A Push–Pull Mechanism Helps Design Highly Competent G-Quadruplex-DNA Catalysts

Open AccessCCS ChemistryRESEARCH ARTICLE1 Aug 2021A Push–Pull Mechanism Helps Design Highly Competent G-Quadruplex-DNA Catalysts Jielin Chen†, Jiawei Wang†, Stephanie C. C. van der Lubbe†, Mingpan Cheng†, Dehui Qiu, David Monchaud, Jean-Louis Mergny, Célia Fonseca Guerra, Huangxian Ju and Jun Zhou Jielin Chen† State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 †J. Chen, J. Wang, S.C.C. van der Lubbe, and M. Cheng contributed equally to this work.Google Scholar More articles by this author , Jiawei Wang† State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 †J. Chen, J. Wang, S.C.C. van der Lubbe, and M. Cheng contributed equally to this work.Google Scholar More articles by this author , Stephanie C. C. van der Lubbe† Theoretical Chemistry, Department of Chemistry and Pharmaceutical Sciences, AIMMS, Vrije Universiteit Amsterdam, Amsterdam 1081 HV †J. Chen, J. Wang, S.C.C. van der Lubbe, and M. Cheng contributed equally to this work.Google Scholar More articles by this author , Mingpan Cheng† State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 †J. Chen, J. Wang, S.C.C. van der Lubbe, and M. Cheng contributed equally to this work.Google Scholar More articles by this author , Dehui Qiu State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 Google Scholar More articles by this author , David Monchaud Institut de Chimie Moléculaire (ICMUB), CNRS, Université de Bourgogne Franche-Comté, Dijon 21078 Google Scholar More articles by this author , Jean-Louis Mergny State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 Google Scholar More articles by this author , Célia Fonseca Guerra *Corresponding authors: E-mail Address: [email protected] E-mail Address: [email protected] E-mail Address: [email protected] Theoretical Chemistry, Department of Chemistry and Pharmaceutical Sciences, AIMMS, Vrije Universiteit Amsterdam, Amsterdam 1081 HV Leiden Institute of Chemistry, Gorlaeus Laboratories, Leiden University, Leiden 2333 CD Google Scholar More articles by this author , Huangxian Ju *Corresponding authors: E-mail Address: [email protected] E-mail Address: [email protected] E-mail Address: [email protected] State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 Google Scholar More articles by this author and Jun Zhou *Corresponding authors: E-mail Address: [email protected] E-mail Address: [email protected] E-mail Address: [email protected] State Key Laboratory of Analytical Chemistry for Life Science, School of Chemistry and Chemical Engineering, Nanjing University, Nanjing 210023 Google Scholar More articles by this author https://doi.org/10.31635/ccschem.020.202000473 SectionsSupplemental MaterialAboutAbstractPDF ToolsAdd to favoritesDownload CitationsTrack Citations ShareFacebookTwitterLinked InEmail Massive efforts are currently being invested to improve the performance, versatility, and scope of applications of nucleic acid catalysts. G-quadruplex (G4)/hemin DNAzymes are of particular interest owing to their structural programmability and chemical robustness. However, optimized catalytic efficiency is still bottleneck and the activation mechanism is unclear. Herein, we have designed a series of parallel G4s with different proximal cytosine (dC) derivatives to fine-tune the hemin-binding pocket for G4-DNAzymes. Combining theoretical and experimental methods, we have assessed the dependence of catalytic enhancement on the electronic properties of proximal dCs and demonstrated how proximal dCs activate catalytic proficiency. These results provide interesting clues in recapitulating the push–pull mechanism as the basis of peroxidase activity and help to devise a new strategy to design highly competent DNA catalysts whose performances are of the same order as protease. Download figure Download PowerPoint Introduction The catalytic capability of nucleic acids was first reported with the study of self-splicing RNAs, or ribozymes, by Kruger et al.1 and Guerriertakada,2 and of RNA-cleaving DNA enzymes, or DNAzymes, by Breaker and Joyce.3 Afterward, both ribozymes and DNAzymes were implemented as efficient biodevices in different fields including biosensing, chemical transformations, and chemical biology.4–7 G-quadruplex (G4)-DNAzymes represent a particular class of DNAzymes in which a four-stranded G4-DNA interacts with the cofactor hemin to perform hemoenzyme-type reactions.8–12 The current popularity of G4-DNAzymes stems from the versatile designability13–17 and excellent biocompatibility of G4 precatalysts.18–21 Unfortunately, the performances of G4-DNAzymes are still below those of corresponding enzymes, leading to new research and strategies for improved performance. To date, optimization strategies can be divided into two types: the addition of exogenous activators,22–29 such as spermine,22 cytidine triphosphate (CTP),23 adenine triphosphate (ATP),23–25 template-assembled synthetic G-quartet (TASQ),26 and self-assembling peptides27–29 on one side, and the modification of the G4 structure30–35 to provide hemin with a more defined and suited binding pocket31,32 or hemin itself36,37 on the other side. The latter mostly focuses on the modification of the so-called parallel G4 structure, in which all constitutive DNA strands are codirectionally oriented and the structure displays fairly accessible G-quartets, the privileged hemin-binding site.38 Efforts have been invested to fine-tune G4 structures to improve their catalytic performances, notably modifying the sequences flanking the external G-quartet,31,34,35,39 as well as the loops and bulges of the G4 core.33 As an example, it was demonstrated that polyadenine and polycytosine tails positively impact catalytic performance.31,33,35 Efforts have also been invested to better understand the mechanisms behind these catalytic improvements. Travascio et al. implemented UV–vis9 and electron paramagnetic resonance (EPR)10 spectroscopies to provide a convincing model for a G4/hemin system. Yamamoto et al.36,40 scrutinized the coordination of the hemin iron atom to gain reliable insights into the hemin/G-quartet interactions by both vibrational and nuclear magnetic resonance (NMR) spectroscopies. We and others have exploited the Lewis acid-base theory to demonstrate the synergistic cooperation of hemin activation between the external G-quartet and its proximal nucleotides.31,34 Herein, we take a further leap toward the elucidation of the actual G4-DNAzyme mechanism, demonstrating how the axial coordination of proximal dC activates hemin in a peroxidase-mimicking manner. Our approach was nature-driven: in the catalytic cycle of the model horseradish peroxidase (HRP), the so-called push–pull mechanism is central.41–44 As shown in Supporting Information Figure S1, the push effect being provided by the histidine 170 (His170) stabilizes the higher oxidation states of the iron atom during catalysis, and the pull effect being mediated by the His42 acting together with the Arg38 facilitates heterolytic O–O bond cleavage of the iron-bound hydrogen peroxide (the stoichiometric oxidant). Thus, we decided to modulate the electron density of the proximal dC to investigate whether and to which extent it influences the G4-DNAzyme activity. Our results demonstrate that electron-rich dCs strengthen the hydrogen bond with H2O2 and boost the Compound 0 → 0* → 1 transfer in a His42-like manner, which leads to notable catalytic enhancements.23,45 Experimental Methods Materials and reagents Oligonucleotides, purified by HPLC, were purchased from Sangon Biotech (Shanghai, China) and Takara Biomedical Technology (Dalian, China), dissolved in ultrapure water (18.2 MΩ·cm) and used without further treatment. The concentrations of oligonucleotides were determined by the UV–vis absorbance at 260 nm, using the molar extinction coefficients provided by OligoAnalyzer 3.1 ( http://sg.idtdna.com/calc/analyzer). About 100 mM Britton–Robinson buffer (B–R buffer) was prepared with 0.5 M H3BO3, 0.5 M H3PO4, and 0.5 M CH3COOH and titrated with 1 M KOH to pH 7.0. All experiments were performed in 10 mM B–R buffer (pH 7.0 except pH-dependence experiments) containing 0.05% Triton X-100, 0.1% DMSO, and 100 mM KCl. G4s (10 μM) were prepared in 10 mM B–R buffer, with 100 mM K+, heated to 95 °C for 5 min, cooled slowly to room temperature, and stored at 4 °C overnight prior to use. Hemin stock solution was prepared with DMSO. About 50 mM 2,2′-azino-bis-(3-ethylbenzothiozoline-6-sulfonic acid) (ABTS), β-nicotinamide adenine dinucleotide (NADH), and H2O2 were freshly prepared every time in ultrapure water and 50 mM 3,3′,5,5′-tetramethylbenzidine (TMB) in DMSO. All chemicals were obtained from Sigma (St. Louis, USA). Circular dichroism measurements About 5 μM G4 solutions were obtained by directly diluting stock solutions with 10 mM B–R buffer. Circular dichroism (CD) spectra were collected with three measurements on a Chirascan CD spectrometer (Applied Photophysics, Leatherhead, UK) in the wavelength range of 220–335 nm at 25 °C. Activity measurements of G4-DNAzymes About 0.4 μM G4s were incubated at 25 °C with 0.8 μM hemin in the B–R buffer (pH 7.0) for 2 h. After the formation of G4/hemin complexes, ABTS (0.6 mM, final concentration) was first added, and then variable concentrations of H2O2 were added, with final concentrations of 0.1, 0.15, 0.2, 0.3, 0.5, 1, 2.5, 5, and 10 mM to start the reactions, where the absorbance of ABTS·+ at 414 nm was monitored for 60 s by a Cary100 (Agilent, Santa Clara, USA) spectrophotometer. To further confirm DNAzyme activity, the other two substrates, TMB and NADH, were chosen. The absorbance of TMB·+ at 652 nm and NADH at 340 nm was recorded for 60 s. The extinction coefficient for ABTS·+ at 414 nm was 36,000 M−1 cm−1; TMB·+ at 652 nm was 39,000 M−1 cm−1; and NADH at 340 nm was 6220 M−1 cm−1.9,34 The initial rate (V0, nM s−1) of the oxidation reaction was calculated with the slope of the plot of absorbance versus reaction time in initial 10 s. The Michaelis constant, Km, was obtained with the Michaelis−Menten model. All kinetic measurements were repeated three times, and the background activity of hemin alone was subtracted. To examine the dependence of G4-DNAzyme activity on pH, the G4-DNAzyme activity was measured in 10 mM B–R buffers at different pHs from 3.0 to 8.0. After G4s (0.4 μM) were incubated with 0.8 μM hemin at 25 °C for 2 h, 0.6 mM ABTS and 0.6 mM H2O2 were added to initiate the reaction. Computational settings All calculations were performed with the density functional theory (DFT)-based program Amsterdam Density Functional (ADF) 2017.208 at the ZORA-BLYP-D3(BJ)/TZ2P level of theory for geometric optimization and binding energies.46–51 Geometries were optimized in implicit chloroform solvation with C1 (i.e., without) symmetry constraints because the dielectric constant ɛ of chloroform (ɛ = 4.8) is close to the experimentally determined dielectric constant of DNA (ɛ ≈ 8).52 All optimized structures have been verified to be true minima through vibrational analysis (zero imaginary frequencies). The binding energies ΔE were obtained by eq 1: Δ E = E dimer − E monomer 1 − E monomer 2 (1)where Edimer and Emonomer are the energies of the optimized dimers and monomers, respectively. Full computational details are given in the Supporting Information. Voronoi deformation density charges The atomic charge distribution was analyzed using the Voronoi deformation density (VDD) method.53 The VDD method partitions the space into so-called Voronoi cells, which are nonoverlapping regions of space that are closer to nucleus A than to any other nucleus. The charge distribution was determined by taking a fictitious promolecule as a reference point, in which the electron density was simply the superposition of the atomic densities. The change in density in the Voronoi cell when going from this promolecule to the final molecular density of the interacting system is associated with the VDD atomic charge (QVDD). Thus, the VDD atomic charge of atom A is given by eq 2: Q A VDD = − ∫ voronoi cell of A [ ρ ( r ) − ρ promolecule ( r ) ] d r (2)Therefore, instead of computing the amount of charge contained in an atomic volume, we computed the flow of charge from one atom to the other upon molecular formation. Therefore, the physical interpretation was straightforward. A positive atomic charge Q A VDD corresponds to a loss of electrons, whereas a negative atomic charge Q A VDD is associated with a gain of electrons in the Voronoi cell of atom A. This effectively means that the more electron-rich N3 atoms have a lower (i.e., more negative) VDD charge. Results and Discussion Proximal dC derivative designation for G4-DNAzymes It is now established that while hemin may bind to the two external G-quartets of a G4, it acquires better catalytic performances atop the 3′-terminal G-quartet.31,34 We also demonstrated that a proximal dC located near the 3′-terminal efficiently enhances the catalytic activity of the corresponding G4.34 Therefore, we decided to further optimize the structure of F3TC (d[T2G3TG3TG3TG3TC]), a parallel G4 tailed with a 3′-terminal dC that can be substituted by a variety of modified nucleobases (Figure 1a). Figure 1 | Schematic illustration of the G4-DNAzyme. (a) F3TC used as a model sequence. (b) Different dC derivatives used in this study. Download figure Download PowerPoint The dC was modified to modulate its electron density to assess the consequence of this modification on G4 catalytic efficiency, performing the oxidation of the dye ABTS as a model reaction in light of its readily monitorable output (Figure 2a). The electron density of dC was modulated by introducing different chemical groups on the dC core, changing the H5 hydrogen atom for OH (OH-dC, F3T-hC), bromine (Br-dC, F3T-brC), iodine (I-dC, F3T-iC), CH2OH (5-OH-Me-dC, F3T-hmC), or CH3 (Me-dC, F3T-mC), C5 for N5 (5-Aza-2′-dC, F3T-azaC), or merging an additional pyrrole ring (Pyrrolo-dC, F3T-pC) (Figure 1b and Supporting Information Table S1). Circular dichroism spectra ( Supporting Information Figure S2) confirmed that all these sequences adopt parallel G4 conformation, indicating that dC derivatives did not affect the G4 formation and its overall topology. Figure 2 | (a) Peroxidation reaction facilitated by G4-DNAzyme for oxidation of ABTS by H2O2. (b) Electrostatic potential surface (at 0.005 a.u.) from −0.1 (red) to 0.1 (blue) a.u. for prepared cytosine derivatives computed at the ZORA-BLYP-D3(BJ)/TZ2P level of theory in implicit chloroform solvation. (c) QVDD (in me–) on N3 atom in different cytosine derivatives in implicit chloroform solvation, obtained at the ZORA-BLYP-D3(BJ)/TZ2P level of theory. (d) Saturation curves for ABTS oxidation catalyzed by different G4-DNAzymes in the presence of H2O2 with increasing concentration. Download figure Download PowerPoint The electronic consequences of these derivatives were studied by analyzing the electrostatic potential surfaces (EPSs) (Figure 2b). The EPSs of the dC derivatives are close to the wild-type dC, with a negative charge accumulation centered on the N3 atom.46–51 To further quantify any differences between the various derivatives, we analyzed the atomic charges by using the VDD method.53 All N3 atoms in all dC derivatives possess a negative charge (Figure 2c), which is in line with the EPSs. The negative VDD charge (QVDD) accumulation is most pronounced in dC [QVDD = −305 milli-electrons (me−)], and the least amount of accumulation occurs in Br-dC and I-dC (−289 me−). The VDD charges of Me-dC (−303 me−) and 5-Aza-2′-dC (−303 me−) are approximately the same as the wild-type dC (−305 me−). Catalytic activity characterization The H2O2-mediated oxidation of ABTS catalyzed by dC-modified G4-DNAzymes was monitored via the time-dependent absorbance change at 414 nm, characteristic of the ABTS·+ formation9 (Figure 2d and Supporting Information Figure S3). All dC-modified G4-DNAzymes were found to be more catalytically proficient than F3T (d[T2G3TG3TG3TG3T]), but F3T-azaC (C5 to N5 substitution) and F3T-mC were found to be more efficient than F3TC. F3T-azaC exhibited the kcat and Km values of 25.39 ± 0.63 s−1 and 7.62 ± 0.28 mM, respectively, versus 12.68 ± 1.58 s−1 and 4.22 ± 1.17 mM for F3T-mC (Table 1), obtained according to Michaelis−Menten model ( Supporting Information Figure S4). The kcat and Km values of F3T-azaC and F3T-mC were 3.4- and 1.7-fold (kcat), and 1.8- and 0.97-fold (Km) those of F3TC, 84.6- and 42.3-fold (kcat), and 4.2- and 2.3-fold (Km) those of F3T. The kcat value of F3T-azaC was close to native enzymes HRP, which varies between 50 and 800 s−1 depending on the circumstances.54,55 Of note, under the same reaction conditions of F3T-azaC, the measured kcat value of HRP was 578 s−1 ( Supporting Information Figure S5). Interestingly, the catalytic activity of these different dC derivatives-based G4-DNAzyme was further confirmed through the oxidation of the other two substrates, TMB and NADH, and showed the same trend with ABTS ( Supporting Information Figure S6). Table 1 | Kinetic Parameters of Various Derivatives DNAzyme with Respect to the Catalysis of ABTS to ABTS·+a Sequencesb Km (mM) Vmax (μM s−1) kcat (s−1) kcat/Km (s−1·mM−1) kcat/k0c F3T 1.80 ± 0.41 0.12 ± 0.01 0.30 ± 0.02 0.17 1.0 F3T-pC 3.70 ± 0.48 0.17 ± 0.01 0.42 ± 0.02 0.11 1.4 F3T-hC 1.87 ± 0.24 0.11 ± 0.01 0.28 ± 0.01 0.15 0.9 F3T-brC 2.26 ± 0.21 0.28 ± 0.01 0.69 ± 0.02 0.30 2.3 F3T-iC 2.17 ± 0.25 0.40 ± 0.02 1.01 ± 0.04 0.47 3.4 F3T-hmC 3.47 ± 0.13 1.64 ± 0.03 4.09 ± 0.07 1.18 13.6 F3TC 4.34 ± 0.20 2.97 ± 0.06 7.44 ± 0.15 1.72 24.8 F3T-mC 4.22 ± 1.17 5.07 ± 0.63 12.68 ± 1.58 3.00 42.3 F3T-azaC 7.62 ± 0.28 10.15 ± 0.25 25.39 ± 0.63 3.33 84.6 aConditions: 0.1–10 mM H2O2, 0.6 mM ABTS, 0.4 μM G4/hemin DNAzyme. bSequences are shown in Supporting Information Table S1. ck0 = (0.30 ± 0.02) s−1, kcat = Vmax/0.4 μM. Potential mechanism for the dependence of G4-DNAzyme activity on proximal dC electron density To gain insights into how the electron density of dC influences catalysis, we next focused on the first catalytic stage of G4-DNAzyme, that is, the conversion of Compound 0 [coordination of H2O2 to the hemin Fe(III) atom] to Compound 0* (heterolytic cleavage of H2O2) to ultimately produce Compound 1 (vide infra). We monitored these steps via UV−vis measurements in the absence of ABTS substrate ( Supporting Information Figures S7 and S8) to quantify the H2O2-mediated oxidative degradation of G4/hemin complexes. G4-DNAzymes showed characteristic peaks at 404, 504, and 630 nm (Soret bands), which decreased rapidly after the addition of H2O2, implying the oxidative decay of the Fe(III) intermediate, concomitantly with the band increase in the 550–620 and 650–700 nm regions, characteristic of Compound 1 formation ( Supporting Information Figure S8).10,31 The plot of initial degradation velocity versus catalytic activity showed linear correlation (Figure 3a), in agreement with the involvement of dC electronic density in the formation of catalytic intermediates. A notable exception was F3T-azaC for which the decay rate was too fast to be reliably quantified. Figure 3 | (a) Correlation between initial degradation velocity and the activity (V0) of G4-DNAzymes at 0.4 mM H2O2. Note: The initial degradation velocity for F3T-azaC was too fast to record, leading to a deviation of its observed velocity from the actual one. (b) Catalytic activity of G4-DNAzymes at different pHs. Download figure Download PowerPoint Next, we focused on the formation of Compound 1, which is produced from Compound 0* after the loss of two electrons, one from the iron atom [which becomes Fe(IV)], and the other from the porphyrin ring. The formation of the catalytically active Fe(IV) is the rate-determining step of the catalysis. In the catalytic cycle of HRP, this step is promoted by His42 with a pKa of 2.5 (His42.H+/His42), suitable to assist H2O2 cleavage since the pKa of the hemin-bound H2O2/HO2− is between 3.2 and 4.0. In G4-DNAzymes, His42 is replaced by dC with a pKa of 4.1 (CTP.H+/CTP), which is also compatible with this transfer.23 We then evaluated the activity of dC-modified DNAzymes at pHs from 3.0 to 8.0 ( Supporting Information Figure S9). The catalytic proficiency of these G4-DNAzymes was found to be sensitive to pH, and F3TC, F3T-mC, F3T-hmC, and F3T-azaC showed relatively high activity with plateaus after 5.0 for F3TC, F3T-mC, and F3T-hmC, and 7.0 for F3T-azaC (Figure 3b). The apparent pKa was found to be 4.2 for F3T-hmC, 4.4 for F3TC, 4.5 for F3T-mC, and 6.4 for F3T-azaC, respectively. The dependence of pKa on dCs was in line with VDD charge (between −300 and −305 me−, Figure 2c), confirming that higher electron density on dCs induce higher apparent pKa, making N3 harder to be deprotonated. These results further confirm the involvement of dC in the rate-determining step of the catalysis, thus mimicking the role of His42 in HRP. The interaction between dCs and H2O2 was further investigated using DFT computations at the ZORA-BLYP-D3(BJ)/TZ2P level of theory in implicit chloroform solvation.46–51 The computed binding energy (ΔE) between dCs and H2O2 ranged between −10.7 and −11.5 kcal·mol−1 (Figure 4a). For most dCs, the binding energy (ΔE) and N3 atomic charge (QVDD) showed positive correlation with experimental activity (Figure 4b), implying a simple electronically driven H2O2 coordination with dCs. However, disconnections were found for Pyrrolo-dC, OH-dC, and 5-Aza-2′-dC. For Pyrrolo-dC, the VDD charge (−294 me−) and binding energy (−10.9 kcal·mol−1) were relatively negative in comparison with its activity of 21.0 nM s−1. This was probably caused by the absence of NH2 group and the presence of NH incorporated in another ring, which affected the size of the Voronoi cell and the VDD charge. OH-dC displayed both more negative (i.e., more stable) binding energy (−11.2 kcal·mol−1) and more negative VDD charge (−296 me−) than anticipated, while 5-Aza-2′-dC displayed a N3 VDD charge of −303 me− but a more positive (i.e., less stable) binding energy than that suggested by the experimental activity (682.6 nM s−1). Figure 4 | (a) Optimized structures, binding energies ΔE (in kcal·mol–1), and N−O distances (in Å) in implicit chloroform solvation, obtained at the ZORA-BLYP-D3(BJ)/TZ2P level of theory. (b) Activity (red columns) and computed binding energy (black line) of different DNAzymes with H2O2 at 0.6 mM, and QVDD of different cytosine derivatives on N3 (blue line). Download figure Download PowerPoint The observed discrepancies between V0 values and the theoretical results for 5-Aza-2′-dC and OH-dC can be explained through their unique capability to form hydrogen (H) bonds with two binding faces. 5-Aza-2′-dC (with high V0 but more positive ΔE) can coordinate with H2O2 via both its N3 atom and its N5 atom (Figures 5a and Supporting Information Figure S10), while OH-dC (with low V0 but more negative ΔE) can coordinate with H2O2 via its additional OH group ( Supporting Information Figure S11). Therefore, we computed H2O2 binding with 5-Aza-2′-dC and OH-dC on their secondary binding face. The H2O2-binding energy of 5-Aza-2′-dC N3-face (−10.7 kcal·mol−1) is only 1.1-fold of its N5-face (−9.5 kcal·mol−1, Supporting Information Figure S10c), while the H2O2-binding energy of OH-dC primary binding face (−11.2 kcal·mol−1) is 2.1-fold of its secondary binding face (−5.4 kcal·mol−1, Supporting Information Figure S11a). The near equivalence of the two 5-Aza-2′-dC faces explains why it is more catalytically proficient than the nonequivalent OH-dC. In addition, the OH group of OH-dC can be involved in an extended H-bond network with surrounding H2O molecules (Figure 5b and Supporting Information Figures S11b and S11c). We computed the influence of additional H bonds with three H2O molecules (two as H-bond acceptor, one as donor) on the N3 electron density. The additional H bonds resulted in the change of QVDD on N3 atom from −296 to −289 me− ( Supporting Information Figures S11c and S11d), which provided a rationale for the lower catalytic proficiency of F3T-hC. Figure 5 | Schematic representation of the double coordination for (a) 5-Aza-2′-dC in G4-DNAzyme with H2O2 and (b) the hydrogen bonds formed between OH-dC in G4-DNAzyme and H2O. Details are shown in Supporting Information Figures S10 and S11. Blue spheres represent Fe(III). Download figure Download PowerPoint From a theoretical point of view, these results indicate that calculations for geometric optimization and binding energy must consider possible structural bias (e.g., nucleobase rotation) and environmental effects (e.g., hydration) to provide more accurate predictive models for DNAzyme catalytic cycles. From an experimental point of view, they cast a bright light on the performances of F3T-azaC, which brings DNA catalysts ever closer to hemoenzymes thanks to a quite unique, double, and synergistic H2O2-coordination site. Overall, we designed the experiments based on theoretical calculations, then obtained a highly active G4-based DNAzyme, and finally continued to verify the construction mechanism for our highly proficient enzymes through the push–pull mechanism. The HRP mechanism has been thoroughly studied, and the central role of two histidine (His) residues, His42 and His170, in the hemin-binding site has been found to synergistically interact with hemin to perform catalytic peroxidation reactions ( Supporting Information Figure S1).41–44 These His, located on both sides of the hemin, activate the cofactor through a push–pull mechanism. The push effect is provided by proximal His170 via direct coordination to the hemin iron atom, and the pull effect is mediated by distal His42 via interaction with the oxidant H2O2. In G4-DNAzymes, the G-quartet on which the hemin stacks plays the role of His170, via a mechanism that is not yet fully understood,30 while the role of His42 can be played by the proximal dCs (Figure 6). Here, we studied dCs as His42 surrogates, and the influence of their electron density on the catalytic activity of the resulting G4-DNAzyme systems. The electron density could be controlled via introduction of various dC derivatives as either electron-donating or electron-withdrawing groups. We demonstrated that the proximal dC with higher electron density led to higher catalytic activity of the G4-DNAzyme. The G4-DNAzymes (F3T-mC and F3T-azaC) containing two dC derivatives, Me-dC and 5-Aza-2′-dC, provided better catalytic performances than F3TC. Interestingly, these two G4s are representative examples exhibiting the intricate relationship between experiments and calculations: the former, F3T-mC, provided a linear correlation between predictions and results, with slightly more stable ΔE than F3TC that led to slightly better catalytic performances. This originated in the simple structure of Me-dC. The latter, F3T-azaC, highlighted that care must be taken in trying to recapitulate experimental conditions in silico, owing to possible structural and environmental bias. 5-Aza-2′-dC indeed provided multiple H2O2 anchoring points that bolstered its catalytic performances, for which a second round of computation could be readily accounted. Figure 6 | Schematic representation of the catalytic cycle of G4-DNAzyme for the oxidation of ABTS to ABTS·+ by H2O2. Blue and red spheres represent Fe(III) and Fe(IV), respectively. Download figure Download PowerPoint Conclusion Globally speaking, we found that dC electron density strongly impacted G4-DNAzyme catalysis, being directly involved in and fostering the formation of Compound 1. Thus, these results confirmed the key role of the push–pull mechanism on the electron-transfer process, in an HRP-like manner. We showed that a combination of experiment results and theoretical calculations provided insights into the fine mechanistic details of G4-DNAzyme inner workings. Each step of the catalytic cycle could be reliably computed, which opened broad perspectives as for the computer-aided design of ever more efficient