RPA transforms RNase H1 to a bidirectional exoribonuclease for processive RNA–DNA hybrid cleavage

Li, Yanan; Liu, Chao; Jia, Xinshuo; Bi, Lulu; Ren, Zhiyun; Zhao, Yilin; Zhang, Xia; Guo, Lijuan; Bao, Yanling; Liu, Cong; Li, Wei; Sun, Bo

doi:10.1038/s41467-024-51984-5

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Published: 29 August 2024

RPA transforms RNase H1 to a bidirectional exoribonuclease for processive RNA–DNA hybrid cleavage

Yanan Li¹^na1,
Chao Liu^2,3^na1,
Xinshuo Jia¹,
Lulu Bi¹,
Zhiyun Ren^1,4,
Yilin Zhao¹,
Xia Zhang¹,
Lijuan Guo¹,
Yanling Bao¹,
Cong Liu ORCID: orcid.org/0000-0003-3425-6672⁵,
Wei Li ORCID: orcid.org/0000-0002-6235-0749^2,3,4 &
…
Bo Sun ORCID: orcid.org/0000-0002-4590-7795¹

Nature Communications volume 15, Article number: 7464 (2024) Cite this article

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Abstract

RNase H1 has been acknowledged as an endoribonuclease specializing in the internal degradation of the RNA moiety within RNA–DNA hybrids, and its ribonuclease activity is indispensable in multifaceted aspects of nucleic acid metabolism. However, the molecular mechanism underlying RNase H1-mediated hybrid cleavage remains inadequately elucidated. Herein, using single-molecule approaches, we probe the dynamics of the hybrid cleavage by Saccharomyces cerevisiae RNase H1. Remarkably, a single RNase H1 enzyme displays 3′-to-5′ exoribonuclease activity. The directional RNA degradation proceeds processively and yet discretely, wherein unwinding approximately 6-bp hybrids as a prerequisite for two consecutive 3-nt RNA excisions limits the overall rate within each catalytic cycle. Moreover, Replication Protein A (RPA) reinforces RNase H1’s 3′-to-5′ nucleolytic rate and processivity and stimulates its 5′-to-3′ exoribonuclease activity. This stimulation is primarily realized through the pre-separation of the hybrids and consequently transfers RNase H1 to a bidirectional exoribonuclease, further potentiating its cleavage efficiency. These findings unveil unprecedented characteristics of an RNase and provide a dynamic view of RPA-enhanced processive hybrid cleavage by RNase H1.

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Introduction

Nucleases are a family of phosphodiesterases tasked with catalyzing the cleavage of the phosphodiester bonds that link nucleotides within nucleic acids¹. Based on the cleavage mode, nucleases can be classified into endonucleases and exonucleases. The main distinction lies in that endonucleases internally introduce cleavage sites along nucleic acids, whereas exonucleases continuously digest nucleic acids from their ends, producing mononucleotides or oligonucleotides as products^1,2. RNase H are a class of evolutionarily conserved nucleases that appear ubiquitous in bacteria, archaea, and eukaryotes^3,4. Since their discovery, RNase H have been widely recognized as endoribonucleases that specifically catalyze the elimination of the RNA moiety of RNA–DNA hybrids and R-loops in a sequence non-specific manner^5,6. RNase H are generally involved in the maintenance of genome stability, such as R-loop suppression, DNA replication, homologous recombination (HR)-directed DNA double-strand break (DSB) repair, and ribonucleotide excision repair. Their significance is underscored by the genetic linkage to defective mitochondrial DNA replication and a severe neurological disorder called Aicardi-Goutières syndrome (AGS)^7,8. Among their multifaceted cellular functions, RNase H must deal with hybrids and R-loops with distinct lengths, from single base to several kilobase pairs^9,10,11. Many vital functions and properties of RNase H have been brought to light in the past few decades^3,4,7,12,13. However, one intriguing question that remains unanswered is how RNase H at a physiological concentration of a few nanomolar range effectively degrade a long stretch of an RNA–DNA hybrid or an R-loop in cells¹⁴.

Based on the sequence similarities and biochemical properties, eukaryotic RNase H are classified into two types (RNase H1 and H2)³. While their precise cellular functions may still be under debate, a consensus seems to have emerged that RNase H1 and H2 have overlapping and specialized roles in hydrolyzing illicit hybrids^3,4,7,12,13. In contrast to RNase H2, RNase H1 can work independently of the cell cycle and is more effective in suppressing stable R-loops^15,16. Notably, unlike other RNase H that cleave the hybrids distributively, human RNase H1 is distinct in processing RNA–DNA hybrids processively¹⁷. Structural and biochemical analysis revealed that the unique N-terminal hybrid binding ___domain (HBD) of RNase H1 that is connected to a C-terminal catalytic ___domain (RNase HC) via a linker of variable length may bestow the processivity^17,18. This feature holds promise for comprehending RNase H degrading long hybrids and R-loops in vivo. However, the dynamic molecular mechanism underlining RNase H1’s processive cleavage is still lacking.

It is widely accepted that the nuclease activity of RNase H1 can be regulated by associated proteins^19,20,21,22. A key partner of RNase H1 is the single-stranded DNA binding protein, Replication Protein A (RPA), a heterotrimeric protein complex consisting of the RPA70, RPA32, and RPA14 subunits, which also plays a crucial role in DNA replication and recombination^22,23,24,25. An in vivo study showed that human RPA co-localizes with both RNase H1 and R-loops in cells, suggesting that these two proteins function together in the R-loop regulation²². In addition, biochemical studies have shown that human RPA not only physically interacts with RNase H1 but also stimulates its nuclease activity²². Notably, this interaction is specific as it is undetectable between RNase H2 and RPA²². Consistently, Escherichia coli single-stranded DNA binding protein (SSB) has also proved to interact with RNase HI, the prokaryotic counterpart of RNase H1, and stimulates its nuclease activity^21,22. Therefore, it seems plausible that RPA coordinates with RNase H1 in processing long hybrids. A thorough understanding of the regulatory role of RPA in RNase H1-mediated hybrid cleavage would advance our knowledge in genome maintenance.

In this work, we explore the dynamics of Saccharomyces cerevisiae (sc) RNase H1-mediated RNA–DNA hybrid cleavage using single-molecule approaches. We found that scRNase H1 (hereafter referred to as RNase H1) exhibited concentration-dependent nuclease activities. A high amount of RNase H1 proteins function mainly as endonucleases that sufficiently bind to a long hybrid and introduce multiple cleavage sites, allowing quick RNA and nuclease dissociation from the DNA. On the other hand, following the RNA nicking, a single RNase H1 nuclease assumes a motor-like role to continuously degrade thousands of base pairs of RNA within a hybrid in the 3′-to-5′ direction before dissociation. In each exonucleolytic cycle, unwinding approximately 6-bp hybrids occurs before two consecutive 3-nt RNA excisions and is the major rate-limiting step. Importantly, besides enhancing the 3′-to-5′ hybrid degradation of RNase H1, RPA helps trap thermally frayed ssDNA, allowing RNase H1 to carry out the opposite 5′-to-3′ RNA degradation. The bidirectional exonuclease activity of RNase H1 allows for a processive and fast hybrid degradation. Overall, this work unveils distinctive RNase characteristics and offers insights into the cleavage of long RNA–DNA hybrids and R-loops in cells.

Results

RNase H1 possesses both endoribonuclease and exoribonuclease activities

To explore the dynamics of RNase H1-mediated hybrid cleavage, we used dual optical tweezers to monitor the change in the length of a single RNA–DNA hybrid molecule in real time²⁶. In this experiment, a 7.4-kbp RNA–DNA hybrid molecule was suspended between one fixed trap and one steered trap via two streptavidin-coated microspheres, and a high-frequency feedback system on the steered trap was employed to ensure the force on the hybrid remained constant (Fig, 1a, Supplementary Fig. 1). Once RNase H1 eliminates the RNA within the suspended hybrid under a constant force, the increase in the molecule length would be expected due to the difference in the elastic response between single-stranded (ss) DNA and double-stranded hybrid (Supplementary Fig. 1c). We started this experiment with 2 nM RNase H1 under a force of 10 pN. As expected, the hybrid length continuously increased at a rate of 3.2 ± 0.1 nm/s (mean ± SEM) (Fig. 1b, c, Supplementary Fig. 2a). Similar results were also obtained under 5 nM RNase H1 with a moderately increased rate of 7.8 ± 0.5 nm/s (mean ± SEM) (Fig. 1b, c, Supplementary Fig. 2b). However, upon elevating the RNase H1 concentration to 50 nM, the molecule length dramatically increased, and the rate is approximately 76-fold (243 ± 11 nm/s, mean ± SEM) faster than that under 2 nM RNase H1 (Fig. 1b, c, Supplementary Fig. 2c).

**Fig. 1: Endoribonuclease and exoribonuclease activities of RNase H1 toward the RNA–DNA hybrid.**

The distinct rates detected in the single-molecule assay could be attributable to different types of ribonuclease activities of RNase H1 toward the hybrids. We combined dual optical traps with confocal microscopy to test this hypothesis. Through repeatedly scanning along the suspended hybrid, this combination allowed us to simultaneously record the hybrid length and the fluorescence signals of either the hybrid or RNase H1 during the cleavage (Fig. 1a). Using UTP-Cy3 as one of the nucleotides for RNA synthesis (see Methods), we first constructed a 12.3-kbp fluorescently labelled RNA–DNA hybrid molecule and repeated the optical tweezers assay (Supplementary Fig. 1a). As shown in Fig. 1d and Supplementary Fig. 3a, the transportation of this fluorescently labelled hybrid molecule to a channel containing 50 nM RNase H1 caused an immediate increase in the hybrid length under 10 pN. At the same time, the fluorescence signal of the RNA disappeared drastically, indicating the dissociation of the RNA from the hybrid. We also conducted experiments using 50 nM Cy3-labelled RNase H1 (RNase H1-Cy3). The association of multiple RNase H1 with the hybrid was immediately noticed in the protein channel. However, this association was followed by a quick disappearance of the fluorescence signal and increased hybrid length within a few seconds (Fig. 1e, Supplementary Fig. 3b). The intense binding of RNase H1 to the hybrid and the quick dissociation of both RNase H1 and RNA support the dominant endonuclease activity that adequately introduces multiple internal cleavage sites within the hybrids, causing the simultaneous dissociation of small RNA fragments and RNase H1 proteins from the hybridized DNA and the drastic increase in the hybrid length. However, in contrast, RNase H1 at 5 nM caused a moderate increase in the length of a fluorescently labelled hybrid (Fig. 1f). Concurrently, a dark region in the middle of the hybrid appeared and subsequently expanded in one direction, suggesting directional RNA degradation (Fig. 1f, Supplementary Fig. 3c). Experiments with RNase H1-Cy3 (5 nM) demonstrated that the increased hybrid length was coupled with the directional movement of the nuclease along the hybrid (Fig. 1g, Supplementary Fig. 3d). Meanwhile, replacing Mg²⁺ with Ca²⁺ abolished the movement and the increase in the hybrid length (Supplementary Fig. 4a). The photobleaching assay confirmed the monomeric status of the bound RNase H1 proteins (Supplementary Fig. 4b). Moreover, the waiting time before a single RNase H1-Cy3 protein associating with the hybrid is typically around tens of seconds under this condition, ruling out the possibility that the increase in the hybrid length is a result of multiple RNase H1 proteins (Supplementary Fig. 4a). Based on these findings, the directional RNA degradation observed is attributable to the exonuclease activity of a single RNase H1 protein. Increasing the RNase H1 concentration to 10 nM and 15 nM allowed us to monitor the RNA degradation in company with the simultaneous movements of a few RNase H1-Cy3 proteins in the same direction, further corroborating the exonuclease activity of RNase H1 (Supplementary Fig. 5). It is conceivable that a single RNase H1 enzyme initially functions as an endonuclease, creating a nick in the hybrid, following which it transforms to an exonuclease and directionally degrades RNA from the nick. It is also noteworthy that RNase H1 has a binding preference for the nicked hybrid (Supplementary Fig. 6).

Taken together, we provided direct evidence that scRNase H1 can act as an endonuclease that distributes internal hybrid cleavage and as an exoribonuclease that digests RNA processively. The protein concentration is a determinant of the nucleolytic type of RNase H1. It is imaginable that the adequate coating of RNase H1 along the hybrid would restrict their active movement for exonucleolytic cleavage, wherein their endonuclease activity likely plays a dominant role in RNA degradation.

RNase H1 is a 3′-to-5′ exoribonuclease toward RNA–DNA hybrids

We next asked whether RNase H1 as an exoribonuclease has a confined direction in catalyzing the removal of the RNA in the hybrids. To this aim, we constructed a 12.3-kbp RNA–DNA hybrid template with a digoxigenin at the 3′ end of the DNA and a biotin at its 5′ end. In the optical tweezers assay, a streptavidin-coated microsphere with a diameter of 1.76 μm and an anti-digoxigenin-coated microsphere with a diameter of 2.01 μm were used to suspend this hybrid molecule (Fig. 2a). The distinct size of the two microspheres provides a means to distinguish the directionality of the movement of RNase H1 along the hybrid: RNase H1 would translocate toward the smaller bead if it is a 3′-to-5′ exonuclease or toward the larger bead if the opposite (Fig. 2a). In 20 examined exonucleolytic degradation events, all RNase H1-Cy3 exonucleases exhibited translocation toward the smaller bead, suggesting that RNase H1 is indeed a 3′-to-5′ exoribonuclease (Fig. 2a, Supplementary Fig. 7). Analysis of individual exonucleolytic events revealed its average degradation rate of 29 ± 1 (mean ± SEM) nt/s and processivity of 1.2 ± 0.5 (mean ± SD) knt (Fig. 2b, c) by RNase H1-Cy3 signal. A comparable degradation rate was obtained from the analysis of the fluorescence signal of RNA-Cy3 (Supplementary Fig. 8). Reducing the force applied on the hybrid moderately decreased the cleavage rate (Supplementary Fig. 9). Please note that the processivity is possibly underestimated due to photobleaching of RNase H1-Cy3 and the limited length of the hybrid used in the assay. Our optical tweezers assay demonstrated the internal exonucleolytic degradation of RNA by RNase H1. We next addressed whether RNase H1, like other exonucleases, can also initiate the hybrid cleavage from the end of the RNA. To answer that, we employed a previously developed single-molecule fluorescence resonance energy transfer (smFRET) approach to examine the cleavage of a short RNA–DNA hybrid by RNase H1^27,28,29. In this assay, the DNA in the hybrid was labelled with a donor (Cy3) at the 5′ end and an acceptor (Cy5) at 20 nt from the 5′ end (Supplementary Table 1). The 5′ end of the paired RNA was labelled with biotin for the immobilization on a PEG streptavidin-coated quartz surface (hereafter, this substrate was referred to as 3′-RDH). The 3′-to-5′ degradation of the biotin-labelled RNA strand by RNase H1 would generate a 5′-overhang of the fluorescently labelled ssDNA strand, thereby increasing FRET efficiency (E) due to the intrinsic flexibility of the remaining ssDNA (Fig. 2d). To ensure the exonuclease activity of RNase H1, we chose to first incubate the 3′-RDH substrate with 5 nM RNase H1 and 0.5 mM Ca²⁺ for 3 min at room temperature. According to our fluorophore quenching experiments, this incubation permits nearly all the substrates bound by a single RNase H1 without RNA degradation (Supplementary Figs. 10a and 11), which gave rise to the E efficiency of ~0.18 (Fig. 2d). After that, we washed out free RNase H1 proteins and initiated the RNA degradation by replacing Ca²⁺ with Mg²⁺. Upon adding Mg²⁺, the E histogram showed an apparent shift from 0.18 to higher values in 100 s, with a majority centred at 0.96 and a small portion at 0.45 (Fig. 2d). A 200-s reaction resulted in a similar FRET histogram, indicating adequate reactions (Fig. 2d). Intuitively, the inefficient endonucleolytic cleavage may induce partial RNA elimination, therefore, a gentle increase in E. However, the finding that E increased to 0.96 suggests a thorough digestion of the RNA between the two fluorophores and favours RNase H1′s processive degrading initiated from the 3′ end of the RNA. To further corroborate that, we reversed the hybrid substrate by anchoring the 3′ end of the RNA on the surface (5′-RDH) (Fig. 2e, Supplementary Table 1). In this experimental configuration, the DNA was labelled with a donor (Cy3) at the 3′ end and an acceptor (Cy5) at 20 nt from the 3′ end (5′-RDH). After confirming that most of the substrates were bound by a single RNase H1 (Supplementary Fig. 10b), we examined the cleavage of the 5′-RDH substrate. Unlike the 3′-RDH substrate, adding Mg²⁺ to the RNase H1-bound 5′-RDH substrate caused a negligible increase in E (Fig. 2e). Yet, the total quantity of the monitored substrates disappeared much faster than the 3′-RDH substrate within 70 s (Fig. 2f, g). The distinct outcomes of these two substrates supported the exonuclease activity of RNase H1, as similar FRET distributions would be expected if it were a result of the endonuclease activity. These findings also confirmed the 3′-to-5′ cleavage of RNase H1 initiated from the 3′ end of RNA within the hybrids. The increase in E recorded with the 3′-RDH substrate originates from the directional exoribonucleolytic cleavage by RNase H1. The rapid disappearance of the fluorescence signals of the 5′-RDH can be explained by the preferential 3′ RNA binding and nicking of RNase H1 that leads to the detachment of the substrate from the surface.

**Fig. 2: RNase H1 is a 3′-to-5′ exoribonuclease.**

Stepwise RNA cleavage by RNase H1 reveals a dominant rate-limiting step of hybrid unwinding

Next, we examined the detailed dynamics of RNase H1-mediated 3′-RDH cleavage. By carefully analyzing the real-time FRET trajectories, we uncovered a discrete stepwise increase in FRET efficiency, with each increase cycle constituting a burst and a dwell. (Fig. 3a). Taking advantage of an automated step-finding algorithm³⁰, we pinpointed four dominant FRET states that are unevenly distributed in the range of 0.18 to 0.93 (Fig. 3b). A transition density plot (TDP) demonstrated the two-dimensional step for pairs of FRET values before and after each transition^27,29,30. This plot presents three stepwise increments in the FRET efficiency between the four states (Fig. 3b). Statistical analysis revealed that the dwell-time histogram follows an exponential decay, supporting a single dominant rate-limiting step during the cleavage (Fig. 3c)^27,29,31,32.

Next, we aimed to determine the rate-limiting step in RNase H1-mediated exonucleolytic cleavage. Regarding double-stranded nucleic acids (dsNA) cleavage, exonucleases, such as EXD2 and Rrp44, often involve a major rate-limiting step of dsNA unwinding in their catalytic cycles^27,29. We thus asked whether RNase H1-mediated cleavage necessitates the RNA–DNA hybrid unwinding before nucleotide excision. It is well acknowledged that the ratio of the GC content in the hybrid substrate determines the energy of base stacking and hydrogen bonding^29,33,34. If the hybrid unwinding serves as the rate-limiting step, increasing the GC content of the hybrid substrate would prolong the dwell time in its catalytic cycle and reduce the kinetic rate. With this notion in mind, we prepared another RDH substrate and increased the GC content from 50% (3′-RDH) to 88% (GC-RDH) to attenuate the hybrid unwinding, if there is any (Fig. 3d, Supplementary Table 1). Real-time trajectories and the TDP obtained with this GC-RDH substrate consistently displayed three stepwise increments in E, resembling the observations with the 3′-RDH substrate (Fig. 3d, e). However, although the dwell-time histogram also showed a single exponential decay, the fitting results yielded a reduced kinetic rate compared to the 3′-RDH substrate (Fig. 3f, j). These findings suggest that the hybrid unwinding is likely a prerequisite for the RNase H1 exonuclease to excise nucleotides and possibly dominates the overall catalytic rate. These conclusions are further sustained by the smFRET experiment with an AU-RDH substrate containing 12% of GC pairs (Fig. 3g, Supplementary Table 1). The introduction of RNase H1 to this substrate led to a similar three-step increase in E (Fig. 3g, h). The dwell-time analysis reveals a kinetic rate twice faster than that of the 3′-RDH substrate (Fig. 3i, j). Please note that this rate is underestimated as real-time trajectories containing one-step and two-step increments in E from 0.21 to 0.93 were also detectable with the AU-RDH substrate (Supplementary Fig. 12). Given that the nucleotide excision is an irreversible process, the notion that RNase H1 unwinds the hybrid prior to cleavage is further substantiated by the transient backward stepping in the process of RNase H1 degrading all three substrates (Supplementary Fig. 13) and the sudden FRET bursts monitored when RNase H1-D264N, a nuclease-deficient mutant²², acted on the 3’-RDH substrate (Supplementary Fig. 14).

We further asked how many base pairs of the hybrid duplex are unwound and digested in each catalytic cycle. The increase in E from 0.18 to 0.93 corresponds to approximately 17 ribonucleotides digested by RNase H1 (Supplementary Fig. 15), and the three-step increments thus predict ~ 6-bp (17/3) duplex unwinding in each cycle under the assumption that they are equal. To test this hypothesis, we prepared a tailed RDH substrate containing a 6-nt ssDNA at the 5′ end (6t-RDH) to mimic a hydrolytic intermediate of the 3′-RDH substrate (Supplementary Fig. 16a, Supplementary Table 1). The smFRET experiments with this substrate revealed an initial E state of ~0.42 after the binding of RNase H1. In the process of RNA degradation, two sudden increments in E from 0.42 to 0.96 were monitored with this substrate, which coincided with the last two steps detected with the 3′-RDH substrate (Supplementary Fig. 16b). Consistently, experiments with the 12t-RDH hybrid substrate (12-nt ssDNA at the 5′ end) revealed one sudden increment in E from 0.65 to 0.96 (Supplementary Fig. 16c, d, Supplementary Table 1). The missing steps and the alignment of the remaining states support the cyclic 6-base pair hybrid unwinding by RNase H1. We carried out similar smFRET experiments with differently labelled hybrid substrates and ruled out the possibility that the unwinding step depends on the fluorescence labelling (Supplementary Fig. 17). To further determine the exonucleolytic cleavage products of RNase H1, we performed bulk cleavage assays with a 53-nt RNA–DNA hybrid substrate. Within this substrate, the 5′ end of the ssRNA was labelled with FAM for fluorescence detection in the PAGE gels. To distinguish the intermediate cleavage products, we supplied RNase H1 with 0.07 mM Mg²⁺ to retard the RNA degradation. Under this condition, the cleavage products exhibited a 3-nt periodicity (Fig. 3k). In comparison to the 6-bp base pairs involved in each step detected in the smFRET assay (6 bp), the shorter 3-nt nucleolytic products further corroborate that the hybrid unwinding occurs prior to the RNA cleavage for RNase H1.

Based on these findings, we conclude that scRNase H1 as an exonuclease unwinds 6-bp before two consecutive 3-nt RNA excisions and ascribes the dominant rate-limiting step to the hybrid unwinding (Fig. 3l).

RPA stimulates RNase H1’s 5′-to-3′ exoribonuclease activity

Since RPA is an essential partner of RNase H1, we next investigated how RPA regulates the nuclease activity of RNase H1. We purified the S. cerevisiae (sc) RPA and revisited the experiments with its involvement. As confirmed by the bulk assay, scRPA (hereafter referred to as RPA) also promotes RNase H1 in degrading RNA–DNA hybrid, in agreement with a previous study on human RNase H1 and RPA (Supplementary Fig. 18a, b)²². Consistently, our single-molecule assay substantiated that the presence of RPA increased the nucleolytic rate of RNase H1 (5 nM) under 10 pN and that the depletion of Mg²⁺ abolished its nucleolytic activity (Fig. 4a, b, Supplementary Figs. 18c and 19). Next, we fused the RPA with an enhanced green fluorescent protein (RPA-eGFP) to directly monitor the dynamic nucleolytic process. Taking advantage of the preferential binding of RPA to ssDNA, we harnessed the fluorescence signal of RPA-eGFP along the hybrid substrate as an indicator for RNA elimination. In line with the unidirectional exonucleolytic cleavage by RNase H1, we observed the unidirectional expansion of RPA-eGFP along the hybrid coupled with increased hybrid length, which also proceeded in the 3′-to-5′ directionality (Fig. 4c, Supplementary Fig. 20). To our surprise, the kymograph of the tensioned hybrid molecule also showed a bidirectional expansion of RPA (Fig. 4d). Furthermore, simultaneous monitoring of RNase H1 and RPA revealed that single RNase H1 nucleases persistently associated at the active ss-ds hybrid junctions in both the unidirectional and bidirectional hybrid cleavage events (Supplementary Fig. 21). There findings suggest both the 3′-to-5′ and 5′-to-3′ exonuclease activities of RNase H1. Notably, the processivity and the percentage of the bidirectional exonucleolytic events of RNase H1 increased with RPA concentration (Fig. 4e, f). The average bidirectional exonucleolytic rate is 1.6-fold higher than the unidirectional one, indicating that the exonucleolytic rates for both directions might differ (Fig. 4g). To verify further, we calculated the individual rates in one direction within the bidirectional events and unveiled two distinct populations (Fig. 4h). The faster rate (56 nt/s) is comparable with that from the unidirectional events (49 nt/s). It possibly signifies the 3′-to-5′ exonuclease activity of RNase H1.

**Fig. 4: The regulation of the exonuclease activity of RNase H1 by RPA.**

Pre-separation of the hybrids unleashes the 5′-to-3′ exoribonuclease activity of RNase H1

Having demonstrated that RPA stimulates RNase H1’s 5′-to-3′ exonuclease activity, we next aimed to gain mechanistic insights into this activation. As unveiled by our smFRET assays, RNase H1 necessitates hybrid unwinding before the 3′-to-5′ nucleotide excision (Fig. 3). Moreover, RPA is preferentially associated with thematically frayed ssDNA, relieving the pressure from dsNA reannealing³⁵. These facts motivated us to speculate that RPA may facilitate the transient separation of the 5′ RNA from the paired DNA, facilitating the 5′ RNA degradation by RNase H1. To test that, we prepared a forked RDH substrate, within which the 10-nt ssRNA close to the 5′ biotin end is not matched with the DNA, and the rest of the ssRNA (33 nt) is matched with the ssDNA (Fig. 5a). The partially matched ssDNA was labelled with a donor (Cy3) and an acceptor (Cy5) for the smFRET measurements (Fig. 5a). For this forked substrate, the 3′-to-5′ exonucleolytic cleavage by RNase H1 is supposed to result in a gradual increase in FRET efficiency. In contrast, the 5′-to-3′ exonucleolytic cleavage should cause the detachment of the substrate from the surface, thereby the simultaneous disappearance of both fluorophores (Fig. 5a). Indeed, upon introducing 5 nM RNase H1, we monitored a substantial number of substrates exhibiting the simultaneous disappearance of both fluorophores before or after a gradual increase in E (Fig. 5b, Supplementary Fig. 22). We noticed that the disappearance of both fluorophores could also result from the adequate 3′-to-5′ exonucleolytic cleavage, in which the FRET efficiency was typically over 0.9 before the dissociation (Supplementary Fig. 22). Thus, the sudden fluorophore disappearance before E reaching 0.9 is attributed to RNase H1 degrading RNA from the 5′ end. In statistics, up to 78% of the real-time FRET trajectories showed such type of degradation with the forked substrate, which is nearly negligible with the blunt 3′-RDH substrate (Fig. 5c). These results support that the pre-separation of the hybrid facilitates the 5′ end RNA degradation by RNase H1. To further corroborate that, we examined the cleavage products of the forked substrate by RNase H1 and compared them with the blunt substrate. Consistently, an RNA segment of approximately 16-nt was detected exclusively with the forked substrate in the denaturing PAGE gel, the amount of which increased over time (Fig. 5d).

In addition to RPA, the mechanical force is also known to facilitate the peeling of ssNA from the double-stranded forms^36,37. Therefore, it is speculated that the increased tension on the hybrid may have a chance to stimulate RNase H1’s 5′-to-3′ RNA degradation. To examine the regulatory effect of the force on RNase H1, we revisited the optical tweezers assay with the RNA–DNA hybrid held under a higher force of 15 pN, 20 pN, and 25 pN. The increase in the hybrid length due to the nuclease activity was similarly recorded under these forces and 5 nM RNase H1 (Supplementary Fig. 23a). The cleavage rate and processivity of RNase H1 increased with the force (Supplementary Fig. 23b). As expected, bidirectional exonucleolytic cleavage events on the Cy3-labelled hybrid substrate were monitored under 20 pN and 25 pN, the percentage of which also increased with force (Fig. 5e, f, Supplementary Fig. 23c). The bidirectional cleavage proceeds 1.4-fold faster than the unidirectional cleavage, suggesting inequitable cleavage in opposite directions (Supplementary Fig. 23d). These results confirm the stimulation of the 5′-to-3′ RNA degradation of RNase H1 by force and further support that the pre-separation of the hybrid unleashes this activity.

Discussion

In this work, we investigate the molecular mechanism of S. cerevisiae RNase H1 degrading RNA–DNA hybrids at the single-molecule level. One of the interesting findings is that RNase H1, initially known as an endonuclease, also possesses intrinsic exonuclease activity. RNase H1 continuously eliminates RNA within the hybrid in the 3′-to-5′ direction, which can be initiated internally or from the 3′ end of the RNA (Fig. 6). Interestingly, this distinctive characteristic sets scRNase H1 apart from its E. coli counterpart, RNase HI, which relies on a 5’ end DNA overhang for the 5’-to-3’ exonuclease activity³⁸. Moreover, E. coli RNase HI differs from scRNase H1 in the binding preference for RNA ends with DNA overhangs and yet decreased affinity for blunt ends or internal regions of RNA–DNA hybrids³⁸. This discrepancy is likely due to the marked structural differences between E. coli RNase HI, lacking a hybrid binding ___domain (HBD), and scRNase H1, which features two HBDs^17,18. A few nucleases have been acknowledged to possess both exonuclease and endonuclease activities, and the transitions between these activities are functional in DNA metabolism^39,40,41. For instance, following the introduction of an internal breakage of dsDNA as endonucleases, FEN1 and Mre11 are found to consecutively carry out exonucleolytic cleavage as exonucleases, resulting in a gap or an overhang in the DNA for the following DNA repair^40,41. Therefore, it is plausible that RNase H1 makes full use of its dual enzymatic activities in effectively degrading hybrids without requiring specialized substrates or protein accumulation. In addition, scRNase H1 is capable of recognizing nicks, which may facilitate the identification of the nucleic acids damage and the initiation of the removal of the illicit RNA (Supplementary Fig. 6).

**Fig. 6: A mechanical model of RNase H1-mediated bidirectional processive RNA–DNA hybrids cleavage.**

Taking advantage of the high tempo-spatial resolution of the smFRET method, we also unveiled the dynamics and kinetics of RNA–DNA hybrid cleavage by RNase H1. As an exonuclease, RNase H1 was found to iterate the duplex unwinding and exonucleolytic degradation during processive hybrid cleavage (Fig. 3). A 6-bp duplex is unwound within each catalytic cycle, and instead of mononucleotides, the frayed ssRNA is excised three ribonucleotides at a time for RNase H1 (Fig. 6). This iteration resembles the behaviour of the Rrp44 and EXD2 exonucleases, which also unwind duplexes before nucleotide excision^27,29. This cyclic coupling between duplex unwinding and nucleotide excision could be explained by a spring-loaded mechanism wherein the chemical energy released from the hydrolysis of the nucleic acids chain is converted into elastic energy for unwinding several base pairs in bursts and translocating directionally²⁹. The overall reaction rate of RNase H1 is mainly governed by the hybrid unwinding, coupled with a relatively fast ssRNA excision (Fig. 3). A simple model has been previously proposed to explain the processivity of RNase H1: a single hybrid binding event mediated by the HBD ___domain of RNase H1 permits the engagement of its hydrolyzing ___domain HC with a few adjacent sites on an RNA–DNA hybrid for cleavage⁴². Our findings enrich this model in that RNase H1 is also equipped with a motor-like property that can direct its directional movement along the hybrid after the adjacent RNA cleavage. This property fundamentally enables RNase H1’s processive hybrid cleavage. It is noteworthy that scRNase H1 distinguishes from other RNase H1 nucleases in that it contains two HBD domains^18,43. Whether and if yes, how the two HBD domains contribute to the directional translocation along the hybrid warrants further exploration.

Our work also has important implications for understanding the coordination of RNase H1 and RPA in processing the intracellular RNA–DNA hybrids and R-loops. Based on our findings, two mechanisms underlining the stimulation of RNase H1’s nuclease activity by RPA have been put forth: first, RPA enhances the rate and processivity of the intrinsic 3′-to-5′ exonucleolytic cleavage of RNase H1; second, RPA stimulates RNase H1’s exoribonuclease activity in the 5′-to-3′ direction (Fig. 6). Similar bidirectional stimulation effects of RPA have also been reported on the BLM helicase-mediated dsDNA unwinding⁴⁴. Given that hybrid unwinding is a prerequisite for RNase H1 to cleave ssRNA, it is conceivable that both stimulation pathways for RNase H1 are mainly achieved by RPA destabilizing the hybrid duplex. This notion is further supported by the results from the smFRET assays with the forked substrate and the optical tweezers assays using higher forces (Fig. 5, Supplementary Fig. 23). The direct interactions of RNase H1 with RPA may also help with its exonuclease activity. RPA has been previously demonstrated to promote the association of RNase H1 to RNA–DNA hybrids²². This promotion may help recruit RNase H1 to the hybrid fork and maintain its longstanding association for processive hybrid cleavage (Fig. 4).

The stimulation of RNase H1’s bidirectional hybrid cleavage by RPA provides a clue for understanding the degradation of a long stretch of an RNA–DNA hybrid or an R-loop in DNA repair and recombination^9,10,11. According to the literature, under cellular stress, both RNA–DNA hybrids and R-loops can be quite long and sometimes reach up to 10 kbp in length, while the RNase H1 concentration is estimated to be in a few nM range in cells^11,14. One can imagine that an endonuclease introducing distributive cleavage along such a long substrate would not efficiently remove the RNA. Complete RNA elimination requires multiple rounds of association and dissociation of the endonuclease with the substrate. However, the internal nicking and the bidirectional hybrid cleavage by RNase H1 ensure the best use of the nuclease and allow for the removal of RNA within a long stretch of a hybrid or an R-loop in a timely fashion (Fig. 6). Overall, this work unveiled the dynamics of RNase H1 cleaving hybrid and the regulatory effect of RPA on hybrid cleavage, which help identify the precise cellular functions of RNase H1 and deepen our understanding of the maintenance of genome stability.

Methods

Recombinant protein purification

The plasmid designed for the expression of Yeast RNase H1 was constructed by synthesizing open reading frames and integrating them into pET28a⁴³. The N-terminus of the resulting pET28a-RNase H1 plasmid was fused to a His₆-tag for purification. Staphylococcus aureus sortase A recognition hexa-amino acid sequence (LPETG) was introduced onto the N-terminus of RNase H1 behind the His₆-tag for protein labelling^45,46. RNase H1 mutant, the nuclease-deficient RNase H1-D264N was produced by site-directed mutagenesis (TransGen). The plasmid was transformed into BL21 (DE3) cells (TransGen). This strain was cultured and induced by 1 mM isopropyl-1-thio-D-galactopyranoside (IPTG) at 18 °C for 16 h. Following that, cells were collected by centrifugation and resuspended in the lysis buffer containing 50 mM Tris–HCl (pH 7.5), 300 mM NaCl, 10 mM imidazole, 5% glycerol and 1 mM phenylmethylsulfonyl fluoride, and passed through a homogenizer three times at ~800 bar. The resulting lysate was ultracentrifuged at 11,000 × g for 30 min, and the supernatant was subjected to Ni-NTA resin (TransGen) affinity purification. This bound protein was extensively washed with a solution containing 50 mM Tris–HCl (pH 7.5), 300 mM NaCl, 30 mM imidazole, and 5% glycerol. The protein was eluted using elution buffer containing 50 mM Tris–HCl (pH 7.5), 300 mM NaCl, 300 mM imidazole, and 5% glycerol. The protein sample underwent chromatography for further purification using a 5-ml HiTrap Heparin HP column. Ultimately, the proteins were buffer-exchanged into a storage buffer containing 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, and 10% glycerol and stored at −80 °C. The protein labelling reaction was conducted in 1× sortase buffer containing 50 mM Tris–HCl (pH 7.5), 250 mM NaCl, and 10 mM CaCl₂ at 4 °C overnight with LPETG-tag RNase H1, Staphylococcus aureus sortase A, and the Cy3-CLPETGG (purchased from ChinaPeptides) at a ratio of 1:2:5⁴⁶. Unreacted-free Cy3-LPETGG was removed using a Centrifugal Filters Ultracel - 30 K (Millipore) with a solution containing 50 mM Tris–HCl (pH 7.5), 500 mM NaCl and 5% glycerol. Staphylococcus aureus sortase A was removed through chromatography using a 5-ml Hi-Trap Heparin HP column. Elution of Cy3-labelled RNase H1 occurred around 400–500 mM NaCl, yet the sortase A did not bind to heparin. The labelling efficiency of RNase H1 was typically around 70%. Purified Cy3-labelled RNase H1 was then stored at −80 °C.

For the expression of Yeast RPA, the plasmid was constructed by synthesizing open reading frames of the three units and inserting them into pET28a^47,48. This construct (pET28a-RPA-eGFP) encodes N-terminally His₆-tagged Rfa1 with an eGFP tag at its C-terminus and Rfa2 and Rfa3. Co-expression of these units was carried out in BL21 (DE3) cells (TransGen). RPA-eGFP was purified similarly to a previously described protocol⁴⁹. The expression plasmid was transformed in BL21 (DE3) cells (TransGen), and the expression strain was cultured and induced with 1 mM IPTG at 18 °C for 16 h. Cells were collected by centrifugation and resuspended in a lysis buffer containing 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 10 mM imidazole, 5% glycerol and 1 mM phenylmethylsulfonyl fluoride, and passed through a homogenizer three times at ~800 bar. The lysed dilution was then ultracentrifuged at 11,000 × g for 35 min. The supernatant was applied to a Ni-NTA resin, washed extensively with 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 20 mM imidazole and 5% glycerol, and RPA-eGFP was eluted using elution buffer containing 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 300 mM imidazole and 5% glycerol. The protein sample underwent chromatography for further purification using a 5-ml HiTrap Heparin HP column. Ultimately, the proteins were buffer-exchanged into a storage buffer containing 50 mM Tris–HCl (pH 7.5), 500 mM NaCl, and 10% glycerol and stored at −80 °C.

Preparation of RNA–DNA hybrid templates

The RNA–DNA hybrid templates used in the optical tweezer assays were prepared through the following steps (Supplementary Fig. 1a, b, Supplementary Table 2)⁵⁰. The templates typically consist of a two-sided biotinylated ssDNA and a paired ssRNA. A dsDNA segment was PCR-amplified from the lambda DNA (Thermo). Using this dsDNA segment and a biotin- or digoxigenin-labelled primer, a second asymmetric PCR produced the ssDNA. The resulting dsDNA and ssDNA were separated by agarose gel electrophoresis and purified using GeneJET Gel Extraction Kit (Thermo). The 3′ end biotin labelling of the ssDNA was performed using biotin-dATP (Invitrogen) and terminal transferase (NEB). A dsDNA containing a T7 promoter (T7 DNA) at the 5′ end was PCR-amplified from lambda DNA, from which the ssRNA was transcribed. UTP-Cy3 was used in the transcription for the fluorescence labelling of the template. The final RNA–DNA hybrid templates were produced by annealing the two single-stranded nucleic acids in the TE buffer.

For substrates of smFRET experiments and gel-based bulk cleavage assays, DNA and RNA oligonucleotides with or without labels used to construct the templates were purchased from Sangon Biotech and GenScript, respectively. The RDH substrates for the smFRET experiments and gel-based bulk cleavage assays were produced by mixing fluorophore-labelled DNA or RNA with complementary RNA or DNA at a molar ratio of 1:1.5 in the annealing buffer containing 10 mM Tris–HCl (pH 8.0), 50 mM KCl and 1 mM EDTA. The mixtures were incubated at 95 °C for 3 min and then slowly cooled to room temperature within 3 h. Annealed templates were stored at −80 °C before use. The sequences of the oligonucleotides are listed in Supplementary Table 1.

Single-molecule optical tweezer assay and data analysis

Single-molecule optical tweezer assays were performed at 25 °C on an instrument combining three-colour confocal fluorescence microscopy with dual optical traps (LUMICKS C-trap, Netherland)^51,52. In the assay, an RNA–DNA hybrid molecule was initially captured between two streptavidin-coated polystyrene beads (1.76 μm in diameter, Spherotech) or between one streptavidin-coated polystyrene bead and one anti-digoxigenin-coated bead (2.01 μm in diameter, Spherotech). This hybrid tether was then transferred to channels as specified for each assay. Experiments on RNase H1, RNase H1-Cy3, and/or RPA-eGFP were conducted in the reaction buffer containing 25 mM Tris–HCl (pH 7.5), 50 mM KCl, 1 mM MgCl₂, 0.05 mg/mL BSA, 1 mM DTT, 0.8% d-glucose, 1 mg/ml glucose oxidase, 0.4 mg/ml catalase, and 2 mM Trolox. Imaging of RPA-eGFP utilized a 488-nm excitation laser, while imaging of RNase H1-Cy3 and RNA(Cy3)–DNA hybrid employed a 532-nm excitation laser. The confocal pixel size was set at 50 nm with a pixel dwell time of 1 ms. Custom software from LUMICKS was employed for single-molecule data analysis. Pseudocolors were applied in Zen 3.2 software (Zeiss) for enhanced signal contrast visualization. For fluorescence signals, the total pixel intensities of the DNA–RNA hybrid or ssDNA region were summed up in each frame using Fiji. A linear fitting to the hybrid extension traces measured DNA extension rates in nm/s. The processivity in nm was calculated using fluorescence signals of RNase H1-Cy3 or RPA-eGFP. Based on the elastic parameters of the hybrid and ssDNA, the processivity in nm was converted to knt, and the cleavage rate in nm/s was converted to nt/s (Supplementary Fig. 1b).

Single-molecule FRET assay and data analysis

Single-molecule FRET experiments were performed using TIRF microscopy^27,53. Utilizing an Andor EMCCD camera, fluorescence images were obtained with an integration time of 100 ms (10-Hz frame rate). Quartz slides and coverslips (VWR) were first coated with polyethylene glycol (PEG) for constructing the flow chamber. Next, the coverslip was treated with aminosilane and coated with a mixture of 99% mPEG (m-PEG-5000, Laysan Bio, Inc.) and 1% biotin-PEG (biotin-PEG-5000, Laysan Bio, Inc.). The chamber was then incubated with streptavidin (10 μg/ml) for 5 min. The hybrid substrate, at a concentration of 20 or 30 pM, was introduced into the chamber and immobilized on the quartz surface. After immobilization, free substrates were washed out using 200 μl of the reaction buffer containing 25 mM Tris–HCl (pH 7.5), 50 mM KCl, 0.8% d-glucose, 1 mg/ml glucose oxidase, 0.4 mg/ml catalase, and 2 mM Trolox. Subsequently, 5 nM RNase H1 was injected into the chamber and incubated with the substrates for 5 min. Finally, 1 mM MgCl₂ was introduced into the chamber to wash out free RNase H1 and initiate hybrid cleavage.

A home-built two-colour TIRF microscope was used for imaging. All images were recorded with an exposure time of 100 ms for 600–1000 frames. Each frame was further processed to extract single-molecule fluorescence intensities. The FRET efficiency of a single molecule was approximated as E = I_A/(I_D + I_A), where I_D and I_A are the donor and acceptor’s background and leakage-corrected emission intensities, respectively. Each experiment was performed at least 3 times to ensure reproducibility. Single-molecule trajectories were collected using SPARTAN 3.7.0³⁰. Hidden Markov modelling was performed on smFRET trajectories³⁰. The Baum–Welch algorithm embedded in SPARTAN was used to determine the most likely number of unwinding steps, and the TDPs were correspondingly plotted. Over 70 molecules were analyzed for each experimental condition.

Ensemble hybrid cleavage assays

For the ensemble RNA–DNA hybrid cleavage assay, RNase H1 (15 nM) was incubated with the fluorescently labelled hybrid substrates (15 nM) in the reaction buffer containing 25 mM Tris–HCl (pH 7.5), 50 mM KCl, and Mg²⁺ for the indicated time at room temperature. The reaction was quenched by formamide gel loading buffer supplemented with 50 mM EDTA. The reaction products were resolved by 18% denaturing polyacrylamide gel electrophoresis (7 M urea PAGE) and visualized by phosphorimager (GE Health Care). Each experiment was repeated in triplicate. The intermediate cleavage products were analyzed using Fiji and compared with the 1-nt marker from FAM-RNA by alkaline cleavage.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All data supporting the findings of this study are available within the paper and its Supplementary Information. Source data are provided with this paper.

References

Yang, W. Nucleases: diversity of structure, function and mechanism. Q. Rev. Biophys. 44, 1–93 (2011).
Article PubMed Google Scholar
Lovett, S. T. The DNA exonucleases of Escherichia coli. EcoSal 4, 1128 (2011).
Google Scholar
Hyjek, M., Figiel, M. & Nowotny, M. RNases H: structure and mechanism. DNA Repair 84, 102672 (2019).
Article CAS PubMed Google Scholar
Cerritelli, S. M. & Crouch, R. J. Ribonuclease H: the enzymes in eukaryotes. FEBS J. 276, 1494–1505 (2009).
Article CAS PubMed Google Scholar
Keller, W. & Crouch, R. Degradation of DNA RNA hybrids by ribonuclease H and DNA polymerases of cellular and viral origin. Proc. Natl Acad. Sci. USA 69, 3360–3364 (1972).
Article ADS CAS PubMed PubMed Central Google Scholar
Hausen, P. & Stein, H. Ribonuclease H. An enzyme degrading the RNA moiety of DNA-RNA hybrids. Eur. J. Biochem. 14, 278–283 (1970).
Article CAS PubMed Google Scholar
Cristini, A. et al. RNase H2, mutated in Aicardi-Goutieres syndrome, resolves co-transcriptional R-loops to prevent DNA breaks and inflammation. Nat. Commun. 13, 2961 (2022).
Article ADS CAS PubMed PubMed Central Google Scholar
Misic, J. et al. Mammalian RNase H1 directs RNA primer formation for mtDNA replication initiation and is also necessary for mtDNA replication completion. Nucleic Acids Res. 50, 8749–8766 (2022).
Article CAS PubMed PubMed Central Google Scholar
Garcia-Muse, T. & Aguilera, A. R Loops: from physiological to pathological roles. Cell 179, 604–618 (2019).
Article CAS PubMed Google Scholar
Castillo-Guzman, D. & Chedin, F. Defining R-loop classes and their contributions to genome instability. DNA Repair 106, 103182 (2021).
Article CAS PubMed PubMed Central Google Scholar
Ohle, C. et al. Transient RNA-DNA hybrids are required for efficient double-strand break repair. Cell 167, 1001–1013 (2016).
Article CAS PubMed Google Scholar
Schroeder, J. W. et al. RNase H genes cause distinct impacts on RNA:DNA hybrid formation and mutagenesis genome wide. Sci. Adv. 9, 5945 (2023).
Article Google Scholar
Lai, F., Damle, S. S., Ling, K. K. & Rigo, F. Directed RNase H cleavage of nascent transcripts causes transcription termination. Mol. Cell 77, 1032–1043 (2020).
Article CAS PubMed Google Scholar
Ho, B., Baryshnikova, A. & Brown, G. W. Unification of protein abundance datasets yields a quantitative Saccharomyces cerevisiae proteome. Cell Syst. 6, 192–205 (2018).
Article CAS PubMed Google Scholar
Zhou, H. et al. R-loops mediate transcription-associated formation of human rDNA secondary constrictions. J. Cell Biochem. 122, 1517–1533 (2021).
Article CAS PubMed Google Scholar
Lockhart, A. et al. RNase H1 and H2 are differentially regulated to process RNA-DNA hybrids. Cell Rep. 29, 2890–2900 (2019).
Article CAS PubMed Google Scholar
Gaidamakov, S. A. et al. Eukaryotic RNases H1 act processively by interactions through the duplex RNA-binding ___domain. Nucleic Acids Res. 33, 2166–2175 (2005).
Article CAS PubMed PubMed Central Google Scholar
Gonzalez de Cozar, J. M. et al. A second hybrid-binding ___domain modulates the activity of Drosophila ribonuclease H1. J. Biochem 168, 515–533 (2020).
Article CAS PubMed PubMed Central Google Scholar
Zhang, L. et al. NAT10 and DDX21 proteins interact with RNase H1 and affect the performance of phosphorothioate oligonucleotides. Nucleic Acid Ther. 32, 280–299 (2022).
Article ADS CAS PubMed PubMed Central Google Scholar
Wolak, C. et al. Interaction with single-stranded DNA-binding protein localizes ribonuclease HI to DNA replication forks and facilitates R-loop removal. Mol. Microbiol 114, 495–509 (2020).
Article CAS PubMed PubMed Central Google Scholar
Petzold, C., Marceau, A. H., Miller, K. H., Marqusee, S. & Keck, J. L. Interaction with single-stranded DNA-binding protein stimulates Escherichia coli ribonuclease HI enzymatic activity. J. Biol. Chem. 290, 14626–14636 (2015).
Article CAS PubMed PubMed Central Google Scholar
Nguyen, H. D. et al. Functions of replication protein A as a sensor of R loops and a regulator of RNaseH1. Mol. Cell 65, 832–847 (2017).
Article CAS PubMed PubMed Central Google Scholar
Zou, Y., Liu, Y., Wu, X. & Shell, S. M. Functions of human replication protein A (RPA): from DNA replication to DNA damage and stress responses. J. Cell Physiol. 208, 267–273 (2006).
Article CAS PubMed PubMed Central Google Scholar
Li, S., Dong, Z., Yang, S., Feng, J. & Li, Q. Chaperoning RPA during DNA metabolism. Curr. Genet. 65, 857–864 (2019).
Article CAS PubMed Google Scholar
Feng, S. & Manley, J. L. Replication protein A associates with nucleolar R loops and regulates rRNA transcription and nucleolar morphology. Genes Dev. 35, 1579–1594 (2021).
Article CAS PubMed PubMed Central Google Scholar
Liu, C. et al. RNase H1 facilitates recombinase recruitment by degrading DNA-RNA hybrids during meiosis. Nucleic Acids Res. 51, 7357–7375 (2023).
Article CAS PubMed PubMed Central Google Scholar
Jia, X. et al. Discrete RNA-DNA hybrid cleavage by the EXD2 exonuclease pinpoints two rate-limiting steps. EMBO J. 42, e111703 (2023).
Article CAS PubMed Google Scholar
Roy, R., Hohng, S. & Ha, T. A practical guide to single-molecule FRET. Nat. Methods 5, 507–516 (2008).
Article CAS PubMed PubMed Central Google Scholar
Lee, G., Bratkowski, M. A., Ding, F., Ke, A. & Ha, T. Elastic coupling between RNA degradation and unwinding by an exoribonuclease. Science 336, 1726–1729 (2012).
Article ADS CAS PubMed PubMed Central Google Scholar
Juette, M. F. et al. Single-molecule imaging of non-equilibrium molecular ensembles on the millisecond timescale. Nat. Methods 13, 341–344 (2016).
Article PubMed PubMed Central Google Scholar
Xue, X., Liu, F. & Ou-Yang, Z. C. Single molecule Michaelis-Menten equation beyond quasistatic disorder. Phys. Rev. E Stat. Nonlin Soft Matter Phys. 74, 030902 (2006).
Article PubMed Google Scholar
Lu, H. P., Xun, L. & Xie, X. S. Single-molecule enzymatic dynamics. Science 282, 1877–1882 (1998).
Article ADS CAS PubMed Google Scholar
Han, G. W., Kopka, M. L., Langs, D., Sawaya, M. R. & Dickerson, R. E. Crystal structure of an RNA.DNA hybrid reveals intermolecular intercalation: dimer formation by base-pair swapping. Proc. Natl Acad. Sci. USA 100, 9214–9219 (2003).
Article ADS CAS PubMed PubMed Central Google Scholar
van Oijen, A. M. et al. Single-molecule kinetics of lambda exonuclease reveal base dependence and dynamic disorder. Science 301, 1235–1238 (2003).
Article ADS PubMed Google Scholar
Kemmerich, F. E. et al. Force regulated dynamics of RPA on a DNA fork. Nucleic Acids Res. 44, 5837–5848 (2016).
Article CAS PubMed PubMed Central Google Scholar
King, G. A. et al. Revealing the competition between peeled ssDNA, melting bubbles, and S-DNA during DNA overstretching using fluorescence microscopy. Proc. Natl Acad. Sci. USA 110, 3859–3864 (2013).
Article ADS CAS PubMed PubMed Central Google Scholar
Bosaeus, N., El-Sagheer, A. H., Brown, T., Akerman, B. & Norden, B. Force-induced melting of DNA–evidence for peeling and internal melting from force spectra on short synthetic duplex sequences. Nucleic Acids Res. 42, 8083–8091 (2014).
Article CAS PubMed PubMed Central Google Scholar
Lee, H. et al. RNase H is an exo- and endoribonuclease with asymmetric directionality, depending on the binding mode to the structural variants of RNA:DNA hybrids. Nucleic Acids Res. 50, 1801–1814 (2022).
Article CAS PubMed Google Scholar
Schneider, C., Leung, E., Brown, J. & Tollervey, D. The N-terminal PIN ___domain of the exosome subunit Rrp44 harbors endonuclease activity and tethers Rrp44 to the yeast core exosome. Nucleic Acids Res. 37, 1127–1140 (2009).
Article CAS PubMed PubMed Central Google Scholar
Lukaszewicz, A., Shodhan, A. & Loidl, J. Exo1 and Mre11 execute meiotic DSB end resection in the protist Tetrahymena. DNA Repair 35, 137–143 (2015).
Article CAS PubMed Google Scholar
Balakrishnan, L. & Bambara, R. A. Flap endonuclease 1. Annu. Rev. Biochem. 82, 119–138 (2013).
Article CAS PubMed PubMed Central Google Scholar
Nowotny, M. et al. Specific recognition of RNA/DNA hybrid and enhancement of human RNase H1 activity by HBD. EMBO J. 27, 1172–1181 (2008).
Article CAS PubMed PubMed Central Google Scholar
Cerritelli, S. M. & Crouch, R. J. The non-RNase H ___domain of Saccharomyces cerevisiae RNase H1 binds double-stranded RNA: magnesium modulates the switch between double-stranded RNA binding and RNase H activity. RNA 1, 246–259 (1995).
CAS PubMed PubMed Central Google Scholar
Qin, Z. H. et al. Human RPA activates BLM’s bidirectional DNA unwinding from a nick. Elife 9, 54098 (2020).
Article Google Scholar
Yang, X. W. et al. MutS functions as a clamp loader by positioning MutL on the DNA during mismatch repair. Nat. Commun. 13, 5808 (2022).
Article ADS CAS PubMed PubMed Central Google Scholar
Theile, C. S. et al. Site-specific N-terminal labeling of proteins using sortase-mediated reactions. Nat. Protoc. 8, 1800–1807 (2013).
Article PubMed PubMed Central Google Scholar
Sibenaller, Z. A., Sorensen, B. R. & Wold, M. S. The 32- and 14-kilodalton subunits of replication protein A are responsible for species-specific interactions with single-stranded DNA. Biochemistry 37, 12496–12506 (1998).
Article CAS PubMed Google Scholar
He, Z., Wong, J. M., Maniar, H. S., Brill, S. J. & Ingles, C. J. Assessing the requirements for nucleotide excision repair proteins of Saccharomyces cerevisiae in an in vitro system. J. Biol. Chem. 271, 28243–28249 (1996).
Article CAS PubMed Google Scholar
Modesti, M. Fluorescent labeling of proteins. Methods Mol. Biol. 1665, 115–134 (2018).
Article CAS PubMed Google Scholar
Yang, Y. J. et al. A universal assay for making DNA, RNA, and RNA-DNA hybrid configurations for single-molecule manipulation in two or three steps without ligation. ACS Synth. Biol. 8, 1663–1672 (2019).
Article CAS PubMed Google Scholar
Wang, T. et al. Bloom syndrome helicase compresses single-stranded DNA into phase-separated condensates. Angew. Chem. Int. 61, e202209463 (2022).
Article ADS CAS Google Scholar
Bi, L. et al. The convergence of head-on DNA unwinding forks induces helicase oligomerization and activity transition. Proc. Natl Acad. Sci. USA 119, e2116462119 (2022).
Article CAS PubMed PubMed Central Google Scholar
Ye, S. et al. Proximal single-stranded RNA destabilizes human telomerase RNA G-quadruplex and induces its distinct conformers. J. Phys. Chem. Lett. 12, 3361–3366 (2021).
Article CAS PubMed Google Scholar

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Acknowledgements

We thank all the staff of the molecular and cell biology core facility of the School of Life Science and Technology at ShanghaiTech University for providing technical support. This work was supported by the National Natural Science Foundation of China (32022048 and 32271505 to B.S., 32230029 and 81925015 to W.L.), the Natural Science Foundation of Shanghai (22ZR1441900 and 23ZR1442200 to B.S.), and ShanghaiTech University School of Life Science and Technology Development Fund (to B.S.).

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These authors contributed equally: Yanan Li, Chao Liu.

Authors and Affiliations

School of Life Science and Technology, ShanghaiTech University, Shanghai, China
Yanan Li, Xinshuo Jia, Lulu Bi, Zhiyun Ren, Yilin Zhao, Xia Zhang, Lijuan Guo, Yanling Bao & Bo Sun
Guangzhou Women and Children’s Medical Center, Guangzhou Medical University, Guangzhou, China
Chao Liu & Wei Li
State Key Laboratory of Stem Cell and Reproductive Biology, Institute of Zoology, Chinese Academy of Sciences, Beijing, China
Chao Liu & Wei Li
University of Chinese Academy of Sciences, Beijing, China
Zhiyun Ren & Wei Li
Interdisciplinary Research Center on Biology and Chemistry, Shanghai Institute of Organic Chemistry, Chinese Academy of Sciences, Shanghai, China
Cong Liu

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Contributions

B.S. and W.L. conceived the project and supervised all research. Y.L. and Ch.L. conducted all the experiments. Y.L. Ch.L., W.L., and B.S. analyzed and interpreted the data. X.J., Z.R., Y.Z., X.Z., L.B., Y.B., L.G., and Co.L. helped with the protein purification and hybrid template preparation. B.S. Y.L., Ch.L., and W.L. wrote the manuscript with inputs from all authors.

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Correspondence to Wei Li or Bo Sun.

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Li, Y., Liu, C., Jia, X. et al. RPA transforms RNase H1 to a bidirectional exoribonuclease for processive RNA–DNA hybrid cleavage. Nat Commun 15, 7464 (2024). https://doi.org/10.1038/s41467-024-51984-5

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Received: 04 January 2024
Accepted: 21 August 2024
Published: 29 August 2024
DOI: https://doi.org/10.1038/s41467-024-51984-5