microscopies

Tandem affinity purification

Protein purification for in vitro assays

In vitro pull-down assay

In vitro primer extension assay of Pol α

DNA combing assay

Chromatin immunoprecipitation–quantitative polymerase chain reaction assay

Budding yeast Okazaki fragment purification and in vitro labeling

Statistics and reproducibility

Acknowledgments

Funding:

Author contributions:

Competing interests:

Data, code, and materials availability:

Supplementary Materials

This PDF file includes:

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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Links to NCBI Databases

Fig. 5. The ability of Pol1-N to bind FACT–(H3-H4)₂ is essential for replication elongation and Okazaki fragment synthesis.