A disulfide redox switch mechanism regulates glycoside hydrolase function

Domain: chemistry
Published: 2026-01-05
Editorial outcome: minor_revision

Abstract
Disulfide bonds are a key post-translational modification involved in protein folding, structural stability, and functional regulation. Here, we demonstrate that a glycoside hydrolase from the GH2 family undergoes reversible redox regulation through an intramolecular disulfide bond. The enzyme is inactive in its oxidized state and becomes active when reduced through a fully reversible process. Under oxidative conditions, multiple crystallographic and cryo-EM structures revealed a pronounced structural disorder in the active site, most prominent in the regulatory and catalytic loops, which disrupts the substrate binding site and, remarkably, the configuration of the acidic catalytic residues. Conversely, a high-resolution cryo-EM structure of the active (reduced) state unveiled a well-ordered active site with catalytic residues properly positioned for a classical Koshland retaining mechanism. This reversible order-disorder process based on a disulfide switch provides a mechanism for redox-dependent control of glycoside hydrolase activity, with potential implications for carbohydrate metabolism, microbial adaptation and biotechnological applications.

Introduction
Proteins are remarkably sensitive to environmental modification, including variations in pH, temperature, ionic strength, and oxidative stress. Among these factors, the unique reactivity of sulfhydryl groups in cysteine residues allows reversible formation and disruption of disulfide bonds, in response to the redox environment1. This dynamic behavior underpins critical metabolic processes in biological systems, such as homeostasis2,3, gene expression4 and cellular signaling5.

While the importance of cysteine residues in the metabolic regulation of proteins via reversible disulfide bonds is well documented, their role in carbohydrate enzymology is rather limited. To the best of our knowledge, cysteine residues are described to be functionally associated with (i) substrate recognition, observed in β-galactosidases ArthβDG6 and KL-β-Gal7; (ii) coordination of non-catalytic metals, as described for the GH9 family8; (iii) coordination of catalytic metals in the families GH1279 and GH14610, where cysteine acts as a zinc-mediated nucleophile; and (iv) participation in a redox NAD+-dependent catalytic mechanism in the GH4 family, where a reduced cysteine in the second coordination sphere with substrate coordinate a divalent metal that interacts with both NAD+ and substrate11.

Cysteines in Carbohydrate-Active enZymes (CAZymes) also form disulfide bonds that play a critical role in structural stabilization, including maintaining protein folding and enhancing thermostability. This is exemplified by cellulases from the families GH512, GH1213, and GH4514, a GH11 xylanase15, and lysozymes from the families GH2216 and GH2417. Despite their extensive occurrence, the regulatory potential of disulfide bonds in CAZymes remains undescribed. To date, no studies have addressed whether reversible disulfide bond dynamics contribute to redox-sensitive regulation in these enzymes, leaving a gap in understanding how CAZymes might adapt to oxidative environments and the molecular mechanisms enabling such adaptation.

In this work, by employing an integrative approach including synchrotron X-ray crystallography, single-particle cryo-EM, molecular dynamics simulations, site-directed mutagenesis, enzyme kinetics and mass spectrometry, we uncover a distant GH2 member, whose activity is reversibly regulated by a redox switch involving an intramolecular disulfide bond. Our findings demonstrate that this glycoside hydrolase undergoes reversible disulfide bond formation between two cysteine residues located in the catalytic interface. The oxidized enzyme exhibits a conformational disorder in the active site (regulatory and catalytic loops) that impair enzymatic activity, while reduction promotes active site reorganization, leading to maximum catalytic function. Acting as a regulatory switch, this reversible disulfide bond enables the enzyme to respond dynamically to changes in oxidative environments while simultaneously offering protection against irreversible oxidative damage. These findings expand our current understanding of redox-mediated regulation via disulfide bonds in CAZymes, providing opportunities to explore their functional roles in cellular processes and under oxidative stress conditions.

Results
A redox-regulated glycoside hydrolase
The exploration of the genetic landscape from microbial life specialized in plant biomass deconstruction from Brazilian biodiversity has been demonstrated to be an untapped source of unique enzymatic systems, including several recently discovered CAZy families, such as AA1818, GH17319, CBM8919, and CE2020.

Mining metagenome-assembled genomes (MAGs) recovered from Brazilian biodiversity, we discovered an uncharacterized bacterium, named ‘Candidatus Capybacter oxireducens’ that harbors a distant GH2 member (CapGH2b) that co-localizes with mannan-active enzymes belonging to GH173 and GH5_57 families (Fig. 1a). Sequence similarity analysis (SSN) revealed that this member along with other GH2 sequences, predominantly from gut microbiota Bacteroidales, establishes an uncharacterized cluster (UC) within this family (Fig. 1b, Supplementary Fig. 1). According to the recent classification proposed by the CAZy team, this cluster, based on sequence similarity, would belong to the GH2_13 subfamily21. To gain insights into the function of this UC, we produced and performed the biochemical characterization of recombinant CapGH2b, confirming its β-mannosidase activity (Supplementary Fig. 2), which supports the genomic context along with other mannan-active enzymes.

To further evaluate the enzyme responsiveness to ligands, assays were conducted in the presence of monosaccharides, metals, and reducing agents. As expected, mannose had a negative impact on enzyme activity as well as other monosaccharides and distinct metals, including Co2+, Ni2+, Cu2+, Zn2+, and Fe2+ (Supplementary Fig. 2). The presence of EDTA had no effect, indicating that CapGH2b is not a metal-dependent enzyme. Interestingly, an increase in the catalytic activity was observed upon incubation with DTT, suggesting that the enzyme would be responsive to reducing agents (Fig. 1c).

To further explore this unusual redox behavior, a mild oxidizing condition was employed, utilizing atmospheric oxygen during long-term incubation, which led to an enzymatic activity decrease of 90% (Fig. 1c). Subsequent treatment of the oxidized enzyme with varying DTT concentration and incubation times restored the enzymatic activity, and even enhanced compared to its initial levels (Fig. 1d). All tested DTT concentrations resulted in similar maximum activity values (~100 U g–1), indicating saturation of catalytic activity (Fig. 1d). The recovery of activity with DTT treatment supports a putative reversible redox-dependent regulatory mechanism.

To investigate oxidation over a shorter timescale, the enzyme was treated with the oxidizing agent H2O2, followed by incubation with DTT as a reducing agent. The enzyme demonstrated a behavior consistent with long-term atmospheric oxidation, confirming that oxidation can be chemically induced and subsequently reversed by reducing treatments (Fig. 1e–h, Supplementary Figs. 3, 4). However, as H2O2 concentrations increase, the activity recovery decreases, suggesting that higher levels of H2O2 result in irreversible and non-specific oxidation processes, impairing the full enzymatic activity reestablishment.

After enzyme treatment with H2O2 or DTT, thermostability assays were performed to evaluate whether oxidation and reducing processes can influence structural stability (Supplementary Fig. 5). No remarkable difference in thermostability was observed in the enzyme incubation with H2O2 or DTT, indicating that the redox mechanism does not influence the enzyme thermostability but appears to specifically regulate enzymatic activity.

These findings underscore an unprecedented redox regulatory mechanism for carbohydrate breakdown, in which oxidation impairs activity, whereas reduction effectively restores enzymatic activity, making it a fully reversible process.

The inactive conformation exhibits a disordered active site
To elucidate the structural basis for this redox regulation, we determined the enzyme structure using X-ray crystallography via the single-wavelength anomalous dispersion (SAD) method (Supplementary Table 1). The CapGH2b structure retains the conserved modular architecture typical of GH2 enzymes, comprising five domains (Supplementary Fig. 6). However, the domain 5 in CapGH2b lacks the β-sandwich fold and is consequently shorter than in other GH2 members (Supplementary Fig. 6). This truncated domain likely influences oligomerization22 and could potentially alter the enzyme assembly properties. Indeed, crystal packing analysis revealed a homotrimeric organization of CapGH2b, further confirmed in solution by SEC-MALS and SAXS (Supplementary Fig. 7), which contrasts with other structurally characterized family members featuring a homodimeric organization, such as BtMan2A22 and DtMan23.

Unexpectedly, the refined structure revealed a massive structural disorder in the catalytic domain, specifically in two major loops of the catalytic interface, named here as catalytic (G462-P482) and regulatory (S509-G536) loops (Fig. 2a). These loops contribute to defining the topology of the active site and contain key residues for substrate anchoring (N464 and W532) and catalytic activity (acid/base E465). However, their intrinsic flexibility precludes the visualization of electron density and, therefore, the active site configuration, indicating that this disordered structure likely corresponds to the inactive conformation.

In an effort to obtain the structured active site, the enzyme was crystallized in different conditions, featuring distinct space groups and molecules content within the asymmetric units (Supplementary Fig. 8, Supplementary Tables 1, 2). However, in all structures, consistently, the active site was disordered, in particular these two loops. Mass spectrometric analysis of crystallized samples revealed the presence of the full-length protein (Supplementary Table 3), discarding the possibility of the absence of electron density is due to internal protein cleavage by endogenous proteolytic activity. It reiterates that these loops are inherently disordered regions in the captured conformational state of the enzyme by X-ray crystallography, as also predicted by FoldIndex24 (Supplementary Fig. 9).

Our extensive attempts to resolve the crystal structure with ordered loops were unsuccessful, suggesting that the observed disorder may be related to the oxidation process. The typical incubation time and temperature used for crystallization likely favor oxidation, leading to its inactive state. This hypothesis is supported by the disordered active site observed in all crystal structures and aligns with biochemical data demonstrating that the oxidized state is catalytically inactive. Therefore, the crystallization process itself appears to hinder the elucidation of the active conformation.

The ordered active conformation
To overcome the technical limitations imposed by the crystallization process to solve the active conformation of the enzyme, we employed high-resolution single-particle cryo-EM. In this approach, fresh protein preparations exhibiting full catalytic activity were rapidly vitrified, ensuring the active conformation of the enzyme. The cryo-EM structure achieved a high global resolution of 2.6Å, which allowed the visualization of side chains throughout the trimeric biological unit (Fig. 2b, Supplementary Figs. 10, 11, and Supplementary Table 4). Remarkably, the two flexible (catalytic and regulatory) loops in all crystallographic structures presented a well-defined conformation in the active state captured by cryo-EM. In this structure, we were able to unambiguously identify key residues for catalysis, including the acid/base E465, the catalytically relevant N464, and the +1 aromatic residue W532 (Fig. 2b). To discard that the crystallographic structures featuring disrupted active site is not a crystallization artifact, we also solved the oxidized structure of CapGH2b by cryo-EM. Similar to crystallographic structures, both catalytic and regulatory loops were not observed, supporting their intrinsic disorder in the oxidized (inactive) state (Fig. 2a, c, Supplementary Figs. 10, 11, Supplementary Table 4).

The active conformation of the enzyme revealed that both the catalytic and regulatory loops are essential for defining the architecture of the active site, principally the +1 subsite (Fig. 2d). Interestingly, CapGH2b contains two glycine residues (G473 and G533 in the catalytic and regulatory loops, respectively), which results in an opened active-site pocket architecture compared to structurally characterized GH2 members (Fig. 2e, Supplementary Fig. 12). In those structures, the corresponding residue to G473 is W465 in DtMan (PDB 5N6U23), W470 in BtMan2A (PDB 2JE822) and W485 in XacMan2A (PDB 6BYE25), which serves as a hydrophobic platform for sugar anchoring in the +1 subsite (Supplementary Fig. 12). The next glycine residue in the regulatory loop (G533) corresponds to H490 in DtMan23, Y537 in BtMan2A22, and G557 in XacMan2A25, which also contribute to the +1 subsite topology. Supporting this, the opened +1 subsite of CapGH2b does not impose a steric barrier for accommodating distinct substrate orientations (Supplementary Fig. 12). This adaptability suggests the capacity of the enzyme to recognize and process branched mannooligosaccharides, similar to the role of the co-localized enzyme, CapGH5_57, in cleaving branched hetero-β-mannans26.

Remarkably, the lack of key aromatic residues in the catalytic loop seems to be compensated by the presence of W532 in the regulatory loop. Ligand docking analysis indicates that this residue serves as a hydrophobic platform for the +1 glycoside and forms a hydrogen bond with O5 from mannose in the –1 subsite (Supplementary Fig. 12). Additionally, the residue R535, also located in the regulatory loop, establishes hydrogen bonds with the +1 sugar moiety (Supplementary Fig. 12), an interaction in this subsite not previously reported for GH2 members. These analyses reveal the extensive structural remodeling of the +1 subsite in CapGH2b, driven by the unique molecular properties of both catalytic and regulatory loops.

Following the atypical molecular pattern observed in the +1 subsite, the –1 subsite also exhibits a previously undescribed configuration, featuring I201 and D670 in positions typically occupied by tryptophan residues in all structurally characterized orthologs (Supplementary Figs. 12, 13). Molecular docking analysis indicates that the loss of polar interactions between mannose and the NH of the corresponding tryptophan, which is replaced by I201, is compensated by the presence of the non-conserved residue R605. Moreover, D670 plays a key role in positioning R605, along with R535, which also forms a hydrogen bond with mannose in the –1 subsite of CapGH2b (Supplementary Figs. 12, 13). These structural adaptations in both the negative and positive subsites of CapGH2b underpin an unprecedented molecular basis for substrate recognition within the GH2 family.

Regarding the catalytic residues in the active conformation, the nucleophile and the acid/base are productively positioned with a distance of 4.9Å (Fig. 2d), similar to that observed in structurally characterized GH2 members, such as DtMan, BtMan2A, XacMan2A, and ThMan2A (PDBs 5N6U23, 2JE822, 6BYE25 and 4CVU27, respectively). Therefore, despite a substantial remodeling of both –1 and +1 subsites in the active conformation, the Koshland retaining catalytic mechanism is conserved.

Intramolecular disulfide bond formation
The determination of both active (reduced) and inactive (oxidized) conformations shed light on a redox order-disorder process controlling the catalytic activity, remaining yet unclear the residues involved in this process. A detailed structural inspection of the active conformation revealed two free cysteines exposed to the solvent, potentially capable of forming a disulfide bond, C427 and C517, both located at the catalytic interface (Supplementary Fig. 14). C427 is in the vicinity of the active site, whereas C517 is located in the intrinsically flexible regulatory loop. In the active conformation, these residues are approximately 18Å apart, with the catalytic loop localized between them, imposing a steric barrier for disulfide bond formation (Fig. 3a). Notably, in the inactive structure this physical barrier imposed by the catalytic loop is not present due to the flexibility observed for catalytic and regulatory loops (Fig. 2a). This massive disorder of the catalytic interface in the oxidized state makes feasible the intramolecular disulfide bond formation between these residues. Further support comes from the observation of a continuous electron density emerging from the cysteine C427 side chain in some crystallographic structures and cryo-EM of the oxidized enzyme, suggesting its potential involvement in the formation of a covalent interaction (Supplementary Fig. 15).

The possibility of an intermolecular disulfide bond in the trimeric arrangement was also verified; however, the distance between C427 in adjacent protomers is ~47Å (Supplementary Fig. 16). The best chance for an intermolecular disulfide bond is between two neighboring C517 residues, which are yet approximately ~24Å apart (Supplementary Fig. 16). However, the comparison of both active and inactive structures did not reveal any structural rearrangements that would favor the C517-C517 disulfide bond formation. Therefore, the most plausible hypothesis in light of the structural data is the formation of an intramolecular disulfide bond between C427 and C517, both of which are located in the catalytic interface.

To confirm this interpretation, we employed targeted proteomics to identify and quantify peptides containing these cysteine residues, in both free and covalently bonded states, using a differential cysteine labeling strategy based on the use of two distinct alkylating agents. Free cysteines were labeled with iodoacetamide (IAA) in non-reduced samples, while covalently bonded cysteines were selectively labeled, after reduction, with N-ethylmaleimide (NEM). In the reduced enzyme control, C427 and C517 were primarily identified in their free state. In contrast, the oxidized enzyme predominantly showed C427 and C517 covalently bonded to NEM with similar quantitative values, indicating disulfide bond formation between these residues (Fig. 3b, Supplementary Figs. 17, 18, Supplementary Table 5). Some cysteine residues remained in the free state after oxidation (Fig. 3b), which correlates with the residual activity observed post-oxidation (Fig. 1h). Additionally, the total proportion of free and oxidized cysteines reached approximately 60%, indicating the presence of irreversible oxidizing modifications in cysteine residues caused by the exogenous oxidizing agent (500µM H2O2). These modifications include the formation of sulfinic (-SO2H) and sulfonic (-SO3H) acid species confirmed by mass spectrometry (Fig. 3c, Supplementary Fig. 19).

The redox switch mechanism
To further validate this model, we next evaluated the role of these cysteine residues in the redox-dependent activity regulation by site-directed mutagenesis. Interestingly, the incubation of the C517A variant with DTT resulted in minimal changes in activity (Supplementary Fig. 20), revealing that the absence of this thiol side chain makes the enzyme unresponsive to redox changes. Furthermore, the activity levels of the C517A variant are similar to the wild-type enzyme, indicating that this residue does not directly contribute to the catalytic process but serves as a regulatory element. To assess the oxidation profile, the variant C517A was subjected to mild oxidizing conditions, as in the WT enzyme. Under mild oxidation, the C517A variant exhibited minimal changes in activity, even after DTT treatment, a distinct behavior compared to the WT enzyme (Fig. 4a, Supplementary Fig. 20). This implies that the C517A variant is “locked” in the active conformation, being insensitive to redox changes.

Docking analysis revealed that the cysteine residue C427 makes critical hydrogen bonds at the –1 subsite with O3 from mannose, NH from N464, and OE1 from the acid/base residue E465 (Supplementary Fig. 21). These interactions play a pivotal role in enzymatic activity involving substrate binding, contributing to the correct acid/base positioning for catalysis, and the interaction with N464 further stabilizes the sugar transition state during hydrolysis22. Consequently, as expected, the C427A variant exhibited an expressive reduction in activity (Supplementary Fig. 21). Despite its low residual activity, the C427A variant still exhibited partial responsiveness to redox changes, suggesting that oxidation of C517 favors a structural shift towards the disordered (inactive) state, which could facilitate disulfide bond formation (Supplementary Fig. 22). In contrast, the C427A/C517A double mutant was completely insensitive to redox treatments, even under high peroxide concentrations (500µM), confirming that redox-dependent modulation of activity is mediated by the residues C427 and C517 (Supplementary Fig. 22).

The use of a strong oxidizing agent (H2O2) had a remarkable reduction in the catalytic activity of the variant C517A, and contrary to WT, DTT treatment failed to restore enzymatic activity (Fig. 4b, Supplementary Figs. 23, 24). This indicates an irreversible process mediated by unspecific oxidation events involving the peptide containing C427, evidenced by its low recovery post-oxidation (Supplementary Fig. 25, Supplementary Table 6). Additionally, a considerable formation of overoxidized C427 species, including sulfinic (-SO2H) and sulfonic acids (-SO3H), was observed 6-fold and 7.5-fold higher, respectively, compared to the WT enzyme (Fig. 4c). This result also highlights a potential shielding effect provided by the intramolecular disulfide bond formation, preventing irreversible oxidative damage of the catalytically relevant cysteine residue. Protective disulfide bonds were previously described2,28, acting as a mechanism to shield proteins from irreversible oxidative damage by enabling a reversible redox process that preserves function under oxidative stress.

The negative impact of the disulfide bond between C427 and C517 on activity was further demonstrated through molecular dynamics simulations of the oxidized state. The intramolecular disulfide bond prevents C427 from interacting with the substrate and induces a prominent steric hindrance for substrate binding (Supplementary Fig. 26). Additionally, the optimal catalytic distances between key residues are disrupted, increasing to over 14Å (Supplementary Figs. 26, 27). These data highlight how protein oxidation alters the structural and functional dynamics of the active site, thereby modulating enzymatic activity.

Compared to C427, C517 is more solvent exposed (31% vs 13%) and has a markedly lower predicted pKa (10.3 vs. 15.1)29, indicating it is more prone to initial oxidation. Therefore, we suggest that deprotonation/oxidation of C517 destabilizes the ordered conformation, favoring regulatory loop flexibility, which in turn contributes to the formation of the disulfide bond between C427 and C517.

The proximity of the regulatory loop to the oligomeric interface further raises the possibility that disulfide bond formation could also influence oligomer stability. To test this, we compared the oligomeric states of WT, C517A, and C427A under oxidative and reductive conditions. The WT enzyme exhibited a concentration-dependent, reversible shift from trimer to monomer upon H2O2 treatment, whereas both variants remained exclusively trimeric (Supplementary Fig. 28). These results indicate that once a disulfide bond is formed, structural disorder is propagated to other regions, including the oligomeric interface. This perturbation of the oligomeric arrangement is supported by changes in electrostatic and hydrogen-bonding interactions near the oligomeric interface upon disulfide bond formation (Supplementary Fig. 29).

Taking together, these findings support a reversible redox-regulated mechanism in which the inherent flexibility of the regulatory loop, which can be further favored by C517 oxidation, facilitates the formation of an intramolecular disulfide bond between C427–C517 locking the enzyme in a disordered and inactive conformation. This mechanism might have implications in controlling the enzyme activity under environmental redox changes and as a protein protective effect against oxidative stress.

Methods
SSN and phylogenetic analyses
SSN and phylogenetic analyses were conducted using a set of CapGH2b homologous retrieved from a local version of the CAZy database, built on NCBI NR (accessed August 2024), and from the UniProt database (accessed January 2025). For local searches, DIAMOND v2.1.9.16345 was employed, retaining sequences with ≥30% identity to CapGH2b and an e-value<1×10-10. BLAST search against UniProt database was additionally used to retrieve the top 250 hits with the same thresholds (≥30% identity; e-value<1×10-10). To complement this dataset, ten sequences containing conserved cysteine residues were manually added, along with 215 functionally characterized GH2 sequences from the CAZy database46. Redundancy was subsequently reduced using CD-HIT v4.8.147 to remove identical entries, yielding a final curated dataset of 335 proteins.

SSN analyses were performed using the web-based Enzyme Function Initiative-Enzyme Similarity Tool48. The initial SSN was constructed by setting the edge calculation parameter as an e-value of 1e-10 and an alignment score of 147. The SSN was analyzed using Cytoscape49. Additional filters to remove edges below 35% identity and clusters with fewer than 16 nodes were applied. The final SSN was represented with an organic layout, and the resulting clusters were colored based on characterized GH2’s enzyme commission (EC), according to the CAZy database.

For phylogenetic inference, a multiple sequence alignment was generated using MAFFT (v7.490)50, with the parameters –genefpair and –maxiterate 1000. After alignment inspection, IQ-TREE2 (v2.2.2.7)51 was employed to reconstruct the phylogenetic tree –B 1000 and –m MFP settings. The resulting tree was visualized in iTOL52, incorporating EC annotations from characterized GH2, and branches with ultrafast bootstrap support below 95 were removed.

Mutagenesis, protein expression, and purification
The sequence encoding CapGH2b cloned into pET-28a(+) vector with the N-terminal 6×HisTag, and containing codon optimized for Escherichia coli expression, was synthesized by GenScript. Site-directed mutagenesis of C427A, C517A, C427A/C517A, E465A and E553Q variants was performed using the QuickChange II XL Site-Directed Mutagenesis Kit (Agilent Technologies). The primer pairs employed for each mutation are detailed in Supplementary Table 7 and were chemically synthesized by Exxtend.

Each vector (WT and variants) was transformed into E. coli BL21(DE3) cells by heat shock and grown in auto-induction medium (1% tryptone and 0.5% yeast extract) containing 2mM magnesium sulfate and kanamycin (100mg mL−1), and 2% of solutions B (25mM disodium hydrogen phosphate, 25mM potassium dihydrogen phosphate, 50mM ammonium chloride, 5mM sodium sulfate) and C (0.5% glycerol, 0.05% glucose, 0.2% lactose) at 37°C, 200rpm for 5h. Then, incubation was continued at 18°C, 180rpm for 16h. The cells were harvested by centrifugation at 4°C, 7500g for 15min, and then resuspended in buffer A (20mM sodium phosphate pH 7.4, 500mM sodium chloride, 5mM imidazole) containing 4mM phenylmethylsulfonyl fluoride (PMSF), 0.1mg mL−1 lysozyme and 0.05mg mL−1 DNAse for lysis by sonication.

The lysed cells were clarified by centrifugation (4°C, 30,000g for 1h) and the supernatant was loaded onto a 5mL HiTrap Chelating HP column (GE Healthcare Biosciences) pre-equilibrated with buffer A, coupled to an ÄKTA system (GE Healthcare Biosciences). A nonlinear gradient of buffer B (20mM sodium phosphate pH 7.4, 500mM sodium chloride, and 500mM imidazole) was used for protein elution. The eluted fractions were analyzed by SDS–PAGE under denaturing conditions53. The selected samples were pooled, concentrated by filtration, and submitted to size-exclusion chromatography (SEC) using a Superdex 200 16/60 column (GE Healthcare Biosciences) pre-equilibrated with 20mM sodium phosphate pH 7.4, 150mM NaCl, at a flow rate of 1mL min−1. CapGH2b concentration was estimated measuring the absorbance at 280nm, using the molecular weight and the extinction coefficient.

Enzyme assays
The CapGH2b activity was screened against polysaccharides and synthetics substrates (ρ-nitrophenyl conjugates) (Supplementary Table 8). The assays were performed using 2mg mL−1 of the enzyme, 150mM McIlvaine buffer pH 5.0, and 0.5% (w/v) of polysaccharides or 5mM of synthetic substrates (at final concentrations) at 40°C for 16h. For polysaccharides, the amount of reducing sugar released was determined using the DNS method (absorbance at 540nm)54, while the hydrolysis synthetic ρ-nitrophenyl substrates (Sigma–Aldrich) was determined by the release of ρ-nitrophenolate ion (absorbance at 410nm) using an Infinite M200 Pro (Tecan) with i-Control v.1.10.4.0 software. Graphs were plotted using OriginPro 2025.

The optimum pH and temperature were determined using 10µg enzyme, 5mM ρnp-β-mannopyranoside, 75mM McIlvaine buffer ranging pH from 2.0 to 8.0, and temperature from 10°C to 70°C, for 30min. Enzyme response to ions was determined using 5mM ions, 75mM MES buffer pH 5.0, 3mM ρnp-β-mannopyranoside, and 20µg CapGH2b at 40°C for 15min. The substrate saturation curve was obtained using the enzyme concentration and reaction time in the linear range and under optimum pH and temperature conditions.

Inhibition assays were conducted using 100mM monosaccharides (mannose, glucose, galactose, arabinose, xylose, and N-acetylglucosamine) in enzymatic reactions containing 75mM McIlvaine buffer pH 5.0, 3mM ρnp-β-mannopyranoside, and 20µg CapGH2b at 40°C for 15min. The enzymatic response to dithiothreitol (DTT) was performed through CapGH2b (WT and C517A, 1mg mL−1, in 20mM sodium phosphate pH 7.5 and 150mM NaCl) incubation with 20mM DTT at 25°C, 300rpm, during 10min. Subsequently, 9.5µg of treated enzyme was used in the enzymatic reaction containing 75mM McIlvaine buffer pH 5.0 and 6mM ρnp-β-mannopyranoside at 40°C during 20min.

For reducing kinetics, CapGH2b samples (WT and C517A) were previously subjected to mild oxidation (at 4°C for 200 days). Oxidized samples (1mg mL−1, in 20mM sodium phosphate pH 7.5 and 150mM NaCl) were incubated with varying DTT concentration and incubation time, at 25°C, 300rpm. Following incubation, 9.5µg of treated enzyme was used in the enzymatic reaction containing 75mM McIlvaine buffer pH 5.0 and 6mM ρnp-β-mannopyranoside at 40°C, 20min.

For peroxide oxidation, CapGH2b samples (WT, C517A, C427A and C427A/C517A 1mg mL−1, in 20mM sodium phosphate pH 7.5 and 150mM NaCl) were incubated with 50, 100, 250, or 500µM of peroxide at 25°C, 300rpm. Aliquots were collected after 24h and/or 48h of incubation, treated with 2µg catalase (Sigma–Aldrich) at 25°C for 5min to remove the peroxide excess, and subsequently incubated with 20mM DTT at 25°C, 300rpm for 10min. Enzymatic activities were assessed both after peroxide treatment and following 20mM DTT or 2mM TCEP incubation using 75mM McIlvaine buffer pH 5.0 containing 6mM ρnp-β-mannopyranoside, and 10µg and 8.5µg of CapGH2b, respectively, at 40°C for 20min.

Biophysical characterization by circular dichroism (CD)
CD experiments were carried out on a JASCO J-815 spectropolarimeter equipped with a Peltier temperature controller (Jasco) to acquire measurements in the far UV (195–260nm) at 20°C, 100nm min−1, with 20 accumulations. The samples of CapGH2b WT and C517A variant (at 1mg mL−1) were previously incubated with 20mM of DTT at 25°C, 300rpm, for 10min or with 50, 100, 250, or 500µM of peroxide at 25°C, 300rpm, for 24h. The DTT excess was removed by filtration. Samples were prepared at a final concentration of 0.3mg mL−1 in 2.4mM sodium phosphate pH 7.4, and 18mM sodium chloride. Thermal denaturation was performed with a heating rate of 1°C min−1 and was monitored at 220nm.

Hydrodynamic and oligomeric properties in solution
Size-exclusion chromatography coupled to multi-angle light scattering (SEC-MALS) experiments were performed using an Optilab refractive index monitor and 8-angle static light scattering detector DAWN 8 (Wyatt Technology). Detectors were coupled to Agilent 1260 Infinity II Isocratic Pump and Vialsampler (Agilent Technology) with a WTC-030N5 analytical size-exclusion column (Wyatt Technology). A total of 100µL of CapGH2b WT, C517A or C427A (1mg mL-1), previously treated under oxidizing or reducing conditions, was loaded onto the column pre-equilibrated with 20mM sodium phosphate pH 7.5 and 150mM NaCl in a flow rate of 0.5mL min−1. SEC column was calibrated using BSA as a molecular weight marker, and data was processed using the ASTRA software 8.1.2 (Wyatt Technology).

Small-angle X-ray scattering (SAXS) measurement was performed at the SAXS1 beamline, LNLS (Campinas, Brazil), using a monochromatic X-ray beam (1.488Å) (Supplementary Table 9). The X-ray scattering was recorded using a Pilatus 300K (Dectris). The sample-to-detector distance was adjusted to a scattering-vector range of 0.01 <q< 0.5Å−1, where q is the magnitude of the q-vector defined by q=4πsinθ/λ, and 2θ is the scattering angle. The CapGH2b sample at 7.6mg mL−1 was prepared in 20mM HEPES pH 7.4 and 150mM sodium chloride buffer. Ten successive images of 30 seconds were recorded and used for calculations after subtracting the sample-buffer scattering. The molecular weight was estimated directly from the SAXS data using SAXSMoW55. The distribution curve P(r) was calculated using the GNOM package56 and was used to estimate the radius of gyration (Rg). The molecular envelope was calculated from the experimental SAXS data using DAMMIN57. The theoretical scattering curve of the crystallographic structure, calculated using CRYSOL58, was fitted into the SAXS molecular envelope using the program SUPCOMB59.

Structure determination by X-ray crystallography
Crystals of CapGH2b wild-type (WT) and variants E465A and E553Q were grown by sitting-drop or hanging-drop vapor-diffusion method at 18°C in drops containing the equivalent volume of protein and crystallization condition, detailed in Supplementary Table 2.

X-ray diffraction data were collected at the W01B-MX2 beamline of the Brazilian Synchrotron Light Laboratory (LNLS, Brazil), the BL9-2 beamline from the Stanford Synchrotron Radiation Light source (Stanford, CA), and at the MANACÁ beamline of Sirius (LNLS, Brazil). Data were indexed, integrated, and scaled using the XDS package60. The crystal structure was solved using the single-wavelength anomalous diffraction (SAD) method. The positions of selenium sites using the anomalous differences were obtained with SHELXD61. Phases were calculated using SHELXE61, and the initial model was built using Buccaneer62. The atomic coordinates were refined with PHENIX.REFINE63, alternating manual modeling and inspection cycles using COOT64. The refined models were validated with MolProbity server65. Processing and structure-refinement statistics are shown in Supplementary Table 1.

Structure determination by cryo-electron microscopy
The fresh (reduced) and long-term incubation (oxidized) samples from SEC were submitted to grids preparation for cryo-EM at 0.25mg mL–1 (in 20mM sodium phosphate pH 7.4, 150mM NaCl). The grids were glow-discharged for 50s with 25mA at 0.37mbar using Pelco EasiGlow glow-discharge and prepared using Vitrobot Mark IV (Thermo Fisher Scientific) operated at 22°C and 100% humidity. 3µL of samples were deposited in Quantifoil R 2/2 mesh 200 (Quantifoil), blotted for 4s, and fast vitrified in liquid ethane. Data collections were performed at the Brazilian Nanotechnology National Laboratory, on a transmission electron microscope Titan Krios G3i (Thermo Fisher Scientific), operating at 300kV with ×96,000 of magnification, which corresponds to a sampling of 0.67Å per pixel. Movies were acquired using the software EPU 2.12.1 (Thermo Fisher Scientific) using a Falcon 3EC direct detector, operating in counting mode with 30 frames per movie, and an accumulated dose of 50 e− Å–2, applying a defocus range from −1.3 to −2.8µm.

A total of 3543 (reduced) and 3792 (oxidized) movies were imported to cryoSPARC v4.3.066 to perform the motion and CTF corrections. The initial round of particle picking using blob picker resulted in 10,117 (reduced) and 107,708 (oxidized) selected and extracted particles (box size 400px, Fourier crop box size 200px), aiming to obtain initial 2D classes, that were followed used for the template picker. The resulting 398,599 (reduced) and 760,934 (oxidized) particles, were selected and extracted (box size 400px, Fourier crop box size 200px) to follow 2D classification, with the blurred 2D classes discarded. The remaining 230,960 (reduced) and 300,249 (oxidized) particles were used for ab initio reconstructions, followed by homogeneous and non-uniform refinements.

Local motion correction and local and global CTF refinements were performed to improve the map quality. The global resolutions reported used the FSC method (cutoff of 0.143), achieving 2.62Å (reduced) and 3.41Å (oxidized). The final map was sharpened using DeepEMhancer67. The trimeric structure of CapGH2b obtained by X-ray crystallography was used as a template for loops reconstruction in COOT64. The models were refined by several series of real-space refinement in Phenix63 and visual inspection and model optimization in COOT64.

Molecular docking and molecular dynamics simulations
Molecular docking was conducted using Autodock Vina 1.1.268. The ρnp-β-mannopyranoside and mannobiose ligands were docked into the active cryo-EM structure of CapGH2b within a grid box measuring 14×14x14Å3 centered at the midpoint of NH groups from residues W199 and W532 (x=183.5Å, y=140.4Å, z=155.2Å). The resulting monomeric structure of the Michaelis complex was replicated in all protomers by structural superimposition.

Simulations of the oxidized state of CapGH2b were carried out considering catalytic and regulatory loops modeled using Rosetta 3.1069, and the intramolecular disulfide bond between C427 and C517 residues of all protomers was built using tleap utility from AmberTools 2070,71. The protonation state of all residues was determined at pH 5.0 using PROPKA 3.029. Additionally, the protonation state of all His residues was evaluated by visual inspection based on their chemical environment. Classical molecular dynamics simulations were performed using the cuda-enabled PMEMD module of Amber 20 molecular dynamics package70,71. The system was prepared by retaining all crystallographic waters, and the trimeric structure was solvated in a water box ensuring a 15Å padding from the protein atoms to the edges of the simulation box (Supplementary Table 10). The total charge of the system was neutralized by adding 24 sodium ions. In all simulations, the FF14SB force field was used to describe the protein, along with the TIP3P model for water molecules and GAFF parameters for ions.

Resulting simulation box was subjected to an initial energy minimization procedure with position restraints potential applied only to the protein structure. The system was further relaxed with an additional minimization procedure without restrictions. Several MD equilibration steps using a 1fs timestep were performed as follows: i) the initial heating stage was performed through four consecutive NVT runs of 50ps each. The first run was conducted at 100K with restraints applied to all atoms except for water molecules and ions. The subsequent three runs were carried out without restraints at 100K, 200K and 313K; ii) the density was equilibrated with a short 200ps simulation at the NpT ensemble, followed by iii) final equilibration with another 200ps simulation at NVT ensemble. Finally, the timestep was increased to 2fs, and productive simulations of 500ns were performed using the SHAKE72 algorithm. All MD data were analyzed by CPPTRAJ software from AmberTools 2070,71 and VMD 1.9.373. The generated molecular trajectories were visualized using VMD 1.9.373 and PyMOL (The PyMOL Molecular Graphics System, Version 3.0 Schrödinger, LLC).

Cysteine labeling
Controls of non-oxidized samples (200µg of WT and C517A variant) were performed by alkylation using iodoacetamide (IAA) or N-ethylmaleimide (NEM). For that, initial reduction and denaturation were performed using 50mM dithiothreitol (DTT) in 25mM ammonium bicarbonate, pH 7.8 (ABC) with 5% (w/v) sodium deoxycholate (SDC) at 60°C for 30min. Reduced samples were transferred to a 0.5mL Amicon filter (30kDa MWCO, Merck Millipore) pre-passivated overnight with 1% (v/v) TWEEN-20, then centrifuged at 13,910g for 20min at 20°C. After discarding the flow through, samples were washed three times with 450µL of 8M urea in 25mM ABC buffer, and once with 25mM ABC.

For IAA alkylation, 450µL of IAA 50mM was added and incubated at 25°C for 50min (in the dark), followed by 5 desalting centrifugation washes with 25mM ABC at 13,910g, 20°C for 5min. Trypsin was then added at a 1:25 w/w ratio for overnight digestion at 37°C. Peptides were collected in the flow through after centrifugation at 13,910g, 20°C for 5min, and dried.

Alkylation with NEM was performed after urea washes. For that, one washing step with 450µL of 100mM phosphate buffer pH 6.0 was performed and then, 450µL of 50mM NEM (in the same buffer) was added for alkylation at 25°C for 2h. The desalting washes, trypsin digestion and peptide collection steps were performed in the same manner as for IAA.

For the differential cysteine labeling, 200µg of WT and C517A variant were oxidized with 500µM peroxide at 25°C, 300rpm, for 24h. Samples were then denatured with 5% (w/v) SDC in 25mM ABC at 60°C for 30min. After, samples were cooled down and IAA added to a final concentration of 50mM, followed by incubation at 25°C for 50min (in the dark). Samples were transferred to a MWCO filter, centrifuged, and 450µL of 50mM DTT was added for incubation at 37°C, 45min. Subsequently, samples were washed 3 times with 8M urea in 25mM ABC buffer. Buffer was exchanged to 100mM phosphate pH 6.0 by centrifugation 13,910g at 20°C for 20min, and then, 450µL of 50mM NEM (in the same buffer) was added for alkylation at 25°C for 2hrs74. The desalting washes, trypsin digestion and peptide collection steps were performed as previously described for the standard curves. Both WT and variant C517A samples were initially and processed in triplicate. Peptides were either diluted or analyzed directly, starting from 150pmol, and quantification values were then expressed as a percentage of 150pmol.

Untargeted and targeted proteomics
Both untargeted and targeted proteomics were performed using a Synapt XS (Waters) coupled to ACQUITY Premier UPLC (Waters). The capillary voltage was set to 2.5kV, the sampling cone to 25V, source temperature to 120°C, desolvation temperature to 500°C, desolvation gas flow to 650L h⁻1, cone gas flow to 50L h⁻1, and nebulizer pressure to 6.5bar. Data acquisition was carried out with MassLynx v.4.2. Mobile phases A and B consisted of 0.1% (v/v) formic acid (A) and acetonitrile with 0.1% (v/v) formic acid (B), respectively. Peptide separation was performed using a peptide CSH column (1mm × 100mm, 1.7µm, 130Å) with a 4–35% gradient over 15min at a flow rate of 50µL min−1. The gradient was followed by a 5min wash with 95% B. Column equilibration with 4% B was carried out during the first and last 5min of the run.

Untargeted ion mobility mass spectrometry with data-independent acquisition mode (HDMSE, Waters) was used (50–2000m/z, 0.5s scan cycle), with a low collision energy of 6eV and a ramped collision energy from 15 to 50eV. Lock mass correction was applied every 30s using the peptide standard leucine-enkephalin (100pg µL−1) within a mass window of 0.5Da.

Data processing was performed in Progenesis QI for Proteomics 4.2 (Nonlinear Dynamics), applying a 1% false discovery rate, 20ppm for MS1 error tolerance, and automatic MS2 error adjustment. Protein identification required at least two unique peptides, each supported by a minimum of three fragment ions. The protein database used for identification included the complete E. coli proteome (downloaded from UniProt on January 22, 2024) supplemented with the sequence of the recombinant protein analyzed in this study. Protein was digested with trypsin, allowing up to two missed cleavages at K/R sites, except when followed by proline. Methionine oxidation was set as a variable modification. For controls labeled with IAA or NEM, carbamidomethylation or NEM modification of cysteine was set as a fixed modification, respectively. In contrast to that, cysteine modification with IAA or NEM was set as variable for oxidized samples differentially labeled. Additionally, searches were performed with sulfinic and sulfonic acids as variable cysteine modifications. Label-free quantification was performed using normalized abundance values calculated in Progenesis, based on the sum of intensities within isotope boundaries.

Targeted proteomics data were acquired in a Synapt XS (Waters) using ion mobility mass spectrometry coupled to multiple reaction monitoring (HD-MRM, Waters), with RADAR (Waters) scanning every 5s. A 0.5s scan cycle was used with low collision energy of 6eV. Parameters of targeted analysis are described in Supplementary Tables 5 and 6. Separate standard curves for IAA or NEM were generated using serial dilutions, starting from 150pmol of peptides. Linear response for control standard curves quantification is demonstrated on Supplementary Fig. 17. Lock mass standard was applied every 30s using the peptide leucine-enkephalin (100pg µL−1). Post-processing of continuous lock mass correction was performed using the accurate mass measurement tool in MassLynx 4.2, applying an average of three scans within a mass window of 0.5Da. Extracted ion chromatograms (20ppm) and peak integration were done on TargetLynx v4.2, with Apex Track tool enabled, peak-to-peak baseline threshold of 4 and smoothing by the mean method with 4 iterations. The raw data for mass spectra are detailed in Supplementary Data.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The authors have accumulated structural (X-ray, cryo-EM) and biochemical evidence that the title enzyme, a glycosidase, contains a disulfide-dithiol redox switch that regulates the enzymatic activity. In the oxidized state, a disulfide bond is formed between two cysteines, C427 and C517, that further induces an order-disorder transition of the active site and neighboring loops. In the reduced state, the active site becomes ordered with both cysteines in the thiol state. Apart from the structural analysis, the authors have used targeted proteomics and mutagenesis studies to identify the cysteines involved and to confirm their involvement in redox regulation. The (essentially reversible) redox regulation in the test tube is definitely there, but questions remain regarding several aspects.\n1.) It would be helpful to pinpoint whether this switch is conserved on the sequence level in close and distant orthologs/homologs.\n2.) What could be the biological function of the redox switch, or, paraphrased, why should the glycosidase enzymatic activity be reduced under oxidative conditions in vivo?\n3.) How many cysteines are encoded per chain, just the two mentioned in the text? It would be helpful to provide a list of all cysteines and the individual solvent accessibility. If there are more than the two cysteines, one should explain why they are not expected to be involved in the redox switching.\n4.) In the mutant studies, the authors have analyzed variant C517A regarding its redox sensitivity, showing that it is redox-insensitive. They do not show the corresponding data for the C427A variant. They state that the mutation leads to a loss of 2/3rd activity, but do not show data regarding redox sensitivity. The enzymatic activity seems sufficient to be still detectable, and it would be interesting to learn how the mutant protein responds.\n5.) At some point, the authors speculate that the switching is entropically driven (Introduction). While it is clear that the system has more degrees of freedom in the oxidized state, the question remains whether entropy is the controlling factor. Formation of a disulfide is enthalpically favorable, so it surely drives the reaction, too. We would suggest stating that \"induces a conformational order-disorder transition ...\"\n6.) We would suggest replacing \"superoxidized\" with \"overoxidized\".\n7.) The authors might wish to mention that redox switching based on reversible order-disorder transitions is a distinct mechanism that differs from the classic order-to-order switch that often includes catalytic cysteines, see e.g.\na) Funk et al. Multiple redox switches of the SARS-CoV-2 main protease in vitro provide opportunities for drug design. Nature Communications, 15(1), 411. (2024)\nb) Reinke, P. Y. et al. SARS-CoV-2 Mpro responds to oxidation by forming disulfide and NOS/SONOS bonds. Nature Communications, 15(1), 3827. (2024)", "reviewer_2": "Martins et al. characterized the structure and function of a novel glycosidase hydrolase. They found that the enzyme activity is redox-controlled. The enzyme has two cysteines that form a disulfide bond in oxidative conditions. The disulfide bond formation induces structural perturbation in the active site region and decreases the enzyme activity. The reduction of the disulfide bond restores the enzyme activity. The role of the disulfide bond in the activity was well-presented in a series of biochemical and labeling experiments. Structural studies indicate that the C517-containing loop is disordered in the oxidized structure and the extended extra densities from C427 indicate the disulfide formation. Discovery of a novel redox-controlled enzyme in the glycosidase hydrolase family would open a new paradigm for regulating carbohydrate metabolism. Following are specific comments.\n1. Biochemical data indicate that C427 is the primary cysteine for oxidation, and C517 protects C427 from overoxidation. In the current manuscript, it is unclear how C517 travels the long distance of 18.2 A to form the disulfide bond with C417. Fig.5 explains that the enzyme oxidation enhances the loop flexibility. The authors need to provide a plausible mechanism by which the oxidation of C427 enhances the flexibility of the distance C517-containing loop.\n2. Supp Fig. 12 shows that oxidized structures have extra density for the disulfide bond. Because there is no density for the C517-containing loop, it is unclear whether the extra densities of C427 are for the disulfide bond with C517. For example, are there oxidized-condition structures that do not have extra density? What about the C517 loop flexibility in those structures? Are there other cysteine residues for potential disulfide bonds?\n3. The authors need to elaborate a persuasive explanation for the significance of redox control in this glycosidase family.", "reviewer_3": "The authors focus on an enzyme (named CapGH2b) belonging to GH2 family found in the mannan utilization locus of Candidatus Capybacter oxireducens, and report its activity and structure. This enzyme was found to have a redox-regulated activity control system, which has not been reported for the well-known CAZymes, and is controlled by the formation and destruction of a disulfide bond between two cysteine residues in the catalytic domain and by structural changes in the loops in which they are located. These were supported by multiple experiments, including biochemical, biophysical, and structural biology techniques.\nAlthough the importance of disulfide bonds in protein function is well known, there has been no molecular-level elucidation of the regulation of activity in CAZymes, and the authors report a very interesting case study not only for CAZymologists but also for researchers in the field of protein science. The mechanism proposed by the authors is well supported by experiments, as the properties of the enzyme under redox conditions were examined in detail. There are no major problems with this manuscript and it is worthy of publication, but there are the following minor concerns.\n1. In Fig. 2 legend (lines 871 and 872), “The electron density maps ... contour level.” refers to which map? The middle diagrams in Fig. 2 (two loops) are not the (crystallographic) electron density maps, but the cryo-EM maps?\n2. Please describe how the conformation and map of the side chain of C427 were in the cryo-EM analysis of the oxidized CapGH2b sample.\n3. Line 218: The ligand used in the docking analysis is a non-natural substrate pNP-beta-mannoside, and the moiety that binds to subsite +1 is not sugar but pNP. Even with mannobiose as a ligand, is it possible that R535 forms a hydrogen bond with the ligand? The authors show the data of crystals soaked with mannose or mannooligosaccharides (Supplementary Table 2). Were no ligands found in these crystal structures?\n4. How was the map of C427 in the crystal structure (9NPB) with TCEP? Does TCEP or any other reducing agent other than DTT restore the enzymatic activity?\n5. Please describe whether the regulatory and catalytic loops as well as two cysteine residues (C427 and C517) are conserved among homologs belonging to the uncharacterized cluster (UC). Is this redox regulatory mechanism expected to be common in gut bacterial homologs?\n6. It seems that no reducing agents such as DTT were used during the enzyme purification, but I wonder if the activity of the enzyme was reduced during the purification period.", "reviewer_4": "The manuscript (Hansen et al.) investigates how disulfide bonds can act as a redox switch to regulate the function of Candidatus Capybacter oxireducens glycoside hydrolases CapGH2b, a group of enzymes involved in breaking down β-mannose carbohydrates. They found that this GH2 family enzyme is inactive in its oxidized state but becomes active when reduced. This transition occurs through a reversible process where the disulfide bond between two cysteine residues (C427 and C517) causes structural changes in the enzyme's active site. In the oxidized state, the active site is disordered, preventing substrate binding and catalysis. When reduced, the active site reorganizes into a well-structured form suitable for catalysis. This order-disorder mechanism provides insights into how GH2 enzymes can adapt to varying oxidative environments, with implications for biotechnology and microbial biology. Overall, the study is well done and well written.\nMajor points:\n1 in Figure 1e-1h, the concentrations of H2O2 in the assay are quite high, even the lowest one 50  M. Do such high concentrations of H2O2 mimic any in vivo situations for the transition between oxidized state and reduced state? In fact, the enzymatic activity of CapGH2b had mild decrease with the presence of 50  M H2O2 (Figure 1e), suggesting that the natural oxidation/redox conditions might have little effect to this enzyme.\nMinor points:\n1 in Figure 1c, what is the “Untreated” condition? What is the concentration of DTT and Mild oxidation in the assay? Please indicate them in the figure.\n4 in Supplementary Figure 17b, mild oxidation treatment indeed causes the activity decrease of C517A mutant (compared to untreated), what is potential reason? It suggests other sites (in addition to C517) might be affected by oxidation. Please also show the data for C427 mutant."}, "author_response": "Subject: Revision and resubmission of manuscript NCOMMS-25-23826-T\nResponse to referees' comments\nReviewer #1 (Remarks to the Author)\nThe authors have accumulated structural (X-ray, cryo-EM) and biochemical evidence that the\ntitle enzyme, a glycosidase, contains a disulfide-dithiol redox switch that regulates the\nenzymatic activity. In the oxidized state, a disulfide bond is formed between two cysteines,\nC427 and C517, that further induces an order-disorder transition of the active site and\nneighboring loops. In the reduced state, the active site becomes ordered with both cysteines in\nthe thiol state. Apart from the structural analysis, the authors have used targeted proteomics\nand mutagenesis studies to identify the cysteines involved and to confirm their involvement in\nredox regulation. The (essentially reversible) redox regulation in the test tube is definitely there,\nbut questions remain regarding several aspects.\n1. It would be helpful to pinpoint whether this switch is conserved on the sequence level in\nclose and distant orthologs/homologs.\nWe thank Reviewers #1 and #3 for raising this important consideration. As shown in the new\nSupplementary Fig. 27, several orthologs sequences conserve both cysteines. These\nsequences come specifically from the genus Alistipes, which belongs to the phylum\nBacteroidota, with examples from species such as A. senegalensis and A. timonensis.\nAccording to a recent classification of the GH2 family proposed by the CAZy team (published\nafter the submission of our article), CapGH2b and its orthologs would belong to the GH2_13\nsubfamily based on sequence similarity1.\nInterestingly, flexibility prediction analysis indicates that these orthologs sequences also\nexhibit intrinsic flexibility in the loop containing the second cysteine2, corresponding to the\nregulatory loop in CapGH2b (new Supplementary Fig. 28). As demonstrated in our work, this\nloop flexibility is essential for the formation of the disulfide bond between the two cysteines.\nTherefore, the conservation of the cysteine pair, as well as the loop flexibility, indicate that this\nregulatory switch mechanism would extend to these orthologs.\nWe have modified the Discussion section to include the text below about these orthologs,\nsupported by the new Supplementary Figs. 27 and 28.\n“This mechanism identified in CapGH2b may extend to its closest orthologs that conserve the\ncysteine pair (Supplementary Fig. 27) and display comparable intrinsic flexibility in the\ncorresponding regulatory loop (Supplementary Fig. 28). These two features are critical for\nredox-responsive disulfide bond formation. Notably, these orthologs are predominantly\nrecovered from gut and fecal microbiota of diverse animal hosts, especially within the genus\nAlistipes (phylum Bacteroidota), suggesting that the redox-regulatory mechanism uncovered\nin CapGH2b might be functionally relevant within gut-associated microbial communities.”\nsequence alignment highlights the full conservation of C427 and partial conservation of C517\namong CapGH2b orthologs. The sequences conserving both cysteine residues belong to the\ngenus Alistipes (phylum Bacteroidota), with examples including MCD8032713.1 from\nAlistipes sp., WP_195498682.1 from A. timonensis, and MCI7308998.1 from A. senegalensis.\northologs. PreFold analysis predicts folded (blue) and intrinsically disordered (red) regions in\northologs containing the conserved cysteine pair. The region corresponding to the regulatory\nloop (outlined by a square) is consistently predicted to be flexible. This intrinsic flexibility is a\nkey prerequisite for the redox switch, suggesting the redox switch mechanism would be\nconserved in these orthologs.\n3\n2. What could be the biological function of the redox switch, or, paraphrased, why should the\nglycosidase enzymatic activity be reduced under oxidative conditions in vivo?\nAs discussed above, CapGH2b and its orthologs originate predominantly from the gut\nmicrobiota of diverse animal hosts, specifically associated with the phylum Bacteroidota. This\nsuggests that the redox-regulatory mechanism identified in CapGH2b might be functionally\nrelevant within gut-associated microbial communities.\nConsistent with an intracellular role, CapGH2b and its orthologs are predicted to be\nperiplasmatic or cytoplasmatic, with no evidence of secretion. This is expected, since\nCapGH2b is not active on polymeric substrates and this kind of fine redox regulation is more\nlikely to occur intracellularly. In either compartment, this reversible disulfide bond might\nrepresent an adaptive mechanism, enabling these bacteria to control carbohydrate metabolism\nin response to fluctuating biotic and abiotic conditions.\nThis redox switch allows the CapGH2b activity modulation in response to oxidative stress or\nchanges in the local redox potential. Under oxidative conditions, the formation of the disulfide\nbond between C427 and C517 remarkably reduces the enzyme activity, lowering the\nmetabolism of mannooligosaccharides to prevent potential metabolic imbalance, an essential\ncontrol under stress. Upon restoration of reducing conditions, enzymatic activity is fully\nrecovered, pointing to a reversible protective mechanism for the catalytic machinery.\nFurthermore, both the cytoplasm and periplasm can undergo transient redox fluctuations\ntriggered by the external environment or metabolic signals3. The redox-sensitive regulation of\nCapGH2b may thus reflect an adaptive strategy in gut microbes to temporally control\ncarbohydrate metabolism, improving resistance in fluctuating redox environments. The\nperiplasm exhibits an inherent redox gradient that affects protein folding and other cellular\nprocesses4. This fact, combined with evidence that multiple carbohydrate metabolism steps\noccur in the periplasm of Bacteroidota, would be an indicative that CapGH2b could function\nin this compartment.\nWe have modified the Discussion section to include the potential biological function of this\nredox switch mechanism, as follows:\n“Furthermore, CapGH2b and its closest orthologs are predicted to be intracellular proteins\n(periplasmic or cytoplasmic)5, consistent with the observation that CapGH2b does not act on\npolymeric substrates. Within the Bacteroidota phylum, multiple steps of carbohydrate\nmetabolism occur in the periplasm6, a compartment characterized by inherent redox gradients7.\nSuch an environment provides a natural context for redox-sensitive regulation, enabling\ndynamic control of enzymatic activity in response to biotic and abiotic stress. Importantly,\nintracellular redox homeostasis is highly dynamic and supported by dedicated systems such as\nthiol-disulfide oxidoreductases (e.g., DsbA/DsbC)4 and glutathione-dependent pathways8,\nwhich catalyze the reversible formation or reduction of disulfide bonds in target proteins. These\nsystems, combined with local H O microgradients, could modulate CapGH2b activity through\n2 2\ncoupled oxidation/reduction cycles that are not fully captured in in vitro experiments.”\n4\n3. How many cysteines are encoded per chain, just the two mentioned in the text? It would be\nhelpful to provide a list of all cysteines and the individual solvent accessibility. If there are\nmore than the two cysteines, one should explain why they are not expected to be involved in\nthe redox switching.\nWe thank Reviewers #1 and #2 for this valuable suggestion. Indeed, providing detailed\ninformation on all cysteine residues and their solvent accessibility is essential for evaluating\nthe potential involvement of additional cysteines in redox regulation.\nCapGH2b contains nine cysteine residues per chain. Among these, only three are solvent-\nexposed in the active conformation: C427 (12% solvent-accessible), C517 (31% solvent-\naccessible), and C792 (15% solvent-accessible). C792 is located at the C-terminus within\ndomain 5 and is not involved in oligomerization. Furthermore, C792 is spatially isolated from\nthe only two other solvent-exposed cysteines (C427 and C517), both within and between\nprotomers. Consequently, there is no compelling evidence to support the hypothesis that C792\nforms a disulfide bond or participates in a redox switch.\nTherefore, only C427 and C517 are both solvent-accessible and spatially positioned in a\nmanner compatible with disulfide bond formation, supporting their role in the redox-sensitive\nregulation observed in CapGH2b.\nTo clarify this point, we have added a new Supplementary Fig. 13 that illustrates the solvent\naccessibility of all cysteine residues in the active conformation structure, and we have modified\nthe text accordingly.\nsolvent-accessible surface area (SASA) of each cysteine was calculated for the active\nconformation, with percentages shown relative to the most accessible residue (E153). Although\nC792 is solvent-exposed, its location at the C-terminus (domain 5) is spatially distant from all\nother solvent-accessible cysteines, precluding its involvement in a disulfide bond. In contrast,\nC427 and C517 are the only two solvent-exposed cysteines positioned at the catalytic interface\nin a manner compatible with disulfide bond formation.\n4. In the mutant studies, the authors have analyzed variant C517A regarding its redox\nsensitivity, showing that it is redox-insensitive. They do not show the corresponding data for\nthe C427A variant. They state that the mutation leads to a loss of 2/3rd activity, but do not\nshow data regarding redox sensitivity. The enzymatic activity seems sufficient to be still\ndetectable, and it would be interesting to learn how the mutant protein responds.\nAs requested, we analyzed the C427A mutant and found that, like the wild-type enzyme, its\nactivity is responsive to redox changes (new Extended Fig. 2). Since this mutant cannot form\na disulfide bond, this result reveals that modification of C517 (via deprotonation or oxidation)\ncoupled with the inherent flexibility of the regulatory loop would work as an initial trigger for\ninhibition, favoring a disordered conformation at the catalytic interface, which consequently\ndiminishes catalytic activity. Furthermore, this equilibrium displacement to a disordered state\nwould facilitate the approximation of C517 to C427, promoting the formation of the disulfide\nbond.\nOnce the disulfide bond is formed, the inactive state is locked, and the local disorder is then\npropagated to other regions of the protein. It also affects oligomerization as demonstrated by\nthe presence of a small but detectable monomeric population in the wild-type enzyme under\noxidizing conditions. Supporting the importance of the disulfide bond for disorder propagation,\nthe oligomeric state of the single mutants (C427A and C517A) is not affected by oxidizing\nconditions (new Supplementary Figs. 25 and 26).\nAs a control to ensure that the observed redox sensitivity is associated with these specific\ncysteine residues, we also evaluated the redox response of the double mutant (C427A/C517A).\nThis variant is completely insensitive to redox treatments, even at high peroxide concentrations\n(500 µM), confirming that the redox-dependent modulation of enzymatic activity depends on\nboth cysteine residues, C427 and C517 (new Extended Fig. 2).\nIn the revised manuscript, we have expanded the Results section to incorporate the redox\nexperiments with the C427A and double mutant, as follows:\n“Despite its low residual activity, the C427A variant still exhibited partial responsiveness to\nredox changes, suggesting that oxidation of C517 favors a structural shift towards the\ndisordered (inactive) state, which could facilitate disulfide bond formation (Extended Fig. 2).\nIn contrast, the C427A/C517A double variant was completely insensitive to redox treatments,\neven under high peroxide concentrations (500 µM), confirming that redox-dependent\nmodulation of activity is mediated by the residues C427 and C517 (Extended Fig. 2).”\na-d H O -induced oxidation of C427A at different concentrations progressively diminishes\n2 2\nenzymatic activity, and subsequent DTT treatment restores activity, a similar behavior of the\nwild-type enzyme. This indicates that C517 deprotonation/oxidation favors catalytic interface\ndisorder, decreasing the enzymatic activity. e-h In contrast, the C427A/C517A double mutant\nshowed no change in activity under oxidation and reduction treatments, demonstrating\ncomplete insensitivity to redox changes in the absence of both cysteines. Results are expressed\nas mean ± standard deviations from three independent experiments (n = 3). The statistical\nsignificances were determined by one-way ANOVA with Tukey’s post hoc test (p < 0.001),\nand bars sharing the same letter in each panel do not present statistically significant difference.\n5. At some point, the authors speculate that the switching is entropically driven (Introduction).\nWhile it is clear that the system has more degrees of freedom in the oxidized state, the question\nremains whether entropy is the controlling factor. Formation of a disulfide is enthalpically\nfavorable, so it surely drives the reaction, too. We would suggest stating that \"induces a\nconformational order-disorder transition ...\"\nWe appreciate the Reviewer insightful comment and agree that disulfide bond formation is not\nsolely entropy-driven. Our data indicate that the inherent mobility of the regulatory loop is\nessential for promoting the disulfide bond, but we acknowledge that enthalpic contributions\nalso play a role. Accordingly, we have replaced the term “entropy driven” with the suggested\none.\n7\n6. We would suggest replacing \"superoxidized\" with \"overoxidized\".\nWe appreciate the nice suggestion. We have replaced the term “superoxidized” with\n“overoxidized” in the manuscript and supplementary material.\n7. The authors might wish to mention that redox switching based on reversible order-disorder\ntransitions is a distinct mechanism that differs from the classic order-to-order switch that often\nincludes catalytic cysteines, see e.g.\na) Funk et al. Multiple redox switches of the SARS-CoV-2 main protease in vitro provide\nopportunities for drug design. Nature Communications, 15(1), 411. (2024);\nb) Reinke, P. Y. et al. SARS-CoV-2 Mpro responds to oxidation by forming disulfide and\nNOS/SONOS bonds. Nature Communications, 15(1), 3827. (2024)\nWe do appreciate the nice suggestion to incorporate the redox mechanism of SARS-CoV Mpro,\nwhich also involves order-to-order structural arrangements related to disulfide bond formation.\nIt certainly strengthens our findings. We have added this example into the Discussion section\nas follows:\n“A similar order-to-disorder transition is observed in the SARS-CoV main protease (Mpro),\nwhere oxidation of the catalytic cysteine C145, located 8 Å from C117, disrupts a β-hairpin\nmotif and drives disulfide bond formation9,10.”\nMartins et al. characterized the structure and function of a novel glycosidase hydrolase. They\nfound that the enzyme activity is redox-controlled. The enzyme has two cysteines that form a\ndisulfide bond in oxidative conditions. The disulfide bond formation induces structural\nperturbation in the active site region and decreases the enzyme activity. The reduction of the\ndisulfide bond restores the enzyme activity. The role of the disulfide bond in the activity was\nwell-presented in a series of biochemical and labeling experiments. Structural studies indicate\nthat the C517-containing loop is disordered in the oxidized structure and the extended extra\ndensities from C427 indicate the disulfide formation. Discovery of a novel redox-controlled\nenzyme in the glycosidase hydrolase family would open a new paradigm for regulating\ncarbohydrate metabolism. Following are specific comments.\n1. Biochemical data indicate that C427 is the primary cysteine for oxidation, and C517 protects\nC427 from overoxidation. In the current manuscript, it is unclear how C517 travels the long\ndistance of 18.2 A to form the disulfide bond with C417. Fig.5 explains that the enzyme\noxidation enhances the loop flexibility. The authors need to provide a plausible mechanism by\nwhich the oxidation of C427 enhances the flexibility of the distance C517-containing loop.\nWe thank the reviewer for raising this question. We acknowledge that our initial description of\nthe steps involving the disulfide bond in Fig. 5 was not sufficiently clear.\nAs shown by flexibility prediction analysis and classical molecular dynamics simulations, the\nloop containing C517 is inherently flexible and considering its length (27 residue-long) it could\nvisit a conformation that enables the disulfide bond when exposed to an oxidative environment.\nHowever, this process involves a substantial order-disorder transition at the catalytic interface,\nwhich is coherent with the fact that this process occurs slowly without the presence of oxidizing\nagents in vitro.\nAlthough disulfide bond formation protects C427 from overoxidation, C517 is more prone to\ndeprotonation and oxidation due to its lower pKa (~10) and higher solvent exposure among all\ncysteine residues in the protein. Therefore, we propose that disulfide bond formation is enabled\nby the inherent flexibility of the regulatory loop and this flexibility can be further favored by\nC517 oxidation/deprotonation, shifting the structural equilibrium to a disordered conformation\nof the catalytic interface.\nFurthermore, new experiments suggested by Reviewer #4 revealed that this disorder is likely\npropagated once the disulfide bond is formed. This is evidenced by a small but detectable\nmonomeric population exclusively observed in the oxidized WT enzyme. In contrast, no\nmonomeric populations were observed in C517A and C427A mutants under oxidizing\nconditions. These results indicate that when the disordered conformation is 'locked' by the\ndisulfide bond, it weakens the homotrimeric arrangement due to the proximity of the regulatory\nloop to the oligomeric interface.\nThe Results section was expanded according to this explanation, and Fig. 5 was modified, as\nfollows:\n9\n“Compared to C427, C517 is more solvent exposed (31% vs 13%) and has a markedly lower\npredicted pKa (10.3 vs. 15.1)11, indicating it is more prone to initial oxidation. Therefore, we\nsuggest that deprotonation/oxidation of C517 destabilizes the ordered conformation, favoring\nregulatory loop flexibility, which in turn contributes to the formation of the disulfide bond\nformation between the C427 and C517.\nThe proximity of the regulatory loop to the oligomeric interface further raises the possibility\nthat disulfide bond formation could also influence oligomer stability. To test this, we compared\nthe oligomeric states of WT, C517A, and C427A under oxidative and reductive conditions.\nThe WT enzyme exhibited a concentration-dependent, reversible shift from trimer to monomer\nupon H O treatment, whereas both mutants remained exclusively trimeric (Supplementary\n2 2\nFig. 25). These results indicate that once disulfide bond is formed, structural disorder is\npropagated to other regions, including the oligomeric interface. This perturbation of the\noligomeric arrangement is supported by changes in electrostatic and hydrogen-bonding\ninteractions near the oligomeric interface upon disulfide bond formation (Supplementary Fig.\n26).\nTaking together, these findings support a reversible redox-regulated mechanism in which the\ninherent flexibility of the regulatory loop, which can be further favored by C517 oxidation,\nfacilitates the formation of an intramolecular disulfide bond between C427–C517 locking the\nenzyme in a disordered and inactive conformation. This mechanism might have implications\nin controlling the enzyme activity under environmental redox changes and as a protein\nprotective effect against oxidative stress.”\nadopts an active conformation, with C427 and C517 positioned 18.2 Å apart, separated by the\ncatalytic loop. This structural arrangement maintains active site integrity, allowing proper\nsubstrate binding and catalysis. Upon oxidation, C527 can be initially deprotonated/oxidized,\nfavoring the intrinsic flexibility of the regulatory loop, defining a disordered intermediate state.\nThis conformational change disrupts the active site architecture, impairing substrate\nrecognition. This favored flexibility facilitates disulfide bond formation between C427 and\nC517, locking the enzyme in the inactive (disordered) state. A second structural snapshot from\nmolecular dynamics simulations (green cartoon) highlights the flexibility of the regulatory and\ncatalytic loops in the oxidized (disordered) state. Subsequent reduction breaks the disulfide\nbond, restoring the initial active state. This reversible redox mechanism enables the enzyme to\nadapt dynamically to fluctuations in the redox environment. The mannose positioned in the –1\nsubsite of the active enzyme was obtained by structural superposition with DtMan (PDB\n5N6U12).\n11\n2. Supp Fig. 12 shows that oxidized structures have extra density for the disulfide bond.\nBecause there is no density for the C517-containing loop, it is unclear whether the extra\ndensities of C427 are for the disulfide bond with C517. For example, are there oxidized-\ncondition structures that do not have extra density? What about the C517 loop flexibility in\nthose structures? Are there other cysteine residues for potential disulfide bonds?\nRegarding the potential for other cysteine residues to form disulfide bonds, please see our\nresponse to the specific question from Reviewer #1 on other solvent-accessible cysteines.\nConcerning loop flexibility, all our solved crystallographic structures correspond to a\ndisordered (inactive) state and lack defined electron density for the regulatory and catalytic\nloops. Therefore, we would not expect to consistently observe extra density connected to C427\nin these structures.\nSimilar to crystallographic structures, the cryo-EM structure of the oxidized sample shows a\ndisordered catalytic interface, with no electron density for both regulatory and catalytic loops\nyet exhibiting an extra electron density near C427 (now included in Supplementary Fig. 14).\nIt is worth noting that this feature is preserved in cryo-EM maps of the oxidized sample without\nimposing symmetry correlations during data processing.\nAlthough the disulfide bond formation locks the enzyme in a disordered state, which\nintrinsically precludes observing a defined electron density, we cannot rule out that some\nsolved structures are a result of at least two distinct oxidized states: (i) a disordered state with\ndisulfide bond and (ii) an intermediate disordered state favored by oxidation (without disulfide\nbond), as indicated by our new results included in this revision.\nWe have updated the supplementary material to include details on all cysteine residues and\ntheir potential for disulfide bond formation, as well as the extra electron density at C427 in the\noxidized cryo-EM structure, as follows:\ndisulfide bond formation. (a-d) Crystallographic structures showing extended electron\ndensity at C427. 2Fo-Fc (blue) and Fo-Fc difference maps (green) are contoured at 1.0σ and\n2.0σ level, respectively. (e-f) Cryo-EM map of the oxidized CapGH2b showing the extended\ndensity at C427, contoured at 0.28. The multiple conformations observed for C427 highlight\nits intrinsic flexibility, which is essential for disulfide bond formation.\n3. The authors need to elaborate a persuasive explanation for the significance of redox control\nin this glycosidase family.\nPlease see our response to question #2 from Reviewer #1, which addresses the biological\nrelevance of this redox switch mechanism.\nThe authors focus on an enzyme (named CapGH2b) belonging to GH2 family found in the\nmannan utilization locus of Candidatus Capybacter oxireducens, and report its activity and\nstructure. This enzyme was found to have a redox-regulated activity control system, which has\nnot been reported for the well-known CAZymes, and is controlled by the formation and\ndestruction of a disulfide bond between two cysteine residues in the catalytic domain and by\nstructural changes in the loops in which they are located. These were supported by multiple\nexperiments, including biochemical, biophysical, and structural biology techniques.\nAlthough the importance of disulfide bonds in protein function is well known, there has been\nno molecular-level elucidation of the regulation of activity in CAZymes, and the authors report\na very interesting case study not only for CAZymologists but also for researchers in the field\nof protein science. The mechanism proposed by the authors is well supported by experiments,\nas the properties of the enzyme under redox conditions were examined in detail. There are no\nmajor problems with this manuscript and it is worthy of publication, but there are the following\nminor concerns.\n1. In Fig. 2 legend (lines 871 and 872), “The electron density maps ... contour level.” refers to\nwhich map? The middle diagrams in Fig. 2 (two loops) are not the (crystallographic) electron\ndensity maps, but the cryo-EM maps?\nWe thank the Reviewer for pointing out this inconsistency. The electron density maps shown\nin the middle panel of Fig. 2a correspond to crystallographic data, while those in Fig. 2b are\nfrom cryo-EM reconstructions. To address this issue, we have revised the legend of Fig. 2a to\nclarify that the maps shown in the middle panel are from crystallographic data, as follows:\n“Fig. 2. Structural order/disorder transitions in the active site drive active/inactive\nconformations. a Structural disorder of the catalytic and regulatory loops in the active site\nobserved in oxidized samples via crystallography and cryo-EM. The left panel shows the cryo-\nEM map (deepEMhancer, contoured at a level of 0.11), while the middle panel shows the\ncorresponding crystallographic electron density maps (contoured at 1.0σ). b Ordered\nconformation of the loops, captured by cryo-EM of fresh samples (predominantly in the\nreduced form), with the deepEMhancer map represented at a 0.20 level.”\n2. Please describe how the conformation and map of the side chain of C427 were in the cryo-\nEM analysis of the oxidized CapGH2b sample.\nPlease see our response to question #2 from Reviewer #2, which addresses the cryo-EM\nstructure of the oxidized enzyme.\n3. Line 218: The ligand used in the docking analysis is a non-natural substrate pNP-beta-\nmannoside, and the moiety that binds to subsite +1 is not sugar but pNP. Even with mannobiose\nas a ligand, is it possible that R535 forms a hydrogen bond with the ligand? The authors show\n14\nthe data of crystals soaked with mannose or mannooligosaccharides (Supplementary Table 2).\nWere no ligands found in these crystal structures?\nWe performed additional docking analysis using mannobiose instead of pNP-β-mannoside.\nThe results revealed that R535 is indeed capable of forming a hydrogen bond with mannose at\nthe -1 and +1 subsites, suggesting a possible role for this residue in substrate recognition and/or\nstabilization during catalysis. These results are now included in the revised manuscript\n(Supplementary Fig. 11), as follows:\nSupplementary Fig. 11. The +1 subsite architecture of other structurally characterized\nGH2 members is narrowed compared to CapGH2b. (a-c) Structural views of the +1 subsite\nin (a) DtMan (PDB 5N6U12), (b) BtMan2A (PDB 2JE813) and (c) XacMan2A (PDB 6BYE14),\nhighlighting their restricted product release area. The mannose positioned in the –1 subsite of\n(a), (b), and (c) was obtained by structural superposition with DtMan (PDB 5N6U). (d) The\nexpanded catalytic interface of CapGH2b accommodates distinct spatial orientations of 4-\nnitrophenol-ring, as assessed by molecular docking. (e) The flexibility of the ligand at the +1\nsubsite is associated with the limited stabilizing interactions with surrounding residues. (f) The\n–1 subsite preserves canonical interactions of GH2 family, while incorporating novel\ninteractions mediated by non-conserved residues unique to CapGH2b, such as D670, R605 and\nR535. (g) Molecular docking with mannobiose reveals further hydrogen bonds at the +1 subsite\ninvolving R535 and Q468.\nRegarding ligands, several crystals were soaked or co-crystallized with different mono- and\noligosaccharides at 100 mM, including mannose, mannobiose, mannotriose, mannopentaose,\nand glucose, as listed in Supplementary Table 2. However, no ligands were detected in the\nresulting structures, likely because all crystallographic structures correspond to a disordered\n(inactive) conformation either locked by disulfide bond formation or favored by oxidation. To\nclarify this point, we have added a corresponding note to Supplementary Table 2, as follows:\n15\n“No ligands were observed, as all solved crystallographic structures captured a disordered\n(inactive) conformation of the enzyme”\n4. How was the map of C427 in the crystal structure (9NPB) with TCEP? Does TCEP or any\nother reducing agent other than DTT restore the enzymatic activity?\nThe 9NPB map displays an undefined electron density continuously connected to C427, which\ncould not be properly modeled. We attribute this to re-oxidation of the sample, likely occurring\nduring the crystallization process and the subsequent waiting period before data collection (as\ncrystals were cryo-cooled immediately prior to data collection). This is evidenced by the local\nstructural disorder observed at the catalytic interface in the resulting structure. We have\nupdated Supplementary Fig. 14 to highlight the extended electron density in the 9NPB map.\nRegarding the role of other reducing agents in modulating this redox switch, we also tested\nTCEP and found that it had an effect similar to that of DTT. These results have been included\nin the new Supplementary Fig. 4, as follows:\nSupplementary Fig. 4. TCEP restores enzyme activity similarly to DTT. Oxidation was\nperformed with 500 µM H O over 24h and the sample was then incubated with 2 mM TCEP\n2 2\nfor (a) 100 min, (b) 180 min and (c) 1190 min, highlighting the enzyme responsiveness to\nTCEP. Results are expressed as mean ± standard deviations from three independent\nexperiments (n = 3). Statistical significance was determined by one-way ANOVA with Tukey’s\npost hoc test (p < 0.001); bars in each panel sharing the same letter are not significantly\ndifferent.\n5. Please describe whether the regulatory and catalytic loops as well as two cysteine residues\n(C427 and C517) are conserved among homologs belonging to the uncharacterized cluster\n(UC). Is this redox regulatory mechanism expected to be common in gut bacterial homologs?\nPlease see our response to question #1 from Reviewer #1, which addresses the conservation of\nthese cysteine residues in CapGH2b orthologs.\n16\n6. It seems that no reducing agents such as DTT were used during the enzyme purification, but\nI wonder if the activity of the enzyme was reduced during the purification period.\nIndeed, no reducing agents were used during purification, and as a result, the enzyme was\npartially oxidized by the end of the process, most likely due to “natural” oxidation events during\ncell lysis and subsequent purification steps. Consistent with our biochemical data, chemical\nreduction of the untreated enzyme (post-purification) resulted in an increase in the activity (Fig.\n1c), reaching its maximum activity level that corresponds to the fully reduced state.\nThe manuscript (Hansen et al.) investigates how disulfide bonds can act as a redox switch to\nregulate the function of Candidatus Capybacter oxireducens glycoside hydrolases CapGH2b, a\ngroup of enzymes involved in breaking down β-mannose carbohydrates. They found that this\nGH2 family enzyme is inactive in its oxidized state but becomes active when reduced. This\ntransition occurs through a reversible process where the disulfide bond between two cysteine\nresidues (C427 and C517) causes structural changes in the enzyme's active site. In the oxidized\nstate, the active site is disordered, preventing substrate binding and catalysis. When reduced,\nthe active site reorganizes into a well-structured form suitable for catalysis. This order-disorder\nmechanism provides insights into how GH2 enzymes can adapt to varying oxidative\nenvironments, with implications for biotechnology and microbial biology. Overall, the study\nis well done and well written.\nMajor points:\n1. In Figure 1e-1h, the concentrations of H2O2 in the assay are quite high, even the lowest one\n50 M. Do such high concentrations of H2O2 mimic any in vivo situations for the transition\nbetween oxidized state and reduced state? In fact, the enzymatic activity of CapGH2b had mild\ndecrease with the presence of 50 M H2O2 (Figure 1e), suggesting that the natural\noxidation/redox conditions might have little effect to this enzyme.\nWhile our in vitro assays establish a clear biochemical mechanism, translating these findings\nto a physiological context involves a deeper understanding of the bacterial intracellular\nenvironment, which differs substantially from in vitro assays.\nA key factor is the subcellular location of the enzyme. The putative cytoplasmic or periplasmic\nlocalization of CapGH2b would place it in compartments with distinct and highly dynamic\nredox potentials. These environments are known to undergo significant fluctuations in response\nto various biotic or abiotic stresses, establishing conditions where the enzyme activity could be\ndirectly and rapidly modulated.\nBeyond the general redox environment, the in vivo context is populated by a suite of dedicated\nredox-active systems, such as the thiol-disulfide oxidoreductases (e.g., the DsbA/DsbC\nsystems) and glutathione-dependent pathways, which specifically evolved to catalyze disulfide\nbond formation and breakdown in target proteins. The presence of these systems could possibly\nbe involved in the regulation of this redox switch.\nFinally, the intrinsic properties of the mechanism itself highlight its potential biological\nrelevance. The demonstrated reversibility is a hallmark of a functional regulatory switch,\nallowing the enzyme to switch between active and inactive states in response to redox changes.\nFurthermore, the role of disulfide bond formation in protecting an essential cysteine residue\nfrom irreversible overoxidation points to a sophisticated protective strategy, preserving the\nenzyme function under oxidative stress.\nof manuscript, as follows:\n“Furthermore, CapGH2b and its closest orthologs are predicted to be intracellular proteins\n(periplasmic or cytoplasmic)5, consistent with the observation that CapGH2b does not act on\npolymeric substrates. Within the Bacteroidota phylum, multiple steps of carbohydrate\nmetabolism occur in the periplasm6, a compartment characterized by inherent redox gradients7.\nSuch an environment provides a natural context for redox-sensitive regulation, enabling\ndynamic control of enzymatic activity in response to biotic and abiotic stress. Notably,\nintracellular redox homeostasis is highly dynamic, involving dedicated systems such as thiol-\ndisulfide oxidoreductases (e.g., DsbA/DsbC)4 and glutathione-dependent pathways8, which\ncatalyze the reversible formation or reduction of disulfide bonds in target proteins. These\nsystems, combined with local H O microgradients, could modulate CapGH2b activity through\n2 2\ncoupled oxidation/reduction cycles that are not fully captured in in vitro experiments.”\n2. Since structural order/disorder transitions in the active site drive active/inactive\nconformation of CapGH2b are the key focus of the current study, one of the major concerns is\nthat the structures of CapGH2b were solved without any substrates. The sufficient amounts of\nsubstrate binding might stabilize the catalytic loop and regulatory loop, in either oxidized or\nreduced conditions. Please test and show the structure of CapGH2b with different substrates.\nWe would like to clarify that we performed extensive crystallization trials over two years, with\na variety of ligands (mannose, mannobiose, mannotriose, mannopentaose, and glucose at 100\nmM; Supplementary Table 2) and employing both co-crystallization and soaking approaches.\nFurthermore, these trials included the wild-type enzyme and its E553Q (nucleophile) and\nE465A (acid/base) variants to maximize the chances of obtaining ligand-bound complexes.\nDespite these efforts, no ligands were detected in the electron density maps from multiple\ndatasets collected at different synchrotron beamlines. Additional trials with other ligands not\nincluded in this manuscript (such as 1,4-β-glucosyl-mannose and mannotetraose) also yielded\nthe same outcome (Table 1). We attribute it to enzyme oxidation during the crystallization\nprocess, which our biochemical data demonstrate is detrimental to substrate binding.\nTable 1. Crystallographic datasets that were not deposited in the Protein Data Bank, including\ncrystallization conditions, protein concentration, ligand incubation and enzyme variant.\nProtein variant Spatial Ligand\nResolution Buffer composition\nand concentration group addition\n1M di-ammonium hydrogen\n50 mM 1,4-β-\nE553Q Variant; phosphate, 0.1M sodium acetate pH\nP1 2.6 Å glucosyl-\n31.6 mg mL-1 4.5, 0.1M praseodyminium (III)\nmannose\nacetate hydrate\n1M di-ammonium hydrogen\nE553Q Variant; 100 mM\nP1 2.5 Å phosphate, 0.1M sodium acetate pH\n31.6 mg mL-1 mannotetraose\n4.5\n19\n1M di-ammonium hydrogen\nE553Q Variant; 100 mM\nP1 2.1 Å phosphate, 0.1M sodium acetate pH\n31.6 mg mL-1 mannose\n4.5\n0.75M di-ammonium hydrogen\nE465A Variant; 100 mM\nP1 2.8 Å phosphate, 0.1M imidazole pH 8,\n31.6 mg mL-1 glucose\n0.2M sodium chloride\nMinor points:\n1. In Figure 1c, what is the “Untreated” condition? What is the concentration of DTT and Mild\noxidation in the assay? Please indicate them in the figure.\nWe would like to clarify that the “untreated” condition refers to the enzyme immediately after\npurification, collected directly from the final size-exclusion chromatography step, without any\nsubsequent oxidizing or reducing treatment. Its incubation with DTT demonstrated that it was\npartially oxidized after purification.\nFor the reducing experiments, we optimized the reaction using 20 mM DTT for 10 min at 25\n°C, as detailed in the “Enzyme assays” within the Methods section. The mild oxidation\ncondition was obtained by long-term incubation of the enzyme with atmospheric oxygen at 4\n°C, also as described in the “Enzyme assays” section.\nTo make this information immediately accessible to the reader, we have now included these\nexperimental details in the legend of Figure 1, as follows:\n“Fig. 1. CapGH2b is catalytically active under reducing conditions and inactive upon\noxidation. (a) Genetic organization of the β-heteromannan Polysaccharide Utilization Locus\n(PUL) from 'Candidatus Capybacter oxireducens', identified in the capybara gut microbiome.\n(b) Sequence similarity network (SSN) analysis of CapGH2b and its closest orthologs from\nCAZy and Uniprot. Functionally characterized enzymes are colored according to their activity\nand organized into clusters. (c) Enzymatic activity assay showing that the fresh enzyme\n(untreated) is active under reducing conditions (20 mM DTT incubation), while mild oxidation\n(long-term incubation at 4 ºC) leads to activity reduction. (d) Reduction saturation curve with\nthe post-mild oxidation enzyme incubated under different DTT concentrations and time points.\nMaximum activity restoration upon DTT treatment confirms reversible redox regulation. (e-f)\nH O -induced oxidation over 24 hours at different concentrations progressively diminishes\n2 2\nenzymatic activity. Subsequent DTT treatment fully restores activity, supporting a redox-\nsensitive regulatory mechanism. Results are expressed as mean ± standard deviations from\nthree independent experiments (n = 3). The statistical significances in (c), (e), (f), (g), and (h)\nwere determined by one-way ANOVA with Tukey’s post hoc test (p < 0.001), and bars sharing\nthe same letter in each panel do not present statistically significant difference.”\n20\n2. What are the oligomeric states of CapGH2b protein in Size-exclusion chromatography in\ndifferent conditions (untreated, DTT treatment, with different concentrations of H2O2)? Is this\nprotein always in trimeric state as illustrated by X-ray and Cryo-EM structures? Please present\nthe data.\nAs requested, we performed SEC-MALS experiments to assess the oligomeric state of\nCapGH2b under different redox conditions. Treatment with H O induced a discrete shift in\n2 2\nthe wild-type enzyme from the predominant trimeric arrangement to a detectable monomeric\npopulation, with the fraction of monomers progressively increasing at higher peroxide\nconcentrations (new Supplementary Fig. 25). Subsequent DTT treatment fully restored the\ntrimeric assembly.\nTo explore the contribution of each cysteine to this event, we analyzed the C517A and C427A\nvariants under the same conditions. In contrast to the WT enzyme, the C517A and C427A\nvariants remained exclusively trimeric upon oxidation and reduction, with no detectable\nmonomer formation (Supplementary Fig. 25).\nAs previously discussed, the initial deprotonation/oxidation of C517 can shift the structural\nequilibrium towards a disordered conformation, which in turn facilitates disulfide bond\nformation between C427 and C517. The disulfide bond formation then allows the disorder\npropagation from the catalytic interface to other parts of the biological unit, including the\nhomotrimeric interface. The structural context supports this interpretation, since the loop\ncontaining C517 (the regulatory loop) lies near this oligomeric interface (Supplementary Fig.\n26).\nWe have included these new results in the revised manuscript, as follows:\n“The proximity of the regulatory loop to the oligomeric interface further raises the possibility\nthat disulfide bond formation could also influence oligomer stability. To test this, we compared\nthe oligomeric states of WT, C517A, and C427A under oxidative and reductive conditions.\nThe WT enzyme exhibited a concentration-dependent, reversible shift from trimer to monomer\nupon H O treatment, whereas both variants remained exclusively trimeric (Supplementary\n2 2\nFig. 25). These results indicate that once disulfide bond is formed, structural disorder is\npropagated to other regions, including the oligomeric interface. This perturbation of the\noligomeric arrangement is supported by changes in electrostatic and hydrogen-bonding\ninteractions near the oligomeric interface upon disulfide bond formation (Supplementary Fig.\n26).”\nOxidation of the WT enzyme with 50, and 250 µM H O resulted in the formation of a minor\n2 2\nfraction of monomers. (b) Subsequent reduction with DTT fully restored the homotrimeric\narrangement. (c-f) In contrast, oxidation or reduction of the C427A and C517A variants did\nnot alter the oligomeric state. These results highlight that the structural disorder observed in\nthe inactive conformation is only propagated to neighboring regions, like the oligomeric\ninterface, upon disulfide bond formation.\nproximity results, upon disulfide bond formation, in a propagation of the local structural\ndisorder to other regions of the biological unit as evidenced by changes in electrostatic and\nhydrogen-bonding interactions near the oligomeric interface.\n3. Can the deep 3D classification in cryo-EM studies of CapGH2b in oxidized find any\nconformer with ordered catalytic loop and regulatory loop?\nWe appreciate the Reviewer thoughtful question. To explore potential conformational\nheterogeneity, we performed 3D classification of the oxidized cryo-EM dataset without\nimposing symmetry, despite the enzyme C3 architecture (Supplementary Fig. 10). This\napproach aimed to capture distinct conformational states of the catalytic and regulatory loops\nacross different protomers. However, we did not observe any reconstructions with well-ordered\ndensity for these loops. It indicates that the predominant molecules on the grid are in the\ndisordered (inactive) conformation, although we cannot rule out that a minor population of\nmolecules adopting the ordered conformation is present.\n4. In Supplementary Figure 17b, mild oxidation treatment indeed causes the activity decrease\nof C517A mutant (compared to untreated), what is potential reason? It suggests other sites\n(in addition to C517) might be affected by oxidation. Please also show the data for C427\nmutant.\nAlthough the mild oxidation of C517A appeared to cause a slight reduction in activity\ncompared to the untreated sample, this effect was not statistically significant. We attribute the\napparent decrease to two factors: (i) the prolonged storage period, which naturally reduces\nprotein stability, and (ii) the limited oxidation of C427, which becomes more solvent exposed\nin the absence of C517. As discussed in the manuscript, C517 protects C427 from\noveroxidation by forming the intramolecular disulfide bond. Under mild oxidation with\natmospheric oxygen, C427 oxidation is limited and thus only slightly impacts activity over\nextended storage.\nRegarding the activity of the C427A mutant, please see the response to question #4 from\nReviewer #1.\nReferences\n1. Lebreton, A. et al. Division of the large and multifunctional glycoside hydrolase family\n2: high functional specificity and biochemical assays in the uncharacterized subfamilies.\nBiotechn. Biofuels and Bioprod. 18, 68 (2025).\n2. Prilusky, J. et al. FoldIndex©: A simple tool to predict whether a given protein sequence\nis intrinsically unfolded. Bioinformatics 21, 3435–3438 (2005).\n23\n3. Neish, A. S. Redox signaling mediated by the gut microbiota. Free Radic. Res. 47, 950–\n957 (2013).\n4. Dutton, R. J., Boyd, D., Berkmen, M. & Beckwith, J. 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Chem. 293, 13636–13649 (2018).\nResponse to Reviewer’s comment\nReviewer #3 comment 1: Are all the characterized β-mannosidases shown in Fig. 1b and\nSupplementary Fig. 1 exo-β-mannosidases belonging to GH2_13, and do they not\ninclude the plant GH2_17 endo-β-mannosidases?\nR1: Fig. 1b includes all characterized GH2 members, including GH2_13 exo-β-\nmannosidases and the plant GH2_17 endo-β-mannosidases. It is worth noting that the\ncharacterized GH2_17 endo-β-mannosidases formed an isolated cluster containing only\nthe three characterized members, which did not meet our cutoff of at least 16 nodes for\nrepresentation in Fig. 1b.\nReviewer #3 comment 2: Is it correct that there are two β-mannosidase (β-man) clusters\nand an uncharacterized (Unc) cluster within GH2_13?\nR2: Yes, it is correct. The CAZy team reports that subfamily-level clustering is defined\nusing closeness centrality rarefaction curves, with subdivisions primarily reflecting\nenzymatic function (Hornung and Terrapon, 2023). As discussed in the referred work,\nfurther subdivision within subfamilies is possible and may capture differences at the\nstructural level in addition to functional ones. In our analysis, we used full-length protein\nsequences rather than only the conserved domain and applied more stringent sequence\nidentity thresholds, which allowed us to resolve finer structural patterns within the\nsubfamily. Consequently, GH2_13 β-mannosidases were separated into distinct clusters,\nwhich reflect structural and/or taxonomic differences. This refined subdivision was\nparticularly relevant for our study, revealing an unprecedented redox order-disorder\nmechanism governing the catalytic activity of a carbohydrate-active enzyme.\nAdditionally, we thank the Reviewer for carefully noting the missing digit in the EC\nnumber for β-mannosidase in Supplementary Fig. 1. We have corrected it to EC 3.2.1.25\nin the revised version.\nReference:\nHornung, B. V. H, Terrapon, N. An objective criterion to evaluate sequence-similarity\nnetworks helps in dividing the protein family sequence space. PLoS Comput. Biol. 19,\ne1010881 (2023)."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have convincingly and carefully addressed all points raised by the reviewers such that acceptance is supported.", "reviewer_2": "All comments of this reviewer were addressed well.", "reviewer_3": "The authors have properly addressed my concerns through the additional experiments and this revision. While a very minor concern has been raised, I believe the authors have done excellent work and the manuscript is generally of publishable quality.\nAccording to the recent subfamily classification by the CAZy team, CapGH2b belongs to GH2_13. Are all the characterized β-mannosidases shown in Fig. 1b and Supplementary Fig. 1 exo-β-mannosidases belonging to GH2_13, and do they not include the plant GH2_17 endo-β-mannosidases? Is it correct that there are two β-mannosidase (β-man) clusters and an uncharacterized (Unc) cluster within GH2_13?\nAdditionally, please correct the EC number for β-mannosidase in Supplementary Fig. 1 to EC 3.2.1.25.", "reviewer_4": "The revised manuscript added some new data to clarify their findings and adequately addressed most of my initial concerns."}, "author_response": ""}]

Decision letter