Structural basis of the promiscuity of the unusual Fe(II) and 2-oxoglutarate dependent human aspartate/asparagine-β-hydroxylase

Domain: chemistry
Published: 2026-02-25
Editorial outcome: minor_revision

Abstract
Protein-hydroxylation catalysed by Fe(II) and 2-oxoglutarate (2OG) dependent oxygenases is an important regulatory mechanism in human biology. Such oxygenases typically coordinate their Fe(II) cofactor via a conserved triad of an aspartate- or glutamate- and two histidine-residues. By contrast, aspartate/asparagine β-hydroxylase (AspH), which catalyses asparagine/aspartate-residue oxidation in epidermal growth factor-like domains (EGFDs), has only two histidine-residues (H679, H725), with a water occupying the site normally occupied by an aspartate- or glutamate-residue. We describe mechanistic studies with catalytically active AspH crystals. Turnover studies with single crystals under cryogenic conditions give (3R)-hydroxylated EGFDs with the product alcohol coordinating Fe(II) trans to H725. Time-resolved serial crystallography of microcrystals using an acoustic droplet ejection system, coupled to X-ray emission analyses, demonstrate turnover within 1.5 s, giving a product complex in which Fe(II) is regenerated. Solution and crystallographic studies with the O2 surrogate nitric oxide imply O2 binds to Fe(II) trans to H725. The additional Fe-chelating water is maintained throughout AspH catalysis and is not directly involved in substrate hydroxylation, because O2 is the sole oxygen source in alcohol products, as shown by 18O labelling studies. The results reveal how AspH accommodates both aspartate- and asparagine-substrates and will assist in efforts targeting AspH for cancer treatment.

Introduction
2-Oxoglutarate (2OG)- and Fe(II)-dependent oxygenases catalyse a range of oxidative reactions, including hydroxylations, halogenations, demethylations, and oxidative ring closures.1 In humans, they have roles in transcriptional regulation, lipid metabolism, epigenetics, collagen biosynthesis, and hypoxia sensing.2, 3, 4, 5 Inhibitors of the 2OG-dependent hypoxia-inducible factor-α (HIF-α) prolyl hydroxylases (PHDs) and γ-butyrobetaine hydroxylase (BBOX) are clinically used.6, 7, 8 2OG oxygenases other than the PHDs and BBOX are current medicinal chemistry targets for cancer treatment, including JmjC histone demethylases9 and aspartate/asparagine β-hydroxylase (AspH).10, 11, 12, 13, 14

AspH is a 2OG- and Fe(II)-dependent oxygenase that catalyses C3 hydroxylation of aspartate- and asparagine-residues in multiple epidermal growth factor-like domains (EGFDs) in the endoplasmic reticulum (ER).15, 16 EGFDs are compact ~30–50 residue domains that typically contain three intradomain disulfide bonds.17 Recent studies have revealed that AspH only accepts EGFDs as substrates when they have a non-canonical disulfide bond connectivity (C1-C2, C3-C4, C5-C6), rather than the canonical C1-C3, C2-C4, C5-C6 disulfide connectivity that is observed in most EGFD structures, suggesting a role for AspH in regulating thiol/disulfide biochemistry in the ER.18 The results of crystallographic studies on AspH complexes using Mn(II) as a catalytically inactive Fe(II) surrogate and a derivative of the EGFD1 of human coagulation factor X (hFX) with the non-canonical C3-C4 disulfide (hFX-Asp) show that substrate binding not only involves the AspH 2OG oxygenase domain, but also an adjacent tetratricopeptide repeat (TPR) domain located N-terminal to the oxygenase domain (Fig. 1).18, 19 The ability of AspH to catalyse hydroxylation of multiple EGFD substrates resembles that of several other human 2OG oxygenases, including factor inhibiting HIF-α (FIH) and Jumonji-C domain-containing protein 6 (JMJD6).20, 21

Many 2OG oxygenases appear to employ a conserved catalytic cycle, wherein coordination of 2OG to Fe(II) is normally followed by substrate binding at the active site.22 O₂ binding to Fe(II) then generates a Fe(III):2OG:superoxide intermediate that undergoes oxidative decarboxylation to yield CO₂ and, at least in one case, a succinyl peroxide intermediate.23, 24, 25 The latter fragments to produce an Fe(IV)=O (ferryl) species,26, 27, 28, 29, 30, 31 which abstracts a hydrogen from the substrate;32 radical rebound reaction with Fe(III)–OH forms the hydroxylated product and regenerates Fe(II) (Fig. 1, Supplementary Fig. 1).32 Variations on the consensus mechanism can occur, including with respect to the coordination sites to which O2 and the 2OG C1 carboxylate bind.33, 34

Unlike most human 2OG oxygenases which typically coordinate Fe(II) via a conserved HXD/E…H triad of residues,1, 35 the Fe(II) in AspH is coordinated by two histidine-residues (H679 and H725), with a water molecule (W1) occupying the normal aspartate-/glutamate-carboxylate site.18 This Fe(II)-binding motif resembles that of the 2OG-dependent halogenases (Supplementary Fig. 2),36, 37 suggesting that AspH may employ an atypical mechanism, although it apparently does not catalyse halogenation of EGFDs.18 Computational and biochemical studies indicate the importance of second coordination sphere residues in stabilizing the water (W1) coordinating Fe(II).38, 39

The unusual Fe(II) coordination chemistry of AspH raises the question as to what extent it follows the consensus mechanism for 2OG dependent hydroxylases.1, 39 In particular, it is unknown to which Fe(II) coordination site O2 binds to and in which direction the Fe(IV)=O intermediate is formed, which is usually responsible for hydrogen abstraction and subsequent hydroxylation.

A recently reported catalytically inactive AspH:Mn:2OG:hFX-Asp complex crystal structure revealed two conformations of Asp103hFX (i.e., the hydroxylation site of hFX-Asp): one in which it interacts with the Q627 sidechain, leaving the coordination site trans to H725 vacant for potentially O₂ binding, and one in which the Asp103hFX carboxylate coordinates directly to Mn, potentially blocking O₂ access (Fig.1).40 Since the substitution of Fe(II) for inactive Mn(II) inhibits catalysis,41 investigating the native AspH:Fe(II) complex is important to determine whether this non-productive Asp103hFX conformation is relevant to Fe-dependent catalysis.

Time-resolved serial femtosecond crystallography (SFX) studies using an X-ray free electron laser (XFEL) enable mechanistic investigations on enzymes at atomic resolution in real time at ambient temperature,42, 43, 44 including on redox-labile intermediates which are difficult to analyse using traditional synchrotron radiation due to reactions induced by X-ray photoelectric effects.45, 46 By integrating acoustic droplet ejection (ADE)-type sample delivery with femtosecond crystallography,47, 48 experimental throughput and temporal precision have been significantly enhanced, enabling precise initiation of reaction and synchronized capture of intermediate states. The combination of SFX with complementary techniques, such as X-ray emission spectroscopy (XES), enables time-resolved analysis of oxidation states, adding a distinct dimension to the mechanistic understanding of catalytic cycles. These approaches have provided insight into the mechanisms and structures of metalloenzymes,49, 50, 51 including 2OG oxygenases.25, 52

Here we report investigations on the AspH mechanism, using both conventional cryogenic X-ray crystallography of catalytically active single crystals and room temperature substrate and product complex structures, obtained using SFX at an XFEL source. The combined crystallographic results provide information on conformational changes involving a hydrogen bonding network at the AspH active site during catalysis. Solution-phase studies, including EPR spectroscopy and nitric oxide (NO) binding experiments, together with isotopic labelling and X-ray emission spectroscopy (XES) analyses reveal how AspH accommodates both aspartate- and asparagine-substrates for stereoselective hydroxylation and provide insight into the activation of O2 and substrate hydroxylation within the atypical AspH active site.

Results
Crystallographic analysis of catalytically active AspH complexes

Reported AspH complex crystal structures have employed catalytically inert Mn(II) or Ni(II) as Fe(II) substitutes (Fig. 1a),18, 19 a difference that may perturb the fold and/or active site metal coordination geometry. We thus worked to obtain crystal structures of catalytically active AspH complexed with Fe(II). We obtained >500 mg of homogenous recombinant untagged AspH315-758 in a single preparation following our recently reported optimized protocol.53 This material was used to optimise AspH crystallization under anaerobic conditions with Fe(II), 2OG, and hFX-Asp, yielding needle-shaped crystals (precipitation buffer: 0.1 M bis-tris propane, pH 7.5, 0.2 M NaBr, 18% v/v PEG 3350). Data collection at Diamond Light Source (DLS) under cryogenic conditions and subsequent elucidation of the AspH structure revealed the presence of Fe, 2OG, and hFX-Asp (AspH:Fe:2OG:hFX-Asp; PDB: 9FVZ [http://doi.org/10.2210/pdb9FVZ/pdb], 1.95 Å resolution). The overall fold in the AspH:Fe:2OG:hFX-Asp complex structure is similar to that observed in the corresponding reported aerobic AspH:Mn:2OG:hFX-Asp complex structure (PDB: 8RE9 [http://doi.org/10.2210/pdb8RE9/pdb] , RMSD = 0.156 Å, Supplementary Fig. 3).40

The Fe-ion in the AspH:Fe:2OG:hFX-Asp complex structure is coordinated via the H679 and H725 sidechains (2.3 and 2.1 Å, respectively), the 2OG C1 carboxylate and C2 carbonyl (2.1 and 2.3 Å, respectively), and two H2O molecules: W1 located trans to the 2OG C2 carbonyl (2.1 Å), i.e. the site typically occupied by an aspartate/glutamate carboxylate in 2OG oxygenases,1, 35 and W2 located trans to H725 (2.2 Å) (Fig. 2a). 2OG binding is stabilized via hydrogen bonding and electrostatic interactions with the sidechains of S668, H690 and R735, as observed in the corresponding AspH:Mn:2OG:hFX-Asp complex structure (Supplementary Fig. 3c).19, 40 W1 binding is stabilised by hydrogen bonding with the sidechain carboxylate of the second coordination sphere D721 and the backbone carbonyl of the substrate Asp103hFX (2.9 and 2.6 Å, respectively; Fig 2a).

In the AspH:Fe:2OG:hFX-Asp structure, modelling of W2 (the water located trans to H725) indicated partial occupancy (~50%) (Fig. 2a, Supplementary Fig. 3d+e). In the refined model, W2 interacts with a nearby water (W3; 3.1 Å), which interacts with the Q627 sidechain amide (2.6 Å) and the Asp103hFX sidechain carboxylate (2.9 Å). These interactions form a Fe-W2-W3-Q627-Asp103hFX hydrogen bond network (Supplementary Fig. 3d+f). Importantly, the Fe-coordination site trans to H725 is only partially occupied in the Fe-bound structure, whereas in the reported AspH:Mn:ligand:hFX-Asp complexes,18, 19, 40 it is filled, either by a water molecule or the substrate Asp103hFX carboxylate. In the AspH:Fe:2OG:hFX-Asp complex structure, no evidence for coordination of the Asp103hFX sidechain to the metal was observed (Fig. 2a). These differences suggest that the Fe(II) centre possesses greater coordination flexibility than the analogous Mn(II) complex. The observed six-coordinate, slightly distorted octahedral geometry with a weakly bound W2 in the Fe-bound complex supports the proposal that the W2 occupying coordination site becomes available for O2 binding, consistent with reported computational predictions.39

The corresponding AspH:Fe:2OG:hFX-Asn structure (PDB 9FVX [http://doi.org/10.2210/pdb9FVX/pdb], 1.90 Å resolution, Fig. 2b, Supplementary Fig. 3a+b) was obtained using identical conditions with the hFX-Asn substrate to inform on the effect of substituting Asp103hFX for Asn103hFX. The overall fold (RMSD 0.152 Å) and active site geometry resemble those of the hFX-Asp complex (Supplementary Fig. 3f). Notably, however, only weak positive electron density (<20%; Supplementary Fig. 3e) was observed for an Fe-bound water molecule located trans to H725 (corresponding to W2), and this water was therefore not refined (Fig. 2b, Supplementary Fig. 3).

The combined results imply that W2 is relatively weakly bound to Fe, suggesting that binding of the EGFD substrate may influence W2 binding and dissociation, thereby potentially modulating O2 binding and catalytic efficiency. Such a mechanism may help explain the wide variation in the observed extent of EGFD hydroxylation (0-100%) even when the preferred substrate consensus sequence for AspH is present.54, 55, 56

The absence of W2 in the hFX-Asn complex affects the conformation of the Q627 sidechain, which adopts a conformation enabling a stronger interaction with the Asn103hFX sidechain amide (2.8 Å) than with the Asp103hFX carboxylate (3.0 Å) in the corresponding hFX-Asp structure. W3 is absent in the hFX-Asn complex, thus the Fe–W2–W3–Q627–Asp103hFX hydrogen-bond network is not formed (Supplementary Fig. 3e+f). These observations suggest that the intact hydrogen bond network is not essential for hydroxylation, but may contribute to tuning substrate selectivity and reactivity. Notably, no evidence was obtained for direct Fe-coordination by the Asn103hFX carboxamide, contrasting with the AspH:Mn:2OG:hFX-Asp structures where the Asp103hFX carboxylate was modelled to coordinate the Mn with 50% occupancy (Supplementary Fig. 3i).19, 40 Collectively, the differences between the Fe and Mn structures highlight the importance of employing catalytically-relevant Fe(II) when investigating the coordination chemistry of 2OG oxygenases and, by implication, related metalloenzymes.

To investigate the effects of the hFX-Asp to hFX -Asn substitution on AspH catalysis, we performed solution-based kinetic studies with hFX-Asn using solid phase extraction coupled to mass spectrometry (SPE-MS) assays, comparing the results to those with hFX-Asp.57 The results reveal that the AspH 𝑘 /𝐾 value for Fe(II) with hFX-Asn was similar, within experimental error, to that reported with hFX-Asp.57 For 2OG, the 𝑘 /𝐾 for hFX-Asn was ~4-fold lower than that reported for hFX-Asp.57 Interestingly, direct competition studies of hFX-Asn and hFX-Asp reveal that AspH catalysed hFX-Asp oxidation occurs more efficiently than with hFX-Asn (Supplementary Fig. 4), suggesting that the Fe-W2-W3-Q627-Asp103hFX hydrogen bond network supports catalytic efficiency. These results imply that turnover of Asn-containing EGFDs is less efficient than Asp-containing EGFDs, under equivalent conditions. Although the biological significance of this difference remains to be established, it raises the possibility that substrate-dependent variations in 2OG affinity may contribute to the variable extents of EGFD hydroxylation observed in cells. Future work is required to determine whether such kinetic differences play a regulatory role in AspH activity in vivo.

Investigations on the O binding site of AspH

Anaerobic AspH:Fe:2OG:hFX-Asp single crystals were exposed to nitric oxide (NO), a catalytically inactive O2 mimic, to investigate whether O2 binds to the Fe(II) coordination site partially occupied by W2 in the AspH:Fe:2OG:hFX-Asp structure. Incubation of crystals in NO saturated precipitation solution resulted in an AspH:Fe:2OG:NO:hFX-Asp structure (PDB: 9HO3 [http://doi.org/10.2210/pdb9HO3/pdb], 2.39 Å resolution; Fig. 3a, Supplementary Fig. 5a). NO displaces W2 to coordinate to Fe(II) trans to H725 (2.0 Å) with the oxygen of NO 3.0 Å from the α-carbon of Asp103hFX; note that electron density for W3 was not observed. To test for the presence of NO at the active site, modelling of H2O or a product alcohol [(3R)-OH-Asp103hFX] was attempted: neither model reflected the observed electron density (Supplementary Fig. 5c).

EPR was then employed to investigate NO binding to AspH in solution. Samples were prepared by exposing a solution of the anaerobic AspH:Fe:2OG:hFX-Asp complex to 0.1% NO in N2. The results show that NO binds to form a single characteristic high-spin (hs) [FeNO]7 S = 3/2 state, simulated with effective g-values of g1 = 4.15, g2 = 3.915, and g3 = 2.0017 (Fig. 3b, Supplementary Fig. 6).58 Without hFX-Asp, evidence for a mixture of nitrosyl iron centre species was observed, in particular dinitrosyl iron complexes (DNIC, Supplementary Fig. 7), indicating that NO occupies more than one Fe(II) coordination site, possibly in a concentration-dependent manner. UV-Vis analyses of the same samples further supports binding of NO to the AspH:Fe:2OG:hFX-Asp complex, as shown by increased absorption in the 300-400 nm region, an observation more pronounced without hFX-Asp (Supplementary Fig. 8). This absorption band is characteristic of metal–nitrosyl charge-transfer transitions,59 reflecting Fe–NO bond formation and changes in the Fe(II) coordination environment.

Further evidence for an AspH:Fe:2OG:NO:hFX-Asp complex was obtained by the observation that irradiation with unfiltered white light resulted in apparent NO dissociation from the complex, as implied by the disappearance of the hs [FeNO]7 signal at temperatures of 10 K, 15 K, 25 K, and 30 K, which was recovered upon removal of the light source and/or increasing of the temperature to 77 K (Supplementary Fig. 9). The photolysis-recovery EPR experiments imply an activation energy (Ea) for re-binding of NO of ~0.12 kJ/mol (Supplementary Fig. 10-11). This low activation barrier value is substantially lower than those reported for analogous studies with other metalloenzymes; for instance, with cytochrome c oxidase, Ea values for NO re-binding measured at 50 K are an order of magnitude higher, with reported ranges between 1–5 kJ/mol.60, 61 These comparisons imply that the [FeNO]7 species in AspH exists in a labile coordination state, supporting its role as a mimic of a transient Fe–O₂ complex. The combined results support the likely site of O2 binding as the Fe(II) coordination site trans to H725, i.e., the site partially occupied by W2 in the AspH:Fe:2OG:hFX-Asp complex structure (Fig. 2a), a configuration that mirrors the proposed O2 binding modes observed in the 2OG oxygenases TauD and PHD2.62, 63, 64

We assessed the binding of NO to the microcrystalline sample of AspH:Fe:2OG:hFX-Asp and AspH:Fe:2OG:hFX-Asn by EPR spectroscopy, using FeSO₄ as an Fe(II) source. In the microcrystalline state, NO did not appear to coordinate to the iron after exposure to 0.1% NO in N2. Following exposure to an NO saturated precipitation solution, however, the microcrystalline samples resulted in formation of the hs [FeNO]7 S = 3/2 spin state (Fig. 3b, trace iia+b, Supplementary Table 1). Overall, in frozen solution, the rhombicity of the [FeNO]⁷ complex was approximately twice that of the microcrystalline state, indicating a more symmetric ligand field in the latter which may influence the energetic landscape of the catalytic intermediate. Interestingly, an anomalous NO species at g ~ 2 was also observed in microcrystals, something reminiscent of observations with other non-heme iron systems.58

Binding of isothiocyanate to AspH

We investigated the ability of (pseudo)halides to bind with AspH using X-ray crystallography to inform on possible reasons why AspH does not catalyse EGFD halogenation despite sharing a similar Fe(II) coordination geometry with 2OG-dependent halogenases.18, 65 Crystals grown in the presence of potassium thiocyanate (KSCN) revealed electron density for AspH, Fe, 2OG, and hFX-Asn (PDB: 9FVU [http://doi.org/10.2210/pdb9FVU/pdb] , resolution 1.70 Å). Notably, additional positive electron density was observed near the Fe W2/O₂ binding site trans to H725: The electron density was modelled with isothiocyanate (70% occupancy) coordinating to Fe via its nitrogen (M-N=C=S) with an Fe–N–C angle of ~137° (AspH:Fe:2OG:hFX-Asn:NCS complex; Fig. 3c+d, Supplementary Fig. 12a+b), consistent with reported isothiocyanate complexes.66 We also obtained the corresponding AspH:Mn:2OG:hFX-Asn:NCS complex, which showed a similar AspH fold as in the corresponding Fe structure (RMSD = 0.135 Å), but with full isothiocyanate occupancy (PDB: 9FVV [http://doi.org/10.2210/pdb9FVV/pdb] , Supplementary Fig. 12c+d).

Superimposition of the hFX-Asn complex structures, either with or without isothiocyanate, reveals binding of isothiocyanate induces local conformational changes. In the presence of isothiocyanate, the Q627 sidechain amide reorients away from the carboxamide of Asn103hFX, increasing the separation beyond hydrogen bonding distance (>5 Å). Q627L adopts a conformation that enables a potential hydrogen bond with the sidechain hydroxyl of S688 (2.9 Å), which forms a hydrogen bond with the 2OG C5 carboxylate (2.7 Å) (Supplementary Fig. 12e,f). Interestingly, while isothiocyanate coordination was observed in the AspH:hFX-Asn structures, it was not evident when substituting hFX-Asn for hFX-Asp. This observation may reflect a stronger interaction between the Q627 sidechain and the Asp103hFX sidechain carboxylate than that with the Asn103hFX sidechain amide, as supported by the observed substrate preference of AspH for hFX-Asp over hFX-Asn (Supplementary Fig. 4).

The observation that a pseudohalogen can bind to AspH-complexed Fe(II) is of interest, because, although isothiocyanate binds in proximity of the EGFD substrate, turnover studies with AspH in the presence of excess thiocyanate/other pseudohalogens did not reveal evidence for EGFD (pseudo)halogenation.18 Although the AspH Fe(II) coordination geometry resembles that of 2OG dependent halogenases,36, 37 isothiocyanate binding is observed at the W2/O2 binding site rather than the W1 Fe(II) coordination site (trans to the 2OG C2 carboxylate), where halides typically bind in 2OG halogenases to enable substrate halogenation. The location of isothiocyanate at W2 therefore represents a non-productive geometry, which likely impairs both O2 activation and subsequent potential substrate (pseudo)halogenation.

Crystallographic analysis of AspH product complexes

Exposure of AspH:Fe:2OG:hFX-Asp/hFX-Asn crystals to air for ≥20 min at room temperature gave product complex structures with (3R-OH)-Asp103hFX (PDB: 9FVY [http://doi.org/10.2210/pdb9FVY/pdb], 1.90 Å resolution) and (3R-OH)-Asn103hFX (PDB: 9FVW [http://doi.org/10.2210/pdb9FVW/pdb], 1.90 Å resolution), respectively, both refined with complete product occupancy (data collected under cryogenic conditions at the Diamond Light Source), implying single, O2-dependent substrate turnover in crystallo (Fig. 2c+d, Supplementary Fig. 13). Both product-complex structures reveal near identical AspH folds (Fig. 2c+d, Supplementary Fig. 13c), very similar to those of the corresponding substrate complexes. A mixture of 2OG and succinate (each modelled with 50% occupancy based on best agreement with electron density and B-factors) was refined in both product structures, consistent with partial re-binding of 2OG to the active site after substrate hydroxylation, as 2OG was present in excess during crystallization. Because AspH performs only a single turnover in crystallo, the large hydroxylated EGFD product remains bound due to lattice and domain interactions, whilst the smaller 2OG cosubstrate and succinate coproduct can exchange with the surrounding mother liquor via solvent channels.

The cryogenic AspH:Fe:2OG/succinate:hFX-(3R-OH)-Asp/hFX-(3R-OH)-Asn product structures imply stereospecific oxidation to the (3R)-OH-Asp103hFX and (3R)-OH-Asn103hFX products (Fig. 2c+d), consistent with pioneering work on assigning the configuration of 3-OH-Asn/Asp in EGFDs.15, 67 We validated the product stereochemistry by refining the structures with the (3S)-OH-Asp103hFX and (3S)-OH-Asn103hFX, resulting in models that did not reflect the observed electron density (Supplementary Fig. 13d+e).

The C3 hydroxy group of (3R)-OH-Asp/Asn103h FX was observed to coordinate to Fe trans to H725 in both AspH:Fe:2OG/succinate:hFX-(3R-OH)-Asp/Asn product structures (2.5 Å; Fig. 2c+d), replacing the Fe-bound W2 observed in the AspH:Fe:2OG:hFX-Asp substrate structure and occupying the vacant Fe-coordination site in the AspH:Fe:2OG:hFX-Asn structure. Note the absence of W3 in both product structures (Fig. 2c+d). The orientations of AspH active site residues in both product complex structures are similar to those in the substrate complex structures with the notable exception of the Q627 sidechain, which adopts a conformation that likely enhances interactions with the sidechain carboxylate/amide of Asp/Asn103hFX. The distorted octahedral Fe-coordination geometry observed in the product structures likely affects the electronic and steric properties of their Fe centres, possibly modulating the stability of the product complex and promoting product release in solution. The observed lack of product release in crystallo is likely a result of the interactions of the hydroxylated EGFD product with both the 2OG oxygenase and TPR domains in the crystalline lattice and the reduced conformational flexibility of AspH in crystallo; by contrast, the smaller 2OG cosubstrate remains exchangeable on the experimental timescale; note that AspH binds its substrates via an induced fit mechanism associated with substantial conformational changes involving both the 2OG oxygenase and the TPR domains.18

Optimisation for serial crystallography

Given its ability to probe reactions in crystals with minimal radiation damage, we explored the potential of serial femtosecond crystallography (SFX) to capture AspH substrate and product complexes at physiological temperature and pressure. Crystallisation conditions were optimized to enable robust microcrystal slurry formation (~2 μm x 2 μm x 60 μm) of the AspH:Fe(II):2OG:hFX-Asp complex under strictly anaerobic conditions (4 °C, <2 ppm O2). The first SFX data set was collected at the Macromolecular Femtosecond crystallography (MFX)68 instrument of the Linac Coherent Light Source (LCLS) using the drop-on-demand sample delivery system48 and demonstrated that well-ordered anaerobic AspH microcrystals manifest diffraction suitable for time-resolved analysis (PDB: 9FW0 [http://doi.org/10.2210/pdb9FW0/pdb], 1.95 Å resolution; Fig. 4a, Supplementary Fig. 15a). The structure was very similar to its cryogenic analogue, including with respect to the Fe, 2OG and hFX-Asp binding modes (RMSD = 0.254 Å, Supplementary Fig. 16a). Whilst full occupancy of W2 was observed in the room temperature structure, W3 which was observed in the analogous cryogenic structure, was not detected. This change in the solvent network did not affect the conformation of Q627. These results confirmed the stability of the AspH complex under near physiologically relevant temperature and established a robust foundation for subsequent SFX and XES experiments.

Subsequent optimization, including washing of microcrystals to remove excess Fe(II), which may interfere with simultaneously performed XES analysis, yielded an improved structure (PDB: 9HO2 [http://doi.org/10.2210/pdb9HO2/pdb], 1.83 Å resolution; Fig. 4b, Supplementary Fig. 15b), with a fold being highly similar to that of the previously obtained room temperature structure (RMSD = 0.159 Å, Supplementary Fig. 16b). The apparent full occupancy of Fe(II) and 2OG confirmed that washing of the crystals with precipitant did not affect Fe(II)/2OG binding. However, the Fe-W2-W3-Q627-Asp103hFX hydrogen bonding network was partly restored in the improved room temperature hFX-Asp complex: Although both W2 and W3 were observed in full occupancy, the Q627 sidechain was observed in two conformations: one in which the sidechain amide is positioned to interact with the Asp103hFX carboxylate (3.0 Å) and W3 (2.9 Å) and one positioned to interact with the W625 (2.5 Å) and S668 (3.1 Å) sidechains.

Monitoring AspH catalysis in crystallo using serial crystallography and emission spectroscopy

To monitor AspH catalysis over different time scales at physiologically relevant temperatures, complementary experimental procedures were employed: (i) rapid, millisecond–second O2 exposure of microcrystals at the LCLS using the drop-on-demand system,48 enabling precise control of reaction time before diffraction (“in crystallo turnover” experiments); and (ii) slower, passive air exposure of crystallisation samples for 2–24 h at 4 °C, enabling observation of late-stage or equilibrium product states.69, 70 The controlled O₂ delivery at LCLS was chosen to capture early catalytic intermediates and single-turnover events that cannot be resolved through bulk air exposure.

We first characterised a relatively short-timescale reaction by exposing the microcrystal slurry to O₂ for 1.5 s using the drop-on-demand tape-drive sample-delivery system at LCLS. Regulation of the O2 exposure time by altering the tape drive speed48 yielded a room temperature AspH product structure with clear density for (3R)-OH-Asp103hFX (PDB: 9HO1 [http://doi.org/10.2210/pdb9HO1/pdb], 1.85 Å resolution; Fig. 4c, Supplementary Fig. 15c), indicating complete substrate turnover. The fold of this structure closely resembles that of the cryogenic structure (RMSD = 0.296 Å), with minor differences in the conformation of the Q627 sidechain (Supplementary Fig. 16c). Only one conformation of the Q627 sidechain was apparent in which it is positioned to interact with the (3R)-OH-Asp103hFX sidechain carboxylate (2.9 Å). By contrast, two conformations of the Q627 sidechain were observed in the corresponding anaerobic room temperature structure with the substrate (Supplementary Fig. 16c).

Isomorphous difference maps, which inform on local chemical transformations, were calculated between the anaerobic AspH:Fe(II):2OG:hFX-Asp room temperature complex and the product complex obtained after 1.5 s O2 exposure, to visualize site-specific changes accompanying turnover (Fig. 4d,f,h). The isomorphous difference maps reveal negative density for W2 and W3 indicating their absence in the O2-exposed structure. Clear positive density was observed corresponding to a (3R)-OH group on Asp103hFX (Fig. 4d). Negative density was also apparent around the 2OG C1 carboxylate, reflecting conversion of 2OG into succinate. 2OG and succinate were modelled in a 1:1 ratio, an observation suggesting re-binding of 2OG after turnover, as observed for the cryo-product structures, suggesting that the smaller 2OG cosubstrates can exchange with the surrounding mother liquor via solvent channels. Analysis of normalised B-factors in the anaerobic SFX structure and the O2-exposed structure revealed high similarity, suggesting little dynamic variance in the AspH fold following completion of the single turnover event (Supplementary Fig. 17).

Simultaneous collection of X-ray emission spectroscopy (XES) data on the anaerobic crystals and the 1.5 s O2-exposed crystals informed on the Fe-oxidation state in the AspH complex before and after O2 reaction (Fig. 4i+j, Supplementary Fig. 18). The XES analyses reveals little differences in the spectra of the anaerobic and the 1.5 s O2-exposed crystals. This observation contrasts with observations for related systems, e.g. IPNS or sMMO, where a distinct feature in the XES difference spectrum was observed upon reaction of samples in the Fe(II) state when O2 exposed,25, 71 corresponding to a change from an Fe(II) to an Fe(III) state. The observed full width at half maximum (FWHM) results are indicative of an Fe(II) species in both the anaerobic AspH:Fe:2OG:hFX-Asp and the O2-exposed AspH:Fe:2OG/succinate:(3R)-OH-Asp room temperature complexes based on comparison with data obtained with the anaerobic IPNS:Fe(II):substrate and O2-exposed IPNS:Fe(III):substrate:superoxide complexes.25 The slight broadening of the FWHM of ~50 meV upon O2 mediated oxidation is likely not indicative of Fe-oxidation, as in that case a narrowing of the FWHM would be expected.25, 71, 72 The broadening might be caused by a slight change in the ligand environment of the Fe site instead of a change in oxidation state. These results support the conclusion that, following a single catalytic turnover, AspH returns to the Fe(II) state, consistent with a mechanism involving Fe(II) regeneration (Fig. 4i+j, Supplementary Fig. 18).

We next examined longer equilibration time points by extending the O₂ exposure time, in order to investigate differences in the active site 2OG (present in excess under crystallisation conditions) to succinate ratios.18 AspH:Fe:2OG:hFX-Asp microcrystals were exposed to air at 4 °C for ≥2 h with diffraction data collection using fixed target methods (both with serial synchrotron crystallography (SSX) at DLS and SFX at PAL-XFEL). Both the SSX (PDB: 9NHZ [http://doi.org/10.2210/pdb9NHZ/pdb], 2.40 Å resolution; Fig. 4e+f, Supplementary Fig. 15d) and SFX (PDB: 9HO1 [http://doi.org/10.2210/pdb9HO1/pdb], 2.14 Å resolution; Fig. 4g+h, Supplementary Fig. 15e) AspH structures could be refined with full occupancy for (3R)-OH-Asp103hFX and manifested highly similar overall folds (RMSD = 0.196 Å). Similar to the cryogenic product structure, replacement of the Fe-coordinating W2 by the product hydroxyl-group was observed along with a lack of W3. As observed in the 1.5 s O2 exposed structure, B-factor analysis showed high overall similarity to the anaerobic structure (Supplementary Fig. 17).

Evidence for 2OG was not observed in the SSX and SFX structures obtained at longer time points (≥ 2 h O2 exposure), contrasting with the structure obtained after 1.5 s O2 exposure (Supplementary Fig. 16d); complete EGFD hydroxylation was observed for both 1.5 s and ≥ 2 h O2 exposure. The presence of 2OG and succinate in the short time point may reflect (re)binding of 2OG subsequent to hydroxylation. At the longer time points, 2OG in solution may be depleted and succinate increased due to uncoupled solution catalysis arising from solvated AspH outside the crystal lattice or AspH at crystal surfaces, consistent with the observation of only succinate at the active site. Thus, while each AspH molecule within the crystal undergoes a single catalytic event, solution-phase turnover progressively alters the small-molecule composition available for exchange with the crystal (Supplementary Fig. 19).Data for the SSX structure were collected ~2-6 h post exposure to air reveal full occupancy of succinate, which was positioned to interact with Fe in a bidentate manner via a single carboxylate (Fig. 4e+f). By contrast, data for the SFX structure collected ~12-24 h post air exposure reveal ~0.85 occupancy of succinate (Fig. 4g+h), with the conformation of succinate differing from that observed in the SSX structure: succinate is positioned to coordinate the Fe in a monodentate mode, trans to H679. The Fe site trans to W1 featured additional density for which a H2O molecule was modelled (Supplementary Fig. 16e). Analysis of the electron density for the >2 h O2 exposed structure also indicates that the Fe-occupancy decreases upon exposure to air (Fig. 4f+h).

The combined results inform on a possible mechanism promoting succinate displacement from the active site following substrate hydroxylation, i.e., binding of the Fe-binding succinate carboxylate alters from a bidentate to a monodentate being mode, possibly correlating with strengthening of the interaction of the other succinate carboxylate with the R735 guanidinium group. The hydroxylated product and succinate can then dissociate enabling water and 2OG rebinding.

Mass spectrometry studies of oxygen-atom incorporation

Given the potential for O2 ligand exchange between Fe-coordinated waters (W1, W2) and the differing binding modes of Asp- and Asn-containing substrates, the source of the hydroxyl oxygen in the AspH products is of interest (Fig. 5). To investigate whether O2 is the source of the O atom incorporated into the EGFD substrate, we incubated mixtures containing AspH, Fe(II), 2OG, LAA, and hFX-Asp or hFX-Asn with 18O2 or H2 18O using a customised Schlenk-line setup25 and analysed the samples by LC coupled to a QTOF mass spectrometer (Fig. 5, Supplementary Fig. 20). Controls without 2OG revealed no turnover with only the substrate mass being detected, i.e., for hFX-Asp: 1047.2 Da (M+4H]4+). Incubation of the reaction mixture with 16O2 gave the (3R)-16OH-Asp103hFX product with a mass of 1051.2 Da (M+4H]4+), i.e., 4 mass shifts in the m/z = 4 charge state with respect to the substrate (Supplementary Fig. 20-21). Incubation of the mixture under 18O2 gave the (3R)-18OH-Asp103hFX product with a mass of 1051.7 Da (M+4H]4+), i.e., with relative mass increments of 0.51 in the m/z = 4 charge state compared to the 16O2 results (Fig. 5b); the 1051.2 Da (M+4H]4+) masses were not observed in the 18O2 incubation mixtures. Analogous results were obtained when substituting hFX-Asp with hFX-Asn (Fig. 5c). Variations in the 2OG concentration (from 1 to 5 equiv.) and pH (7.0 – 8.5) did not affect the (3R)-OH-Asp103hFX product masses (Supplementary Fig. 22-23). Analysis of the mass shifts associated with the conversion of 2OG (146.03 Da) to succinate (120.04 Da; mass difference of 25.99 Da) in mixtures incubated with 18O2 demonstrate that one 18O atom from 18O2 is incorporated into succinate (Fig. 5d). Comparison of the calculated and observed mass envelopes for 18O- and 16O-incorporation revealed 90-95% incorporation of 18O in the products using 97% 18O2 gas (Supplementary Fig. 21).

Incubations were performed using 16O2 in buffer carefully made with H218O. The results reveal formation of the hydroxylated (3R)-OH-Asp103hFX product with a mass of 1051.21 Da (M+4H]4+), showing no evidence for 18O incorporation from H218O (Fig. 5e, Supplementary Fig. 20). The combined observations clearly indicate that O2 is the sole source of the O atom incorporated into EGFD substrates during AspH catalysis, irrespective of whether an aspartate- or asparagine-residue undergoes hydroxylation.

Methods
AspH production and purification

Purified (>95% by SDS-PAGE and MS) recombinant human AspH315-758 was produced using E. coli BL21 (DE3) pLysS competent cells (Agilent Technologies) as reported.53 In brief, AspH315-758 was produced from a pET-28a(+) construct encoding N-terminally His6-tagged AspH315-758 (UniProt Q12797), transformed into BL21 (DE3) pLysS cells. Single colonies were used to inoculate 2×YT starter cultures containing kanamycin (50 µg·mL-1), which were then used to seed large-scale expression cultures in autoinduction medium supplemented with kanamycin. Cultures were grown at 37 °C until an OD600 of 0.8, then at 18°C overnight. Cells were harvested by centrifugation (≈11,800 × g, 4°C) and stored at −80°C. For purification, frozen cell pellets were resuspended in binding buffer (50 mM HEPES, pH 7.5, 500 mM NaCl, 5 mM imidazole) supplemented with EDTA-free protease inhibitor cocktail and DNase I. Cells were lysed using a high-pressure cell disruptor, and insoluble material was removed by centrifugation (≈58,000 × g, 30 min, 4°C). The clarified lysate was filtered and loaded onto a Ni(II)-affinity column pre-equilibrated in binding buffer. After washing with buffer containing 40 mM imidazole, His6-tagged AspH315–758 was eluted using an imidazole gradient up to 500 mM. To obtain untagged AspH315–758, pooled Ni(II)-affinity fractions were treated with thrombin (0.11 units per mg His6-tagged AspH315–758) and dialysed overnight at 4°C against binding buffer to remove imidazole and enable tag cleavage. The reaction mixture was applied to reverse Ni(II)-affinity chromatography, and AspH315–758 was collected in the flow-through and wash fractions. For preparation of metal-free protein, the combined AspH315–758 fractions were dialysed into gel filtration buffer (50 mM HEPES, pH 7.5, 150 mM NaCl) supplemented with EDTA (30 mM) and 1,10-phenanthroline (5 mM). The sample was then concentrated and further purified by size-exclusion chromatography on a Superdex 75 column (Cytiva) using Chelex-treated gel filtration buffer. Purified AspH315–758 was concentrated (typically 50–350 µM), aliquoted, flash-frozen in liquid N2, and stored at −80°C.

Preparation of AspH crystals for cryo-crystallography

An AspH crystallization solution was prepared by mixing AspH (~370 μM in 50 mM HEPES, pH 7.5, 150 mM NaCl), 2OG (2 mM), and ferrous ammonium sulphate x 6 H2O (FAS) (1 mM), followed by the addition of hFX-Asp or hFX-Asn (~800 μM) at 4 °C. A crystallization screen was conducted by varying the pH (0.1 M bis-tris propane, pH 6.5–8.5 in 0.5 pH increments, vertical axis) and PEG concentration (PEG 3350, 18–22 in 1% increments, horizontal axis), with 0.2 M NaBr or KSCN. AspH crystals were grown using the hanging drop method by combining 1 μL of reservoir solution with 2 μL of protein solution, using 500 μL of precipitant solution in 24-well hanging drop VDX plates (Hampton Research) within an anaerobic chamber (Belle Technologies; <2 ppm O₂). The plate was then transferred to a fridge and maintained at 4 °C. Crystals formed within 24 to 72 h. Crystals were supplemented with cryoprotectant (mother liquor with 25% v/v PEG 400, mixed 1:1 with the crystal-containing drop), harvested using a nylon loop, and cryocooled by rapid plunging into liquid nitrogen before data collection. Diffraction data for single crystals were collected at 100 K using synchrotron radiation at the Diamond Light Source beamline I03. The data were indexed, integrated, and scaled using the Xia2 pipeline (Supplementary Table 2). Structures were solved by molecular replacement with Phaser,83 using PDB entry 8RE9 [http://doi.org/10.2210/pdb8RE9/pdb]40 as the search model. Structures were iteratively refined using PHENIX (version 1.21.1.5286)84 and manual model building using COOT (WinCOOT 0.9.8.7).85 For product complex structures, 2-oxoglutarate (2OG) and succinate were modelled as alternative ligands occupying the same cosubstrate binding site and refined with reciprocal occupancies constrained to a combined occupancy of 1.0, with final values selected based on best agreement with electron density, ligand and neighbouring residue B-factors, and overall refinement statistics.

Sample preparation for SFX and SSX experiments

AspH315-758 seed crystals were prepared in aerobic conditions using a batch method; AspH (100 μL of ~20 mg·mL-1 AspH315-758 in 50 mM HEPES, pH 7.5 and 150 mM NaCl) was mixed with MnCl2 (2.5 μL of 100 mM in Milli-Q), 2OG (2.5 μL of 100 mM in Milli-Q), and hFX-Asp (1.2 mg), and incubated at 4 °C for 5 min prior to mixing with precipitant solution (640 μL of 0.1 M bis-tris propane pH 7.5, 0.1 M KSCN, 16% v/v PEG 3350) and single crystal seed stock solution (10 μL). The crystallisation solution was mixed carefully and pipetted into the wells (60 μL) of a 96-well plate (PS, half area, clear, microlon, med. binding, Greiner Bio-One). The plate was sealed (Polyolefin StarSeal, Starlab, UK) and incubated at 4 °C for at least 14 h. The wells were then combined and prepared as seeds using the Seed Bead Kit (Hampton Research, USA) following the manufacturer’s instructions.

Anaerobic (<2 ppm O2) microcrystallisation was performed using a batch method; AspH (500 μL of ~20 mg·mL-1 AspH315-758 in 50 mM HEPES pH 7.5 and 150 mM NaCl) was mixed with FAS (12.5 μL of 100 mM in Milli-Q), 2OG (12.5 μL of 100 mM in Milli-Q), and hFX-Asp (3 mg) in an anaerobic chamber (Belle Technology, UK) and incubated at 4 °C for 5 min prior to mixing with the precipitant solution (3.2 mL of 0.1 M bis-tris propane pH 7.5, 0.1 M KSCN, 16%v/v PEG 3350) and a seed stock solution (10 μL). The resultant crystallisation solution was mixed carefully and divided into the wells (60 μL) of a small volume 96-well plate (PS, half area, clear, med. binding plates, Greiner Bio-One). The plate was sealed (Polyolefin StarSeal, Starlab, UK) and incubated at 4 °C for at least 48 h. The microcrystal slurry was pooled, allowed to settle at 4 °C, and subsequently washed 2x with precipitant solution to remove excess Fe. Microcrystals were stored at 4 °C.

Data collection of SFX and SSX datasets

Room temperature diffraction data for microcrystal slurries were collected at the Macromolecular Femtosecond Crystallography (MFX) beamline of LCLS, i24 of the Diamond Light Source (DLS), or at the nano-crystallography and coherent imaging (NCI) beamline of PAL-XFEL.86, 87

At the LCLS, the ADE drop-on-tape method was used to collect anaerobic diffraction data for the AspH:Fe:2OG:hFX-Asp complex.48 Droplets were ejected onto the conveyor belt at room temperature in a He atmosphere at a flow rate of 6 μL/min. The X-ray wavelength was 1.27 Å (9.83 keV) with a data collection rate of 30 Hz, a pulse photon energy of 1.5-2 mJ, a pulse length of 35 fs and a beam size of 2.5 x 2.5 μm2. X-ray diffraction data (XDR) was collected with a Rayonix MX340-HS detector. A structure of AspH:Fe:2OG:hFX-Asp that was exposed to O2 for 1.5 s was obtained using the same conditions, in addition to passing the droplets through a 100% O2 reaction chamber at a conveyor belt speed of 99 mm/s. The AspH:Fe:2OG/succinate:(3R)-OH-Asp103hFX complex structures were obtained by exposing the anaerobically prepared crystal slurries to air. At the PAL-XFEL, fixed-target data collection was used. Samples (60 μL) were loaded on a 41 μm mesh, and data was collected at a step size of 50 μm. The X-ray wavelength was 1.30 Å (9.5 keV) with a data collection rate of 30 Hz, and a detector distance of 123 mm. XDR was collected with a Rayonix MX225-HS detector. At DLS, fixed-target data collection was used, samples (100 μL) were loaded onto a silicon chip.88 The X-ray wavelength was 1.00 Å (12.4 keV) with a pulse photon energy of 12.4 keV, and a beam size of 7 μm x 7 μm. XDR was collected with a Dectris PILATUS3 6M detector.

X-ray emission spectroscopy

X-ray emission data were collected in tandem with diffraction data using a multicrystal wavelength-dispersive hard X-ray spectrometer in von Hamos geometry with the analyser crystals above the X-ray interaction point and the position sensitive detector at 90 degrees from the beam direction in the horizontal plane.89, 90 An array of three cylindrically bent (R = 250 mm) LiNbO3 (234) analyser crystals was placed 250 mm above the interaction point with the centre of the crystals at 80.69 degrees with respect to the interaction point, collecting both Fe Kα lines on an ePix 100 detector with its centre located ~82 mm to the side of the X-ray interaction point. Spectra from a Fe-metal foil were collected for calibration. The XES data were pedestal corrected to account for differences in noise of the detector pixels and for background subtraction, a slice on either side of the region of interest (ROI) was selected. A row-by-row first order polynomial fitting scheme was utilized for the initial two-dimensional background subtraction. To bring the baseline of the spectra to zero, a one-dimensional background subtraction (also utilizing a first order polynomial fitting scheme) was also deployed. Spectra were smoothed using a Savitzky-Golay filter with a window length of 9 and polynomial order of 3 and subsequently were area normalized. Full Width Half Max (FWHM) values for the K α1 line were calculated numerically using an energy window of 6399 to 6406.5 eV. Error bars for the FWHM were calculated using a bootstrapping procedure as described.91

Data processing and model building of SFX datasets

Data processing was carried out as described.47 In brief, during the SFX experiments, AspH data were monitored and processed using dials.stills_process-based scripts (in the case of PAL-XFEL data)92 and cctbx.xfel (in the case of LCLS data93) using dials.stills_process to index and integrate diffraction images. The initial estimation of detector geometry (Beam centre and distance) was measured from Ag(I) behenate (in the case of LCLS data) or ceric oxide (in the case of PAL-XFEL data) powder diffraction patterns. A first round of detector metrology refinement was performed,94 followed by dials.stills_process based strong spots indexing. One or more rounds of geometry refinement were carried out until no significant increasing of the indexing rate was observed. The detector geometry was checked and refined periodically during experiments to identify any shifts of the sample position. Integrated data were scaled and merged using cctbx.xfel.merge, with a reference model that has unit cell parameters of a=50.796 Å, b=88.16 Å, c=125.161 Å, α=β=γ=90° (space group P212121). The merging resolution was selected based on several criteria, including where the multiplicity drops below ten-fold, the CC1/2 no longer decreases uniformly and the completeness beyond 99%.

Structures were solved by molecular replacement with Phaser,83 using PDB entry 8RE9 [http://doi.org/10.2210/pdb8RE9/pdb] as the search model. Structures were iteratively refined using PHENIX (version 1.21.1.5286)84 and manual model building using COOT (WinCOOT 0.9.8.7).85

B-factor analyses were performed using PyMOL (version 3.0, Schödinger, LLC). For each refined structure, main chain atoms were selected and B-factors were extracted. The B-factors were normalised using a Z-score approach, computing the mean and the standard deviation of the B-factors. If the standard deviation was zero (indicating no variance), we used 1 as a default value to avoid division by zero. B-factors were then normalised by subtracting the mean and dividing by the standard deviation. Then, the normalised B-factors were applied to the corresponding main chain atoms, and the structure was rebuilt. B-factors were visualised in a blue-white-red spectrum between -2 and 2 to highlight the variations. The same protocol was applied separately to sidechain atoms.

Sample preparation for UV-Vis experiments and solution EPR experiments

Samples were prepared under anaerobic conditions (<2 ppm O2) in an anaerobic chamber (Belle Technology, UK). Stock solutions of UV-Vis buffer (50 mM HEPES, pH 7.5, 150 mM NaCl, 20% glycerol) and solids (FAS, 2OG, and hFX-Asp) were placed in the anaerobic chamber to remove residual O2. Stock solutions of apo-AspH (~68 mg·mL–1 AspH315-758, 1.3 mM) were transferred into the anaerobic chamber prior to use. Solutions of 2OG (100 mM in H2O) and FAS (100 mM in H2O), were prepared in the anaerobic chamber. The total volume was 75 µL and contained 1 mM AspH, 4.75 mM FAS (4.75 equiv.), 5 mM hFX-Asp (5 equiv.) and residual UV-Vis buffer. Nitric oxide (NO; 1,000 ppm in N2, 60 min) exposure of samples involved the use of a custom glass apparatus as described.25

UV-Vis experiments

UV-Vis spectra were recorded using a Carry 3500 UV-Vis Compact Peltier spectrometer (Agilent, US). Samples were prepared and data collected in a rectangular, quartz, ultra-micro, open top cuvette (2.5 x 5 mm, 10 mm pathlength, minimal volume: 70 μL), and sealed with a rubber septum in an anaerobic chamber (<2 ppm O2, Belle Technology, UK). Data were recorded at 293 K, with a scan range from 200-800 nm (1 nm data interval). The UV-Vis spectra were processed with the Carry UV workstation (Agilent, US).

Sample preparation for microcrystal EPR experiments

Anaerobic (< 2 ppm O2) microcrystallisation was performed using the batch method as described above, using either hFX-Asp or hFX-Asn, and FeSO4·7H2O instead of FAS. The microcrystal slurry was pooled and allowed to settle at 4 °C, then washed 2x with precipitant buffer to remove excess Fe. The precipitant buffer (1 mL of: 0.1 M bis-tris propane, pH 7.5, 0.1 M KSCN, 16% v/v PEG 3350) was incubated for 30 min with pure NO gas (>99.9%) using a custom glass apparatus, as described,25 to saturate NO in the buffer. Excess precipitant solution was removed from the microcrystal slurry and NO-saturated precipitant solution was added in a 1:1 (v/v) ratio.

EPR experiments

Samples were prepared and transferred into a 1.2 ID x 1.6 mm OD clear-fused quartz tube ("Ilmasil" quartz from Qsil GmbH), followed by cryo-cooling and storage in liquid N2. X-band continuous wave (CW) EPR data were collected using a Bruker BioSpin EMXmicro spectrometer equipped with a Premium bridge, a Bruker ER4122-SHQE-W1 resonator, an Oxford Instruments ITC-503S temperature controller, and ESR900 helium flow cryostat. Photolysis was achieved with unfiltered white light LED of a Schott KL2500 source of irradiance 670 mW/cm2 delivered by a liquid light guide to the 66% transmission resonator window grill. The EPR simulation and data processing were performed in the MATLAB 24.1.0 scripting environment (The MathWorks, Inc., Natick, NJ) with the simulation routines from the EasySpin package, version 6.0.0.95

AspH kinetics

Kinetic studies with isolated recombinant AspH315-758 were performed using solid phase extraction coupled to mass spectrometry (SPE-MS) assays, as described.57 In brief, AspH-catalysed hydroxylation of hFX-Asn/hFX-Asp was monitored in real time using SPE-MS. The AspH reaction was started by addition of an appropriate amount of AspH315-758 to the substrate mixture, containing hFX-Asn/hFX-Asp, LAA, 2OG, and Fe(II), in buffer (25 mM HEPES, pH 7.5, 50 mM NaCl, 20 °C) in a 2 mL 96 deep well plate (Greiner). Final concentrations: AspH315-758 (0.05 μM), hFX-Asn (4.0 μM), LAA (100 μM), and the shown concentrations of 2OG and FAS (Supplementary Fig. 4). Samples were purified using a C4 cartridge and analysed using a RapidFire RF 365 high-throughput sampling robot (Agilent) attached to an iFunnel Agilent 6550 accurate-mass quadrupole time-of-flight mass spectrometer operated in the positive ionization mode. The m/z +4 charge states of hFX-Asn/hFX-Asp was used to extract ion chromatogram data, peak areas were integrated using RapidFire Integrator 4.3.0 (Agilent). Data was exported into Microsoft Excel and used to calculate the percentage conversion; data was analysed using GraphPad prism to determine turnover numbers (kcat) and Michaelis constants (Km) for 2OG and Fe(II) with hFX-Asn (Fig. 1).

18O and H 18O AspH assays

Labelling experiments were conducted under controlled 18O2 (97% 18O, Merck) conditions using a Schlenk line setup. Initially, all to-be-used solutions and solids were transferred into an anaerobic chamber (Belle Technology, O2 concentration: <2 ppm) and equilibrated overnight. AspH315-758 (1 μM) was mixed with FAS (final concentration: 20 μM), 2OG (20 μM, 50 μM, or 100 μM), L-ascorbic acid (100 μM) and hFX-Asp (20 μM) or hFX-Asn (20 μM) in 50 mM HEPES (pH 7.0, 7.5, 8.0, or 8.5) with 50 mM NaCl (total volume: 200 μL). The anaerobic sample was placed into a gas-tight customized 96-well plate holder with a gas inlet. This plate holder setup was removed from the anaerobic chamber and connected to the Schlenk line, which was attached to an 18O2 source (97% 18O, Merck). Residual O2 was purged from the system with alternating Ar and vacuum cycles. To create a mild vacuum in the plate holder setup, the pressure was adjusted to 700 mbar, and the gas inlet was opened. The system was then filled with 18O2 and equilibrated for 5 min. The setup was sealed, and samples were exposed to 18O2 for at least 60 min; the reaction was subsequently quenched by addition of 10% v/v formic acid. MS-analyses were performed by LC-MS using a Xevo G2-XS (Waters) equipped with an ACE 5-AQ (Avantor), and analysed using MassLynx. For the control experiments in 18O-water, AspH was buffer exchanged into HEPES buffer (50 mM in 18O-water (>98 atom-% 18O), pH 7.5) using Zeba™ Spin Desalting Columns (ThermoFisher, UK). All cofactor stock solutions were prepared in 18O-water and then diluted to the final working concentration using HEPES (50 mM in 18O-water (>98 atom % 18O), pH 7.5). The assay mixtures were then prepared as described above.

Detection of 2OG and succinate in assay samples was performed by Liquid Chromatography Mass Spectrometry (LCMS) using an Agilent 1290 infinity II LC system connected to an Agilent 6550 accurate mass iFunnel quadrupole time of flight (QTOF) mass spectrometer. Assay samples were diluted 10-fold in 100% acetonitrile; 4 μL of the sample was injected and loaded onto a InfinityLab Poroshell 120 HILIC-Z, 2.1 x 50 mm, 2.7 μm column (Agilent). 2OG and succinate were separated using a step wise gradient (0 min – 100% Solvent B, 5.0 min - 100% Solvent B, 15.0 min - 0% Solvent B, 20 min - 0% Solvent B, 25 min - 100% Solvent B, 40.0 min - 100% Solvent B), using a flow rate of: 0.25 mL/min; Solvent A: LCMS grade water containing ammonium acetate (10 mM, pH 9.0), Solvent B: acetonitrile:solvent A (90:10 v/v). The mass spectrometer was operated in the negative ionisation mode with a drying gas temperature (200 °C), drying gas flow rate (13 L/min), nebulizer pressure (40 psig), sheath gas temperature (300 °C), sheath gas flow rate (12 L/min), capillary voltage (3000 V), nozzle voltage (0 V), fragmentor voltage (125 V). Acquired data were analysed using Agilent MassHunter Qualitative Analysis (Version B.07.00) software.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "I think this is an interesting study that is worthy of publication. Some comments for the authors are listed below.\nIn the abstract, the observation of a water ligand throughout catalysis does not immediately rule out the possibility of oxygen exchange through coordination of additional water ligands or other mechanisms. Also – if the 18O data has not be analyzed quantitatively – can it be concluded that O2 is the sole oxygen source? Some Fe/2OG enzymes undergo exchange of the oxygen in the ferryl or ferric hydroxide state, making it difficult to conclude the source of the O atom from the isotope labeling alone. While I think the underlying conclusion here is probably correct – I’m not sure that the data analysis is sufficiently rigorous to make the exact claim stated in the abstract.\nIn the Fe•2OG•Asp complex, why is the Q627 side chain modeled with the carbonyl interacting with the D103 side chain? Wouldn’t it make more sense to flip the side chain to have the amide nitrogen interacting with the substrate instead?\nIt would also be nice to show the electron density map for the Q627 and S668 side chains. Are these well-ordered with both substrates?\nOn line 202, I was not sure what the statement “suggesting that 201 the Fe-W2-W3-Q627-Asp103hFX hydrogen bond network does not compromise catalytic efficiency” meant. Are the authors trying to say that the additional interactions observed between the enzyme and substrate in the Asp complex would slow the reaction? My intuition is that it would be the opposite. Tighter substrate binding should accelerate the reaction between iron and O2 as observed in many other Fe/2OG enzymes.\nIn Fig. 3A, it would be nice to show a difference or omit map for the NO and NCS models.\nIn the 18O isotope labeling experiments, it would be ideal to analyze the data quantitatively to show the extent of isotope incorporation.\nSome aspects of the discussion (first 5 paragraphs) are redundant with the results.", "reviewer_2": "This is an extensive and immensely detailed manuscript looking at structures of aspartate/asparagine b-hydroxylase (AspH). The authors have used time resolved serial crystallography and rapid mixing methods to study reactivity in crystals. A large amount of other data accompany the structural work, including NO binding, kinetics and EPR in the early part of the paper. These studies have been carried out to a very high standard but do distract slightly from the more interesting turnover work in crystallo. The product formation (page 10) is presented as a somewhat incremental advance on previous work (refs 15,65). Throughout the paper, there are discussions on small movements of waters and h-bond/conformational changes, with many technical details included that, to me, look like they should be in the Methods. The mechanistic implications are described, but not clearly depicted.\nBearing in mind the outstanding and extensive structural work already published on 2OG enzymes (including from this group), the paper as presented appears more suitable for specialist audience. Shortening and simplification of the descriptions, and a better focus on the main headlines, would be needed if the paper were to be suitable for publication in Nature Comm.", "reviewer_3": "The manuscript by de Munnik and colleagues uses careful structural biology experiments to rationalize the ability of the aspartate/asparagine-β-hydroxylase (AspH) to hydroxylate both Asp- and Asn- residues and defines the key interactions that drive catalysis. AspH targets epidermal growth factor-like domains (EGFDs) for C3 hydroxylation. Major outstanding questions regarding the mechanism of AspH that were addressed by this work include: how AspH can target both Asp- and Asn- residue types, why AspH is incapable of catalyzing halogenation of the substrate, clarifying the source of the oxygen for hydroxylation, and visualizing the key interactions required for substrate turnover and release. Firstly, the manuscript corrects an artifactual interaction of the substrate Asp sidechain carboxylate observed in the inactive Mn-bound structure – the true interaction shows a solvent water molecule that is the likely site for O2 incorporation. The structure of AspH with Asn- and Asp- substrates highlights two key interactions which are further explored by the authors – that of the Gln627 with the substrate carboxylates, the solvent network, and with a nearby Ser668 residue, and that of the consistent W1 water which appears to be indispensable and also provides a rationale for why the enzyme does not halogenate the substrate. The serial crystallography experiments unfortunately don’t add too much nuance to the story, but are carefully analyzed to reveal the appearance of C3 hydroxylation density, disappearance of the solvent network, and disappearance of the 2OG C1 carboxylate. Overall, there are significant insights into mechanistic detail in this work that will help with drug discovery efforts using this enzyme, as well as possibilities for enzyme engineering.\nOverall, the manuscript is well-written, though the nomenclature used to distinguish the different structures is onerous (see comment below). It answers some outstanding questions about AspH mechanism and highlights both the importance of using the biologically relevant metals for structural elucidation as well as complementary spectroscopic techniques. The structures generated by the authors are of sufficient quality to enable molecular dynamics simulations to clarify some of the questions they discuss (i.e. why the varying degrees/rates of Asp-/Asn- hydroxylation for different substrates?) I recommend publication of the manuscript with minor revisions.\nComments:\n1) In the Introduction, it would be useful to discuss the consequence of non-canonical disulfide bonded EGFD hydroxylation.\n2) In the results section, part 1 (analysis of complexes), paragraph 3 – at the end of the paragraph, please add a sentence that summarizes the differences between the structures, in an overarching way that relates to the biochemical questions being addressed.\n3) In the same section, last paragraph, the authors mention a change in Km for 2OG between the two substrates (Asp/Asn). Maybe I missed this, but it would be great to have a biological link to this change in kinetics – is there a preference in cells for a given substrate? Is this a regulatory mechanism somehow?\n4) In the second section of the results, paragraph 2 – please define DNIC for non-experts and why the increased absorption at 300-400 nm is a characteristic of?\n5) Why is isothiocyanate used to mimic the halogenation complex? An active site comparison between AspH and a related halogenase for which this approach was used in the supplement would be helpful.\n6) Can the authors expand on why NCS may be present at full occupancy in the Mn structure but not the Fe structure?\n7) In the last paragraph of the NCS section in the results, there is a mention that the NCS binds to the predicted O2 site rather than the site trans to 2OG – was this expected? Please elaborate on what was meant by this note.\n8) Results, analysis of product complexes: The 2OG and succinate molecules were modelled at 50% in both the Asn and Asp structures. Is this what is expected? Is the hydroxylated product modelled at 100%? Since there is a difference in the Km, should these values be different? How was the occupancy refined?\n9) Same section, paragraph 2, relationship to FIH – the stereocenter is opposite, it is unclear why this is specifically mentioned here. Is this strange? Or does the architecture of the two enzymes effectively determine the stereochemistry, and this is as expected?\n10) Same section, paragraph 3 – missing W3 – this is noted in the text, but there is no follow up on its significance.\n11) Results, microcrystallography. Please explain why serial femtosecond crystallography was used instead of serial synchrotron crystallography. The time delays are more than sufficient to perform these experiments at a synchrotron.\n12) The discussion on the optimization of the first SFX to the second SFX structure (also depicted in figure 4) is not entirely relevant to the biochemical discussion in this paper and adds to an already long results section. I would suggest removing this from the main text and focusing on the structure that is relevant (optimized) and moving this optimization description to the supplementary information.\n13) Same section, and figure 4. The description of the experiments becomes very confusing between describing a 1.5s O2 exposure vs a multi-hour air exposure. The section needs a sentence that describes the experimental pipeline – samples exposed to O2 in a chamber as well as samples left in air for 2-24h. It is unclear to me why choose an O2 chamber rather than just air exposure for the first experiment.\n14) It was unclear to me why the authors are comparing the O2 and air exposures by difference maps (not fully explained in the main text). The figure legend has to be highly simplified by agglomerating the repeated protocols to make the figures and instead pointing at what the maps are showing. It is unclear from the text what conclusions from these different exposures bring.\n15) Same section, last paragraph – “w2/W3 binding may vary in a context dependent manner” – please define what is meant by this in biological terms to explain why this is relevant.\n16) Same section, 2OG and succinate in structures – also confusing. The succinate should be the product of the turnover (according to figure 1). So why is it only at 50% occupancy with the short O2 exposure (1.5 seconds), whereas the product is modelled at 100% occupancy but then full occupancy of succinate is visible after 2 hours? And why does it drop after 24h? Is it substituted by 2OG? Or just unbound? From my interpretation of the results shown, the structures do not really match a coherent biochemical pathway of binding, turnover, unbinding of co-product and rebinding of substrate.\n17) In the discussion, it is alluded that AspH is overexpressed in cancers, but based on the explanation of its function, I was unclear as to how this can impact cancer pathogenesis. A small expansion on the topic would help set the stage for drug discovery\n18) The authors allude to future experiments using time-resolved crystallography. I agree that such experiments are exciting, but such onerous experiments should be justified with a clear statement of what information is missing from the current studies. What would TR-SFX bring beyond this already thorough structural study? How would this impact our understanding of AspH function and medicinal chemistry developments? What are the outstanding questions?\nMinor concerns, for readability\n1) Through the text, the use of the full complex composition for the structures made it harder to follow, especially since for some of the comparisons the change was from the Asp to the Asn substrate (one letter in a long structure acronym). I would urge the authors to consider a change in how these complexes are referred to in the results section, maybe reducing the number of times the full acronyms are used and simplifying the sentences to highlight the differences between the structures being compared.\n2) The results section has an enormous amount of detail on the sometimes very small changes observed in the structures that distracts from the main changes that drive the mechanistic conclusions. I would advise reducing information that can be seen in figures (for example, adding hydrogen bond lengths to the figures rather than the main text) as well as removing information such as overall RMSD values and repeated notes of the similarity of the overall fold of the protein. All these structures are from the same protein, so the overall fold is expected to be the same, and this information does not add to the discussion. A single sentence in the conclusion that mentions this similarity would be sufficient.\n3) Results, Mass spectrometry section – similar concerns as in comment 2. All the mass values are stated in the text. A table would be better suited, with the text mentioning the species identified, rather than the masses observed. The figure legend for 5 should also be simplified to avoid the repetition, and the figure may even be more suitable as supplementary. The masses are visible in the main figure, charge states can be added to the figure.\n4) Discussion, paragraph 1 – please correct Figure X.\n5) The authors should ensure that relevant figure sub-panels are mentioned in the text, rather than only the figure number.\n6) The authors should also try to minimize sentence length, with some sentences being very long and complex.\n7) Figure 2 – the chemical drawings are too small and barely readable\n8) Conclusions, microcrystal optimization. The first paragraph should probably be moved to the discussion section.\n9) Please add to the SI a figure showing the two setups at MFX and PAL", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "POINT-TO-POINT RESPONSE TO THE REVIEWERS' COMMENTS:\nWe thank all the reviewers for their positive concerning our scientific results and very helpful\ncomments to improve the manuscript, which we have addressed as described below.\nReviewer #1\nI think this is an interesting study that is worthy of publication. Some comments for the authors are\nlisted below.\nIn the abstract, the observation of a water ligand throughout catalysis does not immediately rule out\nthe possibility of oxygen exchange through coordination of additional water ligands or other\nmechanisms. Also – if the 18O data has not be analyzed quantitatively – can it be concluded that O2\nis the sole oxygen source? Some Fe/2OG enzymes undergo exchange of the oxygen in the ferryl or\nferric hydroxide state, making it difficult to conclude the source of the O atom from the isotope\nlabeling alone. While I think the underlying conclusion here is probably correct – I’m not sure that\nthe data analysis is sufficiently rigorous to make the exact claim stated in the abstract.\nResponse: We thank the reviewer for pointing this out, and agree that the statement would hold\nmore weight with additional quantitative analysis. Therefore, we have calculated the expected mass\nenvelopes for both full 18O incorporation, full 16O incorporation, and 5%, 10%, and 20% 16O\n2 2 2\nincorporation. A comparison with the experimental data reveals incorporation of 5-10% 16O into the\nalcohol product. Given the purity level of the isotopically labelled 18O (95% purity and lack of\n2\nincorporation of 18O from H 18O) the experimental data supports our conclusion that O is the sole\n2 2\nsource of oxygen incorporation. We have modified the results section and SI to reflect the additional\ndata analysis.\nIn the Fe•2OG•Asp complex, why is the Q627 side chain modeled with the carbonyl interacting with\nthe D103 side chain? Wouldn’t it make more sense to flip the side chain to have the amide nitrogen\ninteracting with the substrate instead?\nResponse: We thank the reviewer for pointing out this modelling detail. To further investigate the\nside-chain orientation of Q627 we include 2mFₒ–DFc density maps contoured at 1.0σ and 3.0σ for the\nQ627/Asn103 region. The maps support the modelled orientation in which the carbonyl of Q627\naccepts a hydrogen bond from the substrate Asn103 N–H. The amide N of Q627 and of the Asn103\nN–H is less well defined in the density (consistent with higher mobility and weaker scattering for\nhydrogen-bearing moieties), which disfavours a flipped conformation in the present maps.\nAdditionally, the substrate Asn carbonyl group is positioned to hydrogen bond with K666, providing a\ncoherent network of interactions that is consistent with the modelled geometry. For clarity in the\nmain text figures, we have omitted the K666–Asn carbonyl contact to highlight the Fe coordination\ngeometry and the principal substrate/protein contacts; the full network is shown in the\nSupplementary Information.\nIt would also be nice to show the electron density map for the Q627 and S668 side chains. Are these\nwell-ordered with both substrates?\nResponse: We observed well-ordered electron density for the Q627 and S668 side chains in our\nstructures. As representative examples, the electron density maps for these residues are shown in\nSupplementary Figure S3a and S3d. To avoid diverting attention from the (co)substrates, products,\nand cofactor, we chose not to include these maps in the main text figures.\nOn line 202, I was not sure what the statement “suggesting that 201 the Fe-W2-W3-Q627-\nAsp103hFX hydrogen bond network does not compromise catalytic efficiency” meant. Are the\nauthors trying to say that the additional interactions observed between the enzyme and substrate in\nthe Asp complex would slow the reaction? My intuition is that it would be the opposite. Tighter\nsubstrate binding should accelerate the reaction between iron and O2 as observed in many other\nFe/2OG enzymes.\nResponse: We agree that the phrasing in the original manuscript was unclear. The sentence has been\nrevised to enhance clarity, by changing “suggesting that the Fe-W2-W3-Q627-Asp103hFX hydrogen\nbond network does not compromise catalytic efficiency” to “suggesting that the Fe-W2-W3-Q627-\nAsp103hFX hydrogen bond network supports catalytic efficiency”.\nIn Fig. 3A, it would be nice to show a difference or omit map for the NO and NCS models.\nResponse: We agree with the reviewer that an isomorphous difference map would improve the\nrepresentation of our data. We have adapted figure 3c to reflect the difference map of NCS in\ncomparison to the NO density.\nIn the 18O isotope labeling experiments, it would be ideal to analyze the data quantitatively to show\nthe extent of isotope incorporation.\nResponse: We thank the reviewer for this valuable suggestion. To address it, we performed a\nquantitative analysis of the 18O isotope-labeling experiments to assess the extent of isotope\n2\nincorporation into the substrate. Using the substrate’s isotopic envelope as a reference, we\ncalculated the expected mass envelopes for reactions carried out under 16O (m/z increase of 4.0 Da\n2\nat charge state 4) and 18O (m/z increase of 4.5 Da at charge state 4) conditions. We also generated\n2\nsimulated envelopes representing 5%, 10%, and 20% admixtures of 16O to model potential\n2\ncontamination from atmospheric oxygen. Comparison of these calculated distributions with our\nexperimental spectrum obtained at pH 7.5 shows that the data are best explained by approximately\n5–10% 16O contribution.\n2 2\nconsistent with minor contamination from the gas source and ambient air. Therefore, we conclude\nthat incorporation of 18O into the substrate is, at least near, complete under our experimental\n2\nconditions. In support of this conclusion, reactions conducted with 18OH (water) showed no isotope\n2\nincorporation. A representative dataset and the quantitative analysis are now included in the revised\nSupplementary Information.\nSome aspects of the discussion (first 5 paragraphs) are redundant with the results.\nResponse: We thank the reviewer for this helpful suggestion. We have revised the discussion to\nreduce redundancy with the results. However, instances where we believe small redundancy is\nnecessary to place the results in a wider context remain unchanged.\nReviewer #2:\nThis is an extensive and immensely detailed manuscript looking at structures of\naspartate/asparagine b-hydroxylase (AspH). The authors have used time resolved serial\ncrystallography and rapid mixing methods to study reactivity in crystals. A large amount of other\ndata accompany the structural work, including NO binding, kinetics and EPR in the early part of the\npaper. These studies have been carried out to a very high standard but do distract slightly from the\nmore interesting turnover work in crystallo. The product formation (page 10) is presented as a\nsomewhat incremental advance on previous work (refs 15,65). Throughout the paper, there are\ndiscussions on small movements of waters and h-bond/conformational changes, with many\ntechnical details included that, to me, look like they should be in the Methods. The mechanistic\nimplications are described, but not clearly depicted.\nBearing in mind the outstanding and extensive structural work already published on 2OG enzymes\n(including from this group), the paper as presented appears more suitable for specialist audience.\nShortening and simplification of the descriptions, and a better focus on the main headlines, would\nbe needed if the paper were to be suitable for publication in Nature Comm.\nResponse: We thank the reviewer for their comments. We have revised the manuscript to improve\noverall readability and accessibility for a broad scientific audience. The text has been streamlined to\nfocus on the key findings—namely, the unique Fe(II) coordination chemistry of AspH, its complete\nturnover observed in crystallo, and the mechanistic insight into oxygen binding and product\nformation. Given the general relevance of these findings to Fe/2OG oxygenase catalysis, oxygen\nsensing, and biomedical targeting of AspH, we believe the study now meets the interdisciplinary\nscope and high impact expected for Nature Communications.\nReviewer #3:\nThe manuscript by de Munnik and colleagues uses careful structural biology experiments to\nrationalize the ability of the aspartate/asparagine-β-hydroxylase (AspH) to hydroxylate both Asp-\nand Asn- residues and defines the key interactions that drive catalysis. AspH targets epidermal\ngrowth factor-like domains (EGFDs) for C3 hydroxylation. Major outstanding questions regarding the\nmechanism of AspH that were addressed by this work include: how AspH can target both Asp- and\nAsn- residue types, why AspH is incapable of catalyzing halogenation of the substrate, clarifying the\nsource of the oxygen for hydroxylation, and visualizing the key interactions required for substrate\nturnover and release. Firstly, the manuscript corrects an artifactual interaction of the substrate Asp\nsidechain carboxylate observed in the inactive Mn-bound structure – the true interaction shows a\nsolvent water molecule that is the likely site for O2 incorporation. The structure of AspH with Asn-\nand Asp- substrates highlights two key interactions which are further explored by the authors – that\nof the Gln627 with the substrate carboxylates, the solvent network, and with a nearby Ser668\nresidue, and that of the consistent W1 water which appears to be indispensable and also provides a\nrationale for why the enzyme does not halogenate the substrate. The serial crystallography\nexperiments unfortunately don’t add too much nuance to the story, but are carefully analyzed to\nreveal the appearance of C3 hydroxylation density, disappearance of the solvent network, and\ndisappearance of the 2OG C1 carboxylate. Overall, there are significant insights into mechanistic\ndetail in this work that will help with drug discovery efforts using this enzyme, as well as possibilities\nfor enzyme engineering.\nOverall, the manuscript is well-written, though the nomenclature used to distinguish the different\nstructures is onerous (see comment below). It answers some outstanding questions about AspH\nmechanism and highlights both the importance of using the biologically relevant metals for\nstructural elucidation as well as complementary spectroscopic techniques. The structures generated\nby the authors are of sufficient quality to enable molecular dynamics simulations to clarify some of\nthe questions they discuss (i.e. why the varying degrees/rates of Asp-/Asn- hydroxylation for\ndifferent substrates?) I recommend publication of the manuscript with minor revisions.\nComments:\n1) In the Introduction, it would be useful to discuss the consequence of non-canonical disulfide\nbonded EGFD hydroxylation.\nResponse: The functional consequences of this are not known buy may reflect a role for AspH in\nredox homeostasis - we have added a comment on this point.\n2) In the results section, part 1 (analysis of complexes), paragraph 3 – at the end of the paragraph,\nplease add a sentence that summarizes the differences between the structures, in an overarching\nway that relates to the biochemical questions being addressed.\nResponse: We thank the reviewer for this helpful suggestion. We have revised the description of the\nAspH:Fe:2OG:hFX-Asp and AspH:Fe:2OG:hFX-Asn complex structures to improve clarity and\nreadability and to make the differences between the structures more apparent. In particular, we have\nsimplified the description of W2 and the associated hydrogen-bonding network, and clarified how\nthese structural differences relate to Fe coordination flexibility, O binding, and substrate-dependent\n2\neffects on catalysis. While we did not add a separate summary sentence, the revised text now\nintegrates these key points directly into the flow of the paragraph, making the biochemical\nimplications clearer and easier to follow.\n3) In the same section, last paragraph, the authors mention a change in Km for 2OG between the\ntwo substrates (Asp/Asn). Maybe I missed this, but it would be great to have a biological link to this\nchange in kinetics – is there a preference in cells for a given substrate? Is this a regulatory\nmechanism somehow?\nResponse: We thank the reviewer for this helpful comment. We agree that the observed difference in\nKₘ for 2-oxoglutarate between hFX-Asp and hFX-Asn substrates may have biological relevance. We\nhave now added a statement in the Results section noting that, while the functional implications of\nthis kinetic difference remain unclear, it raises the possibility that substrate-dependent variations in\n2OG affinity could influence the extent of EGFD hydroxylation in cells. We also note that further work\nwill be required to test whether such differences play a regulatory role in AspH activity in vivo.\n4) In the second section of the results, paragraph 2 – please define DNIC for non-experts and why the increased absorption at 300-400 nm is a characteristic of?\nResponse: We thank the reviewer for this helpful comment. We have revised the text to define DNIC\n(dinitrosyl iron complex) as a nitrosyl–iron species that forms when NO binds to Fe(II) at morethan\none coordination site. We have also added an explanation that the increased absorption at 300–400\nnm is characteristic of Fe–NO charge-transfer transitions, consistent with formation of metal–nitrosyl\ncomplexes and changes in the Fe(II) coordination environment,\n5) Why is isothiocyanate used to mimic the halogenation complex? An active site comparison\nbetween AspH and a related halogenase for which this approach was used in the supplement would\nbe helpful.\nResponse: We thank the reviewer for this comment. We have added citations to relevant literature\ndescribing how pseudohalides (N , NCS, and NCO) are used to mimic halide binding in structural and\n3\nmechanistic studies. We have also revised the text to make it clearer for non-expert readers that, in\nAspH, isothiocyanate binds at the O -binding site rather than at the W1 site (the position trans to the\n2\n2OG C2 carbonyl) where halides typically coordinate in halogenases. This clarification has been\nadded to both the Results and Discussion sections.\nA comparative figure illustrating the active sites of a representative halogenase, a canonical\noxygenase, and AspH is already included in the manuscript (Fig. S2), highlighting the unique AspH\ncoordination environment and explaining why it supports hydroxylation rather than halogenation\nchemistry.\n6) Can the authors expand on why NCS may be present at full occupancy in the Mn structure but not\nthe Fe structure?\nResponse: We thank the reviewer for this observation. We note that the Mn(II) ion in AspH typically\nexhibits a slightly larger ionic radius and more rigid coordination geometry compared to Fe(II), which\nmight result in tighter ligand binding and reduced coordination flexibility. This may favour full NCS\noccupancy in the Mn complex. In contrast, the catalytically relevant Fe(II) centre displays a more\ndynamic coordination environment, as also reflected by the partial occupancy of W2 in the Fe(II)\nsubstrate complexes and its ability to exchange ligands during turnover. This increased lability likely\naccounts for the lower apparent occupancy of NCS in the Fe(II) complex. Together, these differences\nfurther emphasize the importance of using the native Fe(II) metal when interpreting AspH’s\ncoordination chemistry and catalytic mechanism. We have added a comment in Figure S12 legend to\nkeep the text concise.\n7) In the last paragraph of the NCS section in the results, there is a mention that the NCS binds to the\npredicted O2 site rather than the site trans to 2OG – was this expected? Please elaborate on what\nwas meant by this note.\nResponse: We agree this point required clarification. In 2OG-dependent halogenases, halide ions (or\npseudohalides) typically bind at the metal coordination site trans to the 2OG C2 carbonyl group,\nwhere they are positioned to enable halogen transfer, after Fe(IV)=O equivalent reaction . In\ncontrast, in AspH the isothiocyanate ligand binds at the O binding site trans to His725 (the W2\n2\nposition), not the site trans to the 2OG carbonyl (W1 position). This was unexpected because the\nFe(II) coordination geometry of AspH superficially resembles that of halogenases. The observed NCS\nbinding at W2 instead of W1 indicates that AspH’s primary coordination environment is not pre-\norganized for halogen transfer chemistry, consistent with its exclusive hydroxylase activity. We have\nclarified this point in both the Results and Discussion sections.\n8) Results, analysis of product complexes: The 2OG and succinate molecules were modelled at 50%\nin both the Asn and Asp structures. Is this what is expected? Is the hydroxylated product modelled at\n100%? Since there is a difference in the Km, should these values be different? How was the\noccupancy refined?\nResponse: We thank the reviewer for this helpful comment. It was indeed unexpected that both 2OG\nand succinate refined to approximately 50% occupancy in the product complex structures. This likely\nreflects partial re-binding of 2OG after turnover, which is plausible given that 2OG was present in\nexcess relative to substrate during crystallization. Since AspH catalyses only a single turnover within\nthe crystal lattice, the large 39-mer EGFD peptide substrate and its hydroxylated product remain\nbound in the active site (modelled at 100% occupancy), whereas the smaller 2OG/succinate\nmolecules can exchange within the solvent channels.\nWe therefore interpret the mixed 2OG/succinate occupancy as evidence of 2OG re-binding to the\nactive site following hydroxylation, rather than as an indicator of incomplete turnover. This\ninterpretation is consistent with the serial crystallography data collected after extended O exposure\n2\nat room temperature, where only succinate was observed, indicating that all residual 2OG had been\nconverted over time. We have modified the text to make this clear.\nFor refinement, the occupancies of 2OG and succinate were adjusted reciprocally (in 0.1 increments)\nuntil their combined occupancy totalled 1.0 (100%), with corresponding restraints applied to\nmaintain stable B-factors. The hydroxylated EGFD products were refined at full (100%) occupancy in\nall product complex structures.\n9) Same section, paragraph 2, relationship to FIH – the stereocenter is opposite, it is unclear why this\nis specifically mentioned here. Is this strange? Or does the architecture of the two enzymes effectively determine the stereochemistry, and this is as expected?\nResponse: We thank the reviewer for this comment. We have removed this statement in the text to\nincrease the flow and readability.\n10) Same section, paragraph 3 – missing W3 – this is noted in the text, but there is no follow up on\nits significance.\nResponse: We thank the reviewer for this observation. W3 is indeed not observed in any of the\nproduct complex structures. This likely reflects a genuine loss of that water molecule following\nhydroxylation rather than a refinement artefact. As W3 is part of the hydrogen-bonding network\nobserved in the substrate complexes, its absence in the product structures is consistent with local\nrearrangements of the active site after turnover. Beyond this, at present we cannot assign a specific\nmechanistic role to W3. Since the text already notes its absence, we have not added further\ndiscussion to avoid overinterpretation.\n11) Results, microcrystallography. Please explain why serial femtosecond crystallography was used\ninstead of serial synchrotron crystallography. The time delays are more than sufficient to perform\nthese experiments at a synchrotron.\nResponse: We thank the reviewer for this insightful comment and agree that the time delays used in\nour study could, in principle, also be accessed at a synchrotron source. We chose to perform the\nexperiments at an XFEL in part because the diffraction quality obtained under identical conditions\nwas significantly higher at the XFEL than at a synchrotron, resulting in improved data completeness\nand signal-to-noise.\nThe anaerobic data sets were collected using the acoustic droplet ejection (ADE) sample delivery\nsystem available at the LCLS, which at the time of data collection was the only setup that allowed us\nto maintain strict anaerobic conditions during sample delivery. For these reasons, we selected serial\nfemtosecond crystallography, even though the reaction times studied here could also be examined at\na synchrotron.\n12) The discussion on the optimization of the first SFX to the second SFX structure (also depicted in\nfigure 4) is not entirely relevant to the biochemical discussion in this paper and adds to an already\nlong results section. I would suggest removing this from the main text and focusing on the structure\nthat is relevant (optimized) and moving this optimization description to the supplementary\ninformation.\nResponse: We thank the reviewer for this constructive comment. We agree that the detailed\noptimization steps between the two SFX datasets were overly described and have shortened this\nsection accordingly. However, we have retained both SFX structures in the main text because they\nrepresent two independent beam times and demonstrate the reproducibility of our anaerobic\ncrystallography approach. Importantly, the second SFX dataset (PDB: 9HO2) was collected after\nwashing excess Fe(II) from the crystals, which not only improved resolution and data quality but also\nenabled simultaneous X-ray emission spectroscopy (XES) measurements that were not feasible in the\nfirst experiment. For this reason, we now emphasize the second dataset as the primary basis for\ndiscussion, while keeping the first structure (PDB: 9FW0) as validation of experimental\nreproducibility.\n13) Same section, and figure 4. The description of the experiments becomes very confusing between\ndescribing a 1.5s O2 exposure vs a multi-hour air exposure. The section needs a sentence that\ndescribes the experimental pipeline – samples exposed to O2 in a chamber as well as samples left in\nair for 2-24h. It is unclear to me why choose an O2 chamber rather than just air exposure for the first\nexperiment.\nResponse: We thank the reviewer for pointing out that the description of the 1.5 s O exposure and\n2\nmulti-hour air exposure experiments was unclear. We have revised the beginning of this section to\nexplicitly describe the two complementary experimental pipelines — rapid, controlled O exposure at\n2\nemployed to capture early single-turnover events not accessible by bulk air exposure. We also added\nphrasing (“longer equilibration time points”) to indicate that the 2–24 h datasets represent later\nstages of the same catalytic process. These changes clarify the workflow and rationale for each\nexperimental condition.\n14) It was unclear to me why the authors are comparing the O2 and air exposures by difference\nmaps (not fully explained in the main text). The figure legend has to be highly simplified by\nagglomerating the repeated protocols to make the figures and instead pointing at what the maps are\nshowing. It is unclear from the text what conclusions from these different exposures bring.\nResponse: We thank the reviewer for this helpful suggestion. We have clarified in the main text that\nthe isomorphous difference maps were used to visualize local structural changes at the AspH active\nsite during catalytic turnover, highlighting loss of coordinated waters, substrate hydroxylation, and\n2OG-to-succinate conversion. The figure legend was simplified to focus on the key structural and\nspectroscopic findings rather than repeating experimental protocols.\n15) Same section, last paragraph – “w2/W3 binding may vary in a context dependent manner” –\nplease define what is meant by this in biological terms to explain why this is relevant.\nResponse: By stating that “W2/W3 binding may vary in a context-dependent manner,” we refer to\nthe observation that the positions and occupancies of the Fe-coordinating water molecules (W2 and\nW3) differ depending on the catalytic state and local ligand environment of AspH. In biological terms,\nthis implies that these solvent ligands are dynamic and can be displaced or replaced by substrates,\nintermediates, or products during the reaction cycle. We have clarified the original text in the revised\nmanuscript.\n16) Same section, 2OG and succinate in structures – also confusing. The succinate should be the\nproduct of the turnover (according to figure 1). So why is it only at 50% occupancy with the short O2\nexposure (1.5 seconds), whereas the product is modelled at 100% occupancy but then full occupancy\nof succinate is visible after 2 hours? And why does it drop after 24h? Is it substituted by 2OG? Or just\nunbound? From my interpretation of the results shown, the structures do not really match a\ncoherent biochemical pathway of binding, turnover, unbinding of co-product and rebinding of\nsubstrate.\nResponse: We thank the reviewer for this comment. As outlined in our response to comment 8, the\nmixed 2OG/succinate occupancy observed after 1.5 s O exposure reflects partial re-binding of 2OG\n2\nfollowing a single catalytic turnover, consistent with the excess of 2OG present during crystallisation.\nAt longer equilibration times, only succinate is observed, confirming complete conversion of residual\n2OG, while the reduced succinate occupancy after 12–24 h likely reflects gradual dissociation of\nsuccinate and/or Fe from the active site. Together, these occupancies represent sequential stages of\nsubstrate hydroxylation, co-product formation, and eventual release within the crystal lattice.\n17) In the discussion, it is alluded that AspH is overexpressed in cancers, but based on the\nexplanation of its function, I was unclear as to how this can impact cancer pathogenesis. A small\nexpansion on the topic would help set the stage for drug discovery\nResponse: We thank the reviewer for this valid suggestion; in hindsight, we did not summarize the\nrelationship of AspH with cancer in sufficient detail. We have elaborated on this point in the revised\nmanuscript. Note, however, that the mechanism how AspH and/or AspH catalysis affect cancer\npathogenesis is not well defined.\n18) The authors allude to future experiments using time-resolved crystallography. I agree that such\nexperiments are exciting, but such onerous experiments should be justified with a clear statement of\nwhat information is missing from the current studies. What would TR-SFX bring beyond this already\nthorough structural study? How would this impact our understanding of AspH function and\nmedicinal chemistry developments? What are the outstanding questions?\nResponse: We thank the reviewer for this thoughtful comment. A clarifying sentence has been added\nto justify the need for time-resolved SFX. The current study defines substrate, product, and resting\nstates of AspH but does not capture the short-lived Fe–O or Fe(IV)=O intermediates that mediate\n2\nhydroxylation. TR-SFX will allow direct observation of these transient states, providing missing\nmechanistic insight into O activation and informing future inhibitor design targeting specific\n2\ncatalytic intermediates.\nMinor concerns, for readability\n1) Through the text, the use of the full complex composition for the structures made it harder to\nfollow, especially since for some of the comparisons the change was from the Asp to the Asn\nsubstrate (one letter in a long structure acronym). I would urge the authors to consider a change in\nhow these complexes are referred to in the results section, maybe reducing the number of times the\nfull acronyms are used and simplifying the sentences to highlight the differences between the\nstructures being compared.\nResponse: We thank the reviewer for this helpful suggestion. To improve readability, we have\nsimplified the naming of AspH complexes throughout the Results section by reducing repetition of full\ncomplex compositions (e.g., AspH:Fe:2OG:hFX-Asp) and instead referring to them in a more concise\nand consistent manner (e.g., hFX-Asp complex) where context permits. These changes improve\nclarity and help the reader focus on the structural differences being discussed.\n2) The results section has an enormous amount of detail on the sometimes very small changes\nobserved in the structures that distracts from the main changes that drive the mechanistic\nconclusions. I would advise reducing information that can be seen in figures (for example, adding\nhydrogen bond lengths to the figures rather than the main text) as well as removing information\nsuch as overall RMSD values and repeated notes of the similarity of the overall fold of the protein.\nAll these structures are from the same protein, so the overall fold is expected to be the same, and\nthis information does not add to the discussion. A single sentence in the conclusion that mentions\nthis similarity would be sufficient.\nResponse: We thank the reviewer for this comment - we have worked to simplify the text wherever\npossible. We appreciate the concern regarding the level of detail in the Results section; however, we\nbelieve that, at least in some cases, retaining detailed structural descriptions, including hydrogen-\nbonding distances and RMSD values, is important for completeness and transparency. These details\nsupport the mechanistic conclusions and allow readers to independently assess subtle but\nfunctionally relevant differences among structures. For this reason, we have chosen to maintain the\ncurrent level of detail in the main text wherever necessary.\n3) Results, Mass spectrometry section – similar concerns as in comment 2. All the mass values are\nstated in the text. A table would be better suited, with the text mentioning the species identified,\nrather than the masses observed. The figure legend for 5 should also be simplified to avoid the\nrepetition, and the figure may even be more suitable as supplementary. The masses are visible in the\nmain figure, charge states can be added to the figure.\nResponse: We thank the reviewer for this helpful suggestion. We agree that the mass spectrometry\ndata presentation could have been simplified, and we have accordingly streamlined the Figure 5\nlegend to avoid repetition. We have also added the assay conditions in the figure legend which were\nmissing. However, we have retained the numerical mass values in the main text to ensure\ncompleteness and facilitate comparison with previously reported AspH mass spectrometry data. We\nbelieve this format provides clarity while preserving transparency of the experimental results.\n4) Discussion, paragraph 1 – please correct Figure X.\nResponse: Thank you - we have corrected this error.\n5) The authors should ensure that relevant figure sub-panels are mentioned in the text, rather than\nonly the figure number.\nResponse: We have carefully addressed this in the revised manuscript.\n6) The authors should also try to minimize sentence length, with some sentences being very long\nand complex.\nResponse: We have carefully increased readability and flow of the text by simplifying sentences\nwherever possible.\n7) Figure 2 – the chemical drawings are too small and barely readable\nResponse: We thank the reviewer for this comment. We have increased the size of the ChemDraw for\neasier readability in the revised manuscript.\n8) Conclusions, microcrystal optimization. The first paragraph should probably be moved to the\ndiscussion section.\nResponse: We thank the reviewer for this comment. We have revised the section and have moved\nparts of the text to the discussion to increase the overall readability and accessibility for a broad\nscientific audience.\n9) Please add to the SI a figure showing the two setups at MFX and PAL\nResponse: The two sets ups are already described in the literature for the tape drive setup at LCLS (F.\nD. Fuller, Nature Methods 2017, 14, 443–449) and for the fixed target setup at Diamond Light Source\n(Horrell, S., et al. J. Vis. Exp. 2021, 168, e62200) and PAL-XFEL (D. Lee, et al. Scientific Reports 2019,\n9, 6971) - since these were not directly part of our work we think citations are most appropriate, but\nwe are happy to take editorial advice on this issue.\nReviewer #4:\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part\nof the Nature Communications initiative to facilitate training in peer review and to provide\nappropriate recognition for Early Career Researchers who co-review manuscripts.\nWe thank this reviewer for valuable contributions.\nPOINT-TO-POINT RESPONSE TO THE REVIEWERS' COMMENTS:\nResponse to reviewer 3:\nComment: As a continued discussion of comment 8, Please can the authors indicate how the single\nturnover with retention of the large substrate but exchange with excess mobile substrate has been\nclarified in the text? The explanation given is a good hypothesis on why these substrates are visible in\n50% occupancy, but the second explanation – the full conversion of 2OG upon extended O2 exposure\nis still not clear to me – as a non-expert in these specific enzymes. If the protein can only turnover\nonce, then why is 2OG further converted over time? Does 2OG easily oxidize in solution when not\nunder inert conditions? If so, this needs to be clarified in the text for the non-expert audiences that\nread Nature Communications.\nResponse:\nWe thank the reviewer for this good point and for highlighting where additional clarification was\nneeded for a non-expert audience.\nRegarding single turnover in crystallo versus exchange of small cosubstrates: the statement that AspH\nundergoes a single turnover in crystallo refers specifically to the large 39 residue EGFD peptide\nsubstrate. After hydroxylation this peptide product remains bound and cannot diffuse out of the\ncrystal lattice because it is constrained by lattice contacts and interactions with both the oxygenase\nand TPR domains. By contrast, the small cosubstrate 2OG and succinate are mobile and can readily\nexchange through solvent channels within the crystal. A single turnover leads to 'trapped'\nhydroxylated-EGFD and succinate. In some active sites succinate can leave enabling rebinding of 2OG\nfrom the surrounding mother liquor, while others retain succinate, leading to the mixed\n2OG/succinate occupancies observed in the product structures.\nRegarding the apparent complete conversion of 2OG at longer O₂ exposure times: this does not\ncontradict the single turnover observed within the crystals. In addition to enzyme molecules\nincorporated into the crystal lattice, there is always a population of AspH present in the surrounding\nsolution due to residual soluble protein and partial resolvation of crystal surfaces. This soluble enzyme\ncan continue to turnover in solution over extended exposure times, progressively consuming free 2OG\nand generating succinate. As a result, at later time points the solution becomes depleted in 2OG and\nenriched in succinate, and (only) succinate is available to diffuse into and occupy the cosubstrate\nbinding site within the crystals.\nWe now explicitly state this in the Results section of the revised manuscript. Note that we do not\ninvoke spontaneous oxidation of 2OG in solution; its depletion is likely due to enzyme mediated\nturnover outside the crystal lattice.\nComment:\nThe explanation of the adjustment of the occupancies and restraints of 2OG and succinate used needs\nfurther clarification. Why do the authors assume that the combined final occupancy should be 1 –\nespecially if there is a claim that succinate gently unbinds over time? When determining sub-total\noccupancies, what was the criteria that lead to the decision of 0.5 for each? Best refinement statistics?\nBest b-factor correlation to the surrounding residues? Did the authors compare their protocols to an\nautomated method, such as that used in phenix?\nResponse:\nTo clarify the refinement procedure in the expanded Methods section. Briefly, 2OG and succinate were\nmodelled as alternative ligands occupying the same cosubstrate binding site, and their occupancies\nwere refined reciprocally with the combined occupancy constrained to 1.0. This reflects exclusive\nbinding of a single small molecule at that site rather than simultaneous partial occupancy by two\nligands. Models with combined occupancies significantly below 1.0 resulted in positive residual\nelectron density, indicating incomplete modelling of ligand density at the site.\nThe final approximately 0.5 / 0.5 occupancies were selected based on agreement with minimal\ndifference features in the electron density, ligand and neighbouring residue B factors, and overall\nrefinement statistics. Automated occupancy refinement using PHENIX was not required in this case,\nas the reciprocal manual refinement approach yielded a consistent solution that is well supported by\nthe crystallographic data.\nComment:\nRegarding the comment 12\nFor readability and to improve on a complex manuscript that should be geared to general audiences,\nI still advise that the discussion of the first, unoptimized structure be moved to supplementary\ninformation. A couple of sentences highlighting the considerations that lead to the high resolution\nstructure and XES measurements (i.e. the washing of the crystals) should be mentioned in the main\ntext, but this structure should not crowd a figure as it does not add any mechanistic insights.\nResponse:\nWe thank the reviewer for the suggestion. Since the initial submission, we have substantially\nshortened the discussion of the first SFX dataset and moved extensive optimisation details to the\nSupplementary Information. In its current form, we believe the remaining text is concise and does not\ndistract from the biochemical narrative.\nWe respectfully disagree that the remaining description should be moved entirely to the\nSupplementary Information. The brief discussion of the initial SFX structure provides important\ncontext for the subsequent optimisation, in particular the rationale for washing the microcrystals to\nremove excess Fe(II), which was critical for enabling higher-quality data and the combined time-\nresolved crystallography and X-ray emission spectroscopy experiments. We therefore believe it\nremains appropriate to retain this material in the main text and have not made further changes.\nComment:\nRegarding comment 16 – thank you for clarifying the interpretation and the catalytic steps. Can the\nauthors specify where in the main text this was clearly stated? A supplementary figure showing this\nexperimental-condition-driven mechanism would be very helpful for the unambiguous and easy\ninterpretation of the results described. Do the authors have a measurement for the affinity of\nsuccinate to the enzyme-product complex? This would substantiate the final hypothesis that there is\nslow unbinding over time.\nResponse:\nWe thank the reviewer for this further request for clarification and agree additional explanation was\nrequired; the requested changes have now been made, as already partially outlined in our response\nto comment 8.\nIn the revised manuscript, this interpretation is now stated explicitly in the main text for both the\ncryogenic and room-temperature datasets. We clarify that the mixed 2OG/succinate occupancy\nreflects partial re-binding of excess 2OG following a single catalytic turnover, whereas at longer\nexposure times the surrounding solution becomes depleted in 2OG and enriched in succinate due to\nsolution-phase turnover by enzyme molecules outside the crystal lattice. We further state that the\nreduced succinate occupancy observed after 12–24 h reflects gradual dissociation of succinate and/or\nFe from the active site, rather than re-binding of 2OG. These additions are intended to make the\nsequence of events unambiguous for non-expert readers.\nWe agree that a schematic summary aids interpretation and have therefore added a supplementary\nfigure 19 illustrating the experimentally driven progression from the substrate complex to mixed\n2OG/succinate occupancy, followed by exclusive succinate binding and eventual partial Fe dissociation\nat extended time points.\nRegarding succinate affinity measurements, such experiments are not currently feasible. At present,\nchemical synthesis of the hydroxylated 39-residue EGFD product is not possible, and isolation of\nsufficient quantities of the hydroxylated product following enzymatic turnover is likewise impractical,\nas the amounts required for quantitative binding studies would be prohibitively large and resource-\nintensive. Moreover, based on the induced-fit binding mode observed in crystallo, the hydroxylated\nproduct is expected to have low intrinsic affinity for AspH once released, making formation of a stable\nenzyme–product complex in solution unlikely. As such, quantitative measurements of succinate\nbinding to the enzyme–product complex would be experimentally challenging and fall outside the\nscope of the present study.\nWe believe that the clarified text and the added supplementary schematic sufficiently address this\npoint."}]

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