Structural and molecular basis for allosteric regulation and catalytic coupling of human phosphoribosylformylglycinamidine synthase

Domain: neuroscience
Published: 2026-02-13
Editorial outcome: minor_revision

Abstract
Purine nucleotides are ubiquitous molecules essential for all life. The de novo biosynthesis of purines is a metabolic dependency that is frequently reprogrammed in cancers and is a well-established target for chemotherapies, immune modulation and antivirals. Here, we report cryo-electron microscopy structures of the multi-domain human phosphoribosylformylglycinamidine synthase, a central purine biosynthetic enzyme and foundational feature of the purinosome metabolon. These data capture, the proposed iminophosphate intermediate and provide the structural elucidation of an ammonia channel connecting the active sites of the glutaminase and synthase domains. Analysis of this series of structures and the accompanying biochemical data also reveal the molecular features and transient conformational changes that underlie allosteric regulation and catalytic coupling of the domains. This data resolves several longstanding mechanistic questions about this enzyme class and provides a strong foundation for therapeutic development.

Introduction
Purine nucleotides play essential cellular roles including the transfer of genetic information, energy storage, signaling, and as cofactors in enzyme-catalyzed reactions. They are critical to numerous physiological processes such as neurotransmission, the immune response, the regulation of cardiac function, and growth and development1–8. Purine derivatives and antimetabolites that target purine metabolism have long been used successfully as antivirals, immune modulators and chemotherapeutics9–12. The reliance on nucleotide triphosphates (NTPs) and their deoxy counterparts (dNTPs) for rapidly proliferating cells is due to the fact that these molecules are rate-limiting for numerous physiological processes, including transcription, energy storage, DNA replication and repair, ribosome biogenesis, and post-translational protein glycosylation. Increased biosynthesis of (d)NTPs is a universal, and highly druggable, feature of cancer13,14. Not surprisingly, many oncogenic drivers upregulate nucleotide biosynthesis, and these increases have been shown to promote immune evasion, metastasis, and the development of resistance to chemotherapy14–16.

Inhibitors of nucleotide biosynthetic enzymes were among the first antineoplastic agents developed and remain the core therapy for many cancer indications10,17. Despite the critical role that they play, drugs that inhibit their biosynthesis are clinically well tolerated, likely due to the presence of multiple, independent pathways to synthesize and recycle nucleotides. Numerous recent, unbiased genomic, chemical and metabolomic screening approaches have validated nucleotide metabolism as a crucial cancer vulnerability, sparked renewed interest in the field, and helped motivate many clinical trials and drug development campaigns18–26.

Purine levels are carefully regulated by coordinated control over two complementary pathways, the de novo biosynthetic pathway and the salvage pathway. The salvage pathway recycles preformed purine bases and ribose sugars to form nucleotides, while the de novo pathway builds the purine ring from small building blocks in a multi-step, energy-intensive pathway. Compared to purine salvage, the de novo biosynthetic pathway is more highly regulated and has a greater effect on cell growth27,28. Regulation of de novo biosynthesis occurs at the gene and protein level through several feed-forward and feedback mechanisms27–30. Six cytosolic enzymes catalyze the ten highly conserved steps of this pathway31,32. In addition to generating nucleotide building blocks, an increasing number of intermediates from this pathway have been implicated in important signaling cascades and non-metabolic processes33–37.

Phosphoribosylformylglycinamidine synthase (gene name PFAS, also referred to as FGAM synthase, FGAMS or PurL) catalyzes the fourth step in the de novo biosynthesis of purine nucleotides. This enzyme is directly involved in regulating purine biosynthetic rate and is known to be a core structural element of the purinosome, a biomolecular condensate that provides spatiotemporal control over purine metabolism31,32,38–41. Increased human FGAMS (HsFGAMS) expression has also been shown to promote cancer growth. In liver cancer, specifically, high expression of this protein is strongly correlated with lower survival rates13,42. In addition, FGAMS has been directly implicated in RAS-dependent cancer cell colony formation and tumor growth18,43.

Purine metabolism is regulated post-translationally by several distinct mechanisms, with HsFGAMS playing a major role in a number of these. As with other metabolic pathways, several of the nucleotide products regulate early steps in the pathway through feedback inhibition44–46. Biosynthetic flux, spatiotemporal regulation of activity, and control over reactive intermediates are also regulated, in part, by the formation and localization of the purinosome32,38–40,47. Control over purine biosynthetic enzyme kinetics, at the individual protein level, also occur through allosteric mechanisms whereby ligand binding at a location distal to the active site induces conformational changes that influence activity48–50. All but one of the enzymes of the de novo purine biosynthetic pathway are multi-domain proteins that catalyze multi-step reactions4. The activity of several of these multi-domain proteins is regulated through interdomain allostery4,44,51,52. The enzyme that catalyzes the first committed step in the pathway, phosphoribosylpyrophosphate amidotransferase, couples activity at two distinct active sites through the allosteric regulation of a channel connecting the two sites53. Such interdomain channeling functions as a means to protect labile intermediates and facilitate direct handoff of reaction products54. Parallel allosteric control of the distinct binding sites and modulation of structural features of the channel can promote efficient catalytic coupling and be a means to subtly regulate enzyme activity.

Eukaryotic FGAMS is a multi-domain protein, representative of the large PurL (lgPurl) variants of this protein, as compared to the small PurL (smPurL) proteins, which are expressed as three distinct gene products (Fig. 1a)4,55. FGAMS catalyzes the conversion of formylglycinamide ribonucleotide (FGAR) to formylglycinamidine ribonucleotide (FGAM) (Fig. 1b). In the first, amidotransferase, reaction, ammonia is liberated from glutamine and directed to the synthase domain, where it is added to FGAR in an ATP-dependent reaction56. The transfer of ammonia between the two active sites is facilitated by a putative ammonia channel56. The presence of the channel is supported by computational and kinetic analyses, although it has not yet been physically observed nor is it clear how channel entry and exit are regulated57. FGAMS also harbors an N-terminal domain with an, as yet, unestablished role.

In this work, we describe three Cryo-EM structures of HsFGAMS, from 3.2 to 3.3 Å resolution, that capture distinct functional states and important pathway intermediates. These structures of FGAMS reveal the location and architecture of the ammonia channel and capture the proposed iminophosphate intermediate. These data also implicate the N-terminal domain as a key element involved in allosteric control of FGAMS activity. The structures and supporting kinetic analyses reveal a putative mechanism that couples the chemistry of the active sites through substrate-induced conformational changes, controlling formation of the ammonia channel as well as substrate entry and exit.

Results
Expression and initial characterization of HsFGAMS
Full details about the cloning, expression, and purification of HsFGAMS are included in the supplementary information. Briefly, HsFGAMS was expressed as a C-terminally FLAG-tagged protein from HEK 293T cells and purified using immobilized affinity chromatography (Fig. 1c). Unlike the homologs from E. coli and S. typhimurium, which show a high level of basal glutaminase activity55,58–60, HsFGAMS does not exhibit observable glutaminase activity in the absence of the co-substrate FGAR (Fig. 1d). With FGAR present, the enzyme turns over glutamine with a KM of 189 ± 20 μM and kcat of 1.0 ± 0.04 s−1. Although the synthase activity of FGAMS is difficult to measure directly, we recently developed a mechanism-based assay that measures dephosphorylation of 6,8-difluoro-4-methylumbelliferyl phosphate as a surrogate for synthase activity42. Using this assay, we determined that an anomeric mixture of the substrate, α/β-FGAR, competitively binds to HsFGAMS with an IC50 of 148 ± 4 μM. As FGAMS is known to be selective for the β-FGAR anomer59, the KM for this substrate is expected to be approximately half this value.

Cryo-EM structure of the HsFGAMS-AMPPNP binary complex
To elucidate the molecular features underlying FGAMS function and as a basis for future drug discovery efforts, we sought to solve structures of FGAMS in functionally relevant states. Prior studies of S. typhimurium FGAMS (StPurL) include unliganded (3UGJ), glutamine-bound (1T3T), and glutamine/AMP-PNP (non-hydrolyzable ATP analogue) bound (3UMM) X-ray crystal structures60. Using Cryo-EM, we solved the structures of (1) HsFGAMS with AMP-PNP bound (referred to as the binary complex); (2) HsFGAMS in complex with the iminophosphate reaction intermediate and adenosine diphosphate (ADP) (referred to as the ternary complex); and 3) HsFGAMS with glutamyl thioester, AMP-PNP and FGAR bound (referred to as the quaternary complex) (Fig. 2). The ligand combinations were chosen to map the distinct steps of the catalytic mechanism, stabilize transition/intermediate states and to identify the long-range allosteric regulation network essential for the enzyme to reach its catalytically active state.

The binary complex was prepared by incubating HsFGAMS with 5 mM AMP-PNP on ice for 4 hours prior to grid preparation. We obtained an initial 3.35 Å map using C1 symmetry (Supplementary Figs. 1–3), with further 3D classification followed by non-uniform refinement yielding an improved 3.31 Å map. The final model was built using the predicted HsFGAMS structure from AlphaFold61 as a starting model. Full data collection parameters and refinement statistics are provided in Supplementary Table 1. Analysis of the binary complex structure confirms that HsFGAMS is a single polypeptide organized into three domains (Fig. 2a, b), similar to the homologous prokaryotic lgPurL structures (Supplementary Fig. 4)56. In addition to the glutaminase domain (GD, residues 1005–1335, Fig. 2 and Supplementary Fig. 5a) and synthase domain (SD, residues 173–1004, Fig. 2 and Supplementary Fig. 5b), is an N-terminal domain (NTD, Fig. 2 and Supplementary Fig. 5c) comprising residues 1–164. The SD has the expected pseudo 2-fold symmetry characteristic of this domain, consistent with a gene duplication event during the evolutionary history of this protein56. The synthase active site is found in one half of the SD, situated in a cleft formed by three subdomains (Supplementary Figs. 5b). In the other half of the SD, occupying a symmetry-related nucleotide-binding site, is an auxiliary ADP molecule (Supplementary Fig. 5b). The map shows clear density for the ADP and three associated Mg2+ ions (Fig. 3a and Supplementary Fig. 3). As there is no evidence linking this nucleotide to an enzymatic or regulatory function, the ADP is believed to play only a structural role or be a vestigial remnant of the putative duplication event that led to the formation of the SD62. The clear density distinguishing this molecule as an ADP in the binary complex, the enclosed nature of the binding site, and the fact that ADP was not present in any solutions used during purification or structural work, suggests that this molecule is incorporated during protein folding and cannot be competitively displaced by similar nucleotides, such as AMP-PNP (Fig. 3a and Supplementary Fig. 5b). The identity of the bound nucleotide was confirmed as ADP in HsFGAMS by analysis of denatured apoprotein by HPLC (Supplementary Fig. 5b).

In the binary complex of HsFGAMS, the ATP-binding site is occupied by AMP-PNP (Fig. 3b and Supplementary Fig. 3). The nucleotide binding pocket, which sits at the interface of two subdomains (Supplementary Fig. 5b), is composed of a hydrophobic pocket surrounding the adenine base (Fig. 3b, F235, F251, and F269) abutted by a loop (267–274) that forms hydrogen bonds with ribose hydroxyls and phosphate oxygen atoms. A comparison to the AMP-PCP bound PurL structure from T. maritima (TmPurL, PDB code 2HS4) suggests that additional interactions with, and charge stabilization of, the phosphates may be contributed by a metal cation that is not observable in our maps (Supplementary Fig. 6a). While the overall architecture of both the auxiliary ADP binding site and ATP binding site in the HsFGAMS SD is relatively well conserved with those of S. typhimurium and T. maritima PurL, there are several subtle but clear differences (Supplementary Fig. 6a, b). In the ATP binding site, interactions mediated by residues on the loop spanning 267-274 are distinct and dissimilar to those observed in the prokaryotic homologs (Supplementary Fig. 6a). In addition, a partially resolved, unstructured catalytic loop (C-loop, 461–479) is observed near the synthase site in the binary complex (Fig. 2a). For the prokaryote structure, this region was observed to become ordered only in the presence of FGAR at the synthase site55. Computational studies performed with the full length StPurL proposed that this loop plays a role in signal transmission and catalytic coupling between the distant active sites57.

The GD of HsFGAMS (Fig. 2b and Supplementary Fig. 5a) contains the conserved catalytic triad characteristic of the class I amidotransferase superfamily63. The catalytic cysteine (C1158) resides on a structural motif known as a “nucleophilic elbow”63. The GD interacts with the NTD primarily through hydrophobic contacts (W62, N103, I104, V107, and T111 of the NTD and V1122, G1128, A1131, and F1135 of the GD) while a series of hydrogen bonds span the GD-SD interface (N95, P99, and N102 of the NTD and D488, E490, G545, and D546 of the SD). Close examination of the HsFGAMS binary complex reveals a loop region (designated loop L1) from the NTD blocking the glutaminase active site (Supplementary Fig. 6c). The location of the L1 loop in the absence of substrate appears to be driven by an interaction between H14 on L1 and Y1237 of the GD. The presence of this 11-residue loop, and an additional, longer, solvent exposed loop (designated loop L4), are major differences setting HsFGAMS apart from the prokaryotic homologs, which do not possess these features (Supplementary Fig. 6d). However, despite low sequence identity and the presence of the loop inserts, the NTD of HsFGAMS has an overall fold that is structurally homologous to the prokaryotic PurLs of known structure (Supplementary Fig. 6e). The position of loop L1 in HsFGAMS, and its absence in prokaryotes, is suggestive of a putative role in regulating access to the glutaminase site that is distinct to the human enzyme.

Cryo-EM structure of the HsFGAMS ternary complex with trapped iminophosphate intermediate
Production of the FGAM product by FGAMS requires two separate reactions at distinct active sites (Fig. 1b). Ammonia liberated in the GD is added to FGAR in an ATP-dependent reaction. The mechanism of the synthase reaction is proposed to proceed via an activated iminophosphate intermediate55. To evaluate the architecture and allosteric regulation of the HsFGAMS SD, we prepared samples and collected Cryo-EM data for the protein with FGAR and ATP, but lacking the co-substrate glutamine. Details about sample preparation and processing are provided in the supplementary information and methods. After processing we obtained a 3.22 Å map (Fig. 2c and Supplementary Figs. 2b, 7 and 8), that was further post-processed using DeepEMhancer with individual half maps as input64, revealing additional features (Supplementary Fig. 9). The resulting map showed clearly resolved coulombic potential for a trapped iminophosphate intermediate together with an ADP molecule (Figs. 2c, d, and 3c), and a well ordered C-loop (Fig. 3d and Supplementary Fig. 8). Other alternative ligand binding modes, including ATP and FGAR or FGAR, ADP and phosphate, were not consistent with the observed density (Supplementary Figs. 10a, b). A comparison of the HsFGAMS ternary complex synthase site to that of TmPurL in complex with FGAR and AMP-PCP (PDB code 2HS4) reveals a similar binding position and orientation of the ligands and C-loop (Supplementary Fig. 10c). The ordered C-loop adopts a closed conformation in this ternary complex, shielding the synthase site and the reactive iminophosphate intermediate (Fig. 3e). This intermediate and the ADP molecule were observed to be present perpendicular to each other in the synthase site (Fig. 3e). The density also revealed the likely presence of an alternate conformer of ADP in the active pocket (Fig. 3c, e). Q485 on the C-loop makes key H-bonding interactions with the H311 and the formylglycinamide of the trapped intermediate (Fig. 3e). This interaction forms a putative anchor point that could drive the C-loop into a closed conformation. Prior computational studies using StPurL also implicated these two residues in a putative gating mechanism that controls transient channel formation65.

The SD of FGAMS belongs to the PurM superfamily of proteins, which includes selenophosphate synthase (SelD), iron-nickel hydrogenase maturation protein (HypE), and thiamine phosphate kinase (ThiL), among others66. These proteins are related by their signature DX4GAXPATP binding motif and the putative formation of an iminophosphate intermediate during catalysis. The trapped iminophosphate in the structure of the HsFGAMS ternary complex provides structural confirmation of this reaction intermediate. For FGAMS specifically, two alternate enzyme mechanisms have been proposed (Supplementary Fig. 11)67. These involve either a tetrahedral intermediate (Supplementary Fig. 11; path a) or the iminophosphate intermediate (Supplementary Fig. 11; path b) as observed in the HsFGAMS ternary complex structure. Our data, collected in the absence of an ammonia source, are consistent with phosphorylation of the δ-carbonyl of FGAR to form an iminophosphate intermediate.

Cryo-EM structure of the HsFGAMS quaternary complex with Glutamine, AMP-PNP, and FGAR
To further evaluate substrate binding, mechanism, and cooperative regulation of the distinct active sites, we incubated the protein with L-glutamine, AMP-PNP, and α/β-FGAR prior to grid preparation and data collection. From a total of 184,653 particles, we obtained a map at 3.19 Å using C1 symmetry (Fig. 2e and Supplementary Figs. 2c, 12 and 13). Further post-processing was performed using DeepEMHancer64, leading to improvements in the map, particularly in a hydrophobic linker, 3-helix bundle near the synthase site and the partially disordered C-loop (Figs. 2f, 4a and Supplementary Fig. 14). In the glutaminase active site the map was similarly well resolved to enable modeling of the substrate, which was trapped as a glutamyl thioester intermediate (Fig. 4b and Supplementary Fig. 13). In the synthase active site, there was clearly discernible coulomb potential to build the AMP-PNP and FGAR ligands (Fig. 4c, and Supplementary Fig. 13), as well as the majority of the C-loop structure (Fig. 4a–c). Similar to what was observed for the ternary complex, the AMP-PNP ligand could be built into two alternate conformations (Supplementary Fig. 13). The FGAR molecule sits perpendicular to the nucleotide, and is held in place primarily through H-bonding interactions from both the synthase binding site residues and the C-loop (Fig. 4c). When compared to the structure of the T. maritima PurL (PDB code 2HS4), the C-loop of HsFGAMS is in a more open conformation (Fig. 4d), potentially due to substrate binding at the distant GD. This configuration of the C-loop repositions Q485 away from H311 (Fig. 4c), disrupting the H-bond interaction between these two residues. This is consistent with a putative role for Q485 in regulation of intraprotein signal transmission, as proposed in previous studies65. The glutamyl thioester intermediate observed in the glutaminase active site is engaged in the primarily hydrophobic binding pocket that is highly conserved across species (Fig. 4d). Clear density was also observed at H-bonding distance from the OE2 atom of the thioester, modeled here as a water molecule (Fig. 4b, W). This marks the putative location of the initial site of the ammonia molecule immediately after liberation from glutamine.

Formation of the iminophosphate intermediate signals the GD via structural rearrangements in the NTD
The function of the NTD in FGAMS has long been unresolved. Comparison of the structures of the different states of HsFGAMS reveals distinct differences, particular between the binary (AMP-PNP bound) and ternary (iminophosphate and ADP) complexes (Fig. 5). Upon binding of the synthase substrates and formation of the iminophosphate intermediate, the NTD shifts substantially, rotating approximately 10° with respect to the GD (Fig. 5a). Accompanying this domain motion is a similar shift in a 3-helix bundle of the SD that abuts the C-loop (Fig. 5a). Conversely, other than a slight disordering of the C-loop, the overall structure of HsFGAMS in the quaternary complex was very similar to that of the binary complex (Fig. 5b).

Further analysis of these structural differences reveals several functionally relevant implications. In the ternary complex, one edge of the 3-helix bundle moves into proximity with the C-loop, forming new H-bonding interactions, specifically between G462, A465, and S468 of the C-loop and Q224 and E228 of the 3-helix bundle (Supplementary Fig. 15). The ordering and repositioning of the C-loop in the ternary complex result in a well-structured pocket surrounding the iminophosphate (Fig. 3e), presumably to protect the reactive intermediate. In addition, residues of the C-loop, which spans the region between the NTD and the 3-helix bundle, form a cation-π stacking interaction with conserved residues of the NTD (Supplementary Fig. 16). Based on the structures and supported by previous analysis of mutants57, this network plays a crucial role in intraprotein signaling between the NTD and C-loop.

This newly created NTD:C-loop interface is accompanied by conformational changes in the putative oxyanion loop, a recurring and short-lived feature common to nearly all glutaminases that stabilizes the oxyanion transition state (Fig. 6a and Supplementary Figs. 17a–d)68. Upon formation of the ternary complex, the interactions between the oxyanion loop and nearby Ω-loop, which harbors the signaling residue N107457,65, are disrupted. This results in an approximately 90° flip of the oxyanion loop and creation of the putative oxyanion hole (Fig. 6b–d). Based on the expected position of the oxyanion, movement of a section of the oxyanion loop would be necessary to accommodate the tetrahedral transition state. Our structures indicate that stabilizing interactions would be contributed through hydrogen bonding interactions with the amide nitrogen atoms of N1159 and one of the loop residues. Formation of the oxyanion hole in this manner is a proposed mechanism for substrate-dependent allosteric regulation in other glutamine amidotransferases and related enzymes, such as imidazole glycerol phosphate synthase69,70. The HsFGAMS structures suggest a critical role for the G1115-N1074 interaction in stabilizing the closed conformation of the oxyanion loop (Fig. 6b). This is consistent with previous mutational studies performed with the analogous residues in StPurL65. Upon glutamyl thioester intermediate formation, as observed in the quaternary structure of HsFGAMS, the oxyanion loop reverts back to the resting state, orienting residues G1115, G1116, and F1117 to help position and stabilize the intermediate (Fig. 6e, f).

In addition to the allosteric control through transient oxyanion stabilization, we also observed that entry to the glutaminase active site is regulated by two gates. Gate 1 is composed of H14 on the L1 loop of the NTD and Y1237 of the GD, while Y1202 and F1117 of the GD comprise gate 2 (Fig. 6a). The changes in the NTD of the ternary complex result in disorder in loop L1 (Fig. 6c), providing the needed flexibility for gating motions of H14. Similarly, these changes also impact the positioning of Y1237, yielding an open state of one of the gates (Fig. 6c, gate 1). At the same time, the loop flipping that controls transient oxyanion hole formation moves F1117 and Y1202. The latter, in particular, swings approximately 90° away from Y1202, opening this second gate (Fig. 6c, gate 2). Both of these gates return to the closed state in the quaternary complex of HsFGAMS (Fig. 6e). Note that the mechanism for gating through gate 1 is distinct to mammals, as the loop L1 region is not conserved in lower organisms (Supplementary Fig. 6c, d).

Catalytic coupling through transient ammonia channel formation
The chemistry at the synthase site of FGAMS requires delivery of the ammonia molecule produced at the distinct and distant glutaminase site. Delivery of the ammonia has been proposed to travel between the two active sites through a putative ammonia channel in the enzyme (Fig. 1b). To date, however, this channel has not been experimentally observed. In the HsFGAMS quaternary complex structure, we elucidate the architecture of this hydrophobic transient channel that passes near the auxiliary ADP binding site (Fig. 7a and Supplementary Fig. 18) using CAVER 3.0.2 PyMOL plugin71. The channel location and composition are consistent with what was predicted by MD simulations for the StPurL system (Supplementary Fig. 18)57.

Comparison of the quaternary structure to the binary and ternary complex structures reveal several key structural changes driving transient channel formation. Structure analysis showed that after glutamyl thioester intermediate formation in GD, the oxyanion loop flipped back to regain its original conformation and helped to stabilize the carbonyl group of the glutamyl moiety via hydrogen bonding interaction through its G1116 residue (Fig. 6e, f) and by closing gate 1 and gate 2 to protect the thioester intermediate in the GD (Fig. 6e). This oxyanion loop resting conformation further reverted the signal to the NTD by rotating it back by ~10o to go back in its previous conformation (Fig. 5b) via disrupting the GD and NTD interface (Fig. 6a and Supplementary Figs. 17a–d). This NTD/GD interface disruption forced the C-loop to move back in its unstructured conformation, similar to the HsPFAS-AMPPNP binary state. From moving fully ordered to fully disordered state, there is a stage when the C-loop is partially disordered. At this stage, two other gates; gate 3 (R1300-D685) at the top of the channel (Fig. 7b) and gate 4 (Q485-H311) at the bottom of the channel (Fig. 7c), are opened, which were observed to be closed in other two HsPFAS binary and ternary complexes solved here and in the previously solved StPurL and smPurL crystal structures also55,56. In brief, a salt bridge between R1300 and D685 reorganizes to form the opening of the channel at the glutaminase active site (Fig. 7b). At the synthase end of the channel, H311 is observed to block the exit in both the binary and ternary complexes, but subtle changes to Q485 disrupt the H311-Q485 interaction, opening the gate at the synthase end of the channel (Fig. 7c). The opening and closing events of these conserved gates were also highlighted in previous MD simulation studies57,65. Moreover, the mutational studies performed on gate 3 in StPurL showed a high ammonia channeling efficiency, highlighting the role of this gate in controlling ammonia release into the channel, while the mutants of gate 4 showed its dual role in gating and signal transduction to the glutaminase site65. In addition to these dual gates, additional motions were observed in the secondary structure elements on which the conserved channel lining residues reside. These residues include N394, P399, V400, S390, E398, D333, A348, T327, Y350, P423, T325, R330, T310 and M425. Additional bottleneck residues M425, Y350, and E398 obstruct the channel in the binary and ternary complexes, but change positions subtly in the quaternary complex structure to provide a clear path for the ammonia to traverse (Fig. 7a, d, and e). Mutational and crystallographic studies were previously performed on the conserved channel lining residue (V400 in HsFGAMS and V333 in StPurL) that showed a remarkable effect on the channel efficiency72. In this case, a valine to isoleucine mutation constricted the channel by ~2 Å, leading to a decrease in the synthase activity by ~20%72. Collectively, the structural changes precipitated by substrate binding and formation of the quaternary complex lead to the transient formation of the ammonia channel and provide additional gating mechanisms to regulate FGAMS activity. This type of enzyme control is not without precedent. Similar substrate-driven domain motions lead to the formation of an ammonia channel in the bacterial glucosamine-6-phosphate synthase73,74.

To confirm the role of the gating residues and transient channel formation, we generated a series of HsFGAMS mutants and characterized their relative kinetics. For all of the mutants, we determined the basal glutaminase activity, the ATP and FGAR dependent glutamate production, and overall FGAM synthesis rate (Supplementary Fig. 19a–c, respectively). While the basal rate of glutaminase activity of HsFGAMS is very low, removal of a short section of loop L1 containing gate 1 residue H14 (Δ12–15) or mutation of either of the gate 2 residues (Y1202F, F1117A), leads to a substantial increase in background glutaminase activity (Supplementary Fig. 19a). Leakage of ammonia through gate 2 is at a much higher rate, consistent with an overall motive force favoring ammonia flow from the GD to the synthase active site. As expected, the presence of ATP and FGAR does not have a large impact on the glutaminase rate for these mutants (Supplementary Fig. 19b). These mutants, however, do substantially decrease the overall rate of FGAM synthesis (Supplementary Fig. 19c). This disruption in the gating and coupling efficiency of the two domains through these subtle changes (Supplementary Fig. 19d) illustrates the importance of the tight control over substrate access and transit in FGAMS (Supplementary Figs. 19e, f). As a comparison, mutation of one of the bottleneck residues (M425K), which sits in the interior of the channel, has little impact on the glutaminase or synthase activity (Supplementary Figs. 19a–c).

Methods
General methods
Unless otherwise stated, all chemicals and reagents were from Fisher bioreagents, Merck-Millipore, Sigma-Aldrich, Fluka, and Acros Organics. All restriction enzymes were purchased from New England Biolabs or Thermo Scientific. The C-term DDK(Flag)-tagged human FGAMS (HsFGAMS) (accession #NM_012393.3) plasmid in pCMV6-entry vector was purchased from OriGene. HEK-293 cells (CRL-3216) used for protein expression were purchased from ATCC®.

Expression and FLAG affinity purification of human FGAMS
The C-terminally flag-tagged HsFGAMS protein was expressed in HEK-293 cells and purified using Flag-affinity purification as reported previously42. In brief, five 182 cm2 polystyrene tissue culture flasks having 0.22 μm vented filter cap (Genesee Scientific) were each seeded with 1 × 106 HEK-293 cells in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS) and 1% penicillin streptomycin (PS). Cells were grown at 37 °C in 5% CO2. The next day, when the cell confluence was ~80%, the media was replaced, and each flask was transfected with a complex of HsFGAMS plasmid and PEI transfection reagent (in a ratio of 1:3 in serum free media). The media was again replaced 24 h’ post-transfection and incubated at 37 °C in 5% CO2. Cells were harvested after 72 h and stored at −80 °C until purification. The protein was purified using 1X PBS (phosphate buffered saline), pH 7.4, running buffer. Cells were re-suspended in running buffer having 1X protease inhibitor cocktail (Roche) and sonicated twice for 25 s each with a 1-min pause and 40% amplitude. The cell lysate was centrifuged at 19840 rcf for 30 min and the supernatant was incubated with pre-equilibrated Flag affinity resin at 4 °C. After 2 h, the protein bound resin was collected by centrifugation at 770 rcf for 15 min and the supernatant was transferred into another vial. The Flag resin was washed 4 times with 1X PBS to remove loosely bound impurities. The protein was eluted by incubating the resin with 1 mL elution buffer (100 μg/mL Flag peptide in running buffer) at 4 °C for 30 min. The purity of the eluted protein was determined using 10% SDS-PAGE. Fractions deemed to be greater than 95% purity were pooled, concentrated using a 50k MWCO centrifugal concentrator (Millipore), and the concentration was measured using a nanodrop spectrophotometer at 280 nm.

Size exclusion chromatography (SEC)
SEC was performed at 25 °C using an Agilent 1100 series HPLC instrument with an analytical Zenix SEC-300 column (4.6 × 300 mm, Sepax). The column was pre-equilibrated and eluted in filtered and degassed SEC buffer (1X PBS; pH 7.4). HsFGAMS protein (0.6 mg/mL) in 20 μL of buffer was injected and eluted with a flow rate of 0.350 mL/min. A standard curve, generated using Thyroglobulin (669 kDa), Ferritin (443 kDa), Aldolase (158 kDa), Conalbumin (75 kDa), and RNase A (13.7 kDa), was generated and used to determine the approximate values for molecular weight.

Preparation of HsFGAMS complexes
The binary complex of HsFGAMS with AMPPNP was generated by incubating purified protein (0.7 mg/mL final concentration) with 10 mM AMPPNP. The ternary complex was generated by incubating purified HsFGAMS (0.7 mg/mL final concentration) with 10 mM ATP and 2 mM α/β-FGAR (1:1 anomeric mixture). The quaternary complex was prepared by incubating 0.5 mg/mL HsFGAMS with 10 mM L-Glutamine, 10 mM AMPPNP, and 2 mM α/β-FGAR. The reactions were setup in 1X PBS pH 7.4 buffer at 4 °C for 4 h followed by a short spin at 770 rcf, followed by further incubation on ice for 30 min.

Grid preparation and Cryo-EM data collection
4–5 μL of each complex was applied to a glow discharged Quantifoil R 1.2/1.3, UT, 300 mesh gold grid (Electron Microscopy Sciences) at a concentration of 0.5 to 0.7 mg/mL on a Vitrobot Mark IV system (Thermo Fisher Scientific) set at 4 °C, 100% humidity, blot time of 5 s and blot force of 0. The grids were screened for ice quality, as well as particle distribution and density, before setting up data collection. Images were collected on a 300 kV Titan Krios G2 microscope equipped with K3 Summit detector (Gatan) in super-resolution mode with binning 2 and CDS, along with a Gatan BioContinuum GIF energy filter (slit width of 20 eV) at The Hormel Institute with an exposure of 1.7–2 s. Image stacks comprising 40 frames were recorded at an estimated total dose of 50 e-/Å2. Automated data collection was carried out using the EPU 3.2.0.4776REL software (Thermo Fisher Scientific) at a nominal magnification of 130000X corresponding to a calibrated pixel size of 0.652 Å with a defocus range from -0.75 to -2.5 μm.

Data processing, class reconstruction, and model building of HsFGAMS complexes
Dose-fractionated movies were gain normalized, drift corrected, summed, and dose weighted using Patch motion correction module in CryoSPARC75. The contrast transfer function (CTF) was estimated for each summed image using the Patch CTF module. For HsFGAMS/AMPPNP binary complex, particles were picked and extracted from the dose-weighted images with box size of 320 px using CryoSPARC blob picker and particle extraction. The particles were curated using two rounds of CryoSPARC 2D classification (number of classes, N=50), resulting in 3,595,274 particles. The templates selected using Select 2D module were further used for particle picking using template picker module and extracted using the same box size resulting in 7,325,050 particles. High quality particles were selected using 4 cycles of 2D classification and selection. Initial models were generated using CryoSPARC ab initio reconstruction on full set of good 1,010,335 particles using C1 symmetry (N=3). The largest class of curated particles were used for homogenous refinement. This was further refined using non-uniform refinement that resulted in a map of 3.35 Å76. This map was used as a template for 3D classification (N=4) out of which particles from classes 1–3 were pooled and refined using non-uniform refinement with C1 symmetry, resulting in a 3.31 Å map. Further local map sharpening and denoising were performed using deepEMhancer tool in CryoSPARC. The alpha-fold predicted structure of HsFGAMS was used as a primary model and docked in the final map using ChimeraX77. For the HsFGAMS/Iminophosphate/ADP ternary complex, particles were picked and extracted from the dose-weighted images with box size of 320 px using the HsFGAMS/AMPPNP binary complex 2D templates. The particles were curated using 4 rounds of CryoSPARC 2D classification (number of classes, N=50). Initial models were generated using CryoSPARC ab initio reconstruction on a full set of 1,137,400 particles using C1 symmetry (N=5). The largest class of curated particles was used for non-uniform refinement that resulted in a map of 3.22 Å. This map was used as a template for 3D classification (N=4), out of which, the largest class was refined using non-uniform refinement with C1 symmetry, resulting in a 3.22 Å map. The HsFGAMS/AMPPNP binary complex structure solved above was used as a primary model and docked in the final map using ChimeraX. For the HsFGAMS/L-Glutamine/AMPPNP/FGAR quaternary complex, particles were picked and extracted from the dose-weighted images with box size of 320 px using the HsFGAMS/AMPPNP binary complex 2D templates and particle extraction. The particles were curated using 2 rounds of CryoSPARC 2D classification (number of classes, N=50). Initial models were generated using CryoSPARC ab initio reconstruction on the full set of 1,366,039 particles using C1 symmetry (N=4). The largest curated class was used for homogeneous refinement and followed by non-uniform refinement that resulted in a map of 3.21 Å. This map was used as a template for 3D classification (N=10), out of which three classes were pooled and refined using homogeneous followed by non-uniform refinement with C1 symmetry, resulting in a 3.19 Å map. In this case also, the HsFGAMS/AMPPNP binary complex structure solved above was used as a primary model and docked in the final map using ChimeraX. The manual model building for all three complexes was done in WinCoot78, and the models were refined in Phenix79. The refined models were validated with MolProbity80 within Phenix. Figures were prepared using UCSF Chimera, ChimeraX, and Pymol 2.4 (The PyMOL Molecular Graphics System, Version 1.2r3pre, Schrödinger, LLC).

Mutagenesis
All mutants were designed either by Gibson’s method81 or by traditional cloning. Native plasmids served as a template, and gene fragments were amplified using Phusion or Q5 high fidelity DNA polymerase. For the Δ12_15 and M425K mutants, the amplified fragments were assembled using 2X HiFi DNA assembly master mix (New England Biolab) for 1 h at 50 °C. For the F1117A and Δ12_15/Y1202F mutants, the amplified gene was digested with Mfe-I and Mlu-I restriction enzymes and ligated into the pCMV6-ENTRY vector using T4 DNA ligase. DH5α cells were directly transformed using the assembled and ligated genes to obtain the mutant plasmid. The inserted mutations were confirmed by Sanger sequencing.

Assay for glutamate production
The glutamate production of HsFGAMS was coupled with a standard glutamate dehydrogenase assay, where glutamate produced from HsFGAMS is converted to α-ketoglutarate in the presence of NAD+. Here, we used 3-acetylpyridiene adenine dinucleotide (APAD) in place of NAD+ because APAD has a more significant oxidation potential than NAD+, is more effectively reduced, and is more stable than NAD+. A 50 μL reaction was setup containing 20 mM Tris pH 7.5, 20 mM MgCl2, 80 mM KCl, 10 mM ATP, 1 mM α/β-FGAR, 1 mM APAD, 2 U L-Glutamate dehydrogenase, 1 mM L-Glutamine, and 1 μM test enzyme according to the previously reported protocol59. The glutamate production was determined using a coupled glutamate dehydrogenase assay82 by monitoring the reduction of APAD at 363 nm in Spectramax M2 plate reader for 30 min. To determine the kinetic parameters, the concentration of L-Glutamine was varied from 30 μM to 1 mM. The concentration of glutamate produced was determined by Beer Lambert’s equation (A=εCl where A is absorbance, ε is molar extinction coefficient of product, C is concentration of the product and l is path length in mm) using a molar extinction coefficient of 9100 M−1 cm−1.

Assay for FGAM synthesis
FGAM production was determined using a previously established protocol59. In brief, the conversion of FGAR to FGAM by HsFGAMS was coupled with TrifGART enzyme that further converts FGAM to aminoimidazole ribonucleotide (AIR). A reaction mixture of 100 μL having 20 mM HEPES buffer pH 7.5, 20 mM MgCl2, 80 mM KCl, 10 mM ATP, 1 μM α/β-FGAR, 5 mM L-Glutamine, 0.5 μM test enzyme, and 1 μM TrifGART enzyme was incubated at 37 °C for 30 min. The reaction was quenched by adding 15 μL of 20% trichloroacetic acid (pH 1.4). Precipitates were removed by centrifugation, and AIR formation was determined by coupling using a standard Bratton–Marshall’s assay83. The intensity of pink color, measured at 500 nm on a Spectramax M2 plate reader, was used to quantify product formation.

Transient ammonia channel identification
The transient ammonia channel has been identified using CAVER 3.0.2 PyMOL plugin using default parameters, i.e., probe radius of 0.9 Å, shell depth 4 Å, shell radius of 3 Å, and clustering threshold of 3.5. The channel connecting the two distant active sites in GD and FD was identified only in HsFGAMS quaternary complex cryo-EM structure reported here.

HPLC analysis of nucleotide binding in apo FGAMS
To determine the identity of the bound nucleotide in the apo FGAMS protein an aliquot of HsFGAMS protein purified as described above (in the absence of any nucleotides) was boiled for 20 min and centrifuged to remove precipitates. This sample was evaluated with ion exchange HPLC (Agilent 1100 system) with a 4.6 × 250 mm BioSAX NP5 column (Agilent) running a gradient from 25 to 300 mM NaCl in 20 mM Tris, pH 8, over 20 min at 1 mL/min. The elution time of the sample was compared to authentic standards of ATP, ADP, and AMP-PNP.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript by Sharma and colleagues reports the structures of three complexes of human PFAS that elucidate different steps in the catalytic cycle. Importantly, these structures reveal the ammonia tunnel between the glutaminase and FGAMS domain and elucidate the gating mechanism. The structures also appear to contain a metastable intermediate. These are very important findings. The experiments are carefully performed and for the most part clearly described. A few issues could be further clarified.\nMajor points\n1. The authors claim to capture the iminophosphate intermediate for the first time. However, they really only observe electron density that they have modeled as the iminophosphate intermediate. Given the resolution is only ~3Å, can they really rule out other potential complexes? Fits to the electron density (e.g., Fig. S8) should be included in the main manuscript, with the intermediate and ADP displayed as they appear in the active site, not separately as in Fig. S8, so that the reader can assess the goodness of the fit. Alternative complexes should be included in the SI to show that other reasonable possibilities do not fit as well (FGAR•ADP•Pi, FGAR•ATP, also alpha FGAR). In the absence of other evidence confirming the iminophosphate intermediate, given the resolution, it might be good to moderate their claim.\n2. ADP is a common contaminant of AMPPNP and ATP- have the authors confirmed that the apoenzyme contains the second ADP molecule?\n3. Tunnel mutants. The key issue in these experiments is whether the glutaminase reaction has become uncoupled from the FGAMS reaction. This is very difficult to assess from the data as presented. One wants to directly compare the rate of ammonia production to the rate of FGAM production, which is most easily accomplished in a Table. What is the “relative basal glutamate production” in Fig. S18A? there is no wild-type basal NH3 production…\nMinor issues\n1. The introduction refers to Fig. 1A, but never discusses lgPurLs and smPurLs.\n2. The manuscript text should report kcat for glutaminase reaction in addition to Km. Note that the kcat reported int Fig 1 legend has units of uM/min- these are units for Vmax. The value of kcat should be in time -1 (e.g., s-1): Vmax = kcat•[E]\n3. Fig. 1B would be easier to read if the structures had thicker lines/bold fonts.\n4. Fig. 1C the y-axis should be converted to concentration of product (or the assay should be described since it is not clear what absorbs at 363 nm). The x-axis should be converted to minutes.\n5. Fig. 1D legend- “kinetic analysis of the GD” can be taken to mean that the GD domain is assayed independently. Better to “kinetic analysis of the glutaminase reaction”.\n6. The IC50 = 148 uM for FGAR requires more explanation- as written, it implies that the authors have developed a new assay for the FGAMS reaction. However, the new assay is for a promiscuous dephosphorylation reaction.\n7. “Cryo-EM of the HSFGAMS ternary” sectin, 3rd line: “Glutamine” should be “Ammonia”. Second to last line “formylglycinamide” is misspelled.\n8. The two conformations of ADP should be discussed when the ternary complex is discussed.\n9. More information about how the composition of the ammonia tunnel resembles the MS simulations would be welcome.\n10. Gates 3 and 4 in Fig. 7 are not discussed in the text.\n11. No red C-loop fragment is apparent in Fig. 2A although red loops fragments are visible in Fig. 2B.\n12. Fig. 6 talks about Gates 1 and 2 controlling entry into the glutaminase site, but the ammonia should be leaving the glutaminase site and going into the tunnel. It might make more sense to number the gates in the order that the ammonia would encounter them.", "reviewer_2": "In this manuscript, Sharma et al. have structurally characterized catalytic cycle of human phosphoribosylglycinamidine synthase (FGAMS). This is in line with previous results of Nandini Sharma on the topic of FGAMS enzymatic mechanism, e.g. Sharma et al. Acs Catal 2022 and Science advances 2020.\nThe manuscript describes detailed conformational steps in the enzymatic reaction performed by FGAMS on the example of the human enzyme. This brings insights into allosteric regulation and the related confirmation changes of this class of enzymes. In particular, identification of the reaction intermediate, formation of the ammonia channel or the catalytic coupling of individual enzyme domains. The authors should however bring these results better in perspective to the current knowledge (largely produced by the first author) rather than claiming future possible use of the results in therapeutic development, which is not the scope of the manuscript.\nThe structural data is of good quality to answer the relevant biological questions. Structural figures are very informative. The final figure of the proposed model could be improved.\nOverall, the study by Sharma et al. is undoubtedly a significant contribution to our understanding of amidotransferase enzymes and will be interesting to a broad audience of readers. However, some adjustments of the manuscript can be done to improve the interpretation of the very complex mechanism the authors describe.\nComments:\n1. The authors discuss reaction pathway of the FGAR to FGAM conversion and structural confirmation of reaction intermediate, described also in figure S10. The discussion should clearly mark which pathway is supported by the structural data. In figure S10 is should be marked which intermediate is captured in the cryo-EM structure. From my understanding of results in ref. 57, the 018 labeling experiments only confirms the transfer of amide oxygen of FGAR to the released phosphate group, not the reaction intermediate itself.\n2. The authors claim ‘structural elucidation of an ammonia channel connecting the active sites of the glutaminase and synthase domains’. It might be beneficial to compare their finding with the previously observed ammonia channels, e.g. in work of Stephane Mouilleron (J. Biol. Chem. 2006).\n3. In the final model Figure 8 there are many individual residues marked, however, their function is not mentioned in neither the discussion or figure legends. It might be beneficial to mention their function while describing the figure. Overall the figure could better depict the individual confirmation changes and their respective functions. Omit marking of individual domains in individual states but rather describe what they do/how they change in individual steps. E.g. glutamine entry gating is completely omitted in the Figure 8.\nMinor comments:\n4. The authors should include line and page numbering in the submitted manuscript.\n5. The first part of the introduction is dedicated to relevance of purine metabolism to disease, which is not the main topic of the study. Part of the introduction should be dedicated to the current knowledge of ‘allosteric regulation and catalytic coupling’ of amidotransferase-like enzymes.\nThe ‘Increased biosynthesis of (d)NTPs is a universal, and highly druggable, feature of cancer.’ and ‘The transfer of ammonia between the two active sites is facilitated by a putative ammonia channel.’ sentences should be referenced.\n6. The manuscript should avoid ‘first time’ claims as they do not add any useful meaning.\n7. The first paragraph of results should be largely transferred to methods section.\n8. The ‘as a basis for future drug discovery efforts’ should be omitted, as there is no reference to practical drug discovery in the manuscript. Or the authors should make some discussion on the possible drug discovery in the discussion section.\n9. In the second paragraph of the results, authors might add some details to the description of the observed structure.\nParticularly to comment about the ligand combinations.\n10. In the third paragraph of the results the sample preparation and cryo-EM details should be transferred into methods section.\n11. Figure S4B is referenced but does not exist.\n12. Is glutamine correct in this sentence: ‘Glutamine liberated in the GD is added to FGAR in an ATP-dependent reaction. The mechanism of the synthase reaction is proposed to proceed via an activated iminophosphate intermediate (51).’?\n13. Is this a surprise or rather expected outcome: ‘Surprisingly, the resulting maps showed clearly resolved coulombic potential for a trapped iminophosphate intermediate, and ADP molecule, and a well ordered C-loop (Figs. 2C, 2D, 3C and S8).’?\n14. Sentence ‘This is consistent with previous computational studies that implicated these two residues in a gating mechanism for transient channel formation (54).’ probably needs to be introduced to be in context with the text before.\n15. The authors used the non-uniform refinement implemented in the cryoSPARC software, which generated over sharpened regions of the obtained cryo-EM maps. To mitigate his phenomenon they utilized the DeepEMhancer post-processing. Please verify individual half-maps were used as inputs to DeepEMhancer post-processing. The authors should also mention DeepEMhancer utilization in any cryo-EM map figures, e.g. Figure S3 or S8 o S12. The authors should also make sure they deposit the raw cryo-EM maps along with the DeepEMhancer post-processed maps into the EMDB database.\n16. Did the authors use any data aberration correction algorithms, reference-based motion correction and frame weighing to improve the final cryo-EM maps?\n17. Could authors comment on the very low Q-scores of some residues of the deposited PDB model? In some places the model could be improved to corelate better with the obtained cryo-EM map. Please consider adding all side chain residues.\n18. Some water molecules are modelled. Are the modeled water molecules confirmed by occurrence in multiple cryo-EM structures? The water molecules are not visible in the DeepEMhanced maps.\n19. In the cryo-EM methods the authors mention utilization of Gatan K3 camera. In other parts they mention Falcon 4i detector. Please check this discrepancy.\nMinor comments to Figures:\n1. In Figure 1 the sticking of PAGE analysis should be marked.\n2. In Figure 2 the overall cryo-EM maps might be omitted, as they do not add to much additional information. They look in different orientation in respect to the presented models.\n3. In Figure 4C it might be helpful to directly compare details of C-loop in ternary complex and quaternary complex. Not only refencing to TmPurL.\n4. Figure 7 legend. In line 5, there is probably a word missing after ‘The’ in the beginning of sentence.\n5. In Figure 8, the titles of described states should be referred to observed structures.\n6. In Figure 8, the position of release ammonia should correlate with the predicted position in the structure study.", "reviewer_3": "The manuscript describes structures of the human de novo purine biosynthetic enzyme phosphoribosylglycinamidine synthase (hFGAMS) from cryo-EM analysis.\nThe 3-domain monomeric enzyme (>1300 residues) catalyzes a multi-step reaction: ATP-dependent formation of an imino-phosphate intermediate of the FGAR substrate in a synthase active site, release of ammonia by glutamine hydrolysis in a glutaminase active site, diffusion of ammonia through a channel in the enzyme, and attack of ammonia on the imino-phosphate to release the FGAM product. The synthase and glutaminase active sites are >20 Å apart. A long-standing question has been the coordination of the catalytic steps, as no glutamine hydrolysis occurs in absence of the FGAR substrate.\nThe authors have considerable experience with FGAMS, and here they cryo-trapped 3 substrate complexes on EM grids by judicious choices of reaction conditions. The EM maps are of high quality (resolution 3.1 – 3.3 Å by 0.143 FSC). They include\nbinary complex of AMP-PNP in the synthase acive site,\nternary complex with the imino-phosphate intermediate & ADP in the synthase active site,\nquaternary complex with AMP-PNP & FGAR in the synthase active site and the acyl-enzyme intermediate (Cys-Glu thioester) in the glutaminase active site.\nMost of the Results section is devoted to a detailed analysis of small structural features of the 3 complexes. Nearly all the structure regions described have good density in the EM maps.\nFGAMS is well conserved from bacteria to humans, so some of the substrate poses can be validated in published higher-resolution crystal structures of prokaryotic homologs. This aspect of the interpretation seems very solid. The changes in structure with addition of substrates are very small. Visualizing the human enzyme in several catalytic states is a substantial advance in our understanding of catalytic coupling in hFGAMS, but – in my view – less of an advance than the authors claim.\nThe structural details were challenging to understand, even after superposition of the 3 models, because several elements of structure are not defined (loop 1, loop 4, C-loop, catalytic triad, oxyanion loop, Ω-loop, 3-helix bundle, gate 1, gate 2, gate 3, gate 4). This makes it very challenging to look at the structures and evaluate the authors’ model for catalytic coupling and ammonia travel."}, "author_response": "Point-by-point responses to reviewer comments.\n(Reviewer comments are shown as italicized text and our responses are given in blue text.)\nReviewer 1:\nThe manuscript by Sharma and colleagues reports the structures of three complexes of human\nPFAS that elucidate different steps in the catalytic cycle. Importantly, these structures reveal the\nammonia tunnel between the glutaminase and FGAMS domain and elucidate the gating\nmechanism. The structures also appear to contain a metastable intermediate. These are very\nimportant findings. The experiments are carefully performed and for the most part clearly\ndescribed. A few issues could be further clarified.\nWe thank the reviewers for their positive comments and constructive feedback. Our responses\nare provided for all issues raised.\n1. The authors claim to capture the iminophosphate intermediate for the first time. However,\nthey really only observe electron density that they have modeled as the iminophosphate\nintermediate. Given the resolution is only ~3Å, can they really rule out other potential\ncomplexes? Fits to the electron density (e.g., Fig. S8) should be included in the main\nmanuscript, with the intermediate and ADP displayed as they appear in the active site, not\nseparately as in Fig. S8, so that the reader can assess the goodness of the fit. Alternative\ncomplexes should be included in the SI to show that other reasonable possibilities do not fit as\nwell (FGAR•ADP•Pi, FGAR•ATP, also alpha FGAR). In the absence of other evidence\nconfirming the iminophosphate intermediate, given the resolution, it might be good to moderate\ntheir claim.\nThe observed coulomb potential at the binding site in that structure shows clear density that is\nconsistent with the proposed iminophosphate intermediate. We have moved this figure panel to\nFigure 3 in the revised manuscript (shown below). In order to rule out other possibilities, we\nhave fit the other possible ligand combinations and included these results in a new figure in the\nSI (shown below). As illustrated, neither of the putative alternative substrate combinations\nmatches the observed density. Binding of the alpha anomer of FGAR has been previously ruled\nout (Stubbe et al. Biochemistry 1986, 25(8): 2256) and is similarly not consistent with the\nobserved density.\nFig. 3C from the revised manuscript New SI Figure. Shows the fit to the\nshowing the fit to the iminophosphate alternative substrates, ATP and FGAR\nintermediate and ADP. (A), and FGAR, ADP and phosphate (B).\n2. ADP is a common contaminant of AMPPNP and ATP- have the authors confirmed that the\napoenzyme contains the second ADP molecule?\nAs illustrated in the SI (Figs S3, S8 and S13), there is clear density consistent with an ADP\nmolecule coordinated to three magnesium ions. This is also consistent with what was observed\nfor homologous FGAMS proteins reported to bind ADP at this site. However, as the reviewer\nsuggested, we have conducted additional experiments to confirm the identity of this molecule.\nNew SI figure S5b shows results from HPLC analysis of denatured apo protein, purified in the\nabsence of any additional nucleotides. There is a clear peak for a molecule present in the apo\nprotein, and this elutes with a profile and UV spectrum matching that of the ADP standard.\n3. Tunnel mutants. The key issue in these experiments is whether the glutaminase reaction has\nbecome uncoupled from the FGAMS reaction. This is very difficult to assess from the data as\npresented. One wants to directly compare the rate of ammonia production to the rate of FGAM\nproduction, which is most easily accomplished in a Table. What is the “relative basal glutamate\nproduction” in Fig. S18A? there is no wild-type basal NH3 production…\nWe thank the reviewer for this suggestion. To better make the point, as the reviewer suggested we have\nadded a table to Fig. S18 that shows the ratio of ammonia production to FGAM production. We have also\nchanged the plots to all have the same scale, which additionally helps to clarify the point. Regarding the\nquestion about relative glutamate production in S18A, we have updated the description to now be “%\nBasal glutamate production”, which is defined as the rate of glutamate production in the absence of co-\nsubstrates as compared to the WT rate in the presence of all substrates (set to 100%). As suggested by\nthe reviewer, this new representation of the data clearly shows a substantial reduction (20- to 100-fold) in\ncoupling efficiency.\nMinor issues\n1. The introduction refers to Fig. 1A, but never discusses lgPurLs and smPurLs.\nWe have included additional text in the introduction discussing the distinction between large\nPurL and small PurL protein variants.\n2. The manuscript text should report kcat for glutaminase reaction in addition to Km. Note that\nthe kcat reported int Fig 1 legend has units of uM/min- these are units for Vmax. The value of\nkcat should be in time -1 (e.g., s-1): Vmax = kcat•[E]\nCorrected.\n3. Fig. 1B would be easier to read if the structures had thicker lines/bold fonts.\nWe have updated Fig. 1B as suggested.\n4. Fig. 1C the y-axis should be converted to concentration of product (or the assay should be\ndescribed since it is not clear what absorbs at 363 nm). The x-axis should be converted to\nminutes.\nWe have included additional description to the online methods under the section ‘Assay for\nglutamate production’ to improve clarity, as suggested. The glutamate production of HsFGAMS\nwas coupled with a standard glutamate dehydrogenase assay where glutamate produced from\nHsFGAMS is converted to α-ketoglutarate in the presence of NAD+. In the assay reported, we\nused 3-acetylpyridiene adenine dinucleotide (APAD), in place of NAD+. APAD has a more\nsignificant oxidation potential than NAD+ and is more effectively reduced than NAD+. Also, the\nstability of its reduced product (which is monitored at 363 nm) is significantly higher than the\nreduced product of NAD+ (NADH).\n5. Fig. 1D legend- “kinetic analysis of the GD” can be taken to mean that the GD domain is\nassayed independently. Better to “kinetic analysis of the glutaminase reaction”.\nCorrected.\n6. The IC50 = 148 uM for FGAR requires more explanation- as written, it implies that the\nauthors have developed a new assay for the FGAMS reaction. However, the new assay is for a\npromiscuous dephosphorylation reaction.\nWe have corrected the main text to clarify these details about the new assay, as suggested.\n7. “Cryo-EM of the HSFGAMS ternary” sectin, 3rd line: “Glutamine” should be “Ammonia”.\nSecond to last line “formylglycinamide” is misspelled.\nCorrected.\n8. The two conformations of ADP should be discussed when the ternary complex is discussed.\nWe have included discussion of the two conformations of ADP in the revised manuscript.\n9. More information about how the composition of the ammonia tunnel resembles the MS\nsimulations would be welcome.\nWe have included two additional paragraphs of text in the results section (under the subheading\n“Catalytic coupling through transient ammonia channel formation”) further describing the\nammonia channel and prior MD simulations.\n10. Gates 3 and 4 in Fig. 7 are not discussed in the text.\nWe thank the reviewer for pointing this out. While we had discussed gates 3 and 4 in the text,\nthey were not clearly defined as such. We have clearly defined gates 3 and 4 in the revised\nmanuscript prior to the discussion of these gates.\n11. No red C-loop fragment is apparent in Fig. 2A although red loops fragments are visible in\nFig. 2B.\nThe red C-loop is now visible, and clearly labeled, in the revised figure 2A.\n12. Fig. 6 talks about Gates 1 and 2 controlling entry into the glutaminase site, but the ammonia\nshould be leaving the glutaminase site and going into the tunnel. It might make more sense to\nnumber the gates in the order that the ammonia would encounter them.\nWe chose to number the gates following the order that the ammonia source enters the glutaminase site\nvia ammonia release from the glutaminase site followed by its entry to the synthase site for reaction. The\ngate 1 (loop L1; at the exterior of the protein), gate 2 (Y1202-F1117; next to the gate 1) allow the\nammonia source (glutamine) to enter to the synthase site. Then the ammonia produced leaves the\nglutaminase site via rearrangements in gate 3 (R1300-D685; at the exit of glutaminase site) at GD and\nenters to the synthase site via opening of the gate 4 (H311-Q485; at the entry of synthase site) at SD. We\nfeel that this convention is the most suitable in this case.\nReviewer 2:\nIn this manuscript, Sharma et al. have structurally characterized catalytic cycle of human\nphosphoribosylglycinamidine synthase (FGAMS). This is in line with previous results of Nandini\nSharma on the topic of FGAMS enzymatic mechanism, e.g. Sharma et al. Acs Catal 2022 and\nScience advances 2020. The manuscript describes detailed conformational steps in the\nenzymatic reaction performed by FGAMS on the example of the human enzyme. This brings\ninsights into allosteric regulation and the related confirmation changes of this class of enzymes.\nIn particular, identification of the reaction intermediate, formation of the ammonia channel or the\ncatalytic coupling of individual enzyme domains. The authors should however bring these\nresults better in perspective to the current knowledge (largely produced by the first author)\nrather than claiming future possible use of the results in therapeutic development, which is not\nthe scope of the manuscript. The structural data is of good quality to answer the relevant\nbiological questions. Structural figures are very informative. The final figure of the proposed\nmodel could be improved. Overall, the study by Sharma et al. is undoubtedly a significant\ncontribution to our understanding of amidotransferase enzymes and will be interesting to a\nbroad audience of readers. However, some adjustments of the manuscript can be done to\nimprove the interpretation of the very complex mechanism the authors describe.\nWe thank the reviewer for their thorough evaluation of this work and recognition of its significance. We\nhave attempted to address these suggestions, specifically regarding the scope of the work and model, as\ndescribed in the point-by-point response below.\nComments:\n1. The authors discuss reaction pathway of the FGAR to FGAM conversion and structural\nconfirmation of reaction intermediate, described also in figure S10. The discussion should\nclearly mark which pathway is supported by the structural data. In figure S10 is should be\nmarked which intermediate is captured in the cryo-EM structure.\nThe proposed intermediate captured in the cryoEM ternary complex is now highlighted in Fig. S10.\nFrom my understanding of results in ref. 57, the 018 labeling experiments only confirms the\ntransfer of amide oxygen of FGAR to the released phosphate group, not the reaction\nintermediate itself.\nWe thank the reviewer for catching this overstatement of the published data. We have removed this\nstatement.\n2. The authors claim ‘structural elucidation of an ammonia channel connecting the active sites of\nthe glutaminase and synthase domains’. It might be beneficial to compare their finding with the\npreviously observed ammonia channels, e.g. in work of Stephane Mouilleron (J. Biol. Chem.\n2006).\nAs suggested by the reviewer, we have included additional discussion of ammonia channel formation in\nbacterial glucosamine-6-phosphate synthase as described in Teplyakov et al (JMB 2001) and Mouilleron\net al (JBC 2006).\n3. In the final model Figure 8 there are many individual residues marked, however, their function\nis not mentioned in neither the discussion or figure legends. It might be beneficial to mention\ntheir function while describing the figure. Overall the figure could better depict the individual\nconfirmation changes and their respective functions. Omit marking of individual domains in\nindividual states but rather describe what they do/how they change in individual steps. E.g.\nglutamine entry gating is completely omitted in the Figure 8.\nTo better depict the observed changes and their functions we have updated the figure and clarified the\ndescription of individual steps in the text. Specifically, we have removed the domain names, labeled the\nindividual residues in the figure and labeled specific features, such as gates and loops, that are discussed\nmore fully in the text. In addition, we have substantially expanded our discussion of the mechanism,\nclearly referencing the structural features in Fig. 8, and describing the relevant functional changes that\naccompany the structural reorganizations.\nMinor comments:\n4. The authors should include line and page numbering in the submitted manuscript.\nCorrected.\n5. The first part of the introduction is dedicated to relevance of purine metabolism to disease,\nwhich is not the main topic of the study. Part of the introduction should be dedicated to the\ncurrent knowledge of ‘allosteric regulation and catalytic coupling’ of amidotransferase-like\nenzymes.\nAs suggested we have included an additional paragraph of text describing regulatory mechanisms of\npurine metabolic enzymes and, in particular, catalytic coupling of amidotransferases.\nThe ‘Increased biosynthesis of (d)NTPs is a universal, and highly druggable, feature of cancer.’\nand ‘The transfer of ammonia between the two active sites is facilitated by a putative ammonia\nchannel.’ sentences should be referenced.\nCorrected.\n6. The manuscript should avoid ‘first time’ claims as they do not add any useful meaning.\nCorrected.\n7. The first paragraph of results should be largely transferred to methods section.\nAs suggested, we have removed extraneous information about the purification from the first paragraph of\nthe results section.\n8. The ‘as a basis for future drug discovery efforts’ should be omitted, as there is no reference to\npractical drug discovery in the manuscript. Or the authors should make some discussion on the\npossible drug discovery in the discussion section.\nOur group is currently using these results as a basis for drug discovery (anti-metabolite chemotherapy for\nliver cancer). We have included additional text near the end of the discussion section to clarify the\nimportance of this protein target for drug discovery.\n9. In the second paragraph of the results, authors might add some details to the description of\nthe observed structure.\nParticularly to comment about the ligand combinations.\nIn the second paragraph of the results section, we introduce the three structures and describe\nthe combinations of ligands bound for each. The text reads: “Using Cryo-EM, we solved the\nstructures of 1) HsFGAMS with AMP-PNP bound (referred to as the binary complex); 2)\nHsFGAMS in complex with the iminophosphate reaction intermediate and adenosine\ndiphosphate (ADP) (referred to as the ternary complex); and 3) HsFGAMS with glutamyl\nthioester, AMP-PNP and FGAR bound (referred to as the quaternary complex) (Fig. 2).”\nDetailed descriptions of each structure are provided in the sections following.\n10. In the third paragraph of the results the sample preparation and cryo-EM details should be\ntransferred into methods section.\nAs suggested, we have removed some Cryo-EM details to the methods section.\n11. Figure S4B is referenced but does not exist.\nOur apologies for the mistake. This has been corrected to cite the appropriate figure, Fig. S5A.\n12. Is glutamine correct in this sentence: ‘Glutamine liberated in the GD is added to FGAR in an\nATP-dependent reaction. The mechanism of the synthase reaction is proposed to proceed via\nan activated iminophosphate intermediate (51).’?\nOur apologies for the mistake. This has been corrected to read “Ammonia liberated…is added to\nFGAR…”\n13. Is this a surprise or rather expected outcome: ‘Surprisingly, the resulting maps showed\nclearly resolved coulombic potential for a trapped iminophosphate intermediate, and ADP\nmolecule, and a well ordered C-loop (Figs. 2C, 2D, 3C and S8).’?\nWhile, as the reviewer indicates, this result was what we were seeking to achieve with the experiment, the\nflexibility and dynamics of protein structures always makes solving a structure with a trapped intermediate\na pleasant surprise. Having said that, given what was known about the protein, this result was not\nunexpected. We have removed the qualifier at the beginning of this sentence in the revised text. It now\nbegins “The resulting maps showed…”\n14. Sentence ‘This is consistent with previous computational studies that implicated these two\nresidues in a gating mechanism for transient channel formation (54).’ probably needs to be\nintroduced to be in context with the text before.\nWe have updated this text to put it better into context with the previous statements.\n15. The authors used the non-uniform refinement implemented in the cryoSPARC software,\nwhich generated over sharpened regions of the obtained cryo-EM maps. To mitigate his\nphenomenon they utilized the DeepEMhancer post-processing. Please verify individual half-\nmaps were used as inputs to DeepEMhancer post-processing.\nIndividual half-maps were used as inputs to DeepEMhancer. We have clarified this in the text.\nThe authors should also mention DeepEMhancer utilization in any cryo-EM map figures, e.g.\nFigure S3 or S8 o S12. The authors should also make sure they deposit the raw cryo-EM maps\nalong with the DeepEMhancer post-processed maps into the EMDB database.\nWe have revised the captions for these figures to describe that individual half-maps were used. The raw\nmaps and post-processed maps have been deposited to the EMDB database.\n16. Did the authors use any data aberration correction algorithms, reference-based motion\ncorrection and frame weighing to improve the final cryo-EM maps?\nNo. No further improvements were made beyond what is mentioned in the text.\n17. Could authors comment on the very low Q-scores of some residues of the deposited PDB\nmodel? In some places the model could be improved to corelate better with the obtained cryo-\nEM map. Please consider adding all side chain residues.\nWe built the model into the map using the iterative process described in the methods section until, in our\nestimation, no further residues or side chains could be definitively built into the maps. The fit to the maps\nis given in SI figures S3, 8 and 13. While additional side chains could be added, it is our convention to\nonly add side chain density when the placement of the side chain is unambiguous. As discussed, we\nhave deposited the raw data for the maps and the models to the EMDB for later analysis by anyone\ninterested.\n18. Some water molecules are modelled. Are the modeled water molecules confirmed by\noccurrence in multiple cryo-EM structures? The water molecules are not visible in the\nDeepEMhanced maps.\nYes, we have confirmed that the ordered water molecules placed in the maps are accurate and consistent\nacross multiple structures. The enhancement algorithm used by DeepEMhancer often suppresses the\nsignals from non-protein features.\n19. In the cryo-EM methods the authors mention utilization of Gatan K3 camera. In other parts\nthey mention Falcon 4i detector. Please check this discrepancy.\nThank you for catching this typo. We have corrected and further clarified this in the text (the Images were\ncollected on a 300kV Titan Krios G2 microscope equipped with K3 Summit detector).\nMinor comments to Figures:\n1. In Figure 1 the sticking of PAGE analysis should be marked.\nWe have updated this in the revised manuscript.\n2. In Figure 2 the overall cryo-EM maps might be omitted, as they do not add to much additional\ninformation. They look in different orientation in respect to the presented models.\nThe models are presented in the exact same orientation with respect to the maps. We appreciate the\nreviewer’s viewpoint, but respectively disagree that the maps should be omitted. It is our opinion that they\nare a vital accompaniment to the models by providing support for the quality of the data.\n3. In Figure 4C it might be helpful to directly compare details of C-loop in ternary complex and\nquaternary complex. Not only refencing to TmPurL.\nWe thank the reviewer for the suggestion and have included in the revised Fig. 4 the comparison with\nboth the ternary and quaternary complex.\n4. Figure 7 legend. In line 5, there is probably a word missing after ‘The’ in the beginning of\nsentence.\nCorrected.\n5. In Figure 8, the titles of described states should be referred to observed structures.\nCorrected.\n6. In Figure 8, the position of release ammonia should correlate with the predicted position in\nthe structure study.\nWe have modified the position of the ammonia release in the figure to more accurately reflect the position\nin the structure.\nReviewer 3:\nThe manuscript describes structures of the human de novo purine biosynthetic enzyme\nphosphoribosylglycinamidine synthase (hFGAMS) from cryo-EM analysis.\nThe 3-domain monomeric enzyme (>1300 residues) catalyzes a multi-step reaction: ATP-\ndependent formation of an imino-phosphate intermediate of the FGAR substrate in a synthase\nactive site, release of ammonia by glutamine hydrolysis in a glutaminase active site, diffusion of\nammonia through a channel in the enzyme, and attack of ammonia on the imino-phosphate to\nrelease the FGAM product. The synthase and glutaminase active sites are >20 Å apart. A long-\nstanding question has been the coordination of the catalytic steps, as no glutamine hydrolysis\noccurs in absence of the FGAR substrate.\nThe authors have considerable experience with FGAMS, and here they cryo-trapped 3\nsubstrate complexes on EM grids by judicious choices of reaction conditions. The EM maps are\nof high quality (resolution 3.1 – 3.3 Å by 0.143 FSC). They include\nbinary complex of AMP-PNP in the synthase acive site,\nternary complex with the imino-phosphate intermediate & ADP in the synthase active site,\nquaternary complex with AMP-PNP & FGAR in the synthase active site and the acyl-enzyme\nintermediate (Cys-Glu thioester) in the glutaminase active site.\nMost of the Results section is devoted to a detailed analysis of small structural features of the 3\ncomplexes. Nearly all the structure regions described have good density in the EM maps.\nFGAMS is well conserved from bacteria to humans, so some of the substrate poses can be\nvalidated in published higher-resolution crystal structures of prokaryotic homologs. This aspect\nof the interpretation seems very solid. The changes in structure with addition of substrates are\nvery small. Visualizing the human enzyme in several catalytic states is a substantial advance in\nour understanding of catalytic coupling in hFGAMS, but – in my view – less of an advance than\nthe authors claim.\nThe structural details were challenging to understand, even after superposition of the 3 models,\nbecause several elements of structure are not defined (loop 1, loop 4, C-loop, catalytic triad,\noxyanion loop, Ω-loop, 3-helix bundle, gate 1, gate 2, gate 3, gate 4). This makes it very\nchallenging to look at the structures and evaluate the authors’ model for catalytic coupling and\nammonia travel.\nTo address this point and help clarify the catalytic coupling mechanism, we have made substantial\nchanges to the model figure and included additional text in the figure caption and body of the manuscript.\nIn particular (as suggested by this and the other reviewers), we have included additional labels on the\nfigure, changed the titles of the states and clearly described each of the labeled structural elements in the\ncaption and body of the manuscript. We feel that this improved figure and additional details provide a\nmuch more thorough and clearer description of the mechanism and features that drive it.\nMoreover, the structural differences among the 3 complexes are very small – this is apparent in\nthe superposition. For example, the salt bridge between D685 and R1300 occurs in all 3\ncomplexes, belying the claim that it “reorganizes to form the opening of the channel at the\nglutaminase active site”. The very small structural changes may be amplified by use of a too-\nsmall probe sphere (radius not stated) in the CAVER calculation of an interior channel surface.\nWe have included a detailed description of the methods used to identify the transient ammonia channel,\nincluding the probe size (0.9 Å), in the revised online methods section. We have also updated the figure\nshowing the proposed gating residues to include bond distances. With respect to the scale of the\ndifference between the complexes, the maps were of sufficient quality to clearly place the sidechains of\nthe residues lining the ammonia channel (including D685 and R1300) in all three complexes with a high\ndegree of certainty. Comparison of the quaternary structure to that of the ternary structure shows a clear\n1 Å shift in the gating residue (D685) coordinated with a shift in R1300. This opens the distance in the\nchannel from 3 Å to greater than 4 Å (as shown in figure 7B – reproduced below). While this is a relatively\nsmall change, considering the effective Van der Waals diameter of ammonia is estimated to be 3.7 Å, the\nscale of the change is certainly consistent with the role of these residues in controlling access to and\nthrough this transient channel. Note also that, as mentioned in the text, the involvement of these residues\nwas predicted and then confirmed by MD simulations and mutagenesis studies, respectively, in the\nhomologous S. typhimurium enzyme StPurL (Science Advances, 2020, 6, eaay7919; ACS Catalysis\n2022, 12: 1930).\nFig. 7B: A superposition of the HsFGAMS structures shows changes in the gating residues at the top of the channel.\nThe binary complex is colored white, ternary complex is colored gray and for the quaternary complex, the GD\nresidues are colored magenta, SD residues are cyan. The ammonia channel is shown by dark blue spheres.\nThe ammonia channel is certainly near where the authors have placed it, but close examination\nof the structures does not support a unique position for the channel. Mutagenesis of residues\nlining the proposed ammonia channel (especially those in secondary structures where\nsubstitution is less likely to alter other structure elements) would be a good test of the proposed\nchannel location.\nAs mentioned in the previous comment and described in the text, previous computational and\nmutagenesis experiments using StPurL are consistent with the description of the ammonia channel\nprovided herein. The channel lining residues are highly conserved from human to S. typhimurium (as\nillustrated in SI Fig. S18). Additional structural and mutagenesis experiments in StPurL on residues lining\nthe channel similarly demonstrated a profound impact on activity (ACS Chem Biol 2014, 10: 698). The\nfurther mutagenesis work described herein provides additional strong evidence for the location of the\nchannel and role of the residues lining this structure. While additional mutations across the enzyme could\nbe evaluated, considering the preponderance of structural and functional data, all consistent with the\nmodel proposed herein, we feel that further mutagenesis experiments will not substantially alter the\noutcome and, as such, are beyond the scope of this current work.\nThe mutagenesis data in Fig S18 are a nice addition to the paper. It would be interesting to\ndetermine whether the loss of catalytic coupling in some of the variants affects only glutamine-\ndependent FGAM production (can hFGAMS use ammonia directly?).\nHsFGAMS does not accept ammonia directly as a substrate for the synthase activity and, as such, it is\nnot possible to determine a rate for glutamine independent FGAM production. Note however, that\nmutations that effect the gating residues impact the coupling efficiency (how effectively the synthase site\nuses the ammonia produced at the glutaminase site), as described in the revised figure S19.\nTechnical comments\n“formation of the ternary complex…results in a 90° flip of the oxyanion loop and creation of the\noxyanion hole”, but the “flip” looks far less than 90°.\nThe change in angle was 67°, 75° and 120° for the three flipped residues, G1115, G1116 and F1117,\nrespectively. As such, we reported this as the approximate average value of 90° for the loop residues. We\nhave updated the text to reflect that this is an approximate value for the three residues.\nI disagree that this “flip” of the 1115-1116 peptide, whatever its magnitude, forms the oxyanion\nhole. The oxyanion hole should consist of 2 H-bond donors, presumably the backbone NHs of\nG1116 and N1159. In the ternary complex where an oxyanion hole is proposed to exist, the\nG1116 NH is not pointed in a direction that can partner with the backbone NH of N1159 to\nstabilize a transient O-. However, this pair of NHs is beautifully positioned to stabilize an\noxyanion in the binary and quaternary complexes.\nThe reviewer’s latter point is correct. It was not very clearly stated in the manuscript, but the oxyanion\nhole is formed by two of these residues, including N1159 and one of the residues in the loop 1115-1117.\nThe point that we wished to make was that there is a flexible loop region that is clearly involved in\nstabilizing the oxyanion that is formed during the reaction. Our structural data shows a transition from a\nclosed/narrow to open to closed state as the reaction proceeds. While it is not possible to definitively\nassign which residues stabilize the transition state, the peptide flip is consistent with a reordering of this\nregion to accommodate and stabilize the oxyanion. This type of oxyanion loop reorganization has been\npreviously observed for other enzymes including glutamine amidotransferases and glycerol phosphate\nsynthase (as described in the text). We have updated the text in the revised manuscript to more clearly\ndescribe this facet of the observed structural reorganization and putative implications for mechanism.\n“an enantiomeric mixture of the substrate, α/β-FGAR” – shouldn’t this be an anomeric mixture\nWould the density have been cleaner if a pure anomer (or stable analog thereof) were\navailable?\nThe reviewer is correct; we have changed this to “an anomeric mixture”. Since the protein will only bind\none anomer, it is unlikely that the purity of the anomeric mixture would impact the results.\n“Glutamine liberated in the GD is added to FGAR” – should be ammonia liberated in the GD\nCorrected.\nSeveral waters have only a whisper of density – why are they in the models?\nThe water molecules in the model are fitted based on the density observed in the refined sharpened\nmaps of the complexes. As described in our response to reviewer 2, these densities are not clearly\nobservable in the deepEMhanced maps because the algorithm is trained primarily on protein-only\ndatasets, making it unable to recognize or preserve the much weaker, diffuse signals characteristic of\ndynamic small molecules like water. The enhancement process prioritizes sharpening protein density to\naid in modeling, which can actively suppress or remove the signals from non-protein features. As such,\nwhile we have modeled waters only into density that was clearly visible in the sharpened maps, the\ndensity may not be clear in the enhanced maps. Both the sharpened and deepEMhanced maps have\nbeen uploaded in the database for each structure and will be accessible for further analysis by all\ninterested parties."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have addressed all my concerns.", "reviewer_2": "The authors have addressed all concerns raised. They have improved the discussion section and the description of the final model in Figure 8. In particular, the revised manuscript provides a clearer description of the catalytic cycle of human FGAMS and the associated structural reorganizations of the enzyme.\nThe cryo-EM methodology follows standard approaches for 3D structure determination.\nMinor comments:\nThe authors should refrain from making a “first time” claim in the discussion section (line 393).\nIn the superposition figures (e.g., Figures 5 and S17), it would be helpful to specify which residues were used for the structural alignment.", "reviewer_3": "The authors have responded appropriately to my comments."}, "author_response": "Response to reviewer comments (our response follows each reviewer comment):\nThe authors should refrain from making a “first time” claim in the discussion section (line 393).\nWe have removed all “first time” claims from the manuscript.\nIn the superposition figures (e.g., Figures 5 and S17), it would be helpful to specify which residues\nwere used for the structural alignment.\nWe have now specified the numbering of the aligned residues and rmsd values in the figure\ncaptions of these respective figures.\nThe manuscript describes structures of the human de novo purine biosynthetic enzyme\nphosphoribosylglycinamidine synthase (hFGAMS) from cryo-EM analysis.\nThe 3-domain monomeric enzyme (>1300 residues) catalyzes a multi-step reaction: ATP-\ndependent formation of an imino-phosphate intermediate of the FGAR substrate in a synthase\nactive site, release of ammonia by glutamine hydrolysis in a glutaminase active site, diffusion of\nammonia through a channel in the enzyme, and attack of ammonia on the imino-phosphate to\nrelease the FGAM product. The synthase and glutaminase active sites are >20 Å apart. A long-\nstanding question has been the coordination of the catalytic steps, as no glutamine hydrolysis\noccurs in absence of the FGAR substrate.\nThe authors have considerable experience with FGAMS, and here they cryo-trapped 3\nsubstrate complexes on EM grids by judicious choices of reaction conditions. The EM maps are\nof high quality (resolution 3.1 – 3.3 Å by 0.143 FSC). They include\nbinary complex of AMP-PNP in the synthase acive site,\nternary complex with the imino-phosphate intermediate & ADP in the synthase active site,\nquaternary complex with AMP-PNP & FGAR in the synthase active site and the acyl-enzyme\nintermediate (Cys-Glu thioester) in the glutaminase active site.\nMost of the Results section is devoted to a detailed analysis of small structural features of the 3\ncomplexes. Nearly all the structure regions described have good density in the EM maps.\nFGAMS is well conserved from bacteria to humans, so some of the substrate poses can be\nvalidated in published higher-resolution crystal structures of prokaryotic homologs. This aspect\nof the interpretation seems very solid. The changes in structure with addition of substrates are\nvery small. Visualizing the human enzyme in several catalytic states is a substantial advance in\nour understanding of catalytic coupling in hFGAMS, but – in my view – less of an advance than\nthe authors claim.\nThe structural details were challenging to understand, even after superposition of the 3 models,\nbecause several elements of structure are not defined (loop 1, loop 4, C-loop, catalytic triad,\noxyanion loop, Ω-loop, 3-helix bundle, gate 1, gate 2, gate 3, gate 4). This makes it very\nchallenging to look at the structures and evaluate the authors’ model for catalytic coupling and\nammonia travel. Moreover, the structural differences among the 3 complexes are very small –\nthis is apparent in the superposition. For example, the salt bridge between D685 and R1300\noccurs in all 3\ncomplexes, belying the claim that it “reorganizes to form the opening of the channel at the\nglutaminase active site”. The very small structural changes may be amplified by use of a too-\nsmall probe sphere (radius not stated) in the CAVER calculation of an interior channel surface. The ammonia channel is certainly near where the authors have placed it, but close examination of the structures does not support a unique position for the channel. Mutagenesis of residues lining the proposed ammonia channel (especially those in secondary structures where substitution is less likely to alter other structure elements) would be a good test of the proposed channel location.\nThe mutagenesis data in Fig S18 are a nice addition to the paper. It would be interesting to\ndetermine whether the loss of catalytic coupling in some of the variants affects only glutamine-\ndependent FGAM production (can hFGAMS use ammonia directly?).\nTechnical comments\n“formation of the ternary complex…results in a 90° flip of the oxyanion loop and creation of the\noxyanion hole”, but the “flip” looks far less than 90°.\nI disagree that this “flip” of the 1115-1116 peptide, whatever its magnitude, forms the oxyanion\nhole. The oxyanion hole should consist of 2 H-bond donors, presumably the backbone NHs of\nG1116 and N1159. In the ternary complex where an oxyanion hole is proposed to exist, the\nG1116 NH is not pointed in a direction that can partner with the backbone NH of N1159 to\nstabilize a transient O-. However, this pair of NHs is beautifully positioned to stabilize an\noxyanion in the binary and quaternary complexes.\n“an enantiomeric mixture of the substrate, α/β-FGAR” – shouldn’t this be an anomeric mixture\nWould the density have been cleaner if a pure anomer (or stable analog thereof) were\navailable?\n“Glutamine liberated in the GD is added to FGAR” – should be ammonia liberated in the GD\nSeveral waters have only a whisper of density – why are they in the models?"}]

Decision letter