Evolutionary repurposing of a metabolic thiolase complex enables antibiotic biosynthesis

Domain: neuroscience
Published: 2026-01-30
Editorial outcome: minor_revision

Abstract
The functional diversification of biosynthetic enzymes underlies the chemical richness of natural products, yet how primary metabolic enzymes evolve to acquire specialized functions in secondary metabolism remains elusive. Here, we report a tripartite enzyme complex from oral Streptococcus species—comprising 3-hydroxy-3-methylglutaryl (HMG)-CoA synthase (HMGS), acetyl-CoA acetyltransferase (ACAT), and a DUF35 protein—that catalyzes an unusual Friedel–Crafts C-acetylation on a pyrrolidine-2,4-dione scaffold, completing the biosynthesis of the antibiotic reutericyclin A. Cryo-electron microscopy of the S. macacae-derived thiolase complex (SmaATase) reveals a conserved architecture resembling the archaeal HMGS/ACAT/DUF35 complex involved in the mevalonate pathway, yet with key catalytic residues rewired to reprogram substrate specificity. Biochemical characterization, molecular modeling, and evolutionary analysis confirmed that the ancestral activity of HMG-CoA synthesis has been lost, while the complex has been repurposed to mediate Friedel–Crafts C-acylation of small molecule acceptors. These findings reveal a rare example of thiolase complex neofunctionalization, shedding light on an underexplored trajectory in enzyme evolution and offering a template for engineering C–C bond-forming catalysts in synthetic biology.

Introduction
Natural products, also known as secondary or specialized metabolites, confer selective advantage to their producing organisms by mediating ecological interactions such as competition, defense, and communication1. In contrast to the relatively conserved structures of primary metabolites, natural products display remarkable chemical diversity, largely driven by the evolutionary plasticity of their biosynthetic enzymes2. This diversity often arises through the recruitment and functional repurposing of primary metabolic enzymes into specialized metabolic pathways2,3. Beyond the well-known fatty acid synthase–polyketide synthase (FAS–PKS) paradigm4,5, notable examples include the evolution of chalcone isomerase from a noncatalytic fatty acid-binding protein6,7, and the emergence of β-lactam synthetases from asparagine synthase8,9 (Supplementary Fig. 1). While the catalytic repertoire of biosynthetic enzymes continues to expand, the evolutionary trajectories by which primary metabolic enzymes—especially multimeric enzyme complexes—acquire new functions in secondary metabolism remain poorly understood.

The thiolase superfamily comprises highly conserved enzymes present across all domains of life and plays essential roles in central metabolic pathways10,11. For instance, acetyl-CoA acetyltransferase (ACAT) catalyzes the formation of acetoacetyl-CoA, a key intermediate in cholesterol and steroid hormone biosynthesis, while its reverse reaction is critical for fatty acid β-oxidation12,13. In archaea, a tripartite enzyme complex composed of 3-hydroxy-3-methylglutaryl (HMG)-CoA synthase (HMGS), ACAT, and a DUF35 protein (Pfam family PF01796, Domain of Unknown Function) enables mevalonate biosynthesis by coupling a thermodynamically unfavorable thiolase reaction with a favorable HMG-CoA condensation, thereby facilitating intermediate channeling (Fig. 1a)14. In previous work, we identified a HMGS homolog mucA, a ACAT homolog mucB, and a DUF35 homolog mucC within the muc biosynthetic gene cluster (BGC) in Streptococcus mutans (Fig. 1a), which directs the production of the antimicrobial compound reutericyclin A (RTC-A) and the immunomodulatory molecule mutanocyclin15–17. The muc BGC is widely distributed among Firmicutes, particularly in Streptococcus species16. Based on sequence homology and biosynthetic logic, MucABC was hypothesized to be an acyltransferase (ATase) essential for RTC-A biosynthesis15. Its divergence from the canonical role in mevalonate biosynthesis indicates a previously unrecognized evolutionary trajectory within the thiolase superfamily.

Here, we report functional and structural characterization of the MucABC enzyme complex from oral Streptococcus, exemplified by the S. macaca-derived thiolase complex. We show that this enzyme complex catalyzes a non-canonical Friedel–Crafts C-acetylation on a pyrrolidine-2,4-dione scaffold during RTC-A biosynthesis. Cryo-electron microscopy (cryo-EM) and biochemical analyses reveal that the neofunctionalization of this complex involves the loss of conserved catalytic motifs—such as the E–C–H–N tetrad in HMGS and the HDxF motif in ACAT—along with active-site remodeling for specialized substrate recognition. Furthermore, we demonstrate that this acetylation step is indispensable for the antimicrobial activity of RTC-A, providing Streptococcus a competitive advantage in its native niche. Together, our findings reveal a previously unrecognized evolutionary trajectory in which multimeric enzyme complexes from primary metabolism are repurposed for secondary metabolic functions, offering opportunities for engineering modular biocatalysts in synthetic biology.

Results
Characterization of Streptococcal ATases as Friedel-Crafts acylases

A BLASTP search using the MucABC sequence from S. mutans as the query showed 31–37% sequence identity with PhlACB from Pseudomonas protegens (known as PpATase), a rare Friedel-Crafts acylase known to mediate the reversible disproportionation of phloroglucinol (PG), monoacetylphloroglucinol (MAPG), and 2,4-diacetylphloroglucinol (DAPG; Supplementary Fig. 2a)18,19. Although their overall sequence similarity is low, the individual components of MucABC retain conserved domains found in both PpATase and the archaeal HMGS/ACAT/DUF35 complex (Supplementary Fig. 3), suggesting a potential conservation of thiolase-related chemistry. To investigate the catalytic activity of mucABC-encoded proteins, we selected three highly similar mucABC operons (89–100% identity) from host-adapted oral streptococci: S. mutans B30 (human), S. macacae NCTC11558 (monkey), and S. troglodytae TKU31 (chimpanzee; Supplementary Fig. 2a). Given that PpATase is functional only when the entire phlACB operon is co-expressed18–20, we constructed a plasmid expressing the full mucABC operon with a C-terminal His6-tag on MucB to facilitate purification (Supplementary Fig. 2b). Of the three variants, S. macacae mucABC exhibited superior expression in Escherichia coli, yielding the highest protein amounts and purity after affinity chromatography (Supplementary Fig. 2c), and was therefore chosen for biochemical characterization.

To perform in vitro enzymatic assays, we chemically synthesized preRTC-A (Supplementary Fig. 4), the presumed nonribosomal peptide synthetase-polyketide synthetase (NRPS-PKS)-derived biosynthetic precursor of RTC-A16. Four unnatural acetyl donors previously shown to be accepted by PpATase—N-acetylimidazole (N-AcIm), isopropenyl acetate (IPEA), ethyl thioacetate (EtSAc), and phenyl acetate (PA)—were tested with MucABC (Fig. 1b). Ultra-performance liquid chromatography (UPLC) analysis confirmed the formation of a product peak with identical retention time for all four acetyl donors (Fig. 1b). High-resolution mass spectrometry (HRMS) analysis of the enzymatic reaction using PA revealed a 42 Da shift from the substrate ion [M−H]- at m/z 306.2087 (Cald. 306.2075) to the production at m/z 348.2187 (Cald. 348.2180), consistent with an acetylation event (Fig. 1b). Scaled-up enzymatic reactions enabled product isolation and confirmation of its identity as RTC-A by 1H and 13C NMR spectroscopy (Supplementary Table 1 and Supplementary Fig. 5), with the acetyl group installed at the C-3 position of the pyrrolidine-2,4-dione core (Fig. 1b). No O-acetylated byproducts were detected in time-course analyses (Supplementary Fig. 6a). Since the chemically synthesized preRTC-A was obtained as a racemic mixture (due to the acidity of the α-proton), we further isolated the individual (R)- and (S)-preRTC-A via chiral separation. UPLC analysis confirmed that MucABC could utilize both enantiomers as substrate, as RTC-A was produced in both incubations with the same retention times compared to the standard (Supplementary Fig. 6b and 6c). Additionally, MucABC-mediated reverse conversion of RTC-A back to preRTC-A was observed, indicating that the C–C bond formed by MucABC is cleavable by the enzyme, thereby enabling RTC-A to serve as an acetyl donor in a transacetylation reaction (Supplementary Fig. 7). Collectively, these findings establish that MucABC from S. macacae functions as an acyltransferase (hereafter termed SmaATase) that catalyzes C3-acetylation of preRTC-A, thereby expanding the family of acyltransferases capable of mediating non-canonical Friedel-Crafts acylation.

Acetyl-CoA serves as acetyl donor for SmaATase and PpATase

Although various phenyl esters and thioesters have been employed as acetyl donors in in vitro assays of PpATase19,21,22, its physiological acetyl donor has remained unknown. To identify a biologically relevant donor for SmaATase, we evaluated several endogenous acyl donors, including acetyl-CoA, malonyl-CoA, acetoacetyl-CoA, and acetyl-phosphate. Intriguingly, RTC-A formation was clearly observed in the presence of acetyl-CoA, whereas no signal was observed with acetyl-P and trace signals were detected with either acetoacetyl-CoA or malonyl-CoA (Fig. 1b). To determine whether this feature extends to PpATase, we expressed and purified PhlACB using the same strategy as employed for SmaATase (Supplementary Fig. 8a and b). Consistent with the results for SmaATase, reactions containing PpATase and PG yielded both MAPG and DAPG when acetyl-CoA was supplied as the donor, with only trace-level signals were detected when acetoacetyl-CoA or malonyl-CoA were used as alternative donors (Supplementary Fig. 8c). The identity of DAPG was confirmed by co-elution with an authentic standard (Supplementary Fig. 8c), while MAPG was verified by HRMS (Supplementary Fig. 8d) and NMR spectroscopy (Supplementary Table 2 and Supplementary Fig. 9). Together, these results support acetyl-CoA as the potential physiological acetyl donor for both SmaATase and PpATase, aligning their function with that of canonical acetyltransferases23,24.

SmaATase requires tripartite assembly for catalysis

SDS-PAGE analysis confirmed successful co-purification of all three components of the SmaATase complex purified via affinity chromatography, despite only MucB carrying a C-terminal His6-tag (Supplementary Fig. 2c). Mass spectrometry further validated the presence of MucA, MucB, and MucC units in the enzyme solution, with experimentally determined masses matching theoretical predictions (Supplementary Fig. 10a). Size-exclusion chromatography indicated enzyme complex formation by showing a peak corresponding to approximately 400–600 kDa (Supplementary Fig. 10b). To determine the structural basis of SmaATase function, we resolved its cryo-EM structure at 2.32 Å resolution, revealing a dodecameric architecture composed of four MucA, four MucB, and four MucC units (Fig. 2a, Supplementary Table 3, Supplementary Figs. 11–13). This organization closely resembles the reported crystal structures of PpATase20 (Supplementary Fig. 2a) and the archaeal HMGS/ACAT/DUF35 complex14 (Fig. 1a). In addition to the dodecamer, we observed an alternative octameric assembly (2.16 Å resolution), consisting of a MucA4 and two MucB units (Fig. 2b, Supplementary Table 3, Supplementary Figs. 11–13). Structural superposition of the dodecameric and octameric forms revealed a root-mean-square deviations (RMSD) of 0.160 Å over 2254 superposed Cα atoms, with active sites displaying comparable configuration (Supplementary Fig. 14). Furthermore, native mass spectrometry revealed additional oligomeric species (Fig. 2c), including 11-mer (MucA4B3C4), 10-mer (MucA4B2C4), 9-mer (MucA4BC4), and 8-mer (MucA4C4). These findings were corroborated by native gel electrophoresis (Supplementary Fig. 15a), indicating that SmaATase exists as a heterogeneous mixture of assemblies under the in vitro conditions tested.

Similar heterogeneity was observed for PpATase, which was previously characterized as a 12-mer [PhlA4C4B4] by crystallography20. Additional 10-mer (PhlA4C2B4) and 8-mer (PhlA4B4) were detected by native mass spectrometry (Supplementary Fig. 15b), consistent with the three distinct bands observed by native PAGE (Supplementary Fig. 15a). Quantitative mass photometry analysis revealed their relative distribution as ~4:4:2 (8-mer:10-mer:12-mer, Supplementary Fig. 15c).

To assess the requirement for complex formation in catalytic activity, we co-expressed pairwise combinations of MucA, MucB, and MucC in E. coli. Co-expression of MucAB or MucBC yielded poorly soluble proteins (Supplementary Fig. 16a and b), and purified MucAC lacked detectable C-acetylation activity (Supplementary Fig. 16c and d). Given that previous studies implicated residues within the PhlC (ACAT) subunit are essential for catalysis in PpATase20,25, we evaluated the potential of MucB (the SmaATase ortholog of PhlC) to function independently. However, the individual expression of MucB in E. coli resulted in insoluble aggregates (Supplementary Fig. 17a). Although fusion to an N-terminal trigger factor (TF) improved solubility and enabled purification (Supplementary Fig. 17a), neither TF-tagged nor cleaved MucB displayed detectable C-acetyltransferase activity (Supplementary Fig. 17b and c). Surface plasmon resonance (SPR) showed that both the full SmaATase complex (KD = 2.52 × 10⁻⁶ M) and the isolated MucB subunit (KD = 6.04 × 10⁻⁶ M) are capable of binding the substrate, with the intact complex exhibiting slightly higher affinity (Supplementary Fig. 18a). However, because the catalytic pocket of MucB is situated at the interface with MucA and MucC (Supplementary Fig. 18b), and given that individual subunits of the related archaeal HMGS/ACAT/DUF35 complex are either insoluble (ACAT) or inactive (HMGS) when expressed alone14, we hypothesize that MucA and MucC help constrain MucB into a catalytically competent conformation.

Structural comparison of HMGS/ACAT/DUF35 complexes

Despite low sequence identity and distinct catalytic functions, SmaATase shares striking structural similarity with both PpATase and the archaeal HMGS/ACAT/DUF35 complex at multiple levels. First, in addition to their shared dodecameric architecture, the spatial arrangement of subunits in SmaATase closely resembles that observed in PpATase and the archaeal complex (Fig. 2d). Structural alignment of the HMGS-ACAT-DUF35 modules between SmaATase and either PpATase or the archaeal homology yields RMSD of 1.245 Å (585 Cα atoms) and 1.454 Å (363 Cα atoms), respectively, underscoring the high degree of structural conservation across these systems (Supplementary Fig. 19). Second, the DUF35 subunit architecture and positioning are also conserved. MucC, a family member of DUF35 protein, features an N-terminal rubredoxin-like zinc ribbon (Supplementary Fig. 20) and a C-terminal oligonucleotide/oligosaccharide-binding (OB) fold domain. Within SmaATase, MucC is embedded in the multimeric assembly and establishes multiple contacts with neighboring MucA and MucB subunits (Supplementary Fig. 21). This arrangement resembles that of PhlB in PpATase, which bridges PhlA and PhlC, as well as the DUF35 homolog in the archaeal complex that connects the core HMGS and ACAT units (Fig. 2e). Finally, conservation is observed in the thiolase fold and the CoA-binding chamber. Both MucA and MucB adopt a conserved five-layered αβαβα core architecture characteristic of thiolase superfamily enzymes (Supplementary Fig. 22)11,26. This structural signature is also preserved in PhlA and PhlC of PpATase and the HMGS and ACAT units of the archaeal complex (Supplementary Fig. 22). Additionally, a hallmark of the archaeal HMGS/ACAT/DUF35 complex is the presence of a fused CoA-binding site at the HMGS–ACAT interface, which facilitates shuttling of the pantetheinyl arm between active sites during catalysis (Fig. 2f). Notably, a similar interfacial chamber is retained in both SmaATase and PpATase (Fig. 2f, supplementary Fig. 23), despite the absence of the HDxF motif in the thiolase domains and the key Claisen condensation residues in the HMGS units (details will be discussed in the following section).

Together, these structural observations indicate that the HMGS/ACAT/DUF35 complex represents a deeply conserved architectural module that may have been evolutionarily repurposed from primary metabolism to secondary metabolism.

Key catalytic residue mutations support functional divergence

Given the striking structural conservation of archaeal and bacterial HMGS/ACAT/DUF35 complexes as acetyl-CoA-utilizing C-acetyltransferases, we conducted a phylogenetic analysis comparing their ACAT and HMGS units with representative members of the thiolase superfamily to elucidate the evolutionary origins of SmaATase and PpATase. As expected, enzymes are clustered according to their functions (Fig. 3a). MucA/PhlA and their homologs formed a clade closely related to archaeal HMGSs, while MucB/PhlC and their homologs grouped near archaeal ACATs (Fig. 3a), supporting their evolutionary relationship with the archaeal HMGS/ACAT/DUF35 complex involved in mevalonate biosynthesis. Beyond the αβαβα structural motif, a hallmark of the thiolase superfamily is a catalytic cysteine (Cys) residue that mediates CoA or acyl carrier protein (ACP) substrate attachment27. However, multiple sequence alignment revealed the absence of the typical E-C-H-N tetrad conserved among HMGSs in MucA/PhlA and related sequences (Fig. 3b). Similarly, although MucB/PhlC and their homologs retain the CxS and GHx motifs characteristic of ACATs, they lack the HDxF motif present in archaeal ACATs (Fig. 3b). These sequence deviations likely contribute to the functional divergence of bacterial HMGS/ACAT/DUF35 complexes from their archaeal counterparts.

To test the functional significance of these divergent residues, we first assessed whether SmaATase and PpATase could catalyze the Claisen condensation of acetyl-CoA to form HMG-CoA. LC-MS analysis revealed the formation of CoA, but not HMG-CoA, from incubations with either enzyme (Fig. 3c), indicating that both utilize acetyl-CoA solely as a donor. To further validate this, we prepared the archaeal HMGS/ACAT/DUF35 complex using the same purification protocols and generated two variants: one carrying an H284A substitution in the ACAT unit’s HDxF motif, and another with substitutions in the HMGS unit’s E-C-H-N catalytic tetrad entirely (Supplementary Fig. 24). While the wild-type complex produced HMG-CoA from acetyl-CoA, neither mutant yielded detectable product (Fig. 3c), confirming the essentiality of these residues for canonical HMGS activity.

To gain deeper insights into the sequence diversity of SmaATase-like enzyme complexes, we mined bacterial genomes in the NCBI database and identified 1,030 BGCs containing co-localized HMGS, ACAT, and DUF35 genes (Fig. 4a). The vast majority of the identified BGCs (1,001) lacked adjacent biosynthetic machinery for canonical secondary metabolism, only 29 BGCs were predicted to be associated with secondary metabolism (Fig. 4b). These include muc-like clusters from Streptococcus species, featuring hybrid NRPS-PKS assembly lines, and phl-like clusters from Pseudomonas species harboring type III PKSs (Fig. 4b).

Given the dominance of this uncharacterized category, and its lack of obvious biosynthetic partners, we next asked whether these minimal HMGS–ACAT–DUF35 loci might encode functionally distinct or evolutionarily intermediate enzyme complexes. To explore this possibility, we selected a representative candidate from Pseudomonas fluorescens for further analysis. This PfACAT/HMGS-DUF35 complex features a fused HMGS-DUF35 protein and, importantly, lacks any neighboring PKS or NRPS genes (Fig. 4b, Supplementary Figs. 25 and 26). Sequence alignment showed that its HMGS domain lacks the E-C-H-N tetrad, whereas its ACAT unit retains the catalytic Cys and conserved HDxF and GHx motifs (Fig. 3b), suggesting that the HMGS unit is inactive whereas the ACAT component remains functionally intact.

In vitro biochemical assays confirmed this functional divergence: the PfACAT/HMGS-DUF35 complex did not produce HMG-CoA or acetoacetyl-CoA from acetyl-CoA, resembling SmaATase and PpATase (Fig. 3c). In contrast, it catalyzed the thiolytic cleavage of acetoacetyl-CoA to generate acetyl-CoA, a reaction not observed for SmaATase and PpATase (Fig. 4c). Introducing an H294Y mutation in PfACAT nearly abolished this activity (Fig. 4c and Supplementary Fig. 25), confirming the essential catalytic role of the His residue within the HDxF motif. This finding is consistent with observations in Geobacter metallireducens BbsAB, a DUF35-associated sterol-carrier-protein (SCP2)-type thiolase where the HDxF motif mediates reversible benzylsuccinyl-CoA formation from benzoyl-CoA and succinyl-CoA28.

Taken together, these findings suggest that the PfACAT/HMGS-DUF35 complex may represent an evolutionary intermediate that has lost its primary metabolic function but has not yet been recruited into a defined specialized pathway. In contrast, SmaATase and PpATase have undergone further evolution, characterized by the loss of the HDxF motif and remodeling of the active-site architectures within the ACAT domain that enables specific recognition of small-molecule acetyl acceptors (Fig. 5). To elucidate the substrate-binding mode of SmaATase, we performed molecular docking and dynamics simulations (Fig. 5 and Supplementary Fig. 27). A comparison with the well-established catalytic mechanism of PpATase20,25 reveals that key catalytic residues are largely conserved: the catalytic nucleophile (Cys88 in PhlC vs Cys87 in MucB); the general base (Glu116 vs Asp115); the proton source residue and proton shuttling residues (Asp137-Thr132-His144 vs Asp136-Thr131-His143, Fig. 5 and Supplementary Fig. 28). Notably, the catalytic cysteine is acetylated in both the PpATase20 (Cys88, 5MG5) and SmaATase (Cys87) structures; this was evidenced by residual electron density in both cases and confirmed for SmaATase by LC-MS/MS analysis (Supplementary Fig. 29).

Despite the conserved catalytic core, clear differences are observed in the substrate-binding region. For example, His56 and Tyr124 in PhlC are replaced by Ala55 and Phe123 in MucB, respectively, and the flexible lid residue Trp211 in PhlC29 is substituted by Ala210 in MucB (Fig. 5 and Supplementary Fig. 28). Sequence alignment further shows that key catalytic and substrate-binding residues are well conserved within PhlC or MucB homologs, yet markedly divergent from those in archaeal ACATs (Supplementary Fig. 28). Together, these observations indicate that although SmaATase and PpATase retain a shared catalytic apparatus inherited from their ancestral thiolase scaffolds, they have undergone functional repurposing through reconfiguration of their substrate-binding sites, enabling them to recognize chemically distinct acetyl acceptors.

Acetylation confers antimicrobial activity of RTC-A and DAPG

RTC-A is a tetramic acid-derived antibiotic with broad-spectrum activity against gram-positive bacteria15,30,31. To evaluate the biological relevance of the acetylation modification catalyzed by streptococcal ATases, we compared the antimicrobial activities of RTC-A and its unacetylated precursor, preRTC-A, against both oral streptococcal competitors and clinically relevant pathogens. Strikingly, RTC-A exhibited potent antibacterial effects against all tested strains, whereas preRTC-A showed negligible inhibition (Fig. 6 and Supplementary Fig. 30). A similar structure–activity relationship was observed for the DAPG/PG pair, with DAPG displayed markedly stronger antimicrobial activity than PG against most strains (with the exception of S. gordonii and S. oralis) (Fig. 6 and Supplementary Fig. 30). Together, these results establish acetylation as a critical modification for the antimicrobial potency of both RTC-A and DAPG, underscoring the ecological significance of evolved ATases from Streptococcus and Pseudomonas species in interspecies competition.

Methods
General methods

All reagents were purchased from Sigma-Aldrich, Thermo Fisher Scientific, Bidepharm, and Aladin. Gene synthesis, plasmid construction, and point mutation were carried out by Tsingke Biotechnology Company (Supplementary Table 4). All codon-optimized DNA and protein sequences used in this study are provided as a Source Data file. Luria-Bertani (LB, 1% NaCl, 1% tryptone, and 0.5% yeast extract) medium was used for culturing E. coli. Chloramphenicol (25 μg/mL), ampicillin (50 μg/mL), and kanamycin (50 μg/mL) were used for the selection of recombinant E. coli strains. Antibacterial activity assays were performed using indicator strains cultured in Mueller-Hinton broth (MHB, OXOID), except for S. oralis and S. gordonii, which were maintained in brain-heart infusion (BHI, OXOID) broth due to poor growth in MHB.

Protein expression and purification

Codon-optimized genes encoding SmaATase (mucABC), PpATase (phlACB), archaeal HMGS/ACAT/DUF35 complex, pfACAT/HMGS-DUF35 complex, and individual combinations of MucABC subunits were cloned into the pACYCDuet-1 and transformed into E. coli BL21(DE3) for protein expression. Codon-optimized mucB gene was cloned into both pET28a(+) and pCold-TF (TF = trigger factor) for enhanced soluble expression. Cells harboring the expression plasmid were grown overnight in LB supplemented with appropriate antibiotic at 37 °C and 220 r.p.m. to prepare the preculture. Five milliliters of preculture were then inoculated into 500 mL of LB medium containing the same antibiotic and cultured under identical conditions. Protein expression was induced with 0.2 mM isopropyl β-D-thiogalactopyranoside (IPTG) at OD600 ~0.6, followed by incubation at 30 °C (21 h, 140 rpm). For the expression of MucB, the culture was incubated at 16 °C, 160 rpm for 16 h after adding 0.2 mM IPTG. Cells were harvested and resuspended in lysis buffer (50 mM Tris-HCl, 500 mM NaCl, 10 mM imidazole, 10% glycerol, pH 8.0). The suspension was disrupted by ultrasonication (40% amplitude, 10 min, pulse 3 sec, pause 6 sec) and then centrifuged at 13,000 × g for 30 min to spin down the cell debris. Recombinant proteins were purified using an ÄKTA go system (Cytiva) equipped with HisTrap HP columns (Cytiva). Eluted proteins were buffer-exchanged into 50 mM Kpi buffer (pH 7.5) containing 10% glycerol using PD-10 desalting columns (Cytiva). Protein concentrations were determined after concentration with Amicon Ultra filters (Merck Millipore). Size-exclusion chromatography was performed using an ÄKTA go system equipped with a Superdex 200 Increase 10/300 GL column (Cytiva) pre-equilibrated with 50 mM Kpi buffer (pH 7.5) containing 150 mM NaCl. Purity of proteins was assessed by SDS-PAGE analysis (Supplementary Fig. 32) and imaged using the GelDoc Go system (Bio-Rad). Freshly purified enzyme complexes were used for all in vitro assays to ensure activity.

Activity assays

Enzymatic assays for functional characterization of SmaATase were carried out in 100 μL reaction mixtures containing 50 mM Kpi buffer (pH 7.5), 12 μM purified enzyme, 1 mM preRTC-A, and various acetyl donors at the following concentrations: 1.5 mM PA, 25 mM N-AcIm, 10 mM IPEA (supplemented with 10 mM imidazole), 10 mM EtSAc, 1.5 mM acetyl-CoA, 1.5 mM malonyl-CoA, 1.5 mM acetoacetyl-CoA, 1.5 mM acetyl-P. Reactions were incubated at 30 °C for 16 h and quenched by the addition of 100 μL methanol. After centrifugation at 21,000 × g for 30 min, supernatants were analyzed by ultra-performance liquid chromatography (UPLC) using LC method 1 (Supplementary Table 5). UPLC was performed on an Agilent 1290 system equipped with a Phenomenex Kinetex XB-C18 column (4.6 mm × 100 mm, 2.6 μm), and data were processed using Agilent OpenLab CDS ChemStation Edition C.01.07.

For PpATase activity assays, 100 μL reaction mixtures contained 50 mM Kpi buffer (pH 7.5), 12 μM purified enzyme, 1 mM PG, 1.5 mM of either PA, or acetyl-CoA, malonyl-CoA, acetoacetyl-CoA, acetyl-P. Reactions were incubated at 30 °C for 16 h, and quenched with 100 μL methanol, and centrifuged at 21,000 × g for 30 min. Supernatants were analyzed by UPLC using LC method 2 (Supplementary Table 5).

For in vitro HMG-CoA formation assays, 50 μL reaction mixtures were prepared containing 50 mM Kpi buffer (pH 7.5), 15 μM purified enzyme, 1 mM acetyl-CoA, and 1 mM CoA. After incubation at 30 °C for 2.5 h, reactions were quenched with 2.5 μL formic acid and 50 μL methanol. Samples were centrifuged at 21,000 × g for 30 min, and supernatants were analyzed by liquid chromatography-mass spectrometry (LC-MS) using LC method 3 (Supplementary Table 5). Thiolytic cleavage activity of acetoacetyl-CoA was assessed in 50 μL reaction mixtures containing 50 mM Kpi buffer (pH 7.5), 15 μM purified enzyme, 1 mM acetoacetyl-CoA, and 1 mM CoA. After incubation at 30 °C for 30 min, reactions were quenched with 2.5 μL formic acid and 50 μL methanol. Following centrifugation (21,000 × g, 30 min), supernatants were analyzed by LC-MS using LC method 4 (Supplementary Table 5) on a Phenomenex Kinetex XB-C18 column (4.6 mm × 100 mm, 2.6 μm).

All LC-MS analyses were performed on the 1290 infinity UPLC system coupled to an Agilent Technologies 6470 triple quadrupole mass spectrometer. The LC/TQ used a Jet Stream Electrospray Ionization (AJS-ESI) source operated in the negative ionization mode (capillary voltage, 3.5 kV; gas temperature, 300 °C; drying gas, 5 L·min-1; fragmentor voltage: 135 V; nebulizer, 45 psig; sheath gas flow, 11 L·min-1; acquisition range, 100–1500 m/z). MS data were processed using Agilent MassHunter Qualitative Analysis B.06.00.

Isolation and characterization of the enzymatic products

To determine the structure of the enzymatic product of SmaATase, the in vitro reaction was scaled up to 50 mL reaction volume containing 50 mM Kpi buffer (pH 7.5), 1 mM preRTC-A, 1.5 mM PA, 15 μM purified enzyme, and incubated at 30 °C for 16 h. For MAPG isolation, the reaction was similarly scaled up to 100 mL using 1 mM PG and the same buffer and enzyme conditions, followed by incubation at 30 °C for 16 h. Reaction mixtures were extracted three times with equal volume of ethyl acetate (EtOAc), and the organic phases were combined and concentrated under vacuum. RTC-A (tR = 10.6 min) was purified via semi-preparative HPLC on a Phenomenex Kinetex XB-C column (250 mm × 10 mm, 5 μm) using isocratic elution with acetonitrile (ACN)/H2O (95:5, v/v) containing 0.1% TFA at a flow rate of 2.5 mL/min. MAPG (tR = 6.9 min) was purified under similar conditions using ACN/H2O (40:60, v/v) with 0.1% TFA. The 1H and 13C NMR spectra were recorded on a Bruker Advance Neo 400 MHz or 600 MHz spectrometer and processed by MestReNova 6.1.0 (Metrelab). NMR data are presented in Supplementary Tables 1 and 2, and NMR spectra are provided in Supplementary Figs. 4, 5, 7, and 9.

Cryo-EM structural determination

For Cryo-EM sample preparation, Holey carbon grids (Au R1.2/1.3, 300 mesh, Quantifoil) were used with 30 s glow-discharge using medium power of Plasma Cleaner PDC-32G (Harrick). The Vitrobot Mark IV (Thermo Fisher Scientific) was maintained at the conditions of 8 °C and 100% humidity during sample preparation. For each sample, 3.5 μL of protein (3.8 mg/mL of SmaATase in 50 mM potassium phosphate, pH 7.5) were loaded on the grid, blotted for 4.5 s with filter paper, and plunged into liquid ethane. The vitrified grids were subsequently transferred to liquid nitrogen for storage.

Data acquisition was performed on the 300 kV Titan Krios electron microscope (Thermo Fisher Scientific) equipped with a Falcon 4i detector and GIF Quantum energy filter. A total of 3089 micrographs were collected automatically using EPU, with a preset defocus range of −1.2 to −1.8 μm. Each image stack received a total dose of 50 e-/Å², and the final images were binned to a pixel size of 0.927 Å.

Data processing flow shown in Supplementary Fig. 11 was performed on cryoSPARC v4.7. Initial particle picking using blob picking yielded 1,155,067 particles. Following 2D classification, 973,419 particles were retained and re-extracted at bin 1 for subsequent processing. Ab-initio reconstruction of these particles produced an octameric structure, and the resolution of this structure reaches 2.16 Å after non-uniform refinement. During 2D classification, we observed particles exhibiting higher-order polymeric assemblies. Through iterative ab-initio reconstruction and 3D classification, we isolated 71,520 particles for subsequent processing. Non-uniform refinement followed by local refinement of these particles yielded a dodecameric structure at 2.32 Å resolution.

Structural models for the octameric and dodecameric complexes were built based on their respective cryo-EM maps. The initial monomeric models of MucA, MucB, and MucC, predicted by AlphaFold2, were docked into the density maps using ChimeraX to generate initial complex models. These models were then manually adjusted and refined in Coot, followed by automated real-space refinement using phenix.real_space_refine in PHENIX. The stereochemical quality of the final models was validated with MolProbity. Data collection and refinement statistics are summarized in Table S3. The atomic coordinates and structure factors have been deposited in the Protein Data Bank under accession codes 9VBO (octamer) and 9VBT (dodecamer).

Mass spectrometry

For TOF mass spectrometry, SmaATase (n = 3) was buffered to 50 mM ammonium acetate (pH 7.5) before applying on SYNAPT XS (Waters) mass spectrometer equipped with an ACQUITY UPLC I-Class PLUS system. A 2 μL aliquot of protein solution (1 mg/mL) was injected onto a MAbPacTM RP column (3.0 mm × 100 mm, 4 μm, Thermo Fisher Scientific) and separated using LC method 5 (Supplementary Table 5). Mass spectrometry was performed on a Waters SYNAPT XS operating in positive ESI mode over the range of 200−4000 m/z.

Chromatography-coupled mass spectrometry was performed at 1 mm scale with the following optimized conditions: Electrospray voltage: 3.0 kV, ion source: 120 °C, desolvation: 450 °C, cone gas: 50 L/h, desolvation gas: 800 L/h, nebulizer pressure: 6.5 bar. Raw spectra were acquired using MassLynx 4.2 (Waters) and processed through Maximum Entropy (MaxEnt) deconvolution to generate the molecular mass spectra of injected proteins.

Native MS analysis was performed using a Q-Exactive Plus Hybrid Quadrupole-Orbitrap Mass Spectrometer (Thermo Fisher Scientific). Samples (n = 3) were buffer-exchanged into 0.2 M ammonium acetate (pH 7.5) and loaded on a gold-coated capillary installed on a nESI source. Spectra were acquired with 1.4 kV spray voltage, 20 ms maximum inject time, and 50−200 V collisional activation in the HCD cell. Mass and charge assignments were determined using an Excel script kindly provided by the Benesch Laboratory, University of Oxford.

For protein posttranslational modification analysis, SmaATase (n = 3; 2.5 mg/mL in 50 mM Kpi buffer, pH = 7.5) were reduced with TCEP [Tris(2-carboxyethyl)phosphine, 10 mM] at 37 °C with agitation (800 rpm) for 60 min, alkylated with iodoacetamide (IAA, 40 mM) in the dark at room temperature for 30 min, and digested in-solution with a combination of trypsin and chymotrypsin, both at a ratio of 1:50 (w/w) to SmaATase, in 50 mM ammonium bicarbonate at 37 °C for overnight. The digestion was quenched with 0.1% formic acid, and the resulting peptides were desalted prior to LC-MS/MS analysis. Approximately 1 μg of sample was injected into a Vanquish Neo nanoflow LC system equipped with a Orbitrap Exploris 480 Mass Spectrometer (Thermo Fisher Scientific). Peptides were separated on an Acclaim PepMap 100 C18 column (250 mm × 75 μm, 3 μm, Thermo Fisher Scientific) by using LC method 6 (Supplementary Table 5). Full-scan MS spectra (m/z 350–1500) were acquired in the Orbitrap in DDA mode at a resolution of 120,000 (at m/z 200), with an RF lens setting of 50%, an automatic gain control (AGC) target of 300%, and a max IT of 25 ms. MS/MS analysis was performed with the following settings: MS2 resolution of 15,000 (at m/z 200); the AGC target was set to 200% and the max IT to 22 ms; dynamic exclusion was set to 30 s; MS2 scan range was set to m/z 150–2000. The most intense ions fulfilling the predefined criteria were isolated with a 1.6 Th window and fragmented via higher-energy collisional dissociation (HCD) with a normalized collision energy (NCE) of 28%. The cycle time was 2 s, the dynamic exclusion duration was 30 s, and all other parameters were set to default values. Database searching was performed using Peaks Studio 13 software. The mass error tolerances were set to 10 ppm for precursor ions and 0.02 Da for fragment ions; enzyme specificity was set to semi-specific for both trypsin and chymotrypsin, allowing for a maximum of two missed cleavages and a minimum of 6 amino acid residues for the peptide sequence length. Protease cleavage sites were defined for trypsin and chymotrypsin at lysine (K) and arginine (R) residues. The following variable modifications were included in the search: Oxidation (M: +15.9949 Da); Acetylation (C: +42.0106 Da); Acetylation (Protein N-terminus: +42.0106 Da); Carbamidomethylation (+57.0215 Da); Carboxymethylation (+58.0055 Da). Filter parameters were set as follows: PSM -10LgP ≥ 20.0, Proteins -10LgP ≥ 15.0. PTM localization was assessed using an Ascore threshold of 15, and a minimum of one unique peptide was required for protein identification.

Mass photometry measurements

Protein samples were diluted in phosphate-buffered saline (PBS) to a final concentration of 200 nM for mass photometry analysis. Measurements were performed on a OneMP instrument (Refeyn) using pre-cleaned glass coverslips. Following the standard protocol, 10 μL of PBS was loaded into the sample well for system focusing, followed by the addition of 10 μL diluted protein sample (1:1 mixing ratio). Data acquisition commenced immediately after mixing, recording for 60 s at 100 frames/s using AcquireMP software v2.3. The system was calibrated with bovine serum albumin (BSA) standards. Raw data were processed using DiscoverMP software v2.1.

Phylogenetic analysis

Protein sequences of thiolase superfamily members were compiled from published datasets. Homologs of MucA, MucB, PhlA, and PhlC were obtained via BLASTP searches against the NCBI non-redundant protein database (nr). All sequences used in the analysis are provided as a Source Data file. Multiple sequence alignments were performed using MUSCLE with default parameters. Phylogenetic reconstruction was conducted in MEGA 11 using the Neighbour-Joining method with the Poisson model, incorporating 500 bootstrap replicates to assess node support. The resulting Newick-formatted tree was visualized using the Interactive Tree of Life (iTOL) server tool (https://itol.embl.de/).

Genome mining methods

To systematically identify multi-component enzyme complexes with C-acyltransferase activity across the bacterial kingdom, a domain-centric genome mining approach was employed. Based on the biosynthetic gene cluster organization of muc and phl BGC, hidden Markov model (HMM) search were conducted for Pfam domain corresponding to HMGS (MucA and PhlA), ACAT (MucB and PhlC), DUF35 proteins (MucC and PhlB). Specifically, the domains corresponding to PF08541 [3-oxoacyl-(acyl-carrier-protein) synthase III C-terminal], PF00108 (thiolase), and PF01796 (DUF35 OB-fold domain), respectively, served as probes for further genome mining. The thiolase domain was used as the anchor, and genomic regions encompassing 10 genes upstream and downstream were scanned for the presence of the other two domains. Loci containing all three domains were retained, and redundancy was eliminated using FastANI. The resulting BGCs were analyzed using antiSMASH and visualized with BiG-SCAPE.

Molecular docking and molecular dynamics simulation

The ligand structure was first optimized at the B3LYP/def2-TZVP level of theory using the ORCA 5.0.4 quantum chemistry package to obtain a reliable equilibrium geometry and electronic structure. The optimized ligand was subsequently docked into the target protein using the DiffDock. The top-ranked docking pose, based on DiffDock’s confidence score and binding-site complementarity, was selected for further molecular dynamics (MD) simulations. Docking poses were visualized using PyMOL v3.0 and ProLIF.

To investigate the interaction between the ligand and the MucA-B-C protein complex, classical molecular dynamics (MD) simulations were carried out. In the constructed systems, the protonation states of all ionizable residues were determined using PROPKA, based on empirical pKa predictions under physiological pH conditions. The following residues were modeled in their protonated form: His305 and Asp35 in MucA; Glu57, Asp134, and Glu296 in MucB; and Glu70 in MucC. All remaining residues retained their standard protonation states at neutral pH. All MD simulations were performed with GROMACS (version 2025.03) using an all-atom force field for the protein (AMBER ff14SB) and a compatible ligand topology generated via appropriate parameterization (GAFF2). RESP charges were generated using Psi4 for all ligands as well as for the acetylated Cys87 residue. The protein–ligand complex was solvated in a cubic box with explicit TIP3P water molecules, and counter ions (Na⁺/Cl⁻) were added to neutralize the total charge.

Prior to dynamics, the system underwent energy minimization using the steepest descent algorithm until the maximum force fell below 1000 kJ·mol⁻¹·nm⁻¹, ensuring removal of steric clashes. Subsequently, two equilibration phases were performed: The system was equilibrated for 1 ns under the canonical ensemble using the V-rescale thermostat to maintain the temperature at 300 K, while restraining the heavy atoms of the protein and ligand with a force constant of 1000 kJ·mol⁻¹·nm⁻¹. The simulation was continued for 1 ns under the isothermal–isobaric ensemble (NPT), maintaining a pressure of 1 bar using the Parrinello–Rahman barostat. Position restraints were retained to allow solvent and ions to relax around the solute. Finally, to improve statistical sampling, three independent 200 ns production MD simulations were conducted in the NPT ensemble without positional restraints, starting from distinct randomized initial velocity distributions. A 2 fs integration time step was employed, and all covalent bonds involving hydrogen atoms were constrained using the LINCS algorithm. Long-range electrostatic interactions were treated with the Particle Mesh Ewald (PME) method, and van der Waals interactions were truncated at 1.0 nm. Trajectories were saved every 10 ps for subsequent analyses. The binding free energy between the ligand and protein was calculated using the MM/GBSA method.

Antibacterial activity and MIC assays

Stock solutions of preRTC-A, RTC-A, PG, and DAPG were prepared in DMSO at concentrations of 6.4 mg/mL or 8 mg/mL. Ampicillin sodium salt, dissolved in water at the same concentrations, served as a positive control. The following indicator strains were used: Streptococcus salivarius DA547, Streptococcus oralis DA1241, Streptococcus gordonii ATCC10558, Bacillus subtilis 168, Staphylococcus aureus ATCC29213, Methicillin-resistant Staphylococcus aureus 544, Methicillin-resistant Staphylococcus aureus 103 and Streptococcus pneumonia ATCC49619. For MIC assays, compounds were serially 2-fold diluted in MHB to 50 μL per well in 96-well plates. Bacterial cultures in mid-log phase were diluted to 1 × 106 CFU/mL in MHB, and 50 μL of this suspension was added to each well, yielding a final inoculum of 5 × 105 CFU/mL (validated range: 2 × 105 CFU/mL to 8 × 105 CFU/mL). Streptococcus oralis DA1241 and S. gordonii ATCC10558 were cultured in BHI due to poor growth in MHB. Each plate included antimicrobial-free medium as a negative control and untreated bacterial culture as a growth control. Plates were incubated at 37 °C for 18 h, and absorbance was measured at 625 nm using a BioTek Epoch 2 (Agilent). MIC values were defined as the lowest concentration that completely inhibited visible growth (OD625 ≤ 0.02 compared to the negative control). All assays were performed in duplicate and independently repeated three times.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In their manuscript, Liao et al study bacterial tripartite complex systems analogous to the thiolase/HMG-CoA synthase/DUF-35 protein used by archaea to catalyse the first steps of isoprenoid synthesis. Through an impressive panel of methods ranging from enzymology, structural biology, biochemistry, bioinformatics, phylogeny, and even genome mining, they found that the thiolase subunit from the bacterial homologues from Streptomyces macacae (SmaATase) and from Pseudomonas protegens (PpATase) catalyse a rare Friedel-Crafts reaction involving acetyl-CoA as acetyl donor. The end products of these reactions ultimately generate molecules with antibiotic activities. It is without any doubt a study of high quality, bringing to light an enzymatic complex involved in secondary metabolism that most probably originates from a central carbon metabolism involved in isoprenoid biosynthesis. The reviewer congratulates the authors on this excellent study that reaches the standard of Nature Communications, and would strongly support its publication upon addressing these minor comments.\n1- Abstract: “Cryo-electron microscopy of the S. macaca-derived thiolase complex”.\nShouldn´t it be S. macacae instead of S. macaca?\n2- Title of the result first subsection. “Characterization of Streptococcal ATases as non-canonical Friedel-Crafts acylases”. For clarification, it would be helpful to state the meaning of “ATases” here or before instead of using the abbreviation. Moreover, since the physiological donor is acetyl-CoA, authors should consider using the term acetylases instead of acylases.\n3- Legend of Supplementary Fig. 2c. “SDS-PAGE analysis of the heterologous expression and purification results of SmaATase, SmuATase, and StrATase, respectively. SmaATase, SmuATase, and StrATase represent MucABC enzyme complex from S. mutans, S. macacae, S. troglodytae, respectively.”\nBased on the text and results, it should be: “SDS-PAGE analysis of the heterologous expression and purification results of SmaATase, SmuATase, and StrATase, respectively. SmaATase, SmuATase, and StrATase represent MucABC enzyme complex from S. macacae, S. mutans, S. troglodytae, respectively.”\n4- 2nd paragraph results: “Additionally, MucABC-mediated reverse conversion of RTC-A back to preRTC-A was observed, confirming the bidirectionality of the reaction (Supplementary Fig. 6d).” Would it be possible to provide the Standard Gibbs free energy for the reaction? That would provide some insights on its expected directionality.\n5- 3rd paragraph results: “Notably, earlier studies reported no detectable product formation when PpATase was incubated with acetyl-CoA”. By curiosity, do the authors have an explanation for why the activity was not observable before with acetyl-CoA?\n6- Supplementary Fig. 9. Please define “Cald.” in the legend. Could the authors explain the differences between observed and calculated masses for MucB and MucA (e.g., 44481 Da versus 44351 Da)?\n7- Supplementary Fig. 10. Has the mask used to generate the local refinement been deposited in the EMDB? Adding an extra panel with the mask on Fig. S10 would be informative and provide more transparency on the model-building process. Moreover, for Figures S10 and S11, is the map based on raw data or a sharpened one? This should be stated in the legend.\n8- Figure 2. According to the legend, panels d and e must be swapped.\n9- 4th paragraph results: “In addition to the dodecamer, we observed an alternative octameric assembly (2.16 Å resolution), consisting of a MucA4C4 core”. The nomenclature seems wrong and should be MucA4C2 according to the deposited model.\n10- 4th paragraph results: “Structural superposition of the dodecameric and octameric forms revealed a root-mean-square deviations (r.m.s.d.) of 0.173 Å”. Please add the number of superposed Calpha.\n11- 4th paragraph results: “…indicating that SmaATase exists as a heterogeneous mixture of assemblies under near-physiological conditions.”. This is a very dangerous statement that must be reformulated because the tested conditions in vitro are expected to be very different compared to the physiological ones (molecular crowding, physicochemical conditions, protein concentrations…). Rather, use “under in vitro conditions” or “in solution”.\n12- 7th paragraph results: “Structural alignment of the HMGS-ACAT-DUF35 modules between SmaATase and either PpATase or the archaeal homolog yields r.m.s.d. of 1.245 Å and 1.466 Å,” Please add the number of superposed Calpha.\n13- 7th paragraph results: “DUF35 subunit architecture and positioning: MucC, a family member of DUF35 protein, features an N-terminal rubredoxin-like zinc ribbon and a C-terminal oligonucleotide/oligosaccharide-binding (OB) fold domain.”. IMPORTANT POINT. Unfortunately, MucC has been modelled without the 4 conserved Cys binding the Zn. The Zn is observable in the EM map together with the four Cys binding to it. See the attached picture.\nZn has been modelled in all structural homologues, and must be modelled in both structures from this work. Presenting an additional supplementary figure with a close-up of the Zn binding site and the electron density would be informative.\nMoreover, Cys87 in ACAT subunit of SmaATase could be acetylated. Mass spectrometry might provide some clues if this residue is truly acetylated in the protein in solution (e.g., with the acetyl-group coming from acetyl-CoA from E. coli). Based on the electron density, there is high suspicion that Cys87 is indeed acetylated. Therefore, the referee suggests adding an acetyl group on this cysteine (as it has been done for PpATase structure), and if possible, obtain a confirmation by mass spectrometry. Presenting an additional supplementary figure with a close-up of the acetylated Cys87 and the electron density would be informative. If mass spectrometry, or author´s conclusions point to a non-acetylated form, please add a snapshot of the Cys87 with the surrounding electron density for each chain.\nTo conclude on the model, the referee would suggest adding the Zn, the acetyl group and polishing the model further for a better fit in the electron density. For instance, some loops and sidechains are observable in the electron density, but were not modelled (see below). Please correct the residue presenting a very high clash score (higher than 0.8).\n14- Supplementary Fig. 17. Presented residues should be labelled. A second snapshot with a 90° orientation could be useful to show the different residues. Also, hydrogen bond distances are impossible to read. Please remove the distances label, keep the dashes (in another colour than yellow) and add a general statement such as: all hydrogen bond distances below 3.5 Å are displayed as dashes.\n15- 7th paragraph results: “). Additionally, a hallmark of the archaeal HMGS/ACAT/DUF35 complex is the presence of a fused CoA-binding site at the HMGS–ACAT interface, which facilitates shuttling of the pantetheinyl arm between active sites during catalysis (Fig. 2f).”\nA supplementary figure comparing the CoA binding site between the archaeal ACAT/HMGS/DUF-35 protein (PDB 6ESQ) and SmaATase would reinforce the observation made on Fig. 2f. Are most of the residues conserved? Also, please correct the typo on Fig. 2f “fused Co-A binding” should be “fused CoA-binding”.\n16- 11th paragraph results: “Introducing an H294Y mutation in PfACAT nearly abolished this activity (Fig. 4c and Supplementary Fig. 20), confirming the essential catalytic role of the His residue within the HDxF motif.”. This is a very interesting point. Are the authors expecting a role in antibiotic biosynthesis for this enzyme? It is unlikely that its physiological function would be simply the formation of acetyl-CoA from acetoacetyl-CoA. The latter might simply be a side reaction.\n17- Figure 5: Cys88 is acetylated, please label the residue accordingly (e.g., Acetyl-C88). D137 cited in the text is not shown on the figure. Importantly, the difference in the active site pose confuses the reader as it seems that the orientation between PpATase and SmaATase is the same, but the position of the residues (particularly C88/H347 and C87/H349) is different. Please take a new picture with the exact same orientation between PpATase and SmaATase active sites. Highlight the key residues C88/H347 and C87/H349 with a bold label.\n18- Supplementary Fig. 23: Why is the hydrogen bond not shown for Q302? Is there a possibility to switch the H143 rotamer to gain a hydrogen bond with the ligand without generating steric clashes? Here, 4.1 A seems too distant for efficient\n19- Supplementary Fig. 24: The final concentration of the used compounds must be stated in the figure or legend.\n20- Last paragraph of the result section: “Although preRTC-A showed modest activity against S. gordonii ATCC 10558 (MIC = 32 μg/mL), RTC-A demonstrated a 32-fold enhancement in potency (MIC = 1 μg/mL; Fig. 6). A similar trend was observed for the phenolic polyketide DAPG and its biosynthetic precursor PG. DAPG exhibited significantly higher antimicrobial activity against most tested strains (except S. oralis DA1241 and S. gordonii ATCC 10558) compared to PG, recapitulating the structure–activity relationship observed for RTC-A and preRTC-A pair (Fig. 6 and Supplementary Fig. 24).”. The whole sentence should be clarified. The results obtained are excellent, and authors should rather use a general statement to describe the results, for instance: All strains showed inhibition for DAPG compared to PG, except S. gordonii and S. oralis. The effect is even more pronounced for preRTC-A versus RTC-A, where all tested strains showed inhibition for RTA-A compared to preRTC-A.”\n21- 1st paragraph discussion: “In contrast, SmaATase is remarkable for its ability to catalyze Friedel–Crafts acylation independently of CoA or ACP-activated acyl donors, highlighting its potential as a versatile biocatalyst for synthetic applications.” The acetylation is CoA-dependent. Please rephrase.\n22- 2nd paragraph discussion: “…enzyme complexes in secondary metabolism are more commonly associated with scaffold formation (such as PKS37 and NRPS38) than with structural modification”. What is the meaning of structural modification here? Please reformulate.\n23- Supplementary Fig. 25: It would help the reader to add the acetylated cysteine residue position in the three studied systems (in the figure or in the legend).\n24- Methods, protein expression and purification: “and imaged using the DelDoc Go system (Bio-Rad).”. It should be GelDoc Go system.\n25- Methods, Cryo-EM structural determination: Please provide the following information.\nProtein concentration and buffer used for the grid.\nHow did authors build, refine, and validate the model?\n“Ab-initio reconstruction of these particles produced an octameric structure at 2.16 Å resolution.” Ab initio is not a refinement technique, so it does not make sense to associate ab initio with resolution. Please reformulate.", "reviewer_2": "Xiaoyu Tang and colleagues present a manuscript detailing the biochemical characterization of a tripartite enzymatic complex catalyzing Friedel-Crafts-Acylation in the biosynthesis of the antibiotic tetramate reutericyclin A. This enzymatic complex bears similarities to a previously characterized system able to catalyze acetylation of phloroglucinol. The protein complex was structurally characterized via Cryo-EM, revealing a dodecameric structure, analogous to a previously characterized pathway in archaeal mevalonate biosynthesis. Bioinformatic analyses showed that key catalytic residues of the primary metabolism enzyme were lost, and new, potentially critical residues for a new catalytic functon(acetylation) acquired. Based on these findings, the authors pose the hypothesis that the enzymatic complex was repurposed during evolution to acquire a new catalytic function. Finally, the authors show that acetylation is an essential feature for antibiotic activity in both reutericyclins and phloroglucinols, highlighting the biological importance of this modification.\nThe paper is very well and clearly written and contains a lot of data. The experiments were carried out with a range of state-of the-art methodologies, and the results are nicely presented in main and SI Figures. Overall, this study deserves publication in Nature Communications. However, there are a couple of issues that should be improved or taken care of before publication:\nMajor:\nl.265: SmaATase is praised having potential as a “versatile biocatalyst for synthetic applications.” Indeed, the possible use of different acetyl donors has been demonstrated, but the substrate scope has not been investigated at all to my understanding, except for the acetylation of the native substrate pre-RTC and its short chain analogue. Of course, a comprehensive enzymatic study is beyond the scope of this paper, but it would be nice to provide some initial characterization: e.g. on simplified pyrrolidones with a few different substituents. Also, I wonder, if propionyl chains could be accepted?\nFigure 3b: The loss of key catalytic residues of primary metabolic enzymes has been shown, but I am missing a Figure, where the acquisition of key residues for the new catalytic functon(see Figure 5, Fig. S23) is analyzed. It would be good to provide a similar Figure and underlying alignments for these residues in primary vs. secondary metabolism. Do all 31 enzymes contain the same sequence motifs in this region or are there variations?\nFigure 4b,c and corresponding main text l.218-230: This part is not entirely clear to me: The first sentence “While no other function predictable Friedel–Crafts acylases were identified” makes no sense to me. Also, to me it is not clear, why the enzyme from P. fluorescens was selected for further analysis. The rationale behind this experiment should be better explained, as well the meaning of the obtained results.\nMinor:\nMethods: In the genome mining section, a reference for BiG-SCAPE should be added.\nFig. 3a: Why are MucC/PhlB homologues not included in the phylogenetic tree?\nFig. S7: What is the meaning of PA in this Figure?\nFig: S24: The photos are barely visible and recognizable. Please improve Figure quality.\nFig. S26: In this Figure, it is referred to proteins in “Extended Data Fig. 2,4,6”. I don´t see these data anywhere in the submitted files.\nSource data Fig. b (BGCs): It is nice, that the BGCs are displayed. However, these collections in the Excel sheet confuse me: Either, the arrow colors must be clearly explained, or better, cleaner SI Figures visualizing and aligning the BGCs should be provided.", "reviewer_3": "The manuscript entitled \"Evolutionary repurposing of a metabolic thiolase complex enables antibiotic biosynthesis\" by Liao et al. presents an interesting and informative structural and functional investigation of SmaATase, a multimeric thiolase enzyme complex from S. macaca involved in the biosynthesis of the antimicrobial compound reutericyclin A. This secondary metabolic pathway is proposed to have evolved from the central metabolic pathway for mevalonate biosynthesis in archaea by functionally repurposing the tripartite enzyme complex that couples 3-hydroxy-3-methylglutaryl-CoA synthase (HMGS) and acetyl-CoA acetyltransferase (ACAT) reactivity. Biochemical characterization of SmaATase revealed the HMGS activity of the proposed archaeal ancestor was lost, and instead of Claisen condensation activity the ACAT homolog catalyzes a non-canonical Friedel-Crafts C-acetylation on a pyrrolidine-2,4-dione scaffold. This is similar to another bacterial tripartite enzyme complex, PpATase from P. protogens, that performs a Friedel-Crafts C-acylation of monoacetylphloroglucinol (MAPG) to diacetylphloroglucinol (DAPG, an antimicrobial compound) and phloroglucinol (PG). Notably, although the HMGS activity of the archaeal ancestor is not preserved in SmaATase or PpATase, the authors were only able to observe the C-acetylation activity in the presence of the complete tripartite complex, suggesting the evolutionary trajectory may have prioritized chemical specialization over quaternary structural modifications. This point is further illustrated by comparison of experimental structures, including new cryo-EM results of SmaATase, that show strong structural similarities in the overall subunit organization and Co-A binding site, and all complexes contain the canonical thiolase ⍺β⍺β⍺ fold. Multiple sequence alignment reveals SmaATase and PpATase retain some, but not all, sequence motifs characteristic of ACATs, and they lack a conserved sequence essential for HGMS activity. Biochemical activity assays of wild type and generated mutants shed light on the functional roles of various sequence motifs. Finally, they find that the acetylation modification to form RTC-A and DAPG is critical for antimicrobial potency against competitors, implicating the vital importance of these bacterial ATases. The results have broad importance across fields of biosynthesis and bioengineering, and I recommend for publication after addressing the following issues/comments that would improve this manuscript primarily in terms of clarity, discussion depth, and reproducibility.\nMajor Comments\n1. Docking studies are performed, but interpretation and presentation of the results are lacking. No data/results are presented besides SI Fig. 23 which does not provide substantial information. Line 237 states “Docking results showed that conserved C87 and H349 may coordinate catalysis, while H143, Y300, and Q302 support substrate positioning”. How? C87 and H349 are not depicted in SI Fig. 23a, and H143, Y300, and Q302 are only depicted as “contact residues”. Are there truly no more hydrogen bonds to the ligand besides with Tyr300 (SI Fig. 23a)? Do the docking results show many or few binding conformations? This manuscript could be greatly benefit from an explanation of the reaction mechanism catalyzed by PpATase (steps detailed in the manuscript Ref. 25) and based on that the authors can discuss the likely residues of the SmaATase active site that are involved in the reaction. The differences between these two active sites, and their deviation from archaeal ACAT discussed from the mechanistic perspective would certainly enhance the manuscript’s argument of functional repurposing.\n2. MD simulations are performed, but interpretation and presentation of the results are lacking. SI Fig 23b is not discussed anywhere. In particular, the per-residue RMSF appears to show that many active site residues have high fluctuations. Typically, when a binder is present we might expect active site rigidity, especially if the molecule is stably bound.\n3. Simulations were performed with only MucB, not the full complex, which was shown to be required for activity. Did the authors perform any binding kinetics with only MucB, or have reason to believe MucB can effectively bind the substrate, and in the same way it would if the complex was formed? It is interesting that the MM/GBSA binding free energy calculations show favorable binding to MucB, but experimental results (line 166) “suggest that while MucA and MucC may not directly participate in catalysis, they are essential for proper folding and assembly of the tripartite complex, thereby enabling its enzymatic activity”. Some discussion about the implications of these two results should be included to strengthen the manuscript.\n4. The energy minimized coordinates of the ligand used in docking should be provided for reproducibility, as well as at least one set of xyz coordinates from the docking/simulation results of the holo enzyme.\n5. The assigned protonation states used for simulations should be stated in the SI for the method to be transparent and reproducible. Additionally, the protonation state of residues in the binding site are likely significant. Some justification of choice should be provided, or evaluation of various possible states to ensure results are robust, but since these results are not critical for the manuscripts main objectives additional calculations could be avoided.\nMinor Comments\n1. DUF35 is not defined.\n2. Line 91: Why was the MucABC sequence from S. mutans was used for the query instead of S. macaca? Since the paper focuses on the S. macaca complex it would be more consistent to use those results and likely does not alter any discussion.\n3. Lines 128-129: Results for acetyl-CoA as a donor are stated. The reader would benefit from the results of the other physiologically relevant donors also being quickly stated.\n4. Lines 134-135: “Together, these results support acetyl-CoA as the physiological acetyl donor…”. This may be too strong of a statement. First, it appears there is some, albeit very little, product formation with donors acetoacetyl-CoA and malonyl-CoA (Fig. 1b). Second, are the donors evaluated the only physiological possibilities? Third, a previous study found no product with PpATase and acetyl-CoA. Either more justification should be included in the text to convince the reader that acetyl-CoA is THE donor, or the statement can be modified. Additionally, some discussion on why product was found using PpATase with acetyl-CoA when a previous study found this did not work is warranted.\n5. “r.m.s.d” should be RMSD throughout manuscript.\n6. Line 179: The use of the word “mirrors” here could cause confusion as it implies chiral partners. “Resembles” would be a safer choice.\n7. Line 192: Replace “substitutions” with “modifications” or similar, since the differences are more signification than single AA substitutions.\n8. Line 218: “While no other function predictable…” This sentence was awkward to understand- could be written more clearly.\n9. Line 239: E124 and S142 are not hydrophobic residues.\n10. Line 240: Reference needed for W211 in Ph1C proposed to act as conformational lid.\n11. Line 275: The reactions of SmaATase and PpATase are not “single-step” (particularly because they involve acylation and deacylation phases that are likely both multi-step reactions). Do the authors mean single reactions, as opposed to coupled reactions?\n12. Line 435: What pH was assumed for protonation state assignment?\n13. Line 435: Is “hydrogenated” an appropriate term here? Do the authors mean they modeled in unresolved H atoms to the structure?\n14. Line 440: Can the authors provide more information beyond “Phad Calculation Co., Ltd”? I tried searching for this company and could not find any results.\n15. Line 442: ff19SB needs reference and AMBER Tools needs reference.\n16. Line 443: How were the atomic charges derived for GAFF parameters of the ligand?\n17. Line 448: Says SHAKE algorithm was used in heating and equilibration- but what about during the production runs?\n18. Line 451: PME needs reference.\n19. Line 452: MM/GBSA needs reference.\n20. Line 617: “DUF35 unites highlighted in cartoon format”- it looks like more than just DUF35 is in cartoon… are the captions for d and e switched?\n21. Fig. 5:\na. Is the PpATase Cys88 acylated? This should be stated somewhere, as well as where this structure came from.\nb. The ACAT represented after step2 for both ATases has a H x-ed out, but this is a bit confusing because the actives sites of both have numerous histidines.\nc. Why are some H-bonds listed in this SmaATase active site that are not shown in SI Fig. 23?\nd. It should be clarified somewhere that the H-bonds are measure from the heavy atoms (at least they appear to be), or those H atoms should be included in the representation and the true H-bond labeled.\n22. Fig. S16: Typo “ligntblue”.\n23. Fig. S23: Text of b, c, d, e, f too small. Why is Tyr300 in ball and stick representation? It makes it seem like it is also a ligand and is confusing to the reader. See comment 21 about H-bonds. Cys88 and His349 should be included.\nRespectfully,\nHeidi Klem", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response to Reviewers' Comments to Author:\nReviewer #1 (Remarks to the Author)\nComments:\nIn their manuscript, Liao et al study bacterial tripartite complex systems analogous to the\nthiolase/HMG-CoA synthase/DUF-35 protein used by archaea to catalyse the first steps of\nisoprenoid synthesis. Through an impressive panel of methods ranging from enzymology,\nstructural biology, biochemistry, bioinformatics, phylogeny, and even genome mining, they\nfound that the thiolase subunit from the bacterial homologues from Streptomyces macacae\n(SmaATase) and from Pseudomonas protegens (PpATase) catalyse a rare Friedel-Crafts\nreaction involving acetyl-CoA as acetyl donor. The end products of these reactions\nultimately generate molecules with antibiotic activities. It is without any doubt a study of\nhigh quality, bringing to light an enzymatic complex involved in secondary metabolism that\nmost probably originates from a central carbon metabolism involved in isoprenoid\nbiosynthesis. The reviewer congratulates the authors on this excellent study that reaches\nthe standard of Nature Communications, and would strongly support its publication upon\naddressing these minor comments.\n1- Abstract: “Cryo-electron microscopy of the S. macaca-derived thiolase complex”.\nShouldn´t it be S. macacae instead of S. macaca?\nResponse:\nWe have corrected the species name to S. macacae in the abstract.\n2- Title of the result first subsection. “Characterization of Streptococcal ATases as non-\ncanonical Friedel-Crafts acylases”. For clarification, it would be helpful to state the meaning\nof “ATases” here or before instead of using the abbreviation. Moreover, since the\nphysiological donor is acetyl-CoA, authors should consider using the term acetylases\ninstead of acylases.\nResponse:\nClarification of the ATase as acyltransferase was added in the title of first result subsection:\n“Characterization of Streptococcal acyltransferases (ATases) as non-canonical\nFriedel-Crafts acylases”.\nWe agree that “acetylase” is a more precise term when considering that acetyl-CoA is the\nprimary physiological donor. However, our enzyme assays demonstrate that SmaATase\nexhibits activity with acyl donors of up to C8 chain length (as detailed in the attached file\nfor reviewers), indicating its catalytic scope extends beyond acetylation. Therefore, we\nhave chosen to use the broader and established term “acylase”, which encompasses\nacetylase and is consistent with the nomenclature used for the related enzyme PpATase\n(ChemBioChem 2019, 20, 88–95).\n3- Legend of Supplementary Fig. 2c. “SDS-PAGE analysis of the heterologous expression\nand purification results of SmaATase, SmuATase, and StrATase, respectively. SmaATase,\nSmuATase, and StrATase represent MucABC enzyme complex from S. mutans, S.\nmacacae, S. troglodytae, respectively.”\nBased on the text and results, it should be: “SDS-PAGE analysis of the heterologous\nexpression and purification results of SmaATase, SmuATase, and StrATase, respectively.\nSmaATase, SmuATase, and StrATase represent MucABC enzyme complex from S.\nmacacae, S. mutans, S. troglodytae, respectively.”\nResponse:\nThe correction has been made as suggested.\n4- 2nd paragraph results: “Additionally, MucABC-mediated reverse conversion of RTC-A\nback to preRTC-A was observed, confirming the bidirectionality of the reaction\n(Supplementary Fig. 6d).” Would it be possible to provide the Standard Gibbs free energy\nfor the reaction? That would provide some insights on its expected directionality.\nResponse:\nWe thank the reviewer for raising this point regarding the reaction’s bidirectionality. Our\nfollow-up investigations indicate that determining a ΔG°' value is not feasible because the\nreverse reaction is kinetically blocked by the enzyme’s substrate specificity.\nIn the forward direction, SmaATase accepts acetyl-CoA or phenyl acetate (PA) as acetyl\ndonors and transfers the acetyl group to preRTC-A. In contrast, the enzyme cannot\nefficiently use CoA or phenol as acetyl acceptors in the reverse direction. Incubation of\nSmaATase with RTC-A and either CoA or phenol did not yield more preRTC-A than the\ncontrol (see new Supplementary Fig. 7a).\nThe “reverse” activity we observed is in fact a transacetylation reaction. As demonstrated\nin revised Supplementary Fig. 7b and 7c, when provided with a suitable structural analog\n(preRTC-C2) as an acetyl acceptor, SmaATase catalyzes acetyl group transfer from RTC-\nA to form preRTC-A and RTC-C2. This demonstrates that the enzyme can cleave the C–C\nbond in RTC-A, thereby enabling it to function as an acetyl donor. A similar capability has\nbeen reported for PpATase when using MAPG and DAPG as acetyl donors.\nAccordingly, we have revised the statement to: “Additionally, MucABC-mediated\nconversion of RTC-A to preRTC-A was observed, indicating that the C–C bond\nformed by MucABC is cleavable by the enzyme, thereby enabling RTC-A to serve as\nan acetyl donor in a transacetylation reaction (Supplementary Fig. 7).”\n5- 3rd paragraph results: “Notably, earlier studies reported no detectable product formation\nwhen PpATase was incubated with acetyl-CoA”. By curiosity, do the authors have an\nexplanation for why the activity was not observable before with acetyl-CoA?\nResponse:\nWe thank the reviewer for their curiosity regarding this discrepancy. We agree that\nproviding a possible explanation is important.\nIn the first study (Biosci. Biotech. Bioch. 2012, 76: 559), enzyme activity was monitored\nspectrophotometrically at 370 nm—an indirect and relatively insensitive approach that may\nnot detect low turnover rates. In the later study (ChemCatChem 2019, 11: 1064), HPLC\ndetection was performed at 254 nm. Our data show that this wavelength is suboptimal for\nspecifically detecting MAPG and DAPG, whereas our UPLC-based assay at 280 nm\nprovides a substantially stronger and more selective signal (see the figure below).\nAdditionally, the HMGS/ACAT/DUF35 complex is known to be unstable during storage; for\nexample, the archaeal homolog retains only ~15% activity after two weeks at –80 °C\n(PNAS 2018, 115: 3380). The use of suboptimally preserved enzyme or insufficiently\nsensitive detection methods in previous studies could therefore have resulted in a false-\nnegative outcome.\nTo avoid over-interpreting these methodological differences and to keep the focus on our\nfindings, we have removed the comparative statement about earlier studies from the\nrevised manuscript (see below).\n'##\n#\n! \"# \"$ \"% \"&\n'##\n#\n! \"# \"$ \"% \"&\nAs illustrated in the figure, the absorbance maxima of both MAPG and DAPG occur at 280\nnm; detection at this wavelength is therefore optimal.\n6- Supplementary Fig. 9. Please define “Cald.” in the legend. Could the authors explain the\ndifferences between observed and calculated masses for MucB and MucA (e.g., 44481 Da\nversus 44351 Da)?\nResponse:\nWe have now defined “Cald.” as “calculated” in the legend of new Supplementary Figure\n10.\nRegarding the discrepancy between the observed and calculated masses, this difference\narises because the N-terminal methionine is post-translationally removed during\nexpression in E. coli, a common phenomenon for many recombinant proteins (Mol. Cell.\nProteomics 2006, 5:2336). When the mass is calculated for the mature protein lacking the\ninitiator methionine, the value matches the experimentally observed mass. In the revised\nfigures, we have therefore labeled the calculated masses corresponding to M -Met and\nMucA\nM -Met.\nMucB\nWe sincerely apologize for an error we made in the original figure, in which the calculated\nand experimental mass values for MucA and MucB were inadvertently swapped. We have\ncorrected this mistake in the revised version. We have also thoroughly re-checked the\nentire manuscript to ensure that no similar errors remain.\n7- Supplementary Fig. 10. Has the mask used to generate the local refinement been\ndeposited in the EMDB? Adding an extra panel with the mask on Fig. S10 would be\ninformative and provide more transparency on the model-building process. Moreover, for\nFigures S10 and S11, is the map based on raw data or a sharpened one? This should be\nstated in the legend.\nResponse:\nWe thank the reviewer for these suggestions on adding details about data processing and\nmodel building.\nIn our cryo-EM processing pipeline, local refinement was performed using the mask\nautomatically generated during the non-uniform refinement procedure in CryoSPARC. This\nmask was not manually edited or modified. Following the standard practice for this method,\nthe primary output deposited in the EMDB is the final sharpened density map, which\nreflects the results of the complete refinement process. The internal mask used by\nCryoSPARC during automated refinement is not typically deposited as a separate file.\nIn response to the reviewer’s request, we have added a new supplementary Figure\n(Supplementary Figure 13) that displays a transparent surface representation of the map\noverlaid on the atomic model.\nWe also clarify that all cryo-EM maps shown in the manuscript and supplementary figures\nare sharpened maps, which were used directly for model building and analysis. This\ninformation has now been explicitly stated in the revised figure legends: “The cryo-EM\ndensity map shown is the sharpened map.”\n8- Figure 2. According to the legend, panels d and e must be swapped.\nResponse:\nWe have swapped the captions for (d) and (e) in the figure legend to ensure they now\naccurately correspond to the correct images in Figure 2.\n9- 4th paragraph results: “In addition to the dodecamer, we observed an alternative\noctameric assembly (2.16 Å resolution), consisting of a MucA4C4 core”. The nomenclature\nseems wrong and should be MucA4C2 according to the deposited model.\nResponse:\nWe thank the reviewer for identifying this error. The nomenclature has been corrected to\nMucA C core in the revised manuscript.\n4 2\n10- 4th paragraph results: “Structural superposition of the dodecameric and octameric\nforms revealed a root-mean-square deviations (r.m.s.d.) of 0.173 Å”. Please add the\nnumber of superposed Calpha.\nResponse:\nThe sentence has been updated to reflect the RMSD value (0.160 Å) obtained from the re-\npolished structures and to include the number of superposed Cα atoms (2254): “Structural\nsuperposition of the dodecameric and octameric forms revealed a root-mean-square\ndeviations (RMSD) of 0.160 Å over 2254 superposed Cα atoms”.\nAll similar descriptions throughout the manuscript have been revised accordingly for\nconsistency.\n11- 4th paragraph results: “…indicating that SmaATase exists as a heterogeneous mixture\nof assemblies under near-physiological conditions.”. This is a very dangerous statement\nthat must be reformulated because the tested conditions in vitro are expected to be very\ndifferent compared to the physiological ones (molecular crowding, physicochemical\nconditions, protein concentrations…). Rather, use “under in vitro conditions” or “in\nsolution”.\nResponse:\nWe fully agree that our original wording overstated the physiological relevance of the in\nvitro observations. The sentence has been revised to accurately reflect the experimental\ncontext: “...indicating that SmaATase exists as a heterogeneous mixture of\nassemblies under the tested in vitro conditions”.\n12- 7th paragraph results: “Structural alignment of the HMGS-ACAT-DUF35 modules\nbetween SmaATase and either PpATase or the archaeal homolog yields r.m.s.d. of 1.245\nÅ and 1.466 Å,” Please add the number of superposed Calpha.\nResponse:\nThe sentence has been revised to include the number of superposed Cα atoms:\n“Structural alignment of the HMGS-ACAT-DUF35 modules between SmaATase and\neither PpATase or the archaeal homolog yields r.m.s.d. of 1.245 Å (585 Cα atoms)\nand 1.454 Å (363 Cα atoms), respectively”.\n13- 7th paragraph results: “DUF35 subunit architecture and positioning: MucC, a family\nmember of DUF35 protein, features an N-terminal rubredoxin-like zinc ribbon and a C-\nterminal oligonucleotide/oligosaccharide-binding (OB) fold domain.”. IMPORTANT POINT.\nUnfortunately, MucC has been modelled without the 4 conserved Cys binding the Zn. The\nZn is observable in the EM map together with the four Cys binding to it. See the attached\npicture.\nZn has been modelled in all structural homologues, and must be modelled in both\nstructures from this work. Presenting an additional supplementary figure with a close-up of\nthe Zn binding site and the electron density would be informative.\nMoreover, Cys87 in ACAT subunit of SmaATase could be acetylated. Mass spectrometry\nmight provide some clues if this residue is truly acetylated in the protein in solution (e.g.,\nwith the acetyl-group coming from acetyl-CoA from E. coli). Based on the electron density,\nthere is high suspicion that Cys87 is indeed acetylated. Therefore, the referee suggests\nadding an acetyl group on this cysteine (as it has been done for PpATase structure), and if\npossible, obtain a confirmation by mass spectrometry. Presenting an additional\nsupplementary figure with a close-up of the acetylated Cys87 and the electron density\nwould be informative. If mass spectrometry, or author´s conclusions point to a non-\nacetylated form, please add a snapshot of the Cys87 with the surrounding electron density\nfor each chain.\nTo conclude on the model, the referee would suggest adding the Zn, the acetyl group and\npolishing the model further for a better fit in the electron density. For instance, some\nloops and sidechains are observable in the electron density, but were not modelled (see\nbelow). Please correct the residue presenting a very high clash score (higher than 0.8).\nResponse:\nWe are deeply grateful to the reviewer for these exceptionally careful and insightful\ncomments regarding the structural models.\nThe reviewer is absolutely correct that the Zn²⁺ ion in the DUF35 (MucC) subunit\nshould be modeled. We have now incorporated the zinc ion coordinated by the four\nconserved cysteine residues in both structures. A new supplementary figure\n(Supplementary Figure 20) has been added to provide a close-up view of the zinc-binding\nsite alongside with the corresponding density.\nRegarding the putative acetylation of Cys87 in the ACAT (MucB) subunit, we agree\nwith the reviewer that the observed electron density is strongly suggestive of an acetyl\nmodification, consistent with observations reported for related complexes (ChemBioChem\n2019, 20: 88; PNAS 2018, 115: 3380). To further validate this, we performed mass\nspectrometric analysis on the purified SmaATase complex. The results unambiguously\nidentified peptides containing an acetylated Cys87, confirming that this modification is\npresent in solution. We have therefore added the acetyl group to Cys87 in the refined\nstructural model. A new supplementary figure (Supplementary Figure 29) has been\nincluded to show a close-up view of the acetylated Cys87 alongside with the corresponding\nelectron density. Variability in the density among different chains likely reflects partial\nacetylation, which is consistent with our MS results.\nIn addition, a thorough model polishing was performed to ensure an optimal fit to the\nelectron density. All structural figures throughout the manuscript have been updated to\nreflect the final refined models. We believe these revisions have substantially strengthened\nthe accuracy, completeness, and reliability of our structural interpretations, and we are\nsincerely grateful to the reviewer for the detailed guidance.\n14- Supplementary Fig. 17. Presented residues should be labelled. A second snapshot\nwith a 90° orientation could be useful to show the different residues. Also, hydrogen bond\ndistances are impossible to read. Please remove the distances label, keep the dashes (in\nanother colour than yellow) and add a general statement such as: all hydrogen bond\ndistances below 3.5 Å are displayed as dashes.\nResponse:\nThe figure (now is Supplementary Fig. 21) has been corrected as suggested.\n15- 7th paragraph results: “). Additionally, a hallmark of the archaeal HMGS/ACAT/DUF35\ncomplex is the presence of a fused CoA-binding site at the HMGS–ACAT interface, which\nfacilitates shuttling of the pantetheinyl arm between active sites during catalysis (Fig. 2f).”\nA supplementary figure comparing the CoA binding site between the archaeal\nACAT/HMGS/DUF-35 protein (PDB 6ESQ) and SmaATase would reinforce the observation\nmade on Fig. 2f. Are most of the residues conserved? Also, please correct the typo on Fig.\n2f “fused Co-A binding” should be “fused CoA-binding”.\nResponse:\nWe agree with the reviewer that a detailed structural comparison of the CoA-binding sites\nwould strengthen the interpretation presented Fig. 2f.\nThe previous study on the archaeal complex (PNAS 2018, 115: 3380) revealed the novel\narchitectural principle of a fused CoA-binding site that couples the ACAT (thiolase)\nand HMGS reactions (see below Fig. i). Although the study illustrated a few candidate\nresidues (C206, K332 in ACAT; R203, K243, K247 in HMGS) proximal to CoA in a figure\n(see below Fig. ii) through comparison with the CoA-substrate binding sites of lone-standing\nthiolase and HMGS, their specific functional roles were not experimentally validated and\nwere not discussed in detail in the publication.\nOur residue-level comparison indicates that while some residues (e.g., K332, R203, K243)\nare conserved in archaea, others (such as C206 and K247) are not (see below Fig. iii).\nThe corresponding positions in SmaATase also differ, suggesting that although the spatial\narchitecture of the binding pocket is conserved, the exact residue composition is not\nidentical between the two complexes.\nTo provide a clear and objective visualization of these similarities and differences, we have\nadded a new supplementary figure (Supplementary Fig. 23) showing a side-by-side\ncomparison of the CoA-binding interface in SmaATase and the archaeal homolog. This\nfigure highlights both the conserved structural arrangement and the variations in residue\nidentity.\nFinally, as suggested, the typo in Fig. 2f has been corrected from “fused Co-A binding” to\n“fused CoA-binding”.\n16- 11th paragraph results: “Introducing an H294Y mutation in PfACAT nearly abolished\nthis activity (Fig. 4c and Supplementary Fig. 20), confirming the essential catalytic role of\nthe His residue within the HDxF motif.”. This is a very interesting point. Are the authors\nexpecting a role in antibiotic biosynthesis for this enzyme? It is unlikely that its physiological\nfunction would be simply the formation of acetyl-CoA from acetoacetyl-CoA. The latter\nmight simply be a side reaction.\nResponse:\nWe thank the reviewer for raising this intriguing question regarding the physiological\nfunction of the PfACAT/HMGS-DUF35 complex.\nWe agree that its in vivo function is unlikely to be simply the thiolytic cleavage of\nacetoacetyl-CoA. Instead, we propose that the thiolase activity observed in vitro may\nrepresent a residual biochemical capability of the ACAT domain, rather than its primary\nphysiological role.\nAlthough PfACAT/HMGS-DUF35 retains the ancestral HMGS/ACAT/DUF35 architectural\nframework, key mutations in catalytic tetrad E-C-H-N have inactivated the HMGS domain.\nThis loss of HMGS activity disrupts the canonical function of the complex in the mevalonate\npathway. The reaction catalyzed by ACAT (2×Acetyl-CoA ⇌ Acetoacetyl-CoA + CoA) is\nhighly endergonic (ΔG°' = +26.1 kJ/mol) with a small equilibrium constant [Keq = (1.1 ±\n0.2)×10-5] (PNAS 2018, 115: 3380), strongly favoring the thiolytic direction. Thus, in the\nabsence of a functional HMGS domain capable of consuming acetoacetyl-CoA, the\nreaction defaults toward thiolysis—consistent with what we observed in vitro.\nGenomic context also argues against a role in classical antibiotic biosynthesis. Unlike\nPpATase or SmaATase, the PfACAT/HMGS–DUF35 locus lacks adjacent PKS, NRPS, or\nhybrid biosynthetic genes typically associated with HMGS–ACAT–DUF35 complexes\nengaged in secondary metabolism. This genomic isolation makes its direct participation in\ncanonical natural product biosynthesis unlikely. While the available data do not support\nrecruitment of PfACAT/HMGS–DUF35 into a known antibiotic biosynthetic pathway, we\ncannot exclude the possibility that it participates in an uncharacterized specialized\nmetabolic process that operates independently of PKS/NRPS systems.\nIn conclusion, we propose that PfACAT/HMGS–DUF35 represents an evolutionary\nintermediate: a complex that has diverged from its primary metabolic role but has not been\nco-opted into a specialized biosynthetic pathway like its bacterial homologs. We have\nrevised the manuscript to incorporate this evolutionary interpretation.\n17- Figure 5: Cys88 is acetylated, please label the residue accordingly (e.g., Acetyl-C88).\nD137 cited in the text is not shown on the figure. Importantly, the difference in the active\nsite pose confuses the reader as it seems that the orientation between PpATase and\nSmaATase is the same, but the position of the residues (particularly C88/H347 and\nC87/H349) is different. Please take a new picture with the exact same orientation between\nPpATase and SmaATase active sites. Highlight the key residues C88/H347 and C87/H349\nwith a bold label.\nResponse:\nWe thank the reviewer for these precise suggestions to improve the clarity of the figure. All\nsuggested changes have been implemented in the revised Figure 5.\n18- Supplementary Fig. 23: Why is the hydrogen bond not shown for Q302? Is there a\npossibility to switch the H143 rotamer to gain a hydrogen bond with the ligand without\ngenerating steric clashes? Here, 4.1 A seems too distant for efficient\nResponse:\nThe discrepancy in hydrogen bond depiction between Fig. 5 and the original\nSupplementary Fig. 23 was due to differences in the visualization algorithms used by the\ntwo software packages.\nTo establish a more reliable substrate-binding model, we re-performed the molecular\ndocking and dynamics simulations using an improved system. Specifically, we employed\nthe complete MucA-B-C complex bound to the native substrate preRTC-A, rather than the\npreviously used standalone MucB subunit with a truncated analog. The updated results\ndemonstrate that H143 mediates hydrophobic contacts with the substrate rather than\nforming a direct hydrogen bond (Fig. 5); the overall binding of preRTC-A is stabilized by\nhydrogen bonds with Y300 and H349 (Fig. 5), supplemented by hydrophobic and van der\nWaals interactions with other surrounding residues (revised Supplementary Fig. 27).\nTo present these findings clearly, hydrogen bonds are depicted in the 3D structural\noverview (Fig. 5), and we now provide a dedicated 2D ligand–interaction diagram in\nrevised Supplementary Fig. 27 summarizing all hydrophobic and van der Waals contacts.\n19- Supplementary Fig. 24: The final concentration of the used compounds must be stated\nin the figure or legend.\nResponse:\nThe final concentration of the tested compounds (400 μg/mL) has been added to the\nlegend of revised Supplementary Fig. 30.\n20- Last paragraph of the result section: “Although preRTC-A showed modest activity\nagainst S. gordonii ATCC 10558 (MIC = 32 μg/mL), RTC-A demonstrated a 32-fold\nenhancement in potency (MIC = 1 μg/mL; Fig. 6). A similar trend was observed for the\nphenolic polyketide DAPG and its biosynthetic precursor PG. DAPG exhibited significantly\nhigher antimicrobial activity against most tested strains (except S. oralis DA1241 and S.\ngordonii ATCC 10558) compared to PG, recapitulating the structure–activity relationship\nobserved for RTC-A and preRTC-A pair (Fig. 6 and Supplementary Fig. 24).”. The whole\nsentence should be clarified. The results obtained are excellent, and authors should rather\nuse a general statement to describe the results, for instance: All strains showed inhibition\nfor DAPG compared to PG, except S. gordonii and S. oralis. The effect is even more\npronounced for preRTC-A versus RTC-A, where all tested strains showed inhibition for\nRTA-A compared to preRTC-A.”\nResponse:\nWe thank the reviewer for this helpful suggestion. The text has been revised for clarity and\nconciseness (see below).\n21- 1st paragraph discussion: “In contrast, SmaATase is remarkable for its ability to\ncatalyze Friedel–Crafts acylation independently of CoA or ACP-activated acyl donors,\nhighlighting its potential as a versatile biocatalyst for synthetic applications.” The\nacetylation is CoA-dependent. Please rephrase.\nResponse:\nWe thank the reviewer for this important clarification. We agree that, in vivo, acetyl-CoA is\nthe physiological acetyl donor for SmaATase and therefore the native acetylation reaction\nis CoA-dependent.\nOur intention in the original sentence was to emphasize a biochemical capability of\nSmaATase revealed by our in vitro assays that SmaATase is not strictly restricted to acetyl-\nCoA as an acetyl donor. As shown in Fig. 1b, the enzyme can utilize several alternative\ndonors, including N-acetylimidazole, isopropenyl acetate, ethyl thioacetate, and phenyl\nacetate. This donor promiscuity distinguishes SmaATase from canonical acyltransferases\nthat rely exclusively on CoA or ACP-activated acyl donors.\nTo avoid ambiguity and better reflect this point, we have reivsed the sentence to: “In\ncontrast, SmaATase exhibits remarkable promiscuity toward acetyl donors,\ncatalyzing Friedel–Crafts acylation with not only the canonical donor acetyl-CoA but\nalso with unnatural donors such as phenyl acetate.”\n22- 2nd paragraph discussion: “…enzyme complexes in secondary metabolism are more\ncommonly associated with scaffold formation (such as PKS37 and NRPS38) than with\nstructural modification”. What is the meaning of structural modification here? Please\nreformulate.\nResponse:\nWe thank the reviewer for pointing out the ambiguity. In the original sentence, “structural\nmodification” referred to post-assembly tailoring reactions in secondary metabolism—such\nas methylation, prenylation, hydroxylation, and glycosylation—that decorate or remodel the\ncore scaffold after its initial construction.\nOur intention was to contrast the quaternary structure complexity of enzyme complexes\ninvolved in scaffold assembly (e.g., PKSs and NRPSs, which are large, multi-domain\nmegasynthases) with the generally simpler architecture of enzymes responsible for\ntailoring modifications, which are typically monomeric, dimeric, or form only small\noligomers. In this context, the discovery of SmaATase as a hetero-oligomeric complex\ndedicated to an acetylation tailoring reaction is therefore noteworthy.\nTo remove ambiguity, we have reformulated the sentence in the manuscript to: “... enzyme\ncomplexes in secondary metabolism are more commonly associated with the\nprimary assembly of carbon skeletons scaffold formation (such as PKS37 and\nNRPS38) than with the post-assembly tailoring reactions (e.g., methylation,\nprenylation, hydroxylation).”\n23- Supplementary Fig. 25: It would help the reader to add the acetylated cysteine residue\nposition in the three studied systems (in the figure or in the legend).\nResponse:\nWe thank the reviewer for this suggestion. The positions of the acetylated cysteine residues\nin the three studied systems have been added to the figure (now is Supplementary Fig.\n31).\n24- Methods, protein expression and purification: “and imaged using the DelDoc Go\nsystem (Bio-Rad).”. It should be GelDoc Go system.\nResponse:\nDone.\n25- Methods, Cryo-EM structural determination: Please provide the following information.\nProtein concentration and buffer used for the grid.\nHow did authors build, refine, and validate the model?\n“Ab-initio reconstruction of these particles produced an octameric structure at 2.16 Å\nresolution.” Ab initio is not a refinement technique, so it does not make sense to associate\nab initio with resolution. Please reformulate.\nResponse:\nAs requested, the methods section has been updated to include the protein concentration\n(3.8 mg/mL) and buffer composition (50 mM potassium phosphate, pH 7.5) used for grid\npreparation.\nWe have expanded the description of the model building, refinement, and validation\nprocedures to provide a more comprehensive account of the workflow.\nBesides, the sentence “Ab-initio reconstruction of these particles produced an octameric\nstructure at 2.16 Å resolution.” has been revised to: “Ab-initio reconstruction of these\nparticles produced an octameric structure, and the resolution of this structure\nreaches 2.16 Å after non-uniform refinement.”\nReviewer #2 (Remarks to the Author):\nXiaoyu Tang and colleagues present a manuscript detailing the biochemical\ncharacterization of a tripartite enzymatic complex catalyzing Friedel-Crafts-Acylation in the\nbiosynthesis of the antibiotic tetramate reutericyclin A. This enzymatic complex bears\nsimilarities to a previously characterized system able to catalyze acetylation of\nphloroglucinol. The protein complex was structurally characterized via Cryo-EM, revealing\na dodecameric structure, analogous to a previously characterized pathway in archaeal\nmevalonate biosynthesis. Bioinformatic analyses showed that key catalytic residues of the\nprimary metabolism enzyme were lost, and new, potentially critical residues for a new\ncatalytic functon(acetylation) acquired. Based on these findings, the authors pose the\nhypothesis that the enzymatic complex was repurposed during evolution to acquire a new\ncatalytic function. Finally, the authors show that acetylation is an essential feature for\nantibiotic activity in both reutericyclins and phloroglucinols, highlighting the biological\nimportance of this modification.\nThe paper is very well and clearly written and contains a lot of data. The experiments were\ncarried out with a range of state-of the-art methodologies, and the results are nicely\npresented in main and SI Figures. Overall, this study deserves publication in Nature\nCommunications. However, there are a couple of issues that should be improved or taken\ncare of before publication:\nMajor:\nl.265: SmaATase is praised having potential as a “versatile biocatalyst for synthetic\napplications.” Indeed, the possible use of different acetyl donors has been demonstrated,\nbut the substrate scope has not been investigated at all to my understanding, except for\nthe acetylation of the native substrate pre-RTC and its short chain analogue. Of course, a\ncomprehensive enzymatic study is beyond the scope of this paper, but it would be nice to\nprovide some initial characterization: e.g. on simplified pyrrolidones with a few different\nsubstituents. Also, I wonder, if propionyl chains could be accepted?\nResponse:\nWe sincerely thank the reviewer for their thoughtful comments regarding the biocatalytic\npotential of SmaATase.\nOur description of SmaATase as a “versatile biocatalyst” was motivated by two\nobservations: (1) its ability to catalyze Friedel–Crafts C-acylation, a reaction scarcely\nobserved in enzymology; and (2) its capacity to utilize simple and inexpensive acetyl\ndonors, including phenyl acetate, in contrast to most acyltransferases that rely strictly on\nCoA-activated donors.\nAdditionally, compared with PpATase—the first reported Friedel-Crafts acylase, which\ndisplay strict to resorcinol-containing substrates—SmaATase accepts preRTC-A, a\nchemically distinct pyrrolidine-2,4-dione scaffold. This distinction suggests that the two\nenzymes may offer complementary opportunities for biocatalysis.\nWe fully agree with the reviewer that a systematically evaluation of substrate scope would\nbe essential for defining the enzyme’s synthetic utility. In our preliminary assays,\nSmaATase showed notable flexibility: (a) it accepts preRTC-A analogs with varying N-acyl\nchain lengths (C2–C8) or different amino-acid-derived side chains as acceptors; (b) it also\nutilizes phenyl esters with acyl chains up to C8 as donors. These data have been provided\nto the reviewers in the attached file.\nHowever, conducting a comprehensive substrate-scope study is beyond the scope of the\ncurrent work, which focuses primarily on the evolutionary repurposing of the thiolase\ncomplex. To avoid overstating the biocatalytic implications, we have removed the phrase\n“versatile biocatalyst for synthetic applications” from the revised manuscript, while\nmaintaining a factual description of the enzyme’s observed donor flexibility.\nFigure 3b: The loss of key catalytic residues of primary metabolic enzymes has been\nshown, but I am missing a Figure, where the acquisition of key residues for the new\ncatalytic function(see Figure 5, Fig. S23) is analyzed. It would be good to provide a similar\nFigure and underlying alignments for these residues in primary vs. secondary metabolism.\nDo all 31 enzymes contain the same sequence motifs in this region or are there variations?\nResponse:\nWe thank the reviewer for this excellent suggestion. We agree that examining the\nacquisition of key residues—in addition to the loss of ancestral primary-metabolism\nresidues—provides a more complete understanding of the functional repurposing process.\nIn response, we conducted a new comparative sequence analysis focusing on the active-\nsite residues that may contribute to the evolved acylation activity and substrate specificity.\nThe analysis highlights both the conserved catalytic machinery and the divergent\nsubstrate-binding features that underlie functional specialization (Supplementary Figure\n28, and we also included this figure on below).\nCatalytically essential residues in PpATase map directly onto those in SmaATase: the\nnucleophile C88 (PhlC) corresponds to C87 (MucB); the general base E116 corresponds\nto D115; and the proton-relay network D137-T132-H144 corresponds to D136-T131-H143.\nThis conservation supports a shared catalytic mechanism between the two bacterial\nenzymes that differs from the archaeal ACAT/HMGS complex.\nBy contrast, the substrate-binding pocket shows clear reconfiguration. Key PhlC substrate-\nbinding residues (H56, Y124, W211, Y298, H347) are either conserved (Y300, H349) or\nreplaced (A55, F123, A210) in MucB, whereas residues crucial for preRTC-A binding in\nMucB (N86, M127, I144, Y300, H349) are conserved (N87, Y298, H347) or substituted\n(I128, T145) in PhlC. These differences illustrate how the binding pocket was remodeled,\nenabling SmaATase and PpATase to recognize distinct chemical scaffolds. We have\nbegun exploring several site-directed mutants in this region; however, these did not alter\nsubstrate preference, suggesting that the molecular determinants of specificity are more\ncomplex than initially anticipated and warrant further investigation.\nFigure 4b,c and corresponding main text l.218-230: This part is not entirely clear to me:\nThe first sentence “While no other function predictable Friedel–Crafts acylases were\nidentified” makes no sense to me. Also, to me it is not clear, why the enzyme from P.\nfluorescens was selected for further analysis. The rationale behind this experiment should\nbe better explained, as well the meaning of the obtained results.\nResponse:\nWe thank the reviewer for pointing out the lack of clarity in this section and for prompting\nus to improve the explanation.\nThe original sentence “While no other function predictable Friedel–Crafts acylases were\nidentified” has been removed from the manuscript. Our intention was simply to convey that\nour genome-mining survey did not reveal any additional, easily predictable Friedel–Crafts\nacylases beyond the known PpATase- and SmaATase-type enzymes.\nWe selected the PfACAT/HMGS–DUF35 complex from the 1,032 identified BGCs for\nfurther analysis because it represents a major but previously uncharacterized class. It is a\nminimal cluster that lacks canonical secondary metabolic partners (PKS/NRPS) but\nfeatures a fused HMGS-DUF35 protein (Fig. 4b). More importantly, sequence alignment\nrevealed a strikingly asymmetric conservation of catalytic features: (a) the HMGS domain\nlacks the essential E–C–H–N catalytic tetrad, indicating that this module is inactive, while\n(b) the ACAT domain retains the catalytic cysteine as well as the conserved HDxF and GHx\nmotifs (Fig. 3b).\nOur biochemical data confirmed this prediction. The PfACAT/HMGS–DUF35 complex does\nnot catalyze the HMGS reaction, but retains thiolase activity, converting acetoacetyl-CoA\nto acetyl-CoA—an activity not observed in SmaATase or PpATase (Fig. 4c).\nTogether, these findings support the interpretation that the PfACAT/HMGS–DUF35\ncomplex represents a plausible evolutionary intermediate—one that has diverged from its\nrole in primary metabolism but has not yet been recruited into a canonical secondary\nmetabolic pathway.\nThe narrative in this section has been revised accordingly in the manuscript (Please see\nlines 218 to 237).\nMinor:\nMethods: In the genome mining section, a reference for BiG-SCAPE should be added.\nResponse:\nDone.\nFig. 3a: Why are MucC/PhlB homologues not included in the phylogenetic tree?\nResponse:\nWe thank the reviewer for pointing this out. The MucC/PhlB homologs were included in our\nphylogenetic analysis, but their positions were not explicitly labeled in the original figure.\nWe have now revised Fig. 3a to clearly label the corresponding branch as “MucC/PhlB and\nhomologs”, ensuring that the figure is fully consistent with the description in the main text.\nFig. S7: What is the meaning of PA in this Figure?\nResponse:\nPA refer to phenyl acetate. We have revised the figure legends to include the definition\nexplicitly (“PA = phenyl acetate”). The original Fig. S7 has been updated and is now\npresented as Fig. S8 in the revised manuscript.\nFig: S24: The photos are barely visible and recognizable. Please improve Figure quality.\nResponse:\nWe have redesigned the figure with an optimized layout and high-resolution imagery to\nensure immediate visual recognition. The original Fig. S24 has been updated and is now\npresented as Fig. S30 in the revised version.\nFig. S26: In this Figure, it is referred to proteins in “Extended Data Fig. 2,4,6”. I don´t see\nthese data anywhere in the submitted files.\nResponse:\nWe thank the reviewer for pointing out this labeling inconsistency. We have corrected the\nlabels in the revised figure, which is now provided as revised Supplementary Fig. 32, and\nall corresponding references have been updated accordingly.\nSource data Fig. b (BGCs): It is nice, that the BGCs are displayed. However, these\ncollections in the Excel sheet confuse me: Either, the arrow colors must be clearly\nexplained, or better, cleaner SI Figures visualizing and aligning the BGCs should be\nprovided.\nResponse:\nWe thank the reviewer for this constructive suggestion. To improve clarity, we have\ngenerated new supplementary figures that provide a cleaner and more intuitive\nvisualization of the BGC alignments (see below). Additionally, during this revision we\nidentified two BGCs (one muc-like and one phl-like) with incomplete gene annotations.\nThese BGCs have been excluded from the analysis, and all corresponding descriptions in\nthe manuscript have been corrected accordingly.\n$%&SMT P !\"U !\"#SMT P !\"U\n5)$(7 + -,) G #\" \n!\"#$ \" ' ''()*+(\",-)!!\"# !\"# >\n!\"# \"\n!\"#$ ! \" \" # ' ' ' ( ) * # ) , ) - \" . # 5) ! $ \" ( # 7 + - , ) * : 0 - $ - ) )\n!\"#$ \" ' ! \" ' # ' ( ) * 0( \" $ \" $ - ) ) \" @ A C ? # 5)$( ! 7 \" # + - , ) > *9 > 0( #$ ) ' $ - ! ) ! # . < ;\n!\"#$ \" ' ! \" ' G . # ?#\n!\"# > A\n5)$(7 + -,)*90( #$)'$-) \" #\n!\"# @\n,\" 0,'\" 4,9 00()*),:$ J G\"\n!\"# C #\n5)$(7 + -,) G P \";\n!\"# > A\n,'\" 0,-\" 4,' 00()* 0,-\",#(+ C\n!\"# >\n=2# + 4,'\"$# (+*) 2, - < ;\n!\"# C \"\n. 9 7 4,'\"$# (+*0 - (+ K G M ;#\n!\"# ??\n5)$(7 + -,)* - $# !\n!\"# > >\n+ ) 0,'\" 4,' 00()*#$(\"$# < \"\n!\"# > ?@ >\n5)$(7 + -,) G ;>!\n!\"# >\n,'\" ' ''()*2 #' 0,'\" ) . \" #\n!\"#\n5)$(7 + -,)*,$\")() #\n,'\" 0,-\" 4,' 00()* 0,-\",#(+ \" !\"# > > A\n!\"# ! !\n=2# + 4,'\"$# (+* ,'' - J \"#\n.,' 00()* )$(7 +8' 7$) >?.# \" !\"# >\n!\"# > A >\n!\"# . , ' 00 ( ) *' $ # $ ( ) . =2# + ! 4 \" , # ' \"$ # ( + * ) ' - , $ ! !\n! + - \"'2 ,*\"$##,$ . !\"#$ \" +8'$) G N . P;\n!\"# !\"# > ##?@\n!\" 5 # # $ ) \" , * + $ , \" $# ( + \" \" ; \" !\"# > ! \" # $ \" + 8 ' $ ) ## ? G N . P ;\nC!? \" # ? KKiM P P\n; ',#7 7$) G >## \"\n!\"#\n; ',#7 ,*<,00,'$ ? !\n!\"#\nC!? \" # ? KKiM P P\nReviewer #3 (Remarks to the Author)\nThe manuscript entitled \"Evolutionary repurposing of a metabolic thiolase complex enables\nantibiotic biosynthesis\" by Liao et al. presents an interesting and informative structural and\nfunctional investigation of SmaATase, a multimeric thiolase enzyme complex from S.\nmacaca involved in the biosynthesis of the antimicrobial compound reutericyclin A. This\nsecondary metabolic pathway is proposed to have evolved from the central metabolic\npathway for mevalonate biosynthesis in archaea by functionally repurposing the tripartite\nenzyme complex that couples 3-hydroxy-3-methylglutaryl-CoA synthase (HMGS) and\nacetyl-CoA acetyltransferase (ACAT) reactivity. Biochemical characterization of SmaATase\nrevealed the HMGS activity of the proposed archaeal ancestor was lost, and instead of\nClaisen condensation activity the ACAT homolog catalyzes a non-canonical Friedel-Crafts\nC-acetylation on a pyrrolidine-2,4-dione scaffold. This is similar to another bacterial\ntripartite enzyme complex, PpATase from P. protogens, that performs a Friedel-Crafts C-\nacylation of monoacetylphloroglucinol (MAPG) to diacetylphloroglucinol (DAPG, an\nantimicrobial compound) and phloroglucinol (PG). Notably, although the HMGS activity of\nthe archaeal ancestor is not preserved in SmaATase or PpATase, the authors were only\nable to observe the C-acetylation activity in the presence of the complete tripartite complex,\nsuggesting the evolutionary trajectory may have prioritized chemical specialization over\nquaternary structural modifications. This point is further illustrated by comparison of\nexperimental structures, including new cryo-EM results of SmaATase, that show strong\nstructural similarities in the overall subunit organization and Co-A binding site, and all\ncomplexes contain the canonical thiolase ⍺β⍺β⍺ fold. Multiple sequence alignment reveals\nSmaATase and PpATase retain some, but not all, sequence motifs characteristic of ACATs,\nand they lack a conserved sequence essential for HGMS activity. Biochemical activity\nassays of wild type and generated mutants shed light on the functional roles of various\nsequence motifs. Finally, they find that the acetylation modification to form RTC-A and\nDAPG is critical for antimicrobial potency against competitors, implicating the vital\nimportance of these bacterial ATases. The results have broad importance across fields of\nbiosynthesis and bioengineering, and I recommend for publication after addressing the\nfollowing issues/comments that would improve this manuscript primarily in terms of clarity,\ndiscussion depth, and reproducibility.\nMajor Comments\n1. Docking studies are performed, but interpretation and presentation of the results are\nlacking. No data/results are presented besides SI Fig. 23 which does not provide\nsubstantial information. Line 237 states “Docking results showed that conserved C87 and\nH349 may coordinate catalysis, while H143, Y300, and Q302 support substrate\npositioning”. How? C87 and H349 are not depicted in SI Fig. 23a, and H143, Y300, and\nQ302 are only depicted as “contact residues”. Are there truly no more hydrogen bonds to\nthe ligand besides with Tyr300 (SI Fig. 23a)? Do the docking results show many or few\nbinding conformations? This manuscript could be greatly benefit from an explanation of the\nreaction mechanism catalyzed by PpATase (steps detailed in the manuscript Ref. 25) and\nbased on that the authors can discuss the likely residues of the SmaATase active site that\nare involved in the reaction. The differences between these two active sites, and their\ndeviation from archaeal ACAT discussed from the mechanistic perspective would certainly\nenhance the manuscript’s argument of functional repurposing.\nResponse:\nWe sincerely thank the reviewer for this thorough and constructive feedback. It has\nprompted a substantial revision of our computational analyses.\nWe acknowledge the limitations of our initial docking model and have therefore re-\nperformed all docking and molecular dynamics simulations with major methodological\nimprovements:\n(1) Full quaternary context: The system was reconstructed to include the MucB subunit\nalong with its neighboring MucA and MucC subunits, providing a more realistic active-site\nenvironment (details were given in question 3).\n(2) Incorporation of acetylated Cys87: Cys87 was modeled in its acetylated state as a\nnon-standard residue, consistent with the electron density and confirmed by LC–MS/MS\n(new Supplementary Fig. 29). This correction was essential for accurate mechanistic\ninterpretation.\n(3) Full-length substrate modeling: preRTC-A was modeled in its full native form rather\nother a truncated analogue.\n(4) Improved docking methodology: Docking was re-performed using DiffDock, and the\nmost representative poses were selected for analysis.\n(5) Replicated dynamics simulations: Subsequent analyses were based on three\nindependent and replicated MD simulations to ensure statistical reliability.\nThe results of this refined workflow are now presented as follows: (1) Fig. 5 shows key\npolar interactions identified by PyMOL analysis of the optimal docking pose; (2)\nSupplementary Fig. 27 provides a comprehensive 2D interaction map (hydrophobic\ncontacts and van der Waals interactions) generated with ProLIF.\nImportantly, following the reviewer’s suggestion, we have added a mechanistic comparison\nbetween SmaATase and PpATase in the revised manuscript (lines 242–256). We\nsummarize the established catalytic steps of PpATase, map their equivalents in SmaATase,\nand discuss how divergence from the archaeal ACAT active site supports functional\nrepurposing. This substantially strengthens the mechanistic argument of our study.\n2. MD simulations are performed, but interpretation and presentation of the results are\nlacking. SI Fig 23b is not discussed anywhere. In particular, the per-residue RMSF appears\nto show that many active site residues have high fluctuations. Typically, when a binder is\npresent we might expect active site rigidity, especially if the molecule is stably bound.\nResponse:\nWe thank the reviewer for this insightful comment. To improve transparency and\ninterpretation of the MD results, we have added the following explanatory sentence to the\nFig. 5 legend regarding the updated Supplementary Fig. 27 (formerly Fig. S23):\n“The reliability of the simulated substrate-binding mode is supported by the\nidentification of specific protein-ligand interactions, alongside with the convergence\nof the protein-ligand RMSD, the low residual flexibility (RMSF) in the binding pocket,\nand the stable compactness (Rg) and solvent accessibility (SASA) (Supplementary\nFig. 27).”\nRegarding the reviewer’s concern about the per-residue RMSF values: in the updated\nsimulations, we indeed observe measurable flexibility in several active-site residues. This\nmay reflect the intrinsic property of the binding pocket in MucB, which is situated at the\ninterface with MucA and MucC subunits (as shown in Supplementary Fig. 18b). Our in\nvitro results also demonstrate that SmaATase accepts a broad spectrum of preRTC-A\nanalogs featuring varying acyl chain lengths and distinct amino acid side chains (These\nexperimental data have been provided to the reviewers in the attached file.). This evidence\nalso supports the conformational flexibility of the substrate-binding pocket.\n3. Simulations were performed with only MucB, not the full complex, which was shown to\nbe required for activity. Did the authors perform any binding kinetics with only MucB, or\nhave reason to believe MucB can effectively bind the substrate, and in the same way it\nwould if the complex was formed? It is interesting that the MM/GBSA binding free energy\ncalculations show favorable binding to MucB, but experimental results (line 166) “suggest\nthat while MucA and MucC may not directly participate in catalysis, they are essential for\nproper folding and assembly of the tripartite complex, thereby enabling its enzymatic\nactivity”. Some discussion about the implications of these two results should be included\nto strengthen the manuscript.\nResponse:\nWe sincerely thank the reviewer for raising this important point. Our initial simulations\nfocused on MucB extracted from the full complex under the assumption that it retained the\nfunctional conformation of the catalytic pocket. We agree that this required refinement.\nTo directly address the reviewer’s question, we examined whether isolated MucB is\ncapable of binding the substrate. Surface plasmon resonance (SPR) measurements\nshowed that MucB alone binds preRTC-A with a K of 6.04 × 10⁻⁶ M, which is comparable\nD\nto the affinity of the full SmaATase complex (2.52 × 10⁻⁶ M, Supplementary Fig. 18a).\nHowever, isolated MucB is catalytically inactive, despite its ability to bind the substrate.\nThis apparent discrepancy can be explained by the structural organization of the active\nsite: as shown in Supplementary Fig. 18b, the catalytic pocket resides at the interface of\nMucA, MucB, and MucC. Thus, MucB alone can bind the ligand, but cannot achieve the\ncatalytic geometry without the contributions of MucA and MucC, which help shape the\nactive-site architecture. In light of this, we have re-performed all molecular docking and MD\nsimulations using a model that includes MucB together with its adjacent MucA and MucC\nsubunits, providing a more realistic representation of the enzymatically competent state.\nThese revised results are now included in Fig. 5 and Supplementary Fig. 27.\nTo reconcile the biochemical and computational findings, we propose that SmaATase\nexhibits assembly-induced activation, a phenomenon well documented in other multimeric\nsystems. For example:\n• In the archaeal HMGS/ACAT/DUF35 complex, isolated subunits are insoluble or inactive.\n• In the proteasome, β-subunits are minimally active alone but become fully active only\nupon correct assembly.\nWe have incorporated a description of the SPR results and the mechanistic implication into\nthe revised manuscript (lines 167–173), highlighting that while substrate binding can occur\nin MucB alone, proper assembly of the tripartite complex is required to establish the\ncatalytically competent active-site configuration.\n4. The energy minimized coordinates of the ligand used in docking should be provided for\nreproducibility, as well as at least one set of xyz coordinates from the docking/simulation\nresults of the holo enzyme.\nResponse:\nWe thank the reviewer for this important suggestion regarding reproducibility. As requested,\nthe energy-minimized coordinates of the ligand (optimized at the B3LYP/def2-TZVP level\nusing ORCA) and representative XYZ coordinates from the DiffDock simulation of the holo-\nenzyme have been provided in the figshare repository associated with this manuscript.\n5. The assigned protonation states used for simulations should be stated in the SI for the\nmethod to be transparent and reproducible. Additionally, the protonation state of residues\nin the binding site are likely significant. Some justification of choice should be provided, or\nevaluation of various possible states to ensure results are robust, but since these results\nare not critical for the manuscripts main objectives additional calculations could be avoided.\nResponse:\nIn the revised Methods section, we have added a detailed description of the protonation-\nstate assignment. Protonation states for all ionizable residues were determined using\nPropKa at neutral pH. Specifically, His305 and Asp35 in MucA, Glu57, Asp134, and Glu296\nin MucB, and Glu70 in MucC were predicted to be protonated, while all other residues were\nkept in their standard protonation states.\nMinor Comments\n1. DUF35 is not defined.\nResponse:\nWe have revised the main text to include a definition upon first use as: “DUF35 (Pfam\nfamily PF01796, Domain of Unknown Function)”.\n2. Line 91: Why was the MucABC sequence from S. mutans was used for the query instead\nof S. macaca? Since the paper focuses on the S. macaca complex it would be more\nconsistent to use those results and likely does not alter any discussion.\nResponse:\nWe used the S. mutans MucABC sequence as the initial BLASTP query because this study\nbuilds directly on our previous work on the S. mutans muc BGC (ACS Infect. Dis. 2020,\n6:563; Microbiome 2023, 11:97), making it the natural starting point for our analysis. For\nbiochemical and structural characterization, however, we prioritized the S. macacae\nhomolog due to its superior protein purity and yield. Importantly, the two homologs share\nextremely high sequence identity (89-99% for MucA/MucB and 100% for MucC), so the\nchoice of query sequence does not alter the BLASTP mining results or affect any\nconclusions in the manuscript.\n3. Lines 128-129: Results for acetyl-CoA as a donor are stated. The reader would benefit\nfrom the results of the other physiologically relevant donors also being quickly stated.\nResponse:\nWe thank the reviewer for this suggestion. In response, we have revised the main text as\nfollowing:\n4. Lines 134-135: “Together, these results support acetyl-CoA as the physiological acetyl\ndonor…”. This may be too strong of a statement. First, it appears there is some, albeit very\nlittle, product formation with donors acetoacetyl-CoA and malonyl-CoA (Fig. 1b). Second,\nare the donors evaluated the only physiological possibilities? Third, a previous study found\nno product with PpATase and acetyl-CoA. Either more justification should be included in\nthe text to convince the reader that acetyl-CoA is THE donor, or the statement can be\nmodified. Additionally, some discussion on why product was found using PpATase with\nacetyl-CoA when a previous study found this did not work is warranted.\nResponse:\nWe thank the reviewer for raising this important point. We agree that our original statement\nthat acetyl-CoA is “the physiological acetyl donor” was too strong and have revised the text\naccordingly.\nOur data indicates that acetyl-CoA is the most likely physiological donor, as it produces\nclear and reproducible product formation in SmaATase-catalyzed reactions. In contrast,\nother potential donors—acetoacetyl-CoA and malonyl-CoA—generate only trace signals\n(Fig. 1b). These minimal peaks may reflect very weak enzymatic activity toward these\ndonors or low-level non-enzymatic breakdown of acetoacetyl-CoA/malonyl-CoA into\nacetyl-CoA. Given the extremely small magnitude of these signals and the robust activity\nobserved with acetyl-CoA, we consider the physiological relevance of the alternative\ndonors to be negligible. Accordingly, we have modified the relevant sentence to the more\ncautious: “Together, these results support acetyl-CoA as the potential physiological\nacetyl donor for both SmaATase and PpATase.”\nRegarding the discrepancy with earlier studies reporting no detectable PpATase activity\nwith acetyl-CoA, the cited work provides insufficient methodological detail to identify the\nprecise cause. However, methodological differences offer a plausible explanation. In the\nfirst study (Biosci. Biotech. Bioch. 2012, 76: 559), enzyme activity was monitored\nspectrophotometrically at 370 nm—an indirect and relatively insensitive approach that may\nnot detect low turnover rates. In the later study (ChemCatChem 2019, 11: 1064), HPLC\ndetection was performed at 254 nm. Our data show that this wavelength is suboptimal for\nspecifically detecting MAPG and DAPG, whereas our UPLC-based assay at 280 nm\nprovides a substantially stronger and more selective signal (see the figure below).\n'##\n#\n! \"# \"$ \"% \"&\n'##\n#\n! \"# \"$ \"% \"&\nAdditionally, the HMGS/ACAT/DUF35 complex is known to be unstable during storage; for\nexample, the archaeal homolog retains only ~15 activity after two weeks at –80 °C\n(PNAS 2018, 115: 3380). The use of suboptimally preserved enzyme or insufficiently\nsensitive detection methods in previous studies could therefore have resulted in a false-\nnegative outcome.\nTo avoid over-interpreting these methodological differences and to keep the focus on our\nfindings, we have removed the comparative statement about earlier studies from the\nrevised manuscript (see below).\n5. “r.m.s.d” should be RMSD throughout manuscript.\nResponse:\nDone.\n6. Line 179: The use of the word “mirrors” here could cause confusion as it implies chiral\npartners. “Resembles” would be a safer choice.\nResponse:\nRevision has been made as suggested.\n7. Line 192: Replace “substitutions” with “modifications” or similar, since the differences\nare more signification than single AA substitutions.\nResponse:\nWe agree with the reviewer that “substitutions” may imply simple single-amino-acid\nchanges. We have chosen “mutations” instead of “modifications” for the revised heading,\nas the latter may refers to post-translational or chemical alterations of amino acids.\n8. Line 218: “While no other function predictable…” This sentence was awkward to\nunderstand- could be written more clearly.\nResponse:\nThis sentence was intended to convey that our genome mining did not identify any new,\npredictable Friedel-Crafts acylases beyond the known types (SmaATase and PpATase).\nWe have revised the sentence in the manuscript to clarify this point and remove the\nawkward original phrasing (see below).\n9. Line 239: E124 and S142 are not hydrophobic residues.\nResponse:\nThe paragraph (line238–line256) has been revised to correct the inaccurate description.\n10. Line 240: Reference needed for W211 in Ph1C proposed to act as conformational lid.\nResponse:\nDone.\n11. Line 275: The reactions of SmaATase and PpATase are not “single-step” (particularly\nbecause they involve acylation and deacylation phases that are likely both multi-step\nreactions). Do the authors mean single reactions, as opposed to coupled reactions?\nResponse:\nWe thank the reviewer for this important clarification. We agree that describing the\nSmaATase and PpATase reactions as “single-step” was inaccurate, given that both the\nacylation and deacylation phases involve multiple mechanistic steps. As suggested, we\nhave revised the sentence to state that SmaATase and PpATase mediate single reactions\nthat do not require substrate channeling.\n12. Line 435: What pH was assumed for protonation state assignment?\nResponse:\nWe assigned the protonation states at the canonical physiological pH of 7.4 in the revised\nsimulations.\n13. Line 435: Is “hydrogenated” an appropriate term here? Do the authors mean they\nmodeled in unresolved H atoms to the structure?\nResponse:\nWe thank the reviewer for catching this inaccuracy. The term “hydrogenated” was\ninappropriate, as AutoDock does not systematically complete all missing hydrogens. We\nhave replaced AutoDock with DiffDock in our revised workflow, which does not require pre-\nadded hydrogens for the docking procedure.\n14. Line 440: Can the authors provide more information beyond “Phad Calculation Co.,\nLtd”? I tried searching for this company and could not find any results.\nResponse:\nWe thank the reviewer for raising this point. For clarification, the official website of “Phad\nCalculation Co., Ltd” is https://phadcalc.com/.\nHowever, in the revised version, we have removed all simulations previously performed by\n“Phad Calculation Co., Ltd” and deleted the corresponding acknowledgement. The\nupdated molecular dynamics data were performed and validated by Dr. Zhuanglin Shen\n(Chinese Academy of Sciences), who has been added to the author list in recognition of\nthis contribution.\n15. Line 442: ff19SB needs reference and AMBER Tools needs reference.\nResponse:\nIn the revised manuscript, we have updated the simulation methodology to reflect our new\ncomputational workflow. All molecular dynamics simulations were re-performed using\nGROMACS with the AMBER ff14SB force field, which offers improved compatibility with\nthe TIP3P water model. The appropriate references have now been added to the Methods\nsection.\n16. Line 443: How were the atomic charges derived for GAFF parameters of the ligand?\nResponse:\nIn the revised manuscript, RESP charges were generated using Psi4 for the ligand as well\nas for the acetylated Cys87 residue. This approach ensures a consistent and accurate\ndescription of the electrostatics, which improves the reliability of our GAFF\nparameterization for subsequent simulations.\n17. Line 448: Says SHAKE algorithm was used in heating and equilibration- but what about\nduring the production runs?\nResponse:\nIn the revised manuscript, the molecular dynamics simulations were re-performed using\nGROMACS, and the bond constraints were handled with the more efficient LINCS\nalgorithm. This was applied throughout both the equilibration and production phases of the\nsimulations to ensure consistency and stability.\n18. Line 451: PME needs reference.\nResponse:\nDone.\n19. Line 452: MM/GBSA needs reference.\nResponse:\nDone.\n20. Line 617: “DUF35 unites highlighted in cartoon format”- it looks like more than just\nDUF35 is in cartoon… are the captions for d and e switched?\nResponse:\nWe apologize for the mismatch between the figure panels d and e and their descriptions in\nthe legend. The captions for Figure 2d and 2e have been swapped in the revised version.\n21. Fig. 5:\na. Is the PpATase Cys88 acylated? This should be stated somewhere, as well as where\nthis structure came from.\nb. The ACAT represented after step2 for both ATases has a H x-ed out, but this is a bit\nconfusing because the actives sites of both have numerous histidines.\nc. Why are some H-bonds listed in this SmaATase active site that are not shown in SI Fig.\n23?\nd. It should be clarified somewhere that the H-bonds are measure from the heavy atoms\n(at least they appear to be), or those H atoms should be included in the representation and\nthe true H-bond labeled.\nResponse:\nWe thank the reviewer for these detailed comments on Figure 5. We have addressed each\npoint as follows:\na, We have stated in the main text that PhlC-Cys88 is acetylated in PpATase (PDB: 5MG5)\nand added the corresponding reference.\nb, We have labeled the relevant histidine motifs as “HDxF” and “GHx” in Fig. 5 for clarity.\nc, We have resolved the depiction discrepancy by showing hydrogen bonds in the 3D\noverview (Fig. 5) and hydrophobic/van der Waals forces in the 2D diagram (Supplementary\nFig. 27).\nd, We thank the reviewer for pointing out the precise depiction of hydrogen bonds. To\nensure clarity, we have revised the figure captions to state that hydrogen bonds are inferred\nfrom donor-acceptor heavy atom distances.\n22. Fig. S16: Typo “ligntblue”.\nResponse:\nDone.\n23. Fig. S23: Text of b, c, d, e, f too small. Why is Tyr300 in ball and stick representation?\nIt makes it seem like it is also a ligand and is confusing to the reader. See comment 21\nabout H-bonds. Cys88 and His349 should be included.\nResponse:\nWe have fully regenerated this figure (now is Supplementary Fig. 27) to address the\nreviewer’s concerns. The updated version incorporates a ProLIF analysis to display\nhydrophobic contacts and Van der Waals interactions between the active site residues\n(including His349 and acetylated Cys88) and preRTC-A. All residues are consistently\nlabeled with color-coding. The figure was also updated with new MD data and the text size\nwas increased for enhanced clarity.\nReviewer(s)' Comments to Author:\nWe thank the reviewers for their time and constructive comments, which have helped us\nsignificantly improve the manuscript. We have addressed the remaining concerns raised\nby the reviewers as detailed below.\nReviewer #1 (Remarks to the Author)\nThe reviewer and co-reviewer read the rebuttal letter and the manuscript modifications\nindependently. Both arrived at the unbiased conclusion that the manuscript gained\nsufficient clarity while presenting additional experiments that strengthen the story.\nTherefore, we are delighted to recommend its publication and congratulate the authors\nonce more on this excellent work. The final editing during the proofs should be sufficient to\nremove the last typos (e.g., Fig S10 panel b, the axis should be volume and not volumn).\nResponse:\nWe are grateful for the reviewer’s positive feedback and their recommendation for\npublication. As noted, we have corrected the typo (“volumn” to “volume”) in Fig. S10b.\nReviewer #2 (Remarks to the Author):\nThe authors have addressed most of the comments by me and also the other reviewers\n(as far as I can judge) comprehensively and convincingly. There is only one remark to one\nof my initial minor comments that has not been addressed, probably due to a\nmisunderstanding in nomenclature.\nThe comment \"Fig. 3a: Why are MucC/PhlB homologues not included in the phylogenetic\ntree?\" was related to the lack of exactly these enzymes (DUF35 homologues) in the\nphylogenetic tree. In the revised version, these are still missing in the tree, which has not\nchanged, as far as I can see. My question was pointing to if there is a particular reason for\nexcluding these from the phylogenetic analysis or if it would make sense to include them.\nResponse:\nWe sincerely apologize for the oversight of having confused MucC/PhlB with MucB/PhlC\nin our previous response. The phylogenetic analysis in Fig. 3a was intended to reconstruct\nthe evolutionary relationships within the thiolase superfamily. MucA/PhlA belong to the\nHMGS (hydroxymethylglutaryl-CoA synthase) clade, and MucB/PhlC are members of the\nACAT (acetyl-CoA C-acetyltransferase) clade, both of which are established members of\nthe thiolase superfamily. In contrast, MucC/PhlB are classified under the DUF35 family\n(domain of unknown function), which falls outside the thiolase superfamily. Therefore,\nMucC/PhlB were not included in this analysis."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The reviewer and co-reviewer read the rebuttal letter and the manuscript modifications independently. Both arrived at the unbiased conclusion that the manuscript gained sufficient clarity while presenting additional experiments that strengthen the story. Therefore, we are delighted to recommend its publication and congratulate the authors once more on this excellent work. The final editing during the proofs should be sufficient to remove the last typos (e.g., Fig S10 panel b, the axis should be volume and not volumn).", "reviewer_2": "The authors have addressed most of the comments by me and also the other reviewers (as far as I can judge) comprehensively and convincingly. There is only one remark to one of my initial minor comments that has not been addressed, probably due to a misunderstanding in nomenclature.\nThe comment \"Fig. 3a: Why are MucC/PhlB homologues not included in the phylogenetic tree?\" was related to the lack of exactly these enzymes (DUF35 homologues) in the phylogenetic tree. In the revised version, these are still missing in the tree, which has not changed, as far as I can see. My question was pointing to if there is a particular reason for excluding these from the phylogenetic analysis or if it would make sense to include them.", "reviewer_3": "The authors have satisfactorily addressed all of my previous comments. The manuscript is now clearer, the data are well supported, and the conclusions are appropriately framed. I have no further concerns.\nI recommend the manuscript for publication in its current form.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "We thank the reviewers for their time and constructive comments, which have helped us significantly improve the manuscript. We have addressed the remaining concerns raised by the reviewers as detailed below.\nReviewer #1 (Remarks to the Author)\nThe reviewer and co-reviewer read the rebuttal letter and the manuscript modifications independently. Both arrived at the unbiased conclusion that the manuscript gained sufficient clarity while presenting additional experiments that strengthen the story. Therefore, we are delighted to recommend its publication and congratulate the authors once more on this excellent work. The final editing during the proofs should be sufficient to remove the last typos (e.g., Fig S10 panel b, the axis should be volume and not volumn).\nResponse:\nWe are grateful for the reviewer’s positive feedback and their recommendation for publication. As noted, we have corrected the typo (“volumn” to “volume”) in Fig. S10b.\nReviewer #2 (Remarks to the Author):\nThe authors have addressed most of the comments by me and also the other reviewers (as far as I can judge) comprehensively and convincingly. There is only one remark to one of my initial minor comments that has not been addressed, probably due to a misunderstanding in nomenclature.\nThe comment \"Fig. 3a: Why are MucC/PhlB homologues not included in the phylogenetic tree?\" was related to the lack of exactly these enzymes (DUF35 homologues) in the phylogenetic tree. In the revised version, these are still missing in the tree, which has not changed, as far as I can see. My question was pointing to if there is a particular reason for excluding these from the phylogenetic analysis or if it would make sense to include them.\nResponse:\nWe sincerely apologize for the oversight of having confused MucC/PhlB with MucB/PhlC in our previous response. The phylogenetic analysis in Fig. 3a was intended to reconstruct the evolutionary relationships within the thiolase superfamily. MucA/PhlA belong to the HMGS (hydroxymethylglutaryl-CoA synthase) clade, and MucB/PhlC are members of the ACAT (acetyl-CoA C-acetyltransferase) clade, both of which are established members of the thiolase superfamily. In contrast, MucC/PhlB are classified under the DUF35 family (domain of unknown function), which falls outside the thiolase superfamily. Therefore, MucC/PhlB were not included in this analysis."}]

Decision letter