Molecular insights into fungal inositol phosphorylceramide synthesis and its inhibition by antifungal aureobasidin A

Domain: chemistry
Published: 2026-02-19
Editorial outcome: major_revision

Abstract
Fungal inositol phosphorylceramide (IPC) synthase is an essential enzyme complex that catalyzes a critical step in sphingolipid biosynthesis. It is the molecular target of potent antifungal aureobasidin A (AbA). Despite its therapeutic relevance, the lack of structural and mechanistic insights into IPC synthase function and inhibition has impeded rational antifungal drug development. Here, we present cryo-EM structures of Saccharomyces cerevisiae IPC synthase in two distinct functional states: a ceramide-bound form and an AbA-inhibited complex. Our study reveals a conserved heterodimeric architecture formed by Aur1 and Kei1, stabilized through extensive protein-protein and lipid-mediated interactions. Within catalytic Aur1, we identify a membrane-embedded reaction chamber harboring a conserved H-H-D catalytic triad (H255, H294, and D298) essential for IPC synthesis. Structural comparisons illuminate the mechanism of ceramide recognition and reveal how AbA acts as a competitive inhibitor by occupying the substrate-binding pocket. Further analyses identify key residues involved in AbA binding and explain the molecular basis of drug resistance. Together, these findings advance the mechanistic understanding of fungal IPC biosynthesis and inhibition, and establish a foundation for developing new antifungal drugs targeting IPC synthase.

Introduction
Invasive fungal infections (IFIs) pose a serious global health threat, particularly among immunocompromised individuals, accounting for an estimated 3.8 million deaths annually1. The clinical management of IFIs is increasingly challenged by the limited arsenal of antifungal drugs, the growing prevalence of antifungal resistance and the emergence of multidrug-resistant fungal pathogens2,3. These pressing issues underscore the urgent need for developing novel antifungal drugs3,4.

A particularly promising target for antifungals is the biosynthesis of inositol phosphorylceramide (IPC), a critical component of fungal membranes that is absent in mammalian host4-7. IPC is synthesized by fungal IPC synthase, a membrane-bound enzyme complex composed of the catalytic subunit Aur1 and the regulatory subunit Kei18-11. IPC synthase as a key enzyme in fungal sphingolipid biosynthetic pathway, catalyzes the transfer of inositol phosphate from phosphatidylinositol (PI) to ceramide, generating IPC and diacylglycerol (DAG)8,10 (Fig. 1A). Crucially, IPC synthase is highly conserved across prominent fungal pathogens and is indispensable for fungal viability and virulence12-14. Genetic disruption of AUR1 is lethal in Candida albicans and S. Cerevisiae, due to ceramide accumulation and sphingolipid depletion9,15,16. The mould Aspergillus nidulans with AUR1 deficiency suffers from impaired microtubule assembly and cell death17. KEI1 defects cause mislocalization of IPC synthase and severe growth defects in S. Cerevisiae11. Furthermore, down-regulation of IPC synthase in Cryptococcus neoformans significantly attenuates cell growth and pathogenicity13,18.

The therapeutic potential of targeting IPC synthase is exemplified by the Aureobasidin A (AbA), a cyclic depsipeptide fungicide that specifically inhibits this enzyme19. AbA and its derivatives exhibit potent broad-spectrum antifungal activity, with nanomolar minimum inhibitory concentrations (MICs) against key pathogens such as Candida albicans and Aspergillus fumigatus20-24. Notably, in murine models of disseminated candidiasis, AbA exhibited superior fungicidal activity compared to conventional antifungals such as fluconazole and amphotericin B23. Furthermore, AbA demonstrates excellent selectivity, showing no cytotoxicity against mammalian cells at therapeutic concentrations25. Recent studies have also revealed synergistic interactions between AbA and established antifungals, highlighting its potential for combination therapy approaches26. The genetic basis of AbA resistance and susceptibility further underscores the specificity of IPC synthase targeting. Mutations in AUR1 or its overexpression confer AbA resistance in Aspergillus nidulans, S. cerevisiae and S. pombe9,27-29. Conversely, KEI1 deficiency hypersensitizes fungi cells to AbA, reinforcing the critical role of IPC synthase activity in antifungal susceptibility11. Despite the clear clinical relevance of fungal IPC synthase, detailed molecular insights into this essential antifungal drug target remain limited, hindering rational drug design.

Here, we report the cryo-electron microscopy (cryo-EM) structures of S. cerevisiae IPC synthase in both ceramide-bound and Aureobasidin A (AbA)-inhibited states. Further structure-guided functional characterizations enable us to elucidate the molecular basis of IPC biosynthesis, uncover AbA-mediated inhibition, and rationalize drug-resistant mutations within the enzyme complex.

Results
Functional characterization of IPC synthase
To enable in vivo functional characterization of fungal IPC synthase (IPCS), we generated a conditional AUR1 knockdown strain to circumvent the lethal phenotype associated with AUR1 disruption in S. cerevisiae30. Under repressing conditions with glucose, silencing of AUR1 expression resulted in growth arrest (Fig. 1B). This phenotype was rescued by reintroducing wild-type AUR1, confirming the essential role of AUR1 in fungal viability (Fig. 1B). In contrast, complementation with AUR1 H294A, a catalytically inactive mutation identified from our structural analysis (see the section ‘The reaction chamber and catalytic mechanism’), failed to restore growth (Fig. 1B). These findings demonstrate that the enzymatic activity of IPC synthase is indispensable for cell survival.

To biochemically characterize IPC synthase activity, we immunopurified the wild-type enzyme complex and performed in vitro assays using phosphatidylinositol (PI) and a fluorogenic NBD-labeled ceramide as substrates. The enzyme efficiently catalyzed the formation of NBD-labelled inositol phosphorylceramide (IPC), confirming its inositol phosphoryl transferase activity (Fig. 1C). Interestingly, SDS-PAGE combined with western-blot analyses of the purified IPCS complex revealed that Aur1, but not Kei1, displayed several high–molecular-weight species (Supplementary Fig. 1G-H). Treatment of the purified complex with PNGase F eliminated these higher–molecular-weight forms of Aur1, confirming that they arise from N-linked glycosylation (Supplementary Fig. 1G-H). Enzymatic assays further showed that removal of N-glycans had minimal impact on IPC synthase activity (Supplementary Fig. 1I).

Given that certain sphingolipid synthases or variants exhibit bifunctional or trifunctional synthase activities to produce IPC, sphingomyelin (SM), or ethanolamine phosphorylceramide (EPC) depending on the phospholipid donor31-33, we investigated the substrate specificity of fungal IPC synthase. Substitution of PI with phosphatidylcholine (PC) or phosphatidylethanolamine (PE) failed to support the synthesis of SM or EPC (Fig. 1C). Although faint product bands were initially observed, their mobility on thin-layer chromatography (TLC) matched that of IPC and further time-course analyses revealed that these signals likely arose from residual endogenous PI co-purified with the enzyme rather than authentic utilization of PE or PC (Fig. 1C), highlighting the strict substrate specificity of fungal IPC synthase.

Structural determination of IPC synthase
To elucidate mechanistic insights into fungal IPC synthase, we set out to determine the structure of the S. cerevisiae enzyme complex. However, initial cryo-electron microscopy (cryo-EM) structural analysis using the wild-type enzyme is difficult due to the small molecular size (~71 kd) and the lack of prominent structural features beyond the transmembrane regions (Fig. 1E). To overcome these limitations, we employed a molecular weight augmentation strategy by introducing a helical ALFA epitope tag (SRLEEELRRRLTE) along with its high-affinity binding nanobody (Nb) as a fiducial marker34,35. To preserve enzymatic function, the ALFA tag was inserted in replace a flexible loop region (residues 127-131) of the regulatory subunit Kei1 (Fig. 1E). The engineered ALFA-tagged IPC synthase with or without the bound Nb were prepared (Supplementary Fig. 1A-F). Subsequent activity assays confirmed that they both retained comparable catalytic activity to the wild-type enzyme (Fig. 1D), demonstrating that neither ALFA insertion nor nanobody binding compromised the functional integrity of the complex.

The incorporation of the ALFA-Nb system markedly improved particle alignment, as evidenced by the well-defined structural features in both 2D class averages and 3D reconstructions (Fig. 1F, Supplementary Fig. 1J-L). Using a dataset of 162,123 particles, we obtained a final cryo-EM map of ceramide-incubated IPC synthase at an overall resolution of 3.5 Å (Fig. 1F, Supplementary Fig. 3). This high-quality density map enabled unambiguous modeling of Aur1, Kei1, ceramide and several well-ordered bound lipids (Fig. 1G, Supplementary Fig. 3-4), yielding critical molecular insights into the fungal IPC synthase complex. Notably, although we confirmed the presence of N-glycosylation on Aur1 (Supplementary Fig. 1G-H), the specific glycosylation site could not be unambiguously assigned. Among the three predicted glycosylation sites (N132, N243, and N367)36, N367 is located in a disordered region of the structure. In contrast, while N132 and N243 lie in ordered regions, neither exhibits clear glycan density. This suggests that these sites are either unmodified or contain highly flexible glycans, with the latter consistent with the minimal enzymatic effects observed upon glycan removal (Supplementary Fig. 1I).

Architecture of fungal IPC synthase
The fungal IPC synthase is organized in a heterodimeric assembly comprising Aur1 and Kei1, with overall dimensions of approximately 112 Å × 96 Å × 89 Å. Both the N- and C-termini of Aur1 and Kei1 are oriented toward the cytosol (Fig. 1E-G).

Aur1, the larger subunit (401 amino acids; ~45kd), contains eight transmembrane helices (TM1-TM8) (Fig. 1G). Among these, TM3-TM8 form the core transmembrane domain, whereas TM1 and TM2 are positioned peripherally (Fig. 1G). On one side of the Aur1 core domain, TAM8 approaches Kei1 to mediate inter-subunit interaction, while the opposite side forms the substrate-binding chamber that is enclosed by TM3–TM6 (see below) (Fig. 1G). Within the Aur1 core domain, TM3 and TM4 are discontinuous, with TM3 interrupted close to its cytosolic end and TM4 is split in the middle (Fig. 1G). Another distinctive feature of Aur1 is the presence of two extended luminal loops: EL2 (residues 106–143) between TM3 and TM4, and EL3 (residues 198–253) between TM5 and TM6 (Fig. 1E, 1G). These two ~40-residue loops harbor short helices and interweave to form a cap-like subdomain beneath the core transmembrane region, playing crucial roles in complex assembly and active site formation with short luminal loop EL4 (see below).

Kei1, the smaller subunit (221 amino acids; ~25kd), consists of four elongated transmembrane helices (TM1-TM4). These helices all adopt tilted configurations within the membrane and collectively form a four-helix bundle that opens toward the luminal side (Fig. 1G). At the C-terminus, Kei1 features an amphipathic lateral helix (LH1) that interacts with and stabilizes the juxtaposed cytosolic ends of TM3 and TM4. Two loop regions, EL2 (residues 112–139) on the luminal side and CL2 (residues 185–199) on the cytosolic side, were not resolved in the structure, indicating their intrinsic conformational flexibility.

Aur1 topology and the LPT (lipid phosphatase/phosphotransferase) fold
A DALI server search for the regulatory subunit Kei1 of the IPC heterodimer did not yield any structurally similar homologs37. In contrast, for the catalytic subunit Aur1, although the overall architecture of Aur1 differs from previously reported structures, a DALI homology search identified several membrane-protein structures that resemble a portion of Aur1’s transmembrane topology (residues 143-314) (Supplementary Fig. 5). These include (in a descending order of Z-score): Lipid A 1-Phosphatase (LpxE)38, Glucose-6-phosphatase (G6PC1)39, Phosphatidylglycerophosphatase B (PgpB)40, and Sphingomyelin synthase-related protein 1 (SMSr)41. Despite their low sequence conservation (<10% identities), structural superposition revealed that a cluster of five successive transmembrane helices of these proteins adopt comparable topologies, whereas the remaining regions diverge in both fold and number of transmembrane helices (Supplementary Fig. 5). Notably, all these structurally related proteins belong to lipid phosphatase/phosphotransferase (LPT) family, most members of which are integral membrane enzymes catalyzing the dephosphorylation or phosphotransfer reactions of diverse phospho-lipid or phospho-carbohydrate substrate42. This suggests that these enzymes (including Aur1, LpxE, G6PC1, PgpB, SMSr) share a conserved structural module, which we term the LPT fold. This fold exhibits several common features (Supplementary Fig. 5): it comprises five transmembrane helices, often situated in the C-terminal region (except in G6PC1); the luminal loop (e.g., EL4 in Aur1) connecting the last two helices harbors a catalytically essential histidine residue (e.g., H294 in Aur1; see Discussions for further mechanistic details). Despite this conserved core, notable structural variations exist. The orientation and length of the topologically equivalent transmembrane helices differ among family members. For instance, the first helix of the module is continuous in SMSr (TM5) but kinked in Aur1 (TM4), LpxE (TM1), G6PC1 (TM1), and PgpB (TM2), reflecting structural variability within this shared scaffold (see Discussion for further comparison).

Assembly of the IPC synthase complex
Though Kei1 has been proposed as the non-catalytic subunit, our analyses clearly demonstrate its essential role in IPC synthase function. Repression of KEI1 expression resulted in severe growth defects (Fig. 2A), comparable to those observed upon AUR1 knockdown (Fig. 1B). Notably, while Aur 1 could be expressed and purified in the absence of Kei1, the resulting protein exhibited no detectable IPC synthase activity (Fig. 2B), underscoring the indispensable requirement for Kei1 for enzymatic function.

To elucidate the molecular basis of this functional interdependence, we examined the assembly of the Aur1–Kei1 complex. The two subunits form extensive contacts spanning both membrane-embedded and luminal regions (Fig. 2C). Within the membrane, the inter-subunit interface is predominantly mediated by van der Waals interactions between TM8 of Aur1 and TM1 of Kei1 (Fig. 2D). Key residues involved include Y315, Y312, Y307, V304, M300 and F296 in Aur1, and M16, L20, I24, I28, L31, and Y38 in Kei1. On the luminal side, the tilted arrangement of Kei1 TM1–TM4 creates a spacious inter-helical cavity that accommodates the elongated EL2 loop of Aur1 (Fig. 2E). This loop engages Kei1 through a dense network of interactions in mixed nature: polar contacts include Aur1 D131 with Kei1 H47, Aur1 Y129 with Kei1 N32, and the Aur1 D131 backbone with Kei1 Y57; representative hydrophobic contacts involve L137, A123 and G130 of Aur1 with L41, W107, and F103 of Kei1 (see Supplementary Fig. 6 for a full list of interacting residues). Notably, this interface exhibits high evolutionary conservation. Its functional significance is underscored by the previously reported activity-impaired kei1-1 mutant that contains F103I substitution11. Consistently, our co-immunoprecipitation assays confirmed that this altered Kei1 significantly disrupts complex assembly (Fig. 2J).

Strikingly, our structural analysis also revealed a direct role for membrane lipids in stabilizing the complex. The high-resolution cryo-EM map revealed several tightly bound lipid molecules at the cytosolic membrane leaflet of the inter-subunit interface, including one phospholipid and two lipid alkyl chains (Fig. 2C, Supplementary Fig. 4A). Among the modeled lipids, the two lone alkyl chains are likely derived from phospholipid with disordered head group, possibly due to a lack of stabilizing interactions in this region as documented in several other structural studies43-45. The assignment of phospholipid is supported by multiple lines of evidence. No exogenous phospholipids were added during purification, yet the observed phospholipid-like density could not be reasonably fitted to either of the detergents used—neither LMNG (with its bulky maltose headgroup) nor CHS (with its distinct sterol scaffold) (Supplementary Fig. 4A, 4D). Moreover, we performed lipidomics analysis via mass spectrometry of lipids co-purified with IPC synthase. This result confirmed the presence of several lipid species, most notably phospholipid, providing additional support for our lipid assignment (Supplementary Fig. 4C). Notably, the phospholipid is deeply embedded between Aur1 and Kei1, filling a ~12 Å inter-subunit gap (Fig. 2F-G). Its headgroup is poised to contact several highly conserved charged residues, including K274 and K314 of Aur1, and H75 of Kei1 (Fig. 2F-H). Its alkyl tails are nestled among hydrophobic residues from TM6-TM7 of Aur1 and TM1-TM2 of Kei1 (Fig. 2F). These observations suggest a lipid-mediated mechanism for heterodimer stabilization. To test this model, we introduced alanine substitutions at key lipid-coordinating residues. Co-immunoprecipitation assays revealed that K274A or K314A mutations in Aur1 markedly impaired complex formation (Fig. 2I), while alanine substitution of Kei1 H75 also compromised assembly (Fig. 2J).

The reaction chamber and catalytic mechanism
Opposite its Kei1 binding site, the core transmembrane domain (TM3–TM8) of catalytic subunit Aur1 forms a groove-shaped reaction chamber that is laterally accessible to the surrounding lipid bilayer (Fig. 3A-B). This membrane-embedded chamber is highly conserved among Aur1 homologs from yeast and pathogenic fungi (Fig. 3B). It is primarily delineated by TM3–TM6 and capped on the luminal side by two short lateral helices (EL3-h2 and EL3-h3) within the EL3 loop. A notable feature of the chamber architecture is the kinked TM4 by a conserved ‘Gly-Ala-Pro’ loop (159-Gly-Ala-Pro-161), substantially expanding the chamber near the luminal membrane leaflet. In addition, TM6 is unusually short (only 15-residue long) and transitions into EL3 before reaching the lumen, allowing the chamber to extend deeper into the protein core (Fig. 3A-B).

In the cryo-EM map of ceramide-bound IPC synthase, a distinct non-protein density was observed within the reaction chamber (Fig. 3A). Its shape and location are consistent with a bound ceramide molecule, the only exogenous lipid added throughout the purification procedure (see Methods section). The well-defined density closely matches the ceramide structure and cannot be reasonably fitted to the two purification detergents used (Supplementary Fig. 4B and 4D): neither to CHS with a distinct sterol scaffold, nor to LMNG with bulky maltose-based headgroup that is also sterically incompatible with the narrow, groove-shaped active site. One alkyl chain of the bound ceramide is embedded in the membrane-facing chamber, contacting conserved hydrophobic residues from TM3b, TM4b and TM5 (Fig. 3C, Supplementary Fig. 4B). The other chain lies between two conserved hydrophobic residues, F162 on TM4b and F158 in the preceding loop (Fig. 3C). The hydroxyl group of ceramide extends toward the luminal base of the reaction chamber and is situated near the conserved residue H255 (Fig. 3D).

Our analysis revealed that IPC synthase Aur1 shares a conserved fold with canonical LPT family enzymes (Supplementary Fig. 5), whose catalytic mechanism usually involves a His-His-Asp triad42. Guided by this, we identified five conserved candidate catalytic residues positioned adjacent to the ceramide hydroxyl group in our structure (Fig. 3E-F, Supplementary Fig. 7): H255 from EL3; H293, H294 and D298 from EL4; H157 from the loop preceding TM4b. Considering that Ser-His-Asp catalytic triad are also characteristic of several lipases/transferases46,47, we additionally evaluated S253 from EL3, due to its spatial proximity to the putative active site (Fig. 3E). To assess the functional relevance of these residues, we generated single-point mutants of each residue and assessed their enzymatic activity in vitro. All mutants exhibited wild-type-like expression levels, subunit stoichiometry, and size-exclusion chromatography profiles, indicating their proper folding (Supplementary Fig. 2B-D). However, the functional consequences of these mutations varied markedly. Mutations H255A, H294A, and D298A each completely abolished IPC synthase activity (Fig. 3G-H), supporting them as essential constituents of the catalytic triad. In contrast, other active-site mutations (H293A, H157A, and S253A) led to varying degrees of partial activity loss, with H157A, located most distantly from the catalytic triad, showing the least impairment (~50% residual activity) (Fig. 3G-H). To specifically evaluate the potential catalytic role of S253, we generated S253C and S253T mutants. Activity assays revealed that both mutants retained substantially higher activity than S253A (Supplementary Fig. 8A), suggesting that S253 is not directly involved in catalysis. This conclusion was further supported by structural comparison: the Aur1 active site—containing S253 and the His-His-Asp triad (H255, H294, D298)—showed no equivalence to the canonical Ser-His-Asp triad found in several enzymes46-48 (Supplementary Fig. 8B-D). Collectively, this gradation of mutational effects suggests that while the H255/H294/D298 triad is indispensable for core enzymatic function, neighboring residues such as H157, S253, and H293 also contribute to activity. These latter auxiliary residues may help maintain optimal catalytic geometry, fine-tune substrate orientation, or stabilize the active-site environment, though their precise mechanistic roles await further elucidation.

We demonstrated the strict substrate specificity of fungal IPC synthase for phosphatidylinositol (PI) over phosphatidylethanolamine (PE) or phosphatidylcholine (PC) (Fig. 1C). To locate the PI-binding site and investigate the basis of this specificity, we performed molecular docking of PI into the Aur1 structure and showed that PI occupies the reaction chamber identified in our ceramide-bound IPCS structure49 (Supplementary Fig. 9A). Notably, the inositol group of PI is nestled deep within a pocket formed by transmembrane helices TM5–TM7 and luminal loops EL3 and EL4 (Supplementary Fig. 9B-C). The shape of this pocket appears to optimally accommodate the bulkier inositol group of PI (Supplementary Fig. 9C-D), likely underlying the observed phospholipid preference (Fig. 1C). Importantly, in this binding pose, the phosphate group of PI is positioned in close proximity to H255 and H294 (Supplementary Fig. 9D), both identified as essential catalytic residues (Fig. 3G-H).

Based on our structural and functional evidence, we propose a two-step catalytic mechanism for IPC synthesis (Fig. 3I). In the first step, D298 activates H294 to initiate a nucleophilic attack on the phosphate group of PI, leading to the formation of an enzyme–phosphoinositol intermediate and the release of diacylglycerol (DAG). In the second step, H255 activates the hydroxyl group of an incoming ceramide, which attacks the intermediate and transfers the phosphorylinositol moiety to ceramide, ultimately forming inositol phosphorylceramide (IPC). Notably, this mechanism is consistent with of canonical LPT enzymes38-40,42, yet it differs markedly from that of SMSr41 (see Discussion for further details), indicating a catalytic divergence within the conserved H-H-D catalytic triad.

Given that IPC synthase has been highlighted as a potential target for anti-protozoal drug development30,32, we extended our analysis to TbSLS1, the Trypanosoma brucei ortholog of Aur131. Despite the overall low sequence conservation between Aur1 and TbSLS1, we found the core domain of Aur1 (TM3-TM8) resembles the overall topologies of TbSLS1, with a root-mean-square-deviation (RMSD) of 5.2 Å (Supplementary Fig. 10A-B). Notably, comparison of their ceramide-binding pockets revealed substantial divergence, largely due to structural adaptions in transmembrane segments TM1-TM2 of TbSLS1 relative to TM3-TM4 of Aur1 (Supplementary Fig. 10A). In contrast, their catalytic centers show remarkable conservation, with close spatial alignment of the catalytic triad residues (Aur1 H255/H294/D298 and TbSLS1 H210/H253/D257), as well as their surrounding structural elements, including TM6–TM8 in Aur1 and TM4–TM6 in TbSLS1 (Supplementary Fig. 10C-D). Together, these observations suggest that IPC synthases from fungi and protozoan parasites, despite evolutionary divergence, retain a conserved catalytic architecture centered on the His–His–Asp triad, while allowing structural flexibility in substrate recognition.

Molecular basis of IPC synthase inhibition by Aureobasidin A and implications for drug resistance
Aureobasidin A (AbA) is an important antifungal agent that targets fungal IPC synthase9 (Fig. 4A). Our biochemical assays show that detergent-purified IPC synthase retains robust inositol phosphoryl transferase activity and exhibits high sensitivity to AbA, confirming the drug’s potent and specific action (Fig. 4B). Notably, our engineered ALFA-tagged IPC synthase constructs preserved wild-type level of catalytic efficiency and AbA sensitivity regardless of Nb binding status (Fig. 4B), establishing their suitability for subsequent structural investigations.

Based on the functionally validated Nb-bound ALFA-tagged complex, we aimed to determine cryo-EM structure of IPC synthase in complex with AbA drug. Through systematic optimization, we found that extending the AbA incubation time from 1 to 3 hours substantially enhanced drug occupancy, ultimately enabling us to resolve the complex structure at an overall resolution of 3.5 Å (Supplementary Fig. 11A–H). The resulting map showed well-defined density for the intact AbA molecule (Supplementary Fig. 11I). AbA binds at the luminal bottom of the enzyme’s reaction chamber, with binding site framed by TM3b and TM4a on two sides and enclosed from below by the extended luminal loop EL3 and its two constituent helices, EL3-h2 and EL3-h3 (Fig. 4C). Notably, H157 and F158, located on the loop preceding TM4a, appear as a key anchoring site for the drug: H157 forms a hydrogen bond with the HoMeVal moiety of AbA, while F158 engages in a π-stacking interaction with AbA Phe residue (Fig. 4D). Additional stabilization is provided by extensive van der Waals contacts with surrounding hydrophobic residues, including W98, V99, Y102, and F103 on TM3b; L254, V246, F241, A240, Y237, M236, and L225 on EL3 and its two constituent helices EL3-h2 and EL3-h3; and Y153 on TM4a (Fig. 4D).

The AbA occupies the reaction chamber and its upper portion sterically overlaps with part of the ceramide, indicating that AbA competes directly with this substrate (Fig. 4E-F). Consistently, the AbA density extends toward the H-H-D catalytic triad and occludes the catalytic residue H255 and H294, effectively blocking access to the active site (Fig. 4G). Comparison between the ceramide- and AbA-bound structures revealed that while the Kei1 subunit and the C-terminal region of Aur1 (TM5–8) remain largely unchanged, pronounced rearrangements occur in Aur1 elements surrounding the drug, including TM3, EL3, the TM4a–TMA4b linker, and the adjacent TM2 (Fig. 4E). Detailed inspection identified key conformational adaptations in four aromatic residues that directly contact AbA (Fig. 4F, Supplementary Fig. 11J-L): Y102 (TM3), F158 (TM4a–TM4b linker), and F241 (EL3-h3) reorient their side chains to relieve steric clashes and form favorable contacts with AbA, whereas Y153 (TM4a–TM4b linker) rotates to establish an additional interaction. Notably, the reorientation of F158 creates a favorable π-stacking interface with AbA but would sterically clash with ceramide (Fig. 4F), suggesting a mutually exclusive binding mode. Moreover, the AbA-induced downward rotation of Y102 appears to act as a lever, propagating into a broader structural shift to drive inward movement of TM3 and its connected TM2 toward the drug (Fig. 4E-F). Together, these structural rearrangements indicate a coordinated allosteric response that may underlie AbA binding and its competitive inhibition mechanism.

Our structural insights also shed light on the molecular basis of drug resistance related to IPC synthase. AUR1 mutations F158Y and H157Y/L137F have been identified to confer AbA-resistant phenotype9,27,28. F158 and H157 cluster around our identified AbA binding pocket, contributing directly to AbA binding (Fig. 4D). Biochemical characterization confirmed that F158Y and H157Y/L137F mutants maintain basal IPC synthase activity but show markedly reduced AbA sensitivity (Fig. 4H). Further functional dissection revealed that H157Y - which directly disrupts the inhibitor-binding pocket - confers strong resistance, while the distantly located L137F mutation retains drug sensitivity (Fig. 4H). These observations underscore the critical role of H157 and F158 in both AbA binding and resistance acquisition.

Methods
Nanobody expression and purification
The coding sequence encoding the ALFA-binding nanobody (Nb) was subcloned into the pSecTag2 vector with an N-terminal 6×His tag. For expression, Human embryonic kidney (HEK) 293F cells (Invitrogen) were cultured in Freestyle 293 medium at 37°C under 5% CO₂ in a shaking incubator at 120 r.p.m. to a density of ~2 × 10⁶ cells per milliliter before transfection with expression plasmid. For 1-liter culture, 1 mg plasmid was pre-mixed with 3 mg polyethyleneimine (PEI, Polysciences, 23966-100) in 30ml of fresh medium at room temperature for 30 minutes for transfection. After transfection, the cells were cultured at 37°C for 48 hours.

The secreted nanobody protein was purified from the culture medium using Ni-NTA affinity chromatography. After centrifugation at 4,000g for 10 minutes, the collected medium was applied to Ni-NTA resin (Thermo Fisher). After washing with washing buffer (25 mM HEPES pH 7.5, 150 mM NaCl, 20 mM imidazole), the nanobody protein was eluted with washing buffer supplemented 500 mM imidazole. The eluate was further purified by gel filtration chromatography in buffer: 25 mM HEPES pH 7.5 and 150 mM NaCl. The central fractions of monodisperse peak were collected and concentrated for later preparation of the nanobody-bound form of IPC synthase.

IPC synthase expression and purification
The codon-optimized full-length cDNA constructs of S. cerevisiae AUR1 (UniProt ID: P36107) and KEI1 (UniProt ID: Q06346) were subcloned into the pCAG vectors. A FLAG tag was fused to the C-terminus of Aur1 and the ALFA tag was inserted to replace the Kei1 loop (residues 127–131). Human embryonic kidney (HEK) 293F cells (Invitrogen) were cultured in Freestyle 293 medium at 37°C under 5% CO in a shaking incubator at 120 r.p.m. For the expression of IPC synthase, cells at a density of ~2 × 10⁶ cells per milliliter were co-transfected with AUR1 and KEI1 plasmids at a 1:1 mass ratio. For 1-liter cell culture, ~2 mg of plasmids was pre-mixed with 6 mg of polyethyleneimine (PEI, Polysciences, 23966-100) in 40 mL of fresh medium at room temperature for 30 minutes before transfection. After transfection, the cells were cultured for 48 hours at 37°C and harvested by centrifugation at 4,000 g for 10 min.

For the purification of IPC synthase complex, the harvested cells were resuspended in lysis buffer: 25 mM HEPES pH 7.5, 150 mM NaCl and 1% protease-inhibitor cocktail (Roche), and lysed by sonication on ice. The membrane was solubilized by the addition of 1% (w/v) lauryl maltose neopentyl glycol (LMNG, Anatrace) and 0.1% cholesteryl hemisuccinate Tris salt (CHS, Anatrace) at 4°C for 2 hours. After centrifugation at 20,000 g for 1 hour, the supernatant was collected and loaded into pre-equilibrated anti-Flag M2 affinity resin (Sigma). The resin was washed with washing buffer containing 25 mM HEPES pH 7.5, 150 mM NaCl, 0.01% LMNG and 0.001% CHS, and eluted with washing buffer supplemented with 250 μg/ml 3 × FLAG peptide.

To obtain the nanobody-bound IPC synthase, the purified ALFA-tagged IPC synthase was mixed with purified nanobody at a 1:1.2 molar ratio on ice for 1 hour. The resulting complex was further purified by gel filtration chromatography (Superose 6 10/300 GL, GE Healthcare) with buffer containing 25 mM HEPES pH 7.5, 150 mM NaCl, 0.01% LMNG and 0.001% CHS. The preparation of the ceramide-bound IPC synthase sample follows the procedure described above, with ceramide (APExBIO) at a final concentration of 10 µM added during membrane solubilization and in all subsequent purification buffers.

Cryo-EM data acquisition
For single-particle cryo-EM analysis, a 3 µl aliquot of the ceramide-bound IPC synthase (ALFA-tagged with bound nanobody) at a concentration of 5mg/ml was applied to glow-discharged Quantifoil carbon grids (R1.2/1.3 Au, 300 mesh). The grids were blotted for 4 s at 100% humidity and vitrified in liquid ethane using a FEI Vitrobot Mark IV. For the AbA-bound IPC synthase sample, IPC synthase (ALFA-tagged with bound nanobody) was incubated with 50µM AbA (Med Chem Express, Cat No: HY-P1975) on ice for 1 to 3 hours before freezing the cryo-EM grids.

Cryo-EM data were acquired on a Titan Krios electron microscope (Thermo Fisher) operated at 300 kV, equipped with a Gatan K3 Summit direct electron detector and a GIF Quantum energy filter. Automated data collection was performed using EPU software (Thermo Fisher). Micrographs were recorded under super-resolution counting mode with a physical pixel size of 0.74 Å or 0.926 Å. Defocus values ranged from -1.1 μm to -2.8 μm, with a total accumulated dose of 50 electrons per Å2.

Image processing and 3D reconstruction
The dose-fractionated movies were subjected to motion correction and dose weighting using MotionCor2 51. All subsequent image processing steps were performed using cryoSPARC (v4.7.0)52.

For the IPC synthase-ceramide dataset, 8,250 drift-corrected micrographs were imported into cryoSPARC for patch CTF estimation and template picking. A total of 7,447,058 particles were extracted 4 × binning and subjected to multiple rounds of 2D classification. Ab initio maps were generated and used as initial references for heterogeneous refinement. A cleaned subset of 1,006,952 particles were selected and re-extracted without binning. These particles were further subjected to iterative rounds of ab initio reconstruction and heterogeneous refinement. A final subset of 162,123 particles was selected for non-uniform refinement, resulting a map with an overall resolution of 3.5 Å. A detailed flowchart of the data process is presented in Supplementary Fig. 3. The overall resolution estimates were based on the gold standard Fourier shell correlation (FSC) 0.143 criterion calculations53. The local resolution distribution was estimated using cryoSPARC.

For AbA-incubated IPC synthase sample, we collected two datasets after one-hour and three-hour incubation periods, respectively. Processed analogously to the ceramide-incubated dataset, it was found that the three-hour incubation yielded a structure with higher AbA occupancy. For this long incubation dataset, a total of 10,346 movie stacks were processed and the final reconstruction map with an overall resolution of 3.5 Å from 228,212 particles was obtained. A detailed flowchart of the data processing is presented in Supplementary Fig. 11.

Model building and refinement
The atomic model of ceramide-bound IPC synthase complex was first built. An initial model was generated using AlphaFold3 and rigid-body fitted into the cryo-EM map using Chimera54,55. It was further improved by manual rebuilding in Coot56, guided by the well-defined densities for transmembrane helices and distinctive side-chain features for bulky residues such as Phe, Tyr, Trp, and Arg. Iterative real-space refinement was conducted in PHENIX, with secondary structure and geometry restraints applied57. The resulting model served as the template for building the AbA-bound IPC synthase structure, following a similar procedure. The final models were assessed using MOLPROBITY58. Chimera, ChimeraX59 and PyMOL (Schrödinger, LLC.) were used to prepare the illustrated figures. Statistics of the 3D reconstructions and model refinements were provided in Supplementary Table 1.

Construction of conditional AUR1 or KEI1 knockdown strains
To circumvent the lethality associated with AUR1 deletion, we constructed a conditional AUR1 knockdown strain (YPH499-HIS-GAL-AUR1) by placing AUR1 expression under the control of the tightly regulated GAL1 promoter, which is repressed in glucose-containing medium30. A HIS3-selectable GAL1 promoter replacement cassette (HIS-GAL1) was assembled and introduced into the YPH499 strain via homologous recombination to replace the endogenous AUR1 promoter30,60. Transformants were selected on SGal-His medium (synthetic dropout medium lacking histidine, supplemented with 5% galactose). Correct integration of the HIS-GAL1 cassette was verified by genomic PCR and DNA sequencing, yielding the conditional knockdown YPH499-HIS-GAL-AUR1 strain.

To construct the conditional KEI1 knockdown strain (YPH499-HIS-GAL-KEI1), the endogenous KEI1 promoter was similarly replaced with the HIS-GAL1 cassette using the same strategy. Transformants were selected on SGal-His medium and verified by genomic PCR and DNA sequencing.

Spot assay for growth phenotypic analysis of AUR1 and KEI1 variants
To assess the functional contribution of AUR1 to IPC synthase, the conditional AUR1 knockdown strain (YPH499-HIS-GAL-AUR1) was transformed with either the empty vector pRS424, pRS424 carrying wild-type AUR1, or pRS424 carrying AUR1 variants. The parental YPH499 strain transformed with empty pRS424 was used as the control. Transformants were selected on SLGal-Leu (synthetic dropout medium lacking leucine and containing 5% galactose). Spot assays were performed by first cultivating the cells in liquid SGal-Leu at 30°C with shaking (180 r.p.m.) until reaching exponential phase (OD600 = 0.5). Four 10-fold serial dilutions were prepared, and 5 μL of each dilution was spotted on SGal-Leu or SGlu-Leu (containing 5% glucose instead of galactose). Plates were incubated at 30 °C for 72 hours, and photographed with the 5200CE Image System (TANON). Each assay was performed in triplicate with similar results.

To investigate the role of KEI1, the same assay was carried out using the conditional KEI1 knockdown strain (YPH499-HIS-GAL-KEI1), transformed with either empty pRS424, pRS424 carrying wild-type KEI1, or the parental YPH499 with pRS424 as control. The procedure for growth assays was identical to that described above, and each experiment was independently repeated three times with similar results.

Activity assay via thin-layer chromatography
AUR1 or KEI1 point mutations were introduced by site-directed mutagenesis and confirmed by sequencing. Wild-type and mutated IPC synthases with indicated Aur1 or Kei1 mutations were expressed and purified as described above. The IPC synthase activity was assessed by monitoring fluorescent product formation via incorporation of inositol phosphate from phosphatidylinositol (PI) into NBD-labeled ceramide31. 20µM phosphatidylinositol PI (MCE) or phosphatidylcholine PC (MCE) or phosphatidylethanolamine PE (MCE) was added to 200µl reaction mixture in total containing 25 mM HEPES pH 7.5, 150 mM NaCl and 40µM C6-NBD-ceramide (MCE). The reaction was initiated by adding 2 µg of purified IPCS synthase variants. The reaction was carried out at 30˚C for 30min or 120min. Reactions were terminated by adding 600 μl of chloroform: methanol (2:1, v/v) mixture. Following centrifugation at 1,300g for 2 min, the organic phase was collected and dried under high-purity nitrogen. The reaction products were analyzed by thin-layer chromatography (TLC silica gel 60 F254, Merck) in developing solvent containing chloroform: acetone: acetic acid: methanol: H2O (at volume ratio of 50:20:10:10:5). Fluorescent bands were visualized using a Champ Chemi 610 imaging system (Sinsage).

Immunoprecipitation and immunoblotting
For Aur1 and Kei1 interaction studies, AUR1 and KEI1 genes were subcloned into the pCAG vectors, with a C-terminal FLAG tag fused to AUR1 and an N-terminal Strep tag fused to KEI1. Mutations were introduced via site-directed mutagenesis and confirmed by DNA sequencing. HEK293T cells were cultured and co-transfected with 10 µg of expression plasmids per 10-cm dish using PEI (Polysciences, 23966-100), following the manufacturer’s instructions. For co-transfection, wild-type or mutant AUR1 and KEI1 plasmids were added at a 1:1 mass ratio. After transfection, the cells were cultured at 37 °C for 48 h, harvested and washed with TBST buffer (Tris Buffered Saline with Tween-20).

The harvested cells were resuspended in lysis buffer: 1% (w/v) lauryl maltose neopentyl glycol (LMNG, Anatrace), 0.1% cholesteryl hemisuccinate Tris salt (CHS, Anatrace), 25 mM HEPES pH 7.5, 150 mM NaCl and 1% protease-inhibitor cocktail (Roche), lysed by sonication on ice and centrifuged at 20,000 g for 30 minutes at 4 °C. Immunoprecipitations were performed by subjecting supernatants to either anti-Flag M2 affinity resin (Sigma) or Strep-Tactin resin (IBA), with bound beads washed three times with lysis buffer. Samples were resuspended in SDS-loading buffer and analyzed by 15% SDS-PAGE. Proteins were transferred to PVDF membranes and immunoblotted with the indicated antibodies. Membranes were imaged using the 5200CSE Imaging System (TANON). Sources and dilutions for primary antibodies were as follows: anti-Flag antibody for Flag-tagged Aur1 (Proteintech, 66008-4-lg, 1:5000); anti-Strep antibody for Strep-tagged Kei1 (Solarbio, K114183P, 1:5000). The secondary antibodies used were anti-mouse IgG (Proteintech, SA00001-1, 1:10000). All antibodies were diluted in 3% (w/v) BSA in TBST.

Lipidomics analysis
Lipids were extracted from the protein solution using 750 µL of a chloroform:methanol:Milli-Q water mixture (3:6:1, v/v/v). The mixture was homogenized and centrifuged at 231 g for 1 h at 4 °C. After centrifugation, 350 µL of deionized water and 250 µL of chloroform were added to induce phase separation. The sample was centrifuged again, and the lower organic phase containing lipids was collected into a clean tube. To maximize lipid recovery, 450 µL of chloroform was added to the remaining aqueous phase, and lipid extraction was repeated once. The combined organic phases were dried using a SpeedVac under OH mode, and the resulting lipid extracts were stored at −80 °C for further analysis. The upper aqueous phase and the residual protein pellet were dried in a SpeedVac under H2O mode. The dried protein pellet was subsequently used for total protein quantification. Lipidomic analysis was performed using a Shimadzu Nexera 20AD-HPLC/ExionLC-AD system coupled to a Sciex QTRAP 6500 PLUS mass spectrometer. Polar lipid classes were separated by normal phase (NP) HPLC using a TUP-HB silica column (150 × 2.1 mm, 3 µm). The mobile phases consisted of (A) chloroform:methanol:ammonium hydroxide (89.5:10:0.5, v/v/v) and (B) chloroform:methanol:ammonium hydroxide:water (55:39:0.5:5.5, v/v/v/v). Multiple reaction monitoring (MRM) transitions were optimized for the comparative quantification of various polar lipid species. Quantification of individual lipid species was performed relative to spiked internal standards, including d9-PC32:0(16:0/16:0), d9-PC36:1p(18:0p/18:1), d7-PE33:1(15:0/18:1), d9-PE36:1p(18:0p/18:1), d31-PS(d31-16:0/18:1), d7-PA33:1(15:0/18:1), d7-PG33:1(15:0/18:1), d7-PI33:1(15:0/18:1), C17-SL, d5-CL72:8(18:2)₄, Cer d18:1/15:0-d7, C12:0 Cer-1-P, d9-SM d18:1/18:1, C8-GluCer, C8-GalCer, d3-LacCer d18:1/16:0, Gb3 d18:1/17:0, Pd7-LPC18:1, d7-LPE18:1, C17-LPI, C17-LPA, C17-LPS, C17-LPG, d17:1 Sph, and d17:1 S1P (all from Avanti Polar Lipids). GM3-d18:1/18:0-d3 was obtained from Matreya LLC. Free fatty acids were quantified using d31-16:0 (Sigma-Aldrich) and d8-L20:4 (Cayman Chemical) as internal standards. Free cholesterol and cholesteryl esters were analyzed in atmospheric pressure chemical ionization (APCI) mode using a Jasper HPLC system coupled to a Sciex 4500 MD mass spectrometer, as previously described. Quantification was performed with d6-cholesterol and d6-C18:0 cholesteryl ester (CE) (CDN Isotopes) as internal standards.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript by Hingjun Yu and colleagues describes the cryo-EM structure of the fungal inositol phosphorylceramide (IPC) synthase from S. cerevisiae, the enzyme responsible for inositol phosphorylceramide synthesis. IPC is a key component of fungal membranes, and is absent from mammalian cells. It is therefore an important anti-fungal drug target. The authors find that IPC synthase is a heterodimer of the Aur1 and Kei1 subunits. The dimer interface is stabilised by a phospholipid and two lipid alkyl chains. The authors identify H255, H294 and D98 of the Aur1 subunit as being essential for catalysis and suggest that these three residues form a catalytic triad. A HHD triad is also found in other fungal IPC synthases despite low sequence similarity. The authors determined a structure of the complex with the antifungal agent aureobasidin (AbA) and suggest AbA acts as a competitive inhibitor to block ceramide binding. Finally, the authors test models of the Aur1-Kei dimer interface, phospholipid binding and catalysis using mutagenesis.\n\nIn general this is an interesting paper. The figures are clear and the cryo-EM work is sound. Use of the ALFA-binding nanobody to increase size and help alignment was innovative.\n\n1. When describing the structure, the authors do not compare and contrast with other membrane protein structures. A brief comment on the relatedness of IPC synthase to other membrane protein structures would be informative and useful, or is this a novel structure?\n2. The identification of a phospholipid and two alkyl chains at the Aur1-Kei subunit interface is of interest. However, the two lone alkyl chains are a little unusual. Are these attached to a phospholipid whose head group is disordered?\n3. The authors suggest that the catalytic triad of His255, His294 and Asp298 catalyse inositol phosphate transfer. A reaction schematic to show the catalytic reaction mechanism would be helpful.\n4. Where is the binding site for phosphatidylinositol in Aur1? How is IPC synthase specific for phosphatidylinositol and not able use phosphatidylcholine (PC) or phosphatidylethanolamine?\n5. The authors suggest that in the presence of AbA, the aromatic side chain of Phe 158 rotates 90°. It is not entirely obvious that this would be clear at 3.5 A resolution. The authors should show cryo-EM density that supports this conclusion.\n6. I didn’t see reference in Supplementary Fig. 2.\n7. ‘Comparation’ is not an English word", "reviewer_2": "This study presents cryo-EM structures of IPC synthase in complex with its substrate ceramide and the inhibitor AbA. The authors successfully identified the substrate-binding cavity and the AbA inhibitor pocket, providing critical insights into the enzyme’s catalytic mechanism. Complementary biochemical and cell-based assays robustly validate the essential role of IPC synthase catalytic activity in fungal sphingolipid biosynthesis. Notably, the resolution of a high-resolution structure for this ~71 kDa membrane protein—achieved through the combination of an ALFA epitope tag and a high-affinity nanobody—represents a significant methodological advancement in the field of membrane protein structural biology.\n\nOverall, the topic is of interest given the essential role of IPC synthase in fungal sphingolipid metabolism and its potential as an antifungal drug target. However, I note that a closely related study by Wu et al. (Nature Communications, 2025) has already reported high-resolution structures of the Aur1–Kei1 complex bound to AbA, identifying the catalytic H–H–D triad, key resistance mutations, and the dimeric architecture. Therefore, my comments will focus more on the novelty part of this work relative to Wu’s paper.\n\nMajor points:\n1. A major concern with the current manuscript is the insufficient acknowledgment of critical related literature. Wu et al. reported cryo-EM structures of the S. cerevisiae Aur1–Kei1 complex bound to Aureobasidin A, describing the same protein target and inhibitor, and this work should be explicitly cited in both the Introduction and Discussion. The authors need to carefully compare their findings with those of Wu et al. to highlight both consistencies (such as the catalytic H–H–D triad, AbA resistance mutations, and overall architecture) and differences (including the ceramide-bound state and lipid-mediated assembly) to avoid overstating novelty. In addition, because IPC synthase belongs to the sphingomyelin synthase (SMS) family, readers would benefit from a broader discussion that integrates previous structural and biochemical studies on SMS enzymes. Such a comparison could illuminate how the H–H–D catalytic triad and substrate positioning in IPC synthase relate to SMS enzymes, and whether conserved or divergent features explain differences in substrate specificity. Incorporating these key references and analyses will provide proper context, clarify the unique contributions of the present study, and strengthen its impact.\n2. The capture of ceramide in the active site is indeed a valuable addition not present in Wu et al., and the authors support their proposed structural model and catalytic mechanism with mutagenesis of key residues in the reaction chamber, which resulted in reduced activity. However, to strengthen this conclusion, they should provide evidence that the observed effects are not due to global misfolding of the enzyme but rather specific to the catalytic center—for example, by assessing the stability or expression of the purified mutant proteins with size-exclusion FPLC. Moreover, the H157A mutant retains ~50% activity, showing the least impairment among the catalytic site variants; a brief discussion of why different mutations within the catalytic center yield varied activity reductions would be helpful. Finally, it remains uncertain whether the ceramide density reflects a true catalytic intermediate or a trapped state introduced during purification and exposure to detergents or exogenous lipids. The authors should discuss the likelihood of ceramide incorporation under their preparation conditions and address how well this state represents in vivo substrate binding.\n3. Another point that warrants attention is the interpretation of the lipid-mediated heterodimer stabilization of the Aur1–Kei1 complex. The identification of a phospholipid density bridging the interface is intriguing. However, several concerns remain. First, the exact identity of the lipid is not determined, and it is unclear whether this represents a physiologically specific cofactor, a detergent-derived molecule, or a general phospholipid from the purification system. The authors should clarify the lipid assignment and, if possible, provide mass spectrometry or biochemical analysis to strengthen this claim. Second, it is important to discuss whether this lipid-mediated interaction is unique to IPC synthase or could represent a more general feature among the sphingomyelin synthase family. Finally, the authors should place their findings in context with prior work: Gong et al. noted lipid-like densities at the interface but did not ascribe functional significance, so a more explicit comparison would help delineate what is novel here. Overall, while the lipid-mediated stabilization model is a potentially important advance, its physiological specificity and mechanistic implications require more careful discussion and validation.\n4. In the AbA-bound IPC synthase complex (Fig. 4), the reported unassigned density within the catalytic cavity does not convincingly fit the 3D structure of the AbA inhibitor. This discrepancy raises critical concerns regarding the interpretation of the AbA-bound state. Specifically: (1) does the IPCS–ALFA construct reduce AbA binding affinity compared to the wild-type enzyme, and (2) could AbA be hydrolyzed within the catalytic cavity, thereby explaining the incomplete density? If either hypothesis is plausible, the authors should provide biochemical validation: for example, quantifying AbA binding affinity (e.g., Kd) for IPCS–ALFA versus wild-type using SPR or ITC, or testing for AbA hydrolysis by LC–MS to detect cleavage products. If neither explanation holds, the authors must rigorously demonstrate that the unassigned density corresponds to bound AbA rather than endogenous lipids co-purified with the protein. The current assertion that AbA binding is “dynamic” is not sufficiently supported. Rigorous validation of this density is essential to substantiate the proposed inhibition mechanism.\n\nSome minor points:\nFig. 2B (SDS-PAGE) and Figs. 2I-J (Co-IP):\nIn the SDS-PAGE results of Fig. 2B (lane for purified Aur1), what do the high-molecular-weight bands above the Aur1 band represent? Similar bands are observed in the co-immunoprecipitation assays (Figs. 2I-J). Do they correspond to glycosylated forms of Aur1? If so, glycan density should be detectable in your cryo-EM maps, why is this density absent in the manuscript? What is the biological significance of Aur1 glycosylation?\nLine 246: While the Ser residue in the canonical Ser–His–Asp catalytic triad typically provides the nucleophilic hydroxyl oxygen essential for catalysis, the specific role of S253 (located in EL3) in Aur1 during sphingolipid synthesis remains unclear. Notably, mutagenesis of S253 does not completely abolish enzymatic activity, raising questions about whether it truly functions as the nucleophile in the reaction. To clarify this discrepancy, the authors could strengthen their study by (i) providing a comparative analysis of S253’s position and function relative to the catalytic Ser residues in classical triads, and (ii) performing additional mutagenesis, such as S253C or S253T substitutions, to quantify enzymatic activity and evaluate whether side-chain chemistry influences catalysis. Such data would help resolve whether S253 is directly catalytic or plays an auxiliary structural/positioning role in Aur1’s mechanism.", "reviewer_3": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Point-to-point responses to the reviewers’ comments\nWe greatly thank the insightful and valuable suggestions from reviewers. Our responses\nbelow are in blue.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThe manuscript by Hingjun Yu and colleagues describes the cryo-EM structure of the\nfungal inositol phosphorylceramide (IPC) synthase from S. cerevisiae, the enzyme\nresponsible for inositol phosphorylceramide synthesis. IPC is a key component of\nfungal membranes, and is absent from mammalian cells. It is therefore an important\nanti-fungal drug target. The authors find that IPC synthase is a heterodimer of the Aur1\nand Kei1 subunits. The dimer interface is stabilised by a phospholipid and two lipid\nalkyl chains. The authors identify H255, H294 and D98 of the Aur1 subunit as being\nessential for catalysis and suggest that these three residues form a catalytic triad. A\nHHD triad is also found in other fungal IPC synthases despite low sequence similarity.\nThe authors determined a structure of the complex with the antifungal agent\naureobasidin (AbA) and suggest AbA acts as a competitive inhibitor to block ceramide\nbinding. Finally, the authors test models of the Aur1-Kei dimer interface, phospholipid\nbinding and catalysis using mutagenesis.\nIn general this is an interesting paper. The figures are clear and the cryo-EM work is\nsound. Use of the ALFA-binding nanobody to increase size and help alignment was\ninnovative.\nResponse: We thank the reviewer for the support and the valuable comments. They are\nso helpful to improve our manuscript. We have revised our manuscript accordingly.\n1. When describing the structure, the authors do not compare and contrast with other\nmembrane protein structures. A brief comment on the relatedness of IPC synthase to\nother membrane protein structures would be informative and useful, or is this a novel\nstructure?\nResponse: We thank the reviewer for this insightful suggestion. In response, we have\nadded a section comparing the IPC synthase structure with other known membrane\nproteins identified by DALI analysis [1]. This analysis not only underscores the\nstructural novelty of both Aur1 and Kei1 but also reveals a conserved LPT (lipid\nphosphatase/phosphotransferase) fold within the catalytic subunit Aur1. The related\ndescription has been incorporated into the revised manuscript (Lines 190-219 on Page\n7-8; Supplementary Fig. 5) as follows:\n“A DALI server search for the regulatory subunit Kei1 of the IPC heterodimer did not\nyield any structurally similar homologs [1]. In contrast, for the catalytic subunit Aur1,\nalthough the overall architecture of Aur1 differs from previously reported structures, a\nDALI homology search identified several membrane-protein structures that resemble a\nportion of Aur1’s transmembrane topology (residues 143-314) (Supplementary Fig.\n5). These include (in a descending order of Z-score): Lipid A 1-Phosphatase (LpxE),\nGlucose-6-phosphatase (G6PC1), Phosphatidylglycerophosphatase B (PgpB), and\nSphingomyelin synthase-related protein 1 (SMSr) [2-5]. Despite their low sequence\nconservation (<10% identities), structural superposition revealed that a cluster of five\nsuccessive transmembrane helices of these proteins adopt comparable topologies,\nwhereas the remaining regions diverge in both fold and number of transmembrane\nhelices (Supplementary Fig. 5). Notably, all these structurally related proteins belong\nto lipid phosphatase/phosphotransferase (LPT) family, most members of which are\nintegral membrane enzymes catalyzing the dephosphorylation or phosphotransfer\nreactions of diverse phospho-lipid or phospho-carbohydrate substrate [6]. This suggests\nthat these enzymes (including Aur1, LpxE, G6PC1, PgpB, SMSr) share a conserved\nstructural module, which we term the LPT fold. This fold exhibits several common\nfeatures (Supplementary Fig. 5): it comprises five transmembrane helices, often\nsituated in the C-terminal region (except in G6PC1); the luminal loop (e.g., EL4 in Aur1)\nconnecting the last two helices harbors a catalytically essential histidine residue (e.g.,\nH294 in Aur1; see Discussions for further mechanistic details). Despite this conserved\ncore, notable structural variations exist. The orientation and length of the topologically\nequivalent transmembrane helices differ among family members. For instance, the first\nhelix of the module is continuous in SMSr (TM5) but kinked in Aur1 (TM4), LpxE\n(TM1), G6PC1 (TM1), and PgpB (TM2), reflecting structural variability within this\nshared scaffold (see Discussion for further comparison).”\nNotably, this analysis also provides critical clues for elucidating its catalytic\nmechanism, as further discussed in our comparative analysis of the active sites in the\nrevised manuscript (Lines 509-529 on Page 18-19; Supplementary Fig. 13).\nSupplementary Figure 5. Structural comparison of IPC synthase Aur1 with\nmembers of the lipid phosphatases/phosphotransferases (LPT) family: LpxE,\nG6PC1, PgpB and SMSr.\nA-E, Comparison of the putative shared folds between IPC synthase Aur1 (A) and four\nmembrane-integrated LPT family enzymes identified as top DALI matches (listed in\ndescending order of Z-score) [1,6]: the lipid A 1-phosphatase LpxE (PDB ID: 6EBU)\n(B), the glucose-6-phosphatase G6PC1 (PDB ID: 9J7V) (C), the\nphosphatidylglycerolphosphate phosphatase PgpB (PDB ID: 5JKI) (D) and\nsphingomyelin synthase-related protein 1 SMSr (PDB ID: 8IJQ) (E). The superimposed\nstructures are shown separately in panels (A) to (E). Their corresponding elements are\nin the same color scheme, whereas elements out of the shared fold are in either grey\nsurface or cartoon representation as indicated. The TM helices are denoted by numbers.\nTop (each panel): side view parallel to the membrane plane.\nBottom (each panel): bottom view from the luminal side, with transmembrane (TM)\nsegments within the shared fold highlighted in gray shading and the absolutely\nconserved catalytic histidine residue shown as magenta sticks.\nReference:\n[1] Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351–W355\n(2016).\n[2] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[3] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[4] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\n[5] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[6] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n2. The identification of a phospholipid and two alkyl chains at the Aur1-Kei subunit\ninterface is of interest. However, the two lone alkyl chains are a little unusual. Are these\nattached to a phospholipid whose head group is disordered?\nResponse: We thank the reviewer for raising this important point. We agree that the\ntwo lone alkyl chains likely belong to phospholipid with disordered head group, which\nmay result from a lack of stabilizing interactions for the head group. Such phenomena\nhave been documented in previous structural studies [1-3]. Accordingly, we have\nrevised the manuscript to explicitly mention this possibility (Line 252-255 on Page 9).\nReference:\n[1] Yu, J. et al. Hippocampal AMPA receptor assemblies and mechanism of allosteric\ninhibition. Nature 594, 448-453 (2021).\n[2] Frieg, B. et al. Cryo-EM structures of lipidic fibrils of amyloid-β (1-40). Nature\nCommunications 15, 1297 (2024).\n[3] Srivastava, D. K. et al. Structure of the human dopamine transporter and\nmechanisms of inhibition. Nature 632, 672-677 (2024).\n3. The authors suggest that the catalytic triad of His255, His294 and Asp298 catalyse\ninositol phosphate transfer. A reaction schematic to show the catalytic reaction\nmechanism would be helpful.\nResponse: We thank the reviewer for this valuable suggestion. Accordingly, we have\nadded a reaction schematic (Figure 3I) to illustrate the proposed catalytic mechanism\ninvolving the His255-His294-Asp298 triad. The mechanism has been incorporated into\nthe revised manuscript (Lines 348-355, Page 12-13) as follows:\n“We propose a two-step catalytic mechanism for IPC synthesis (Fig. 3I). In the first\nstep, D298 activates H294 to initiate a nucleophilic attack on the phosphate group of\nPI, leading to the formation of an enzyme–phosphoinositol intermediate and the release\nof diacylglycerol (DAG). In the second step, H255 activates the hydroxyl group of an\nincoming ceramide, which attacks the intermediate and transfers the phosphorylinositol\nmoiety to ceramide, ultimately forming inositol phosphorylceramide (IPC).”\nFigure 3I. Proposed two-step catalytic mechanism mediated by the H-H-D triad in\nthe Aur1 subunit of IPC synthase. In the first step, D298 activates H294 for\nnucleophilic attack on the phosphate group of phosphatidylinositol (PI), leading to the\nformation of an enzyme-phosphoinositol intermediate and the release of diacylglycerol\n(DAG). In the second step, H255 activates the hydroxyl group of ceramide, which\nattacks the enzyme-bound intermediate to generate inositol phosphorylceramide (IPC).\n4. Where is the binding site for phosphatidylinositol in Aur1? How is IPC synthase\nspecific for phosphatidylinositol and not able use phosphatidylcholine (PC) or\nphosphatidylethanolamine?\nResponse: We thank the reviewer for raising these important questions. To address\nthese points, we performed molecular docking of PI into the Aur1 structure\n(Supplementary Fig. 9A). The result indicates that PI occupies the reaction chamber\nidentified in our ceramide-bound IPCS structure. Specifically, the inositol group of PI\nis nestled deep within a pocket formed by transmembrane helices TM5–TM7 and\nluminal loops EL3 and EL4 (Supplementary Fig. 9B-C). The shape of this pocket\nappears to optimally accommodate the bulkier inositol group of PI (Supplementary\nFig. 9C-D), likely underlying the observed phospholipid preference (Fig. 1C).\nImportantly, in this binding pose, the phosphate group of PI is positioned in close\nproximity to the catalytically essential residues H255 and H294 (Supplementary Fig.\n9D), which is consistent with our proposed reaction mechanism described above (Fig.\n3I).\nThe corresponding descriptions have been incorporated into the revised manuscript\n(Lines 334-347, Page 12; Supplementary Fig. 9).\nSupplementary Figure 9. Phosphorylinositol (PI) binding in IPC synthase (IPCS)\nanalyzed by molecular docking.\nA, Structural model of the PI-bound IPCS complex generated by docking PI into the\nIPCS Aur1 structure. Aur1 is shown as blue cartoon, Kei1 as red surface, and PI as\nyellow sticks.\nB, Cutaway view of the electrostatic surface representation of the PI binding pocket.\nC, Close-up view of the binding pocket for the PI head group, formed by\ntransmembrane helices TM5-TM7 and luminal loops EL3 and EL4. The catalytic triad\nresidues (Aur1 H255, H294 and D298) are shown as sticks. The phosphate group of PI\nis positioned in close proximity to H255 and H294.\nD, Cytoplasmic view of the binding pocket accommodating the inositol moiety of PI.\nThe pocket shape appears fit the bulkier inositol group.\nE, Chemical structure of phospholipids: phosphorylinositol (PI),\nphosphotidylethanolamine (PE), and phosphotidylcholine (PC). The distinct\nphosphoryl head groups are highlighted with dashed boxes.\n5. The authors suggest that in the presence of AbA, the aromatic side chain of Phe 158\nrotates 90°. It is not entirely obvious that this would be clear at 3.5 A resolution.\nThe authors should show cryo-EM density that supports this conclusion.\nResponse: We thank the reviewer for this important comment. In response, we have\nnow included the corresponding cryo-EM densities in Supplementary Fig. 11J–L to\ndirectly support the observed conformational change of Phe158 and other key residues.\nNotably, this is done with our new, high-occupancy AbA-bound complex structure\nthrough prolonged drug incubation under the same conditions, which is obtained as\nsuggested by Reviewer 2 (Point #4). This improved structure exhibits clearer density\nfor the intact drug molecule, which confirms our identified drug-binding site and\nsupports our proposed mechanism of drug inhibition and resistance. Therefore, we\nbwe\nbelieve that this would be better to demonstrate the conformational change resulting\nfrom AbA binding. Based on this high-occupancy structure, we have re-analyzed the\nconformational changes of AbA-binding residues between the AbA-bound and\nceramide-bound states (Fig. 4F) and have demonstrated the corresponding densities in\nSupplementary Fig. 11J-L.\n6. I didn’t see reference in Supplementary Fig. 2.\nResponse: We apologize for the oversight. We have added the references in\nSupplementary Fig. 2.\n7. ‘Comparation’ is not an English word\nResponse: We applogize for this oversight. We have corrected this typo throughout the\nmansucript.\nReviewer #2 (Remarks to the Author):\nThis study presents cryo-EM structures of IPC synthase in complex with its substrate\nceramide and the inhibitor AbA. The authors successfully identified the substrate-\nbinding cavity and the AbA inhibitor pocket, providing critical insights into the\nenzyme’s catalytic mechanism. Complementary biochemical and cell-based assays\nrobustly validate the essential role of IPC synthase catalytic activity in fungal\nsphingolipid biosynthesis. Notably, the resolution of a high-resolution structure for this\n~71 kDa membrane protein—achieved through the combination of an ALFA epitope tag\nand a high-affinity nanobody—represents a significant methodological advancement in\nthe field of membrane protein structural biology.\nOverall, the topic is of interest given the essential role of IPC synthase in fungal\nsphingolipid metabolism and its potential as an antifungal drug target. However, I note\nthat a closely related study by Wu et al. (Nature Communications, 2025) has already\nreported high-resolution structures of the Aur1–Kei1 complex bound to AbA, identifying\nthe catalytic H–H–D triad, key resistance mutations, and the dimeric architecture.\nTherefore, my comments will focus more on the novelty part of this work relative to\nWu’s paper.\nResponse: We thank the reviewer for the support and the important comments to\nimprove the manuscript. We have revised our manuscript accordingly.\nMajor points:\n1. A major concern with the current manuscript is the insufficient acknowledgment of\ncritical related literature. Wu et al. reported cryo-EM structures of the S. cerevisiae\nAur1–Kei1 complex bound to Aureobasidin A, describing the same protein target and\ninhibitor, and this work should be explicitly cited in both the Introduction and\nDiscussion. The authors need to carefully compare their findings with those of Wu et al.\nto highlight both consistencies (such as the catalytic H–H–D triad, AbA resistance\nmutations, and overall architecture) and differences (including the ceramide-bound\nstate and lipid-mediated assembly) to avoid overstating novelty.\nResponse: We thank the reviewer for this important comment. We fully agree that the\nstudy by Wu et al. should be appropriately acknowledged and compared. As\nrecommended, we have revised the manuscript to explicitly compare our study with\nthis recently published work and to clarify our unique contributions (see Lines 530-563\non Page 19-20) as mentioned below:\nDuring the preparation of this manuscript, Wu et al. reported the cryo-EM structure\nof the S. cerevisiae Aur1–Kei1 IPC synthase complex (IPCS) bound to Aureobasidin A\n(AbA) [1]. To overcome challenges posed by the small size of IPCS (~71 kd) and the\nabsence of prominent soluble domain for particle alignment, these two studies used\ndistinct strategies. Wu et al. achieved structure determination by imposing C2\nsymmetry on a dimerized complex, but beyond the AbA-bound form, they were unable\nto resolve additional functional states due to the lack of AbA-mediated stabilization [1].\nIn contrast, we employed an independent approach utilizing an ALFA tag and a\nnanobody as fiducial markers (Supplementary Fig. 1), which enabled us to capture\nmultiple functional states of IPCS, including both the AbA- and ceramide-bound forms\n(Fig. 3, Fig. 4).\nDespite being determined independently, the structural insights from AbA-bound\nstates from both studies are in excellent agreement, uncovering consistent overall\narchitecture, drug-binding site, and mechanisms of drug inhibition and resistance.\nNotably, our structural and functional analyses revealed a stabilizing role of lipids at\nthe Aur1-Kei1 interface (Fig. 2F-J), a feature implicated but not explored in another\nstudy [1].\nBeyond these consensus findings, our work provides several unique insights. While\nanalysis of catalytic mechanism was almost not included in their study, Wu et al.\nproposed that Aur1 and SMSr share similar substrate-binding pockets and catalytic\nmechanism based on analogy with the substrate-bound SMSr structure [1]. In contrast,\nour ceramide-bound Aur1 structure—exclusively obtained in this study—reveals an\nadapted reaction chamber and ceramide-binding mode (Supplementary Fig. 12).\nMoreover, this divergence also extends to the catalytic mechanism (Supplementary\nFig. 13): while SMSr relies on additional catalytic residues (E205 and D295) absent in\nAur1 and canonical LPT members [2-3], our data supports an alternative mechanism\nconsistent with that of canonical LPT enzymes (see discussions below for more detailed\ncomparison between Aur1 and SMSr).\nImportantly, by comparing our AbA-bound and ceramide-bound states, we directly\nvisualize the competitive relationship between substrate and drug (Fig. 4E). This\ncomparison analysis further captures the conformational dynamics within Aur1,\nrevealing how AbA binding induces pronounced rearrangements in key drug-\ninteracting residues and the adjacent TM2–TM3 region (Fig. 4E-F). The exclusive\ncomparison of multiple functional states advances the molecular insights into AbA\nbinding and AbA-mediated inhibition.\nReference:\n[1] Wu, X., Gong, X. & Xie, T. Mechanisms of aureobasidin A inhibition and drug\nresistance in a fungal IPC synthase complex. Nature Communications 16, 5010 (2025).\n[2] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[3] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\nIn addition, because IPC synthase belongs to the sphingomyelin synthase (SMS) family,\nreaders would benefit from a broader discussion that integrates previous structural and\nbiochemical studies on SMS enzymes. Such a comparison could illuminate how the H–\nH–D catalytic triad and substrate positioning in IPC synthase relate to SMS enzymes,\nand whether conserved or divergent features explain differences in substrate specificity.\nIncorporating these key references and analyses will provide proper context, clarify the\nunique contributions of the present study, and strengthen its impact.\nResponse: We thank the reviewer for this insightful suggestion. Following this\ncomment and a related point from Reviewer 1 (Point #1), we have now incorporated a\nthorough comparison between IPC synthase and structural homologs (with low\nsequence homologies) identified through DALI analysis [1], including the human\nsphingomyelin synthase (SMS) family member-SMSr [2]. This analysis not only\nhighlights the structural novelty of Aur1 and Kei1 but also identifies a conserved LPT\n(lipid phosphatase/phosphotransferase) fold within the catalytic subunit Aur1, offering\nkey mechanistic clues for understanding its catalytic mechanism (see also our response\nto Reviewer 1, Point #1).\nFurthermore, in direct response to this suggestion, we have conducted a detailed\ncomparative analysis between IPCS Aur1 and SMSr [2]. These results help delineate\ntheir conserved and notable divergent features (as described in the revised manuscript\non Lines 488-529, Pages 17–19), as listed below:\n(1) Structural comparison between IPCS Aur1 and SMSr:\nDespite their limited sequence identity (<10%) between IPCS Aur1 and SMSr,\ntheir\nstructures can be superimposed with an RMSD of 3.5 Å over 264 aligned residues\n(Supplementary Fig. 12A-B). Beyond the conserved LPT fold (Supplementary Fig.\n5), TM1 of SMSr can also align with TM3 of Aur1, though the latter become\ndiscontinuous in Aur1. Despite their overall resemblance, key differences emerge near\nthe reaction chamber (Supplementary Fig. 12C-E). First, in Aur1, TM4—which lines\nthe side wall of the reaction chamber and corresponds to the continuous TM2 in\nSMSr—is interrupted and kinked by a unique ‘Gly-Ala-Pro’ loop. Second, the extended\nluminal EL3 loop of Aur1, which caps the reaction chamber, adopts a markedly\ndifferent conformation from the corresponding EL2 of SMSr. Consistent with these\nstructural variations, although ceramide occupies comparable positions within the\nreaction chambers, its binding pose differs between the two enzymes. Given their\ndistinct substrate specificities (PI for Aur1 versus PE for SMSr), it is conceivable that\nthe structural differences surrounding the reaction chamber may underlie this\ndivergence, a hypothesis that merits further investigation.\n(2) Comparison of the catalytic mechanisms between IPCS Aur1 and SMSr\nConsistent with the shared LPT module (Supplementary Fig. 5), detailed structural\ncomparisons of Aur1 with other LPT members further reveal that the H-H-D catalytic\ntriad (comprising one histidine from the C2 motif and a histidine-aspartate pair from\nthe ‘HXXXD’ C3 motif) is conserved and arranged in a remarkably similar spatial\nconfiguration [3] (Supplementary Fig. 13). In particular, the catalytic histidine within\nthe C3 motif (e.g., H294 in Aur1) is absolutely conserved, suggesting a unified catalytic\nmechanism. In our proposed catalytic model for Aur1 (Fig. 3I), the invariant C3\nhistidine (H294), activated by D298 in the same motif, acts as a nucleophile to form a\ntransient phosphohistidine intermediate, while the C2 histidine (H255) facilitates\nfurther phosphate transfer. This mechanism is consistent with those established for LPT\nenzymes such as LpxE, G6PC1, and PgpB, among others [3-6]. In contrast, SMSr has\nbeen proposed to employ a distinct catalytic strategy, wherein two catalytic residues\n(first, C2 histidine H301, activated by E205; second, D295, activated by C3 H344 and\nD348) mediates two sequential catalytic steps [2]. It is noted that these additional\ncatalytic residues in SMSr (E205 and D295) are outside the C2 and C3 motifs and are\nnot conserved in Aur1 and several LPT members such as LpxE, G6PC1 and PgpB\n(Supplementary Fig. 13B-F). These functional distinctions suggest that divergent\ncatalytic mechanism may occur in the context of conserved H-H-D catalytic triad,\nwarranting further investigation.\nSupplementary Figure 12. Structural comparison of fungal IPC synthase Aur1\nand human sphingomyelin synthase-related protein 1 (SMSr).\nA-B, Structural superposition of Aur1 (blue cartoon with ceramide in deep-blue sticks;\nthis study) and SMSr (yellow cartoon with ceramide in orange sticks; PDB ID: 8IJQ),\nwith an RMSD of 3.5 Å. The dashed line denotes a similar overall architecture between\nAur1 and SMSr, comprising the LPT fold (see Supplementary Fig. 5 for details) and\none N-terminal TM helix (Aur1 TM3 and SMSr TM1). Beyond this shared fold, Aur1\nhas two additional transmembrane helices (TM1-TM2) when compared to SMSr.\nC-D, Close-up view of the reaction chambers between Aur1 (C) and SMSr (D), as\nmarked by the box in (A). Residues proposed for catalysis are shown as sticks, with\nblue indicating residues conserved in both enzymes and yellow indicating residues\nuniquely proposed for SMSr catalysis [2].\nE, Structure-based sequence alignment of the shared fold between Aur1 and SMSr as\noutlined in (A). The alignment was first generated in DALI server and then illustrated\nusing ESPript3 server. Catalytic residues in Aur1 (red triangles) and SMSr [2] (red\nsquares) are labeled. Shown above and below the alignment are TM helices derived\nfrom Aur1 structure (blue bar) and SMSr structure (yellow bar), respectively.\nSupplementary Figure 13. Comparison of catalytic sites among top structural\nmatches of IPCS Aur1 identified by DALI analysis.\nA, Sequence alignment of the C2 and C3 catalytic motifs in IPCS Aur1 and its structural\nhomologs identified by DALI analysis (listed in descending order of Z-score): LpxE\n(Uniprot ID: O67603), G6PC1 (Uniprot ID: P35575), PgpB (Uniprot ID: O34349) and\nSMSr (Uniprot ID: Q96LT4). Residues of the conserved H-H-D calytic triad are marked\nby triangles.\nB-F, Structural superposition of IPC synthase Aur1 (B) and its structural homologs\nidentified by DALI analysis [1] (listed in descending order of Z-score): LpxE (PDB ID:\n6EBU) (C), G6PC1 (PDB ID: 9J7V) (D), PgpB (PDB ID: 5JKI) (E) and SMSr (PDB\nID: 8IJQ) (F). The superimposed structures are displayed separately in cartoon\nrepresentation, following the same style as in Supplementary Fig. 5, with\ntransmembrane segments of the shared LPT fold highlighted in gray shading. Using the\nfive proposed catalytic residues of SMSr (F) as reference, spatially equivalent residues\nin other structures (B–E) are colored according to conservation status: yellow for\nconserved, blue for non-conserved. Notably, while the H-H-D catalytic triad is highly\nconserved [3], the two additional residues (E205 and D295) proposed to be unique to\nSMSr catalysis [2] (F) are not present in the other LPT members (B-E).\nReference:\n[1] Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351-W355\n(2016).\n[2] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[3] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n[4] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[5] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[6] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\n2. The capture of ceramide in the active site is indeed a valuable addition not present\nin Wu et al., and the authors support their proposed structural model and catalytic\nmechanism with mutagenesis of key residues in the reaction chamber, which resulted in\nreduced activity. However, to strengthen this conclusion, they should provide evidence\nthat the observed effects are not due to global misfolding of the enzyme but rather\nspecific to the catalytic center—for example, by assessing the stability or expression of\nthe purified mutant proteins with size-exclusion FPLC.\nMoreover, the H157A mutant retains ~50% activity, showing the least impairment\namong the catalytic site variants; a brief discussion of why different mutations within\nthe catalytic center yield varied activity reductions would be helpful.\nFinally, it remains uncertain whether the ceramide density reflects a true catalytic\nintermediate or a trapped state introduced during purification and exposure to\ndetergents or exogenous lipids. The authors should discuss the likelihood of ceramide\nincorporation under their preparation conditions and address how well this state\nrepresents in vivo substrate binding.\nResponse: We thank the review for these insightful comments, which have been\naddressed accordingly in our revised manuscript.\nRegarding the concern about potential misfolding of the mutants, we first verified\nthat all key mutants were expressed at levels comparable to the wild-type (WT) enzyme,\nas shown by western blot analysis (Supplementary Fig. 2C). Furthermore, in direct\nresponse to the reviewer's suggestion, we purified these mutants and assessed their\nstates by size-exclusion FPLC (Superose 6 Increase 10/300 GL column). The elution\nprofiles of all mutants were similar to that of the WT complex (Supplementary Fig.\n2D), which is consistent with their maintained subunit stoichiometry (Supplementary\nFig. 2B). Together, these data support the proper folding of all mutants analyzed. We\nhave incorporated these results and corresponding discussion into the revised\nmanuscript (Lines 312-315, Page 11).\nAbout the varied activity impairment of different catalytic site mutants, it shows a\nhierarchy of functional contributions that offers valuable insights into the catalytic\nmechanism. Mutations H255A, H294A, and D298A each completely abolished IPC\nsynthase activity (Fig. 3G-H), supporting them as essential members of the catalytic\ntriad. In contrast, other active-site mutations (H293A, H157A, and S253A) led to\nvarying degrees of partial activity loss, with H157A, located farthest from the catalytic\ntriad, showing the least impairment (~50% residual activity) (Fig. 3G-H). This\ngradation in mutational effects suggests that while the catalytic triad (H255, H294,\nD298) is indispensable for the core enzymatic reaction, neighboring residues such as\nH157, S253, and H293 also contribute to activity. These latter auxiliary residues may\nhelp maintain optimal catalytic geometry, fine-tune substrate orientation, or stabilize\nthe active-site environment, though their precise mechanistic roles await further\nelucidation. We have included a discussion of this interpretation in the revised\nmanuscript (Lines 315-333, Page 11-12).\nAbout ceramide, following reviewer’s suggestion, we have considered it carefully\nand would like to present the following evidence supporting its incorporation and\nfunctional relevance. As detailed in the Methods section (Lines 615-622, Page 22),\nceramide was the only exogenous lipid added throughout the purification procedure for\npreparing the ceramide-bound complex. While LMNG (lauryl maltose neopentyl glycol)\nand CHS (cholesteryl hemisuccinate) were employed as two essential purification\ndetergents, the well-defined electron density closely matches the ceramide structure and\ncannot be reasonably fitted to either detergent (Supplementary Fig. 4B and 4D):\nneither to CHS with a distinct sterol scaffold, nor to LMNG with bulky maltose-based\nheadgroup that is also sterically incompatible with the narrow, groove-shaped active\nsite. Importantly, the ceramide headgroup is positioned in proximity to the catalytic\ntriad, consistent with the proposed reaction mechanism of IPC synthase and canonical\nLPT enzymes [1] (Fig. 3D and 3I). Moreover, the bound ceramide conformation\nsterically clashes with AbA, suggesting a competitive inhibition mechanism for this\ndrug (Fig. 4E). Together, these evidences strongly support ceramide incorporation as\nwell as the functional relevance of the observed ceramide-bound state. We have\nincluded a discussion of this interpretation in the revised manuscript (Lines 287-295,\nPage 10-11).\nSupplementary Figure 2B-D. Expression level and state analysis of IPCS and its\nmutants of candidate catalytic sites.\nB, SDS-PAGE analysis of purified wild-type IPCS and mutants of candidate catalytic\nsites.\nC, Western blot analysis of the expression levels of Aur1 (Flag-tagged) and its mutants\nfrom (B), using anti-Flag antibody. GAPDH was used as load control.\nD, Size-exclusion chromatography (SEC) profile of wild-type IPCS and its mutants\nfrom (B), analyzed on a Superose 6 Increase 10/300 GL column.\nSupplementary Figure 4. Ceramide and lipid interactions of IPC synthase.\nA, Top, the lipid binding site at the interface between Aur1 and Kei1 in electrostatic\nsurface representation. One phospholipid and two lipid alkyl chains (shown as yellow\nsticks) are identified at this interface. Bottom, the density of the lipids.\nB, Top, the ceramide binding site in Aur1 in electrostatic surface representation. Bottom,\nthe density of the ceramide.\nC, Mass-spectrometric analysis of lipids co-purified with IPC synthase. PG,\nphosphatidylglycerols; BMP, bis(monoacylglycerol)phosphate; SL, sulfatides; CL,\ncardiolipins; FA, Fatty acid; PEO, alkyI PE; PE, phosphatidylethanolamines; PI,\nphosphatidylinositols; GM3, monosialogangliosides; LPI, lyso-PI; LPE, lyso-PE; LPA,\nlyso-PA; PA, phosphatidic acids; LPS, lyso-PS; PS, phosphatidylserines; Sph,\nsphingosine; Cer, ceramides; GlcCer, Glucosylceramide; GalCer, galactosylceramides;\nLacCer, lactosylceramides; Gb3, Ceramide trihexoside; LPC, lyso-PC; PCO, alkyl PC;\nPC, phosphatidylcholines; SM, sphingomyelins.\nD, Chemical structure of two detergent molecules used for the purification of IPC\nsynthase: Lauryl Maltose Neopentyl Glycol, LMNG; Cholesteryl hemisuccinate, CHS.\nReference:\n[1] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n3. Another point that warrants attention is the interpretation of the lipid-mediated\nheterodimer stabilization of the Aur1–Kei1 complex. The identification of a\nphospholipid density bridging the interface is intriguing. However, several concerns\nremain. First, the exact identity of the lipid is not determined, and it is unclear whether\nthis represents a physiologically specific cofactor, a detergent-derived molecule, or a\ngeneral phospholipid from the purification system. The authors should clarify the\nlipid assignment and, if possible, provide mass spectrometry or biochemical analysis to\nstrengthen this claim.\nSecond, it is important to discuss whether this lipid-mediated interaction is unique\nto IPC synthase or could represent a more general feature among the sphingomyelin\nsynthase family.\nFinally, the authors should place their findings in context with prior work:\nGong et al. noted lipid-like densities at the interface but did not ascribe functional\nsignificance, so a more explicit comparison would help delineate what is novel here.\nOverall, while the lipid-mediated stabilization model is a potentially important advance,\nits physiological specificity and mechanistic implications require more careful\ndiscussion and validation.\nResponse: We thank the review for these insightful suggestions. We have revised\naccordingly.\nAbout the assignment of the phospholipid, we have considered it carefully and\nwould like to present the following consideration. It is important to note that no\nexogenous phospholipids were added during the purification of either the ceramide-\nbound or AbA-bound complexes. The consistent presence of similar lipid-like densities\nin both structural states supports the identification as an endogenous phospholipid.\nFurthermore, although LMNG and CHS were used as detergents during purification,\nthe observed density matches a phospholipid structure and cannot be reasonably fitted\nto either CHS (with a distinct sterol scaffold) or LMNG (with bulky maltose-based\nheadgroup) (Supplementary Fig. 4A, 4D). Moreover, following reviewer’s suggestion,\nwe also performed lipidomics analysis via mass spectrometry of lipids co-purified with\nIPC synthase. The result confirmed the presence of several lipid species, most notably\nphospholipid, providing additional support for our lipid assignment (Supplementary\nFig. 4C). We have included a discussion of this interpretation in the revised manuscript\n(Lines 255-263, Page 9-10).\nAs for the uniqueness of this lipid-mediated interaction, the key lipid-interacting\nresidues in Aur1 (K274 and K314) are highly conserved across IPC synthases from\nmajor fungal pathogens (Supplementary Fig. 7), yet are not conserved in\nsphingomyelin synthase family members such as SMSr (Supplementary Fig. 12E).\nFurthermore, the lipid-mediated stabilization occurs specifically between the catalytic\nAur1 and regulatory Kei1 subunits. This heteromeric architecture appears to be a\ndistinctive feature of IPC synthase, as sphingomyelin synthases like SMSr function\nwithout regulatory partners [1] (Supplementary Fig. 5A, 5E). Therefore, this lipid-\nmediated inter-subunit interaction appear a general mechanism among fungal IPC\nsynthases but is absent in the sphingomyelin synthase family. We have included a\ndiscussion of this interpretation in the revised manuscript (Lines 457-463, Page 16).\nAs for the third suggestion, in response, we have now added an explicit comparison\nwith the study by Gong et al. in the revised manuscript (Lines 530-563, Pages 19-20;\nplease see also our response to Reviewer 2, Point #1), including a specific discussion\nof these interface lipids (Lines 544-546, Page 19). While Gong et al. briefly mentioned\nlipid-like densities at the Aur1–Kei1 interface, they did not assign these densities or\nexplore their functional relevance [2]. In contrast, our work combines structural and\nconservation analysis (Fig. 2F–H) and functional/biochemical assays (Fig. 2I–J;\nSupplementary Fig. 4C) to assign these lipid densities and to identify lipid-interacting\nresidues, highlighting lipid-mediated heterodimer stabilization at the Aur1–Kei1\ninterface. Notably, our study and Gong et al. employed independent structural\ndetermination strategies (please see more details about this in our response to Reviewer\ninterfacial lipid density [2]. This consistency reinforces the validity of the lipid-\nmediated stabilization mechanism.\nSupplementary Figure 4. Ceramide and lipid interactions of IPC synthase.\nA, Top, the lipid binding site at the interface between Aur1 and Kei1 in electrostatic\nsurface representation. One phospholipid and two lipid alkyl chains (shown as yellow\nsticks) are identified at this interface. Bottom, the density of the lipids.\nB, Top, the ceramide binding site in Aur1 in electrostatic surface representation. Bottom,\nthe density of the ceramide.\nC, Mass-spectrometric analysis of lipids co-purified with IPC synthase. PG,\nphosphatidylglycerols; BMP, bis(monoacylglycerol)phosphate; SL, sulfatides; CL,\ncardiolipins; FA, Fatty acid; PEO, alkyI PE; PE, phosphatidylethanolamines; PI,\nphosphatidylinositols; GM3, monosialogangliosides; LPI, lyso-PI; LPE, lyso-PE; LPA,\nlyso-PA; PA, phosphatidic acids; LPS, lyso-PS; PS, phosphatidylserines; Sph,\nsphingosine; Cer, ceramides; GlcCer, Glucosylceramide; GalCer, galactosylceramides;\nLacCer, lactosylceramides; Gb3, Ceramide trihexoside; LPC, lyso-PC; PCO, alkyl PC;\nPC, phosphatidylcholines; SM, sphingomyelins.\nD, Chemical structure of two detergent molecules used for the purification of IPC\nsynthase: Lauryl Maltose Neopentyl Glycol, LMNG; Cholesteryl hemisuccinate, CHS.\nReference:\n[1] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[2] Wu, X., Gong, X. & Xie, T. Mechanisms of aureobasidin A inhibition and drug\nresistance in a fungal IPC synthase complex. Nature Communications 16, 5010 (2025).\n4. In the AbA-bound IPC synthase complex (Fig. 4), the reported unassigned density\nwithin the catalytic cavity does not convincingly fit the 3D structure of the AbA inhibitor.\nThis discrepancy raises critical concerns regarding the interpretation of the AbA-bound\nstate. Specifically: (1) does the IPCS–ALFA construct reduce AbA binding affinity\ncompared to the wild-type enzyme, and (2) could AbA be hydrolyzed within the catalytic\ncavity, thereby explaining the incomplete density? If either hypothesis is plausible, the\nauthors should provide biochemical validation: for example, quantifying AbA binding\naffinity (e.g., Kd) for IPCS–ALFA versus wild-type using SPR or ITC, or testing for AbA\nhydrolysis by LC–MS to detect cleavage products. If neither explanation holds, the\nauthors must rigorously demonstrate that the unassigned density corresponds to bound\nAbA rather than endogenous lipids co-purified with the protein. The current assertion\nthat AbA binding is “dynamic” is not sufficiently supported. Rigorous validation of this\ndensity is essential to substantiate the proposed inhibition mechanism.\nResponse: We thank the reviewer for raising this important point. Our biochemical data\nshowed that the ALFA-tagged IPCS construct maintained catalytic efficiency and AbA\nsensitivity comparable to the wild-type enzyme, regardless of whether the Nb was\nbound (Fig. 4B). This indicates that neither the tag nor Nb binding impaired enzyme\nfunction or inhibitor binding. In addition, the compound we used consistently exhibited\nstable inhibitory activity in our assays and to our knowledge, there appears no previous\nstudies reporting the hydrolysis of AbA.\nTo directly address the reviewer’s concern regarding the incomplete density, we\nsystematically optimized the sample preparation procedure. By extending the AbA\nincubation time from 1 to 3 hours, we achieved substantially improved drug occupancy,\nwhich allowed us to determine the complex structure at an overall resolution of 3.5 Å\n(Supplementary Fig. 11A–H). The resulting cryo-EM map displayed clear and\ncontinuous density corresponding to the intact AbA molecule (Supplementary\nFig. 11I), indicating that the previously observed partial density likely resulted from a\nlow-occupancy state of the inhibitor.\nThis high-occupancy AbA-bound structure reveals detailed interactions of AbA\nwithin the Aur1 reaction chamber, involving multiple residues, and confirms that H157\nand F158 (located on the linker between TM4a and TM4b) constitute the key binding\nsite (Fig. 4D), consistent with our original proposal. These findings further underscore\nthe functional importance of these two residues (H157 and F158) in the development\nof AbA resistance (Fig. 4H), as proposed in our original and current manuscript.\nNotably, by comparing this new AbA-bound structure with our ceramide-bound\nIPCS structure, we directly demonstrate the competitive relationship between substrate\nand inhibitor (Fig. 4E), as well as pronounced conformational rearrangements in key\ndrug-interacting residues and the adjacent TM2–TM3 region (Fig. 4E-F). These\nstructural insights substantially advances the molecular insights into AbA binding and\nAbA-mediated inhibition..\nThe related description of this new AbA-bound state has been incorporated into the\nrevised manuscript (Lines 388-426, Page 14-15).\nFig. 4B, Inhibition of IPC synthase activity by AbA, analyzed by TLC using fluorescent\nC6-NBD- ceramide. ALFA-tagged IPCS (IPCS-ALFA) and its nanobody-bound form\n(IPCS-ALFA + Nb) show comparable activity and drug sensitivity to the wild-type\nIPCS.\nSupplementary Figure 11. Cryo-EM structural determination of S. cerevisiae IPC\nsynthase (IPCS-ALFA + Nb) complexed with Aureobasidin A (AbA).\nA, A representative cryo-EM micrograph.\nB, Representative 2D class averages.\nC, Flow chart of cryo-EM data acquisition and data processing.\nD, Local resolution distribution of the final cryo-EM map.\nE, The gold-standard Fourier shell correlation curve of the reconstructed map.\nF, Angular particle distribution of particles for the final map reconstruction.\nG-H, Fit of cryo-EM map with structure models of Aur1 (G) and Kei1 (H).\nI, Two views of the modeled AbA superimposed with its corresponding density, with\nthe chemical structure of AbA shown on the right.\nSome minor points:\nFig. 2B (SDS-PAGE) and Figs. 2I-J (Co-IP):\nIn the SDS-PAGE results of Fig. 2B (lane for purified Aur1), what do the high-\nmolecular-weight bands above the Aur1 band represent? Similar bands are observed\nin the co-immunoprecipitation assays (Figs. 2I-J). Do they correspond to glycosylated\nforms of Aur1? If so, glycan density should be detectable in your cryo-EM maps, why\nis this density absent in the manuscript? What is the biological significance of Aur1\nglycosylation?\nResponse: We thank the reviewer for this insightful comment. In response, we have\nperformed additional analyses of Aur1 glycosylation, which are now included in the\nrevised manuscript (Lines 107-114, Page 4; Lines 151-159, Page 6).\nSDS–PAGE and western blot analyses of the purified IPCS complex revealed that\nAur1, but not Kei1, displayed several high–molecular–weight species (Supplementary\nFig. 1G-H). Treatment of the purified complex with PNGase F eliminated these higher–\nmolecular-weight forms of Aur1, confirming that they arise from N-linked\nglycosylation (Supplementary Fig. 1G-H). Enzymatic assays further showed that\nremoval of N-glycans had minimal impact on IPC synthase activity (Supplementary\nFig. 1I). We have included a discussion of this interpretation in the revised manuscript\n(Lines 107-114, Page 4).\nNotably, although we confirmed the presence of N-glycosylation on Aur1\n(Supplementary Fig. 1G-H), the specific glycosylation site could not be\nunambiguously assigned. Among the three predicted glycosylation sites (N132, N243,\nand N367), N367 is located in a disordered region of the structure. In contrast, while\nN132 and N243 lie in ordered regions, neither exhibits clear glycan density. This\nsuggests that these sites are either unmodified or contain highly flexible glycans, with\nthe latter consistent with the minimal enzymatic effects observed upon glycan removal\n(Supplementary Fig. 1I). We have included a discussion of this interpretation in the\nrevised manuscript (Lines 151-159, Page 6).\nSupplementary Figure 1G-I. Analysis of Aur1 glycosylation.\nG-H, SDS-PAGE (G) and corresponding immunoblot (H) analysis of purified IPC\nsynthase, treated with or without PNGase F (N-Glycosidase F) as indicated.\nI, In vitro activity analysis of IPC synthase treated with or without PNGase F.\nLine 246: While the Ser residue in the canonical Ser–His–Asp catalytic triad typically\nprovides the nucleophilic hydroxyl oxygen essential for catalysis, the specific role of\nS253 (located in EL3) in Aur1 during sphingolipid synthesis remains unclear. Notably,\nmutagenesis of S253 does not completely abolish enzymatic activity, raising questions\nabout whether it truly functions as the nucleophile in the reaction. To clarify this\ndiscrepancy, the authors could strengthen their study by (i) providing a comparative\nanalysis of S253’s position and function relative to the catalytic Ser residues in classical\ntriads, and (ii) performing additional mutagenesis, such as S253C or S253T\nsubstitutions, to quantify enzymatic activity and evaluate whether side-chain chemistry\ninfluences catalysis. Such data would help resolve whether S253 is directly catalytic or\nplays an auxiliary structural/positioning role in Aur1’s mechanism.\nResponse: We thank the reviewer for these valuable suggestions. In response, we\ngenerated S253C and S253T mutants and quantitatively compared their enzymatic\nactivities with those of the wild-type (WT) and S253A variants. Both S253C and S253T\nmutants retained substantially higher activity than S253A (Supplementary Fig. 8A),\nindicating that S253 is not directly involved in catalysis.\nFurthermore, we subsequently compared the active site of Aur1—comprising S253\nand the His–His–Asp triad (H255, H294, D298)—with those of representative enzymes\ncontaining a canonical Ser–His–Asp triad, and found no structural equivalence between\nthem [1-3] (Supplementary Fig. 8B-D). Notably, we found that IPCS Aur1 shares a\nconserved LPT fold and conserved His-His-Asp catalytic triad with several canonical\nLPT family members [4] (Supplementary Fig. 5, Supplementary Fig. 13). These\nenzymes have been proposed to typically rely on a His–His–Asp triad for catalysis,\nwithout the involvement of a serine residue [4-7].\nTaken together, our mutagenesis and structural analyses indicate that S253 does not\nplay a direct catalytic role in Aur1. These findings have been incorporated into the\nrevised manuscript (Lines 321-328, Page 12).\nSupplementary Figure 8. Analysis of S253 in the reaction chamber of IPCS Aur1.\nA, In vitro activity of wild-type IPC synthase and Aur1 S253 mutants (S253A, S253C,\nand S253T). Left panel: Representative TLC analysis of IPC synthase activity. Right\npanel: Quantitative assessment of enzyme activity derived from TLC results. Activities\nare normalized to that of the wild-type enzyme. Data are presented as mean ± s.d. from\nthree independent experiments (n = 3).\nB, Close-up view of the reaction chamber in IPCS Aur1, with the H-H-D catalytic triad\n(H255, H294 and D298) and the adjacent residue S253 shown as sticks.\nC, Close-up view of the reaction chamber in Vibrio dual lipase/transferase (ValDLT),\nwhich employs a Ser-His-Asp catalytic triad [1] (S153, H393 and D390, shown as\nsticks).\nD, Close-up view of the reaction chamber in lysosomal phospholipase A2 (LPLA2),\nwhich employs a Ser-His-Asp catalytic triad [2] (S165, H359 and D327, shown as\nsticks).\nReference:\n[1] Wang, C., Liu, C., Zhu, X., Peng, Q. & Ma, Q. Catalytic site flexibility facilitates\nthe substrate and catalytic promiscuity of Vibrio dual lipase/transferase. Nat Commun\n14, 4795 (2023).\n[2] Glukhova, A. et al. Structure and function of lysosomal phospholipase A2 and\nlecithin:cholesterol acyltransferase. Nature Communications 6, 6250 (2015).\n[3] Rauwerdink, A. & Kazlauskas, R. J. How the Same Core Catalytic Machinery\nCatalyzes 17 Different Reactions: the Serine-Histidine-Aspartate Catalytic Triad of α/β-\nHydrolase Fold Enzymes. ACS Catalysis 5, 6153-6176 (2015).\n[4] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n[5] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[6] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[7] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nResponse: We appreciate this note and thank all reviewers for their valuable comments\nand suggestions to improving our manuscript.\nPoint-to-point responses to the reviewers’ comments\nWe greatly thank the insightful and valuable suggestions from reviewers. Our responses\nbelow are in blue.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThe manuscript by Hingjun Yu and colleagues describes the cryo-EM structure of the\nfungal inositol phosphorylceramide (IPC) synthase from S. cerevisiae, the enzyme\nresponsible for inositol phosphorylceramide synthesis. IPC is a key component of\nfungal membranes, and is absent from mammalian cells. It is therefore an important\nanti-fungal drug target. The authors find that IPC synthase is a heterodimer of the Aur1\nand Kei1 subunits. The dimer interface is stabilised by a phospholipid and two lipid\nalkyl chains. The authors identify H255, H294 and D98 of the Aur1 subunit as being\nessential for catalysis and suggest that these three residues form a catalytic triad. A\nHHD triad is also found in other fungal IPC synthases despite low sequence similarity.\nThe authors determined a structure of the complex with the antifungal agent\naureobasidin (AbA) and suggest AbA acts as a competitive inhibitor to block ceramide\nbinding. Finally, the authors test models of the Aur1-Kei dimer interface, phospholipid\nbinding and catalysis using mutagenesis.\nIn general this is an interesting paper. The figures are clear and the cryo-EM work is\nsound. Use of the ALFA-binding nanobody to increase size and help alignment was\ninnovative.\nResponse: We thank the reviewer for the support and the valuable comments. They are\nso helpful to improve our manuscript. We have revised our manuscript accordingly.\n1. When describing the structure, the authors do not compare and contrast with other\nmembrane protein structures. A brief comment on the relatedness of IPC synthase to\nother membrane protein structures would be informative and useful, or is this a novel\nstructure?\nResponse: We thank the reviewer for this insightful suggestion. In response, we have\nadded a section comparing the IPC synthase structure with other known membrane\nproteins identified by DALI analysis [1]. This analysis not only underscores the\nstructural novelty of both Aur1 and Kei1 but also reveals a conserved LPT (lipid\nphosphatase/phosphotransferase) fold within the catalytic subunit Aur1. The related\ndescription has been incorporated into the revised manuscript (Lines 190-219 on Page\n7-8; Supplementary Fig. 5) as follows:\n“A DALI server search for the regulatory subunit Kei1 of the IPC heterodimer did not\nyield any structurally similar homologs [1]. In contrast, for the catalytic subunit Aur1,\nalthough the overall architecture of Aur1 differs from previously reported structures, a\nDALI homology search identified several membrane-protein structures that resemble a\nportion of Aur1’s transmembrane topology (residues 143-314) (Supplementary Fig.\n5). These include (in a descending order of Z-score): Lipid A 1-Phosphatase (LpxE),\nGlucose-6-phosphatase (G6PC1), Phosphatidylglycerophosphatase B (PgpB), and\nSphingomyelin synthase-related protein 1 (SMSr) [2-5]. Despite their low sequence\nconservation (<10% identities), structural superposition revealed that a cluster of five\nsuccessive transmembrane helices of these proteins adopt comparable topologies,\nwhereas the remaining regions diverge in both fold and number of transmembrane\nhelices (Supplementary Fig. 5). Notably, all these structurally related proteins belong\nto lipid phosphatase/phosphotransferase (LPT) family, most members of which are\nintegral membrane enzymes catalyzing the dephosphorylation or phosphotransfer\nreactions of diverse phospho-lipid or phospho-carbohydrate substrate [6]. This suggests\nthat these enzymes (including Aur1, LpxE, G6PC1, PgpB, SMSr) share a conserved\nstructural module, which we term the LPT fold. This fold exhibits several common\nfeatures (Supplementary Fig. 5): it comprises five transmembrane helices, often\nsituated in the C-terminal region (except in G6PC1); the luminal loop (e.g., EL4 in Aur1)\nconnecting the last two helices harbors a catalytically essential histidine residue (e.g.,\nH294 in Aur1; see Discussions for further mechanistic details). Despite this conserved\ncore, notable structural variations exist. The orientation and length of the topologically\nequivalent transmembrane helices differ among family members. For instance, the first\nhelix of the module is continuous in SMSr (TM5) but kinked in Aur1 (TM4), LpxE\n(TM1), G6PC1 (TM1), and PgpB (TM2), reflecting structural variability within this\nshared scaffold (see Discussion for further comparison).”\nNotably, this analysis also provides critical clues for elucidating its catalytic\nmechanism, as further discussed in our comparative analysis of the active sites in the\nrevised manuscript (Lines 509-529 on Page 18-19; Supplementary Fig. 13).\nSupplementary Figure 5. Structural comparison of IPC synthase Aur1 with\nmembers of the lipid phosphatases/phosphotransferases (LPT) family: LpxE,\nG6PC1, PgpB and SMSr.\nA-E, Comparison of the putative shared folds between IPC synthase Aur1 (A) and four\nmembrane-integrated LPT family enzymes identified as top DALI matches (listed in\ndescending order of Z-score) [1,6]: the lipid A 1-phosphatase LpxE (PDB ID: 6EBU)\n(B), the glucose-6-phosphatase G6PC1 (PDB ID: 9J7V) (C), the\nphosphatidylglycerolphosphate phosphatase PgpB (PDB ID: 5JKI) (D) and\nsphingomyelin synthase-related protein 1 SMSr (PDB ID: 8IJQ) (E). The superimposed\nstructures are shown separately in panels (A) to (E). Their corresponding elements are\nin the same color scheme, whereas elements out of the shared fold are in either grey\nsurface or cartoon representation as indicated. The TM helices are denoted by numbers.\nTop (each panel): side view parallel to the membrane plane.\nBottom (each panel): bottom view from the luminal side, with transmembrane (TM)\nsegments within the shared fold highlighted in gray shading and the absolutely\nconserved catalytic histidine residue shown as magenta sticks.\nReference:\n[1] Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351–W355\n(2016).\n[2] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[3] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[4] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\n[5] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[6] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n2. The identification of a phospholipid and two alkyl chains at the AurKei subunit\ninterface is of interest. However, the two lone alkyl chains are a little unusual. Are these\nattached to a phospholipid whose head group is disordered?\nResponse: We thank the reviewer for raising this important point. We agree that the\ntwo lone alkyl chains likely belong to phospholipid with disordered head group, which\nmay result from a lack of stabilizing interactions for the head group. Such phenomena\nhave been documented in previous structural studies [1-3]. Accordingly, we have\nrevised the manuscript to explicitly mention this possibility (Line 252-255 on Page 9).\nReference:\n[1] Yu, J. et al. Hippocampal AMPA receptor assemblies and mechanism of allosteric\ninhibition. Nature 594, 448-453 (2021).\n[2] Frieg, B. et al. Cryo-EM structures of lipidic fibrils of amyloid-β (1-40). Nature\nCommunications 15, 1297 (2024).\n[3] Srivastava, D. K. et al. Structure of the human dopamine transporter and\nmechanisms of inhibition. Nature 632, 672-677 (2024).\n3. The authors suggest that the catalytic triad of His255, His294 and Asp298 catalyse\ninositol phosphate transfer. A reaction schematic to show the catalytic reaction\nmechanism would be helpful.\nResponse: We thank the reviewer for this valuable suggestion. Accordingly, we have\nadded a reaction schematic (Figure 3I) to illustrate the proposed catalytic mechanism\ninvolving the His255-His294-Asp298 triad. The mechanism has been incorporated into\nthe revised manuscript (Lines 348-355, Page 12-13) as follows:\n“We propose a two-step catalytic mechanism for IPC synthesis (Fig. 3I). In the first\nstep, D298 activates H294 to initiate a nucleophilic attack on the phosphate group of\nPI, leading to the formation of an enzyme–phosphoinositol intermediate and the release\nof diacylglycerol (DAG). In the second step, H255 activates the hydroxyl group of an\nincoming ceramide, which attacks the intermediate and transfers the phosphorylinositol\nmoiety to ceramide, ultimately forming inositol phosphorylceramide (IPC).”\nFigure 3I. Proposed two-step catalytic mechanism mediated by the H-H-D triad in\nthe Aur1 subunit of IPC synthase. In the first step, D298 activates H294 for\nnucleophilic attack on the phosphate group of phosphatidylinositol (PI), leading to the\nformation of an enzyme-phosphoinositol intermediate and the release of diacylglycerol\n(DAG). In the second step, H255 activates the hydroxyl group of ceramide, which\nattacks the enzyme-bound intermediate to generate inositol phosphorylceramide (IPC).\n4. Where is the binding site for phosphatidylinositol in Aur1? How is IPC synthase\nspecific for phosphatidylinositol and not able use phosphatidylcholine (PC) or\nphosphatidylethanolamine?\nResponse: We thank the reviewer for raising these important questions. To address\nthese points, we performed molecular docking of PI into the Aur1 structure\n(Supplementary Fig. 9A). The result indicates that PI occupies the reaction chamber\nidentified in our ceramide-bound IPCS structure. Specifically, the inositol group of PI\nis nestled deep within a pocket formed by transmembrane helices TM5–TM7 and\nluminal loops EL3 and EL4 (Supplementary Fig. 9B-C). The shape of this pocket\nappears to optimally accommodate the bulkier inositol group of PI (Supplementary\nFig. 9C-D), likely underlying the observed phospholipid preference (Fig. 1C).\nImportantly, in this binding pose, the phosphate group of PI is positioned in close\nproximity to the catalytically essential residues H255 and H294 (Supplementary Fig.\n9D), which is consistent with our proposed reaction mechanism described above (Fig.\n3I).\nThe corresponding descriptions have been incorporated into the revised manuscript\n(Lines 334-347, Page 12; Supplementary Fig. 9).\nSupplementary Figure 9. Phosphorylinositol (PI) binding in IPC synthase (IPCS)\nanalyzed by molecular docking.\nA, Structural model of the PI-bound IPCS complex generated by docking PI into the\nIPCS Aur1 structure. Aur1 is shown as blue cartoon, Kei1 as red surface, and PI as\nyellow sticks.\nB, Cutaway view of the electrostatic surface representation of the PI binding pocket.\nC, Close-up view of the binding pocket for the PI head group, formed by\ntransmembrane helices TM5-TM7 and luminal loops EL3 and EL4. The catalytic triad\nresidues (Aur1 H255, H294 and D298) are shown as sticks. The phosphate group of PI\nis positioned in close proximity to H255 and H294.\nD, Cytoplasmic view of the binding pocket accommodating the inositol moiety of PI.\nThe pocket shape appears fit the bulkier inositol group.\nE, Chemical structure of phospholipids: phosphorylinositol (PI),\nphosphotidylethanolamine (PE), and phosphotidylcholine (PC). The distinct\nphosphoryl head groups are highlighted with dashed boxes.\n5. The authors suggest that in the presence of AbA, the aromatic side chain of Phe 158\nrotates 90°. It is not entirely obvious that this would be clear at 3.5 A resolution.\nThe authors should show cryo-EM density that supports this conclusion.\nResponse: We thank the reviewer for this important comment. In response, we have\nnow included the corresponding cryo-EM densities in Supplementary Fig. 11J–L to\ndirectly support the observed conformational change of Phe158 and other key residues.\nNotably, this is done with our new, high-occupancy AbA-bound complex structure\nthrough prolonged drug incubation under the same conditions, which is obtained as\nsuggested by Reviewer 2 (Point #4). This improved structure exhibits clearer density\nfor the intact drug molecule, which confirms our identified drug-binding site and\nsupports our proposed mechanism of drug inhibition and resistance. Therefore,nbwe\nbelieve that this would be better to demonstrate the conformational change resulting\nfrom AbA binding. Based on this high-occupancy structure, we have re-analyzed the\nconformational changes of AbA-binding residues between the AbA-bound and\nceramide-bound states (Fig. 4F) and have demonstrated the corresponding densities in\nSupplementary Fig. 11J-L.\n6. I didn’t see reference in Supplementary Fig. 2.\nResponse: We apologize for the oversight. We have added the references in\nSupplementary Fig. 2.\n7. ‘Comparation’ is not an English word\nResponse: We applogize for this oversight. We have corrected this typo throughout the\nmansucript.\nReviewer #2 (Remarks to the Author):\nThis study presents cryo-EM structures of IPC synthase in complex with its substrate\nceramide and the inhibitor AbA. The authors successfully identified the substrate-\nbinding cavity and the AbA inhibitor pocket, providing critical insights into the\nenzyme’s catalytic mechanism. Complementary biochemical and cell-based assays\nrobustly validate the essential role of IPC synthase catalytic activity in fungal\nsphingolipid biosynthesis. Notably, the resolution of a high-resolution structure for this\n~71 kDa membrane protein—achieved through the combination of an ALFA epitope tag\nand a high-affinity nanobody—represents a significant methodological advancement in\nthe field of membrane protein structural biology.\nOverall, the topic is of interest given the essential role of IPC synthase in fungal\nsphingolipid metabolism and its potential as an antifungal drug target. However, I note\nthat a closely related study by Wu et al. (Nature Communications, 2025) has already\nreported high-resolution structures of the Aur1–Kei1 complex bound to AbA, identifying\nthe catalytic H–H–D triad, key resistance mutations, and the dimeric architecture.\nTherefore, my comments will focus more on the novelty part of this work relative to\nWu’s paper.\nResponse: We thank the reviewer for the support and the important comments to\nimprove the manuscript. We have revised our manuscript accordingly.\nMajor points:\n1. A major concern with the current manuscript is the insufficient acknowledgment of\ncritical related literature. Wu et al. reported cryo-EM structures of the S. cerevisiae\nAur1–Kei1 complex bound to Aureobasidin A, describing the same protein target and\ninhibitor, and this work should be explicitly cited in both the Introduction and\nDiscussion. The authors need to carefully compare their findings with those of Wu et al.\nto highlight both consistencies (such as the catalytic H–H–D triad, AbA resistance\nmutations, and overall architecture) and differences (including the ceramide-bound\nstate and lipid-mediated assembly) to avoid overstating novelty.\nResponse: We thank the reviewer for this important comment. We fully agree that the\nstudy by Wu et al. should be appropriately acknowledged and compared. As\nrecommended, we have revised the manuscript to explicitly compare our study with\nthis recently published work and to clarify our unique contributions (see Lines 530-563\non Page 19-20) as mentioned below:\nDuring the preparation of this manuscript, Wu et al. reported the cryo-EM structure\nof the S. cerevisiae Aur1–Kei1 IPC synthase complex (IPCS) bound to Aureobasidin A\n(AbA) [1]. To overcome challenges posed by the small size of IPCS (~71 kd) and the\nabsence of prominent soluble domain for particle alignment, these two studies used\ndistinct strategies. Wu et al. achieved structure determination by imposing C2\nsymmetry on a dimerized complex, but beyond the AbA-bound form, they were unable\nto resolve additional functional states due to the lack of AbA-mediated stabilization [1].\nIn contrast, we employed an independent approach utilizing an ALFA tag and a\nnanobody as fiducial markers (Supplementary Fig. 1), which enabled us to capture\nmultiple functional states of IPCS, including both the AbA- and ceramide-bound forms\n(Fig. 3, Fig. 4).\nDespite being determined independently, the structural insights from AbA-bound\nstates from both studies are in excellent agreement, uncovering consistent overall\narchitecture, drug-binding site, and mechanisms of drug inhibition and resistance.\nNotably, our structural and functional analyses revealed a stabilizing role of lipids at\nthe Aur1-Kei1 interface (Fig. 2F-J), a feature implicated but not explored in another\nstudy [1].\nBeyond these consensus findings, our work provides several unique insights. While\nanalysis of catalytic mechanism was almost not included in their study, Wu et al.\nproposed that Aur1 and SMSr share similar substrate-binding pockets and catalytic\nmechanism based on analogy with the substrate-bound SMSr structure [1]. In contrast,\nour ceramide-bound Aur1 structure—exclusively obtained in this study—reveals an\nadapted reaction chamber and ceramide-binding mode (Supplementary Fig. 12).\nMoreover, this divergence also extends to the catalytic mechanism (Supplementary\nFig. 13): while SMSr relies on additional catalytic residues (E205 and D295) absent in\nAur1 and canonical LPT members [2-3], our data supports an alternative mechanism\nconsistent with that of canonical LPT enzymes (see discussions below for more detailed\ncomparison between Aur1 and SMSr).\nImportantly, by comparing our AbA-bound and ceramide-bound states, we directly\nvisualize the competitive relationship between substrate and drug (Fig. 4E). This\ncomparison analysis further captures the conformational dynamics within Aur1,\nrevealing how AbA binding induces pronounced rearrangements in key drug-\ninteracting residues and the adjacent TM2–TM3 region (Fig. 4E-F). The exclusive\ncomparison of multiple functional states advances the molecular insights into AbA\nbinding and AbA-mediated inhibition.\nReference:\n[1] Wu, X., Gong, X. & Xie, T. Mechanisms of aureobasidin A inhibition and drug\nresistance in a fungal IPC synthase complex. Nature Communications 16, 5010 (2025).\n[2] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[3] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\nIn addition, because IPC synthase belongs to the sphingomyelin synthase (SMS) family,\nreaders would benefit from a broader discussion that integrates previous structural and\nbiochemical studies on SMS enzymes. Such a comparison could illuminate how the H–\nH–D catalytic triad and substrate positioning in IPC synthase relate to SMS enzymes,\nand whether conserved or divergent features explain differences in substrate specificity.\nIncorporating these key references and analyses will provide proper context, clarify the\nunique contributions of the present study, and strengthen its impact.\nResponse: We thank the reviewer for this insightful suggestion. Following this\ncomment and a related point from Reviewer 1 (Point #1), we have now incorporated a\nthorough comparison between IPC synthase and structural homologs (with low\nsequence homologies) identified through DALI analysis [1], including the human\nsphingomyelin synthase (SMS) family member-SMSr [2]. This analysis not only\nhighlights the structural novelty of Aur1 and Kei1 but also identifies a conserved LPT\n(lipid phosphatase/phosphotransferase) fold within the catalytic subunit Aur1, offering\nkey mechanistic clues for understanding its catalytic mechanism (see also our response\nto Reviewer 1, Point #1).\nFurthermore, in direct response to this suggestion, we have conducted a detailed\ncomparative analysis between IPCS Aur1 and SMSr [2]. These results help delineate\ntheir conserved and notable divergent features (as described in the revised manuscript\non Lines 488-529, Pages 17–19), as listed below:\n(1) Structural comparison between IPCS Aur1 and SMSr:\nDespite their limited sequence identity (<10%) between IPCS Aur1 and SMSr,\ntheir\nstructures can be superimposed with an RMSD of 3.5 Å over 264 aligned residues\n(Supplementary Fig. 12A-B). Beyond the conserved LPT fold (Supplementary Fig.\n5), TM1 of SMSr can also align with TM3 of Aur1, though the latter become\ndiscontinuous in Aur1. Despite their overall resemblance, key differences emerge near\nthe reaction chamber (Supplementary Fig. 12C-E). First, in Aur1, TM4—which lines\nthe side wall of the reaction chamber and corresponds to the continuous TM2 in\nSMSr—is interrupted and kinked by a unique ‘Gly-Ala-Pro’ loop. Second, the extended\nluminal EL3 loop of Aur1, which caps the reaction chamber, adopts a markedly\ndifferent conformation from the corresponding EL2 of SMSr. Consistent with these\nstructural variations, although ceramide occupies comparable positions within the\nreaction chambers, its binding pose differs between the two enzymes. Given their\ndistinct substrate specificities (PI for Aur1 versus PE for SMSr), it is conceivable that\nthe structural differences surrounding the reaction chamber may underlie this\ndivergence, a hypothesis that merits further investigation.\n(2) Comparison of the catalytic mechanisms between IPCS Aur1 and SMSr\nConsistent with the shared LPT module (Supplementary Fig. 5), detailed structural\ncomparisons of Aur1 with other LPT members further reveal that the H-H-D catalytic\ntriad (comprising one histidine from the C2 motif and a histidine-aspartate pair from\nthe ‘HXXXD’ C3 motif) is conserved and arranged in a remarkably similar spatial\nconfiguration [3] (Supplementary Fig. 13). In particular, the catalytic histidine within\nthe C3 motif (e.g., H294 in Aur1) is absolutely conserved, suggesting a unified catalytic\nmechanism. In our proposed catalytic model for Aur1 (Fig. 3I), the invariant C3\nhistidine (H294), activated by D298 in the same motif, acts as a nucleophile to form a\ntransient phosphohistidine intermediate, while the C2 histidine (H255) facilitates\nfurther phosphate transfer. This mechanism is consistent with those established for LPT\nenzymes such as LpxE, G6PC1, and PgpB, among others [3-6]. In contrast, SMSr has\nbeen proposed to employ a distinct catalytic strategy, wherein two catalytic residues\n(first, C2 histidine H301, activated by E205; second, D295, activated by C3 H344 and\nD348) mediates two sequential catalytic steps [2]. It is noted that these additional\ncatalytic residues in SMSr (E205 and D295) are outside the C2 and C3 motifs and are\nnot conserved in Aur1 and several LPT members such as LpxE, G6PC1 and PgpB\n(Supplementary Fig. 13B-F). These functional distinctions suggest that divergent\ncatalytic mechanism may occur in the context of conserved H-H-D catalytic triad,\nwarranting further investigation.\nSupplementary Figure 12. Structural comparison of fungal IPC synthase Aur1\nand human sphingomyelin synthase-related protein 1 (SMSr).\nA-B, Structural superposition of Aur1 (blue cartoon with ceramide in deep-blue sticks;\nthis study) and SMSr (yellow cartoon with ceramide in orange sticks; PDB ID: 8IJQ),\nwith an RMSD of 3.5 Å. The dashed line denotes a similar overall architecture between\nAur1 and SMSr, comprising the LPT fold (see Supplementary Fig. 5 for details) and\none N-terminal TM helix (Aur1 TM3 and SMSr TM1). Beyond this shared fold, Aur1\nhas two additional transmembrane helices (TM1-TM2) when compared to SMSr.\nC-D, Close-up view of the reaction chambers between Aur1 (C) and SMSr (D), as\nmarked by the box in (A). Residues proposed for catalysis are shown as sticks, with\nblue indicating residues conserved in both enzymes and yellow indicating residues\nuniquely proposed for SMSr catalysis [2].\nE, Structure-based sequence alignment of the shared fold between Aur1 and SMSr as\noutlined in (A). The alignment was first generated in DALI server and then illustrated\nusing ESPript3 server. Catalytic residues in Aur1 (red triangles) and SMSr [2] (red\nsquares) are labeled. Shown above and below the alignment are TM helices derived\nfrom Aur1 structure (blue bar) and SMSr structure (yellow bar), respectively.\nSupplementary Figure 13. Comparison of catalytic sites among top structural\nmatches of IPCS Aur1 identified by DALI analysis.\nA, Sequence alignment of the C2 and C3 catalytic motifs in IPCS Aur1 and its structural\nhomologs identified by DALI analysis (listed in descending order of Z-score): LpxE\n(Uniprot ID: O67603), G6PC1 (Uniprot ID: P35575), PgpB (Uniprot ID: O34349) and\nSMSr (Uniprot ID: Q96LT4). Residues of the conserved H-H-D calytic triad are marked\nby triangles.\nB-F, Structural superposition of IPC synthase Aur1 (B) and its structural homologs\nidentified by DALI analysis [1] (listed in descending order of Z-score): LpxE (PDB ID:\n6EBU) (C), G6PC1 (PDB ID: 9J7V) (D), PgpB (PDB ID: 5JKI) (E) and SMSr (PDB\nID: 8IJQ) (F). The superimposed structures are displayed separately in cartoon\nrepresentation, following the same style as in Supplementary Fig. 5, with\ntransmembrane segments of the shared LPT fold highlighted in gray shading. Using the\nfive proposed catalytic residues of SMSr (F) as reference, spatially equivalent residues\nin other structures (B–E) are colored according to conservation status: yellow for\nconserved, blue for non-conserved. Notably, while the H-H-D catalytic triad is highly\nconserved [3], the two additional residues (E205 and D295) proposed to be unique to\nSMSr catalysis [2] (F) are not present in the other LPT members (B-E).\nReference:\n[1] Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351-W355\n(2016).\n[2] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[3] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n[4] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[5] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[6] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\n2. The capture of ceramide in the active site is indeed a valuable addition not present\nin Wu et al., and the authors support their proposed structural model and catalytic\nmechanism with mutagenesis of key residues in the reaction chamber, which resulted in\nreduced activity. However, to strengthen this conclusion, they should provide evidence\nthat the observed effects are not due to global misfolding of the enzyme but rather\nspecific to the catalytic center—for example, by assessing the stability or expression of\nthe purified mutant proteins with size-exclusion FPLC.\nMoreover, the H157A mutant retains ~50% activity, showing the least impairment\namong the catalytic site variants; a brief discussion of why different mutations within\nthe catalytic center yield varied activity reductions would be helpful.\nFinally, it remains uncertain whether the ceramide density reflects a true catalytic\nintermediate or a trapped state introduced during purification and exposure to\ndetergents or exogenous lipids. The authors should discuss the likelihood of ceramide\nincorporation under their preparation conditions and address how well this state\nrepresents in vivo substrate binding.\nResponse: We thank the review for these insightful comments, which have been\naddressed accordingly in our revised manuscript.\nRegarding the concern about potential misfolding of the mutants, we first verified\nthat all key mutants were expressed at levels comparable to the wild-type (WT) enzyme,\nas shown by western blot analysis (Supplementary Fig. 2C). Furthermore, in direct\nresponse to the reviewer's suggestion, we purified these mutants and assessed their\nstates by size-exclusion FPLC (Superose 6 Increase 10/300 GL column). The elution\nprofiles of all mutants were similar to that of the WT complex (Supplementary Fig.\n2D), which is consistent with their maintained subunit stoichiometry (Supplementary\nFig. 2B). Together, these data support the proper folding of all mutants analyzed. We\nhave incorporated these results and corresponding discussion into the revised\nmanuscript (Lines 312-315, Page 11).\nAbout the varied activity impairment of different catalytic site mutants, it shows a\nhierarchy of functional contributions that offers valuable insights into the catalytic\nmechanism. Mutations H255A, H294A, and D298A each completely abolished IPC\nsynthase activity (Fig. 3G-H), supporting them as essential members of the catalytic\ntriad. In contrast, other active-site mutations (H293A, H157A, and S253A) led to\nvarying degrees of partial activity loss, with H157A, located farthest from the catalytic\ntriad, showing the least impairment (~50% residual activity) (Fig. 3G-H). This\ngradation in mutational effects suggests that while the catalytic triad (H255, H294,\nD298) is indispensable for the core enzymatic reaction, neighboring residues such as\nH157, S253, and H293 also contribute to activity. These latter auxiliary residues may\nhelp maintain optimal catalytic geometry, fine-tune substrate orientation, or stabilize\nthe active-site environment, though their precise mechanistic roles await further\nelucidation. We have included a discussion of this interpretation in the revised\nmanuscript (Lines 315-333, Page 11-12).\nAbout ceramide, following reviewer’s suggestion, we have considered it carefully\nand would like to present the following evidence supporting its incorporation and\nfunctional relevance. As detailed in the Methods section (Lines 615-622, Page 22),\nceramide was the only exogenous lipid added throughout the purification procedure for\npreparing the ceramide-bound complex. While LMNG (lauryl maltose neopentyl glycol)\nand CHS (cholesteryl hemisuccinate) were employed as two essential purification\ndetergents, the well-defined electron density closely matches the ceramide structure and\ncannot be reasonably fitted to either detergent (Supplementary Fig. 4B and 4D):\nneither to CHS with a distinct sterol scaffold, nor to LMNG with bulky maltose-based\nheadgroup that is also sterically incompatible with the narrow, groove-shaped active\nsite. Importantly, the ceramide headgroup is positioned in proximity to the catalytic\ntriad, consistent with the proposed reaction mechanism of IPC synthase and canonical\nLPT enzymes [1] (Fig. 3D and 3I). Moreover, the bound ceramide conformation\nsterically clashes with AbA, suggesting a competitive inhibition mechanism for this\ndrug (Fig. 4E). Together, these evidences strongly support ceramide incorporation as\nwell as the functional relevance of the observed ceramide-bound state. We have\nincluded a discussion of this interpretation in the revised manuscript (Lines 287-295,\nPage 10-11).\nSupplementary Figure 2B-D. Expression level and state analysis of IPCS and its\nmutants of candidate catalytic sites.\nB, SDS-PAGE analysis of purified wild-type IPCS and mutants of candidate catalytic\nsites.\nC, Western blot analysis of the expression levels of Aur1 (Flag-tagged) and its mutants\nfrom (B), using anti-Flag antibody. GAPDH was used as load control.\nD, Size-exclusion chromatography (SEC) profile of wild-type IPCS and its mutants\nfrom (B), analyzed on a Superose 6 Increase 10/300 GL column.\nSupplementary Figure 4. Ceramide and lipid interactions of IPC synthase.\nA, Top, the lipid binding site at the interface between Aur1 and Kei1 in electrostatic\nsurface representation. One phospholipid and two lipid alkyl chains (shown as yellow\nsticks) are identified at this interface. Bottom, the density of the lipids.\nB, Top, the ceramide binding site in Aur1 in electrostatic surface representation. Bottom,\nthe density of the ceramide.\nC, Mass-spectrometric analysis of lipids co-purified with IPC synthase. PG,\nphosphatidylglycerols; BMP, bis(monoacylglycerol)phosphate; SL, sulfatides; CL,\ncardiolipins; FA, Fatty acid; PEO, alkyI PE; PE, phosphatidylethanolamines; PI,\nphosphatidylinositols; GM3, monosialogangliosides; LPI, lyso-PI; LPE, lyso-PE; LPA,\nlyso-PA; PA, phosphatidic acids; LPS, lyso-PS; PS, phosphatidylserines; Sph,\nsphingosine; Cer, ceramides; GlcCer, Glucosylceramide; GalCer, galactosylceramides;\nLacCer, lactosylceramides; Gb3, Ceramide trihexoside; LPC, lyso-PC; PCO, alkyl PC;\nPC, phosphatidylcholines; SM, sphingomyelins.\nD, Chemical structure of two detergent molecules used for the purification of IPC\nsynthase: Lauryl Maltose Neopentyl Glycol, LMNG; Cholesteryl hemisuccinate, CHS.\nReference:\n[1] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n3. Another point that warrants attention is the interpretation of the lipid-mediated\nheterodimer stabilization of the Aur1–Kei1 complex. The identification of a\nphospholipid density bridging the interface is intriguing. However, several concerns\nremain. First, the exact identity of the lipid is not determined, and it is unclear whether\nthis represents a physiologically specific cofactor, a detergent-derived molecule, or a\ngeneral phospholipid from the purification system. The authors should clarify the\nlipid assignment and, if possible, provide mass spectrometry or biochemical analysis to\nstrengthen this claim.\nSecond, it is important to discuss whether this lipid-mediated interaction is unique\nto IPC synthase or could represent a more general feature among the sphingomyelin\nsynthase family.\nFinally, the authors should place their findings in context with prior work:\nGong et al. noted lipid-like densities at the interface but did not ascribe functional\nsignificance, so a more explicit comparison would help delineate what is novel here.\nOverall, while the lipid-mediated stabilization model is a potentially important advance,\nits physiological specificity and mechanistic implications require more careful\ndiscussion and validation.\nResponse: We thank the review for these insightful suggestions. We have revised\naccordingly.\nAbout the assignment of the phospholipid, we have considered it carefully and\nwould like to present the following consideration. It is important to note that no\nexogenous phospholipids were added during the purification of either the ceramide-\nbound or AbA-bound complexes. The consistent presence of similar lipid-like densities\nin both structural states supports the identification as an endogenous phospholipid.\nFurthermore, although LMNG and CHS were used as detergents during purification,\nthe observed density matches a phospholipid structure and cannot be reasonably fitted\nto either CHS (with a distinct sterol scaffold) or LMNG (with bulky maltose-based\nheadgroup) (Supplementary Fig. 4A, 4D). Moreover, following reviewer’s suggestion,\nwe also performed lipidomics analysis via mass spectrometry of lipids co-purified with\nIPC synthase. The result confirmed the presence of several lipid species, most notably\nphospholipid, providing additional support for our lipid assignment (Supplementary\nFig. 4C). We have included a discussion of this interpretation in the revised manuscript\n(Lines 255-263, Page 9-10).\nAs for the uniqueness of this lipid-mediated interaction, the key lipid-interacting\nresidues in Aur1 (K274 and K314) are highly conserved across IPC synthases from\nmajor fungal pathogens (Supplementary Fig. 7), yet are not conserved in\nsphingomyelin synthase family members such as SMSr (Supplementary Fig. 12E).\nFurthermore, the lipid-mediated stabilization occurs specifically between the catalytic\nAur1 and regulatory Kei1 subunits. This heteromeric architecture appears to be a\ndistinctive feature of IPC synthase, as sphingomyelin synthases like SMSr function\nwithout regulatory partners [1] (Supplementary Fig. 5A, 5E). Therefore, this lipid-\nmediated inter-subunit interaction appear a general mechanism among fungal IPC\nsynthases but is absent in the sphingomyelin synthase family. We have included a\ndiscussion of this interpretation in the revised manuscript (Lines 457-463, Page 16).\nAs for the third suggestion, in response, we have now added an explicit comparison\nwith the study by Gong et al. in the revised manuscript (Lines 530-563, Pages 19-20;\nplease see also our response to Reviewer 2, Point #1), including a specific discussion\nof these interface lipids (Lines 544-546, Page 19). While Gong et al. briefly mentioned\nlipid-like densities at the Aur1–Kei1 interface, they did not assign these densities or\nexplore their functional relevance [2]. In contrast, our work combines structural and\nconservation analysis (Fig. 2F–H) and functional/biochemical assays (Fig. 2I–J;\nSupplementary Fig. 4C) to assign these lipid densities and to identify lipid-interacting\nresidues, highlighting lipid-mediated heterodimer stabilization at the Aur1–Kei1\ninterface. Notably, our study and Gong et al. employed independent structural\ndetermination strategies (please see more details about this in our response to Reviewer\ninterfacial lipid density [2]. This consistency reinforces the validity of the lipid-\nmediated stabilization mechanism.\nSupplementary Figure 4. Ceramide and lipid interactions of IPC synthase.\nA, Top, the lipid binding site at the interface between Aur1 and Kei1 in electrostatic\nsurface representation. One phospholipid and two lipid alkyl chains (shown as yellow\nsticks) are identified at this interface. Bottom, the density of the lipids.\nB, Top, the ceramide binding site in Aur1 in electrostatic surface representation. Bottom,\nthe density of the ceramide.\nC, Mass-spectrometric analysis of lipids co-purified with IPC synthase. PG,\nphosphatidylglycerols; BMP, bis(monoacylglycerol)phosphate; SL, sulfatides; CL,\ncardiolipins; FA, Fatty acid; PEO, alkyI PE; PE, phosphatidylethanolamines; PI,\nphosphatidylinositols; GM3, monosialogangliosides; LPI, lyso-PI; LPE, lyso-PE; LPA,\nlyso-PA; PA, phosphatidic acids; LPS, lyso-PS; PS, phosphatidylserines; Sph,\nsphingosine; Cer, ceramides; GlcCer, Glucosylceramide; GalCer, galactosylceramides;\nLacCer, lactosylceramides; Gb3, Ceramide trihexoside; LPC, lyso-PC; PCO, alkyl PC;\nPC, phosphatidylcholines; SM, sphingomyelins.\nD, Chemical structure of two detergent molecules used for the purification of IPC\nsynthase: Lauryl Maltose Neopentyl Glycol, LMNG; Cholesteryl hemisuccinate, CHS.\nReference:\n[1] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[2] Wu, X., Gong, X. & Xie, T. Mechanisms of aureobasidin A inhibition and drug\nresistance in a fungal IPC synthase complex. Nature Communications 16, 5010 (2025).\n4. In the AbA-bound IPC synthase complex (Fig. 4), the reported unassigned density\nwithin the catalytic cavity does not convincingly fit the 3D structure of the AbA inhibitor.\nThis discrepancy raises critical concerns regarding the interpretation of the AbA-bound\nstate. Specifically: (1) does the IPCS–ALFA construct reduce AbA binding affinity\ncompared to the wild-type enzyme, and (2) could AbA be hydrolyzed within the catalytic\ncavity, thereby explaining the incomplete density? If either hypothesis is plausible, the\nauthors should provide biochemical validation: for example, quantifying AbA binding\naffinity (e.g., Kd) for IPCS–ALFA versus wild-type using SPR or ITC, or testing for AbA\nhydrolysis by LC–MS to detect cleavage products. If neither explanation holds, the\nauthors must rigorously demonstrate that the unassigned density corresponds to bound\nAbA rather than endogenous lipids co-purified with the protein. The current assertion\nthat AbA binding is “dynamic” is not sufficiently supported. Rigorous validation of this\ndensity is essential to substantiate the proposed inhibition mechanism.\nResponse: We thank the reviewer for raising this important point. Our biochemical data\nshowed that the ALFA-tagged IPCS construct maintained catalytic efficiency and AbA\nsensitivity comparable to the wild-type enzyme, regardless of whether the Nb was\nbound (Fig. 4B). This indicates that neither the tag nor Nb binding impaired enzyme\nfunction or inhibitor binding. In addition, the compound we used consistently exhibited\nstable inhibitory activity in our assays and to our knowledge, there appears no previous\nstudies reporting the hydrolysis of AbA.\nTo directly address the reviewer’s concern regarding the incomplete density, we\nsystematically optimized the sample preparation procedure. By extending the AbA\nincubation time from 1 to 3 hours, we achieved substantially improved drug occupancy,\nwhich allowed us to determine the complex structure at an overall resolution of 3.5 Å\n(Supplementary Fig. 11A–H). The resulting cryo-EM map displayed clear and\ncontinuous density corresponding to the intact AbA molecule (Supplementary\nFig. 11I), indicating that the previously observed partial density likely resulted from a\nlow-occupancy state of the inhibitor.\nThis high-occupancy AbA-bound structure reveals detailed interactions of AbA\nwithin the Aur1 reaction chamber, involving multiple residues, and confirms that H157\nand F158 (located on the linker between TM4a and TM4b) constitute the key binding\nsite (Fig. 4D), consistent with our original proposal. These findings further underscore\nthe functional importance of these two residues (H157 and F158) in the development\nof AbA resistance (Fig. 4H), as proposed in our original and current manuscript.\nNotably, by comparing this new AbA-bound structure with our ceramide-bound\nIPCS structure, we directly demonstrate the competitive relationship between substrate\nand inhibitor (Fig. 4E), as well as pronounced conformational rearrangements in key\ndrug-interacting residues and the adjacent TM2–TM3 region (Fig. 4E-F). These\nstructural insights substantially advances the molecular insights into AbA binding and\nAbA-mediated inhibition..\nThe related description of this new AbA-bound state has been incorporated into the\nrevised manuscript (Lines 388-426, Page 14-15).\nFig. 4B, Inhibition of IPC synthase activity by AbA, analyzed by TLC using fluorescent\nC6-NBD- ceramide. ALFA-tagged IPCS (IPCS-ALFA) and its nanobody-bound form\n(IPCS-ALFA + Nb) show comparable activity and drug sensitivity to the wild-type\nIPCS.\nSupplementary Figure 11. Cryo-EM structural determination of S. cerevisiae IPC\nsynthase (IPCS-ALFA + Nb) complexed with Aureobasidin A (AbA).\nA, A representative cryo-EM micrograph.\nB, Representative 2D class averages.\nC, Flow chart of cryo-EM data acquisition and data processing.\nD, Local resolution distribution of the final cryo-EM map.\nE, The gold-standard Fourier shell correlation curve of the reconstructed map.\nF, Angular particle distribution of particles for the final map reconstruction.\nG-H, Fit of cryo-EM map with structure models of Aur1 (G) and Kei1 (H).\nI, Two views of the modeled AbA superimposed with its corresponding density, with\nthe chemical structure of AbA shown on the right.\nSome minor points:\nFig. 2B (SDS-PAGE) and Figs. 2I-J (Co-IP):\nIn the SDS-PAGE results of Fig. 2B (lane for purified Aur1), what do the high-\nmolecular-weight bands above the Aur1 band represent? Similar bands are observed\nin the co-immunoprecipitation assays (Figs. 2I-J). Do they correspond to glycosylated\nforms of Aur1? If so, glycan density should be detectable in your cryo-EM maps, why\nis this density absent in the manuscript? What is the biological significance of Aur1\nglycosylation?\nResponse: We thank the reviewer for this insightful comment. In response, we have\nperformed additional analyses of Aur1 glycosylation, which are now included in the\nrevised manuscript (Lines 107-114, Page 4; Lines 151-159, Page 6).\nSDS–PAGE and western blot analyses of the purified IPCS complex revealed that\nAur1, but not Kei1, displayed several high–molecular–weight species (Supplementary\nFig. 1G-H). Treatment of the purified complex with PNGase F eliminated these higher–\nmolecular-weight forms of Aur1, confirming that they arise from N-linked\nglycosylation (Supplementary Fig. 1G-H). Enzymatic assays further showed that\nremoval of N-glycans had minimal impact on IPC synthase activity (Supplementary\nFig. 1I). We have included a discussion of this interpretation in the revised manuscript\n(Lines 107-114, Page 4).\nNotably, although we confirmed the presence of N-glycosylation on Aur1\n(Supplementary Fig. 1G-H), the specific glycosylation site could not be\nunambiguously assigned. Among the three predicted glycosylation sites (N132, N243,\nand N367), N367 is located in a disordered region of the structure. In contrast, while\nN132 and N243 lie in ordered regions, neither exhibits clear glycan density. This\nsuggests that these sites are either unmodified or contain highly flexible glycans, with\nthe latter consistent with the minimal enzymatic effects observed upon glycan removal\n(Supplementary Fig. 1I). We have included a discussion of this interpretation in the\nrevised manuscript (Lines 151-159, Page 6).\nSupplementary Figure 1G-I. Analysis of Aur1 glycosylation.\nG-H, SDS-PAGE (G) and corresponding immunoblot (H) analysis of purified IPC\nsynthase, treated with or without PNGase F (N-Glycosidase F) as indicated.\nI, In vitro activity analysis of IPC synthase treated with or without PNGase F.\nLine 246: While the Ser residue in the canonical Ser–His–Asp catalytic triad typically\nprovides the nucleophilic hydroxyl oxygen essential for catalysis, the specific role of\nS253 (located in EL3) in Aur1 during sphingolipid synthesis remains unclear. Notably,\nmutagenesis of S253 does not completely abolish enzymatic activity, raising questions\nabout whether it truly functions as the nucleophile in the reaction. To clarify this\ndiscrepancy, the authors could strengthen their study by (i) providing a comparative\nanalysis of S253’s position and function relative to the catalytic Ser residues in classical\ntriads, and (ii) performing additional mutagenesis, such as S253C or S253T\nsubstitutions, to quantify enzymatic activity and evaluate whether side-chain chemistry\ninfluences catalysis. Such data would help resolve whether S253 is directly catalytic or\nplays an auxiliary structural/positioning role in Aur1’s mechanism.\nResponse: We thank the reviewer for these valuable suggestions. In response, we\ngenerated S253C and S253T mutants and quantitatively compared their enzymatic\nactivities with those of the wild-type (WT) and S253A variants. Both S253C and S253T\nmutants retained substantially higher activity than S253A (Supplementary Fig. 8A),\nindicating that S253 is not directly involved in catalysis.\nFurthermore, we subsequently compared the active site of Aur1—comprising S253\nand the His–His–Asp triad (H255, H294, D298)—with those of representative enzymes\ncontaining a canonical Ser–His–Asp triad, and found no structural equivalence between\nthem [1-3] (Supplementary Fig. 8B-D). Notably, we found that IPCS Aur1 shares a\nconserved LPT fold and conserved His-His-Asp catalytic triad with several canonical\nLPT family members [4] (Supplementary Fig. 5, Supplementary Fig. 13). These\nenzymes have been proposed to typically rely on a His–His–Asp triad for catalysis,\nwithout the involvement of a serine residue [4-7].\nTaken together, our mutagenesis and structural analyses indicate that S253 does not\nplay a direct catalytic role in Aur1. These findings have been incorporated into the\nrevised manuscript (Lines 321-328, Page 12).\nSupplementary Figure 8. Analysis of S253 in the reaction chamber of IPCS Aur1.\nA, In vitro activity of wild-type IPC synthase and Aur1 S253 mutants (S253A, S253C,\nand S253T). Left panel: Representative TLC analysis of IPC synthase activity. Right\npanel: Quantitative assessment of enzyme activity derived from TLC results. Activities\nare normalized to that of the wild-type enzyme. Data are presented as mean ± s.d. from\nthree independent experiments (n = 3).\nB, Close-up view of the reaction chamber in IPCS Aur1, with the H-H-D catalytic triad\n(H255, H294 and D298) and the adjacent residue S253 shown as sticks.\nC, Close-up view of the reaction chamber in Vibrio dual lipase/transferase (ValDLT),\nwhich employs a Ser-His-Asp catalytic triad [1] (S153, H393 and D390, shown as\nsticks).\nD, Close-up view of the reaction chamber in lysosomal phospholipase A2 (LPLA2),\nwhich employs a Ser-His-Asp catalytic triad [2] (S165, H359 and D327, shown as\nsticks).\nReference:\n[1] Wang, C., Liu, C., Zhu, X., Peng, Q. & Ma, Q. Catalytic site flexibility facilitates\nthe substrate and catalytic promiscuity of Vibrio dual lipase/transferase. Nat Commun\n14, 4795 (2023).\n[2] Glukhova, A. et al. Structure and function of lysosomal phospholipase A2 and\nlecithin:cholesterol acyltransferase. Nature Communications 6, 6250 (2015).\n[3] Rauwerdink, A. & Kazlauskas, R. J. How the Same Core Catalytic Machinery\nCatalyzes 17 Different Reactions: the Serine-Histidine-Aspartate Catalytic Triad of α/β-\nHydrolase Fold Enzymes. ACS Catalysis 5, 6153-6176 (2015).\n[4] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n[5] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[6] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[7] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nResponse: We appreciate this note and thank all reviewers for their valuable comments\nand suggestions to improving our manuscript.\nPoint-to-point responses to the reviewers’ comments\nWe greatly thank the insightful and valuable suggestions from reviewers. Our responses\nbelow are in blue.\nREVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nThe manuscript by Hingjun Yu and colleagues describes the cryo-EM structure of the\nfungal inositol phosphorylceramide (IPC) synthase from S. cerevisiae, the enzyme\nresponsible for inositol phosphorylceramide synthesis. IPC is a key component of\nfungal membranes, and is absent from mammalian cells. It is therefore an important\nanti-fungal drug target. The authors find that IPC synthase is a heterodimer of the Aur1\nand Kei1 subunits. The dimer interface is stabilised by a phospholipid and two lipid\nalkyl chains. The authors identify H255, H294 and D98 of the Aur1 subunit as being\nessential for catalysis and suggest that these three residues form a catalytic triad. A\nHHD triad is also found in other fungal IPC synthases despite low sequence similarity.\nThe authors determined a structure of the complex with the antifungal agent\naureobasidin (AbA) and suggest AbA acts as a competitive inhibitor to block ceramide\nbinding. Finally, the authors test models of the Aur1-Kei dimer interface, phospholipid\nbinding and catalysis using mutagenesis.\nIn general this is an interesting paper. The figures are clear and the cryo-EM work is\nsound. Use of the ALFA-binding nanobody to increase size and help alignment was\ninnovative.\nResponse: We thank the reviewer for the support and the valuable comments. They are\nso helpful to improve our manuscript. We have revised our manuscript accordingly.\n1. When describing the structure, the authors do not compare and contrast with other\nmembrane protein structures. A brief comment on the relatedness of IPC synthase to\nother membrane protein structures would be informative and useful, or is this a novel\nstructure?\nResponse: We thank the reviewer for this insightful suggestion. In response, we have\nadded a section comparing the IPC synthase structure with other known membrane\nproteins identified by DALI analysis [1]. This analysis not only underscores the\nstructural novelty of both Aur1 and Kei1 but also reveals a conserved LPT (lipid\nphosphatase/phosphotransferase) fold within the catalytic subunit Aur1. The related\ndescription has been incorporated into the revised manuscript (Lines 190-219 on Page\n7-8; Supplementary Fig. 5) as follows:\n“A DALI server search for the regulatory subunit Kei1 of the IPC heterodimer did not\nyield any structurally similar homologs [1]. In contrast, for the catalytic subunit Aur1,\nalthough the overall architecture of Aur1 differs from previously reported structures, a\nDALI homology search identified several membrane-protein structures that resemble a\nportion of Aur1’s transmembrane topology (residues 143-314) (Supplementary Fig.\n5). These include (in a descending order of Z-score): Lipid A 1-Phosphatase (LpxE),\nGlucose-6-phosphatase (G6PC1), Phosphatidylglycerophosphatase B (PgpB), and\nSphingomyelin synthase-related protein 1 (SMSr) [2-5]. Despite their low sequence\nconservation (<10% identities), structural superposition revealed that a cluster of five\nsuccessive transmembrane helices of these proteins adopt comparable topologies,\nwhereas the remaining regions diverge in both fold and number of transmembrane\nhelices (Supplementary Fig. 5). Notably, all these structurally related proteins belong\nto lipid phosphatase/phosphotransferase (LPT) family, most members of which are\nintegral membrane enzymes catalyzing the dephosphorylation or phosphotransfer\nreactions of diverse phospho-lipid or phospho-carbohydrate substrate [6]. This suggests\nthat these enzymes (including Aur1, LpxE, G6PC1, PgpB, SMSr) share a conserved\nstructural module, which we term the LPT fold. This fold exhibits several common\nfeatures (Supplementary Fig. 5): it comprises five transmembrane helices, often\nsituated in the C-terminal region (except in G6PC1); the luminal loop (e.g., EL4 in Aur1)\nconnecting the last two helices harbors a catalytically essential histidine residue (e.g.,\nH294 in Aur1; see Discussions for further mechanistic details). Despite this conserved\ncore, notable structural variations exist. The orientation and length of the topologically\nequivalent transmembrane helices differ among family members. For instance, the first\nhelix of the module is continuous in SMSr (TM5) but kinked in Aur1 (TM4), LpxE\n(TM1), G6PC1 (TM1), and PgpB (TM2), reflecting structural variability within this\nshared scaffold (see Discussion for further comparison).”\nNotably, this analysis also provides critical clues for elucidating its catalytic\nmechanism, as further discussed in our comparative analysis of the active sites in the\nrevised manuscript (Lines 509-529 on Page 18-19; Supplementary Fig. 13).\nSupplementary Figure 5. Structural comparison of IPC synthase Aur1 with\nmembers of the lipid phosphatases/phosphotransferases (LPT) family: LpxE,\nG6PC1, PgpB and SMSr.\nA-E, Comparison of the putative shared folds between IPC synthase Aur1 (A) and four\nmembrane-integrated LPT family enzymes identified as top DALI matches (listed in\ndescending order of Z-score) [1,6]: the lipid A 1-phosphatase LpxE (PDB ID: 6EBU)\n(B), the glucose-6-phosphatase G6PC1 (PDB ID: 9J7V) (C), the\nphosphatidylglycerolphosphate phosphatase PgpB (PDB ID: 5JKI) (D) and\nsphingomyelin synthase-related protein 1 SMSr (PDB ID: 8IJQ) (E). The superimposed\nstructures are shown separately in panels (A) to (E). Their corresponding elements are\nin the same color scheme, whereas elements out of the shared fold are in either grey\nsurface or cartoon representation as indicated. The TM helices are denoted by numbers.\nTop (each panel): side view parallel to the membrane plane.\nBottom (each panel): bottom view from the luminal side, with transmembrane (TM)\nsegments within the shared fold highlighted in gray shading and the absolutely\nconserved catalytic histidine residue shown as magenta sticks.\nReference:\n[1] Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351–W355\n(2016).\n[2] Zhao, J. et al. The Lipid A 1-Phosphatase, LpxE, Functionally Connects Multiple\nLayers of Bacterial Envelope Biogenesis. mBio 10, 10.1128/mbio.00886-00819 (2019).\n[3] Xia, Z. et al. Structural insights into glucose-6-phosphate recognition and\nhydrolysis by human G6PC1. Proceedings of the National Academy of Sciences 122,\ne2418316122 (2025).\n[4] Ghachi, M. E. et al. Crystal structure and biochemical characterization of the\ntransmembrane PAP2 type phosphatidylglycerol phosphate phosphatase from Bacillus\nsubtilis. Cellular and Molecular Life Sciences 74, 2319-2332 (2017).\n[5] Hu, K. et al. Cryo-EM structure of human sphingomyelin synthase and its\nmechanistic implications for sphingomyelin synthesis. Nat Struct Mol Biol 31, 884-895\n(2024).\n[6] Sigal, Y. J., McDermott, M. I. & Morris, A. J. Integral membrane lipid\nphosphatases/phosphotransferases: common structure and diverse functions. Biochem\nJ 387, 281-293 (2005).\n2. The identification of a phospholipid and two alkyl chains at the AurKei subunit\ninterface is of interest. However, the two lone alkyl chains are a little unusual. Are these\nattached to a phospholipid whose head group is disordered?\nResponse: We thank the reviewer for raising this important point. We agree that the\ntwo lone alkyl chains likely belong to phospholipid with disordered head group, which\nmay result from a lack of stabilizing interactions for the head group. Such phenomena\nhave been documented in previous structural studies [1-3]. Accordingly, we have\nrevised the manuscript to explicitly mention this possibility (Line 252-255 on Page 9).\nReference:\n[1] Yu, J. et al. Hippocampal AMPA receptor assemblies and mechanism of allosteric\ninhibition. Nature 594, 448-453 (2021).\n[2] Frieg, B. et al. Cryo-EM structures of lipidic fibrils of amyloid-β (1-40). Nature\nCommunications 15, 1297 (2024).\n[3] Srivastava, D. K. et al. Structure of the human dopamine transporter and\nmechanisms of inhibition. Nature 632, 672-677 (2024).\n3. The authors suggest that the catalytic triad of His255, His294 and Asp298 catalyse\ninositol phosphate transfer. A reaction schematic to show the catalytic reaction\nmechanism would be helpful.\nResponse: We thank the reviewer for this valuable suggestion. Accordingly, we have\nadded a reaction schematic (Figure 3I) to illustrate the proposed catalytic mechanism\ninvolving the His255-His294-Asp298 triad. The mechanism has been incorporated into\nthe revised manuscript (Lines 348-355, Page 12-13) as follows:\n“We propose a two-step catalytic mechanism for IPC synthesis (Fig. 3I). In the first\nstep, D298 activates H294 to initiate a nucleophilic attack on the phosphate group of\nPI, leading to the formation of an enzyme–phosphoinositol intermediate and the release\nof diacylglycerol (DAG). In the second step, H255 activates the hydroxyl group of an\nincoming ceramide, which attacks the intermediate and transfers the phosphorylinositol\nmoiety to ceramide, ultimately forming inositol phosphorylceramide (IPC).”\nFigure 3I. Proposed two-step catalytic mechanism mediated by the H-H-D triad in\nthe Aur1 subunit of IPC synthase. In the first step, D298 activates H294 for\nnucleophilic attack on the phosphate group of phosphatidylinositol (PI), leading to the\nformation of an enzyme-phosphoinositol intermediate and the release of diacylglycerol\n(DAG). In the second step, H255 activates the hydroxyl group of ceramide, which\nattacks the enzyme-bound intermediate to generate inositol phosphorylceramide (IPC).\n4. Where is the binding site for phosphatidylinositol in Aur1? How is IPC synthase\nspecific for phosphatidylinositol and not able use phosphatidylcholine (PC) or\nphosphatidylethanolamine?\nResponse: We thank the reviewer for raising these important questions. To address\nthese points, we performed molecular docking of PI into the Aur1 structure\n(Supplementary Fig. 9A). The result indicates that PI occupies the reaction chamber\nidentified in our ceramide-bound IPCS structure. Specifically, the inositol group of PI\nis nestled deep within a pocket formed by transmembrane helices TM5–TM7 and\nluminal loops EL3 and EL4 (Supplementary Fig. 9B-C). The shape of this pocket\nappears to optimally accommodate the bulkier inositol group of PI (Supplementary\nFig. 9C-D), likely underlying the observed phospholipid preference (Fig. 1C).\nImportantly, in this binding pose, the phosphate group of PI is positioned in close\nproximity to the catalytically essential residues H255 and H294 (Supplementary Fig.\n9D), which is consistent with our proposed reaction mechanism described above (Fig.\n3I).\nThe corresponding descriptions have been incorporated into the revised manuscript\n(Lines 334-347, Page 12; Supplementary Fig. 9).\nSupplementary Figure 9. Phosphorylinositol (PI) binding in IPC synthase (IPCS)\nanalyzed by molecular docking.\nA, Structural model of the PI-bound IPCS complex generated by docking PI into the\nIPCS Aur1 structure. Aur1 is shown as blue cartoon, Kei1 as red surface, and PI as\nyellow sticks.\nB, Cutaway view of the electrostatic surface representation of the PI binding pocket.\nC, Close-up view of the binding pocket for the PI head group, formed by\ntransmembrane helices TM5-TM7 and luminal loops EL3 and EL4. The catalytic triad\nresidues (Aur1 H255, H294 and D298) are shown as sticks. The phosphate group of PI\nis positioned in close proximity to H255 and H294.\nD, Cytoplasmic view of the binding pocket accommodating the inositol moiety of PI.\nThe pocket shape appears fit the bulkier inositol group.\nE, Chemical structure of phospholipids: phosphorylinositol (PI),\nphosphotidylethanolamine (PE), and phosphotidylcholine (PC). The distinct\nphosphoryl head groups are highlighted with dashed boxes.\n5. The authors suggest that in the presence of AbA, the aromatic side chain of Phe 158\nrotates 90°. It is not entirely obvious that this would be clear at 3.5 A resolution.\nThe authors should show cryo-EM density that supports this conclusion.\nResponse: We thank the reviewer for this important comment. In response, we have\nnow included the corresponding cryo-EM densities in Supplementary Fig. 11J–L to\ndirectly support the observed conformational change of Phe158 and other key residues.\nNotably, this is done with our new, high-occupancy AbA-bound complex structure\nthrough prolonged drug incubation under the same conditions, which is obtained as\nsuggested by Reviewer 2 (Point #4). This improved structure exhibits clearer density\nfor the intact drug molecule, which confirms our identified drug-binding site and\nsupports our proposed mechanism of drug inhibition and resistance. Therefore,nbwe\nbelieve that this would be better to demonstrate the conformational change resulting\nfrom AbA binding. Based on this high-occupancy structure, we have re-analyzed the\nconformational changes of AbA-binding residues between the AbA-bound and\nceramide-bound states (Fig. 4F) and have demonstrated the corresponding densities in\nSupplementary Fig. 11J-L.\n6. I didn’t see reference in Supplementary Fig. 2.\nResponse: We apologize for the oversight. We have added the references in\nSupplementary Fig. 2.\n7. ‘Comparation’ is not an English word\nResponse: We applogize for this oversight. We have corrected this typo throughout the\nmansucript.\nReviewer #2 (Remarks to the Author):\nThe authors have thoroughly addressed all of my previous comments, I am satisfied\nwith the revisions, and fully support the publication of this manuscript in Nature\nCommunications.\nResponse: We thank the reviewer for the positive and encouraging assessment. We\nappreciate the reviewer’s support for publication.\nReviewer #3 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports.\nThis is part of the Nature Communications initiative to facilitate training in peer review\nand to provide appropriate recognition for Early Career Researchers who co-review\nmanuscripts.\nResponse: We appreciate this note and thank all reviewers for their valuable comments\nand careful assessments to improve our manuscript."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have done an excellent job of responding to my comments. I recommend this interesting paper for publication in Nature Communications.", "reviewer_2": "The authors have thoroughly addressed all of my previous comments, I am satisfied with the revisions, and fully support the publication of this manuscript in Nature Communications.", "reviewer_3": "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: We appreciate this note and thank all reviewers for their valuable comments and suggestions to improving our manuscript."}]

Decision letter