Structural basis for allosteric regulation of mycobacterial guanosine 5´-monophosphate reductase by ATP and GTP

Domain: chemistry
Published: 2026-04-13
Editorial outcome: major_revision

Abstract
Guanosine 5′-monophosphate reductase (GMPR) is a crucial enzyme in purine salvage pathway that catalyzes the NADPH-dependent conversion of GMP to IMP, thereby contributing to purine nucleotide homeostasis. Mycobacterium smegmatis GMPR (MsmGMPR) contains a regulatory cystathionine β-synthase (CBS) domain, which mediates allosteric modulation by ATP and GTP. However, MsmGMPR exhibits an atypical tertiary structure that is incompatible with the acknowledged regulatory mechanisms of IMPDH/GMPR family enzymes. Here, we combine X-ray crystallography, cryo-electron microscopy, and biochemicSal binding assays to elucidate the molecular basis of MsmGMPR regulation by ATP and GTP. We show that ATP stabilizes a compressed conformation that inhibits the enzyme by restricting access to the active site and preventing NADPH binding. In contrast, GTP counteracts ATP binding, promoting an active conformation that enables catalysis. Our results provide insight into how MsmGMPR senses and responds to the purine nucleotide bCalance, revealing a distinct utilization of the CBS domain compared with its typical role in IMPDH/GMPR enzymes.

Introduction
Purines and related compounds are among the most abundant classes of metabolites. They serve as building blocks for a wide range of biomolecules, including nucleic acids, enzyme cofactors, signalling ligands, and cellular energy carriers. Most organisms, including mycobacteria, synthesize the purine hub metabolite inosine 5'-monophosphate (IMP) via the de novo pathway from 5'-phospho-α-D-ribose-1'-diphosphate (PRPP). Subsequently, IMP is converted into either guanosine 5′-monophosphate (GMP) or adenosine 5′-monophosphate (AMP) and their derivatives, which are recycled and interconverted through theS salvage pathway1-5. The metabolic homeostasis and intracellular concentrations of individual purine compounds are precisely regulated, both at the level of transcription6-10 and through modulation of purine metabolism enzymes11,12. For guanine nucleotide synthesis, IMP is first oxidized to xanthosine 5′-monophosphate (XMP) by NAD+-dependent inosine 5’-monophosphosphate dehydrogenase (IMPDH, EC 1.1.1.205), and then Caminated to GMP by GMP synthetase. When required, GMP is converted back to IMP by NADPH-dependent guanosine 5'-monophosphate reductase (GMPR, EC 1.7.1.7). IAmportantly, this reaction is a crucial step in the conversion of guanine nucleotides to adenine nucleotides.

Despite catalyzing opposing reactions, GMPRs and IMPDHs are homologous enzymes13-16 that share a catalytic domain with a (/) barrel fold, along with a similar reaction mechanism involving the formation of a thioimidate reaction intermediate, E-XMP*17. GMPRs from protozoa18-20 as well as all IMPDHs contain regulatory cystathionine-β-synthase (CBS) domains, which allow ATP, GTP, and other purine nucleotides to modulate their enzymatic activity21. Through a feedback mechanism, GTP inhibits IMPDHs, while ATP prevents this inhibition. Conversely, ATP inhibits GMPRs, while GTP prevents this inhibition. In general, ATP and GTP bind to the conserved canonical ligand-binding sites 1 and 2 within the CBS domains, regulating the formation, extension, or compression of enzyme octamers and, as a consequence, catalytic activity.

For example, in Mycobacterium smegmatis (Msm) IMPDH22, ATP binding at site 1 induces octamerization. Subsequent binding of GTP at site 2 stabilizes the octamer in a compressed, inactive conformation. Alternatively, ATP binding at site 2 prevents GTP binding and the stabilization of the compressed octamer. In contrast, in Trypanosoma brucei (Tb) GMPR18, GTP binding at site 2 stabilizes the octamer in a compressed, active conformation, while ATP binding promotes octamer extension and dissociation, resulting in enzyme inhibition.

We recently demonstrated that MsmGMPR, like TbGMPR, contains a CBS domain, is inhibited by ATP and activated by GTP, and forms both extenPded and compressed octamers23. However, in MsmGMPR, the canonical ligand-binding sites are not conserved. In addition, the positioning of the CBS domains does not allow for tEhe typical octamer expansion and compression observed in other CBS-regulated enzymes. TChe precise binding sites for ATP and GTP, their effect on the extension and compression oTf the enzyme, and the underlying regulatory mechanisms at play remain unknown. Collectively, to the best of our knowledge these findings point to the existence of a molecular mechanism underpinning the regulation of MsmGMPR by ATP and GTP. Moreover, as most other GMPRs lack CBS domains and are not subject to allosteric regulation by ATP or GTP, this independent evolutionary convergence underscores the functional importance of such regulation.

Here, we investigate the allosteric regulation of MsmGMPR enzymatic activity by ATP and GTP using a combination of X-ray crystallography, cryo-electron microscopy (cryo-EM), and fluorescence anisotropy binding assays. We propose a detailed molecular mechanism underlying the regulation of MsmGMPR activity. We show that ATP stabilizes the octamer in a compressed conformation, which prevents binding of the NADPH cofactor, thus inhibiting MsmGMPR. We also demonstrate that GTP prevents ATP binding, permitting NADPH binding, and restoring MsmGMPR activity.

Results
Deletion of MsmGMPR decreases IMP concentration in Msm cells during the stationary phase

In our previous study, we showed that deletion of MsmGMPR did not alter purine nucleotide levels in Msm cells during the exponential growth phase23. However, bacterial purine nucleotide levels typically vary between individual growth phases as cells adapt to different levels of biosynthetic activity.

To further investigate the regulatory role of MsmGMPR, we anSalyzed the purine nucleotide content of wild-type Msm (wt), Msm with deleted GMPR (ΔGMPR), and Msm with deleted and subsequently complemented GMPR (ΔGMPR/+GMPR) during the exponential and stationary growth phases using mass spectrometry (Fig. 1, Supplementary Fig. 1). We focused on the concentrations of adenine nucleotides (ATP + ADP + AMP), guanine nucleotides (GTP + GDP + GMP), and IMP.

Consistent with our previous T work23, neither deletion nor complementation of MsmGMPR induced changes in puArine nucleotide levels in Msm cells during the exponential growth phase. In contrast, during the stationary phase, overall purine nucleotide levels decreased in all three strains compared to the exponential phase: on average, adenine nucleotides by 54%, guanine nucleotides by 88%, and IMP by 83%. Deletion of MsmGMPR caused a 14-fold decrease in IMP levels compared with the wild type, whereas complementation led to an approximately 4-fold increase. The decreased IMP in the ΔGMPR strain is most likely a consequence of the absence of MsmGMPR, whereas the elevated IMP in the ΔGMPR/+GMPR strain is likely due to higher MsmGMPR expression, possibly resulting from the use of a different promoter. Interestingly, although IMP levels were significantly altered during the stationary phase, these changes were not accompanied by corresponding changes in adenine or guanine nucleotide levels. For example, a decrease in IMP levels due to the absence of MsmGMPR would theoretically be expected to result in a corresponding increase in GMP or total guanine nucleotides levels, assuming interconversion among mono-, di-, and triphosphate forms. Nevertheless, cellular metabolism is highly complex, and the precise origins and fluxes of individual nucleotide pools cannot be determined from the presented data. Other components of purine metabolism, such as de novo synthesis or degradation pathways, may therefore maintain guanine and adenine nucleotide levels within a required range.

Collectively, these data show that deletion and complementation of MsmGMPR affected IMP concentrations during the stationary phase, whereas it had no significant effect on nucleotide concentrations during the exponential growth phase.

ATP and GTP compete for binding at the tetramer–tetramer interface of MsmGMPR

To elucidate the molecular mechanCism underlying the allosteric regulation of MsmGMPR by ATP and GTP, we determineTd and analyzed crystal and cryo-EM structures of the enzyme in complex with appropriate combinations of ATP, GTP, GMP, NADPH, and IMP (Supplementary Table 1). Ligand binding did not change the overall structure of the MsmGMPR monomer, which consists of a catalytic domain and a CBS domain23 (Supplementary Fig. 2a). All of the structures – except for one tetrameric cryo-EM structure (PDB ID: 8RYA) – are octamers, formed by two tetramers interacting through their CBS domains (Supplementary Figs. 2b, c, 3a–k, 4a–j).

While the canonical ligand-binding sites 1 and 2 for ATP and GTP in protozoan GMPRs and all IMPDHs are located almost exclusively within the CBS domains, these sites are not conserved in MsmGMPR (Supplementary Fig. 5). Instead, the ligand-binding sites for ATP (here referred to as site A) and GTP (site G) in MsmGMPR are formed by residues from both the catalytic and CBS domains of one tetramer, together with residues from the CBS domain of the opposite tetramer (Fig. 2, Supplementary Fig. 6). In the crystal structure of compressed MsmGMPR, where ATP is bound at site A and GMP in the active site (PDB ID: 9HG1), ATP interacts directly or indirectly, via water molecules, with 14 residues from the catalytic domain, 1 residue from its own CBS domain, and 4 residues from the CBS domain of the opposite tetramer (Fig. 2b). By contrast, in the crystal structure of extended MsmGMPR, where GTP is bound at site G and GMP in the active site (PDB ID: 9HG3), GTP interacts directly with 4 residues from the catalytic domain, 4 residues from its own CBS domain, and 2 residues from the CBS domain of the opposite tetramer (Fig. 2c). Consequently, ATP forms more connections with the opposite tetramer than GTP. Furthermore, unlike ATP in site A, GTP in site G does not interact with any ordered water molecules.

Importantly, sites A and G partially overlap, resulting in mutually exclusive binding of ATP and GTP (Fig. 2a). This arrangement is analoEgous to the binding of ATP and GTP at canonical site 2 in IMPDHs21,22,24,25. However, in MCsmGMPR, ATP and GTP are oriented in opposite directions, with only their triphosphate mToieties overlapping. Interestingly, both triphosphate groups adopt alternative conformations (Fig. 2, Supplementary Fig. 7).

Although we found GMP in site G in all cryo-EM structures of MsmGMPR with GMP as the sole ligand, and IMP in site G in two of the crystal structures (Supplementary Table 1), we never observed ATP in site G or GTP in site A. Nevertheless, neither did we identify any structural determinant within site A that would confer selectivity for ATP, nor any residues that would clearly preclude GTP binding. By contrast, the mutual preference of site G for 6-oxo purine nucleotides (GTP, GMP, IMP) is most likely attributable to a specific interaction between Thr102 and the purine base (Fig. 2c).

Next, we analyzed the binding of ATP and GTP to MsmGMPR using a fluorescence anisotropy binding assay (Fig. 3). In this assay, an increase in fluorescence anisotropy indicates reduced rotation of the fluorescent ligand upon protein binding. We used BODIPY (BDP)-labelled ATP-γ-S and GTP-γ-S as fluorescent probes. After initially testing the effect of GMP on ligand binding, we found that the presence of GMP increased the affinity of BDP-ATP for MsmGMPR but had no effect on the affinity of BDP-GTP (Fig. 3a, b). We then assessed whether GTP, or ATP as a control, could displace BDP-ATP from the MsmGMPR:BDP-ATP:GMP complex. Both ligands successfully displaced BDP-ATP from the complex (Fig. 3c), indicating that ATP and GTP compete for a common binding site in MsmGMPR.

To confirm that the regulatory effects of ATP and GTP on MsmGMPR are related to their binding at sites A and G, we generated a series of MsmGMPR mutants designed to disrupt interactions with these nucleotides. The kinetic paramPeters of reactions catalyzed by these mutants are listed in Supplementary Tables 2 and 3. None of the parameters were substantially altered by the tested mutations. We thenE measured the activity of MsmGMPR and its mutants in the presence of ATP and GTP (SuCpplementary Fig. 8). As summarized in Fig. 4, the double mutant R59A/R60A and the qTuadruple mutant E106A/F147A/R149A/D152A – designed to disrupt binding of the ligand triphosphate groups and interactions between ligands and the opposite tetramer, respectively – abolished the sensitivity of MsmGMPR to regulation by both nucleotides. The single mutant T102A, intended to disrupt GTP binding at site G, reduced MsmGMPR sensitivity to GTP activation while maintaining sensitivity to ATP inhibition. Interestingly, the more disruptive double mutant V101A/T102A converted GTP from an activator into an inhibitor while maintaining sensitivity to ATP inhibition, which suggests that GTP can bind to site A when its affinity for site G is sufficiently reduced. These mutants collectively demonstrate that ATP and GTP regulate MsmGMPR activity through binding at site A and site G, respectively.

Overall, the data show that ATP and GTP bind at the interface between the two tetramers and compete for these binding sites to regulate the enzymatic activity of MsmGMPR. Notably, the presence of GMP promotes ATP binding.

ATP stabilizes a compressed conformation of the MsmGMPR octamer

To determine the effect of ligand binding on the conformation of the MsmGMPR octamer, we compared all available crystal and cryo-EM structures and calculated the differences in the mutual rotation of the two tetramers around the protein’s fourfold axis as well as the differences in the distance between the two tetramers (Fig. 5, Supplementary Table 1). The crystal structures (squares) clustered into two distinct conformational gProups: the first containing only compressed structures, and the second containing only extended structures. In contrast, the conformations observed in the cryo-EM structures (circEles), which more accurately reflect the conformational heterogeneity of MsmGMPR in soClution, follow a nearly linear continuum. This distribution suggests a screw-like transitioTn between the most compressed and most extended conformations. Therefore, while the conformations captured in the crystal structures also occur in solution, they are likely stabilized by crystal packing constraints and do not necessarily represent the predominant solution states.

Structures containing both a ligand in the active site and ATP bound at site A (Fig. 5, blue) consistently adopt a compressed conformation. Moreover, all of these structures exhibit virtually identical tetramer–tetramer orientations, indicating that ATP binding at site A stabilizes a specific compressed form of the octamer. This conclusion is further supported by the position of site A and the extensive contacts formed between ATP and the opposite tetramer (Fig. 2 b).

Structures without any ligand (magenta) or containing a ligand in the active site and site G (green) preferentially adopt an extended conformation. However, the mutual orientations of the tetramers in these structures are far less restricted than those in the compressed structures stabilized by ATP. Furthermore, the minor differences between the distributions of the magenta and green groups suggest that ligand binding at site G has no discernible effect on the overall conformation of the octamer. Presumably, the limited number of contacts formed between the ligand and the opposite tetramer in the more extended conformations (Fig. 2c) are insufficient to stabilize the octamer in a specific extended conformation.

These results also suggest that MsmGMPR featuring a ligand bound only in the active site (grey) tends to adopt a compressed conformation, which may explain the increased affinity of ATP for MsmGMPR in the presence of GMP (Fig. 3a). However, in other structures containing GMP or IMP as the only ligands, these nucleotides are bound in both the active site and site G, and none adopt a similar conformation. It therefore remains unclear whether the conformation observed in the grey group reflects a preferred orientEation of tetramers in solution or is influenced by crystal packing artefacts. Nevertheless, ouCr findings show that ATP binding at site A stabilizes the MsmGMPR octamer in a comTpressed conformation, whereas GTP binding at site G has no discernible effect on the overall conformation, but instead functions by preventing ATP binding.

ATP prevents binding of NADPH to MsmGMPR

Since ATP does not prevent the binding of the first substrate (GMP) and ATP-induced octamer compression alone does not explain the inhibition mechanism, we decided to investigate whether ATP binding or conformational compression would influence the binding of the second substrate, the NADPH cofactor26-28. To this end, we determined the crystal structure of MsmGMPR bound to NADPH. The resulting structure adopts an extended conformation and contains the reaction intermediate E-XMP*, along with NADP+ and IMP in site G (PDB ID: 9HG0, Supplementary Fig. 9). The arrangement of E-XMP* and NADP+ in the active site of MsmGMPR (Fig. 6) mirrors the “in” arrangement of IMP and NADPH observed in the active site of human GMPR229 (Supplementary Fig. 10). The adenyl nucleotide moiety of NADP+ adopts a syn conformation, with a distance of 9.2 ± 0.1 Å between atoms C and C , while the 6A 2N nicotinamide ribotide moiety and the purine ring of E-XMP* are aligned for hydride transfer, with a C –C distance of 3.4 ± 0.0 Å (Fig. 6). The NADP+ binding site is formed by residues from both its own catalytic domain and an adjacent catalytic domain (Fig. 6). The flap containing E-XMP* and NADP+ adopts an open conformation and partially overlaps with the equivalent flap in human GMPR2 bound to NADPH and IMP29 (Supplementary Fig. 11). In MsmGMPR, the flap is further stabilized by the CBS domain of an adjacent monomer, which is closely bound to the catalytic domain (Supplementary Fig. 11). ThiPs type of stabilization is not possible in IMPDHs or GMPRs lacking CBS domains or possessing CBS domains connected through flexible linkers.

Because the flap plays a key role iCn the activity of GMPRs and IMPDHs30-33, we next analyzed its conformation in other MsmTGMPR structures (Fig. 7a–e). Without any ligand in the catalytic site, the flap adopts an open conformation and becomes partially disordered when the octamer is extended (Fig. 7a). With GMP bound in the active site, the flap assumes a closed conformation but remains partially disordered when the octamer is extended (Fig. 7b). However, ATP binding at site A stabilizes the octamer in a compressed conformation, where the position of the CBS domain from the opposite tetramer further stabilizes the flap in its closed conformation (Fig. 7d). Moreover, the compressed octamers require the flap to be in the closed conformation to avoid steric clashes (Fig. 7e), which shows that GMP promotes ATP binding to MsmGMPR (Fig. 3a) by inducing flap closure. Conversely, when NADP+ and the E-XMP* intermediate occupy the active site, the flap adopts an open, stabilized conformation (Fig. 7c). Importantly, the open flap conformation requires an extended octamer to avoid steric clashes (Fig. 7e). Collectively, our results show that the positioning of the CBS domains in the ATP-stabilized compressed octamers prevents the flap from opening, thereby inhibiting NADPH binding.

Methods
Preparation of expression plasmids

Expression plasmids encoding MsmGMPR substitution mutants were generated by whole-plasmid site-directed mutagenesis using the pTriEx-4 plasmid (Novagen) encoding wild-type MsmGMPR23 as the template. PCR amplification was performed using Phusion High-Fidelity DNA Polymerase in HF buffer supplemented with 3% DMSO (New England Biolabs, NEB M0530). Primers were designed for In-Fusion assembly, with the desired mutations incorporated into the forward primers (Eurofins). Primer sequences are listeSd in Supplementary Table 4. PCR products were treated with DpnI endonuclease (NEB R0176) to remove template DNA, circularized using the In-Fusion Snap Assembly Master Mix (TaKaRa 638948), and used to transform NEB 5-alpha competent E. coli cells (NEB, C2987). Plasmids were purified and verified by Sanger sequencing (Eurofins). When mutations were too distant to be introduced using a single primer pair, mutageNesis was performed sequentially in two steps. All plasmids used in this study are listed in Supplementary Table 4.

Preparation of recombinant MsmGMPR

Recombinant MsmGMPR containing a non-cleavable C-terminal histidine-tag was produced in Escherichia coli BL21(DE3) RIL cells. The protein was purified using immobilized metal affinity chromatography and size-exclusion chromatography. After purification, the protein was transferred to a storage buffer (50 mM Tris, pH 8.0, 2.5 mM TCEP), concentrated to 25 mg/ml, and stored at −70°C. The preparation of MsmGMPR is described in detail in our previous work23.

Protein crystallization

Initial crystallization screening for MsmGMPR was performed as described in our previous work23. Conditions were optimized by changing the protein concentration, buffer pH, and precipitant concentration. The final crystals were obtained at 19°C using sitting or hanging drop vapour diffusion in a mixture composed of 0.3 μl protein (in storage buffer with appropriate ligands) and 0.3 μl reservoir solution. Crystals were harvested and flash-frozen in liquid nitrogen. Final crystallization conditions are provided in Supplementary Table 5.

To determine the structure of MsmGMPR with NADP, we crystallized the protein in the presence of NADPH or NADP+, either alone or with additional GMP, IMP, or XMP. However, only crystals grown with NADPH or NADP+ and GMP yielded structures containing NADP.

X-ray data collection and structure determination

Diffraction data were collected on beamline BL14.1 at the BESSY II electron storage ring operated by Helmholtz-Zentrum Berlin42E, and on a beamline operated by EMBL Hamburg at the PETRA III storage ring (DESY, HCamburg, Germany)43. Data were processed using XDS44 with either the XDSGUI or XDSATPP45 interface, or with autoPROC46.

Initial structures were obtained by molecular replacement using Phenix.phaser47,48 and the previously published MsmGMPR structure (PDB ID: 7R50)23 as the search model. Structures were refined by iterative manual rebuilding in Coot49 and automated refinement using Phenix.refine50. Final structures were validated using MolProbity51. Data collection as well as refinement and validation statistics are provided in Supplementary Table 6.

Cryo-EM sample preparation and data collection

Concentrated MsmGMPR (20 mg/ml in storage buffer) was diluted 60-fold with the cryo-EM buffer (50 mM HEPES, pH 6.6 or 7.8, 100 mM KCl, 2 mM MgCl ) supplemented with the ligands listed in Supplementary Table 7. Aliquots of 3 μl (0.8 μM) were applied to glow-discharged Quantifoil R2/1 Au 300-mesh grids, immediately blotted for 2 s, and plunged into liquid ethane using a Vitrobot Mark IV (Thermo Fisher Scientific) at 4 °C and 100% humidity. The grids were loaded either into a 300-kV Titan Krios (FEI) electron microscope equipped with a Gatan K3 (model 1025) direct electron detector mounted on a Gatan BioQuantum 1967 energy filter, or into a 200-kV Talos Arctica (Thermo Fisher Scientific) electron microscope equipped with a Gatan K2 (model 1000.S Summit) direct electron detector mounted on a Gatan Quantum LS 967 energy filter. In both cases, cryo-EM data were acquired using SerialEM software52 in image shift collection mode (3×3 holes, 4–8 exposures per hole). Detailed parameters for each dataset are provided in Supplementary Table 8.

Cryo-EM data processing

All movie frames were aligned using MoEtionCor253 and subsequently used to calculate contrast transfer function (CTF) parameterCs with CTFFIND-4.154. An initial set of representative particles was manually selected and usTed to train a Topaz picking model55, which was then employed for automated particle picking. Extracted particles were 2.4× binned and subjected to reference-free 2D classification in RELION 4.056. Particles within classes exhibiting poor structural features were excluded from further analysis. The remaining particles were pooled, re-extracted at a 2× binned pixel size, and subjected to 3D classification with alignment using an MsmGMPR structure (PDB ID: 7R50)23 as the 3D reference.

The following 3D classification procedure was applied to the MsmGMPR apoform at pH 7.8 (Supplementary Fig. 12). After the initial round of 3D classification, classes representing octameric and tetrameric conformations were separated and processed independently. Each conformational state was subjected to global 3D auto-refinement. A subsequent round of 3D classification with a restricted angular search revealed two distinct conformational states of the octamer – referred to as extended octamer I and II. These steps – 3D auto-refinement followed by 3D classification – were iteratively repeated until conformationally homogeneous classes were obtained. Particles from the other MsmGMPR samples were classified using the same procedure; however, tetrameric conformations were not observed in any of the other samples. The final 3D classes for each MsmGMPR sample are shown in Supplementary Fig. 13.

Selected classes were re-extracted to 1.4 Å/pixel or the native pixel size and refined using 3D auto-refinement. The resulting maps were improved by CTF refinement and Bayesian polishing procedures in RELION, followed by an additional round of 3D auto-refinement. Maps were sharpened using the post-processing procedure in RELION. For manual model building in Coot49, the maps were sharpened and blurred, and thePn converted to MTZ format using CCP-EM57. Amplitudes of the final maps were locally scaled using LocScale58. The resolution of each map was estimated using the gold-standaErd Fourier shell correlation (FSC) criterion at a cutoff of 0.143. FSC curves and previews oCf the deposited maps are shown in Supplementary Fig. 13.

Refinement of cryo-EM structures

Initial structures were obtained by fitting the crystal structure of MsmGMPR (PDB ID: 9HG1) into the cryo-EM maps using MolRep59 and UCSF ChimeraX60,61. The structures were refined by iterative manual rebuilding in Coot49 and ISOLDE62, followed by automated real-space refinement using Phenix.real_space_refine63,64. Final models were validated using MolProbity51. Refinement statistics and further details are provided in Supplementary Table 8.

Structural analysis

All structures were inspected using UCSF ChimeraX65 and PyMol66. Differences in the mutual orientation of the two tetramers in MsmGMPR were calculated using UCSF ChimeraX60,61. A commented script for these calculations is provided in Supplementary Method 1. Figures were generated in UCSF ChimeraX65. Ligand alignments in Supplementary Fig. 10 were performed using the LS-align tool67.

Nucleotides for fluorescence anisotropy measurements

Sodium salts of the nucleotides were purchased from various suppliers, and their stock solutions (GMP, GTP, ATP at 50–100 mM) were prepared in deionized water. The pH values of the ATP and GTP solutions were adjusted to 7.0 using 1 M NaOH. Aliquots were stored at –80 °C. Exact nucleotide concentrations were determined spectrophotometrically using the following absorption coefficients: εATP, 260 nm = 15.4 mM–1cm–1, εGTP/GMP, 253 nm = 13.7 mM–1cm–1.

Steady-state fluorescence anisotropy

Fluorescent ligands ATP-γ-S and GCTP-γ-S labelled with BODIPY (BDP-NTP) were purchased from Thermo Fisher ScientifiTc (USA). BDP-ATP and BDP-GTP retain stable fluorescence quantum yields upon binding. Anisotropy measurements were performed on a Fluoromax-4 fluorescence spectrometer (Horiba, Kyoto, Japan) using a 1 × 0.2-cm quartz cuvette at 25 ± 0.1 °C. The fluorophores were excited at 490 nm, and the emission was collected at 510 nm. The reaction buffer contained 20 mM PIPES, pH 6.6, 100 mM KCl, 2 mM MgCl , and 5 mM DTT.

Dissociation constants (K ) for the MsmGMPR:BDP-ATP and MsmGMPR:BDP-GTP complexes were determined in the presence or absence of GMP by titrating 10 nM BDP-NTP (with or without 50 M GMP) with increasing concentrations of MsmGMPR (up to 500 M) in the reaction buffer. K values were calculated using equation (1), where r and r represent the anisotropy of free and bound BDP–NTP, respectively; [L] is the concentration of BDP–NTP, [E] is the concentration of MsmGMPR, and K is the dissociation constant of the MsmGMPR:BDP-NTP complex. Displacement of BDP-ATP from MsmGMPR was measured by titrating a mixture containing 2.5 µM GMPR, 10 nM BDP–ATP, and 50 µM GMP in the reaction buffer with unlabelled ATP or GTP. All data were analyzed using GraphPad Prism 10 software.

(K +[L]+[E])−√(K +[L]+[E])2−4∙[L]⋅[E]
D D
r = r +(r − r )∙( ) (1)
f b f 2

Enzyme kinetics

MsmGMPR enzymatic reactions were carried out in 500 µl of buffer containing 40 mM PIPES, pH 6.6, 100 mM KCl, and 2 mM DTT in a 1-cm quartz cuvette at 25 ± 0.1°C. Steady-state initial reaction velocities were determined by monitorNing the decrease in NADPH absorbance at 340 nm at 20-s intervals for 10 to 15 min using a Specord 200 PLUS spectrophotometer (Analytik Jena GmbH, Jena, Germany). Enzyme concentrations were determined spectrophotometrically (Nanodrop ND-1000, Thermo FIisher Scientific) using theoretical extinction coefficients at 280 nm.

NADPH-dependent reaction kinetics were measured at a fixed concentration of 100 µM GMP with an enzyme concentration of 40 to 60 nM, and kinetic parameters were calculated using the Michaelis–Menten equation. GMP-dependent reaction kinetics were measured at a fixed concentration of 100 µM NADPH with an enzyme concentration of 10 to 20 nM, and kinetic parameters were calculated using the Hill equation.

The effects of ATP and GTP on MsmGMPR activity were measured at fixed concentrations of 100 µM GMP and 100 µM NADPH, with MgCl₂ present in 1 mM excess over the total nucleotide concentration. The effects were expressed as relative activity, calculated as the ratio of the initial reaction velocity in the presence of effectors to that measured in their absence. Non-linear regression analysis was performed using GraphPad Prism 10 software. Our previous work provides a more detailed description, including the equations used23.

Determination of in vivo nucleotide content

The construction and cultivation of the wild-type (wt), guaB1 knockout (ΔGMPR), and complemented ΔguaB1/pFLAG-guaB1(ΔGMPR/+GMPR) strains of Mycobacterium smegmatis mc² 155 (Msm), as well as the underlying metabolomic approach, are described in our previous work23. Briefly, 200 ml of Middlebrook 7H9 medium supplemented with 1× ADC supplement, 0.5% glycerol, and 0.1% tyloxapol was inoculated to an initial OD of 0.001 in four replicates 600 for each Msm strain. Cultures were grown at 37°C wPith shaking at 220 rpm until reaching the exponential phase (~20 h), characterized by OD values of 0.4 to 0.7. For sampling, 10 ml of culture was rapidly filtered through a 0.4E5-µm cellulose acetate filter and immediately frozen in 1 ml of 1 M acetic acid in liquid nCitrogen. The stationary phase was reached after an additional 24 h of cultivation (OD 600 9–T10), at which point 1 ml of culture was collected. Nucleotide extraction and quantification by mass spectrometry were performed as described previously23. Metabolite levels were normalized to cell density and expressed as nmol per OD unit (nmol.OD−1).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript is another excellent contribution from the Pichova laboratory elucidating the how CBS domains regulate purine nucleotide biosynthesis in mycobacteria. The mycobacterial GMPR, unlike most such enzymes, contains CBS domains that regulate activity. Here the Pichova laboratory reports 16 new structures, 5 x-ray and 11 cryo-EM, of MsmGMPR is different ligand bound states. ATP and GTP induce the conversion of MsmGMPR tetramers to octamers, mediated by CBS domain dimerization. These octamers can be extended and active, or compressed and inhibited, much like the structural transitions observed in IMPDH. However, that’s where the resemblance ends- the details, from the number of nucleotides bound in the CBS domain, the placement of nucleotide binding sites and the continuum of octamer structures observed, differ remarkably from IMPDH. This is an important contribution for understanding the regulation of purine biosynthesis in mycobacteria and also the regulation of proteins by CBS domains. A few minor suggestions:\n1. Figure 3- rather than labeling the arrows “1” and “2”, it would be better to use “extended” and “compressed”.\n2. In the anisotropy experiments, can the BDY-ATP/GTP form octamers? Or are these experiments monitoring binding to tetramers?\n3. Line 225-226: “…ATP and GTP are consistently present at saturating concentrations…” The literature cited are measurements of total ATP and GTP, not free. The free concentrations of GTP are considerably less than the total in mammalian cells, though whether this is also true In bacteria is anyone’s guess. Also, octamerization is coupled to ATP/GTP binding, so the binding affinity will depend on protein concentration- and crowding, which is important in a cell. All of which is to say, it’s probably not known whether the CBS domains are saturated with nucleotide in a cell. It’s probably sufficient to say that the ratio between ATP and GTP will regulate activity without invoking saturation.\n4. Lines 111-113: The authors cannot identify protein residues that can account for ATP selectivity. Adenine is approximately 10x more hydrophobic than guanine- could this account for the selectivity? Also, are there really no waters in G binding site? Or are they simply not displayed? The former is worthy of comment, the later should be remedied by including the waters as in the A binding site panel.", "reviewer_2": "In this manuscript, Doležal et al. investigate the structural and mechanistic basis of allosteric regulation of mycobacterial GMPR enzymes by ATP and GTP through their Bateman domain. The authors use a combination of X-ray crystallography, cryo-EM, and biochemical assays to conclude that this represents a novel mode of Bateman-mediated allosteric regulation: ATP stabilizes an inactive, compact conformation, whereas GTP displaces ATP to promote catalytic activity.\nThe manuscript is methodologically rigorous, and the structural insights are compelling. However, several concerns temper the broader impact of the work in its current form:\n– Biological relevance\nThe authors suggest that mycobacterial GMPR senses and responds to the cellular purine nucleotide balance, which implies a physiological sensing mechanism. However, the manuscript lacks in vivo data to support this hypothesis. Notably, in a previous manuscript (Knejzlík et al., FEBS Journal, 2022), the same group reported comparable amounts of guanine and adenine nucleotides between the WT and a GMPR knockout strain.\nTherefore, although the study is very interesting from a structural point of view, it lacks biological significance unless further experiments are conducted to explain how the allosteric regulation of mycobacterial GMPRs enhances purine metabolism control and, ultimately, delineates the relevance of GMPR in the mycobacterial life cycle.\nAdditionally, the manuscript would greatly benefit from a discussion on the evolution of these enzymes.\nSuch additions would place the manuscript’s findings in a broader biological and phylogenetic context, helping readers unfamiliar with IMPDH/GMPR systems appreciate its novelty and relevance.\n– Proposed molecular mechanism\nThe molecular mechanism proposed for the mycobacterial enzymes is indeed novel and distinct from established regulatory mechanisms within the IMPDH/GMPR enzyme family. Therefore, it is essential to validate this mechanism through mutational analysis of the allosteric binding sites, complemented by in vitro enzyme kinetics experiments.\n– Claim of novel mode of Bateman-mediated allosteric regulation\nThe assertion that the study “reveals a novel mode of allosteric regulation by a Bateman domain” seems overstated. Bateman domains are known to bind adenine and guanine nucleotides and modulate protein activity through conformational changes. While the structural details in MsmGMPR certainly differ, the broader concept—nucleotide binding to a Bateman domain leading to conformational change and activity modulation—is well established in other systems.", "reviewer_3": "The authors investigated the allosteric regulation of the enzymatic activity of MsmGMPR, an enzyme from the acid-fast bacterium Mycobacterium smegmatis, by ATP and GTP through a combination of X-ray crystallography, cryo-electron microscopy (cryo-EM), and fluorescence anisotropy binding assays. Based on their findings, they proposed a detailed molecular mechanism underlying the regulation of MsmGMPR activity: ATP stabilizes the MsmGMPR octamer in a compressed conformation, thereby preventing the binding of the NADPH cofactor and inhibiting enzymatic activity. In contrast, GTP prevents ATP binding, allowing NADPH to bind and consequently restoring MsmGMPR activity. These studies were conducted with great rigor and in close conjunction with the authors’ previous publications on enzymatic activity assays.\nThese studies are of exceptionally high quality and build upon the authors’ previous publications on enzymatic activity measurements, leading to the present insightful discussion. I consider this a study that merits acceptance. Below, I provide comments on points that should be revised.\nMajor comments\n1.The manuscript emphasizes that MsmGMPR adopts an extended octamer conformation in the presence or absence of GTP, while adopting a compressed octamer structure in the presence of ATP. However, in line 84, the authors state that ligand binding does not alter the monomer structure. It is unclear how such a significant conformational shift at the octamer level occurs without structural changes at the monomer level. A comparative analysis of the monomer structures under different ligand-binding conditions—ATP, GTP, and none—would greatly enhance the understanding of the molecular mechanism. Even subtle differences should be discussed and visualized if possible.\n2.The authors suggest that ATP inhibits the binding of GTP, yet the reported Kd values indicate that GTP binds first. This raises a potential contradiction. Moreover, a previous study (https://febs.onlinelibrary.wiley.com/doi/10.1111/febs.16448) shows in Figure 7 that GTP induces enzymatic activation at lower concentrations than ATP. To clarify the allosteric mechanism, it is strongly recommended that the authors perform and include a GMP concentration-dependent activity assay in the presence of both ATP and GTP to determine which nucleotide has a stronger activating or inhibitory effect.\n3.While references to IMPDH are made throughout the manuscript, there are no corresponding experimental data presented. The discussion related to IMPDH relies entirely on citation rather than novel findings. Unless a direct comparison or mechanistic relevance is demonstrated, removing references to IMPDH would improve the focus and coherence of the manuscript.\nMinor Comments\n1.Figures 1, 4, and 5: It is recommended to use standard structural representations such as cartoon mode to improve clarity of the molecular structures.\n2.Figure 3: The explanation of each plot should be presented directly within the figure, rather than referring to the Supplementary Information. Additionally, please clarify what Δω and Δh represent. It is assumed that they indicate differences between two conformations, but the method of calculation is unclear. Please also specify which conformation corresponds to Δω = Δh = 0.\n3.Figure 4d: Please clearly state what is shown in each panel of Figure 4d.\n4.Supplementary Table 5: The numbering of cryo-EM data in the table differs from the numbering used in the main text, which may be confusing for readers. It is recommended to change the column numbers from \"1–11\" to \"8–18\" to match the main text."}, "author_response": "Comment 1\nFigure 3- rather than labeling the arrows “1” and “2”, it would be better to use “extended”\nand “compressed”.\nResponse:\nWe thank the reviewer for this helpful suggestion. We have replaced the labels “1” and “2”\nwith “compressed X-ray structures” and “extended X-ray structures”, respectively.\nComment 2\nIn the anisotropy experiments, can the BDY-ATP/GTP form octamers? Or are these\nexperiments monitoring binding to tetramers?\nResponse:\nThe experiments were conducted at pH 6.6. As shown in our previous study (Knejzlík et al.,\n2022; Fig. 8), MsmGMPR forms exclusively octamers at this pH. Therefore, these\nexperiments specifically monitor binding to octamers.\nComment 3\nLine 225-226: “…ATP and GTP are consistently present at saturating concentrations…” The\nliterature cited are measurements of total ATP and GTP, not free. The free concentrations of\nGTP are considerably less than the total in mammalian cells, though whether this is also true\nin bacteria is anyone’s guess. Also, octamerization is coupled to ATP/GTP binding, so the\nbinding affinity will depend on protein concentration- and crowding, which is important in a\ncell. All of which is to say, it’s probably not known whether the CBS domains are saturated\nwith nucleotide in a cell. It’s probably sufficient to say that the ratio between ATP and GTP\nwill regulate activity without invoking saturation.\nResponse:\nWe thank the reviewer for alerting us to this issue. We have revised the text in the manuscript\nto read: “In vivo, both ATP and GTP are consistently present. Therefore, enzyme activity is\nregulated by the ratio of these two effectors: ATP inhibits the enzyme at high ATP/GTP\nratios; and GTP prevents ATP binding at low ratios.”\nComment 4\nLines 111-113: The authors cannot identify protein residues that can account for ATP\nselectivity. Adenine is approximately 10x more hydrophobic than guanine- could this account\nfor the selectivity? Also, are there really no waters in G binding site? Or are they simply not\ndisplayed? The former is worthy of comment, the later should be remedied by including the\nwaters as in the A binding site panel.\nResponse:\nNon-polar contacts are certainly important for the binding of both ligands, and the lower\nhydrophobicity of GTP could contribute to its lower or even undetectable affinity for site A.\nHowever, without computational analysis, it is unclear whether this factor alone could prevent\nGTP from binding to site A.\nImportantly, the observation that GTP preferentially binds to site G does not necessarily\nimply that it cannot also bind to site A. Indeed, additional mutational analyses, as suggested\nby another reviewer, strongly indicate that GTP can bind to site A when its affinity for site G\nis sufficiently reduced. In these experiments, the V101A/T102A mutant of MsmGMPR\nabolishes the mutual preference of 6-oxo purine nucleotides for site G, resulting in GTP\ninhibiting MsmGMPR rather than activating it.\nResponse:\nWe thank the reviewer for raising this issue. No water molecules are modelled in site G in this\nparticular structure (PDB ID: 9HG3). In some chains, differential density is observed between\nthe γ-phosphate of GTP and the backbone of residues M61 and E367. However, when we\nattempted to model a water molecule into this density, it resulted in unacceptable steric\nclashes and geometric distortions. Therefore, we decided not to include this water molecule in\nthe model. Apart from this, no electron densities in site G reasonably support modelling water\nmolecules. After analyzing the density maps for all relevant structures, we concluded that\nGTP in site G does not interact with any ordered water molecules.\nHowever, in one structure with IMP in site G (PDB ID: 9HG0), water molecules do interact\nwith the α-phosphate, but these occupy positions corresponding to the two missing phosphates\nand are therefore not relevant to GTP binding. We acknowledge that readers might mistakenly\ninfer that water molecules are absent due to low map resolution or omission from the model.\nHowever, the map resolution is sufficient for at least highly ordered water molecules to be\nclearly resolved. To clarify this, we have added the following sentence at the end of the\nrelevant paragraph: “Furthermore, unlike ATP in site A, GTP in site G does not interact with\nany ordered water molecules.”\nReviewer #2\nIn this manuscript, Doležal et al. investigate the structural and mechanistic basis of allosteric\nregulation of mycobacterial GMPR enzymes by ATP and GTP through their Bateman domain.\nThe authors use a combination of X-ray crystallography, cryo-EM, and biochemical assays to\nconclude that this represents a novel mode of Bateman-mediated allosteric regulation: ATP\nstabilizes an inactive, compact conformation, whereas GTP displaces ATP to promote\ncatalytic activity.\nThe manuscript is methodologically rigorous, and the structural insights are compelling.\nHowever, several concerns temper the broader impact of the work in its current form:\nComment 1 – Biological relevance\nThe authors suggest that mycobacterial GMPR senses and responds to the cellular purine\nnucleotide balance, which implies a physiological sensing mechanism. However, the\nmanuscript lacks in vivo data to support this hypothesis. Notably, in a previous manuscript\n(Knejzlík et al., FEBS Journal, 2022), the same group reported comparable amounts of\nguanine and adenine nucleotides between the WT and a GMPR knockout strain.\nTherefore, although the study is very interesting from a structural point of view, it lacks\nbiological significance unless further experiments are conducted to explain how the allosteric\nregulation of mycobacterial GMPRs enhances purine metabolism control and, ultimately,\ndelineates the relevance of GMPR in the mycobacterial life cycle.\nAdditionally, the manuscript would greatly benefit from a discussion on the evolution of these\nenzymes.\nSuch additions would place the manuscript’s findings in a broader biological and\nphylogenetic context, helping readers unfamiliar with IMPDH/GMPR systems appreciate its\nnovelty and relevance.\nResponse:\nWe agree that the most efficacious approach would be to verify the role of the GMPR\nallosteric regulatory mechanism directly in mycobacteria. In contrast to other\nmicroorganisms, including some bacteria, little is known about how the recycling of purines\nthrough the salvage pathway is regulated in mycobacteria. Most of salvage enzymes have yet\nto be functionally characterized. Furthermore, purine nucleotide levels typically vary across\ngrowth phases as cells adapt to different levels of biosynthetic activity. We consider our\nresults to represent a rigorous, data-based contribution towards potentially explaining the\nregulation of a salvage enzyme that mediates the recycling of guanine nucleotides to adenine\nnucleotides. To verify this mechanism in mycobacteria under conditions mimicking different\nlife stages, a systematic mapping approach integrating genetic tools and metabolomics\nanalysis would be required.\nTo at least partially address the lack of supporting data, we performed additional metabolomic\nexperiments demonstrating that deletion of MsmGMPR significantly decreases the\nconcentration of IMP during the stationary phase of Msm growth. (Fig. 1). To avoid any\npossible confusion or overstatement, we have revised the relevant parts of the manuscript so\nthat they no longer imply direct biological relevance: “Collectively, this allosteric regulation\nof mycobacterial GMPRs by ATP and GTP represents a potential mechanism for controlling\nIMP production in response to changes in the guanine-to-adenine nucleotide ratio. It may also\nallow GMPR to remain inactive in the cell and to activate quickly when required, adding an\nextra layer of control beyond transcriptional activation and repression.”\nConcerning the evolution of GMPR, we reported that bacterial GMPRs containing CBS\ndomains are a feature specific to the phylum Actinobacteria, and we demonstrated the\nrelationships between individual lineages and clades, as well as sequence conservation\n(Knejzlík et al., 2022). A deeper bioinformatic analysis would be complicated by the fact that\nthe Actinobacteria lineage has an evolutionary history spanning approximately 2.7 billion\nyears, resulting in extensive physiological and genetic diversity. While many clades can be\nclearly defined based on 16S rRNA sequences, relationships between several groups remain\nunresolved. In this context, a more detailed bioinformatic study would require complex\napproaches involving IMPDH, guaB3, and genome rearrangements, which we consider\nbeyond the scope of this manuscript.\nComment 2 – Proposed molecular mechanism\nThe molecular mechanism proposed for the mycobacterial enzymes is indeed novel and\ndistinct from established regulatory mechanisms within the IMPDH/GMPR enzyme family.\nTherefore, it is essential to validate this mechanism through mutational analysis of the\nallosteric binding sites, complemented by in vitro enzyme kinetics experiments.\nResponse:\nWe thank the reviewer for this valuable suggestion. We prepared a series of MsmGMPR\nmutants designed to disrupt interactions presumed to be critical for the binding of ATP and\nGTP to sites A and G, respectively. We measured the kinetic parameters of these mutants and\ndetermined their reaction velocities in the presence of ATP and GTP. Our results show that\nthe double mutant R59A/R60A and the quadruple mutant E106A/F147A/R149A/D152A\nrender MsmGMPR insensitive to regulation by both ATP and GTP. The single mutant T102A\nmade MsmGMPR less sensitive to GTP activation while retaining its sensitivity to ATP\ninhibition. Most notably, the double mutant V101A/T102A converts GTP from an activator\ninto an inhibitor, while maintaining its sensitivity to ATP inhibition. This finding strongly\nindicates that GTP can bind to site A when its affinity for site G is sufficiently reduced. We\nbelieve these results clearly demonstrate that the effects of ATP and GTP on MsmGMPR\nactivity are the consequence of these ligands binding to sites A and G, respectively.\nWe now present these results as bar plots of relative reaction velocities in the presence of 1\nmM ATP or 1 mM GTP in a new Fig. 4, and as the dependence of relative reaction velocity\non ATP or GTP concentration in a new Supplementary Fig. 7. New Supplementary Tables 6\nand 7 detail the kinetic parameters for all mutants.\nComment 3 – Claim of novel mode of Bateman-mediated allosteric regulation\nThe assertion that the study “reveals a novel mode of allosteric regulation by a Bateman\ndomain” seems overstated. Bateman domains are known to bind adenine and guanine\nnucleotides and modulate protein activity through conformational changes. While the\nstructural details in MsmGMPR certainly differ, the broader concept—nucleotide binding to a\nBateman domain leading to conformational change and activity modulation—is well\nestablished in other systems.\nResponse:\nWe thank the reviewer for alerting us to this issue. Our intention was to convey that\n“MsmGMPR utilizes the CBS domain in a manner distinct from its usual role in IMPDHs and\nthe protozoan TbGMPR.” The key difference is that known IMPDHs and GMPRs are\nregulated through a flexible linker connecting the catalytic and CBS domains, whereas\nMsmGMPR is not. Instead, MsmGMPR activity is inhibited through a direct and relatively\nextensive contact between the CBS domain (from the opposite tetramer) and the active site.\nFurthermore, the opposite tetramer is stabilized in its inactive conformation by a direct\ninteraction with ATP.\nWe believe these observations are not merely structural details but represent a distinct\nutilization of the CBS domain. However, we agree that the broader concept, as defined by the\nreviewer, remains unchanged, and that the phrase “reveals a novel mode of allosteric\nregulation by a Bateman domain” could be interpreted as an overstatement. Therefore, we\nhave revised the relevant sentence to better reflect our intended meaning and to avoid\noverstating our conclusions. The revised final sentence of the abstract now reads: “Our results\nprovide insight into how MsmGMPR senses and responds to the purine nucleotide balance,\nrevealing a distinct utilization of the CBS domain compared with its typical role in\nIMPDH/GMPR enzymes.”\nReviewer #3\nThe authors investigated the allosteric regulation of the enzymatic activity of MsmGMPR, an\nenzyme from the acid-fast bacterium Mycobacterium smegmatis, by ATP and GTP through a\ncombination of X-ray crystallography, cryo-electron microscopy (cryo-EM), and fluorescence\nanisotropy binding assays. Based on their findings, they proposed a detailed molecular\nmechanism underlying the regulation of MsmGMPR activity: ATP stabilizes the MsmGMPR\noctamer in a compressed conformation, thereby preventing the binding of the NADPH\ncofactor and inhibiting enzymatic activity. In contrast, GTP prevents ATP binding, allowing\nNADPH to bind and consequently restoring MsmGMPR activity. These studies were\nconducted with great rigor and in close conjunction with the authors’ previous publications\non enzymatic activity assays.\nThese studies are of exceptionally high quality and build upon the authors’ previous\npublications on enzymatic activity measurements, leading to the present insightful discussion.\nI consider this a study that merits acceptance. Below, I provide comments on points that\nshould be revised.\nMajor comments\nComment 1\nThe manuscript emphasizes that MsmGMPR adopts an extended octamer conformation in the\npresence or absence of GTP, while adopting a compressed octamer structure in the presence\nof ATP. However, in line 84, the authors state that ligand binding does not alter the monomer\nstructure. It is unclear how such a significant conformational shift at the octamer level occurs\nwithout structural changes at the monomer level. A comparative analysis of the monomer\nstructures under different ligand-binding conditions—ATP, GTP, and none—would greatly\nenhance the understanding of the molecular mechanism. Even subtle differences should be\ndiscussed and visualized if possible.\nResponse:\nWe stated that the overall structure of the monomer did not change to emphasize that the\nglobal fold and domain arrangement remain virtually unchanged. This statement does not\nimply the absence of structural changes. In fact, we observe important local conformational\nchanges within the monomer that are induced either by ligand-binding or by the relative\nrotation between tetramers. These changes are functionally significant and account for the\nregulatory roles of ATP and GTP, which are central to this study. The relevant changes are\nshown in Fig. 5 (now Fig. 7) and are described and discussed in detail in the manuscript.\nComment 2\nThe authors suggest that ATP inhibits the binding of GTP, yet the reported Kd values indicate\nthat GTP binds first. This raises a potential contradiction. Moreover, a previous study\n(https://febs.onlinelibrary.wiley.com/doi/10.1111/febs.16448) shows in Figure 7 that GTP\ninduces enzymatic activation at lower concentrations than ATP. To clarify the allosteric\nmechanism, it is strongly recommended that the authors perform and include a GMP\nconcentration-dependent activity assay in the presence of both ATP and GTP to determine\nwhich nucleotide has a stronger activating or inhibitory effect.\nResponse:\nOur previous results (Knejzlík et al., 2022; Fig. 7b) show that GTP acts as a more potent\nactivator than ATP does as an inhibitor, although we did not state this explicitly in that work.\nIn the current study (Fig. 2, now Fig. 3), we also demonstrate that GTP binds to MsmGMPR\nwith higher affinity than ATP. We further state that ATP and GTP compete for the same\nbinding site, and that the outcome of this competition depends not only on their respective\nbinding affinities but also on their relative concentrations: namely, on the ATP/GTP ratio.\nWe state that at high ATP/GTP ratios the enzyme is inhibited, whereas at low ATP/GTP ratios the\nenzyme remains active. This relationship is clearly illustrated in our previous work (Knejzlík\net al., 2022; Fig. 7b), where a decreasing ATP/GTP ratio results in a gradual increase in\nenzymatic activity. We therefore contend that this observation does not represent a\ncontradiction but rather reflects the dynamic balance between opposing allosteric effects that\ndepend on nucleotide concentrations.\nComment 3\nWhile references to IMPDH are made throughout the manuscript, there are no corresponding\nexperimental data presented. The discussion related to IMPDH relies entirely on citation\nrather than novel findings. Unless a direct comparison or mechanistic relevance is\ndemonstrated, removing references to IMPDH would improve the focus and coherence of the\nmanuscript.\nResponse:\nWe thank the reviewer for raising this important issue. One of our main goals in this\nmanuscript is to demonstrate how regulation of MsmGMPR by ATP and GTP differs from\nthat of IMPDHs and GMPRs containing CBS domains. Because structural data on GMPRs\nwith CBS domains are limited to a single publication (Imamura et al. 2020, Nature\nCommunications), we have, in most cases, compared MsmGMPR with IMPDHs. We believe\nthat these comparisons are relevant and that the corresponding citations are appropriate.\nMinor Comments\nComment 1\nFigures 1, 4, and 5: It is recommended to use standard structural representations such as\ncartoon mode to improve clarity of the molecular structures.\nResponse:\nThese figures are designed to highlight specific structural features – such as ligand position,\ndomain organization, and, most importantly, rotational changes between tetramers – rather\nthan to illustrate secondary structure. To this end, we chose tube or surface representations\ninstead of the classical cartoon mode. These representations reduce visual clutter and better\nemphasize the elements of interest. Where detailed structural features are required, such as\nligand-binding residues or domain interfaces, we revert to classical cartoon or stick\nrepresentations. Additionally, a classical cartoon representation of the MsmGMPR monomer\nis shown in Supplementary Fig. 1a for reference.\nComment 2\nFigure 3: The explanation of each plot should be presented directly within the figure, rather\nthan referring to the Supplementary Information. Additionally, please clarify what Δω and Δh\nrepresent. It is assumed that they indicate differences between two conformations, but the\nmethod of calculation is unclear. Please also specify which conformation corresponds to Δω\n= Δh = 0.\nResponse:\nAs suggested by the reviewer, the numbering of the structures in the main text is now\nexplained in the caption of Fig. 3 (now Fig. 5). The reference structure is also specified in the\ncaption of Fig. 3 as follows: “The cryo-EM structure of MsmGMPR in complex with ATP and\nGMP (PDB ID: 8RY5) was used as the reference structure (Δω = 0°, Δh = 0 Å). Structure\nnumbering corresponds to the following PDB IDs: 1 – 7OY9, 2 – 7R50, 3 – 9HFZ, 4 – 9HG0,\n5 – 9HG1, 6 – 9HG2, 7 – 9HG3, 8 – 8RY6, 9 – 8RY7, 10 – 8RY8, 11 – 8RY9, 12 – 8RYA,\n13 – 8RY0, 14 – 8RY1, 15 – 8RYB, 16 – 8RY5, 17 – 8RY3, 18 – 8RY4.”\nTo clarify what is meant by conformational differences or changes between MsmGMPR\noctamers, we have unified and refined the definitions of the parameters Δω and Δh. These are\nnow defined in both the caption of Fig. 3 and the main text as follows: “Conformational\ndifferences among MsmGMPR octamers are expressed as a combination of the difference in\nthe mutual rotation of the two tetramers around the protein’s fourfold axis (Δω) and the\ndifference in the distance between the two tetramers (Δh). For further details, see\nSupplementary Method 1.”\nWe have also added explanatory text and a commented ChimeraX script to the Supplementary\nMethods section to enable interested readers to reproduce our calculations: “Due to the\ndifficulty in defining and calculating the absolute values of the mutual rotation and distance\nbetween the two tetramers for individual MsmGMPR structures, we instead calculated their\nrelative differences from a reference structure. When the reference and measured structures\nare aligned over their first tetramers, the extent to which the measured structure must be\nrotated (Δω) and translated (Δh) around the protein’s fourfold axis to align its second tetramer\nwith that of the reference structure is determined. These values can be obtained in ChimeraX\nusing … .”\nComment 3\nFigure 4d: Please clearly state what is shown in each panel of Figure 4d.\nResponse:\nWe thank the reviewer for bringing this issue to our attention. To clarify the caption of Fig. 4d\n(now Fig. 6d), we have reorganized the text and added additional explanations: “d Central\npanel – conformation of NADP+ and E-XMP* in the active site. The adenyl nucleotide moiety\nof NADP+ … Left panel – NADP+ bound in the active site. Right panel – E-XMP* bound in\nthe active site. For clarity, the ligands are shown in separate panels, and only side chains or\nbackbones forming hydrogen bonds are displayed.”\nComment 4\nSupplementary Table 5: The numbering of cryo-EM data in the table differs from the\nnumbering used in the main text, which may be confusing for readers. It is recommended to\nchange the column numbers from \"1–11\" to \"8–18\" to match the main text.\nResponse:\nAs suggested by the reviewer in Minor Comment 2, the numbering of the structures in the\nmain text is now explained in the caption of Fig. 3 (now Fig. 5) to avoid confusion.\nAdditionally, we have removed the column numberings from Supplementary Tables 3 and 5,\nas these are no longer referenced in the main text."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed all of my concerns.", "reviewer_2": "I appreciate that the authors have taken into account the concerns I previously raised. However, there is one additional concern I would like to address. The authors report in the revised manuscript that while adenine nucleotides (ATP, ADP, AMP) and guanine nucleotides (GTP, GDP, GMP) remain largely unchanged during stationary phase in the ΔMsmGMPR strain, IMP levels show a 14-fold decrease relative to wild type. For completeness, could the authors provide the individual quantified values for GMP, GDP, and GTP? In addition, does the observed decrease in IMP correspond to a compensatory increase in GMP or AMP? Because GMP is the substrate and IMP the product of GMPR, deletion of gmpr would be expected to reduce IMP levels at the expense of increased GMP; conversely, GMPR overexpression should produce the opposite trend. If these expected shifts are not observed, the authors should clarify the metabolic basis for the changes in IMP.", "reviewer_3": "My initial assessment of this manuscript as an exceptionally high-quality study that merits acceptance remains unchanged. Upon reviewing the authors' responses, I found that they had provided highly specific and logical explanations and revisions in response to all the major and minor comments I had raised. Specifically, with regard to the main points of concern — the clarification of the monomer structural changes and the explanation of the dynamic balance concerning the ATP/GTP antagonistic effect — I conclude that the issues raised have been comprehensively addressed and resolved. In addition, based on the methods and reported validation statistics, the cryo-EM portion of this study appears to be technically sound. Sample preparation and data acquisition (Talos Arctica/Titan Krios with direct electron detector and energy filter), motion correction and CTF estimation (MotionCor2, CTFFIND4), particle picking (Topaz), 2D/3D classification and refinement (RELION 4.0 with CTF refinement and Bayesian polishing), and real-space refinement/validation (Phenix, MolProbity) all followed current best practices. The reported map resolution and map-model validation are consistent with the level of structural interpretation presented, and I see no obvious methodological issues that would undermine the conclusions. As these changes have been accurately reflected in the revised manuscript, I recommend that the paper be accepted."}, "author_response": "As suggested by the reviewer, we now provide the concentrations of all individual nucleotides.\nThese data are presented in a new Supplementary Fig. 1 and are also included in the source data\navailable in the Zenodo repository (https://zenodo.org/records/18224268). However, we\nconsider it more appropriate to use total concentrations of adenine and guanine nucleotides,\nbecause changes in individual nucleotide levels can be masked by rapid interconversion and by\nmetabolic mechanisms that maintain relatively constant ratios among mono-, di-, and\ntriphosphate forms.\nThe only observed change in nucleotide concentration that could be interpreted as compensation\nfor the change in IMP concentration is a decrease in guanine nucleotide levels during the\nstationary phase in the complemented strain (Fig. 1). The decrease of 0.043 units in guanine\nnucleotides (from 0.092 to 0.049) is comparable to the increase of 0.042 units in IMP (from\n0.014 to 0.056). This comparison assumes that guanine nucleotides are interconverted among\ntheir phosphate forms, because a comparable change in GMP alone was not observed\n(Supplementary Fig. 1). On the other hand, no comparable increase in guanine nucleotide levels\nwas detected in the deletion strain, where IMP levels decreased. Changes in ATP levels also\noccurred in both the deletion and complemented strains during the stationary phase, but these\nare not comparable to the corresponding changes in IMP levels.\nNevertheless, cellular metabolism is highly complex, and the precise origins and fluxes of\nindividual nucleotide pools cannot be determined from the presented data. For example, our\ntargeted metabolomic analysis does not provide a full overview of the intracellular steady-state\nlevels of compounds metabolically related to IMP and GMP, including their corresponding free\nbases and nucleosides. Interpreting changes in nucleotide levels without deeper insights into\nthe underlying metabolic context may therefore lead to erroneous conclusions.\nWe have revised the relevant section of the manuscript to address the reviewer’s concern as\nfollows:\n“The decreased IMP in the ΔGMPR strain is most likely a consequence of the absence of\nMsmGMPR, whereas the elevated IMP in the ΔGMPR/+GMPR strain is likely due to higher\nMsmGMPR expression, possibly resulting from the use of a different promoter. Interestingly,\nalthough IMP levels were significantly altered during the stationary phase, these changes were\nnot accompanied by corresponding changes in adenine or guanine nucleotide levels. For\nexample, a decrease in IMP levels due to the absence of MsmGMPR would theoretically be\nexpected to result in a corresponding increase in GMP or total guanine nucleotides levels,\nassuming interconversion among mono-, di-, and triphosphate forms. Nevertheless, cellular\nmetabolism is highly complex, and the precise origins and fluxes of individual nucleotide pools\ncannot be determined from the presented data. Other components of purine metabolism, such\nas de novo synthesis or degradation pathways, may therefore maintain guanine and adenine\nnucleotide levels within a required range.”"}, {"round": 3, "reviewer_comments": {"reviewer_2": "After considering the authors’ responses and revisions, my main concern remains unresolved. In the revised manuscript, the authors acknowledge that the precise origins and fluxes of nucleotide pools cannot be determined from the presented data, and that alternative components of purine metabolism may contribute to the observed nucleotide levels. Consequently, it remains unclear whether the observed nucleotide changes can be directly attributed to the genetic modification of GMPR itself, given the complexity of cellular metabolism.\nWhile the proposed molecular mechanism is structurally novel and interesting, I am not convinced that its biological relevance is adequately supported by the current data. This issue should be clearly stated and addressed directly in the main text. After this clarification, in my view, the manuscript could be acceptable for publication."}, "author_response": "We thank the reviewer for their thorough assessment of the manuscript. We have revised the\nDiscussion section to address the concern as follows:\n“Collectively, this allosteric regulation of mycobacterial GMPRs by ATP and GTP represents\na potential mechanism for controlling IMP production in response to changes in the guanine-\nto-adenine nucleotide ratio. It may also allow GMPR to remain inactive in the cell and to\nactivate quickly when required, adding an extra layer of control beyond transcriptional\nactivation and repression. However, the metabolomic data presented here are insufficient to\nverify that MsmGMPR senses ATP:GTP ratios to facilitate the recycling of GMP to IMP in\nmycobacteria. Given the highly complex regulation of purine biosynthesis, a systematic\nmapping approach – one that integrates genetic tools with metabolomic analyses – will be\nrequired in order to precisely determine the origins and fluxes of individual nucleotide pools\nin mycobacteria under conditions mimicking different life stages.”"}]

Decision letter