Engineering ligands for theophylline riboswitches expands its regulatory dynamic range in prokaryotic and eukaryotic systems

Domain: chemistry
Published: 2026-03-23
Editorial outcome: minor_revision

Abstract
The theophylline riboswitch has been a foundational tool in synthetic biology for three decades, yet its regulatory performance remains constrained by the modest affinity of its native ligand. Enhancing the dynamic range of riboswitches is critical for precise gene regulation in biotechnological applications. Here, we show that synthetic 4-quinazolinone derivatives, designed through a structure-based approach, are significantly better than theophylline in both binding and functional activation across multiple biological systems. We demonstrate that these derivatives bind the theophylline aptamer with up to 30-fold higher affinity, thereby expanding regulatory performance. In the bacterial system, these ligands enhance "ON" gene expression by up to 380-fold, compared to 75-fold with theophylline. This superior control extends to diverse organisms; in mycobacteria, the activation ratio reached 20-fold, and in eukaryotes, expression increased 11-fold. Furthermore, in riboswitch-mediated conditional CRISPR-Cas9 applications, these ligands achieve 70% genome editing efficiency at 10-fold lower concentrations than theophylline. These results demonstrate that ligand optimization is a crucial driver for enhancing riboswitch performance for advanced biomedical engineering.

Introduction
Synthetic biology allows the engineering of biological systems for diverse applications, ranging from the development of novel diagnostic tools1,2 and therapeutics3,4 to the biomanufacturing of drugs5, fuels6, and materials7,8, and bioremediation9,10. The precise induction of gene expression is a key prerequisite for programming the desired cellular function. Therefore, the majority of efforts in synthetic biology have concentrated on developing programmable gene regulatory tools that can work across all forms of life11,12. Ribonucleic acid (RNA) plays a central and multifaceted role in all living organisms, ranging from serving as a template for protein synthesis to influencing gene expression to regulate cell physiology, differentiation, and development13. Among the diverse regulatory RNAs, synthetic riboswitches have emerged as a promising tool for protein-independent spatial, temporal, and ligand dose-dependent gene regulation14. Riboswitches are naturally structured RNA elements found in bacteria15,16 and eukaryotes17,18 that regulate gene expression in response to metabolite binding. A riboswitch composed of two domains: an aptamer domain, which specifically binds to a diverse set of cellular metabolites and induces a conformational change in the adjacent expression platform to alter gene expression at transcription, translation, or mRNA decay19-21.

Researchers have created synthetic counterparts of riboswitches as gene regulatory tools for rewiring cells for therapeutic and synthetic biology applications, given their decreased immunogenicity, smaller genetic footprints, dose-dependent, cis-regulation, and modular characteristics22-26. Synthetic riboswitch-mediated gene regulation has been employed in prokaryotes and eukaryotes for diverse applications, for instance, in enzyme evolution27, metabolic engineering28-31, biosensing32-37, therapeutics38-40, diagnostics2,41,42, etc, highlighting their potential in precision medicine and therapeutic interventions. Synthetic riboswitches have been generally created by fusing the aptamer domain to a natural riboswitch expression platform. Similar to natural riboswitches, the aptamer domain and its cognate ligand are essential building blocks required to develop synthetic riboswitches43,44. To create synthetic riboswitches, the aptamer-ligand pairs are obtained either from SELEX or natural riboswitch aptamer engineering30,45,46. Despite promising, the widespread application of synthetic riboswitches is currently limited by the scarcity of suitable aptamer-ligand pairs47. The existing aptamer-ligand repertoire is primarily limited to a few ligands, such as theophylline48, tetracycline49, paromomycin50, tobramycin51, neomycin52, etc. most of which exhibits toxicity, low affinity, and low dynamic range, requiring high ligand concentrations and restricting their utility for biomedical applications53. One approach to overcome the aforementioned challenges is to create improved ligands for current aptamer domains or entirely novel aptamer-ligand combinations with improved functional characteristics such as high affinity, low toxicity, and high cellular penetration.

For over thirty years, the synthetic riboswitches created have predominantly relied on theophylline aptamer as a sensory domain54. Theophylline aptamers have proven particularly useful across a range of fields, including synthetic biology, diagnostics, therapeutics, and bioremediation. Despite theophylline riboswitch’s wider use, its inducer ligand, theophylline (1,3-dimethylxanthine), is a bronchodilator drug used in clinics and has a narrow therapeutic index55. The higher theophylline levels side effects range from seizures to life-threatening complications56. Furthermore, it is required at a very high concentration, 2-5 mM range, for riboswitch activation, has a low induction range, and low cellular penetration47. From all these observations, we envisioned that developing an improved ligand for the theophylline aptamer with high affinity, low toxicity, and improved cellular penetration could significantly improve the usability and performance of the theophylline riboswitch in synthetic biology. Notably, with such a ligand, we could use the current theophylline-based synthetic riboswitches without changing their sequence or time-consuming optimization of expression platforms required for new riboswitch development. The limitations of the theophylline ligand would be solved by a new ligand; thus, it will greatly increase the practicality of theophylline riboswitches for real-world applications.

Here, we employ a structure-based design approach to develop ligands that bind the theophylline aptamer with greater affinity than the natural theophylline ligand. Rational design of small molecules to target RNA secondary and tertiary structure for biomedical applications is a promising approach57,58. We45,59 and others60,61 have used a structure-based rational design approach to develop small molecules targeting aptamer domains derived from natural riboswitches or from in vitro selection, respectively. Although this strategy can help identify high-affinity aptamer binders, a riboswitch function requires more than just high-affinity binding. For riboswitch function, ligand binding must induce a conformational shift in the downstream expression platform. To address this, we not only develop high-affinity molecules for theophylline aptamers but also test their ability to serve as a riboswitch inducer in both prokaryotic and eukaryotic systems. We therefore report on the development of 4-quinazolinone derivatives, a class of synthetic ligands designed using a structure-based approach that binds the theophylline aptamer with higher affinity than the native theophylline ligand, exhibits improved cellular penetration, and provides a broader dynamic range in both prokaryotic and eukaryotic systems.

Results
Structure-based virtual screening:
The theophylline aptamer (Th_APT), a well-characterized 33-nucleotide synthetic aptamer, adopts a single hairpin secondary structure that creates a flexible binding pocket for theophylline (Theo) through tertiary interactions (Fig. 1a,b)58,62,63. To identify the compounds with potential binding affinity for the Th_APT, we conducted structure-based virtual screening using a comprehensive library of approximately 1 million compounds sourced from PubChem, ZINC, and DrugBank databases. The crystal structure of the Th_APT in complex with Theo (PDB ID: 8D28) served as the template for all docking experiments (Fig. 1b). Initial validation was performed using the native ligand Theo to optimize docking parameters. The computational analysis revealed that Theo establishes hydrogen bond interactions with nucleotides C-22 and U-24, with these interactions being stabilized by two nucleotide stacks: an upper stack comprising A7-C8-G26 and a lower stack consisting of U6-U23-A28 (Fig. 1c). The binding mode predicted by our docking protocol accurately reproduced the interaction pattern observed in the experimental crystal structure58, confirming the reliability of our computational approach.

After protocol validation, we used molecular docking to screen the compound libraries. Analysis of the top 50 compounds revealed a predominant structural motif: 4-(1H)-quinazolinone as the most frequently occurring scaffold among the high-scoring hits (Supplementary Table 1). Based on synthetic accessibility considerations for subsequent hit optimization, we synthesized a focused chemical library comprising 4-(1H)-quinazolinone analogues (Fig. 1e). Molecular docking of these analogues against the Th_APT demonstrated that all designed ligands exhibited binding interactions like those of Theo (Fig. 1d). The binding scores for the developed ligands ranged from -4.11 to -9.05 kcal/mol (Supplementary Table 2), indicating favourable predicted binding affinities comparable to or exceeding that of the native ligand.

In-vitro validation of aptamer ligand interactions for the hits generated from in-silico analysis:
The interaction between synthesized 4-(1H)-quinazolinone analogues and Th_APT was validated using a label-free fluorescence assay with the Thioflavin T (ThT)64. The ThT is a fluorescent dye that exhibits enhanced fluorescence intensity when intercalated into the bulges and loops of RNA secondary structures65(Supplementary Fig. 1a). The Th_APT adopts a hairpin conformation containing two bul ges and a terminal loop, and upon binding of Theo or cognate ligands, these bulges undergo conformational changes to accommodate the ligand, resulting in ThT displacement and subsequent fluorescence reduction (Supplementary Fig. 1a, b). To verify that the observed ThT fluorescence decrease was due to specific ligand-aptamer interactions, a mutant aptamer (Th_APTM) was created by mutating nucleotide residues involved in binding-pocket stabilization via stacking interactions. The introduced mutations were A5C, U6G, A7U, C8U, G26U, and A28U (Fig. 1a). Incubation of Theo with Th_APTM yielded no significant change in ThT fluorescence (Fig. 2a), validating that the signal reduction observed with the wild-type aptamer originates from a specific binding rather than non-specific interference. Fluorescence binding experiments were conducted by incubating both Th_APT and Th_APTM with ThT dye in the presence of 100 μM of each test ligand. Significant fluorescence reduction was observed when Th_APT was treated with Theo, 7ClB, 7MB, 6HB, 7HB, 6MeB, 7MeB, NCB, NFB, HMB, NMB, AMB, and EMB. In contrast, ligands B0, 6NB, 7NB, DHB, and MBB showed no significant fluorescence changes. Caffeine (Caff), differing from theophylline with a single methyl group, was used as a negative control and, consistent with previous reports of its lack of interaction with Th_APT, showed no binding activity as expected. Importantly, Th_APTM showed no fluorescence changes with any of the tested ligands, further supporting the specificity of the binding interactions observed with Th_APT (Fig. 2a). Further, we calculated the binding affinities of compounds with Th_APT using the ThT-based fluorescence assay. Ligands were titrated with a series of two-fold dilutions, and the normalized fluorescence values were plotted against ligand concentrations, with the fluorescence measured at 0 µM ligand concentration defined as 100 %. The fitted K values were calculated and summarized in Supplementary Table 2 (Fig. 2b-e and Supplementary Fig. 1c-k). Among the ligands tested, compounds HMB and NMB exhibited K values of 0.94 µM and 0.88 µM, respectively, which are significantly lower than that of Theo (3.34 µM). Based on this, these compounds were used for further characterizations.

Subsequently, isothermal titration calorimetry (ITC) was performed to determine the thermodynamic properties of binding of the developed ligands, HMB and NMB, alongside Theo and Caff, using Th_APT and Th_APTM. Representative ITC thermograms are presented in Fig. 2f- i, and Supplementary Fig. 2, and Table 1 provides the dissociation constant (K ), along with enthalpy changes (ΔH° ), entropy (ΔS° ), and free energy (ΔG° ) parameters of binding at 25 °C. The binding was observed for Theo, with an estimated K of 0.919 ± 0.015 µM, consistent with prior reports, whereas Caff showed no detectable interaction with Th_APT. The K values for HMB and NMB are 0.096 ± 0.002 µM and 0.031 ± 0.006 µM, respectively. A 9-fold and 30-fold decrease in K values was observed for HMB and NMB, suggesting an improved binding affinity of these compounds towards the aptamer compared to Theo. These results are consistent with those from the ThT-based fluorescence assay. All four ligands are titrated against the Th_APTM, and none of the four ligands interact with Th_APTM, suggesting the specificity of ligand-aptamer interaction (Supplementary Fig. 2).

The favorable enthalpy changes were observed for all Theo, HMRB, and NMB (Table 1). These favorable enthalpy changes are consistent with our in silico docking analysis, which shows that the N1 and N3 positions of the HMB and NMB scaffolds form hydrogen bonds with the U24 and C22 residues in the aptamer, respectively. The unfavorable TΔS values (Table 1) indicate that the aptamer adopts a more ordered structure upon ligand binding66. In the case of Th_APT, the binding site is less organized in the absence of a ligand. In contrast, ligand binding to the residue C27 is flipped out of the pocket, allowing residues C22, U23, and U24 to contribute to an S-turn formation and stacked base triples that form the core of the Theo binding site58. These structural changes contribute significantly to opposing entropy values (Table 1).

The structure-activity relationship analysis of data from fluorescence and ITC experiments revealed key determinants of aptamer-ligand binding. In silico molecular docking studies indicated hydrogen bonding between N1 and N3 of all quinazolinone derivatives and the U24 and C22 residues, respectively. The in silico and in vitro ThT-based fluorescence-binding screening results showed mixed concordance. Some computational hits show binding to theophylline aptamer, yet some compounds predicted to bind did not interact with the aptamer. The unsubstituted 4-(1H)-quinazolinone core (B ), as well as NO₂ substitution at either the 6 (6NB) or 7 (7NB) position, 0 did not exhibit detectable binding. In contrast, OH substitution at position 6 (6HB) or 7 (7HB) resulted in weak but measurable interactions, with binding affinities of 11.10 and 10.26 µM, respectively. Interestingly, simultaneous OH substitution at both the 6 and 7 positions led to complete loss of binding. The lack of interaction observed for NO₂-substituted derivatives may be attributed to reduced π-electron density caused by the strong electron-withdrawing nature of the NO₂ group. Although OH substitution generally increases π-electron density, its strong hydrogen-bonding propensity may induce localized π-electron depletion, thereby diminishing binding interactions67.

Substitution with a CH₃ group at position 6 enhanced binding affinity, with a pronounced position-dependent effect observed when CH₃ was introduced at position 7, resulting in substantially higher affinity compared to substitution at position 6. Similarly, substitution with Cl (7ClB) or –OCH₃ at position 7 (7MB) produced a marked increase in binding affinity relative to the native ligand, theophylline (Supplementary Table 2). This enhancement is likely due to the dual electrostatic nature of these substituents, which facilitates favorable π–π stacking interactions with the nucleobases of the aptamer, indicating that electron-withdrawing substituents (NO ) at these positions may have a negative effect on aptamer ligand interactions68,69. Further, we substituted OH (HMB), NO (NMB), and NH (AMB) at position 6, while keeping -OCH3 at position 7. The data from ThT experiments showed a trend of increasing affinities in the order OH (HMB) > NO (NMB) > NH (AMB) (Table S2). This can be correlated with superior resonance stabilization and reactivity of OH compared to less stabilizing NO₂ and NH₂, respectively 70. Replacement of the -OCH from the 7th position in NMB with halogens Cl (NCB) and F (NFB) showed a decrease in binding affinities in the order –OCH₃ (NMB) > Cl (NCB) > F (NFB) (Table S2). The moderate binding affinities observed with Cl may be due to its polarizable electron cloud71, while a significant decrease in affinity with F might be because of the strong electron-withdrawing nature and low polarizability71.

Characterization of ligand-induced conformational changes using structural probing:
Structural alterations in the aptamer upon ligand binding were investigated using RNase T1 and in-line probing assays. RNase T1, which cleaves specifically single-stranded RNA at the 3' end of guanosine residues, was employed to generate cleavage patterns that reveal accessible versus protected regions within the RNA structure, using RNA labelled at the 3’ end with Cy5 (Fig. 3a upper right panel). The T1 digestion analysis demonstrated distinct cleavage patterns at specific nucleotide positions in the presence of ligands (Supplementary Fig. 3). Ligand binding enhanced cleavage at G14, indicating increased nucleotide accessibility, while G19 was protected, reflecting structural restriction. HMB and NMB showed similar patterns but with stronger cleavage at G14 and greater protection at G19 compared to Theo. The greater magnitude of these band-intensity changes indicates higher binding affinities for HMB and NMB than for Theo, consistent with the ITC data.

To complement the RNase T1 analysis, in-line probing was performed using the same 3’-Cy5-labelled RNA to monitor spontaneous RNA cleavage patterns. This approach allows for the assessment of ligand-mediated conformational changes in a non-enzymatic context, effectively eliminating the risk of enzymatic interference. In-line probing revealed similar modulation patterns in band intensities upon exposure of the aptamer to Theo (Fig. 3a), HMB (Fig. 3b), and NMB (Supplementary Fig. 4a). The changes in cleavage susceptibility at specific nucleotide positions indicated regions of the aptamer that undergo structural reorganization upon ligand binding. Across all three ligands, we observed consistent reductions in cleavage susceptibility at nearly identical nucleotide positions. This high degree of overlap in the protection patterns suggests that Theo, HMB, and NMB occupy a common binding pocket within the aptamer. Notably, the band intensities corresponding to nucleotide positions C22, U23, U24, and G25 decrease upon an increase in ligand concentration, suggesting protection of these residues in the presence of the ligand. Molecular docking revealed that the nucleotides C22 and U24 form hydrogen bonds with all three ligands (Fig. 3c-d and Supplementary Fig. 4b), consistent with previous studies for Theo58,62,72. Ligand interaction at positions C22 and U24 induces structural compaction, promoting S-turn formation by nucleotides C21 to G25 (Fig. 3a), which transitions from an initially unorganized state58 where these nucleotides are more accessible for cleavage. Titration experiments conducted with increasing concentrations of each ligand demonstrated that these conformational changes were dose-dependent (Fig. 3a- b, and Supplementary Fig. 4c). As ligand concentrations increased, we observed proportional changes in band intensities at key nucleotide positions (Fig. 3c-d and Supplementary Fig. 4). This dose-response relationship provides evidence for specific binding interactions between the aptamer and each of the tested ligands. While Theo, HMB, and NMB all induce structural changes in the aptamer at similar nucleotide residues, the magnitude of these changes varies between ligands (Fig. 3c-d and Supplementary Fig. 4c).

The ligand HMB showed significantly improved binding affinities towards the aptamer. To further investigate the kinetic parameters of ligand binding, we performed surface plasmon resonance (SPR) analysis of HMB and Theo in the presence of Th_APT. Representative sensograms for Theo and HMB are shown in Fig. 3e and 3f, respectively. The kinetic data obtained from the SPR analysis showed that HMB interacts with an aptamer with an association rate (k ) of 15.89 ± 4.090 M-1S-1 and dissociation rate (k ) of 15.71 ± 2.12 S-1. While Theo showed significantly slow association with k of 5.72 ± 0.060 M-1S-1 and fast dissociation with k of 84.96 ± 1.45 S-1. Overall, HMB showed a 2.78-fold faster association and a 5.41-fold slower dissociation than Theo. The increase in k and subsequent decrease in k overall reduced the equilibrium dissociation constant (K ) for the ligands HMB (0.105 ± 0.011 µM), suggesting higher binding affinities than Theo (1.469 ± 0.034 µM).

We further evaluated ligand specificity by performing site-directed mutagenesis on the theophylline aptamer, guided by the protection patterns identified in the in-line probing analysis. The probing data highlighted a critical protection pattern at four consecutive nucleotides (C22–G25) situated within the binding pocket. Upon mutating these residues, the binding affinity for theophylline, HMB, and NMB were abolished, as determined by the ThT-based fluorescence assay (Supplementary Table 3). These results confirm that C22–G25 are functionally indispensable for coordinating the ligand scaffold and that the high-potency ligands HMB and NMB bind with high specificity to the wild-type aptamer.

To confirm that the developed ligands bind specifically to the theophylline aptamer rather than nonspecifically to RNA or common RNA secondary structures, FRET, CD spectroscopy, and CD melting studies were conducted. A dual-labelled HpGQ sequence was used to assess potential ligand-induced conformational switching between hairpin (Hp) and G-quadruplex (GQ). No significant FRET changes were observed relative to the DMSO vehicle, while TPE-MePy, A positive control, showed a significant change in FRET signals73 (Supplementary Fig. 5). CD spectrum and melting analyses showed no conformational shifts or stabilization effects (Supplementary Fig. 6). Together, these results indicate that HMB does not interact detectably with ssRNA, GQ, or Hp structures under the tested conditions.

The biophysical characterization was followed by evaluation of antibacterial activity for all ligands that interacted with Th_APT and cytotoxicity for the potent ligands HMB and NMB, along with Theo, before proceeding to gene regulation experiments. Bacterial growth inhibition studies using E. coli DH10β cultured in M9 medium showed no growth suppression at concentrations reaching 2500 µM (Supplementary Fig. 7). To determine bacterial uptake efficiency, E. coli cultures were exposed to 500 µM of each compound 16 hours before extraction and HPLC quantification. Results demonstrated that HMB and NMB achieved 2.88- and 2.61-fold higher intracellular penetration with values of 9.29 µg/108 CFU and 8.43 µg/108 CFU, respectively, compared to Theo (3.23 µg/108 CFU) (Fig. 3g, Supplementary Fig. 8). Mammalian cell toxicity was assessed using MTT viability assay. All compounds maintained complete cell viability with no observable cytotoxicity (Supplementary Fig. 9a). Cellular uptake analysis revealed that HMB accumulated to approximately 1.89-fold higher with 26.41 ng/105 cells than the concentration of Theo (13.91 ng/105 cells), whereas NMB showed comparable accumulation levels with 13.11 ng/105 cells (Fig. 3h, Supplementary Fig. 9 b-d). These findings demonstrate that Theo, HMB, and NMB exhibit excellent biocompatibility across both prokaryotic and eukaryotic systems at the concentrations evaluated. Notably, HMB and NMB displayed enhanced bacterial uptake, with HMB additionally showing relatively higher accumulation in mammalian cells.

Effect of developed ligands on the activity of riboswitches in E. coli:
To evaluate the ability of developed ligands to control riboswitch-mediated gene regulation, we established an in vivo screening platform. Synthetic riboswitches were engineered, incorporating the Th_APT with diverse upstream and downstream regulatory sequences that varied in both length and nucleotide composition74,75 (Supplementary Fig. 10, 11, and 12). The aptamer was fused to an expression platform so that the ribosome-binding site (RBS) would be sequestered within a stem-loop structure, restricting ribosome access and maintaining the gene expression in ‘OFF’ state. Upon ligand binding, RNA undergoes conformational changes, making RBS accessible for the ribosome, initiating expression of the downstream reporter gene – RFP, turning ‘ON’ the gene expression (Fig. 4a). To validate that the observed reporter gene activation was specifically due to ligand-aptamer interaction; we constructed a mutant riboswitch with Th_APTM (RS-1_M) with a disrupted aptamer binding site. All riboswitch variants were cloned into the 5′ untranslated regions (UTR) of the RFP reporter gene under the control of the J23100 promoter.

The plasmids consisting of developed riboswitches RS-1 to RS-9 and RS-1_M were transformed into E. coli DH10β cells. The transformed bacterial cells were screened for RFP fluorescence in the presence and absence of the developed ligands at 500 µM. All developed riboswitches, RS-1 to RS-9, showed a significant increase in RFP expression in the presence of ligand Theo, HMB, and NMB, while there was no change in RFP expression reported with RS-1_M. Fold change (Activation ratio) in RFP expression was calculated with respect to the DMSO control, where the expression of RFP in the DMSO control was defined as 1-fold. The results obtained showed that ligands can induce the RFP expression in the range of 1.39 to 110.94-fold (Fig. 4b) (Supplementary Table 4). These findings demonstrate that the riboswitch regulatory dynamic range can be optimized through systematic engineering of riboswitch sequences tailored to diverse experimental requirements. Amongst the riboswitches tested, RS-4 yielded a substantially high activation ratio of 110.94-fold at a 500 μM concentration of HMB, so all the further experiments were performed with RS-4 unless mentioned otherwise. Next, all ligands observed to interact with Th_APT in the ThT-based fluorescent assay were screened against RS-4. The ligands HMB and NMB obtained the highest fold change in RFP expression with 109.44 and 83.45-fold, respectively (Fig. 4b and Supplementary Fig. 13a, Table 2). The half-maximal effective concentration (EC ) values were calculated for 12 ligands, which showed an increase in RFP expression in the preliminary screening. The E. coli DH10β cells harboring RS-4 were grown in the presence of ligands at concentrations ranging from 0 to 2500 μM, and changes in RFP fluorescence were monitored. The EC₅₀ values demonstrated considerable variation among ligands, ranging from 733 µM to 51.50 µM (Table 2, Fig. 4c, and Supplementary Fig. 13b-j).

Flow cytometry analysis of bacterial populations transformed with RS-4 supported the fluorescence screening results (Fig. 4d and Supplementary Fig. 14a). Median fluorescence intensity (MFI) measurements demonstrated ligand-dependent increases in RFP expression (Supplementary Fig. 14b), with HMB and NMB yielding markedly higher responses with 378.06-fold and 237.76-fold compared to Theo (76.81-fold) (Fig. 4e). Complementary confocal fluorescence microscopy was employed to visualize ligand-induced RFP expression. Confocal imaging revealed distinct increases in RFP fluorescence intensity upon ligand treatment compared to DMSO vehicle control (Fig. 4f). A moderate increase in the RFP signal was observed upon treatment with Theo, whereas HMB and NMB induced substantially more pronounced fluorescence increase. These microscopic observations correlated strongly with both dose-response studies and flow cytometric quantification.

Western blot analysis was performed to analyze RFP protein accumulation following ligand treatment. Bacterial lysates were prepared from cells treated with test ligands, subjected to SDS-PAGE, and immunoblotted with anti-RFP antibodies. Significant increases in RFP protein levels in the presence of HMB and NMB were observed (Fig. 4g), consistent with the fluorescence-based measurements. To investigate the mechanism of ligand-induced gene regulation, quantitative reverse transcription PCR (qRT-PCR) was performed to measure RFP transcript levels. Notably, no significant differences in RFP transcript abundance were detected between ligand-treated and untreated samples (Fig. 4h). These results indicate that the observed ligand-dependent increases in RFP expression occur through translational, rather than transcriptional, regulation, consistent with the switch's design.

Subsequently, an OFF-type riboswitch (RS-1L0) was engineered utilizing the Th_APT domain, with strategically modified pre- and post-aptameric sequences, to achieve ligand-dependent gene repression. The RS-10 design positions RBS in an accessible conformation under ligand-free conditions, permitting efficient ribosome recruitment and RFP translation. Upon ligand binding, the conformational changes in the aptamer sequester the RBS within a secondary structure, inhibiting ribosome access and reducing protein expression (Fig. 5a). The activity of riboswitch RS-10 was evaluated in the presence of ligands, which showed interactions with Th-APT in the ThT-based assay. Spectrofluorometric analysis revealed a 32.3% reduction in RFP fluorescence upon treatment with Theo, while HMB and NMB demonstrated substantially greater repression efficiencies of 82.2% and 67.64%, respectively (Fig. 5b and Supplementary Fig. 15a, Table 2). The EC₅₀ values were calculated for the 12 ligands obtained from the screening. Non-linear regression analysis yielded EC₅₀ values ranging from 516 μM to 165 μM (Table 2) (Fig. 5c and Supplementary Fig. 15b-j).

Flow cytometric analysis was performed for the two most potent ligands, HMB and NMB, alongside Theo, to corroborate spectrofluorometric findings (Fig. 5d and Supplementary Fig. 16a). Quantitative analysis revealed that HMB and NMB induced maximal reduction in RFP expression with 80.29% and 70.44%, respectively, exceeding Theo-mediated repression with 50.46% (Fig. 5e and Supplementary Fig. 16b). Confocal fluorescence microscopy of RS-10-transformed bacterial cells treated with ligands provided visual confirmation of reduced RFP expression. Substantial decreases in RFP fluorescence intensity were observed following treatment with HMB and NMB, whereas Theo treatment produced minimal detectable changes compared with the vehicle control (Fig. 5f). The imaging results demonstrated excellent concordance with both spectrofluorometric and flow cytometric quantification.

Further, immunoblot analysis confirmed significant reductions in RFP protein levels in samples treated with HMB and NMB compared with Theo and DMSO controls (Fig. 5g). To elucidate the regulatory mechanism, qRT-PCR was performed to assess RFP transcript levels. No significant differences in RFP transcripts were detected among ligand-treated samples or between treated and DMSO control conditions (Fig. 5h), confirming that transcript levels remained constant regardless of ligand treatment. These findings definitively establish that the developed ligands mediate gene repression through post-transcriptional mechanisms, consistent with the designed riboswitch-based translational control system illustrated in Fig. 5a. The collective data demonstrates that the engineered ligands exhibit greater riboswitch-inducing activity than Theo, enabling efficient, ligand-dependent gene repression.

Robustness of developed ligands under various conditions:
We further assessed the robustness of lead molecules HMB and NMB in activating the riboswitch under diverse growth conditions across different E. coli strains, growth media, temperatures, and pH levels. Five E. coli strains (DH5α, TOP10, BL21, M15, and DH10β) were transformed with a plasmid encoding the reporter protein RFP under RS-4 regulation and subsequently treated with Theo, HMB, and NMB. RS-4 demonstrated significant activation across all strains, with RFP expression increases ranging from 30.67-fold to 113.25-fold (Fig. 6a). HMB achieved the highest activation ratio in E. coli M15, though no significant differences were observed between M15, DH10β, and TOP10 strains for HMB treatment.

E. coli BL21 exhibited the lowest fold change, followed by DH5α. Importantly, HMB and NMB consistently showed significantly higher activation ratios than Theo across all strains tested, confirming their superior performance. Further, we have analyzed the activity of the developed ligands in combination with RS-4 across different growth media. The activity analysis revealed that HMB demonstrated significantly higher activation ratios than Theo across all tested conditions, while NMB showed activity comparable to Theo in Terrific broth (TB), Super optimal broth (SOB), and SOB with catabolite repression (SOC). The varying components of growth media can differentially influence organisms' metabolism, potentially altering ligand molecule metabolism76,77. The altered metabolism of ligand molecules may lead to reduced activation ratios under certain conditions. Although all ligands achieved their highest activation ratios in chemically defined M9 medium, HMB consistently showed a significantly higher activation ratio than Theo across every medium tested. These results indicate that HMB can effectively control gene expression regardless of growth medium composition, making it a more versatile and reliable alternative to Theo for genetic regulation applications (Supplementary Fig. 17a).

As temperature plays an essential role in riboswitch function, we assessed ligand activity across different growth temperatures. A positive correlation between temperature and activation ratio was observed, with minimal activation at 16 °C (5.94-fold for Theo) increasing to maximum levels at 42 °C. Theo consistently showed the lowest activation ratios across all temperatures, while HMB achieved the highest fold change (156.68-fold at 42 °C) (Supplementary Fig. 17b). Decreasing basal expression at higher temperatures contributed to enhanced activation ratios (Supplementary Fig. 17c), likely due to temperature-dependent RNA structural mobility75,78,79. To determine the optimal pH for ligand activity, we tested all compounds in M-9 media over a pH range of 6.5 to 8.5. Each ligand showed a different pH level at which it worked best. Theo and HMB were most active at pH 8.0, with fold changes of 52.34 and 156.48, respectively. In contrast, NMB worked best at pH 7.0, reaching a maximum fold change of 116.18 (Fig. 6b). Statistical analysis (Tukey's test) showed that the ligands responded differently to changes in pH. Theo had a wide effective range, staying consistently active between pH 7.0 and 8.5. HMB and NMB were more sensitive to pH changes: HMB showed strong activity only between pH 7.5 and 8.0, while NMB was most effective at pH 7.0-7.5. Next, to confirm that the pH-dependent changes in ligand activity arise from differences in binding affinity rather than physiological effects, we measured the binding affinities of Theo, HMB, and NMB over a pH range of 7.0–8.5. The results identified distinct optimal pH values for binding: pH 8.0 for Theo and HMB, and pH 7.5 for NMB. Representative thermograms are shown in Supplementary Fig. 18, and the corresponding thermodynamic parameters are summarized in Table S5. The ITC data indicate that the pH-dependent ligand activity observed in expression studies is driven by pH-dependent shifts in binding affinity, rather than by physiological changes within the cells. HMB and NMB maintain robust riboswitch activation across multiple bacterial strains while exhibiting distinct pH optima that can guide their application in different experimental contexts. The expanded regulatory dynamic range and pH-dependent gene regulation of theophylline riboswitches by both HMB and NMB inducers compared to Theo support their development as reliable tools for controlled gene expression systems.

Evaluation of developed ligands for riboswitch function in Mycobacterium smegmatis:
The theophylline-responsive riboswitch enables tunable, reversible gene regulation in mycobacteria without exogenous proteins, overcoming traditional systems' high basal activity, recombination, and optimization challenges80,81. However, theophylline's bronchodilator properties may complicate M. tuberculosis infection studies by interfering with the pathophysiology of the disease81. Given these limitations and our previous identification of alternative ligands that have better activity than Theo in E. coli, we investigated these promising compounds in mycobacteria to develop improved gene-regulation tools specifically suited for mycobacterial research applications.

To assess the potential of alternative ligands for improving gene regulation in mycobacteria, we electroporated M. smegmatis with a plasmid containing the theophylline riboswitch controlling GFP expression. The activities of the ligands Theo, HMB, and NMB were monitored by GFP fluorescence. All three ligands demonstrated increased GFP expression, with HMB and NMB showing significantly higher activation ratios (20.45 ± 2.56 and 18.44 ± 3.69) compared to Theo (6.09 ± 0.39) (Fig. 6c). EC calculations showed Theo requires the highest concentration of 5383 µM, while HMB and NMB demonstrated superior potency (234.8 μM and 287.6 μM, respectively) (Fig. 6d). Anti-GFP immunoblot analysis confirmed fluorescence observations, showing significant GFP increases with HMB and NMB versus Theo, with negligible expression in DMSO controls (Fig. 6e). These results indicate that both alternative ligands substantially showed improved activity than Theo in activating the riboswitch system in mycobacteria.

Effect of ligand activity on aptazyme-mediated gene regulation in mammalian cells:
To evaluate ligand activity in mammalian systems, we employed aptazyme-mediated gene regulation, Th_APT inserted into stem loop I of a hammerhead ribozyme. Without ligand, stems I and II interact, causing cleavage of the 3' poly(A) tail and mRNA degradation. Ligand binding induces structural changes that prevent stem interaction, preserving the poly(A) tail and stabilizing mRNA for translation. Aptazyme was inserted into the mCherry gene's 3' UTR (Fig. 7a), where ligand presence enables mCherry expression measurable by FACS. A constitutive BFP reporter under EF-1α on the same plasmid served as a transfection efficiency control.

The effects of the compounds were validated by confocal microscopy following transient transfection in HEK293T cells. The cells were treated with ligands for 48 hours after 6 hours of transfection and then assessed for mCherry expression. Upon treatment with the ligand molecule Theo, HMB, and NMB, an increase in mCherry expression was observed compared to the DMSO control. HMB and NMB showed a significant increase in mCherry expression. In contrast, Theo showed minimal detectable changes (Fig. 7b and Supplementary Fig. 19). The effect of ligands was also quantified by FACS. It was observed that HMB showed a maximal increase in mCherry expression by 11.27-fold, and NMB showed up to a 6.55-fold increase, while Theo showed minimal effect with a 2.93-fold increase in the expression (Fig. 7c).

These ligands were further characterized to demonstrate a dose-response relationship between relative fluorescence levels and ligand concentration, with EC ranging from 73.90 μM to 1487 μM (Fig. 7d). Further, the change in protein expression was analyzed by immunoblot using an anti-mCherry antibody. A similar trend was observed as that of FACS analysis, where a significant increase in expression of mCherry was observed in the presence of HMB and NMB as compared to Theo (Fig. 7e). The increase in expression was also recorded at transcription level; qRT-PCR data showed consistent increase in transcript levels with 9-fold increase in presence of HMB, and 6-fold increase in presence of NMB while Theo showed around 2.5-fold increase (Fig. 7f). The results obtained are consistent with the trend observed in prokaryotic system and the K values observed in ITC experiments, suggesting that the effect observed was due to the increase in affinity of developed ligand with that of Th_APT.

Advancing temporal control of aptazyme-mediated CRISPR-Cas9 genome editing in mammalian cells:
The requirement for precise temporal control over CRISPR-Cas9 genome editing is crucial to minimize off-target mutagenesis and limit the unintended editing in non-target tissues observed with the constitutive expression of canonical genome-editing agents. This highlights the need for precise regulation of CRISPR-Cas9 activity, particularly through spatiotemporal control mechanisms82,83. One innovative strategy involves small-molecule-controlled genome editing, which utilizes theophylline-based aptazyme-embedded guide RNAs (gRNAs)84. In this approach, the gRNA is sequestered by a complementary blocking sequence, thereby preventing its interaction with its genomic target. Upon ligand binding, the aptazyme undergoes self-cleavage, physically releasing the gRNA from the blocking sequence and allowing the Cas9 complex to bind the target DNA84.

In addition to our initial validation in translation assays, we sought to investigate the efficiency of our developed ligands, HMB and NMB, for a small-molecule-responsive CRISPR-Cas9 genome-editing application. To achieve this, we developed an ‘all-in-one’ pCS2-Cas9-gRNA plasmid construct using the pCS2-Cas9 scaffold, as shown in Figure 8a. In this ‘all-in-one’ construct to integrate both Cas9 and gRNA, we inserted the theophylline-responsive aptazyme-embedded gRNAs84, where the theophylline aptamer is inserted in stem III of the hammerhead ribozyme, while gRNA, designed to target the mCMV promoter, and a blocking sequence were incorporated as an extension in stem I. The genome-editing efficiency of the system was assessed in HEK293T cells using a GFP reporter assay. In this experiment, CRISPR-mediated disruption of the mCMV promoter directly correlates with a decrease in downstream GFP fluorescence. To quantify the efficiency of ligand-responsive genome editing, HEK293T cells were co-transfected with a GFP reporter and either the control pCS2-Cas9 or the all-in-one pCS2-Cas9-gRNA plasmid. At 6 hours post-transfection, cells were treated with varying concentrations of Theo, HMB, or NMB, and GFP fluorescence was monitored five days post-treatment. The percentage of GFP loss was calculated relative to control cells transfected with the GFP reporter and pCS2-Cas9 alone, which were defined as 100% signal.

In the absence of a ligand, the system maintained low leaky genome editing, exhibiting a negligible basal reduction (~5%) in GFP signal. Upon ligand administration, we observed a dose-dependent decrease in GFP fluorescence over a range of 0-200 µM (Fig. 8b). While 200 µM of Theo yielded a modest 25% reduction in signal, notably HMB demonstrated markedly superior performance, achieving a ~70% loss of GFP fluorescence. NMB showed an intermediate effect with a ~40% reduction (Fig. 8b). Notably, while a high dose of the ligand Theo (2 mM) was required to achieve a ~65% reduction in GFP fluorescence, consistent with previous reports (Supplementary Fig. 20)84. HMB achieved comparable or greater editing efficiency at a 10-fold lower concentration than Theo.

To confirm that the observed reduction in GFP fluorescence was a direct result of ligand-induced CRISPR-Cas9 genome editing, we performed a molecular validation across the DNA, RNA, and protein levels. Immunoblotting revealed a significant reduction in GFP protein levels for HMPB and NMB-treated cells, whereas Theo treatment at equivalent concentrations showed no substantial change in protein levels (Fig. 8c). To confirm that the loss of fluorescence resulted from targeted CRISPR-Cas9-mediated genomic modification, we quantified GFP mRNA transcript levels via qRT-PCR. Consistent with our fluorescence and immunoblotting results, treatment with HMB and NMB reduced GFP mRNA levels by up to 97% and 89%, respectively, which is significantly higher than the modest ~25% reduction observed with the Theo ligand (Fig. 8d). Finally, we confirmed the targeted genomic disruption using the T7 endonuclease I (T7E1) assay (Fig. 8e). While DMSO-treated samples showed a baseline indel frequency of 3.21%, Theo treatment increased to 9.69%. On the other hand, HMB and NMB yielded significantly higher indel frequencies of 41.65% and 25.27%, respectively (Fig. 8f). Collectively, these data establish HMB and NMB as highly potent inducers that provide significantly enhanced dynamic range and affinity compared to traditional theophylline-ligand-based switches.

Methods
Molecular Docking: The 3D structure of the Theo-binding RNA aptamer (PDB ID: 8D28) was retrieved from the Protein Data Bank and prepared using Schrödinger’s Protein Preparation Wizard, which involved adding hydrogens, assigning bond orders, removing water, and optimizing hydroxyl orientations. During the preparation step, magnesium and sodium atoms were retained, as observed in the aptamer's crystal structure. Restrained energy minimization was performed with Epik (RMSD cutoff: 0.3 Å). Approximately 1.0 million compounds from two different libraries: FDA-approved drugs from DrugBank and lead-like compounds from ZINC (MW 250–350 Da, LogP ≤ 3.5, ≤ 7 rotatable bonds) were screened. Ligands were prepared using LigPrep with Epik at pH 7.0 ± 0.4, generating ~5 conformers per ligand. The receptor grid was centered on the co-crystallized ligand (Theo) using Receptor Grid Generation, and protocol validation was confirmed by redocking Theo and calculating RMSD. All ligands were screened by high-throughput virtual screening (HTVS), and top hits were selected based on docking scores. The 50 best candidates were analyzed for scaffold diversity.

Aptamer synthesis: DNA templates used in RNA synthesis reactions were generated by primer extension PCR with the Kapa Taq PCR kit (KK1015), and oligonucleotides were obtained from Macrogen (Korea) (Table S7 to S9). In vitro transcription of RNAs was carried out using the HiScribe® T7 High Yield RNA Synthesis Kit (NEB, #N0431), and the transcripts were purified by denaturing polyacrylamide gel electrophoresis.

Thioflavin T (ThT)-mediated ligand-displacement assay: The library of effector ligands was developed, where the ligands were chemically synthesized in our lab (Supplementary Note 1) or obtained from commercial sources. Thioflavin T (ThT) fluorescence was used to monitor the binding of Th_APT RNA with the developed library of compounds. Reactions contained 1 µM RNA, 2 µM ThT, and buffer (10 mM HEPES pH 7.5, 100 mM NaCl, 2 mM MgCl₂, 0.01 % Tween 20) with effector ligands (0-50 µM; 1 % DMSO). Samples were incubated at 70 °C for 5 min, then cooled to 25 °C over 4 hours at 1 °C/sec. Fluorescence was recorded (Ex 440 nm, Em 480 nm), and K values were obtained by fitting corrected data to a 1:1 binding model using GraphPad Prism 8.0. All experiments were performed in triplicate.

Isothermal Titration Calorimetry (ITC): ITC experiments were performed using a MicroCal PEAQ-ITC (Malvern). Unless otherwise stated, buffer conditions were identical to those used in the ThT assay. For pH-dependent binding studies, HEPES was replaced with 10 mM Bis-tris propane adjusted to pH 7.0, 7.5, 8.0, or 8.5. The sample cell (200 μL) contained 5 μM RNA, and the syringe (40 μL) contained 50–100 μM ligand. Following equilibration at 25 °C, titrations were performed with an initial 0.4 μL injection, followed by 19 injections of 2.0 μL at 150 s intervals, with stirring at 750 rpm. Thermograms were integrated to generate binding isotherms, and K values were obtained by fitting to a 1:1 binding model using PEAQ-ITC Analysis Software. Experiments were performed in triplicate.

Fluorescent cyanine 5 (Cy5) labelling of Aptamer: A 300 pmol aptamer was fluorescently labeled at 3’ end using T4 RNA ligase with 20 μM pCp-Cy5 (Jena Bioscience, NU-1706-CY5). The ligation reaction was performed in manufacturer-supplied buffer (New England Biolabs, M0204S) supplemented with 1 mM ATP, 20 % (v/v) DMSO, and 0.6 U/μL T4 RNA ligase, incubated at 16 °C for 15 hours. The Cy5-labeled aptamer was purified using a QIAGEN RNeasy Mini Kit (74104). The purified Cy5-labeled RNA was eluted in nuclease-free water.

T1 Digestion assay: Cy5 labelled aptamer (5 pmol) in a buffer containing 50 mM Tris-Cl (pH 7.5) and 100 mM NaCl was mixed with or without 50 μM ligand and incubated at 70 °C for 5 min, followed by cooling to 25 °C at 1 °C/sec. Refolded samples were then incubated with 2 U of RNase T1 (Sigma) at 37 °C for 15 min, and the reaction was then quenched by an equal volume of 2X loading dye (95 % Formamide, 18 mM EDTA, 0.025 % SDS). Partial alkaline digestion of RNA was performed by incubating 5 pmol of Cy5 labelled RNA in a 10 μL mixture containing 50 mM Na CO and 50 mM NaH CO (pH 9.5 at 25 °С) and 1 mM EDTA, at 90 °С for 5 min, and immediately cooled on ice. RNase T1 ladder was created by incubating 5 pmol Cy5-labeled RNA in a 10 μL mixture containing 5 M urea, 25 mM sodium citrate, 1 mM EDTA (pH 5.0 at 23 °С), and 10 units of RNase T1 at 55 °С for 15 min, followed by immediate cooling on ice. The samples were then electrophoresed on a 15 % denaturing polyacrylamide gel. The Cy5 signal in the gel was detected using Amersham Typhoon (Cytiva).

In-line probing: In-line probing was conducted using the Th_APT aptamer to assess ligand binding. For each reaction, 5 pmol of pCp-Cy5-labeled RNA aptamer was incubated with varying ligand concentrations (0-100 µM) in a 10 µL reaction volume containing 50 mM Tris-HCl (pH 8.3), 2 mM MgCl₂, 100 mM KCl, and 1 % DMSO. The reactions were refolded by heating at 70 °C for 5 min, followed by incubation at 37 °C for 72 hours to allow for spontaneous RNA cleavage. Following incubation, control ladders were prepared for reference. The T1 ladder was generated by treating 5 pmol of labeled RNA with 1 unit RNase T1 (Sigma, R1003-100KU) in a buffer containing 0.25 M sodium citrate (pH 5.0), 5 M urea, and 1 mM EDTA at 55 °C for 15 min. The OH ladder was prepared by incubating 5 pmol of Cy-5-labeled RNA in 50 mM NaHCO₃/Na₂CO₃ (pH 9.5) with 1 mM EDTA at 90 °C for 5 min. All reactions, including the in-line probing samples and ladder controls, were quenched by adding an equal volume of 2X loading buffer. The resulting products were resolved on a 15% denaturing polyacrylamide gel, and band intensities were quantified after gel imaging with an Amersham Typhoon system (Cytiva).

Surface Plasmon Resonance: The real-time binding of Th_APT to Theo and HMB was analyzed using a BIAcore T200 at 25 °C. Biotinylated-(dT)₂₄ was immobilized on a streptavidin sensor chip (SA), and poly(A)-tailed RNA was captured (~1000 RU) in flow cell 2; flow cell 1 served as a reference. Ligands (0.625–10 μM) were injected in HBS-N buffer, and specific binding responses were obtained by subtracting reference signals. The surface was regenerated with 20 mM NaOH. Kinetic constants (k , k , K ) were derived using BIAevaluation 3.0 with a 1:1 model.

Analysis of intracellular compound concentrations: Bacterial growth assays were performed by preparing a 1% inoculum of an overnight-grown culture in M9 medium. 95 µL of inoculum was dispensed into each well of a 96-well plate, followed by the addition of ligands (0–500 µM) with DMSO maintained at 5 %. Plates were incubated at 37 °C with shaking at 800 rpm for 16 hours, and growth was measured at 600 nm. Growth in DMSO alone was defined as 100 %, and relative growth in the presence of ligands was calculated. Bacteria were treated with 500 µM ligand for intracellular concentration analysis under identical conditions. After 16 hours, cells were harvested, washed twice with cold PBS, and lysed in acetonitrile with glass beads (2:1) using a bead beater. Lysates were centrifuged, the supernatants were vacuum-dried, and the residues were resuspended in methanol for HPLC analysis.

For HEK-293T cells, 10,000 cells per well were seeded and treated with 0-200 µM of ligand 24 hours after seeding. After 48 hours of treatment, cell growth was assessed using the MTT assay, with viability in the presence of DMSO considered 100 %. For intracellular ligand accumulation analysis, cells were harvested after 48 hours of treatment, washed twice with cold PBS, and lysed in acetonitrile with glass beads (2:1) using a bead beater. Lysates were centrifuged, the supernatants were vacuum-dried, and the residue were resuspended in methanol for HPLC analysis.

HPLC experiments were conducted on a Waters system with a C18 column and a PDA detector set at 280 nm. The mobile phase comprised solvent A (acetonitrile) and solvent B (water containing 0.1 % trifluoroacetic acid). A binary gradient was applied: 10–100 % B over 20 min, 100 % B from 20 to 25 min, and a return to 10 % B over the following 5 min. Peak areas were quantified using the system’s software, and standard curves generated from known ligand concentrations were used to determine intracellular levels in the test samples. The calculated concentrations were normalized to µg per 108 CFU based on the culture's optical density (OD ).

Plasmid construction: DNA fragments encoding various riboswitches were amplified by PCR using primers listed in Table S6. The resulting PCR products were subjected to restriction digestion with XbaI and SacI enzymes, then ligated into the corresponding restriction sites of the AEB-Theo45 vector plasmid (a gift from Howard Salis; Addgene plasmid #63850; http://n2t.net/addgene:63850; RRID: Addgene_63850). This cloning strategy generated reporter constructs containing riboswitches positioned upstream of the RFP gene.

For aptazyme-based CRISPR-mediated genome editing, the DNA template containing U6 promoter, gRNA sequence with aptazyme, gRNA, and blocking sequence was PCR amplified from theophylline-agRNA-GFP (a gift from David Liu; Addgene plasmid # 96856; http://n2t.net/addgene:96856; RRID: Addgene_96856) and cloned into pCS2-Cas9 (a gift from Alex Schier; Addgene plasmid # 47322; http://n2t.net/addgene:47322; RRID: Addgene_47322) using restriction sites KpnI and NotI. Recombinant plasmid DNA was purified and verified by sequencing to confirm proper riboswitch insertion before proceeding with experimental analyses.

Assessment of riboswitch function in E. coli: Fluorescence measurement experiments were performed with the E. coli DH10β strain unless stated otherwise. Cells transformed with plasmids for the RFP reporter under the control of different riboswitches were cultured overnight in M9 medium (pH 7.0 ± 0.2) supplemented with 25 μg/mL chloramphenicol at 37 °C with shaking at 180 rpm. The overnight cultures were then diluted to 1 % (v/v) into fresh M9 medium containing 25 μg/mL chloramphenicol, and various ligands were added at predefined concentrations; the solvent control and all test samples contained 5 % DMSO. Cultures were incubated for 18 h at 37 °C with orbital shaking at 800 rpm. After incubation, OD and fluorescence values were measured using an I3x SpectraMax plate reader (Molecular Devices). RFP fluorescence was recorded in black-well plates with an excitation wavelength of 555 nm and an emission wavelength of 608 nm. Fluorescence values were normalized to OD to assess ligand-dependent changes in RFP expression. EC values were determined using GraphPad Prism software with a non-linear regression model (log[agonist] vs. response) applying shared top and bottom parameters.

FACS analysis: Bacterial populations transformed with riboswitch constructs were incubated with 500 µM of each test ligand for 16 hours at 37 °C under standard growth condition. After incubation, cells were harvested by centrifugation at 3500 x g for 5 min, washed twice with phosphate-buffered saline (PBS), and resuspended in PBS. Cells were then stained with Hoechst for 5 min at room temperature, washed twice to remove excess dye, and finally resuspended in PBS. Flow cytometric data acquisition was performed using a BD FACS Symphony™ A1 cell analyzer equipped with a YG602A bandpass filter for RFP detection, while UV446A was used for Hoechst. MFI values for Hoechst and RFP were calculated in FlowJo. RFP MFI values were normalized to Hoechst, and fold changes were calculated, with RFP expression in DMSO-treated samples set to 1 for RS-4 and to 100% for RS-10.

Confocal Imaging: Bacterial cells treated with test compounds were harvested by centrifugation at 3500 x g for 5 min and washed with PBS. The cells were then fixed using 4 % paraformaldehyde (PFA) for 20 min at 4 °C. The PFA was removed by centrifugation, and the cells were washed twice with PBS. The nuclear material was counterstained with Hoechst for 5 min, and the cells were washed to remove excess dye. Further, the cells were resuspended in PBS, and 10 µl of the suspension was applied to a 24 × 50-mm coverslip. A 1 × 1-cm agarose pad was laid on top of the cell suspension and covered with a 22 × 22-mm coverslip86. Confocal microscopy was performed using a Zeiss LSM 880 system with a 63X oil-immersion objective and appropriate excitation and emission filters to detect DAPI and RFP.

Western blotting for assessment of RFP expression in E. coli: Bacterial cultures were harvested, lysed by sonication (10 cycles: 20 s pulse/40 s rest, 50 % amplitude), and centrifuged at 18,000 x g for 10 min at 4 °C. Protein concentration in the supernatant was quantified using the Pierce BCA assay. Equal protein amounts (20 μg) were resolved by SDS-PAGE (12 % gels), transferred to PVDF membranes, and probed with anti-RFP (GeneTex, GTX127897) and anti-GAPDH (Biobharti Life Sciences, BB-AB0060) antibodies, followed by HRP-conjugated anti-rabbit secondary antibody (Biobharti Life Sciences, BB-SAB01B). Signals were detected by chemiluminescence on an ImageQuant LAS4000 system.

qRT-PCR analysis: Cultures were prepared under the same growth conditions described in the fluorescence measurement section. Following incubation, bacterial cells were harvested, washed, and resuspended in Tri-Xtract (G-Biosciences, 786-652) for total RNA extraction. Reverse transcription was done using 500 ng of total RNA with the iScript™ cDNA synthesis kit (Bio-Rad-1708890). Gene-specific primers (listed in Table S10) were then used for qRT-PCR. QuantStudio 6 FLEX Real-Time PCR System (Life Technologies) was used to perform the reactions with iTaq Universal SYBR Green Supermix (Bio-Rad-1725121).

Assessment of riboswitch function in Mycobacterium: M. smegmatis was cultured in 7H9 or 7H11 medium (Himedia) with 0.5 % glycerol and 0.05 % Tween-80 at 37 °C. The GFP reporter plasmid pST5552 (Addgene 36255) under theophylline riboswitch control was electroporated, and transformants were selected on 7H11 with 25 μg/mL kanamycin. Cultures (OD₆₀₀ = 0.2) were treated with ligands (0–500 μM) in 24-well plates for 24 hours at 800 rpm. OD₆₀₀ and GFP fluorescence were measured (Spectramax I3x), with fluorescence normalized to OD₆₀₀. EC₅₀ values were determined by non-linear regression in GraphPad Prism. For western blotting, cells treated with 500 μM ligands were processed as described previously, using anti-GFP (GeneTex, GTX113618) and anti-GAPDH (Biobharti Life Sciences, BB-AB0060) primary antibodies with HRP-conjugated secondary antibody (Biobharti Life Sciences, BB-SAB01B).

Cell culture: Human embryonic kidney 293LT (HEK293T) cells (ATCC-CRL-3216) were cultured in Dulbecco's Modified Eagle Medium (DMEM, Sigma Aldrich, D5648) supplemented with 10 % fetal bovine serum (FBS, Himedia, RM10409), 100 U/mL penicillin, 100 μg/mL streptomycin (Himedia, A002A), at 37 °C in a 5 % CO₂ humidified incubator.

Confocal microscopy: HEK293T cells (25000 cells/well) were seeded in 35mm glass-bottom plates and transfected with a plasmid containing theophylline aptazyme controlling expression of mCherry, while BFP is constitutively expressed under EF1α (100 ng) (pCS4092 was a gift from Christina Smolke (Addgene plasmid 131741; http://n2t.net/addgene:131741; RRID: Addgene_131741)) using Lipofectamine transfection reagent (Thermo Fisher Scientific, 11668019) following the manufacturer's protocol. H2B-GFP was used to mark the nucleus. Ligand was added to the plate at a 200 μM concentration 6 hours post-transfection, and fluorescence microscopy images were captured using a Zeiss LSM 880 at 63X magnification after 24 hours of incubation.

FACS analysis of aptazyme-based mRNA destabilization-mediated gene regulation: HEK293T cells (30,000/well, 48-well plate) were transfected with 250 ng plasmid using Lipofectamine 2000 (Thermo Fisher). Six hours later, ligands were added at varying concentrations. After 48 hours, cells were fixed in 4 % PFA, washed, and resuspended in FACS buffer. Flow cytometry was performed on a BD FACS Symphony A1 (mCherry: YG602A; BFP: V450). Data was analyzed in FlowJo, gating viable singlets; transfected cells (BFP > 103 units) were used for downstream analysis. mCherry fluorescence was normalized to BFP, and fold changes were calculated relative to DMSO control. Nonlinear regression in GraphPad Prism obtained dose–response curves and EC₅₀ values.

GFP fluorescence measurement for aptazyme-based CRISPR-mediated genome editing: HEK293T cells (15,000/well in a 48-well plate) were transfected with 200 ng of plasmid (100 ng GFP reporter and 100 ng pCS2-Cas9-gRNA) using Lipofectamine 2000 (Thermo Fisher). Six hours later, ligands were added at varying concentrations. Additional cells were incubated for 5 days with intermittent compound treatment on days 2, 3, and 4 post-transfection. After incubation, cells were washed with 1X cold PBS and lysed with passive lysis buffer (Promega E1941) for 20 minutes at 25°C with shaking at 600 rpm in a Dry bath (Helix Biosciences). The lysate was then clarified by centrifugation at 3500 x g for 5 min, and fluorescence was measured with excitation at 475 nm and emission at 510 nm using an I3x SpectraMax plate reader (Molecular Devices).

Western blot analysis: Western blotting was performed to assess aptazyme-mediated regulation of mRNA stability and CRISPR-based genome editing in HEK293T cells. Cells were seeded in 48-well plates and transfected using Lipofectamine 2000 (Thermo Fisher Scientific). For mRNA destabilization assays, 30,000 cells per well were transfected with 250 ng plasmid. For CRISPR-mediated genome-editing assays, 15,000 cells per well were transfected with 200 ng of plasmid DNA (100 ng of a GFP reporter and 100 ng of pCS2-Cas9-gRNA). Ligands were added 6 h post-transfection at a concentration of 200 µM. Cells were incubated for up to 48 hours post-transfection for aptazyme-mediated regulation of mRNA stability, whereas for long-term genome-editing experiments, cells were incubated for 5 days with intermittent ligand treatment on days 2, 3, and 4 post-transfection. After incubation, cells were lysed using a Bioruptor Pico sonicator (10 cycles, 30 s ON/30 s OFF). Lysates were clarified by centrifugation at 18,000 x g for 10 min at 4 °C, and protein concentrations were determined using a BCA assay. Equal amounts of protein were resolved by 12% SDS–PAGE and transferred onto PVDF membranes. For aptazyme-mediated mRNA destabilization experiments, membranes were probed with anti-mCherry (GeneTex, GTX128508) and anti-β-actin (Cell Signaling Technology, 3700T) antibodies. For CRISPR-mediated genome editing experiments, membranes were probed with OctA-Probe antibody (Santa Cruz Biotechnology, sc-166355) and anti-β-actin (Biobharati Life Science, BB-AB0024). HRP-conjugated secondary antibodies were used as appropriate (Biobharati Life Science, BB-SAB01B; BioLegend, 405306). Protein bands were detected using the Gel Doc XR+ Gel Documentation System.

qRT-PCR analysis: Quantitative RT–PCR was performed to assess aptazyme-mediated regulation of mRNA stability and CRISPR-based genome editing in HEK293T cells. For mRNA destabilization assays, cells were seeded at 50,000 cells per well in 24-well plates 24 h prior to transfection and transfected with 500 ng of plasmid DNA using Lipofectamine 2000 (Thermo Fisher Scientific). For CRISPR-mediated genome-editing assays, cells were seeded at 15,000 cells per well in 48-well plates and transfected with 200 ng of plasmid DNA (100 ng of a GFP reporter and 100 ng of agRNA-pCS2-Cas9). In all experiments, ligands were added 6 h post-transfection at the indicated concentrations. For long-term editing experiments, cells were incubated for up to 5 days with intermittent ligand treatment on days 2, 3, and 4 post-transfection.

Cells were harvested 48 h post-transfection or at the indicated time points, trypsinized, and total RNA was extracted using TRI-extract™ reagent (G-Biosciences, 786-652) according to the manufacturer’s instructions. Reverse transcription was performed using the iScript™ cDNA Synthesis Kit (Bio-Rad, 1708890) with 500 ng total RNA for mRNA destabilization assays and 2.5 μg total RNA for CRISPR-mediated genome editing assays. Quantitative PCR was performed using gene-specific primers (listed in Table S10) and iTaq™ Universal SYBR® Green Supermix (Bio-Rad, 1725121) on a QuantStudio™ 6 FLEX Real-Time PCR System (Life Technologies).

T7E1 cleavage assay for aptazyme-based CRISPR-mediated genome editing: % Indels were assessed using the T7 Endonuclease I (NEB: M0302). Briefly, 2 × 10⁶ cells were lysed in 50 μL Cell Lysis Buffer supplemented with 2 μL Proteinase K, incubated at 68°C for 15 min and 95°C for 10 min, and processed for PCR amplification. PCR products were purified using the NucleoSpin Gel and PCR Cleanup Kit (Takara 740611). PCR amplicons were denatured and re-annealed to generate heteroduplex DNA. For each reaction, 200 ng PCR product was combined with 1 μL 10× Reaction Buffer and nuclease-free water to a final volume of 9 μL. Samples were heated to 95°C for 5 min, cooled from 95°C to 85°C at −2°C s⁻¹, then slowly cooled from 85°C to 25°C at −0.1°C s⁻¹, and held at 4°C. Mismatch-containing heteroduplexes were digested by the addition of 1 μL T7E1 Enzyme and incubation at 37°C for 1 h. Negative enzyme controls were prepared in parallel by substituting nuclease-free water for the enzyme. Digestion products were resolved on 2% agarose gel and visualized by UV transillumination. Band intensities were quantified using Alpha-imager. Cleavage efficiency was calculated as:
Indel% = 100×(1− √1−fraction cleaved)
where fraction cleaved represents the ratio of cleaved band intensity to total (cleaved plus parental) band intensity.

Statistical analysis: Statistical analyses were carried out using one-way ANOVA with Dunnett’s post hoc test, two-way ANOVA with Sidak’s post hoc test, or two-tailed t-tests, as appropriate. All analyses were performed in GraphPad Prism (version 10.4.1; www.graphpad.com). The number of biological replicates is indicated in the figure legends. Data is reported as mean ± standard error of mean, with statistical significance set at α = 0.05

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript presents the development of 4-quinazolinone derivatives as a new class of synthetic ligands for the well-\nknown theophylline aptamer and riboswitch. Through large-scale virtual screening and docking simulations, the authors\nidentify 10 candidates including two promising ligands (HMB and NMB) that exhibit improved binding affinities, enhanced\ncellular penetration, and a broader dynamic range of gene regulation compared to the native theophylline ligand.\nTo validate the performance of these synthetic ligands, the authors performed a comprehensive set of in vitro assays (ITC,\nSPR, and in-line probing), showing approximately 10–25-fold stronger binding affinity relative to the original ligand. They\nfurther characterized ligand-dependent gene expression driven by theophylline riboswitches and aptazymes in both\nbacterial and mammalian systems at translational and transcriptional levels.\nOverall, this work reports a thorough biochemical and cellular characterization of new synthetic theophylline analogs (HMB,\nNMB), demonstrating improved binding affinity and regulatory performance. The concept of enhancing riboswitch behavior\nthrough ligand engineering is compelling; however, the overall novelty remains low.\nSeveral comments should be considered:\nMajor comments:\n1. In Figure 4 e, 5 e, 7 b, all microscopy panels appear low in resolution, making it difficult to observe cells and signal\ndistribution. We recommend repeating imaging using agarose-pad immobilization as described in Skinner et al. (Nat.\nProtoc., 2013, 8:1100–1113) to improve image quality and reproducibility.\n2. The authors showed differential pH sensitivity among the ligands (Theo, HMB, NMB) in riboswitch RS-4-mediated gene\nexpression, with NMB showing maximal activity at pH 7.0, unlike Theo and HMB, which peak at pH 8.0. The authors\nsequentially claimed that HMB and NMB can be used as pH-dependent gene regulation, however, pH changes can\nnormally influence cellular metabolism and growth, thus, in vitro assays (e.g., ITC, SPR, or EMSA) under varying pH\nconditions would provide essential evidence that these effects arise from direct ligand–RNA interactions rather than cellular\nphysiology.\n3. In Figure 3 a and b, the annotation of G positions on the in-line probing gels is unclear. The authors should specify which\nend (5′ or 3′) carries the Cy5 label, as the order of the G positions seems inverted if labelling is at the 5′ end (expected from\nG29). Additionally, the statement that the dose–response relationship “provides strong evidence for ligand binding” (line\n207) may be overstated given the modest quantitative changes observed.\n4. While authors described the docking workflow is generally appropriate for large-scale virtual screening, several\nmethodological aspects could be refined to improve the accuracy and reproducibility of RNA–ligand docking results.\nThe authors removed all crystallographic waters during receptor preparation. However, in RNA–ligand complexes, ordered\nwaters and hydrated ions frequently mediate crucial hydrogen-bonding networks between ligand and nucleobases (e.g.,\nbridging or Mg²⁺-coordinated interactions). Retaining key water molecules within the binding pocket or employing\n“conserved-water” docking protocols available in Schrödinger would yield a more realistic description of the binding\ninterface. The PDB entry 8D28 contains Mg²⁺ ions and Na⁺ that stabilize the binding pocket and aptamer fold. These should\nideally be retained and parameterized in the receptor grid, as they influence local charge distribution and may form direct or\nwater-mediated coordination with the ligand. Neglecting these interactions can lead to an inaccurate description of the\nbinding interface and overly simplified electrostatics.\nMinor comments:\n1. Figures 4e, 5e, and 7b lack scale bars, and the font size in all microscopy images is too small to read clearly.", "reviewer_3": "The manuscript presents the development of a new class of synthetic ligands—4-quinazolinone derivatives—targeting the\ntheophylline riboswitch and explores their potential to modulate aptamer-mediated gene expression in both prokaryotic and\neukaryotic systems. To date, only a limited number of compounds have demonstrated higher binding affinity to the\ntheophylline aptamer than its native ligand, theophylline. In this study, the authors designed and synthesized novel 4-\nquinazolinone derivatives that exhibit strong binding affinity to the theophylline aptamer and function as effective “ON–OFF”\nswitches for translational gene regulation. This work offers valuable insights into enhancing the performance and versatility\nof the theophylline riboswitch in synthetic biology applications by introducing an efficient and tunable inducer with improved\nbinding affinity, low toxicity, and superior cellular uptake.\nThe authors provide thorough biophysical analyses, investigating the structure-activity relationship of the quinazolinone\nderivatives with the theophylline aptamer, alongside biological studies in both prokaryotic and eukaryotic cells to\ndemonstrate translational gene regulation capabilities. While this platform holds significant promise within this field, major\nrevisions are necessary before the manuscript is suitable for publication.\nMajor comments:\n1. In the ligand-aptamer interaction studies, the authors used APT_M as the control aptamer to demonstrate the specific\nbinding of the ligands to Th_APT. However, additional control experiments with diverse RNA structures—such as RNA\nhairpin stem-loop motifs, four-stranded RNA structures that form base stacks (e.g., G-quadruplexes), and single-stranded\nRNA (ssRNA)—should be performed. Since regulatory regions of many genes contain RNA sequences capable of forming\nvarious secondary structures, non-specific binding of the ligands to these motifs could lead to unintended changes in gene\nexpression. The authors are encouraged to include more comprehensive controls for off-target binding and to further discuss\nthe potential impact of such off-target interactions on gene regulatory efficiency.\n2. Figure 2h: The lower ITC thermogram panel does not appear to correspond with the upper panel for the ligand NMB.\nBased on the titration data in the upper panel, NMB seems to exhibit a higher Kd value than HMB. The authors are advised\nto recheck and verify the thermogram data for accuracy.\n3. The authors have used 500 µM ligand concentration in all of the biological studies. This seems high. How did they arrive\nat this concentration.\n4. The flow cytometry results presented in Fig. 4 & 5 are not convincing. The peak width in the histograms for Theo, HMB\nand NMB are not consistent with the claims. In ON type riboswitches, NMB appears better and in OFF type riboswitches,\nTheo works best to reduce the expression. It would be better to use dual staining (e.g., nucleic acid-binding dyes) in\ncombination with the ligand during FACS analysis to obtain a clearer and more reliable representation of the data.\n5. The confocal images presented in Fig. 4, 5 and 7 are of poor quality. The authors are suggested to provide higher-\nresolution and magnified images of both E. coli and HEK293T cells to allow better visualization and interpretation of the\nresults. Furthermore, in Fig. 7, dual staining with nuclear imaging dyes is recommended to help elucidate the spatial\nlocalization of mCherry expression within the cells.\nMinor comments:\n1. Line 98-99: The aptamer nucleotide numbering in Figure 1a does not match that in Figure 1c, which may confuse readers.\nAccording to Figure 1c, theophylline forms hydrogen bonds with C22 and U24 and is stacked between A7–C8–G26 and\nU6–U3–A28. While the colour scheme in Figure 1a is consistent, the nucleotide numbering differs from the 5′ end of the\ntheophylline aptamer sequence. The authors should use consistent numbering across figures.\n2. In Figure 3, a schematic representation of RNAse T1 digestion of Th-APT would be useful for general readership to\nunderstand the assay.\n3. Figure 3a-b and Figure S4: The concentrations of the ligands are not mentioned in the figure/ figure caption.\n4. Line 240: The figure reference should be corrected to Fig. S8. Additionally, it is recommended that the authors use distinct\ncolour codes in the sequences presented in Fig. S8 to clearly differentiate among RS1-6 in regards of the aptamer region\nand its pre- and post-sequence regions. Similar colour codes are also useful for Fig. 4a, 5a and 7a.\n5. Line 264. Figure number is not correct.\n6. In Fig. 7a, the authors are suggested to label the stems I and II region in the hammerhead ribozyme."}, "author_response": ""}, {"round": 2, "reviewer_comments": {"reviewer_1": "Comment 1: In Figure 4 e, 5 e, 7 b, all microscopy panels appear low in resolution, making it difficult\nto observe cells and signal distribution. We recommend repeating imaging using agarose pad\nimmobilization as described in Skinner et al. (Nat. Protoc., 2013, 8:1100–1113) to improve the image\nquality and reproducibility.\nReviewer response: We appreciate the authors’ efforts to implement this suggestion. The revised microscopy images\nobtained using agarose-pad immobilization show improved resolution, and single-cell features are now strongly discernible,\nsubstantially enhancing data interpretability.\nComment 2: The authors showed differential pH sensitivity among the ligands (Theo, HMB, NMB) in\nriboswitch RS-4-mediated gene expression, with NMB showing maximal activity at pH 7.0, unlike\nTheo and HMB, which peak at pH 8.0. The authors sequentially claimed that HMB and NMB can be\nused as pH-dependent gene regulation, however, pH changes can normally influence cellular\nmetabolism and growth, thus, in vitro assays (e.g., ITC, SPR, or EMSA) under varying pH conditions\nwould provide essential evidence that these effects arise from direct ligand–RNA interactions rather\nthan cellular physiology.\nReviewer response: We appreciate the authors’ inclusion of additional in vitro binding measurements. The new ITC data\nclearly illustrate pH-sensitive differences in ligand–aptamer binding affinities among the three ligands. These results provide\nconvincing evidence that the observed regulatory differences result from direct molecular interactions and address the\npromising potential for both in vitro and in vivo pH-responsive gene regulation in future applications.\nComment 3: In Figures 3a and b, the annotation of G positions on the in-line probing gels is unclear.\nThe authors should specify which end (5′ or 3′) carries the Cy5 label, as the order of the G positions\nseems inverted if labelling is at the 5′ end (expected from G29). Additionally, the statement that the\ndose–response relationship “provides strong evidence for ligand binding” (line 207) may be overstated, given the modest quantitative changes observed.\nReviewer response: We thank the authors for addressing these issues. The revised figures now include clear labeling and\ndirectional annotation of the gels, improving readability and interpretation. Also, the corresponding text in line 207 has been\nappropriately moderated, resulting in a more balanced and accurate description of the evidence.\nComment 4: While the authors described the docking workflow is generally appropriate for large-scale\nvirtual screening, several methodological aspects could be refined to improve the accuracy and\nreproducibility of RNA–ligand docking results.\nThe authors removed all crystallographic waters during receptor preparation. However, in RNA–ligand\ncomplexes, ordered waters and hydrated ions frequently mediate crucial hydrogen-bonding networks\nbetween ligand and nucleobases (e.g., bridging or Mg²⁺-coordinated interactions). Retaining key water\nmolecules within the binding pocket or employing “conserved-water” docking protocols available in\nSchrödinger would yield a more realistic description of the binding interface. The PDB entry 8D28\ncontains Mg²⁺ ions and Na⁺ that stabilize the binding pocket and aptamer fold. These should ideally be\nretained and parameterized in the receptor grid, as they influence local charge distribution and may\nform direct or water-mediated coordination with the ligand. Neglecting these interactions can lead to\nan inaccurate description of the binding interface and overly simplified electrostatics.\nReviewer response: We thank the authors for revising the docking simulations with retention of relevant ions and ordered\nwater molecules in the binding pocket. It is encouraging that the core findings remain robust under these more rigorous\nconditions, providing stronger support for the docking results and their consistency with experimental validation.", "reviewer_3": "The authors have satisfactorily addressed the previous concerns by performing adequate additional experiments, and the\nmanuscript is now suitable for acceptance. However, a few minor corrections are still required, as outlined below:\nLine 103: The lower stack depicted in Figures 1c and 1d is composed of U6–U23–A28. However, the text describes it as\nU6–U3–A28. Please revise the text accordingly.\nLine 126: In Fig. S1b, no ThT fluorescence data are shown for Th_APTM. Please also clarify which RNA was used in Fig.\nS1b — was it Th_APT or a different RNA sequence?\nTable S2: For the benefit of general readers, please include the short names of the ligands alongside their full names in the\ntable.\nFigures S5 and S6: The RNA structure and corresponding sequence should be presented in these figures for clarity.\nFigure S18 (a–l): The pH values and ligand names should be indicated in each panel for easier interpretation."}, "author_response": "Comment 1: In Figure 4 e, 5 e, 7 b, all microscopy panels appear low in resolution, making it difficult\nto observe cells and signal distribution. We recommend repeating imaging using agarose-pad\nimmobilization as described in Skinner et al. (Nat. Protoc., 2013, 8:1100–1113) to improve image\nquality and reproducibility.\nAuthor response: We thank the reviewer for this suggestion. As suggested, we used agarose pads to fix\nthe bacterial cells prior to imaging, thereby significantly improving image resolution. Furthermore, we\nreacquired the HEK293T cell images using a high-resolution microscope. These updated images have\nbeen included below as Figures 2A (bacterial), and 2B (mammalian), and updated in the revised\nmanuscript as Figures 4f, 5f, and 7b, respectively.\nComment 2: The authors showed differential pH sensitivity among the ligands (Theo, HMB, NMB) in\nriboswitch RS-4-mediated gene expression, with NMB showing maximal activity at pH 7.0, unlike\nTheo and HMB, which peak at pH 8.0. The authors sequentially claimed that HMB and NMB can be\nused as pH-dependent gene regulation, however, pH changes can normally influence cellular\nmetabolism and growth, thus, in vitro assays (e.g., ITC, SPR, or EMSA) under varying pH conditions\nwould provide essential evidence that these effects arise from direct ligand–RNA interactions rather\nthan cellular physiology.\nAuthor response: We thank the reviewer for this suggestion. To investigate the mechanism of pH-dependent gene\nexpression, we performed Isothermal Titration Calorimetry (ITC) to evaluate the ligand-aptamer\nbinding across a pH range (pH=7.0 to pH=8.5). Representative ITC thermograms and equilibrium\ndissociation constant (KD) are shown in Figure 2 below, as well as supporting information Figure S18.\nThe ligand Theo shows the highest affinity, and the maximal fold changes between pH 7.5 and 8.0\n(Figure 3A). Similarly, HMB showed maximum affinity and fold change at pH 8.0 (Fig 3B), while NMB\ndisplayed higher affinity at pH 7.5, with the most significant fold change occurring between pH 7.0 and\n7.5 (Fig 3C). Collectively, these data suggest that pH-dependent gene regulation is primarily driven by\npH-induced enhancements in aptamer-ligand binding affinity in addition to pH-dependent changes in\ncellular physiology. We have incorporated these findings in the manuscript, Supplementary Figure S18\nand Table S5, and updated the discussion accordingly.\nComment 3: In Figures 3a and b, the annotation of G positions on the in-line probing gels is unclear. The authors should specify which end (5′ or 3′) carries the Cy5 label, as the order of the G positions seems inverted if labelling is at the 5′ end (expected from G29). Additionally, the statement that the dose–response relationship “provides strong evidence for ligand binding” (line 207) may be overstated, given the modest quantitative changes observed.\nAuthor response: We thank the reviewer for this suggestion. The aptamer is labelled with a Cy5 at the\n3’ terminus. The labelling position is now indicated in the graphical representation in Figure 3a, and\nwe have revised the line 207 statement in the manuscript text accordingly.\nComment 4: While the authors described the docking workflow is generally appropriate for large-scale\nvirtual screening, several methodological aspects could be refined to improve the accuracy and\nreproducibility of RNA–ligand docking results.\nThe authors removed all crystallographic waters during receptor preparation. However, in RNA–ligand\ncomplexes, ordered waters and hydrated ions frequently mediate crucial hydrogen-bonding networks\nbetween ligand and nucleobases (e.g., bridging or Mg²⁺-coordinated interactions). Retaining key water\nmolecules within the binding pocket or employing “conserved-water” docking protocols available in\nSchrödinger would yield a more realistic description of the binding interface. The PDB entry 8D28\ncontains Mg²⁺ ions and Na⁺ that stabilize the binding pocket and aptamer fold. These should ideally be\nretained and parameterized in the receptor grid, as they influence local charge distribution and may\nform direct or water-mediated coordination with the ligand. Neglecting these interactions can lead to\nan inaccurate description of the binding interface and overly simplified electrostatics.\nAuthor response: We thank the reviewer for this insightful comment. We agree that these elements are\ncritical for an accurate representation of the binding interface and electrostatics. In response to this suggestion, we repeated the virtual screening using a revised receptor preparation. As recommended,\nwe retained the Mg²⁺ ions, Na⁺ ions, and ordered water molecules within the binding pocket of PDB\nentry 8D28. While the refined protocol resulted in shifts in individual binding scores, the core findings\nremained robust i.e. 4-quinazolinone was identified as the dominant scaffold among the top 50 hits.\nInterestingly, we did not observe direct metal coordination with our ligands. The updated data are now\nincluded in Supplementary Tables S1 and S2, and the Methods section (under 'Molecular docking') has\nbeen revised to reflect these more rigorous parameters.\nMinor comments:\nComment 1: Figures 4e, 5e, and 7b lack scale bars, and the font size in all microscopy images is too\nsmall to read clearly.\nAuthor response: Please refer to the response given to comment 1, where we have also covered the\nanswer for this comment.\nReviewer 3:\nMajor comments:\nComment 1: In the ligand-aptamer interaction studies, the authors used APT_M as the control aptamer\nto demonstrate the specific binding of the ligands to Th_APT. However, additional control experiments\nwith diverse RNA structures-such as RNA hairpin stem-loop motifs, four-stranded RNA structures that\nform base stacks (e.g. G-quadruplexes), and single-stranded RNA (ssRNA)-should be performed. Since\nregulatory regions of many genes contain RNA sequences capable of forming various secondary\nstructures, non-specific binding of the ligands to these motifs could lead to unintended changes in gene\nexpression. The authors are encouraged to include more comprehensive controls for off-target binding\nand to further discuss the potential impact of such off-target interactions on gene regulatory efficiency.\nAuthor response: We thank the reviewer for this insightful comment regarding the molecular\nspecificity of our developed ligands. To thoroughly investigate the selectivity of our lead compounds\nbeyond the initial mutant Th_APT_M described in the manuscript, we generated an additional series\nof theophylline aptamer mutants by systematically mutating single and double nucleotides in and\naround the binding pocket, as shown in Fig 4A. These variants were evaluated using Thioflavin T (ThT)\ndisplacement assays. We observed either no or Weak binding with Theo, HMB, and\nNMB (Fig 4A) (Manuscript supplementary table S3). Furthermore, to investigate the Structure-activity\nrelationship (SAR) of 4-quinazolinone derivatives, we tested derivatives substituted at positions 6\n(Hydroxy) and 7 (Methoxy) of HMB. We tested various 4-quinazolinone derivatives with alkyl, halo, or\nhydroxy groups and, as shown in Figure 4B (Manuscript Figure 1e, Table 2, and Supplementary Table\nS2), their binding affinity and gene-regulatory efficacy. Among the tested, we observed that they have\nlow affinity and lower fold change than HMB and NMB. Collectively, both experimental results\ndemonstrate that the interactions of HMB and NMB are highly specific to the Tht_APT aptamer\nsequence.\nIn response to the reviewer’s suggestion to investigate potential off-target interactions, we further\nevaluated our ligands against single-stranded RNA (ssRNA) as well as a variety of non-target\nsecondary structures, including hairpins (Hp), and G-quadruplexes (GQ). These interactions were\ncharacterized using FRET, Circular Dichroism (CD) spectroscopy, and CD thermal melting\nexperiments (summarized in Figure 5) (Manuscript supplementary Figure S5 and S6). The FRET as\nwell as CD spectral data revealed no significant shifts in FRET or molar ellipticity upon Theo or HMB\naddition, and the CD melting profiles confirmed that the thermal stability (Tm) of the GQ, ssRNA, and\nHp structures remained unperturbed in the presence of the ligands.\nIn summary, the mutation, SAR analysis in combination with FRET, CD, and thermal melting data\nindicate that the developed ligand HMB does not interact detectably with off-target GQ, ssRNA, or Hp\nmotifs. These findings strongly support our conclusion that HMB is a highly selective binder for its\ncognate aptamer. We have updated the manuscript to include additional control experiments that\nfurther validate the specificity of the ligands we developed.\nComment 2: Figure 2h: The lower ITC thermogram panel does not appear to correspond with the upper\npanel for the ligand NMB. Based on the titration data in the upper panel, NMB appears to have a higher\nK than HMB. The authors are advised to recheck and verify the thermogram data for accuracy.\nD\nAuthor response:\nAs noted by the reviewer, we have already shown in the manuscript Table 1 data that NMB exhibits\napproximately a 3-fold higher affinity than HMB. We have re-verified our initial panels and confirmed\ntheir accuracy. Furthermore, we have provided the thermograms for all three experimental repeats for\nboth HMB and NMB ligands to ensure clarity (as shown in Figure 6).\nReviewer Comment 3: The authors have used 500 µM ligand concentration in all of the biological\nstudies. This seems high. How did they arrive at this concentration.\nAuthor response:\nThe 500 µM ligand concentration was selected based on a preliminary dose-response optimization\nstudy. We titrated the ligands over a range of 0 to 2.5 mM to evaluate both biological efficacy and\ncellular viability (as shown in Figure 7 and now also provided in Manuscript Fig 4c, 5c, and\nsupplementary fig S13, and S15). Our results demonstrated that 500 µM is the optimal concentration\nto achieve the maximum fold change (saturation) without detectable cytotoxicity. Consequently, this\nconcentration was standardized across all biological experiments\nComment 4: The flow cytometry results presented in Fig. 4 & 5 are not convincing. The peak width\nin the histograms for Theo, HMB and NMB are not consistent with the claims. In ON type riboswitches,\nNMB appears better and in OFF type riboswitches, Theo works best to reduce the expression. It would\nbe better to use dual staining (e.g., nucleic acid-binding dyes) in combination with the ligand during\nFACS analysis to obtain a clearer and more reliable representation of the data.\nAuthor response:\nWe appreciate the reviewer’s suggestion regarding the flow cytometry data, with the suggested changes\nin analysis, the fold change increased to 380-fold, which was previously observed to be 245-fold. To\naddress the concerns regarding histogram consistency and data reliability, we have repeated the\nexperiments using Hoechst 33342 as a secondary nuclear stain. This dual-staining approach allowed\nus to normalize the mRFP signal relative to Hoechst, providing a more robust and reliable\nrepresentation of expression levels. As shown in Figure 8 (Manuscript figures 4d, 4e, 5d, 5e, and\nsupplementary figures S14 and S16), we have updated the histograms and included normalized Mean\nFluorescence Intensity (MFI) values for both RS-4 (ON-type) and RS-10 (OFF-type). The normalized\ndata clearly demonstrate that, in both switch types, the HMB achieves superior induction compared to\nNMB and Theo. With HMB, we observed approximately. 380-fold induction (ON Switch) and around\n80 % suppression (OFF switch). We have updated this data in the revised manuscript. These refined\nflow cytometry results are now in full agreement with our fluorescence spectroscopy quantification,\nconfirming the consistency across different analytical methods.\nComment 5: The confocal images presented in Fig. 4, 5 and 7 are of poor quality. The authors are\nadvised to provide higher-resolution, magnified images of both E. coli and HEK293T cells to facilitate\nbetter visualization and interpretation of the results. Furthermore, in Fig. 7, dual staining with nuclear\nimaging dyes is recommended to help elucidate the spatial localization of mCherry expression within\nthe cells.\nAuthor response:\nWe appreciate the reviewer’s suggestion regarding the quality of confocal images. To address these\nconcerns and improve clarity, we have replaced the previous images with newly acquired, high-\nresolution micrographs for both E. coli (refer to the response to reviewer comment 1) and HEK293T\ncells (Figure 9). In HEK-293T cells, H2B-GFP was used to mark the nucleus, suggesting that\nexpression is mostly cytoplasmic. The updated results were incorporated in Figure 4f, 5f, 7b, and\nsupplementary figure S19\nMinor comments:\nComment 1: Line 98-99: The aptamer nucleotide numbering in Figure 1a does not match that in Figure\n1c, which may confuse readers. According to Figure 1c, theophylline forms hydrogen bonds with C22\nand U24 and is stacked between A7–C8–G26 and U6–U3–A28. While the colour scheme in Figure 1a\nis consistent, the nucleotide numbering differs from the 5′ end of the theophylline aptamer sequence.\nThe authors should use consistent numbering across figures.\nAuthor response: We have changed the secondary structure of the aptamer in Figure 1, and so the\nnumbering is consistent in Figures 1a, 1c, and 1d, along with Figure 3.\nComment 2: In Figure 3, a schematic representation of RNAse T1 digestion of Th-APT would be useful for the general readership to understand the assay.\nAuthor response: We have added schematics in Figure 3 to clarify the experiment.\nComment 3: Figure 3a-b and Figure S4: The concentrations of the ligands are not mentioned in the\nfigure/ figure caption.\nAuthor response: We have included the ligand concentration in the respective figures.\nComment 4: Line 240: The figure reference should be corrected to Fig. S8. Additionally, it is\nrecommended that the authors use distinct colour codes in the sequences presented in Fig. S8 to clearly\ndifferentiate among RS1-6 in regards of the aptamer region and its pre- and post-sequence regions.\nSimilar colour codes are also useful for Fig. 4a, 5a and 7a.\nAuthor response: We have corrected the figure number, annotated the mutations with colour codes,\nand provided secondary structures for the mutants in supplementary Figures S11 and S12.\nComment 5: Line 264. The figure number is not correct.\nAuthor response: We have corrected the figure number.\nComment 6: In Fig. 7a, the authors are suggested to label the stems I and II region in the hammerhead\nribozyme.\nAuthor response: We agree with the reviewer’s suggestion. We have revised the schematic in Figure\n7a to explicitly label stems I and II of the hammerhead ribozyme.\nBelow, we provide the point-to-point responses to reviewer comments. We greatly appreciate\nthe reviewer's effort to review our manuscript. All modifications in the manuscript text have\nbeen highlighted in yellow.\nReviewer #3\nComment 1: Line 103: The lower stack depicted in Figures 1c and 1d is composed of U6–\nU23–A28. However, the text describes it as U6–U3–A28. Please revise the text accordingly.\nAuthor Response: We have corrected the nucleotide positions in line 103 to U6-U23-A28.\nComment 2: Line 126: In Fig. S1b, no ThT fluorescence data are shown for Th_APTM. Please\nalso clarify which RNA was used in Fig. S1b — was it Th_APT or a different RNA sequence?\nAuthor Response: The Figure citation has been changed to Fig. 2a, where the data for\nTh_APTM are presented. Also, for Fig S1b, we used Th_APT, which is now mentioned in the\ncorresponding figure legend.\nComment 3: Table S2: For the benefit of general readers, please include the short names of\nthe ligands alongside their full names in the table.\nAuthor Response: Supplementary Table 2 has been modified accordingly, with the short names\nused in the manuscript listed in the table.\nComment 4: Figures S5 and S6: The RNA structure and corresponding sequence should be\npresented in these figures for clarity.\nAuthor Response: We have added the secondary structures of the corresponding RNA\nsequences in the Supplementary Figures 5 and 6.\nComment 5: Figure S18 (a–l): The pH values and ligand names should be indicated in each\npanel for easier interpretation.\nAuthor Response: We have mentioned the ligand name and the pH values in each panel of the\nfigure S18 (a-l)."}]

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