A Non‐G‐Quadruplex Hemin Aptamer Forms a Better Peroxidase Mimicking DNAzyme

Abstract

G‐quadruplex DNA is known to bind to hemin, forming a complex that exhibits peroxidase‐like activity. A non‐G‐quadruplex aptamer named Hem1‐2T also exhibits horseradish peroxidase (HRP) like activity upon binding to hemin. Herein, the catalytic characteristics of the Hem1‐2T aptamer are studied and compared with PS2.M, an extensively studied G‐quadruplex. From pH 6–8, the activity of Hem1‐2T decreases with the increase in pH, which is similar to HRP, whereas the activity of PS2.M increases with pH, suggesting that Hem1‐2T might be a better mechanistic mimic of HRP. Additionally, Hem1‐2T is more effective at protecting hemin from degradation by H₂O₂, as evidenced by a slower decrease in the absorbance at 404 nm compared to PS2.M and more sustained catalysis. NMR spectroscopy indicates that hemin promotes ligand‐induced structure formation in the Hem1‐2T aptamer and forms a specific complex, whereas hemin interacts with the PS2.M G‐quadruplex in a way leading to the disappearance of NMR peaks. Overall, the Hem1‐2T‐hemin complex is a better and more stable HRP mimic, supporting its potential applications in bioanalysis and biocatalysis.

A non‐G‐quadruplex aptamer exhibits peroxidase‐like activity when binding to hemin, forming a DNAzyme. This DNAzyme displays a pH‐rate profile similar to horseradish peroxidase, whereas a G‐quadruplex‐based DNAzyme shows an opposite trend. These two aptamers also show distinct hemin‐dependent nuclear magnetic resonance spectroscopy behaviors.

1. Introduction

Peroxidases catalyze H₂O₂‐dependent oxidation of various electron donor substrates, and they play a crucial role in intracellular oxidation.^{[ 1 , 2 ]} Natural peroxidases are heme proteins with Fe(III) protoporphyrin IX (hemin) as a prosthetic group. They help cleave the O—O bond in peroxides, generating four intermediates (compounds I–IV) and stabilizing higher oxidation states of the heme iron during the reaction.^{[ 2 , 3 ]} Due to their versatility in reacting with different substrates, rapid reaction kinetics, and mild reaction conditions, peroxidases have become a valuable tool in bioanalytical chemistry and have numerous applications in biosensors, agriculture and food research, and environmental monitoring.^{[ 4} , ^{5 – 6 ]} However, protein‐based peroxidases are susceptible to stability issues and are expensive to produce, prompting the search for peroxidase mimics. Using catalytic DNA (DNAzymes) with peroxidase‐like activities started with a porphyrin‐binding aptamer isolated by the Sen group.^{[ 7 ]} DNAzymes are not only more affordable and more stable than their protein counterparts but also more versatile in terms of sequence design and incorporation into other functional DNA for applications.^{[ 8} , ^{9 – 10 ]}

Most DNAzymes are known for cleaving, ligating, or phosphorylating nucleic acids, whereas peroxidase DNAzymes do not use a nucleic acid substrate.^{[ 11} , ^{12 – 13 ]} Previously reported peroxidase‐mimicking DNAzymes are guanine‐rich sequences forming G‐quadruplex (G4) structures using hemin as a cofactor.^{[ 8 , 14} , ¹⁵ , ^{16 – 17 ]} The first G4 peroxidase DNAzymes^{[ 18 ]} were derived from the selection of aptamers binding to a porphyrin. Later, many G4 DNAs were found to effectively bind hemin,^{[ 19 , 20 ]} also forming horseradish peroxidase (HRP) mimics.^{[ 21 , 22 ]} A notable application of such DNAzymes is the replacement of HRP as an enzyme label in DNA‐based biosensors.^{[ 23 , 24 ]} However, G4 DNA can bind many planar molecules and thus they lack specificity, and hemin degradation in the presence of G4 DNA also presents challenges for analytical applications.^{[ 25 ]}

Recently, our lab employed the capture‐SELEX method and selected a new hemin binding aptamer with a dissociation constant (K _d ) of 43 nM as measured using isothermal titration calorimetry (ITC), and confirmed that it does not form a G4 structure by analyzing its base composition, salt‐dependent binding, and circular dichroism (CD) spectrum.^{[ 26 ]} This new aptamer also demonstrated peroxidase‐like activity upon binding to hemin. Notably, its activity was high in the presence of Li⁺ instead of K⁺. A systematic comparative study of its analytical applications showed that this non‐G4 aptamer outperforms the G4 aptamer.^{[ 27 ]}

To further advance its applications, fundamental biochemical studies of this new peroxidase DNAzyme are needed. Here, we studied the new aptamer and examined its pH‐dependent catalytic activity. Our results showed that the non‐G4 aptamer is a better mimic of HRP based on the pH‐rate profile. NMR spectroscopy was used to study the structural changes in this aptamer when binding hemin. To our surprise, many new imino proton peaks emerged upon Hem1‐2T binding to hemin, whereas the peaks of a G4‐based aptamer disappeared upon the addition of hemin.

2. Results and Discussion

2.1. The Two Aptamer‐Based Peroxidase‐Mimicking DNAzymes

The secondary structure of the non‐G4 Hem1‐2T aptamer is shown in Figure  1A , and its two conserved loops are connected by two stem regions. Hemin binding is attributed to these two loop regions. Hem1‐2T binds to hemin and exhibits peroxidase activity that increases with the concentration of Mg²⁺ and Li⁺ ions, but not with Na⁺ or K⁺ ions.^{[ 26 ]} Previous CD spectroscopy studies showed a positive peak at 278 nm, characteristic of B‐type DNA, which remains unchanged with the addition of K⁺ ions.^{[ 26 ]} Recently, Tang et al. investigated the Hem1‐2T aptamer structure by computational studies, suggesting strong binding through van der Waals forces with a larger free energy drop of −20.54 kcalmol^{− 1} compared to a G4 aptamer (−5.99 kcal mol^–1).^{[ 28 ]}

Figure 1.

A) The predicted secondary structure of the Hem1‐2T aptamer. B) A scheme of the parallel G4 structure for the PS2.M aptamer bound to K⁺, and the sequence of PS2.M is also supplied.

In this study, we also included a G4‐based DNAzyme for comparison. While many G4 sequences have peroxidase activity upon binding to hemin, we chose to use the PS2.M DNA first reported by Travascio et al., which exhibits a binding affinity for hemin with a K _d value of 0.28 µM.^{[ 29 ]} PS2.M is known to form an antiparallel G4 structure in Na⁺, and it switches to a parallel G4 in the presence of K⁺ _. ^{[ 30 , 31 ]} Since we used 40 mM K⁺ in our PS2.M assays, a parallel G4 scheme is drawn in Figure 1B to represent PS2.M.

2.2. Opposite pH‐Dependent Peroxidase‐Like Activities

One of the most important biochemical properties of enzymes is their pH‐dependent activities, which can provide insights into proton transfer steps. Research by Travascio et al.^{[ 29 ]} found that the hemin‐PS2.M complex is a more efficient catalyst in the pH range of 6–9, compared to unbound hemin, which has limited activity above pH 4.0. Chromogenic substrates such as 2,2^′‐azino‐bis (3‐ethylbenzothiazoline‐6‐sulfonic acid) (ABTS) are most used for activity assays. ABTS is oxidized to a green color product (ABTS•+) by peroxidase reactions (Figure  2C ).^{[ 32 , 33 ]}

Figure 2.

Kinetic traces of the A) Hem1‐2T DNAzyme and B) PS2.M DNAzyme at different pH values. Catalytic activities of DNA–hemin complex (0.5 µM) were measured in 25 mM sodium phosphate buffer, pH 7.5, using ABTS as a substrate (0.5 mM) and monitored at 420 nm. The reaction was initialized by adding H₂O₂ (0.25 mM). C) A scheme showing ABTS peroxidation catalyzed by a DNAzyme producing a colored radical cation (ABTS•+), which can be measured at 420 nm. D) Reaction rates measured in the first minute for the three peroxidases. Each point on the graph represents the mean and standard deviation of three independent samples. The DNA–hemin complex was incubated during 30 min in each pH, at room temperature, before the catalysis.

To characterize the Hem1‐2T DNAzyme, we conducted a thorough analysis of its enzymatic activity at different pH values using ABTS as a substrate. With the increase of pH, the catalytic activity of the Hem1‐2T‐hemin complex gradually decreased in the range of pH 6–8 (Figure 2A and 2D). This pH‐rate trend is the same as that of HRP (Figure 2D). The absorbance of the product reached 0.7 in 20 min and there is no sign of saturation at pH 6.0, indicating a highly stable DNAzyme allowing sustained catalysis. For HRP, pH can influence the ionization of catalytic residues, such as His42 and Arg38.^{[ 3 , 34 ]} At neutral to alkaline conditions, His42 functions as an acid–base catalyst by accepting a proton from a H₂O₂ and transferring a proton to the other oxygen, which aids the heterolytic cleavage of the O—O bond. Arg38 contributes to lowering the pK _a of His42, enabling it to retain its basic function even at lower pH values.^{[ 35 ]} The optimum pH for HRP for ABTS is 6.0.^{[ 36 ]} At this pH, HRP shows stability and promotes the formation of both ferric and ferrous intermediates during the peroxidation process.^{[ 37 ]} At alkaline pH, Arg38 is deprotonated, accompanied with a conformational change in this protein, resulting in a drastic reduction in catalytic activity.^{[ 34 , 38 ]} Without a detailed structure, the exact pH‐dependent catalytic mechanism in Hem1‐2T cannot be precisely described at this moment, but similar mechanisms might take place.

In contrast, the reaction rate of the PS2.M–hemin complex increased in this pH range (Figure 2B,D), and this trend is consistent with previous publications.^{[ 29 , 39 ]} It is important to note that the highest absorbance reached only 0.25 at 20 min for the PS2.M DNAzyme. For its highest activity condition, pH 8.0, the activity plateaued only after 10 min, despite free substrate molecules still available. This result suggested hemin degradation under the oxidative condition, which was reported previously.^{[ 40 , 41 ]} The absorbance of the pH 8.0 sample for the Hem1‐2T also stopped increasing after 10 min, probably suffering from similar hemin degradation. This simple experiment not only suggested that the Hem1‐2T‐hemin complex is a better mimic of HRP in terms of pH dependency, but also Hem1‐2T and PS2.M have distinct catalytic mechanisms.

The lower overall activity of PS2.M can be attributed to the degradation of hemin at high pH and its lower activity at lower pH values. In contrast, the optimal pH of Hem1‐2T is acidic and hemin remained stable under this condition to allow sustained catalysis. In our experimental systems, the catalytic activities of these two DNAzymes happened to crossover at pH 7.5, and thus most of our subsequent experiments were performed at this pH to have a fair comparison of their other properties.

The above experiments were performed in their favorite salt conditions (e.g. 40 mM K⁺ for PS2.M and 100 mM Li⁺ for Hem1‐2T). We also measured pH‐dependent activities of these two DNAzymes without these salts. While the PS2.M–hemin complex lost its activity at all tested pH values, the Hem1‐2T‐hemin complex still had activities following the same pH‐dependency as that in the high salt buffer change (Figure S1, Supporting Information). So, Hem1‐2T is more tolerant to salt concentration.

2.3. Michaelis–Menten Kinetic Comparison

To further compare their catalytic mechanisms, we then assessed the peroxidase activity of Hem1‐2T and PS2.M by varying the substrate concentration of ABTS, initiating the reaction with the addition of H₂O₂. By applying the Michaelis–Menten equation, the Michaelis constant (K _m ) and the catalytic constant (k _cat) were calculated to evaluate their substrate binding affinity and catalytic efficiency. The K _m of the Hem1‐2T‐hemin DNAzyme for ABTS is 69 ± 6 µM (Figure  3A ), in comparison with the affinity of PS2.M for ABTS is 122 ± 5 µM (Figure 3B). Therefore, ABTS has a slightly higher affinity to the Hem1‐2T‐hemin complex than to PS2.M, although such a twofold difference in K _m for different DNAzymes is not very significant. The maximum initial reaction velocity (V _max) is similar for both Hem1‐2T and PS2.M, with values of 4.5 ± 0.2 and 4.4 ± 0.1 µM min⁻¹, respectively. By dividing V _max by the DNAzyme concentrations, the k _cat for Hem1‐2T‐hemin is 8.2 min⁻¹, while for PS2.M‐hemin, it is 9.0 min⁻¹. These similar k _cat values are expected since we carried out this reaction at pH 7.5, where these two DNAzymes have similar activities. A more important conclusion here is that both DNAzymes can fit the Michaelis–Menten equation, confirming their enzyme‐like catalysis.

Figure 3.

The Michaelis–Menten fitting of the two DNAzymes against A) ABTS and B) H₂O₂. The experiments were conducted in 25 mM sodium phosphate buffer, pH 7.5. The DNA‐hemin complexes were incubated for 30 min at room temperature at pH 7.5 before the experiments started. The Hem1‐2T‐hemin complex (0.5 µM) was tested in the presence of 100 mM LiCl, while the PS2.M–hemin complex (0.5 µM) was studied in 40 mM KCl. In all experiments, the activity of the DNA–hemin complex was initiated with the addition of H₂O₂.

We then measured the catalytic constants of DNAzymes at various concentrations of H₂O₂ (Figure 3C,D). The affinity constant (K _m ) of the Hem1‐2T DNAzyme (490 ± 15 µM) was about half that of PS2.M (1003 ± 19 µM), indicating that Hem1‐2T also has a stronger affinity for H₂O₂. In contrast, the V _max for the PS2.M DNAzyme (25.2 µM min⁻¹) was higher, suggesting that with sufficient H₂O₂, the PS2.M DNAzyme can achieve a higher initial catalytic rate. The V _max for the Hem1‐2T DNAzyme was 13 µM min⁻¹. In terms of catalytic efficiency (k _cat), the Hem1‐2T‐hemin complex has a k _cat of 25.6 min⁻¹, while the PS2.M–hemin complex exhibits a k _cat of 49.6 min⁻¹, consistent with the trend of their V _max values. For practical applications, however, H₂O₂ concentration can be a limiting factor in some cases, such as for intracellular applications. In addition, a higher H₂O₂ concentration also leads to more rapid hemin degradation.^{[ 20 , 42 ]} Overall, the Hem1‐2T DNAzyme showed superior performance compared to the PS2.M DNAzyme in low H₂O₂ concentrations.

2.4. UV–Vis Spectroscopic Studies of Reaction Intermediates

The catalytic function of G4‐based DNAzymes was proposed to occur in three primary stages, mirroring that of HRP.^{[ 6 , 22 , 29 , 43 ]} In the initial phase, H₂O₂ binds to hemin Fe(III), breaking the O—O bond and producing an intermediate known as compound I. This intermediate contains a ferryl oxo species (Fe(IV)=O) and a porphyrin cation radical. In the second step, compound I is reduced to compound II. During this process, the ferryl hemin (Fe(IV)=O) receives two electrons from a substrate, such as ABTS²⁻, returning it to its original form as hemin Fe(III). While both compound I and compound II contain Fe(IV), compound II does not have the porphyrin radical. Compound II can oxidize another substrate and return to the resting state (Figure  4 ). Compounds I and II display distinct UV–visible absorption spectra. For compound I, HRP has a reduction of Soret band.^{[ 44 , 45 ]} In contrast, compound II features a Soret band with a slightly higher peak intensity and presents a noticeably different spectrum.^{[ 32 , 46 ]}

Figure 4.

The proposed catalytic cycle of the DNAzymes.

Travascio et al. showed that PS2.M‐hemin produces an intermediate that leads to the creation of compound I, by an immediate decrease in the absorption peaks at 500 and 630 nm. In our work here, the UV–vis spectra of hemin complexed with Hem1‐2T and PS2.M were examined with the introduction of H₂O₂. The absorbance was registered every minute for 4 min (Figure  5 ).

Figure 5.

The Soret absorption band of 10 µM hemin mixed with 10 µM A) Hem1‐2T and C) PS2.M in their respective buffers and then 1 mM H₂O₂ was added. The Q band for the B) Hem1‐2T and D) PS2.M samples. Inset: the arrows indicate the wavelengths at which changes in absorbance are observed.

The initial Soret peak of the Hemin1‐2T‐hemin complex was at 404 nm (Figure 5A), which is similar to that reported for HRP (402 nm).^{[ 47 ]} In the presence of H₂O₂, this system largely retains the peak height and position, suggesting high stability. When examining the PS2.M and hemin complex, its initial Soret band was also at 402 nm (Figure 5B), but a drastic decrease in the intensity of the Soret band was observed and the observed decrease was attributed to degradation of hemin.

We quantified the rate of hemin degradation by following the decrease of the Soret band. The rate of degradation in the PS2.M system was 0.47 min⁻¹, which was ≈95‐fold faster than that with Hem1‐2T (≈0.0049 min⁻¹) (Figure S2, Supporting Information). It is interesting to note that free hemin degradation by H₂O₂ was minimal (Figure S3, Supporting Information), which was consistent with the literature report.^{[ 48 ]} It was quite striking that rapid hemin degradation was observed in the presence of PS2.M, suggesting that PS2.M acted as a catalyst for hemin degradation. In contrast, hemin was very stable in the presence of Hem1‐2T, suggesting that the way of hemin binding by these two aptamers is different.

We then examined the Q‐band in the 450 to 700 nm region (Figure 5C,D). In our experimental conditions, for both DNAzymes, while we observed a decrease in the 500 and 670 nm regions for both DNAzymes, the Hem1‐2T DNAzyme retained better‐defined peaks after the H₂O₂ treatment. This is consistent with hemin being more resistant to degradation by H₂O₂ in the presence of the Hem1‐2T DNAzyme. The Q‐band change appears like that of a heme μ‐oxo dimer,^{[ 49 , 50 ]} and such spectral changes further support the formation of compound I in both DNAzymes.^{[ 29 ]}

2.5. Structural Characterization of the Aptamers by NMR

To characterize the binding and structural changes of the Hem1‐2T and PS2.M aptamers upon hemin binding, we performed 1‐D ¹H NMR spectroscopy (Figure  6 ). The downfield, imino, protons shown in Figure 6 are indicative of base pair formation. In the absence of hemin, the Hem1‐2T aptamer shows four sharp signals indicating the formation of at least four base pairs. With hemin binding, new peaks appear so that a total of ≈15 signals are present in the bound form, indicating that about 10 new base pairs (both Watson–Crick and non‐wecanonical base pairs) are present in the bound form. This result demonstrates that the Hem1‐2T aptamer undergoes ligand‐induced structure formation, confirming hemin binding.^{[ 51 ]}

Figure 6.

Hemin binding monitored by one‐dimensional 1D ¹H NMR by the A) Hem1‐2T and B) PS2.M DNA aptamers. Shown is the downfield imino region of the spectrum. Numbers on the right‐hand side indicates the molar ratio of aptamer: hemin. Data were acquired at 5 °C in 40 mM KCl, 10 mM sodium phosphate (pH 7.5) in 10% ²H₂O/90% H₂0.

For the PS2.M aptamer the free aptamer shows a small number of imino signals in the non‐Watson–Crick region as well as a hump in the baseline often seen with G‐quadruplex structures.^{[ 52 ]} With the addition of hemin, the NMR signals from the PS2.M aptamer broaden and disappear (Figure 6). We attribute this to the PS2.M aptamer binding hemin in many, likely slightly, different conformations with all the conformations in exchange with each other, leading to the broadening out of the NMR signals due to intermediate exchange on the NMR timescale. Paramagnetic Fe³⁺ induced line broadening might also be more pronounced in the PS2.M aptamer due to the way of stacking‐based binding. This is in contrast to the sharp signals of the Hem1‐2T‐hemin complex, which displays the sharp signals consistent with a single‐bound conformation.

3. Conclusions

Our research indicates that the Hem1‐2T‐hemin complex has the same pH‐dependent activity trend as that of HRP, where raising pH dropped the catalytic activity. In contrast, the PS2.M showed an opposite activity trend in the same pH range. Hemin degradation in the presence of PS2.M is more pronounced at higher pH, leading to low catalyst life for the PS2.M DNAzyme when operated at its optimal pH, which is slightly basic. For the Hem1‐2T DNAzyme, the catalysis can be more sustained at its optimal pH, which is slightly acidic. While the substrate concentration‐dependent kinetic data for both DNAzymes can be fitted using the Michaelis–Menten equation, the Hem1‐2T has a lower K _m value for H₂O₂. UV–vis spectroscopy revealed better protection of hemin by the Hem1‐2T aptamer. Despite hemin having a paramagnetic iron, a well‐structured 1:1 aptamer‐hemin structure forms, as revealed by NMR spectroscopy. In contrast, the NMR signals from the PS2.M disappeared upon the addition of hemin. Overall, Hem1‐2T is a good peroxidase mimic and it has stable and excellent catalytic activity at a slightly acidic pH of 6. This non‐G4 aptamer not only provides an interesting ligand to study hemin binding but can also find useful applications to replace HRP.

4. Experimental Section

4.1.

4.1.1.

Chemicals

Integrated DNA Technologies (Coralville, IA) synthesized DNA sequences. PS2.M: GTGGGTAGGGCGGGTTGG; and Hem1‐2T: ACGACTGATTCCGAGATATACTGTACTGAGATTCCAAAGTCGT. HRP was purchased from Merck‐Sigma (Oakville, ON, Canada). Hemin, 2,2′‐azino‐bis(3‐ethylbenzothiazoline‐6‐sulfonic acid) (ABTS), and reagents for preparing the buffers were obtained from Sigma–Aldrich (MO, USA).

Preparation of DNA‐Hemin Complexes

DNA samples were dissolved in water at a stock concentration of 500 µM. A 100 µM DNA solution was prepared in TE buffer (10 mM Tris, 1 mM EDTA, pH 8.0) to obtain the DNA–hemin complex. This solution was incubated at 95 °C for 5 min. After that, the same volume of buffer was added. This buffer contained 50 mM HEPES, pH 7.5, 200 mM NaCl, 1% Triton X‐100, 2% DMSO, and 80 mM KCl (for the PS2.M system) or 200 mM LiCl (for the Hem1‐2T system). In this solution, each DNA is at a concentration of 50 µM. The solution was then incubated at room temperature for 40 min. Following this, an equal volume of hemin solution was added and incubated at room temperature for 2 h. The hemin solution was prepared at a concentration of 5 mM in dimethyl sulfoxide (DMSO) and stored at −20 °C, protected from light. From the hemin stock in DMSO, a 50 µM solution was made in 25 mM HEPES buffer, pH 7.5, 100 mM NaCl, 0.5% Triton X‐100, and 1% DMSO, with an appropriate cation (40 mM KCl or 100 mM LiCl). This final solution was prepared 60 min before mixing with DNA in a 1:1 volume ratio.

Activity Assays at Different pH Levels

Peroxidase activity assays were conducted at room temperature in a 96‐well plate and monitored using a Tecan Spark microplate reader at 420 nm. The buffer for measuring enzyme activity consisted of 25 mM sodium phosphate buffer, 0.05% Triton X‐100, 20 mM cation, and 100 mM NaCl at pH levels of 6.0, 6.5, 7.0, 7.5, and 8.0. The concentration of the DNA–hemin complex was set at 0.5 µM. After 30 min of incubation at room temperature, 0.5 mM ABTS was added as a substrate, and the reaction was initiated with 0.25 mM H₂O₂. The final reaction volume was 100 µL.

Hemin Degradation Kinetics

To analyze the decay kinetics of the DNA–hemin complex, the concentration of the complexes was set at 5 µM in the presence of 2 mM H₂O₂, in 25 mM sodium phosphate buffer, 0.05% Triton X‐100, 20 mM cation, and 100 mM NaCl (pH 7.5), with a final volume of 100 µL. The decay was monitored at an absorbance of 404 nm in 96‐well plates at room temperature using a Tecan Spark microplate reader, with an incubation time of 5 min at 25 °C.

UV–Vis Spectroscopic Studies

The UV–vis spectra of each DNAzyme were collected using the Tecan Spark in 96‐well plates. The sample was prepared with 25 mM sodium phosphate buffer, 0.05% Triton X‐100, 20 mM cation, 100 mM NaCl, DNAzyme (10 µM), and 1 mM H₂O₂, incubated for 5 min at 25 °C. The UV–vis spectra were monitored every minute within a 400–700 nm wavelength range.

NMR Spectroscopy

NMR experiments on the Hem1‐2T and PS2.M DNA aptamer samples were performed using a 600 MHz Bruker Avance spectrometer in a similar manner to what we have previously described.^{[ 53 ]} Experiments were performed with an aptamer concentration of 0.5 mM in an aqueous solution of 10 mM sodium phosphate (pH 7.5), 40 mM KCl, 10% D₂O at 5 °C. Water suppression was achieved using excitation sculpting.^{[ 54 ]} A stock solution of 30 mM Hemin was prepared by dissolving in DMSO‐d₆ and 1% Triton‐X. The stock was kept cool and away from light.

Statistical Analysis

The data are expressed as the mean ± SD of at least three different experiments with triplicate values, and the analysis was performed by the software Origin Pro (2025). Differences were considered statistically significant at p values less than 0.05.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

Claudia Rodríguez‐Almazán performed the kinetics experiments. Yunus A. Kaiyum and Philip E. Johnson performed NMR experiments. Claudia Rodríguez‐Almazán, Yunus A. Kaiyum, Philip E. Johnson, and Juewen Liu conceived and designed the experiments, analyzed the data, and wrote and critically reviewed the paper.

Supporting information

Supplementary Material

Rodríguez‐Almazán Claudia, Kaiyum Yunus A., Johnson Philip E., Liu Juewen, ChemBioChem 2025, 26, e202500486. 10.1002/cbic.202500486

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.