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Transesterification Reaction and the Repair of Embedded Ribonucleotides in DNA are Suppressed upon the Assembly of DNA into Nucleosome Core Particles Mengtian Ren, Yiran Cheng, Qian Duan, and Chuanzheng Zhou Chem. Res. Toxicol., Just Accepted Manuscript • DOI: 10.1021/acs.chemrestox.9b00059 • Publication Date (Web): 16 Apr 2019 Downloaded from http://pubs.acs.org on April 16, 2019
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Transesterification Reaction and the Repair of Embedded Ribonucleotides in DNA are Suppressed upon the Assembly of DNA into Nucleosome Core Particles† Mengtian Ren, Yiran Cheng, Qian Duan and Chuanzheng Zhou*
State Key Laboratory of Elemento-Organic Chemistry and Department of Chemical Biology, College of Chemistry, Nankai University, Tianjin 300071, China * Corresponding author: Chuanzheng Zhou. Email: [email protected]
† Dedicated to 100th anniversary of Nankai University.
Ribonucleotides can be incorporated into DNA through many different cellular processes and abundant amounts of ribonucleotides are detected in genomic DNA. Embedded ribonucleotides lead to
transesterification or by inducing mutagenesis, recombination and chromosome rearrangements. Ribonucleotides misincorporated in genomic DNA can be removed by the ribonucleotide excision repair (RER) pathway in which RNase HII initiates the repair by cleaving the 5′-phosphate of the ribonucleotide. Herein, based on in vitro reconstituted nucleosome core particles (NCPs) containing a single ribonucleotide at different positions, we studied the kinetics of ribonucleotide cleavage via the internal transesterification reaction and repair of the ribonucleotides by RNase HII in NCPs. Our results show that ribonucleotide cleavage via the internal transesterification in NCPs is suppressed compared to that in free DNA. DNA bending and structural rigidity account for the suppressed ribonucleotide cleavage in NCPs. Ribonucleotide repair by RNase HII in NCPs exhibits a strong correlation between the translational and rotational position of the ribonucleotides. An embedded ribonucleotide located at the entry site while facing outward in NCP is repaired as efficiently as that in free DNA. However, the repair of those located in the central part of NCPs and facing inward are inhibited up to 273-fold relative to those in free dsDNA. The difference in repair efficiency appears to arise from their different accessibility to repair enzymes in NCPs. This study reveals that a ribonucleotide misincorporated in DNA assembled into NCPs is protected against cleavage. Hence, spontaneous cleavage of the misincorporated ribonucleotides under physiological conditions is not an essential threat to the stability of chromatin DNA. Instead, their decreased repair efficiency in NCPs may result in numerous and persistent ribonucleotides in genomic DNA, which could exert other deleterious effects on DNA such as mutagenesis and recombination.
INTRODUCTION Abundant amounts of ribonucleotides are embedded in genomic DNA.1-4 Ribonucleotides can be incorporated into DNA through many different cellular processes, including the mis-selection of rNTPs by DNA replicative polymerases,5-7 incomplete removal of Okazaki fragment primers, reverse transcription,8 and DNA repair processes.9,
It is estimated that several million ribonucleotides are
incorporated into the mammalian genome during each round of DNA replication, which is much higher than the abundance of common DNA lesions such as abasic sites and 8-oxo-guanines.5 Ribonucleotides embedded in genomic DNA induce structural alterations,11-14 that can have biological consequences. First, cleavage of the phosphate linkage of ribonucleotides is approximately 105 times faster than that of deoxynucleotides,15-17 thus embedded ribonucleotides significantly increase the genomic instability. Second, ribonucleotides in DNA trigger mutagenesis,18 recombination, chromosome rearrangements and DNA damage response.19 In addition, ribonucleotides in DNA may play a positive role, for instance, acting as strand-discrimination signals in mismatch repair.20, 21 There are efficient repair systems inside cells to counteract the threat of ribonucleotides misincorporation in genomic DNA. The primary pathway is ribonucleotide excision repair (RER) that is initiated by RNase HII.22 RNase HII cleaves the 5’-phosphate of the embedded ribonucleotide, creating a single strand break in which the ribonucleotide is attached to the 5 ′ terminus. Strand displacement synthesis followed by endonuclease processing and ligation restores the doublestranded DNA (dsDNA).23,
Ribonucleotides can also be repaired through topoisomerase I
processing.25, 26 In eukaryotes, nuclear DNA is assembled in nucleosome core particles (NCPs) that are DNAprotein complexes with DNA tightly bound to octameric cores of histone proteins.27,
studies have demonstrated that the formation and repair of DNA in NCPs is different than in free DNA.29-35 Packing DNA in NCPs can protect the DNA from damage.36 However, we have shown that histones can catalyze DNA damage in NCPs through different mechanisms.37-39 In some instances, DNA damage in NCPs leads to DNA-histone cross-links or histone modifications.40-42 DNA repair in NCPs is generally suppressed relative to that in free DNA43-45 and the extent of suppression is highly dependent on the accessibility of DNA lesions to repair enzymes,46, 47 which is, in turn, related to the location of the DNA lesions and the structural dynamics of NCPs.48-51 Hovatter et al. have demonstrated that embedded ribonucleotides distort the structure of DNA and inhibit NCP formation.12 Very recently, the impact of an embedded ribonucleotide on the local NCP structure and dynamics was studied by Broyde based on molecular dynamics simulations.52 Their results showed a large effect of the translational and rotational position of the ribonucleotide on the structural distortions. In the present study, the kinetics of ribonucleotide cleavage via internal transesterification and ribonucleotide repair by RNase HII in NCPs were studied and compared with that in free dsDNAs, with the aim to interrogate how the assembly of DNA into NCPs influence the transesterification reaction and repair of embedded ribonucleotides in genomic DNA.
MATERIALS AND METHODS Materials Native oligodeoxynucleotides were purchased from Shanghai Sangon Biotech Co. Ltd. ssDNAs (90 nt and 145 nt) were prepared by ligating short oligonucleotides (Figure S1), as previously described41, 42. Expression and purification of histones were carried out according to previously reported protocols 53.
Gels were visualized using an Amersham Typhoon Gel and Blot Imaging System with excitation
and emission at 488 and 526 nm, respectively. Preparation of single ribonucleotide modified oligodeoxynucleotides Ribonucleotide modified oligodeoxynucleotides were synthesized on 1 μmol scale on an Applied Biosystems Incorporated 394 oligonucleotide synthesizer using phosphoramidite chemistry, standard DNA synthesis reagents and cycles. 2′-O-TBDMS protected building blocks and extended coupling time (30 min) was employed for ribonucleotides incorporation. After synthesis, the solid supports were treated with a 1 mL mixture of 25% aqueous ammonia/ethanol (v/v, 3/1) at 55°C for 16 h. The supernatant was then separated from the solid supports, and evaporated to dryness. The residue was treated with 0.5 mL of 1 M tetrabutylammonium fluoride in THF at room temperature for 24 h. After quenching with 0.5 mL of distilled water, it was applied to a Sep-Pak C18 column (Waters, WAT020805) according to the manufacturer’s instructions. The obtained crude products were purified by 20% denaturing PAGE (42 × 35 × 0.1 cm), extracted with elution buffer (0.2 M NaCl and 1 mM EDTA), and desalted with Sep-Pak C18 column to give oligos in >99% purity. The integrity of these modified oligonucleotides was characterized by ESI-Q-TOF mass spectrometry. Verification of the ribonucleotides incorporation by NaOH treatment ssDNA substrates (145 nt, 3 pmol) containing a single ribonucleotide at different sites were incubated in 0.3 M NaOH at 55 ºC for 2 h. After neutralization with diluted HCl, the samples were analysed by 8% denaturing PAGE. Reconstitution of NCPs Salmon sperm DNA (1.2 g), dsDNA (145 bp, 25 pmol) and histone octamer (162.5 pmol) were combined in a Slide-A-Lyzer MINI Dialysis Unit (3500 MWCO, Thermo Scientific, Prod. #69550) in a final volume of 50 μL containing 2 M NaCl. The dialysis unit was placed inside another dialysis bag, filled with ~20 mL buffer (2 M NaCl, 10 mM HEPES pH 7.5, 0.1 mM PMSF). This bag was placed into 2 L of low salt buffer (10 mM HEPES, pH 7.5, 0.1 mM PMSF) and dialyzed overnight at 4 ºC. The sample was incubated at 37 ºC for 2 h and any precipitate formed was pelleted by a 10 min spin at 2,000 g. The solution was then transferred to a fresh siliconized tube. NCPs prepared above were used directly in these experiments without adjustments in the concentration following reconstitution. To determine the extent of reconstitution, a small aliquot was removed and analyzed by 5% native
PAGE (10 × 8 × 0.1 cm, acrylamide/bisacrylamide, 59:1, 0.6 × TBE buffer, run at 4 ºC using 0.2 × TBE buffer). The gel was run under limiting voltage (150 V) until the bromophenol blue band migrated to the middle of the gel. All reconstituted NCPs were stored at 4 ºC and used directly in the following studies. Ribonucleotide cleavage in HEPES buffer (pH 7.5) Reconstituted NCP-rA89 (~ 60 mM NaCl, 10 mM HEPES, pH 7.5, 0.1 mM PMSF) was incubated at 37 °C. Aliquots were removed at appropriate times and analyzed by 8% denaturing PAGE. As the control, dsDNA-rA89 was incubated in the same buffer in parallel. Stability of NCP in the borate buffer NCP-rA89 was subjected to buffer exchange using a PD SpinTrapTM G-25 column to provide NCPs in borate buffer (100 mM NaCl, 5 mM MgCl2, 10 mM Borate-NaOH) with different pH values (pH 7.4, 8.0, 9.0, 10.0). The obtained samples were incubated at 37 °C for 4 h followed by EMSA using 5% native PAGE (10 × 8 × 0.1 cm, acrylamide/bisacrylamide, 59:1, 0.6×TBE buffer, run at 4 ºC using 0.2× TBE buffer). Ribonucleotide cleavage in borate buffer (pH 10.0) NCPs in pH 10.0 borate buffer (100 mM NaCl, 5 mM MgCl2, 10 mM borate-NaOH) were obtained through buffer change using PD SpinTrapTM G-25 column. The NCPs were incubated at 37 °C. Aliquots were removed at appropriate times and split into two portions. One was analyzed by 5% native PAGE to examine the extent of NCP disassembly. The other one was analyzed by 8% denaturing PAGE to quantify the strand cleavage in NCP. The percentages of strand cleavage in NCPs were reported without correction for any NCP disassembly. As controls, 145 bp dsDNAs were incubated in the same buffer in parallel. For the reaction with spermine, 1 mM spermine (the final concentration) was added to the buffer directly before incubation. Preparation of 90 nt NC dsDNA The 90 nt 5’-phosphorylated ssDNA (0.1 nmol, 1 eq) and a templating scaffold oligonucleotide (0.3 nmol, 3 eq) were annealed in 0.5 mL of 1× T4 DNA ligase buffer (30 mM Tris-HCl, pH 7.8 at 25 °C, 10 mM MgCl2, 10 mM DTT, and 1 mM ATP) by heating to 95 °C for 10 min followed by cooling to room temperature at a rate of 0.5 °C/min. ATP (0.5 μL, 100 mM) and T4 DNA ligase (1 μL, 40000 U/μL) were added to the mixture followed by incubation at 16 ºC overnight. After ethanol precipitation, the obtained residue was analysed by 8% denaturing PAGE (20 × 16 × 0.1 cm) in 1× TBE. Three major bands were excised and eluted overnight in 0.5 mL of elution buffer (0.2 M NaCl and 1 mM EDTA). After ethanol precipitation, the three products were identified, by DNase I digestion, as 90 nt csDNA, 180 nt ssDNA and 180 nt csDNA respectively (Figure S5). The 90 nt csDNA was annealed with complementary ssDNA to produce NC csDNA. Ribonucleotide cleavage in NC csDNA upon treatment with 0.1 M NaOH
NC csDNA-rA89 (2.5 pmol) was incubated in 0.1 M NaOH at 37 ºC. Aliquots were removed at appropriate times and neutralized with diluted HCl. The samples were analyzed by 8% denaturing PAGE. As the control, dsDNA-rA89 was treated in the same way in parallel. Kinetics of ribonucleotide excision repair by RNase HII NCPs (4 pmol) were treated with RNase HII (3 μL, 15 U) in 1× ThermoPol® Reaction Buffer (20 mM Tris-HCl, 10 mM (NH4)2SO4, 10 mM KCl, 2 mM MgSO4, 0.1% Triton X-100, pH 8.8) at 37 °C. The total reaction volume was 30 µL. Aliquots were removed at appropriate times and quenched by adding 5 μg of proteinase K. All samples were analyzed by 8% denaturing PAGE (20 × 16 × 0.1 cm) in 1× TBE. As controls, dsDNAs were treated the same way in parallel.
RESULTS AND DISCUSSION Preparation of NCPs with a single ribonucleotide at different locations. Given that a single ribonucleotide in DNA imparts obvious an adverse influence on the formation of NCPs,12 the ‘601’ DNA, which has been identified as the one having the highest affinity for histone octamer and formatting well positioned NCPs,54,
was chosen for the preparation of NCPs with a single
ribonucleotide embedded at different positions (Figure 1A). rG73 are located in the dyad axis region (superhelical location [SHL] 0). Both rU86 and rA89 are located near SHL 1.5 where DNA binds tightly to the histone core and closely interacts with the tail of histone H4, but they are in different rotational positions. The sugar-phosphate backbone of rA89 is close to the histone core, and it is designated as “inward” orientation. rU86, whose sugar-phosphate backbone is located far away form the histone core, is referred to as “outward” orientation (Figure 1B). SHL 7 was selected for ribonucleotides incorporation because this region lies at the entry/exit site of the NCPs and has the weakest contact and the most freedom to unwind from the histone core. rG137 and rC141 are oriented outward and inward, respectively, in the region of SHL 7. To this end, chemically synthesized oligonucleotides bearing a single ribonucleotide, together with other native oligo DNA molecules, were ligated to creat 145 nt single-stranded “601” DNA molecules each bearing a 5′-FAM fluorescence label (Figure S1). Incubation of the obtained ssDNAs with 0.3 M NaOH at 55 ℃ for 2 hours resulted in quantitative strand cleavage at the ribonucleotide positions, confirming the correct incorporation of the ribonucleotides (Figure 1C). The ssDNA molecules were annealed with complementary DNA to produce 145 bp dsDNAs, which were employed for NCP reconstitution via the salt dialysis approach in HEPES buffer (pH 7.5). Electrophoretic mobility shift assay (EMSA) showed that stable and homogeneous NCPs were obtained in more than 98% yields regardless of the position of the ribonucleotides incorporation (Figure 1D).
Figure 1. Preparation of NCPs with a single ribonucleotide modification at different positions. (A) Xray crystal structure of NCP showing the locations of the ribonucleotide modifications (PDB: 3LZ0). (B) Positioning of ribonucleotides in DNA on the surface of the histone octamer. The three nucleotides whose sugar-phosphate backbone moieties are close to the histone core in each turn of the helix are referred to as “inward”; The three ones whose sugar-phosphate backbone moieties are far away from the histone core are referred to as “outward”. “Inward” and “outward” locations are marked by gray shadows and yellow shadows respectively. Thus, rA89 and rC141 are located in the “inward” region; rU86 and rG137 are in the “outward” region. (C) An 8% denaturing PAGE analysis of 145 nt ‘601’ ssDNAs with a single ribonucleotide embedded at different positions. For NaOH treatment, the ssDNA molecules (3 pmol) were incubated in 0.3 M NaOH at 55 ℃ for 2 hours. (D) A 5% native PAGE showing the NCPs assembly efficiency. NCPs were obtained in HEPES buffer (60 mM NaCl, 10 mM HEPES, pH 7.5, 0.1 mM PMSF).
Ribonucleotides embedded in NCPs lead to no detectable strand cleavage under physiologically relevant conditions. Ribonucleotides embedded in DNA can lead to spontaneous strand cleavage via an internal transesterification reaction (Figure 2A).16, 17 The cleavage is fulfilled by an “in-line attack”56 of the 2′-hydroxyl group on the vicinal 3 ′ -phosphate to form a pentacoordinate transition state, followed by the departure of the 5′-oxyanion. Specific and general bases can catalyze the reaction by deprotonating the 2′-OH, and acids catalyze the reaction via stabilizing the transition
state and assisting the departure of 5 ′ -oxyanion. Histones are lysine- and arginine-rich proteins. Extensive interactions between Lys and Arg residues with the phosphate linkages were observed in the crystal structures of NCPs. For instance, the 3 ′ -phosphate of A89 is hydrogen bonded with the Arg69 of histone H3 in the “601” NCP.55 We thus speculated that histones may promote the internal transesterification reaction by general acid and base catalysis, making the ribonucleotide cleavage in NCPs faster than in free dsDNAs. To test this hypothesis, the kinetics of spontaneous ribonucleotide cleavage in NCPs were first studied using NCP containing a ribonucleotide at position 89 (NCP-rA89). Incubation of NCP-rA89 in the reconstitution buffer (60 mM NaCl, 10 mM HEPES, pH 7.5, 0.1 mM PMSF) up to 144 hours led to no detectable strand cleavage (Figure 2B). Under the same conditions, 1.5% of strand cleavage was observed for dsDNA-rA89 after incubation for 144 h. These results suggest that, in contrast to our prediction, spontaneous cleavage at ribonucleotides is suppressed in NCP. Misincorporation of ribonucleotides may not be a real threat to the stability of chromatin DNA.
Figure 2. DNA strand cleavage via internal transesterification of ribonucleotide in dsDNA and NCP under physiologically relevant conditions. (A) Mechanism of ribonucleotide cleavage via the internal transesterification reaction. (B) An 8% denaturing PAGE analysis of strand cleavage for dsDNA-rA89 and NCP-rA89 upon incubation in reconstitution buffer (100 mM NaCl, 10 mM HEPES, pH 7.5, 0.1 mM PMSF).
Ribonucleotides cleavage is suppressed in NCPs compared to in free dsDNAs at pH 10.0. The above results show that the assembly of DNA into NCPs protects DNA against cleavage at the incorporated ribonucleotides. The complete stability of ribonucleotides in NCP in neutral buffer makes it impossible to determine to what extent the structure of the NCP suppresses the transesterificationmediated ribonucleotide cleavage. It has been reported that the rate of ribonucleotide cleavage via
internal transesterification increases exponentially with increasing pH.57 We thus intended to compare the kinetics of ribonucleotide cleavage in dsDNA-rA89 and NCP-rA89 at elevated pH. Influence of elevated pH on the stability of NCPs. A prerequisite for studying the kinetics of ribonucleotide cleavage in NCPs at elevated pH is that NCPs should maintain their integrity as much as possible under the conditions. Thus, NCP-rA89 was subjected to buffer exchange, providing NCPs in borate buffer (100 mM NaCl, 5 mM MgCl2, 10 mM borate) with different pH values. Incubation of the obtained NCPs at 37 ℃ for 4 hours clearly showed that the more basic the buffer was, the more the NCPs disassembled (Figure 3A). Incubation of dsDNA-rA89 in borate buffer (pH 10.0) resulted in detectable strand cleavage at rA89 (Figure S2). Meanwhile, approximately 50% of NCP-rA89 remain intact under the same conditions. Therefore, borate buffer (pH 10.0) was chosen for comparing the kinetics of ribonucleotide cleavage in dsDNAs and in NCPs. It should be noted that the partial disassembly of NCPs may lead to an underestimation of the effect of the NCP structure on the rate of ribonucleotide cleavage in NCPs.
Figure 3. Strand cleavage at ribonucleotide modification sites under alkaline conditions. (A) A 5% native PAGE showing the stability of NCP-rA89 after incubation in borate buffer with different pH values for 4 h. (B) Kinetic curves of DNA strand cleavage in NCPs containing a single ribonucleotide at different positions in borate buffer (pH 10.0). (C) Comparison of the strand cleavage of NCPs with that of the dsDNA counterpart after incubation for 66 h in borate buffer (pH 10.0). (D) Comparison of
cleavage kinetics of dsDNA-rA89 in the presence of 1 mM spermine with that of NCP-rA89 and dsDNArA89 in borate buffer (pH 10.0). Borate buffer (pH 10.0): 100 mM NaCl, 5 mM MgCl2, 10 mM borateNaOH, pH 10.0. The percentages of strand cleavage in NCPs were reported without correction for any NCP disassembly.
Kinetics of ribonucleotide cleavage in borate buffer (pH 10.0). We incubated ribonucleotide-embedded NCPs in borate buffer (pH 10.0), and the kinetics of strand cleavage were monitored by denaturing PAGE (Figure S2). The rates of ribonucleotides cleavage were dependent on the location of the ribonucleotide modification, and decreased in the order of NCP-rC141 > NCP-rG137 ≈ NCP-rU86 > NCP-rA89 ≈ NCP-rG73 (Figure 3B). However, ribonucleotides cleavage in free dsDNA showed a similar location-dependent trend (Figure S3). By considering the yields of strand cleavage in NCPs (the percentages of strand cleavage in NCPs were reported without correction for any NCP disassembly) with the counterparts in dsDNA after incubation for 66 h (Figure 3C), we found that ribonucleotide phosphodiester bond cleavage in NCPs was slightly suppressed (less than 2-fold) regardless of modification position. EMSA analysis of the NCP-rA89 revealed that after 66 h incubation in the pH 10.0 buffer, 56% of NCPs disassembled (Figure S4). Taking into account the partial disassembly of NCPs, we can conclude that ribonucleotides cleavage in NCPs should be more significantly suppressed than we observed here. We found that adding 1 mM spermine to the incubation buffer led to a 4-fold increase in the rate of ribonucleotide cleavage for dsDNA-rA89 (Figure 3D), indicating spermine catalyzes the internal transesterification reaction. Given that spermine has a similar pKa value to the -amine of Lys,58 this result supports our previous hypothesis that Lys- and Arg-rich histones can promote ribonucleotide cleavage in NCPs. However, if this hypothesis is true, we should observe a location-dependent increase in the ribonucleotide cleavage rate, keeping in mind that the Lys- and Arg-rich histone tails are not uniformly distributed in the NCP. For instance, position 89 is close to the tail of histone H4 and has been demonstrated to be a reactive site interacting with histone tails.37, 41, 42, 53 NCP-rA89 is thus supposed to be the one showing a high cleavage rate. Unfortunately, the break of rA89 was suppressed the most (approximately 2-fold) upon assembly into NCP. Therefore, there must be some other effects that inhibit ribonucleotide cleavage in NCPs. Inhibition effects outcompeted the catalytic effects, and overall, suppressed cleavage rates for embedded ribonucleotide were observed upon assembly of dsDNA into NCP.
Ribonucleotide phosphodiester cleavage is suppressed by DNA bending. Except for acid and base catalysis, another effect that may dictate the ribonucleotide transesterification reaction is the conformation required for the in-line attack of the 2 ′ -OH to the 3 ′ -phosphate.56,
ribonucleotides in duplexes are more constrained and their cleavage is significantly inhibited compared to that in single-stranded oligos.60,
matters, and a rapid North to South conformational flip followed by internal transesterification via a South-type transition state seems kinetically more favourable for ribonucleotide cleavage.62 Assembly of dsDNA into NCP obviously makes the DNA bent and more rigid, which may account for the decreased cleavage rates for embedded ribonucleotides in NCP. We thus prepared a 90 nt ssDNA-rA89 corresponding to the 31-120 region of the ‘601’ DNA (Figure 4A). The 90 nt ssDNA contains an FAM label tethered to the C5 of T106. After ligation via T4 ligase, a 90 nt cyclical single-stranded DNA (csDNA) was isolated and its structure was verified by DNase I digestion (Figure S5). The csDNA was annealed to complementary DNA to produce a nicked, cyclical dsDNA (NC dsDNA-rA89, Figure 4B)33 that was employed to examine the impact of DNA bending on cleavage of the embedded ribonucleotide. We treated NC dsDNA-rA89 and dsDNA-rA89 with 0.1 M NaOH and the kinetics of strand cleavage were monitored by denaturing PAGE (Figure S6). Cleavage of dsDNA-rA89 was found to be 2 times as fast as the cleavage of NC dsDNA-rA89 (Figure 4C), confirming that DNA bending inhibits strand cleavage at the embedded ribonucleotide. The inhibition effect could be attributed to the fact that DNA bending makes the embedded ribonucleotide more rigid, and thus the conformation alteration required for internal transesterification is more retarded. It is worthy to note that DNA in NCPs is certainly more constrained than in the 90 bp NC dsDNA. Therefore, it is reasonable to presume that rigidity and torsional stress of DNA is the major effector suppressing the ribonucleotide cleavage in NCPs.
Figure 4. Preparating cyclical dsDNA to study the impact of DNA bending on the ribonucleotide cleavage. (A) Strategy for preparing NC dsDNA. csDNA: cyclical ssDNA, NC dsDNA: nicked cyclical dsDNA. (B) A 5% native PAGE analysis of the prepared csDNA and NC dsDNA. The gel was subject
to fluorescence imaging (left panel) followed by YeaGREEN staining (right panel). (C) Kinetic curves of ribonucleotide cleavage in dsDNA and NC dsDNA upon treatment with 0.1 M NaOH.
Repair of ribonucleotide by RNase HII in NCPs. The above results show that assembly of free DNA into NCPs inhibits strand cleavage via ribonucleotide transesterification. This protecting effect improves the stability of genomic DNA. However, if not repaired, the presence of persistent ribonucleotides in DNA may lead to other deleterious consequences such as mutagenesis and recombination.3 In this context, efficient repair is more important for eliminating the negative effects imparted by ribonucleotide misincorporation. Next, we carried out experiments to reveal the impact of NCP structure on the repair of an embedded ribonucleotide by RNase HII. RNase HII initiates the repair of an embedded ribonucleotide via scission of the 5′-phosphate, which allows us to monitor the kinetics of RNase HII repair by analyzing the strand cleavage using denaturing PAGE (Figure S7). dsDNAs with a single ribonucleotide at different positions were treated with RNase HII. All of the substrates were repaired by RNase HII with a similar efficiency (Figure 5A & Table 1). In contrast, the rates of ribonucleotide repair in NCPs varied up to 552-fold with their locations (Figure 5B & Table 1). Repair of rG137 in NCP was slightly faster, at least not less efficient, than in dsDNA. However, repair rates for ribonucleotides at other positions decreased 29- to 273-fold in NCP relative to in dsDNA (Table 1). Ribonucleotide repair in NCPs by RNase HII exhibited a strong correlation between the translational and rotational position of the ribonucleotide. For instance, RNase HII showed higher reactivity for ribonucleotides at SHL 7 (NCP-rC141 and NCP-rG137) than that at the dyad position (NCP-rG73), which, in turn, is a better substrate than ribonucleotides at SHL 1.5 (NCPrA89 and NCP-rU86). At the same translational location, repairing the ribonucleotides facing outward (NCP-rG137 and NCP-rU86) was more efficient than repair of the counterparts facing inward (NCP-rC141 and NCP-rA89).
Figure 5. Kinetics of repair of incorporated ribonucleotides by RNase HII in dsDNAs (A) and NCPs (B).
EMSA showed that all NCPs maintained their integrity after RNase II-mediated strand cleavage (Figure S8). Therefore, the different repair efficiency for ribonucleotides at various positions must stem from their diverse microenvironments in NCPs. Translational and rotational position-dependent repair has been extensively observed for DNA repair by base excision repair enzymes in NCPs, and this is attributed to the fact that lesions at different locations have different accessibility to repair enzymes.31,
Ribonucleotides at SHL 7 (NCP-rG137 and NCP-rC141) have the most freedom for
transient unwrapping from the histone core,47 and thus are the most easily repaired ones. DNA at the dyad axis is generally less accessible to repair enzymes relative to other locations,64, 65 which explains the significantly decreased repair efficiency for NCP-rG73. Previous studies have shown that in NCP, DNA at the SHL 1.5 closely interacts with the tail of histone H4.38, 40, 53, 66, 67 The binding of the H4 tail to this region may hamper the raction of RNase HII, which leads to the extremely low repair efficiency for NCP- rU86 and NCP-rA89, which are located at the SHL 1.5. In addtion, the rigid structure of DNA in this region may also account for the reduced repair efficiency. Recently, Broyde et al., based on molecular dynamics simulations, demonstrated that inward facing ribonucleotides and outward facing ones have different ribose conformations.52 Generally, the ribonucleotides facing outward, similar to a deoxyribonucleotide, adopts a C2′-endo conformation, and, as a result, adjacent deoxynucleotides and the global structure of DNA are not significantly affected. The authors speculated that the abnormal C2′-endo conformation of an embedded ribonucloetide may allow it to escape from the surveillance of RNase HII. In contrast, a ribonucleotide facing inward maintains a more stable C3′-endo conformation through interactions with histones, which causes local structure disturbance including ruptured Watson-Crick pairing and duplex unwinding. Thus, inward-facing ribonucleotides are likely to be recognized by RNase HII. However, their lesser accessibility inhibits the reaction of RNase HII. Our present results corroborate Broyde’s hypothesis that repair of embedded ribonucleotides in DNA is inhibited by chromatin. The observation that ribonucleotides facing outward are more readily repaired than those facing inward suggests that the accessiblility of a ribonucleotide is the major effector determining how fast it can be repaired by RNase HII in NCPs.
CONCLUSION Based on in vitro reconstituted NCPs containing a single ribonucleotide at different positions, we studied the kinetics of transesterification-mediated ribonucleotide cleavage and repair of the ribonucleotides by RNase HII in NCPs. Our results showed that under physiologically relevant conditions, ribonucleotides in NCPs do not lead to obvious strand cleavage via spontaneous transesterification. Increasing the pH to 10.0 allowed us to compare the rates of strand cleavage at ribonucleotides in free DNA and in NCPs. We found the ribonucleotide phosphodiester bond cleavage in NCPs was slightly suppressed (less than 2-fold) compared to that in free DNA. We demonstrated that ribonucleotide cleavage in a cyclical dsDNA was 2 times slower than in its linear dsDNA counterpart, and thus this supports the hypothesis that the bending and rigid structure of nucleosomal DNA accounts for the suppressed ribonucleotide cleavage in NCPs. Unlike free DNA in which embedded ribonucleotides can be repaired by RNase HII with high efficiency regardless of their locations, a 552-fold difference in repair rate was observed for ribonucleotides located at different positions of NCPs. The difference appears to arise from their different accessibility to RNase HII in NCPs. G137 that is located at the entry site while facing outward in NCP is repaired as efficiently as that in free DNA. However, repair of ribonucleotides located in the central part of NCPs and facing inward is significantly suppressed. This study reveals that ribonucleotides misincorporated in nucleosomal DNA are protected against cleavage via internal transesterification. Hence, spontaneous ribonucleotide cleavage under physiological conditions is not a significant threat to the stability of chromatin DNA. Instead, their decreased repair efficiency in NCPs may result in numerous and persistent ribonucleotides in genomic DNA, which could exert other deleterious effects on DNA such as mutagenesis and recombination. These adverse impacts remain to be further addressed in chromatin in the future.
ASSOCIATED CONTENT Supporting Information Supporting Information is available free of charge via the Internet at http://pubs.acs.org. Supplementary figures (PDF)
AUTHOR INFORMATION Corresponding Author *Chuanzheng Zhou. E-mail: [email protected] Funding
This work was supported by the National Natural Science Foundation of China (21572109, 21877064) and the Fundamental Research Funds for the Central Universities, Nankai University (63191523). Notes The authors declare no competing financial interest.
Schroeder, J. W., Randall, J. R., Matthews, L. A., and Simmons, L. A. (2015) Ribonucleotides in bacterial DNA, Crit. Rev. Biochem. Mol. Biol. 50, 181-193. Caldecott, K. W. (2014) Ribose-An internal threat to DNA, Science 343, 260-261. Klein, H. L. (2017) Genome instabilities arising from ribonucleotides in DNA, DNA Repair 56, 26-32. Koh, K. D., Balachander, S., Hesselberth, J. R., and Storici, F. (2015) Ribose-seq: global mapping of ribonucleotides embedded in genomic DNA, Nat. Methods 12, 251-+. McElhinny, S. A. N., Watts, B. E., Kumar, D., Watt, D. L., Lundstrom, E. B., Burgers, P. M. J., Johansson, E., Chabes, A., and Kunkel, T. A. (2010) Abundant ribonucleotide incorporation into DNA by yeast replicative polymerases, Proc. Natl. Acad. Sci. U. S. A. 107, 4949-4954. Brown, J. A., and Suo, Z. C. (2011) Unlocking the sugar "steric gate" of DNA polymerases, Biochemistry 50, 1135-1142. Wanrooij, P. H., Engqvist, M. K. M., Forslund, J. M. E., Navarrete, C., Nilsson, A. K., Sedman, J., Wanrooij, S., Clausen, A. R., and Chabes, A. (2017) Ribonucleotides incorporated by the yeast mitochondrial DNA polymerase are not repaired, Proc. Natl. Acad. Sci. U. S. A. 114, 1246612471. Kennedy, E. M., Amie, S. M., Bambara, R. A., and Kim, B. (2012) Frequent incorporation of ribonucleotides during HIV-1 reverse transcription and their attenuated repair in macrophages, J. Biol. Chem. 287, 14280-14288. Pryor, J. M., Conlin, M. P., Carvajal-Garcia, J., Luedeman, M. E., Luthman, A. J., Small, G. W., and Ramsden, D. A. (2018) Ribonucleotide incorporation enables repair of chromosome breaks by nonhomologous end joining, Science 361, 1126-1129. Zhu, H., and Shuman, S. (2008) Bacterial nonhomologous end joining ligases preferentially seal breaks with a 3′-OH monoribonucleotide, J. Biol. Chem. 283, 8331-8339. Meroni, A., Mentegari, E., Crespan, E., Muzi-Falconi, M., Lazzaro, F., and Podesta, A. (2017) The incorporation of ribonucleotides induces structural and conformational changes in DNA, Biophys. J. 113, 1373-1382. Hovatter, K. R., and Martinson, H. G. (1987) Ribonucleotide-induced helical alteration in DNA prevents nucleosome formation, Proc. Natl. Acad. Sci. USA 84, 1162-1166. Egli, M., Usman, N., and Rich, A. (1993) Conformational influence of the ribose 2'-hydroxyl group: Crystal structures of DNA-RNA chimeric duplexes, Biochemistry 32, 3221-3237.
Evich, M., Spring-Connell, A. M., Storici, F., and Germann, M. W. (2016) Structural impact of single ribonucleotide residues in DNA, Chembiochem 17, 1968-1977. Jenkins, L. A., Bashkin, J. K., and Autry, M. E. (1996) The embedded ribonucleotide assay: A chimeric substrate for studying cleavage of RNA by transesterification, J. Am. Chem. Soc. 118, 6822-6825. Li, Y., and Breaker, R. R. (1999) Kinetics of RNA degradation by specific base catalysis of transesterification involving the 2‘-hydroxyl group, J. Am. Chem. Soc. 121, 5364-5372. Oivanen, M., Kuusela, S., and Lonnberg, H. (1998) Kinetics and mechanisms for the cleavage and isomerization of the phosphodiester bonds of RNA by Bronsted acids and bases, Chem. Rev. 98, 961-990. Shen, Y., Koh, K. D., Weiss, B., and Storici, F. (2012) Mispaired rNMPs in DNA are mutagenic and are targets of mismatch repair and RNases H, Nat. Struct. Mol. Biol. 19, 98-U123. Williams, J. S., and Kunkel, T. A. (2014) Ribonucleotides in DNA: Origins, repair and consequences, DNA Repair 19, 27-37. Ghodgaonkar, Medini M., Lazzaro, F., Olivera-Pimentel, M., Artola-Borán, M., Cejka, P., Reijns, Martin A., Jackson, Andrew P., Plevani, P., Muzi-Falconi, M., and Jiricny, J. (2013) Ribonucleotides misincorporated into DNA act as strand-discrimination signals in eukaryotic mismatch repair, Mol. Cell 50, 323-332. Lujan, Scott A., Williams, Jessica S., Clausen, Anders R., Clark, Alan B., and Kunkel, Thomas A. (2013) Ribonucleotides are signals for mismatch repair of leading-strand replication errors, Mol. Cell 50, 437-443. Sparks, Justin L., Chon, H., Cerritelli, Susana M., Kunkel, Thomas A., Johansson, E., Crouch, Robert J., and Burgers, Peter M. (2012) RNase H2-initiated ribonucleotide excision repair, Mol. Cell 47, 980-986. Rydberg, B., and Game, J. (2002) Excision of misincorporated ribonucleotides in DNA by RNase H (type 2) and FEN-1 in cell-free extracts, Proc. Natl. Acad. Sci. U. S. A. 99, 1665416659. Williams, J. S., Lujan, S. A., and Kunkel, T. A. (2016) Processing ribonucleotides incorporated during eukaryotic DNA replication, Nat. Rev. Mol. Cell Biol. 17, 350-363. Kim, N., Huang, S.-y. N., Williams, J. S., Li, Y. C., Clark, A. B., Cho, J.-E., Kunkel, T. A., Pommier, Y., and Jinks-Robertson, S. (2011) Mutagenic processing of ribonucleotides in DNA by yeast topoisomerase I, Science 332, 1561-1564. Williams, Jessica S., Smith, Dana J., Marjavaara, L., Lujan, Scott A., Chabes, A., and Kunkel, Thomas A. (2013) Topoisomerase 1-mediated removal of ribonucleotides from nascent leading-strand DNA, Mol. Cell 49, 1010-1015. McGinty, R. K., and Tan, S. (2015) Nucleosome structure and function, Chem. Rev. 115, 22552273. Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution, Nature 389, 251-260. Mengtian, R., Jing, B., Zhen, X., and Chuanzheng, Z. (2019) DNA damage in nucleosomes, SCIENCE CHINA Chemistry, DOI: 10.1007/s11426-11018-19421-11425. Hauer, M. H., and Gasser, S. M. (2017) Chromatin and nucleosome dynamics in DNA damage and repair, Genes Dev. 31, 2204-2221. Rodriguez, Y., Hinz, J. M., and Smerdon, M. J. (2015) Accessing DNA damage in chromatin: Preparing the chromatin landscape for base excision repair, DNA Repair 32, 113-119. Taylor, J. S. (2015) Design, synthesis, and characterization of nucleosomes containing sitespecific DNA damage, DNA Repair 36, 59-67. Wang, K., and Taylor, J. S. (2017) Modulation of cyclobutane thymine photodimer formation in T11-tracts in rotationally phased nucleosome core particles and DNA minicircles, Nucleic Acids Res. 45, 7031-7041.
Banerjee, D. R., Deckard, C. E., Elinski, M. B., Buzbee, M. L., Wang, W. W., Batteas, J. D., and Sczepanski, J. T. (2018) Plug-and-play approach for preparing chromatin containing sitespecific DNA modifications: The influence of chromatin structure on base excision repair, J. Am. Chem. Soc. 140, 8260-8267. Zou, T., Kizaki, S., and Sugiyama, H. (2018) Investigating nucleosome accessibility for MNase, Fe(II)peplomycin, and duocarmycinB(2) by using capillary electrophoresis, Chembiochem 19, 664-668. Ljungman, M., and Hanawalt, P. C. (1992) Efficient protection against oxidative DNA damage in chromatin, Mol. Carcinog. 5, 264-269. Zhou, C. Z., and Greenberg, M. M. (2012) Histone-catalyzed cleavage of nucleosomal DNA containing 2-deoxyribonolactone, J. Am. Chem. Soc. 134, 8090-8093. Zhou, C. Z., Sczepanski, J. T., and Greenberg, M. M. (2012) Mechanistic studies on histone catalyzed cleavage of apyrimidinic/apurinic sites in nucleosome core particles, J. Am. Chem. Soc. 134, 16734-16741. Zhou, C. Z., and Greenberg, M. M. (2014) DNA damage by histone radicals in nucleosome core particles, J. Am. Chem. Soc. 136, 6562-6565. Zhou, C. Z., Sczepanski, J. T., and Greenberg, M. M. (2013) Histone modification via rapid cleavage of C4 '-oxidized abasic sites in nucleosome core particles, J. Am. Chem. Soc. 135, 5274-5277. Li, F., Zhang, Y., Bai, J., Greenberg, M. M., Xi, Z., and Zhou, C. (2017) 5-Formylcytosine yields DNA–protein cross-links in nucleosome core particles, J. Am. Chem. Soc. 139, 10617-10620. Bai, J., Zhang, Y., Xi, Z., Greenberg, M. M., and Zhou, C. (2018) Oxidation of 8-Oxo-7,8dihydro-2′-deoxyguanosine Leads to Substantial DNA-Histone Cross-Links within Nucleosome Core Particles, Chem. Res. Toxicol. 31, 1364-1372. Nilsen, H., Lindahl, T., and Verreault, A. (2002) DNA base excision repair of uracil residues in reconstituted nucleosome core particles, EMBO J. 21, 5943-5952. Beard, B. C., Wilson, S. H., and Smerdon, M. J. (2003) Suppressed catalytic activity of base excision repair enzymes on rotationally positioned uracil in nucleosomes, Proc. Natl. Acad. Sci. U. S. A. 100, 7465-7470. Cannan, W. J., Tsang, B. P., Wallace, S. S., and Pederson, D. S. (2014) Nucleosomes suppress the formation of double-strand DNA breaks during attempted base excision repair of clustered oxidative damages, J. Biol. Chem. 289, 19881-19893. Olmon, E. D., and Delaney, S. (2017) Differential ability of five DNA glycosylases to recognize and repair damage on nucleosomal DNA, ACS Chem. Biol. 12, 692-701. Bilotti, K., Tarantino, M. E., and Delaney, S. (2018) Human oxoguanine glycosylase 1 removes solution accessible 8-oxo-7,8-dihydroguanine lesions from globally substituted nucleosomes except in the dyad region, Biochemistry 57, 1436-1439. Menoni, H., Gasparutto, D., Hamiche, A., Cadet, J., Dimitrov, S., Bouvet, P., and Angelov, D. (2007) ATP-dependent chromatin remodeling is required for base excision repair in conventional but not in variant H2A.Bbd nucleosomesv, Mol. Cell. Biol. 27, 5949-5956. Hinz, J. M., Rodriguez, Y., and Smerdon, M. J. (2010) Rotational dynamics of DNA on the nucleosome surface markedly impact accessibility to a DNA repair enzyme, Proc. Natl. Acad. Sci. U. S. A. 107, 4646-4651. Duan, M.-R., and Smerdon, M. J. (2014) Histone H3 lysine 14 (H3K14) acetylation facilitates DNA repair in a positioned nucleosome by stabilizing the binding of the chromatin remodeler RSC (remodels structure of chromatin), J. Biol. Chem. 289, 8353-8363. Rodriguez, Y., Duan, M., Wyrick, J. J., and Smerdon, M. J. (2018) A cassette of basic amino acids in histone H2B regulates nucleosome dynamics and access to DNA damage, J. Biol. Chem.
Fu, I., Smith, D. J., and Broyde, S. (2019) Rotational and translational positions determine the structural and dynamic impact of a single ribonucleotide incorporated in the nucleosome, DNA repair 73, 155-163. Sczepanski, J. T., Zhou, C. Z., and Greenberg, M. M. (2013) Nucleosome core particlecatalyzed strand scission at abasic sites, Biochemistry 52, 2157-2164. Lowary, P. T., and Widom, J. (1998) New DNA sequence rules for high affinity binding to histone octamer and sequence-directed nucleosome positioning, J. Mol. Biol. 276, 19-42. Vasudevan, D., Chua, E. Y. D., and Davey, C. A. (2010) Crystal structures of nucleosome core particles containing the '601' strong positioning sequence, J. Mol. Biol. 403, 1-10. Soukup, G. A., and Breaker, R. R. (1999) Relationship between internucleotide linkage geometry and the stability of RNA, RNA 5, 1308-1325. Li, Y. F., and Breaker, R. R. (1999) Kinetics of RNA degradation by specific base catalysis of transesterification involving the 2'-hydroxyl group, J. Am. Chem. Soc. 121, 5364-5372. Bergeron, R. J., McManis, J. S., Weimar, W. R., Schreier, K., Gao, F., Wu, Q., Ortiz-Ocasio, J., Luchetta, G. R., Porter, C., and Vinson, J. R. T. (1995) The role of charge in polyamine analog recognition, J. Med. Chem. 38, 2278-2285. Emilsson, G. M., Nakamura, S., Roth, A., and Breaker, R. R. (2003) Ribozyme speed limits, RNA 9, 907-918. Zagorowska, I., Mikkola, S., and Lonnberg, H. (1999) Hydrolysis of phosphodiester bonds within RNA hairpin loops in buffer solutions: the effect of secondary structure on the inherent reactivity of RNA phosphodiester bonds, Helv Chim Acta 82, 2105-2111. Mikkola, S., Kaukinen, U., and Lonnberg, H. (2001) The effect of secondary structure on cleavage of the phosphodiester bonds of RNA, Cell Biochem. Biophys. 34, 95-119. Guo, F., Yue, Z., Trajkovski, M., Zhou, X., Cao, D., Li, Q., Wang, B., Wen, X., Plavec, J., Peng, Q., Xi, Z., and Zhou, C. (2018) Effect of ribose conformation on RNA cleavage via internal transesterification, J. Am. Chem. Soc. 140, 11893-11897. Peterson, C. L., and Almouzni, G. (2013) Nucleosome dynamics as modular systems that integrate DNA damage and repair, Cold Spring Harb. Perspect. Biol. 5. Rodriguez, Y., and Smerdon, M. J. (2013) The structural location of DNA lesions in nucleosome core particles determines accessibility by base excision repair enzymes, J. Biol. Chem. 288, 13863-13875. Bilotti, K., Kennedy, E. E., Li, C., and Delaney, S. (2017) Human OGG1 activity in nucleosomes is facilitated, by transient unwrapping of DNA and is influenced by the local histone environment, DNA Repair 59, 1-8. Kuduvalli, P. N., Townsend, C. A., and Tullius, T. D. (1995) Cleavage by calicheamicin gamma 1I of DNA in a nucleosome formed on the 5S RNA gene of Xenopus borealis, Biochemistry 34, 3899-3906. Weng, L. W., Zhou, C. Z., and Greenberg, M. M. (2015) Probing Interactions between Lysine Residues in Histone Tails and Nucleosomal DNA via Product and Kinetic Analysis, ACS Chem. Biol. 10, 622-630.