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Oct 31, 2018 - Department of Pharmacology, School of Pharmacy, China Medical University, Shenyang 110122, Liaoning, China. ∥. Key Laboratory of ...
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Supramolecular Coordination-Directed Reversible Regulation of Protein Activities at Epigenetic DNA Marks Shao-Ru Wang,†,‡ Jia-Qi Wang,†,‡ Bo-Shi Fu,†,§,‡ Kun Chen,† Wei Xiong,† Lai Wei,† Guangyan Qing,∥ Tian Tian,*,† and Xiang Zhou*,†

J. Am. Chem. Soc. 2018.140:15842-15849. Downloaded from pubs.acs.org by UNIV OF RHODE ISLAND on 12/01/18. For personal use only.

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College of Chemistry and Molecular Sciences, Key Laboratory of Biomedical Polymers of Ministry of Education, Wuhan University, Wuhan 430072, Hubei, China § Department of Pharmacology, School of Pharmacy, China Medical University, Shenyang 110122, Liaoning, China ∥ Key Laboratory of Separation Science for Analytical Chemistry, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian 116023, Liaoning, China S Supporting Information *

ABSTRACT: In mammals, 5-formylcytosine (5fC) has been identified as an important mark, which plays significant roles in active DNA demethylation and also in epigenetic regulation. It is therefore important to target this epigenetic mark as well as manipulating DNA−protein interactions at this site. A unique feature of 5fC is the presence of a formyl group at the C-5 position. In the current study, we introduce supramolecular coordination chemistry for reversible regulation of DNA− protein interactions on this mark. We have designed and synthesized the 2-(aminooxy)-N-(quinolin-8-yl)acetamide (AQA), which functions well in selective labeling of 5fC mark. Using this feature, the association and disassociation of metal ion supplementation allow blocking and deblocking of DNA−protein interactions. In addition, we synthesized a close analogue of AQA by replacing the nitrogen atom in the quinoline ring with a CH group. Importantly, the regulatory effects of those metal ion supplementations were completely erased. On the basis of the combined information, we propose a conformational flexibility in a side arm in response to switched coordination. In the absence of coordinating interaction, the flexible side arm probably takes on an extended conformation and points away from the hydrogen bonding cavity. Importantly, coordinating interaction is effective in imposing a restrained geometry to this side arm, with the quinoline ring being oriented opposite the complementary nucleobase. Moreover, the coordination-induced activity control can be reversed by supplementation with a number of chelating agents. The concept described is unique in installing an auxiliary side arm with bending flexibility to control oligonucleotide functions. Finally, these findings show promising potential of supramolecular coordination chemistry for DNA epigenetics.

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INTRODUCTION

Coordination and organometallic chemistry contributed much to the development of synthetic and supramolecular science.21 It therefore has attracted considerable attention in recent years.22 In this research area, the coordinated donors containing typical heteroatom (oxygen or nitrogen) are sufficiently widespread and maintain leadership.23 Although monodentate coordination is generally weaker than a covalent C−C bond, the binding constant can be greatly increased by multidentate coordination.24 It has been shown that there is considerable cooperativity in complexes containing multiple donors and/or acceptors.25 Recently, some significant achievements have been made in protein research using coordination chemistry,26,27 but few have explored nucleotide−metal interactions.28 Since the directional nature of metal-coordinat-

Cytosine can be methylated at the C-5 position to produce 5methylcytosine (5mC in Figure S1), which represents an epigenetic modification.1 The recently identified DNA demethylation pathway involves the stepwise oxidation of 5mC into 5-hydroxymethylcytosine (5hmC),2 5-formylcytosine (5fC),3 and 5-carboxylcytosine (5caC).4 Although 5fC and 5caC can be excised from DNA by the mammalian thymine DNA glycosylase,5−7 they represent important epigenetic marks in mammals, with several known cellular functions.8−11 Among these oxidized forms of cytosine, 5fC is of particular interest because of the presence of the 5-formyl group.12−15 This group is potentially reactive toward cellular nucleophiles such as proteins.16,17 It is therefore important to target 5fC as well as manipulating its functions through chemical interventions.18−20 © 2018 American Chemical Society

Received: August 25, 2018 Published: October 31, 2018 15842

DOI: 10.1021/jacs.8b09113 J. Am. Chem. Soc. 2018, 140, 15842−15849

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Journal of the American Chemical Society

us to explore a related system.32 Having noted the reactive nature of the 5-formyl group, we consider the introduction of an auxiliary side arm at the C-5 position. It is noteworthy that N-(quinolin-8-yl)acetamide (QA), possessing multiple donor atoms, is a suitable candidate to be appended.33 In 5fC nucleobase, the exocyclic C4-NH2 group can also act as a ligating site. We therefore designed the target molecule AQA. In addition to bearing multifunctional ligating sites for supramolecular coordination, AQA comprises the terminal hydroxylamine group for targeting 5-formyl group.11 We anticipate that nucleobase-specific and side arm-directed coordinating interaction could work in concert. Side Arm Installation at Nucleoside and DNA Levels. The synthetic route of AQA was indicated in the Supporting Information (Chemical Synthesis section). The commercially available 8-aminoquinoline was reacted with 2-bromoacetyl bromide, followed by a mild substitution, to give phthalimideN-oxyl derivative, which was then subject to hydrazinolysis, giving the target molecule.33,34 All synthesized compounds were characterized on the basis of spectral analyses, including nuclear magnetic resonance spectroscopy (NMR) and highresolution mass spectrometry (HRMS). We next examined whether AQA can be used to label 5fC nucleoside. As expected, this can be done under moderately acidic conditions with the presence of a nucleophilic catalyst panisidine (5fC-AQA nucleoside in Figure 2A). We further

ing bonds provide potential driving force for structural control of ligands,29 we are interested to investigate the use of supramolecular coordination for nucleotide-specific recognition and functional regulation. In this Article, we present coordination-directed reversible regulation of DNA−protein interactions from viewpoints of supramolecular chemistry (Figure 1). We designed and

Figure 1. Schematic illustration of the design and workflow. (A) Sitespecific labeling of 5fC mark in DNA with AQA. (B) Supramolecular coordination-directed reversible regulation of base pairing and protein−DNA interactions.

synthesized the 2-(aminooxy)-N-(quinolin-8-yl)acetamide (AQA in Figures 1A and S1), which was used to label 5fC in a site-specific manner (Figure 1A). On the basis of the combined information, the flexible side arm in 5fC-AQA nucleotide probably takes on an extended conformation and points away from the hydrogen bonding cavity in the absence of coordinating interaction (left side in Figure 1B). Importantly, coordinating interaction is effective in imposing a restrained geometry to the side arm, with the quinoline ring being oriented opposite the complementary nucleobase (right side in Figure 1B). As a result, base pairing was destabilized by steric hindrance (left-to-right direction in Figure 1B). In contrast, the chelating agents (CA) have been shown to reverse the nucleotide/metal coordination (right-to-left direction in Figure 1B). Using this feature, the association and disassociation of specific metal ions have been applied for reversible regulation of 5fC-targeted base-pairing and DNA− protein interactions (Figure 1B). The current study has general implications for the application of side arm-controlled supramolecular coordination in DNA epigenetics.

Figure 2. Side arm installation at nucleoside and DNA levels. (A) The AQA labeling of 5fC nucleoside. (B) Denaturing electrophoresis of DNA strands before and after AQA treatment. Reaction conditions: 40 mM sodium acetate buffer (pH = 5.0), 10 mM p-anisidine, 0.5 mM AQA. The DNA bands were stained with GelRed. Lanes 1, 3, 5, 7, 9, 11: untreated DNA; lanes 2, 4, 6, 8, 10, 12: AQA-treated DNA.

tested whether AQA is able to label 5fC nucleotide in DNA context. This reaction was done by incubation of AQA with the 5fC-containing DNA (fC-15mer in Table S1) (Figure 2A).34,35 The DNA products were recovered by ethanol precipitation and then analyzed by denaturing electrophoresis. During this process, different size pieces of DNA are routinely separated according to their molecular weights. On the basis of the results, the AQA-reacted fC-15mer migrates more slowly in polyacrylamide gel due to an increased molecular weight (lane 8 in Figure 2B). This result has also demonstrated quantitative yields of AQA labeling. Moreover, the MALDI-TOF assay

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RESULTS Design of Side Arm Strategy with Supramolecular Coordination. The present study aims to target 5fC marks as well as manipulating DNA−protein interactions. We envisioned that molecular recognition properties of nucleobase with metal ions can be exploited.30,31 However, it is difficult to achieve tight binding of metal ion to 5fC mark by itself. The structural versatility of metal-coordinating side arm motivated 15843

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Journal of the American Chemical Society indicated that the AQA-reacted DNA does contain the desired 5fC-AQA nucleotide (Figure S2). In mammalian DNA genome, there are various types of cytosine modifications and DNA lesions, especially the abasic (AP) site.36,37 In a following study, we systematically examined the reaction selectivity of AQA to various DNAs (C-15mer, mC-15mer, hmC-15mer, caC-15mer, or AP-15mer in Table S1), in which 5fC was substituted by cytosine, 5mC, 5hmC, 5caC, or AP site.11 After a long duration of AQA treatment, the DNA products were recovered and analyzed. Importantly, the migrations of these DNA strands did not slow down as compared to the no-treatment controls (lanes 1−6 and lanes 9−12, Figure 2B). These results revealed that AQA shows good selectivity to 5fC nucleotide in DNA. Side Arm Strategy for Reversible Control of DNA Synthesis. Having demonstrated AQA can target 5fC nucleotides in DNA, we set out to manipulate oligonucleotide functions in biological context. Since DNA synthesis is fundamental to the propagation of all living organisms, chemical reversible control of this process is of great importance.38−40 We therefore designed various DNA templates (C-27mer, mC-27mer, hmC-27mer, fC-27mer, and caC-27mer in Table S1) with the same sequence except the indicated site (cytosine, 5mC, 5hmC, 5fC, or 5caC). The extension primer (p1 in Table S1) was designed such that its 3′ end lies two bases upstream to the target residue in template. The Bacillus stearothermophilus DNA polymerase (Bst DNA pol) was examined for the proof of concept study.41,42 It is important to realize that some metal ions are toxic to enzyme activity. To test this, Bst DNA pol was allowed to elongate along various untreated DNA templates in the presence of metal ions supplementation. On the basis of our results (Figure S3), nonspecific inhibitions were consistently found with some metal ions. They were therefore excluded from the following studies. The first issue we studied is whether coordinating interaction at 5fC-AQA site can arrest DNA synthesis (leftto-right direction in Figure 3A). Hence, each AQA-treated template was used to guide DNA synthesis by Bst DNA pol. In the absence of metal ions supplementation, Bst DNA pol efficiently elongated along fC-27mer, and the full extension was reached (lane 3 in Figure 3B). Strikingly, DNA synthesis was impeded at 5fC-AQA (+2 site) by the treatment with some metal ions [Co2+, Ni2+, and Cu2+; lanes 8−10 in Figures 3B]. In contrast, these metal ions did not influence the primer extension along other AQA-treated templates (Figure S4). On the basis of these results, we identified three metal ions, including Co2+, Ni2+, and Cu2+, as being effective in coordination with 5fC-AQA nucleotide. Additionally, single nucleotide incorporation assay suggests that the AQA labeling does not change the base-pairing properties of 5fC toward guanosine (Figure S5). In view of these results, we proceeded to explicitly investigate the effects of those identified metal ions using a couple of DNA primers (p1, p2, and p3 in Table S1). The representative data are shown in Figures 3C and S6. We observed that these three metal ions lead to specific stalling of DNA pol at 5fC-AQA site with a concentration-dependent manner. In particular, the treatment with 12.5 μM Cu2+ evidently inhibited DNA synthesis along fC-27mer, and higher levels of impedance were observed with the 25 and 50 μM treatments. Furthermore, the 100 μM Cu2+ treatment almost completely aborted DNA pol elongation. On the basis of these

Figure 3. Reversible control of DNA synthesis. (A) Schematic illustration of the workflow. (B) The influence of metal ions supplementation on DNA synthesis. Lane 1: DNA size markers (p1, m1, m2, m3, m4, m5, and m6); lane 2: no enzyme control; lane 3: control without metal ion supplementation; lanes 4−16: metal ion supplementation (each at 100 μM). (C) Concentration-dependent influence of coordinating interaction on DNA synthesis. Lane 1: DNA size markers; lane 2: no enzyme control; lane 3: control without metal ion supplementation; lanes 4−8: 12.5, 25, 50, 200, and 800 μM Co2+; lanes 9−13: 50, 100, 200, 400, and 800 μM Ni2+; lanes 14−19: 6.25, 12.5, 25, 50, 100, and 200 μM Cu2+. (D) The influence of chelating on DNA synthesis. Lane 1: DNA size markers; lane 2: no enzyme control; lanes 3, 11: control without metal ion supplementation; lanes 4, 12: 100 μM Cu2+; lanes 5, 13: 100 μM Cu2+, 200 μM DCyTA; lanes 6, 14: 100 μM Cu2+, 200 μM DTPA; lanes 7, 15: 100 μM Cu2+, 200 μM EDTA; lanes 8, 16: 100 μM Cu2+, 200 μM EGTA; lanes 9, 17: 100 μM Cu2+, 200 μM HEDTA; lanes 10, 18: 100 μM Cu2+, 200 μM NTA. For (B−D), reactions were carried out as described in the Experimental Section. The extension products were separated with denaturing electrophoresis.

results, we ultimately identified Cu2+ and Ni2+ as being optimal for coordination with 5fC-AQA. The second issue we studied is whether the site-specific arrest of DNA synthesis by coordinating interaction can be reversed (right-to-left direction in Figure 3A). It is known that 15844

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Journal of the American Chemical Society some CA molecules present high binding affinities with metal ions.28,43 We anticipated that these molecules can function to remove the coordinated roadblock and reactivate DNA synthesis. Our strategic direction is fully supported by the following results (Figures 3D and S7). From these observations, the inhibition of DNA synthesis by Cu2+ and Ni2+ can be almost fully removed by the presence of 2 equiv of various CA molecules. In addition, the treatment with CA molecules does not influence DNA synthesis without 5fC-AQA nucleotide (lanes 11−18 in Figures 3D and S7). Moreover, the CA supplementation recovered DNA synthesis at 5fC-AQA site in a concentration-dependent manner (Figure S8). In our proposed mechanism of action, the coordination of aromatic quinoline ring plays a key role in bending the 5fCAQA side arm and regulating oligonucleotide functions. To functionally validate this mechanism, we consider replacing the nitrogen atom in the quinoline ring with a CH group. On theoretical grounds such a replacement would be expected to disturb coordinating interaction of the side arm considerably due to the lack of ligating sites on naphthalene ring. Hence, 2(aminooxy)-N-(naphthalen-1-yl)acetamide (ANA), a closely related derivative of AQA, has been synthesized (details in Supporting Information). We first examined the reactivity of ANA toward 5fC nucleotide in DNA. As demonstrated in Figure S9, a site-specific labeling of 5fC-containing DNA has been achieved. In addition, the product DNA has a mass consistent with the expected attachment of an ANA group (Figure S2). We further examined the pattern of DNA synthesis along ANA-labeled templates. From our observations, the progress of DNA replication was not stopped at 5fCANA (+2 site) by supplement with various metal ions (Figure S10). Taken together, these results suggest that specific coordinating interaction forces the 5fC-AQA side arm to stay in a bent geometry, presumably because of steric hindrance from the quinoline ring, which lies directly opposite the complementary nucleobase, thereby reducing DNA synthesis. Steady-State Kinetics. To quantitatively address the influence of supramolecular coordination on DNA synthesis, single-turnover and steady-state measurements were performed.44,45 We therefore used the extension primer (p3 in Table S1) with the 3′ end lying immediately adjacent to 5fC residue. In this study, Bst DNA pol was preincubated with a large excess of DNA substrate and rapidly mixed with dGTP for extension. Under these conditions, only one nucleotide is added to the primer because the reaction mixture does not contain the nucleotide complementary to the next position on DNA templates. Initial velocities were calculated in units of nM/s and plotted versus the concentration of dGTP used in each case. This study can readily provide steady-state kinetics parameters Vmax and KM.46 Measurements of kcat (catalytic rate constant) and kcat/KM (catalytic efficiency) were further achieved. A representative range of data is demonstrated (red lines in Figures 4, S11, and S12). On the basis of our results, coordinating interaction (Cu2+, Ni2+, or Co2+) contributes to an evident reduction in catalytic efficiency (kcat/KM) with Bst DNA pol (Table S3). This study quantitatively reflected the impeding impact of coordinating interaction on dGTP incorporation opposite 5fC-AQA nucleotide. We further examined effects of CA supplementation on DNA synthesis. In this study, the AQA-reacted fC-27mer was treated with those identified metal ions (Cu2+, Ni2+ or Co2+) as above before 2 equiv of CA was added. The results

Figure 4. Steady-state kinetics for supramolecular coordination. (A) Representative Michaelis−Menten plots for dGTP incorporation under different conditions. (B) Relative catalytic efficiency (kcat/KM) of Bst DNA pol under different conditions. All the kinetics parameters are summarized in Table S3. The concentrations of dGTP were varied from 0.06 to 1.95 μM.

demonstrated that the CA supplementation significantly reversed the behavior of coordinating interaction on dGTP incorporation (blue lines in Figures 4, S11, and S12). Collectively, these results elucidate the high performance and reversibility of supramolecular coordination for targeting 5fCDNAs with AQA labeling. Side Arm Strategy for Reversible Control of DNA Digestion. In biological study, restriction enzymes have been used as model systems to examine DNA−protein interactions due to their high specificity for DNA sequences and the simplicity of recognition sites.47−49 We next set out to control this process by manipulating supramolecular coordination.50 The SnaBI digestion was studied for the proof of concept. We designed and synthesized a series of double-stranded (ds) DNA substrates (ds-fC-20mer1, ds-fC-20mer2, and ds-fC20mer3 in Tables S1 and S2), which varied depending on the position of the 5fC residue relative to the SnaBI recognition site. The according duplex without 5fC modification (ds-C20mer in Tables S1 and S2) was used as the control. As the first step, we examined the SnaBI digestion of all substrates both with and without AQA treatments in the absence of metal ion supplementation (Figure S13). The results show that the 5fC-AQA modification right in the recognition sequence mildly reduces the rate of cleavage (lane 5 in Figure S13). All other constructs with modifications outside of the recognition site were smoothly cleaved (lanes 7, 9 in Figure S13). Hence, the introduction of AQA side arm alone is relatively mild and does not represent a major roadblock in DNAs. We next examined wether those metal 15845

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5fC site outside of the recognition sequence (ds-fC-20mer2 and ds-fC-20mer3), the cleavage reaction was not substantially affected with Cu2+ and Co2+ at concentrations lower than the threshold of nonspecific inhibition (Figures S15 and S16). Importantly, Ni2+ can regulate the cleavage of ds-fC-20mer2 in a concentration-dependent manner (Figure 5B). Higher concentrations of Ni2+ were required to completely inhibit the digestion of ds-fC-20mer2 (800 μM, lane 15 in Figure 5B) compared to that of ds-fC-20mer1 (150 μM, lane 9 in Figure 5B). From these data analyzed the cleavage of the AQAreacted ds-fC-20mer3 was not influenced by treatment with all those metal ions. According to these results, the steric bulk of the added side arm was significantly increased by coordinating interaction. Importantly, the control of enzymatic action depends on proximity of the 5fC site to the recognition sequence of the endonuclease. Next, we examined whether the coordinating roadblock can be competed away with CA supplementation. In this study, the preparations were made by adding various amounts of CA to the mixture and incubated before endonuclease was added. The control assay demonstrated that CA supplementation did not significantly influence the endonuclease activity (Figure S17). For ds-fC-20mer1, CA supplementation efficiently reversed the blockage of digestion by interruption of coordinating interaction (Figures S18−S20). A similar phenomenon was also observed in the performance of CA supplementation on the cleavage of ds-fC-20mer2 (Figure S20). Moreover, the cleavage of DNA was recovered by CA supplementation with a concentration-dependent manner (Figure 5C and S21). Taken together, the “charm” of the supramolecular approach lies in the reversibility of the roadblock, which can be controlled on demand. Complexation Studies. On the basis of the above studies, the possible role for supramolecular coordination is that of the control of an N−H hydrogen bond critical for Watson−Crick base pairing.50 If so, the coordinating interaction will reduce the effect of hydrogen bonding, leading to decreased stability of DNA duplexes.51,52 To test this, a UV melting assay was performed using different DNA duplexes (ds-C-15mer and dsfC-15mer in Tables S1 and S4).39 Figures S22 and S23A showed normalized UV melting curves of both untreated duplexes under different conditions. For these samples, there were only small variations in Tm values upon Cu2+ treatment (ΔTm ≤ 0.52 °C), displaying no obvious change in duplex stability. We further investigated the stability of AQA-reacted ds-fC-15mer under different conditions. Installation of the auxiliary side arm (AQA) only results in a small depression in Tm (0.12 °C). Importantly, the Cu2+-treated sample showed an evidently lower Tm as compared with that of the no-treatment control (ΔTm = 5.14 °C, Figures 6 and S23B). Moreover, this trend was reversed by subsequent treatment with EGTA chelating. Our study demonstrates that the Cu2+ binding to 5fC-AQA nucleotide significantly destabilizes the Watson− Crick base pairs in paired region of DNA duplexes.

ions exhibit unspecific effects on SnaBI activities. As demonstrated in Figure S14, Ni2+ and Co2+ inhibit SnaBI activity only at concentrations higher than 2 mM, whereas Cu2+ exhibits greater nonspecific toxicity at smaller concentrations (120 μM). We next examined the influence of coordinating interaction on the cleavage of each AQA-treated DNA construct. For the construct with 5fC site within the recognition sequence (ds-fC20mer1), direct and concentration-dependent regulation of cleavage was observed with all those identified metal ions (Figures 5B, S15, and S16). However, for the constructs with

Figure 5. Reversible control of DNA digestion. (A) Schematic illustration of the workflow. The shaded region of DNA substrate indicates the effective area of supramolecular coordination. (B) The influence of Ni2+-coordination on DNA cleavage. Lane 1: no enzyme control; lanes 2, 4, 6, 10, 12, 16, 18: no Ni2+ control; lane 3: 1.0 mM Ni2+; lane 5: 150 μM Ni2+; lane 7: 50 μM Ni2+; lane 8: 100 μM Ni2+; lane 9: 150 μM Ni2+; lane 11: 800 μM Ni2+; lane 13: 400 μM Ni2+; lane 14: 600 μM Ni2+; lane 15: 800 μM Ni2+; lane 17: 1.0 mM Ni2+; lane 19: 600 μM Ni2+; lane 20: 800 μM Ni2+; lane 21: 1.0 mM Ni2+. (C) The influence of chelating on DNA cleavage. Lane 1: no enzyme control; lanes 2, 10: no metal/chelating control; lane 3: 200 μM Ni2+; lane 4: 200 μM Ni2+, 200 μM EDTA; lane 5: 200 μM Ni2+, 300 μM EDTA; lane 6: 200 μM Ni2+, 400 μM EDTA; lane 7: 200 μM Ni2+, 200 μM EGTA; lane 8: 200 μM Ni2+, 300 μM EGTA; lane 9: 200 μM Ni2+, 400 μM EGTA; lane 11: 1.0 mM Ni2+; lane 12: 1.0 mM Ni2+, 250 μM EDTA; lane 13: 1.0 mM Ni2+, 500 μM EDTA; lane 14: 1.0 mM Ni2+, 1.0 mM EDTA; lane 15: 1.0 mM Ni2+, 250 μM EGTA; lane 16: 1.0 mM Ni2+, 500 μM EGTA; lane 17: 1.0 mM Ni2+, 1.0 mM EGTA. For (B) and (C), reactions were carried out as described in the Experimental Section. Denaturing electrophoresis was used to analyze the cleaved products.

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DISCUSSION Supramolecular interactions play essential roles in the transfer of biological information and have received considerable attention in living systems.53 Despite a large number of metal−peptide complexes, the metal−DNA complexes are considerably less studied.54 In chemical structure, DNA nucleobases are heterocyclic compounds.55 From a viewpoint of coordinating interaction, they are excellent moieties because 15846

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metal). In a sense, the side arm behaves as a controller-like tool, providing access to control oligonucleotide functions. Such supramolecular coordination-based tool can also be extended to control some other DNA−protein interactions associated with DNA modifications.

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CONCLUSION In summary, the current study suggests a coordination directional bonding approach to control DNA−protein interactions at epigenetic DNA marks. Given these results, we believe that supramolecular coordination could potentially be exploited in the field of DNA epigenetics.

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Figure 6. Supramolecular coordination-directed reversible control of Watson−Crick base pairs. The Cu2+-coordination leads to an evident decrease in Tm of AQA-reacted ds-fC-15mer as compared with that of the no-treatment control.

EXPERIMENTAL SECTION

Materials. Quinolin-8-amine (CAS# 578−66−5), naphthalen-1amine (CAS# 134−32−7), 2-bromoacetyl bromide (CAS# 598−21− 0), 2-hydroxyisoindoline-1,3-dione (CAS# 524−38−9), hydrazine hydrate (CAS# 10217−52−4), 1,2-cyclohexylenedinitrilotetraacetic acid, (DCyTA, CAS# 13291−61−7), diethylenetriamineN,N,N′,N″,N″-pentaacetic acid (DTPA, CAS# 67−43−6), ethylenediaminetetraacetic acid (EDTA, CAS# 60−00−4), [ethylenebis(oxyethylenenitrilo)]tetraacetic acid (EGTA, CAS# 67−42−5), N-(2hydroxyethyl)ethylenediaminetriacetic acid (HEDTA, CAS# 150− 39−0), and nitrilotriacetic acid (NTA, CAS# 139−13−9) were purchased from Sigma-Aldrich Inc. (St. Louis, MO, USA). The oligonucleotides were synthesized with TaKaRa company (Dalian, China). Bst DNA polymerase, large fragment (M0275), and SnaBI (R0130) were purchased from New England Biolabs, Inc. The Super GelRed (NO.: S-2001) was purchased from US Everbright Inc. (Suzhou, China). The pH was determined with Mettler Toledo, FE20-Five Easy pH (Mettler Toledo, Switzerland). AP-15mer was synthesized through U-15mer (one T site was replaced by uracil) treating with Uracil DNA Glycosylase (Invitrogen, USA). DNA MALDI-TOF mass spectra were measured on MALDI-TOF-MS (Shimadzu, Japan). DNA concentration was determined with NanoDrop 2000c (Thermo Scientific, USA). Visualization of DNA in gel was performed with Pharos FX Molecular imager (Bio-Rad, USA). Chemical Synthesis. AQA and ANA were synthesized using a three-step procedure.33,34 The 5fC nucleoside was synthesized according to a previous literature.60 Full experimental procedures and structure characterization data are provided in the Supporting Information. The HRMS was recorded with Thermo Scientific Dionex Ultimate 3000 hybrid LTQ Orbitrap Elite Velos Pro (Thermo Scientific, USA). The stock solution of AQA or ANA is prepared fresh just prior to each use. Labeling of 5fC Nucleotide in DNA. The labeling reaction was performed in 40 mM sodium acetate buffer (pH = 5.0), 500 μM AQA or ANA (newly synthesized), 10 mM p-anisidine, and 0.5 μM DNA at room temperature for 12 h. Ethanol precipitation was performed to recover the DNA products from the reaction mixture. Electrophoresis. The acrylamide concentration of the separating gel was 20% (19:1 monomer to bis ratio). Electrophoresis was run in a temperature-controlled vertical electrophoretic apparatus (DYCZ22A, Liuyi Instrument Factory, Beijing, China). About 100 ng of DNA with different treatment were loaded on the gel. Electrophoresis was run at 10 °C for 1 h at 200 V and 3 h at 300 V. DNA was stained with GelRed dye. After electrophoresis, in-gel DNA was visualized using a Pharos FX Molecular imager (Bio-Rad, USA) in the fluorescence mode (λex = 590 nm). Reversible Control of DNA Synthesis. The reactions were performed in 1 × ThermoPol reaction buffer, which contained 20 mM Tris-HCl, 10 mM (NH4)2SO4, 10 mM KCl, 2 mM MgSO4, and 0.1% Triton X-100 at pH 8.8 @ 25 °C. To prepare the elongation scaffolds, 5′-fluorophore-labeled primer was hybridized with each template (with or without AQA treatment) at a molar ratio of 1:1.10. For coordinating interaction assay, the assembled primer/template duplexes (100 ng) were incubated in 1 × ThermoPol reaction buffer

their multiple functional groups can serve as potential ligating sites.56 To further enhance the binding affinity of coordination, an auxiliary side arm has been introduced as a bridge and a versatile unit in this study. The exocyclic heteroatoms of 5fC and the aromatic/aliphatic amine of auxiliary side arm independently exhibit affinity for metal ions; moreover, when they are combined, the binding specificity and strength have been significantly enhanced via the synergistic effects. The metal ions with a coordination number of four, such as Cu2+ and Ni2+, probably provide extreme examples of this effect. For these identified metal complexes, two different geometries are possible: tetrahedral or square planar.57 On the basis of all described data, a square planar structure is more favored. The 5fC represents an important epigenetic mark within mammalian DNA genome. Hence, it is important to develop a chemical tool for selective identification of 5fC modifications.58,59 In Watson−Crick base pairing, 5fC forms three hydrogen bonds with guanine. Not surprisingly, the AQA side arm itself does not sterically interfere with the hydrogen bonding between base pairs. Importantly, the coordinating interaction functions well in dictating folding morphology of side arm with precision. In this case, the bending of an auxiliary group may protrude into the Watson−Crick face and thus interfere with the binding to the complementary nucleobase. In coordination complex, the quinoline ring and the complementary guanine are confined in closer proximity to each other, and steric repulsion would become dominant over the hydrogen bonding forces. The beneficial directing effect of AQA side arm provides an inspiration for exploring other biologically relevant applications. For example, the application of this approach for fragment- and site-selective detection of DNA 5fC is performed. Very few studies have been conducted on the effects of supramolecular coordination on the functionality of DNA targeting enzymes. The valued characteristic of supramolecular coordination lies in the inherent property of reversibility. In this study, different examples are presented that demonstrate that the reversible nature of the supramolecular coordination chemistry employed allows blocking and deblocking of DNA− protein interactions: cleavage by an endonuclease and strand elongation by a DNA polymerase. Our strategy relies on the metal-coordinating bond, which is dynamically formed and dissociated between 5fC-AQA and metal ion supplementation. The binding of metal ions to 5fC-AQA effectively acts as a roadblock to different enzymes. The subsequent CA treatment can remove this barrier through competitive binding (CA15847

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Journal of the American Chemical Society

preparation was heated at 90 °C for 5 min. The following steps were similar to the above one. UV Melting Assay. DNA duplex (10 μM) was prepared by annealing equimolar amounts of single-stranded oligonucleotides for 5 min at 90 °C, cooling down to 10 °C over a period of 2 h, and incubation for 30 min at 10 °C in 1 × ThermoPol reaction buffer. The UV melting assay was performed using a Jasco-810 spectropolarimeter (Jasco, Easton, MD, USA) equipped with a water bath temperaturecontrol accessory. The detection is performed with a quartz cell (optical path length at 1 mm). The UV melting profiles were recorded with a heating rate of 0.2 °C/min, and the absorbance values were collected every 1 °C. The melting point (Tm) corresponds to the midtransition temperature, which was determined by the maximum of the first derivative of the absorbance as a function of temperature. Statistical Analysis. Statistical analysis was performed using OriginPro 2016 software. The differences were considered to be significant for P < 0.05.

in the presence or absence of metal ions supplementation for 20 min at room temperature, before adding dNTP mix (100 μM each) and Bst DNA pol (0.5 U) on ice. As a control without metal ions supplementation, the same amount (0.5 μL) of water was added. The following reactions were performed in a Bio-Rad PTC-100 programmable thermal controller (30 °C for 30 min). Enzyme was inactivated by addition of a 4.5-fold excess of quenching solution (95% formamide, 25 mM EDTA at pH 8.0). The products were separated on a denaturing 20% polyacrylamide gel (400 V, 4 h). After electrophoresis, in-gel DNA was visualized using a Bio-Rad Pharos FX Molecular imager in the fluorescence mode (λex = 488 nm). The starting duplex without any treatment is used as the control. For CA competition assay, the duplex DNA was incubated in 1 × ThermoPol reaction buffer in the presence of metal ion supplementation for 20 min, before adding each CA to the given concentration and incubation for an additional 5 min at 90 °C. The following steps are similar to the above one. Steady-State Single-Turnover Kinetics. Steady-state kinetics was performed using a procedure similar to that described in a previous literature.46 Reaction times and enzyme concentrations were predetermined to ensure steady-state conditions. A large excess of primer/template duplex was preincubated with Bst DNA pol. Reactions were initiated by the addition of dGTP, incubated at 30 °C for varying times, followed by quenching with a 4.5-fold excess of quenching solution (95% formamide, 25 mM EDTA at pH 8.0) at 90 °C. Products were separated by denaturing electrophoresis on a polyacrylamide gel. Intensities of bands P and P + 1 (IP and IP+1) were quantified using Image Lab software version 5.1. Equal volumes were used to quantify the densities of both bands P and P + 1 as well as a background volume for each pair of bands. The CP+1 (concentration of strand P + 1) was calculated using eq 1, where Ct is C

C P + 1 = C PIP + 1/(IP + IP + 1)

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S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.8b09113.

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AUTHOR INFORMATION

*[email protected] *[email protected]

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ORCID

Tian Tian: 0000-0002-0340-230X Xiang Zhou: 0000-0002-1829-9368 Author Contributions ‡

These authors contributed equally.

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Notes

The authors declare no competing financial interest.

The catalytic rate constant kcat was determined by dividing Vmax with the total enzyme concentration (Et, 15 nM), which was calculated using the specific activity and the molecular weight of Bst DNA Pol. From the measured kcat and KM values, their ratio was calculated. The kcat/KM of the enzyme is proportional to a rate of dGTP incorporation when the concentration of dGTP approaches 0. kcat = Vmax /Et

General methods; sequences of the used oligomers; more experimental data and more experimental details (PDF)

Corresponding Authors

Initial reaction velocities (V, nM/s) were derived by performing a linear regression with the data (CP+1 versus time). Each experiment was repeated three times. The data (V versus [dGTP]) were then fit to the Michaelis−Menten eq (eq 2) to determine steady-state kinetics parameters Vmax and KM V = Vmax[dGTP] /(KM + [dGTP])

ASSOCIATED CONTENT

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ACKNOWLEDGMENTS The authors thank the National Science Foundation of China (21722803, 21721005, 21572169, 21877086, and 21672165).

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REFERENCES

(1) Feng, S.; Cokus, S. J.; Zhang, X.; Chen, P. Y.; Bostick, M.; Goll, M. G.; Hetzel, J.; Jain, J.; Strauss, S. H.; Halpern, M. E.; Ukomadu, C.; Sadler, K. C.; Pradhan, S.; Pellegrini, M.; Jacobsen, S. E. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 8689. (2) Tahiliani, M.; Koh, K. P.; Shen, Y.; Pastor, W. A.; Bandukwala, H.; Brudno, Y.; Agarwal, S.; Iyer, L. M.; Liu, D. R.; Aravind, L.; Rao, A. Science 2009, 324, 930−935. (3) Ito, S.; Shen, L.; Dai, Q.; Wu, S. C.; Collins, L. B.; Swenberg, J. A.; He, C.; Zhang, Y. Science 2011, 333, 1300. (4) He, Y. F.; Li, B. Z.; Li, Z.; Liu, P.; Wang, Y.; Tang, Q.; Ding, J.; Jia, Y.; Chen, Z.; Li, L.; Sun, Y.; Li, X.; Dai, Q.; Song, C. X.; Zhang, K.; He, C.; Xu, G. L. Science 2011, 333, 1303. (5) Maiti, A.; Drohat, A. C. J. Biol. Chem. 2011, 286, 35334. (6) Zhang, L.; Lu, X.; Lu, J.; Liang, H.; Dai, Q.; Xu, G. L.; Luo, C.; Jiang, H.; He, C. Nat. Chem. Biol. 2012, 8, 328. (7) Maiti, A.; Michelson, A. Z.; Armwood, C. J.; Lee, J. K.; Drohat, A. C. J. Am. Chem. Soc. 2013, 135, 15813. (8) Pfaffeneder, T.; Hackner, B.; Truss, M.; Munzel, M.; Muller, M.; Deiml, C. A.; Hagemeier, C.; Carell, T. Angew. Chem., Int. Ed. 2011, 50, 7008. (9) Song, C. X.; He, C. Trends Biochem. Sci. 2013, 38, 480.

Reversible Control of DNA Digestion. The reactions were performed in 1 × CutSmart buffer, which contained 50 mM potassium acetate, 20 mM Tris-acetate, 10 mM magnesium acetate, and 100 μg/mL BSA at pH 7.9 @ 25 °C. The digestion constructs were prepared by mixing 5′-FAM-labeled strand (with or without AQA treatment) with the complementary strand at equal molar ratio, followed by a heating at 90 °C for 3 min and annealing over a period of 2 h. For coordinating interaction assay, the duplex DNA construct (100 ng) was treated with each metal ion at given concentrations for 20 min at room temperature before adding SnaBI (1.0 U) on ice. The digestion was performed at 37 °C for 1 h. As a control without metal ion supplementation, the same amount (0.5 μL) of water was added to the solution instead. Digestions were terminated by addition of a 4.5-fold excess of quenching solution (95% formamide, 25 mM EDTA at pH 8.0). The products were analyzed on a denaturing 20% polyacrylamide gel (400 V, 4 h). After electrophoresis, in-gel DNA was visualized using a fluorescence imager. For CA competition assay, the sample was prepared in 1 × Tango buffer containing duplex DNA and metal ion supplementation, followed by the addition of CA at given concentrations. The 15848

DOI: 10.1021/jacs.8b09113 J. Am. Chem. Soc. 2018, 140, 15842−15849

Article

Journal of the American Chemical Society (10) Wang, L.; Zhou, Y.; Xu, L.; Xiao, R.; Lu, X.; Chen, L.; Chong, J.; Li, H.; He, C.; Fu, X. D.; Wang, D. Nature 2015, 523, 621. (11) Raiber, E. A.; Beraldi, D.; Ficz, G.; Burgess, H. E.; Branco, M. R.; Murat, P.; Oxley, D.; Booth, M. J.; Reik, W.; Balasubramanian, S. Genome Biol. 2012, 13, R69. (12) Zhu, C.; Gao, Y.; Guo, H.; Xia, B.; Song, J.; Wu, X.; Zeng, H.; Kee, K.; Tang, F.; Yi, C. Cell Stem Cell 2017, 20, 720. (13) Bachman, M.; Uribe-Lewis, S.; Yang, X.; Burgess, H. E.; Iurlaro, M.; Reik, W.; Murrell, A.; Balasubramanian, S. Nat. Chem. Biol. 2015, 11, 555. (14) Raiber, E. A.; Murat, P.; Chirgadze, D. Y.; Beraldi, D.; Luisi, B. F.; Balasubramanian, S. Nat. Struct. Mol. Biol. 2015, 22, 44. (15) Hardwick, J. S.; Ptchelkine, D.; El-Sagheer, A. H.; Tear, I.; Singleton, D.; Phillips, S. E. V.; Lane, A. N.; Brown, T. Nat. Struct. Mol. Biol. 2017, 24, 544. (16) Ji, S.; Fu, I.; Naldiga, S.; Shao, H.; Basu, A. K.; Broyde, S.; Tretyakova, N. Y. Nucleic Acids Res. 2018, 46, 6455. (17) Li, F.; Zhang, Y.; Bai, J.; Greenberg, M. M.; Xi, Z.; Zhou, C. J. Am. Chem. Soc. 2017, 139, 10617. (18) Xu, L.; Chen, Y. C.; Nakajima, S.; Chong, J.; Wang, L.; Lan, L.; Zhang, C.; Wang, D. Chem. Sci. 2014, 5, 567−574. (19) Song, C. X.; Szulwach, K. E.; Dai, Q.; Fu, Y.; Mao, S. Q.; Lin, L.; Street, C.; Li, Y.; Poidevin, M.; Wu, H.; Gao, J.; Liu, P.; Li, L.; Xu, G. L.; Jin, P.; He, C. Cell 2013, 153, 678. (20) Su, M.; Kirchner, A.; Stazzoni, S.; Muller, M.; Wagner, M.; Schroder, A.; Carell, T. Angew. Chem., Int. Ed. 2016, 55, 11797. (21) Cook, T. R.; Stang, P. J. Chem. Rev. 2015, 115, 7001. (22) Tavenor, N. A.; Murnin, M. J.; Horne, W. S. J. Am. Chem. Soc. 2017, 139, 2212. (23) Ma, X.; Jia, J.; Cao, R.; Wang, X.; Fei, H. J. Am. Chem. Soc. 2014, 136, 17734. (24) Haas, K. L.; Franz, K. J. Chem. Rev. 2009, 109, 4921. (25) Li, X. Z.; Zhou, L. P.; Yan, L. L.; Dong, Y. M.; Bai, Z. L.; Sun, X. Q.; Diwu, J.; Wang, S.; Bunzli, J. C.; Sun, Q. F. Nat. Commun. 2018, 9, 547. (26) Eibling, M. J.; MacDermaid, C. M.; Qian, Z.; Lanci, C. J.; Park, S. J.; Saven, J. G. J. Am. Chem. Soc. 2017, 139, 17811. (27) Kiyonaka, S.; Kubota, R.; Michibata, Y.; Sakakura, M.; Takahashi, H.; Numata, T.; Inoue, R.; Yuzaki, M.; Hamachi, I. Nat. Chem. 2016, 8, 958. (28) Engelhard, D. M.; Pievo, R.; Clever, G. H. Angew. Chem., Int. Ed. 2013, 52, 12843. (29) Cook, T. R.; Zheng, Y. R.; Stang, P. J. Chem. Rev. 2013, 113, 734. (30) Lippert, B.; Sanz Miguel, P. J. Acc. Chem. Res. 2016, 49, 1537. (31) Berti, L.; Burley, G. A. Nat. Nanotechnol. 2008, 3, 81. (32) Liao, S.; Sun, X. L.; Tang, Y. Acc. Chem. Res. 2014, 47, 2260. (33) Zhu, J. F.; Yuan, H.; Chan, W. H.; Lee, A. W. Org. Biomol. Chem. 2010, 8, 3957. (34) Guo, P.; Yan, S.; Hu, J.; Xing, X.; Wang, C.; Xu, X.; Qiu, X.; Ma, W.; Lu, C.; Weng, X.; Zhou, X. Org. Lett. 2013, 15, 3266. (35) Larsen, D.; Pittelkow, M.; Karmakar, S.; Kool, E. T. Org. Lett. 2015, 17, 274. (36) Matray, T. J.; Kool, E. T. Nature 1999, 399, 704−708. (37) Greenberg, M. M. Acc. Chem. Res. 2014, 47, 646. (38) Chou, C.; Young, D. D.; Deiters, A. Angew. Chem., Int. Ed. 2009, 48, 5950. (39) Young, D. D.; Edwards, W. F.; Lusic, H.; Lively, M. O.; Deiters. Chem. Commun. (Cambridge, U. K.) 2008, 462. (40) Samanta, B.; Seikowski, J.; Hobartner, C. Angew. Chem., Int. Ed. 2016, 55, 1912. (41) Shi, C.; Shen, X.; Niu, S.; Ma, C. J. Am. Chem. Soc. 2015, 137, 13804. (42) Kiefer, J. R.; Mao, C.; Hansen, C. J.; Basehore, S. L.; Hogrefe, H. H.; Braman, J. C.; Beese, L. S. Structure 1997, 5, 95. (43) Day, H. A.; Wright, E. P.; MacDonald, C. J.; Gates, A. J.; Waller, Z. A. Chem. Commun. (Cambridge, U. K.) 2015, 51, 14099. (44) Einolf, H. J.; Schnetz-Boutaud, N.; Guengerich, F. P. Biochemistry 1998, 37, 13300.

(45) Shibata, A.; Uzawa, T.; Nakashima, Y.; Ito, M.; Nakano, Y.; Shuto, S.; Ito, Y.; Abe, H. J. Am. Chem. Soc. 2013, 135, 14172. (46) Alenko, A.; Fleming, A. M.; Burrows, C. J. Biochemistry 2017, 56, 5053. (47) Schierling, B.; Noel, A. J.; Wende, W.; Hien le, T.; Volkov, E.; Kubareva, E.; Oretskaya, T.; Kokkinidis, M.; Rompp, A.; Spengler, B.; Pingoud, A. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 1361. (48) Mackova, M.; Bohacova, S.; Perlikova, P.; Postova Slavetinska, L.; Hocek, M. ChemBioChem 2015, 16, 2225. (49) Kielkowski, P.; Macickova-Cahova, H.; Pohl, R.; Hocek, M. Angew. Chem., Int. Ed. 2011, 50, 8727−8730. (50) Young, D. D.; Govan, J. M.; Lively, M. O.; Deiters, A. ChemBioChem 2009, 10, 1612. (51) Wang, R.; Jin, C.; Zhu, X.; Zhou, L.; Xuan, W.; Liu, Y.; Liu, Q.; Tan, W. J. Am. Chem. Soc. 2017, 139, 9104. (52) Dohno, C.; Uno, S. N.; Nakatani, K. J. Am. Chem. Soc. 2007, 129, 11898. (53) Uhlenheuer, D. A.; Petkau, K.; Brunsveld, L. Chem. Soc. Rev. 2010, 39, 2817. (54) Boerner, L. J.; Zaleski, J. M. Curr. Opin. Chem. Biol. 2005, 9, 135. (55) Cyranski, M. K.; Gilski, M.; Jaskolski, M.; Krygowski, T. M. J. Org. Chem. 2003, 68, 8607. (56) Wang, F.; Liu, B.; Huang, P. J.; Liu, J. Anal. Chem. 2013, 85, 12144. (57) Rulisek, L.; Vondrasek, J. J. Inorg. Biochem. 1998, 71, 115. (58) Xia, B.; Han, D.; Lu, X.; Sun, Z.; Zhou, A.; Yin, Q.; Zeng, H.; Liu, M.; Jiang, X.; Xie, W.; He, C.; Yi, C. Nat. Methods 2015, 12, 1047. (59) Sun, Z.; Dai, N.; Borgaro, J. G.; Quimby, A.; Sun, D.; Correa, I. R., Jr.; Zheng, Y.; Zhu, Z.; Guan, S. Mol. Cell 2015, 57, 750. (60) Dai, Q.; He, C. Org. Lett. 2011, 13, 3446.

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